This work was supported by Public Health Service grants from the National Institute of Health, NIAID (grants A1065530 and Al124649), to K.A. Fields.

1. Design and Assembly of pSUmC-4.0 with Homology Arms Specific to the Gene of Interest
<ol>
	<li>Identify ~3 kb regions directly upstream and downstream of the gene targeted for deletion to serve as the 5&#39; and 3&#39; homology arms for homologous recombination (Figure 1).</li>
	<li>Design PCR primers to 1) amplify the 3 kb 5&#39; homology arm from chlamydial genomic DNA and 2) contain a 15&#8211;30 bp overhang specific to pSUmC-4.0 when digested at the Sall restriction enzyme site. Ensure that the primer regions overlapping with pSUmC-4.0 have melting temperatures of 55 &#176;C and that hairpin melting temperatures for the entire primer are &#60;35 &#176;C.</li>
	<li>Design PCR primers to 1) amplify the 3 kb 3&#39; homology arm from chlamydial genomic DNA and 2) contain a 15&#8211;30 bp overhang specific to pSUmC-4.0 when digested at the SbfI restriction enzyme site. Ensure that the primer regions overlapping with pSUmC-4.0 have melting temperatures of 55 &#176;C and that hairpin melting temperatures for the entire primer are &#60;35 &#176;C.</li>
	<li>Design PCR screening primers that will amplify the insertion of each homology arm into the pSUmC-4.0 vector after a DNA assembly reaction16. Design these primers to anneal ~500 bp outside the restriction sites to allow easy visualization of a PCR product even if the insertion was unsuccessful (Figure 1).</li>
	<li>Amplify the 3 kb 5&#39; and 3&#39; homology arms from freshly extracted chlamydial genomic DNA by PCR in one to two reactions to yield enough fragment for later DNA assembly. Follow the DNA polymerase manufacturer&#39;s protocol for specific reaction set-up and cycling conditions.</li>
	<li>Verify that both PCR products are the correct size and that the reactions do not contain any contaminating bands. Run 5 &#956;L of each PCR reaction on a 1% DNA agarose gel.</li>
	<li>Purify and concentrate the PCR fragments by phenol/chloroform extraction and ethanol precipitation.
	<ol>
		<li>Pool all 3&#39; PCR reactions together in a 1.5 mL tube and bring to a final volume of 200 &#181;L with nuclease-free H2O. Repeat this process for the 5&#39; PCR reactions.</li>
		<li>Add 200 &#181;L of phenol to each tube and vortex for 30&#8211;60 s. Spin each tube in a microcentrifuge at &#62;21,000 x g for 20 min at room temperature (RT).</li>
		<li>Transfer the aqueous phase top layer of each tube into a new 1.5 mL tube, add one-tenth the volume of 3 M sodium acetate to each tube, then flick to mix.</li>
		<li>Add 3 volumes of 100% ethanol to each tube and flick to mix. Incubate both tubes at -80 &#176;C for 20 min.</li>
		<li>Pellet the DNA by spinning each tube at &#62;21,000 x g for 5 min at RT. Discard the supernatant.</li>
		<li>Wash the DNA pellet with 100 &#181;L of 70 % ethanol. Spin each tube at &#62;21,000 x g for 5 min at RT. Discard the supernatant.</li>
		<li>Wash the DNA pellet again with 100 &#181;L of 100% ethanol. Spin each tube at &#62;21,000 for 5 min at RT. Discard the supernatant.</li>
		<li>Let the pellet fully air-dry, then gently resuspend the DNA in 12 &#181;L of nuclease-free H2O by flicking to mix.</li>
	</ol>
	</li>
	<li>Digest 500 ng of pSUmC-4.0 vector in a 20 &#181;L reaction with 1 unit of Sall and 1 unit of SbfI according to the manufacture&#39;s protocol. Incubate the reaction for approximately 3 h or overnight to ensure complete digestion of the vector.</li>
	<li>Purify and concentrate the digested backbone by phenol/chloroform extraction and ethanol precipitation as described earlier (step 1.7.1&#8211;1.7.9).</li>
	<li>Combine the digested pSUmC-4.0 vector and both the 5&#39; and 3&#39; homology arm PCR products together at a ratio of 0.01 pmol pSUmC-4.0 to 0.09 pmol each homology arm. Assemble the fragments in a DNA assembly reaction according to the manufacture&#39;s protocol.</li>
	<li>Transform 50 &#181;L of electrocompetent E. coli with 1 &#181;L of the DNA assembly reaction. Plate the transformed E. coli onto LB agar plates containing 100 &#181;g/mL spectinomycin by spreading 150 &#181;L per plate. Incubate the plates at 37 &#176;C overnight.</li>
	<li>The next day, screen colonies for green and red fluorescence using an epifluorescence inverted microscope. A total of 5&#8211;30 colonies per agar plate is expected.</li>
	<li>Pick 5&#8211;10 colonies to inoculate 4 mL of LB broth supplemented with 100 &#181;g/mL spectinomycin. Incubate cultures at 37 &#176;C overnight with shaking at 250 rpm.</li>
	<li>Using the overnight E. coli cultures, perform small-scale DNA purification. Elute the DNA with 50 &#181;L of nuclease-free H2O.</li>
	<li>Confirm the insertion of each homology arm into the pSUmC-4.0 vector using the PCR screening primers (designed in step 1.4) to amplify each insertion. If DNA assembly is successful, a ~4 kb PCR product should be observed in a 1% DNA agarose gel for both the 5&#39; and 3&#39; arm regions. A ~1 kb PCR product indicates a lack of insertion. 
	NOTE: If the insertion of the homology arms is unsuccessful in a dual fragment assembly reaction, perform two assembly reactions to insert the 5&#39; arm then 3&#39; arm, individually. Digest the vector with only Sall before assembling with the 5&#39; arm, then digest the vector with SbfI to assemble the 3&#39; arm.</li>
	<li>Transform dam-/dcm- competent E. coli with 200 ng of pSUmc-4.0 assembled with both homology arms. Plate the transformed bacteria onto LB agar plates containing 100 &#181;g/mL spectinomycin by spreading 25 &#181;L per plate. Incubate the plates at 37 &#176;C overnight. A total of 20&#8211;200 colonies are expected.</li>
	<li>Screen the plates for red and green fluorescent colonies as described in step 1.13.</li>
	<li>Set up a culture for a large-scale DNA purification by inoculating 100 mL of LB broth containing 100 &#181;g/mL spectinomycin with red and green colonies. Incubate the culture at 37 &#176;C overnight with shaking at 250 rpm.</li>
	<li>Harvest the culture and purify the DNA using a large-scale DNA purification.
	<ol>
		<li>Resuspend the purified plasmid DNA in 100 &#181;L of NF-H2O. A total DNA yield of at least 2 &#181;g/&#181;L is ideal for C. trachomatis transformation.</li>
		<li>Sequence the 5&#39; and 3&#39; homology arm regions to ensure correct assembly into pSUmC-4.0 before transforming C. trachomatis.</li>
	</ol>
	</li>
</ol>
2. Transformation of C. trachomatis with pSUmC 4.0 + Homology Arms for Gene Deletion by Allelic Exchange
<ol>
	<li>Seed McCoy cells, mouse fibroblasts standardly used for cultivating Chlamydia, into two 6 well plates (1 x 106 cells/well) with growth media #1. Incubate the plates at 37 &#176;C with 5% CO2 until the monolayer is confluent (approximately 24 h).</li>
	<li>In a 1.5 mL tube, add a volume of C. trachomatis WT serovar L2 elementary bodies (EBs) that is sufficient to infect 12 wells of a 6 well plate at an MOI of 2. Pellet the EBs at &#62;20,000 x g for 30 min, then discard the supernatant.</li>
	<li>Initiate chlamydial transformation by gently resuspending the EBs in 600 &#181;L of CaCl2 buffer, and then add 24 &#181;g of pSUmC-4.0 + homology arms to the tube. Gently mix the solution by flicking. Incubate the tube at RTfor 30 min and flick to mix every 10 min.</li>
	<li>Add the transformation solution to 24 mL of Hanks&#39; balanced salt solution (HBSS) and mix by gently pipetting. Transfer 2 mL of the inoculum to each well of confluent McCoy cells in the 6 well plates and infect by centrifugation at 900 x g for 1 h at 20 &#176;C.</li>
	<li>Remove the inoculum and replace with 2 mL/well RPMI growth media #2. Incubate the plates at 37 &#176;C with 5% CO2.</li>
	<li>After a minimum of 7 h, but no more than 12 h, replace the media with 2 mL/well of selection media #1. Continue the incubation at 37 &#176;C for 48 h from the time of infection.</li>
	<li>Harvest and passage the bacteria onto a fresh monolayer of McCoy cells.
	<ol>
		<li>Using a cell scraper, gently scrape the McCoy monolayer to lift the cells into the media. Transfer the material from each well into a 2 mL tube. Pellet the cell material at &#62;20,000 x g in a microcentrifuge for 30 min at 4 &#176;C.</li>
		<li>Remove and discard the supernatant. Resuspend the cell pellet in 1 mL of HBSS by gently pipetting.</li>
		<li>Pellet the cell debris at 200 x g for 5 min at 4 &#176;C. Transfer the supernatant onto a fresh well of confluent McCoy cells. Add an additional 1 mL of HBSS to each well for a total volume of 2 mL/well.</li>
		<li>Infect by centrifugation at 900 x g for 1 h at 20 &#176;C. Replace the inoculum with selection media #1 immediately following infection.</li>
	</ol>
	</li>
	<li>Continue passaging the bacteria onto a fresh monolayer of McCoy cells (section 2.7) every 48 h for three or more passages until red and green inclusions are detected using an epifluorescence inverted microscope 24 h post-infection (24 hpi).</li>
	<li>Once red and green inclusions are detected, continue passaging the monolayer (section 2.7) using selection media #2.</li>
	<li>Continue passaging the monolayer until green-only inclusions are detected 24 hpi (passage #3 or later), which indicates the loss of the pSUmC-4.0 vector and incorporation of the loxP selection cassette into the genome.</li>
	<li>Choose one well of green-only inclusions to enrich by passaging. Harvest and freeze any other wells that also contain green-only inclusions in sucrose-phosphate-glutamate buffer (SPG).
	<ol>
		<li>To enrich green-only inclusions, passage the monolayer from one well of a 6 well plate as done in step 2.7, but after pelleting the cell debris (step 2.7.3), transfer the supernatant into a conical with 12 mL of HBSS. Use this inoculum to infect two 6 well plates by adding 2 mL per well, then spin the plates at 900 x g for 60 min at 20 &#176;C.</li>
		<li>To freeze additional wells, harvest the monolayer as in step 2.7.1, but resuspend the pellets in 1 mL of SPG instead of HBSS. Pellet the cell debris (step 2.7.3) and transfer the supernatant into a 1.5 mL cryotube. Freeze the material at -80 &#176;C.</li>
	</ol>
	</li>
	<li>After enriching green-only inclusions to a MOI of 0.5&#8211;1.0 (one to two more passages after detection), harvest the monolayers into SPG as described in step 2.11.2. Obtain aliquots of 50 &#181;L in 1.5 mL tubes and freeze at -80 &#176;C.</li>
	<li>Titer green-only bacteria to determine inclusion, forming units/&#181;L, according to a standard titration protocol17.</li>
</ol>
3. Clonal Isolation of Green-only C. trachomatis Deletion Mutant Containing the loxP Flanked Selection Cassette by Limiting Dilution
<ol>
	<li>Seed a 384 well tissue culture plate with 50 &#181;L of McCoy cells at ~2,000 cells/well in growth media #1. Incubate at 37 &#176;C with 5 % CO2 for ~24 h or until confluent.</li>
	<li>Dilute an aliquot of green-only bacteria in HBSS to achieve ~50 EBs per 20 mL. Transfer the diluted inoculum to a reservoir tray.
	<ol>
		<li>Remove the media from the 384 well plate by firmly flicking the entire plate upside down into a waste container. Using a multichannel pipette, add 50 &#181;L of C. trachomatis inoculum to each well. Infect by centrifugation at 900 x g for 1 h at 20 &#176;C. 
		NOTE: Be mindful not to flick so hard that the monolayer detaches. If cells are over-confluent, they will detach more easily.</li>
	</ol>
	</li>
	<li>Remove the inoculum by flicking and replace with 50 &#181;L/well growth media #2. Incubate plates at 37 &#176;C with 5% CO2. 
	NOTE: At this stage, integration of the selection cassette into the genome is stable, so antibiotic selection with spectinomycin is no longer required.</li>
	<li>After incubating the plate for 5&#8211;12 days, identify individual wells with green fluorescent inclusions using an epifluorescence inverted microscope or a high content screening platform.</li>
	<li>Harvest wells with green-only inclusions by scraping the monolayer with a p-10 tip and transferring the entire contents of the well into a tube containing 2 mL of HBSS. Gently mix and apply the 2 mL of inoculum to a fresh confluent McCoy monolayer in a 6 well plate for infection. 
	NOTE: When identifying individual wells with inclusions, the inclusions will be in a cluster due to expansion from the initial inclusion. If a well has multiple clusters, it is likely that more than one EB initially infected that well. These wells should not be harvested, and in some cases, multiple rounds of clonal isolation by serial dilution may be necessary.</li>
	<li>Infect the 6 well plate by centrifugation at 900 x g for 1 h at 20 &#176;C. Replace the inoculum with 2 mL/well growth media #2. Incubate at 37 &#176;C with 5 % CO2 for 48 h.</li>
	<li>Repeat steps 2.11&#8211;2.13 to enrich, freeze, and titer clonal populations.</li>
	<li>Confirm the deletion of targeted genes from clonally isolated C. trachomatis using qPCR as previously described12.</li>
</ol>
4. Transformation of C. trachomatis FRAEM Mutant with pSU-Cre to Initiate Removal of loxP Flanked Selection Cassette
<ol>
	<li>Repeat the transformation process in a 6 well plate as previously described (steps 2.1&#8211;2.8) using the clonally isolated C. trachomatis FRAEM mutant, pSU-Cre vector, and selection media #3. 
	NOTE: A successful transformation is indicated by inclusions that are red and green fluorescent.
	<ol>
		<li>Passage the monolayer until red-only inclusions are detected using an epifluorescence inverted microscope, which indicates loss of the selection cassette (two to three passages). Continue passaging the monolayer until red-only inclusions are enriched to a MOI of ~0.5.</li>
	</ol>
	</li>
	<li>Clonally isolate red-only inclusions by limiting dilution in a 384 well plate as previously described (steps 3.1&#8211;3.8); however, use selection media #3 for all steps.</li>
	<li>Enrich clonal populations by passaging until an MOI of 0.1. Once enriched, replace section media with growth media #2 to initiate loss of the pSU-Cre vector (approximately three to four passages).</li>
	<li>Clonally isolate nonfluorescent bacteria (steps 3.1&#8211;3.8); however, manually scan the plate with a brightfield microscope to detect inclusions. Enrich and freeze bacteria (step 2.11&#8211;2.12).</li>
	<li>Screen for the mutant strain using quantitative real-time PCR, Western blotting, and whole genome DNA Sequencing as previously described12.</li>
</ol>

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The authors have no conflicts of interest to disclose.

The protocol described here for the generation of markerless gene deletions in C. trachomatis by FLAEM allows targeted deletion of nonessential genes and eliminates cassette-induced polar effects. The protocol relies upon careful design of 5&#39; and 3&#39; homology arms inserted into the pSUmC 4.0 suicide vector, efficient transformation of C. trachomatis, and careful screening of isolated mutant strains. Successful genome engineering via this method results in bacteria that are nonfluorescent and cont...

Chlamydia trachomatis is the leading cause of bacterial sexually transmitted disease and represents a significant burden to human health. Over 100 million people are infected every year with C. trachomatis1. Approximately 70% of the infections in women are asymptomatic despite detrimental reproductive health effects, such as pelvic inflammatory disease, ectopic pregnancy, and/or infertility. Disease sequela are directly related to immunopathology initiated by C. trachomatis infection2. An efficacious vaccine has yet to be developed; therefore, understanding the function of bacterial virulence factors and other bacterial gene products is an important and urgent research question.
As intracellular bacteria, host cell invasion, intracellular replication, release of progeny, and evasion of host immunological responses are critical processes. C. trachomatis forms a parasitophorous membrane bound vacuole, termed an inclusion, for intracellular development. Establishment of the inclusion and many other critical processes are achieved by secretion of effector proteins via a type III secretion system (T3SS)3. Elucidating the functions of these secreted effectors was limited for many years due to the genetic intractability of C. trachomatis. Unlike E. coli, many classical cloning techniques are not applicable to Chlamydia. A few major limitations involve transformation efficiency, lack of counterselection reporters such as sacB, and plasmid maintenance. Whereas E. coli plasmids can generally be maintained indefinitely with an origin of replication and appropriate selective pressure, C. trachomatis plasmids requires an additional eight open reading frames (pgp1-8) for maintenance that are found on the native pL2 plasmid within the L2 serovar4.
In recent years there have been multiple genetic tools generated that accommodate Chlamydia&#39;s unique biology, yet there are still limitations5,6,7. Chemical mutagenesis by ethyl methanesulfonate (EMS) treatment can introduce missense mutations, or (less frequently) can result in nucleotide transitions introducing a premature stop codon to yield a nonsense mutation8. Transposon insertion is efficient for gene disruption, but current technology in Chlamydia research is laborious and time-consuming9. Both EMS treatment and transposon mutagenesis techniques generate random mutations and require rigorous screening methods to isolate mutant strains. A method to disrupt genes by insertion of group II introns (e.g., TargeTron) allows for directed mutagenesis; however, this method is limited by efficiency, and the insertion site is not always properly predicted10.
Fluorescence-reported allelic exchange mutagenesis (FRAEM) is a strategy used for targeted gene deletion coupled with insertion of a selection cassette providing antibiotic resistance and a fluorescence reporter11. Yet, FRAEM is complicated by the potential of cassette-induced polar effects on downstream genes, especially when targeting genes within polycistronic operons. Floxed cassette allelic exchange mutagenesis (FLAEM) is a novel genetic approach developed to alleviate the cassette-induced polar effects previously observed with the FRAEM selection cassette12. FLAEM utilizes Cre-loxP genome editing to remove the selection cassette and restore expression of downstream genes. The selection cassette containing antibiotic resistance and green fluorescent protein (GFP) is reengineered with flanking loxP sites. These loxP sites can recombine in the presence of Cre recombinase and result in excision of the cassette from the genome13. This strategy has been shown to alleviate cassette induced polar effects when targeting tmeA for deletion12,14.
Both FRAEM and FLAEM methods utilize the same suicide vector, pSUmC 4.0, which can be conditionally maintained through inducible expression of pgp6. Expression of pgp6 has previously been shown to be necessary for plasmid retention and is therefore leveraged to control plasmid maintenance11,15. When C. trachomatis is grown in media supplemented with anhydrous tetracycline (aTc) to induce pgp6 expression, the vector is maintained. In the absence of aTc, the vector is lost. Targeted gene deletion is achieved through allelic exchange of the gene for the selection cassette. The 3 kb regions directly upstream and downstream of the targeted gene serve as homology arms for recombination. These arms are cloned into the pSUmC 4.0 vector flanking the selection cassette. Successful C. trachomatis transformation and recombination events are observed through fluorescence reporting. Expression of mCherry on the vector backbone and gfp within the selection cassette yield red and green fluorescent inclusions. Once aTc is removed from culture media, green-only inclusions indicate successful recombination events with the loss of the suicide vector and integration of the selection cassette into the bacterial genome.
FLAEM represents an extension of FRAEM via subsequent transformation of a Cre recombinase-expressing vector, pSU-Cre, into the newly created mutant strain. Cre recombinase facilitates recombination between loxP sites and excision of the selection cassette. Recombination events are indicated via fluorescence reporting. The pSU-Cre vector encodes mCherry; thus, successful transformation is indicated by addition of red fluorescence to gfp-expressing inclusions. Cultivation in the absence of selective pressure for the cassette results in Cre-mediated recombination at the loxP sites, and loss of the cassette is indicated by red-only inclusions. As with pSUmC-4.0, inducible expression of pgp6 is used to conditionally maintain pSU-Cre. Once aTc and antibiotic selection are removed, the plasmid is cured, and the resulting markerless deletion strain is nonfluorescent. This method addresses the issue of cassette-induced polar effects.

<table><tbody><tr><td>Agarose</td><td>KSE Scientific</td><td>BMK-A1705</td><td>Molecular Biology Grade</td></tr><tr><td>Anhydrotetracycline hydrochloride</td><td>ACROS Organics</td><td>233131000</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt; Buffer</td><td></td><td></td><td>10 mM Tris pH 7.4, 50 mM Calcium Chloride Dihydrate</td></tr><tr><td>Calcium Chloride Dihydrate</td><td>Sigma</td><td>C7902-500G</td><td>Suitable for cell culture</td></tr><tr><td>Cycloheximide</td><td>Sigma</td><td>7698-1G</td><td></td></tr><tr><td>dam&lt;sup&gt;-&lt;/sup&gt;/dcm&lt;sup&gt;-&lt;/sup&gt; Competent &lt;em&gt;E. coli&lt;/em&gt;</td><td>New England BioLabs</td><td>C2925H</td><td></td></tr><tr><td>DMSO</td><td>ATCC</td><td>4-X</td><td>Sterile filtered cell culture tested</td></tr><tr><td>Glutamic acid</td><td>Sigma</td><td>G8415-100G</td><td>L-Glutamic acid</td></tr><tr><td>Growth Media #1</td><td></td><td></td><td>RPMI 1640 media supplemented with 10 % (vol/vol) heat-inactivated fetal bovine serum (FBS).</td></tr><tr><td>Growth Media #2</td><td></td><td></td><td>RPMI 1640 media supplemented with 10 % (vol/vol) heat-inactivated fetal bovine serum (FBS) and 1 &amp;micro;g/mL cycloheximide</td></tr><tr><td>Hanks&#039; Balanced Salt Solution (HBSS) (1x)</td><td>Gibco</td><td>24020-117</td><td></td></tr><tr><td>Heat Inactivated Fetal Bovine Serum Qualified One Shot (FBS)</td><td>Gibco</td><td>A38402-02</td><td></td></tr><tr><td>McCoy Cells</td><td>ATCC</td><td>CRL-1696</td><td></td></tr><tr><td>Monarch Plasmid Miniprep Kit</td><td>New England BioLabs</td><td>T1010S</td><td>Small scale DNA purification</td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>S3139-250G</td><td>Sodium phosphate monobasic</td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>S5136-500G</td><td>Sodium phosphate dibasic</td></tr><tr><td>NEB 10-beta Electrocompetent &lt;em&gt;E. coli&lt;/em&gt; Cells</td><td>New England BioLabs</td><td>C3020K</td><td></td></tr><tr><td>NEBuilder HiFi DNA assembly Cloning Kit</td><td>New England BioLabs</td><td>E5520S</td><td>Gibson Assembly Kit</td></tr><tr><td>Penicillin G sodium salt</td><td>Sigma</td><td>P3032-10MU</td><td>Bioreagent suitable for cell culture</td></tr><tr><td>QIAGEN Plasmid Maxi Kit</td><td>QIAGEN</td><td>12162</td><td>Large scale DNA purification</td></tr><tr><td>Q5 Hot Start High-Fidelity DNA Polymerase</td><td>New England BioLabs</td><td>M0515</td><td>Fragment PCR Polymerase</td></tr><tr><td>RPMI 1640 Medium (1x)</td><td>Gibco</td><td>11875-093</td><td>Containing 2mM L-glutamine</td></tr><tr><td>Sall-HF</td><td>New England BioLabs</td><td>R3138S</td><td></td></tr><tr><td>Sbfl-HF</td><td>New England BioLabs</td><td>R3642S</td><td></td></tr><tr><td>Selection Media #1</td><td></td><td></td><td>RMPI 10 % FBS, 1 &amp;micro;g/mL cycloheximide, 500 &amp;micro;g/mL spectinomycin, and 50 ng/mL anhydrous tetracycline dissolved in DMSO</td></tr><tr><td>Selection Media #2</td><td></td><td></td><td>RMPI 10 % FBS, 1 &amp;micro;g/mL cycloheximide, 500 &amp;micro;g/mL spectinomycin</td></tr><tr><td>Selection Media #3</td><td></td><td></td><td>RPMI 10 % FBS, 1 &amp;micro;g/mL cycloheximide, 50 ng/mL aTc, and 0.6 &amp;micro;g/mL penicillin</td></tr><tr><td>Sodium Acetate Buffer Solution</td><td>Sigma</td><td>S7899-100ML</td><td>3M</td></tr><tr><td>SOC Outgrowth Medium</td><td>New England BioLabs</td><td>B9020SVIAL</td><td></td></tr><tr><td>Spectinomycin dihydrochloride pentahydrate, Cell Culture Grade</td><td>Alfa Aesar</td><td>J61820</td><td></td></tr><tr><td>Sucrose</td><td>Sigma</td><td>S1888-1KG</td><td>Bioreagent suitable for cell culture</td></tr><tr><td>Sucrose-Phosphate-Glutamate Buffer (SPG)</td><td></td><td></td><td>37.5g sucrose, 1.25 g Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;, 0.18 g NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, 0.36 glutamic acid for 500 ml tissue culture grade water</td></tr><tr><td>Tris</td><td>AMRESCO</td><td>0497-5KG</td><td>Ultrapure grade</td></tr><tr><td>Trypsin-EDTA (1x)</td><td>Gibco</td><td>25200-056</td><td>0.25%</td></tr><tr><td>Water</td><td>Sigma</td><td>W3500-500ML</td><td>Sterile-filtered, BioReagent, Suitable for cell culture</td></tr></tbody></table>

markerless gene deletion by floxed cassette allelic exchange mutagenesis in chlamydia trachomatis

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.

The method for markerless gene deletion in C. trachomatis using FLAEM is reliant upon careful cloning and transformation techniques. Successful allelic recombination is an essential first step and requires the identification and insertion of homology arms into the pSUmC-4.0 cloning vector (Figure 1). An essential second step for markerless gene deletion is removal of the fluorescence reporter and antibiotic selection cassette by Cre-lox genome editing, represented in <strong class="...

Watch this Scientific Journal Video about Markerless Gene Deletion by Floxed Cassette Allelic Exchange Mutagenesis in Chlamydia trachomatis at JoVE.com

Markerless Gene Deletion by Floxed Cassette Allelic Exchange Mutagenesis in Chlamydia trachomatis

Described here is a method for targeted, markerless gene deletion in Chlamydia trachomatis using floxed cassette allelic exchange mutagenesis, FLAEM.

Markerless Gene Deletion by Floxed Cassette Allelic Exchange Mutagenesis in Chlamydia trachomatis

Chlamydia trachomatis is an obligate intracellular pathogen that has been historically difficult to genetically manipulate. Definitive progress in ...

markerless-gene-deletion-floxed-cassette-allelic-exchange-mutagenesis

Research

JoVE Journal

Immunology and Infection

6.6K Views.  University of Kentucky. Described here is a method for targeted, markerless gene deletion in Chlamydia trachomatis using floxed cassette allelic exchange mutagenesis, FLAEM.

Video: Markerless Gene Deletion by Floxed Cassette Allelic Exchange Mutagenesis in Chlamydia trachomatis

Genetic Manipulation of the Plant Pathogen Ustilago maydis to Study Fungal Biology and Plant Microbe Interactions

Gene deletion plays an important role in the analysis of gene function. One of the most efficient methods to disrupt genes in a targeted manner is the replacement of the entire gene with a selectable marker via homologous recombination. During homologous recombination, exchange of DNA takes place between sequences with high similarity. Therefore, linear genomic sequences flanking a target gene can be used to specifically direct a selectable marker to the desired integration site. Blunt ends of the deletion construct activate the cell&#39;s DNA repair systems and thereby promote integration of the construct either via homologous recombination or by non-homologous-end-joining. In organisms with efficient homologous recombination, the rate of successful gene deletion can reach more than 50% making this strategy a valuable gene disruption system. The smut fungus Ustilago maydis is a eukaryotic model microorganism showing such efficient homologous recombination. Out of its about 6,900 genes, many have been functionally characterized with the help of deletion mutants, and repeated failure of gene replacement attempts points at essential function of the gene. Subsequent characterization of the gene function by tagging with fluorescent markers or mutations of predicted domains also relies on DNA exchange via homologous recombination. Here, we present the U. maydis strain generation strategy in detail using the simplest example, the gene deletion.

Genetic Manipulation of the Plant Pathogen Ustilago maydis to Study Fungal Biology and Plant Microbe Interactions

We describe a robust gene replacement strategy to genetically manipulate the smut fungus Ustilago maydis. This protocol explains how to generate deletion mutants to investigate infection phenotypes. It can be extended to modify genes in any desired way, e.g., by adding a sequence encoding a fluorescent protein tag.

Gene deletion plays an important role in the analysis of gene function. One of the most efficient methods to disrupt genes in a targeted manner is the ...

Genetics

High-Throughput Robotically Assisted Isolation of Temperature-sensitive Lethal Mutants in Chlamydomonas reinhardtii

Systematic identification and characterization of genetic perturbations have proven useful to decipher gene function and cellular pathways. However, the conventional approaches of permanent gene deletion cannot be applied to essential genes. We have pioneered a unique collection of ~70 temperature-sensitive (ts) lethal mutants for studying cell cycle regulation in the unicellular green algae Chlamydomonas reinhardtii1. These mutations identify essential genes, and the ts alleles can be conditionally inactivated by temperature shift, providing valuable tools to identify and analyze essential functions. Mutant collections are much more valuable if they are close to comprehensive, since scattershot collections can miss important components. However, this requires the efficient collection of a large number of mutants, especially in a wide-target screen. Here, we describe a robotics-based pipeline for generating ts lethal mutants and analyzing their phenotype in Chlamydomonas. This technique can be applied to any microorganism that grows on agar. We have collected over 3000 ts mutants, probably including mutations in most or all cell-essential pathways, including about 200 new candidate cell cycle mutations. Subsequent molecular and cellular characterization of these mutants should provide new insights in plant cell biology; a comprehensive mutant collection is an essential prerequisite to ensure coverage of a broad range of biological pathways. These methods are integrated with downstream genetics and bioinformatics procedures for efficient mapping and identification of the causative mutations that are beyond the scope of this manuscript.

High-Throughput Robotically Assisted Isolation of Temperature-sensitive Lethal Mutants in Chlamydomonas reinhardtii

Temperature-sensitive (ts) lethal mutants are valuable tools to identify and analyze essential functions. Here we describe methods to generate and classify ts lethal mutants in high throughput.

Systematic identification and characterization of genetic perturbations have proven useful to decipher gene function and cellular pathways. However, the ...

Generation of a Gene-disrupted Streptococcus mutans Strain Without Gene Cloning

Typical methods for the elucidation of the function of a particular gene involve comparative phenotypic analyses of the wild-type strain and a strain in which the gene of interest has been disrupted. A gene-disruption DNA construct containing a suitable antibiotic resistance marker gene is useful for the generation of gene-disrupted strains in bacteria. However, conventional construction methods, which require gene cloning steps, involve complex and time-consuming protocols. Here, a relatively facile, rapid, and cost-effective method for targeted gene disruption in Streptococcus mutans is described. The method utilizes a 2-step fusion polymerase chain reaction (PCR) to generate the disruption construct and electroporation for genetic transformation. This method does not require an enzymatic reaction, other than PCR, and additionally offers greater flexibility in terms of the design of the disruption construct. Employment of electroporation facilitates the preparation of competent cells and improves the transformation efficiency. The present method may be adapted for the generation of gene-disrupted strains of various species.

Generation of a Gene-disrupted Streptococcus mutans Strain Without Gene Cloning

We describe a facile, rapid, and relatively inexpensive method for the generation of gene-disrupted Streptococcus mutans strains; this technique may be adapted for the generation of gene-disrupted strains of various species.

Typical methods for the elucidation of the function of a particular gene involve comparative phenotypic analyses of the wild-type strain and a strain in ...

Producing Gene Deletions in Escherichia coli by P1 Transduction with Excisable Antibiotic Resistance Cassettes

A first approach to study the function of an unknown gene in bacteria is to create a knock-out of this gene. Here, we describe a robust and fast protocol for transferring gene deletion mutations from one Escherichia coli strain to another by using generalized transduction with the bacteriophage P1. This method requires that the mutation be selectable (e.g., based on gene disruptions using antibiotic cassette insertions). Such antibiotic cassettes can be mobilized from a donor strain and introduced into a recipient strain of interest to quickly and easily generate a gene deletion mutant. The antibiotic cassette can be designed to include flippase recognition sites that allow the excision of the cassette by a site-specific recombinase to produce a clean knock-out with only a ~100-base-pair-long scar sequence in the genome. We demonstrate the protocol by knocking out the tamA gene encoding an assembly factor involved in autotransporter biogenesis and test the effect of this knock-out on the biogenesis and function of two trimeric autotransporter adhesins. Though gene deletion by P1 transduction has its limitations, the ease and speed of its implementation make it an attractive alternative to other methods of gene deletion.

Producing Gene Deletions in Escherichia coli by P1 Transduction with Excisable Antibiotic Resistance Cassettes

Here we present a protocol for the use of pre-existing antibiotic resistance-cassette deletion constructs as a basis for making deletion mutants in other E. coli strains. Such deletion mutations can be mobilized and inserted into the corresponding locus of a recipient strain using P1 bacteriophage transduction.

A first approach to study the function of an unknown gene in bacteria is to create a knock-out of this gene. Here, we describe a robust and fast protocol ...

TransFLP &#x2014; A Method to Genetically Modify Vibrio cholerae Based on Natural Transformation and FLP-recombination

Several methods are available to manipulate bacterial chromosomes1-3. Most of these protocols rely on the insertion of conditionally replicative plasmids (e.g. harboring pir-dependent or temperature-sensitive replicons1,2). These plasmids are integrated into bacterial chromosomes based on homology-mediated recombination. Such insertional mutants are often directly used in experimental settings. Alternatively, selection for plasmid excision followed by its loss can be performed, which for Gram-negative bacteria often relies on the counter-selectable levan sucrase enzyme encoded by the sacB gene4. The excision can either restore the pre-insertion genotype or result in an exchange between the chromosome and the plasmid-encoded copy of the modified gene. A disadvantage of this technique is that it is time-consuming. The plasmid has to be cloned first; it requires horizontal transfer into V. cholerae (most notably by mating with an E. coli donor strain) or artificial transformation of the latter; and the excision of the plasmid is random and can either restore the initial genotype or create the desired modification if no positive selection is exerted. Here, we present a method for rapid manipulation of the V. cholerae chromosome(s)5 (Figure 1). This TransFLP method is based on the recently discovered chitin-mediated induction of natural competence in this organism6 and other representative of the genus Vibrio such as V. fischeri7. Natural competence allows the uptake of free DNA including PCR-generated DNA fragments. Once taken up, the DNA recombines with the chromosome given the presence of a minimum of 250-500 bp of flanking homologous region8. Including a selection marker in-between these flanking regions allows easy detection of frequently occurring transformants.
This method can be used for different genetic manipulations of V. cholerae and potentially also other naturally competent bacteria. We provide three novel examples on what can be accomplished by this method in addition to our previously published study on single gene deletions and the addition of affinity-tag sequences5. Several optimization steps concerning the initial protocol of chitin-induced natural transformation6 are incorporated in this TransFLP protocol. These include among others the replacement of crab shell fragments by commercially available chitin flakes8, the donation of PCR-derived DNA as transforming material9, and the addition of FLP-recombination target sites (FRT)5. FRT sites allow site-directed excision of the selection marker mediated by the Flp recombinase10.

A quick method to modify the genome of V. cholerae is described. These modifications include the deletion of single genes, gene clusters and genomic islands as well as the integration of short sequences (e.g. promoter elements or affinity-tag sequences). The method is based on the natural transformation and FLP-recombination.

Several methods are available to manipulate bacterial chromosomes1-3. Most of these protocols rely on the insertion of conditionally replicative plasmids ...

Identification of Host Pathways Targeted by Bacterial Effector Proteins using Yeast Toxicity and Suppressor Screens

Intracellular bacteria secrete virulence factors called effector proteins into the host cytosol that act to subvert host proteins and/or their associated biological pathways to the benefit of the bacterium. Identification of putative bacterial effector proteins has become more manageable due to advances in bacterial genome sequencing and the advent of algorithms that allow in silico identification of genes encoding secretion candidates and/or eukaryotic-like domains. However, identification of these important virulence factors is only an initial step. Naturally, the goal is to determine the molecular function of effector proteins and elucidate how they interact with the host. In recent years, techniques like the yeast two-hybrid screen and large-scale immunoprecipitations coupled with mass spectrometry have aided in the identification of protein-protein interactions. Although identification of a host binding partner is the crucial first step toward elucidating the molecular function of a bacterial effector protein, sometimes the host protein is found to have multiple biological functions (e.g., actin, clathrin, tubulin), or the bacterial protein may not physically bind host proteins, depriving the researcher of crucial information about the precise host pathway being manipulated. A modified yeast toxicity screen coupled with a suppressor screen has been adapted to identify host pathways impacted by bacterial effector proteins. The toxicity screen relies on a toxic effect in yeast caused by the effector protein interfering with the host biological pathways, which often manifests as a growth defect. Expression of a yeast genomic library is used to identify host factors that suppress the toxicity of the bacterial effector protein and thus identify proteins in the pathway that the effector protein targets. This protocol contains detailed instructions for both the toxicity and suppressor screens. These techniques can be performed in any lab capable of molecular cloning and cultivation of yeast and Escherichia coli.

Bacterial pathogens secrete proteins into the host that target crucial biological processes. Identifying the host pathways targeted by bacterial effector proteins is key to addressing molecular pathogenesis. Here, a method using a modified yeast suppressor and toxicity screen to elucidate host pathways targeted by toxic bacterial effector proteins is described.

Intracellular bacteria secrete virulence factors called effector proteins into the host cytosol that act to subvert host proteins and/or their associated ...

Forward Genetic Approaches in Chlamydia trachomatis

Chlamydia trachomatis, the etiological agent of sexually transmitted diseases and ocular infections, remains poorly characterized due to its intractability to experimental transformation with recombinant DNA. We developed an approach to perform genetic analysis in C. trachomatis&#160;despite the lack of molecular genetic tools. Our method involves: i.) chemical mutagenesis to rapidly generate comprehensive libraries of genetically-defined mutants with distinct phenotypes; ii.) whole-genome sequencing (WGS) to map the underlying genetic lesions and to find associations between mutated gene(s) and a common phenotype; iii.) generation of recombinant strains through co-infection of mammalian cells with mutant and wild type bacteria. Accordingly, we were able to establish causal relationships between genotypes and phenotypes. The coupling of chemically-induced gene variation and WGS to establish correlative genotype&#8211;phenotype associations should be broadly applicable to the large list of medically and environmentally important microorganisms currently intractable to genetic analysis.

Forward Genetic Approaches in Chlamydia trachomatis

We describe a methodology to perform genetic analysis in Chlamydia based on chemical mutagenesis and whole genome sequencing. In addition, a system for DNA exchange within infected cells is described that can be used for genetic mapping. This method may be broadly applicable to microbial systems lacking transformation systems and molecular genetic tools.

Chlamydia trachomatis, the etiological agent of sexually transmitted diseases and ocular infections, remains poorly characterized due to its ...

Mapping Bacterial Functional Networks and Pathways in Escherichia Coli using Synthetic Genetic Arrays

Phenotypes are determined by a complex series of physical (e.g. protein-protein) and functional (e.g. gene-gene or genetic) interactions (GI)1. While physical interactions can indicate which bacterial proteins are associated as complexes, they do not necessarily reveal pathway-level functional relationships1. GI screens, in which the growth of double mutants bearing two deleted or inactivated genes is measured and compared to the corresponding single mutants, can illuminate epistatic dependencies between loci and hence provide a means to query and discover novel functional relationships2. Large-scale GI maps have been reported for eukaryotic organisms like yeast3-7, but GI information remains sparse for prokaryotes8, which hinders the functional annotation of bacterial genomes. To this end, we and others have developed high-throughput quantitative bacterial GI screening methods9, 10.
Here, we present the key steps required to perform quantitative E. coli Synthetic Genetic Array (eSGA) screening procedure on a genome-scale9, using natural bacterial conjugation and homologous recombination to systemically generate and measure the fitness of large numbers of double mutants in a colony array format. Briefly, a robot is used to transfer, through conjugation, chloramphenicol (Cm) - marked mutant alleles from engineered Hfr (High frequency of recombination) 'donor strains' into an ordered array of kanamycin (Kan) - marked F- recipient strains. Typically, we use loss-of-function single mutants bearing non-essential gene deletions (e.g. the 'Keio' collection11) and essential gene hypomorphic mutations (i.e. alleles conferring reduced protein expression, stability, or activity9, 12, 13) to query the functional associations of non-essential and essential genes, respectively. After conjugation and ensuing genetic exchange mediated by homologous recombination, the resulting double mutants are selected on solid medium containing both antibiotics. After outgrowth, the plates are digitally imaged and colony sizes are quantitatively scored using an in-house automated image processing system14. GIs are revealed when the growth rate of a double mutant is either significantly better or worse than expected9. Aggravating (or negative) GIs often result between loss-of-function mutations in pairs of genes from compensatory pathways that impinge on the same essential process2. Here, the loss of a single gene is buffered, such that either single mutant is viable. However, the loss of both pathways is deleterious and results in synthetic lethality or sickness (i.e. slow growth). Conversely, alleviating (or positive) interactions can occur between genes in the same pathway or protein complex2 as the deletion of either gene alone is often sufficient to perturb the normal function of the pathway or complex such that additional perturbations do not reduce activity, and hence growth, further. Overall, systematically identifying and analyzing GI networks can provide unbiased, global maps of the functional relationships between large numbers of genes, from which pathway-level information missed by other approaches can be inferred9.

Mapping Bacterial Functional Networks and Pathways in Escherichia Coli using Synthetic Genetic Arrays

Systematic, large-scale synthetic genetic (gene-gene or epistasis) interaction screens can be used to explore genetic redundancy and pathway cross-talk. Here, we describe a high-throughput quantitative synthetic genetic array screening technology, termed eSGA that we developed for elucidating epistatic relationships and exploring genetic interaction networks in Escherichia coli.

Phenotypes  are determined by a complex series of physical (e.g. protein-protein) and  functional (e.g. gene-gene or genetic) interactions (GI)1. While ...

Biology

Live-Cell Forward Genetic Approach to Identify and Isolate Developmental Mutants in Chlamydia trachomatis

The intracellular bacterial pathogen Chlamydia trachomatis undergoes a developmental cycle consisting of two morphologically discrete developmental forms. The non-replicative elementary body (EB) initiates infection of the host. Once inside, the EB differentiates into the reticulate body (RB). The RB then undergoes multiple rounds of replication, before differentiating back to the infectious EB form. This cycle is essential for chlamydial survival as failure to switch between cell types prevents either host invasion or replication.
Limitations in genetic techniques due to the obligate intracellular nature of Chlamydia have hampered identification of the molecular mechanisms involved in the cell-type development. We designed a novel dual promoter-reporter plasmid system that, in conjunction with live-cell microscopy, allows for the visualization of cell type switching in real time. To identify genes involved in the regulation of cell-type development, the live-cell promoter-reporter system was leveraged for the development of a forward genetic approach by combining chemical mutagenesis of the dual reporter strain, imaging and tracking of Chlamydia with altered developmental kinetics, followed by clonal isolation of mutants. This forward genetic workflow is a flexible tool that can be modified for directed interrogation into a wide range of genetic pathways.

Live-Cell Forward Genetic Approach to Identify and Isolate Developmental Mutants in Chlamydia trachomatis

This protocol utilizes fluorescent promoter-reporters, live-cell microscopy, and individual inclusion extraction in a directed forward genetic approach to identify and isolate developmental mutants of Chlamydia trachomatis.

The intracellular bacterial pathogen Chlamydia trachomatis undergoes a developmental cycle consisting of two morphologically discrete developmental forms. ...

Generation of Null Mutants to Elucidate the Role of Bacterial Glycosyltransferases in Bacterial Motility

The study of glycosylation in prokaryotes is a rapidly growing area. Bacteria harbor different glycosylated structures on their surface whose glycans constitute a strain-specific barcode. The associated glycans show higher diversity in sugar composition and structure than those of eukaryotes and are important in bacterial-host recognition processes and interaction with the environment. In pathogenic bacteria, glycoproteins have been involved in different stages of the infectious process, and glycan modifications can interfere with specific functions of glycoproteins. However, despite the advances made in the understanding of glycan composition, structure, and biosynthesis pathways, understanding of the role of glycoproteins in pathogenicity or interaction with the environment remains very limited. Furthermore, in some bacteria, the enzymes required for protein glycosylation are shared with other polysaccharide biosynthetic pathways, such as lipopolysaccharide and capsule biosynthetic pathways. The functional importance of glycosylation has been elucidated in several bacteria through mutation of specific genes thought to be involved in the glycosylation process and the study of its impact on the expression of the target glycoprotein and the modifying glycan. Mesophilic Aeromonas have a single and O-glycosylated polar flagellum. Flagellar glycans show diversity in carbohydrate composition and chain length between Aeromonas strains. However, all strains analyzed to date show a pseudaminic acid derivative as the linking sugar that modifies serine or threonine residues. The pseudaminic acid derivative is required for polar flagella assembly, and its loss has an impact on adhesion, biofilm formation, and colonization. The protocol detailed in this article describes how the construction of null mutants can be used to understand the involvement of genes or genome regions containing putative glycosyltransferases in the biosynthesis of a flagellar glycan. This includes the potential to understand the function of the glycosyltransferases involved and the role of the glycan. This will be achieved by comparing the glycan deficient mutant to the wild-type strain.

Here, we describe the construction of null mutants of Aeromonas in specific glycosyltransferases or regions containing glycosyltransferases, the motility assays, and flagella purification performed to establish the involvement and function of their encoded enzymes in the biosynthesis of a glycan, as well as the role of this glycan in bacterial pathogenesis.

The study of glycosylation in prokaryotes is a rapidly growing area. Bacteria harbor different glycosylated structures on their surface whose glycans ...

Markerless Gene Deletion by Floxed Cassette Allelic Exchange Mutagenesis in Chlamydia trachomatis

Summary

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Chapters in this video

Genetic Manipulation of the Plant Pathogen Ustilago maydis to Study Fungal Biology and Plant Microbe Interactions

High-Throughput Robotically Assisted Isolation of Temperature-sensitive Lethal Mutants in Chlamydomonas reinhardtii

Generation of a Gene-disrupted Streptococcus mutans Strain Without Gene Cloning

Producing Gene Deletions in Escherichia coli by P1 Transduction with Excisable Antibiotic Resistance Cassettes

TransFLP — A Method to Genetically Modify Vibrio cholerae Based on Natural Transformation and FLP-recombination

Identification of Host Pathways Targeted by Bacterial Effector Proteins using Yeast Toxicity and Suppressor Screens

Forward Genetic Approaches in Chlamydia trachomatis

Mapping Bacterial Functional Networks and Pathways in Escherichia Coli using Synthetic Genetic Arrays

Live-Cell Forward Genetic Approach to Identify and Isolate Developmental Mutants in Chlamydia trachomatis

Generation of Null Mutants to Elucidate the Role of Bacterial Glycosyltransferases in Bacterial Motility