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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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>CaCl2 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-/dcm- Competent E. coli</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 &#181;g/mL cycloheximide</td>
		</tr>
		<tr>
			<td>Hanks&#39; 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>NaH2PO4</td>
			<td>Sigma</td>
			<td>S3139-250G</td>
			<td>Sodium phosphate monobasic</td>
		</tr>
		<tr>
			<td>Na2HPO4</td>
			<td>Sigma</td>
			<td>S5136-500G</td>
			<td>Sodium phosphate dibasic</td>
		</tr>
		<tr>
			<td>NEB 10-beta Electrocompetent E. coli 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 &#181;g/mL cycloheximide, 500 &#181;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 &#181;g/mL cycloheximide, 500 &#181;g/mL spectinomycin</td>
		</tr>
		<tr>
			<td>Selection Media #3</td>
			<td></td>
			<td></td>
			<td>RPMI 10 % FBS, 1 &#181;g/mL cycloheximide, 50 ng/mL aTc, and 0.6 &#181;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 Na2HPO4, 0.18 g NaH2PO4, 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

6.5K 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

Transfection and Mutagenesis of Target Genes in Mosquito Cells by Locked Nucleic Acid-modified Oligonucleotides

Plasmodium parasites, the causative agent of malaria, are transmitted through the bites of infected Anopheles mosquitoes resulting in over 250 million new infections each year. Despite decades of research, there is still no vaccine against malaria, highlighting the need for novel control strategies. One innovative approach is the use of genetically modified mosquitoes to effectively control malaria parasite transmission. Deliberate alterations of cell signaling pathways in the mosquito, via targeted mutagenesis, have been found to regulate parasite development 1. From these studies, we can begin to identify potential gene targets for transformation. Targeted mutagenesis has traditionally relied upon the homologous recombination between a target gene and a large DNA molecule. However, the construction and use of such complex DNA molecules for generation of stably transformed cell lines is costly, time consuming and often inefficient. Therefore, a strategy using locked nucleic acid-modified oligonucleotides (LNA-ONs) provides a useful alternative for introducing artificial single nucleotide substitutions into episomal and chromosomal DNA gene targets (reviewed in 2). LNA-ON-mediated targeted mutagenesis has been used to introduce point mutations into genes of interest in cultured cells of both yeast and mice 3,4. We show here that LNA-ONs can be used to introduce a single nucleotide change in a transfected episomal target that results in a switch from blue fluorescent protein (BFP) expression to green fluorescent protein (GFP) expression in both Anopheles gambiae and Anopheles stephensi cells. This conversion demonstrates for the first time that effective mutagenesis of target genes in mosquito cells can be mediated by LNA-ONs and suggests that this technique may be applicable to mutagenesis of chromosomal targets in vitro and in vivo.

Oligonucleotides can be used to site specifically substitute a single nucleotide of transfected target genes in both Anopheles gambiae and Anopheles stephensi cells.

Plasmodium parasites, the causative agent of malaria, are transmitted through the bites of infected Anopheles mosquitoes resulting in over 250 million new ...

Differentiating Functional Roles of Gene Expression from Immune and Non-immune Cells in Mouse Colitis by Bone Marrow Transplantation

To understand the role of a gene in the development of colitis, we compared the responses of wild-type mice and gene-of-interest deficient knockout mice to colitis. If the gene-of-interest is expressed in both bone marrow derived cells and non-bone marrow derived cells of the host; however, it is possible to differentiate the role of a gene of interest in bone marrow derived cells and non- bone marrow derived cells by bone marrow transplantation technique. To change the bone marrow derived cell genotype of mice, the original bone marrow of recipient mice were destroyed by irradiation and then replaced by new donor bone marrow of different genotype. When wild-type mice donor bone marrow was transplanted to knockout mice, we could generate knockout mice with wild-type gene expression in bone marrow derived cells. Alternatively, when knockout mice donor bone marrow was transplanted to wild-type recipient mice, wild-type mice without gene-of-interest expressing from bone marrow derived cells were produced. However, bone marrow transplantation may not be 100% complete. Therefore, we utilized cluster of differentiation (CD) molecules (CD45.1 and CD45.2) as markers of donor and recipient cells to track the proportion of donor bone marrow derived cells in recipient mice and success of bone marrow transplantation. Wild-type mice with CD45.1 genotype and knockout mice with CD45.2 genotype were used. After irradiation of recipient mice, the donor bone marrow cells of different genotypes were infused into the recipient mice. When the new bone marrow regenerated to take over its immunity, the mice were challenged by chemical agent (dextran sodium sulfate, DSS 5%) to induce colitis. Here we also showed the method to induce colitis in mice and evaluate the role of the gene of interest expressed from bone-marrow derived cells. If the gene-of-interest from the bone derived cells plays an important role in the development of the disease (such as colitis), the phenotype of the recipient mice with bone marrow transplantation can be significantly altered. At the end of colitis experiments, the bone marrow derived cells in blood and bone marrow were labeled with antibodies against CD45.1 and CD45.2 and their quantitative ratio of existence could be used to evaluate the success of bone marrow transplantation by flow cytometry. Successful bone marrow transplantation should show a vast majority of donor genotype (in term of CD molecule marker) over recipient genotype in both the bone marrow and blood of recipient mice.

Bone marrow transplantation provides a way to change the genotype of the bone marrow derived cells. If the gene of interest is expressed in both bone marrow derived cells and non-bone marrow derived cells, bone marrow transplantation can change the bone marrow derived cells to a different genotype without changing the non-bone marrow derived cell genotype.

To understand the role of a gene in the development of colitis, we compared the responses of wild-type mice and gene-of-interest deficient knockout mice ...

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 ...

Multiplex PCR Assay for Typing of Staphylococcal Cassette Chromosome Mec Types I to V in Methicillin-resistant Staphylococcus aureus

Staphylococcal Cassette Chromosome mec (SCCmec) typing is a very important molecular tool for understanding the epidemiology and clonal strain relatedness of methicillin-resistant Staphylococcus aureus (MRSA), particularly with the emerging outbreaks of community-associated MRSA (CA-MRSA) occurring on a worldwide basis. Traditional PCR typing schemes classify SCCmec by targeting and identifying the individual mec and ccr gene complex types, but require the use of many primer sets and multiple individual PCR experiments. We designed and published a simple multiplex PCR assay for quick-screening of major SCCmec types and subtypes I to V, and later updated it as new sequence information became available. This simple assay targets individual SCCmec types in a single reaction, is easy to interpret and has been extensively used worldwide. However, due to the sophisticated nature of the assay and the large number of primers present in the reaction, there is the potential for difficulties while adapting this assay to individual laboratories. To facilitate the process of establishing a MRSA SCCmec assay, here we demonstrate how to set up our multiplex PCR assay, and discuss some of the vital steps and procedural nuances that make it successful.

Multiplex PCR Assay for Typing of Staphylococcal Cassette Chromosome Mec Types I to V in Methicillin-resistant Staphylococcus aureus

We demonstrate a simple multiplex PCR assay for quick-screening and typing of Staphylococcal Cassette Chromosome mec (SCCmec) types I-V for methicillin-resistant Staphylococcus aureus, and provide some of the vital steps and procedural nuances that make it successful for adapting this assay to individual laboratories.

Staphylococcal Cassette Chromosome mec (SCCmec) typing  is a very important molecular tool for understanding the epidemiology and  clonal strain ...

Cell-based Flow Cytometry Assay to Measure Cytotoxic Activity

Cytolytic activity of CD8+ T cells is rarely evaluated. We describe here a new cell-based assay to measure the capacity of antigen-specific CD8+ T cells to kill CD4+ T cells loaded with their cognate peptide. Target CD4+ T cells are divided into two populations, labeled with two different concentrations of CFSE. One population is pulsed with the peptide of interest (CFSE-low) while the other remains un-pulsed (CFSE-high). Pulsed and un-pulsed CD4+ T cells are mixed at an equal ratio and incubated with an increasing number of purified CD8+ T cells. The specific killing of autologous target CD4+ T cells is analyzed by flow cytometry after coculture with CD8+ T cells containing the antigen-specific effector CD8+ T cells detected by peptide/MHCI tetramer staining. The specific lysis of target CD4+ T cells measured at different effector versus target ratios, allows for the calculation of lytic units, LU30/106 cells. This simple and straightforward assay allows for the accurate measurement of the intrinsic capacity of CD8+ T cells to kill target CD4+ T cells.

This protocol describes a sensitive, cell-based cytotoxicity assay. By enumerating the decrease in frequency of live target CD4+ T cells in the presence of an increasing number of effector CD8+ T cells, this assay allows for the direct assessment of cytolytic activity of antigen-specific CD8+ T cells.

Cytolytic activity of CD8+ T cells is rarely evaluated. We describe here a new cell-based assay to measure the capacity of antigen-specific CD8+ T cells ...

Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

The obligate intracellular bacterium Chlamydia elicits a great burden on global public health. C.&#160;trachomatis is the leading bacterial cause of sexually transmitted infection and also the primary cause of preventable blindness in the world. An essential determinant for successful infection of host cells by Chlamydia is the bacterium's ability to manipulate host cell signaling from within a novel, vacuolar compartment called the inclusion. From within the inclusion, Chlamydia acquire nutrients required for their 2-3 day developmental growth, and they additionally secrete a panel of effector proteins onto the cytosolic face of the vacuole membrane and into the host cytosol. Gaps in our understanding of Chlamydia biology, however, present significant challenges for visualizing and analyzing this intracellular compartment. Recently, a reverse-imaging strategy for visualizing the inclusion using GFP expressing host cells was described. This approach rationally exploits the intrinsic impermeability of the inclusion membrane to large molecules such as GFP. In this work, we describe how GFP- or mCherry-expressing host cells are generated for subsequent visualization of chlamydial inclusions. Furthermore, this method is shown to effectively substitute for costly antibody-based enumeration methods, can be used in tandem with other fluorescent labels, such as GFP-expressing Chlamydia, and can be exploited to derive key quantitative data about inclusion membrane growth from a range of Chlamydia species and strains.

Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

A live cell fluorescent protein based method for illuminating cellular vacuoles (inclusions) containing Chlamydia is described. This strategy enables rapid, automated determination of Chlamydia infectivity in samples and can be used to quantitatively investigate inclusion growth dynamics.

The obligate intracellular bacterium Chlamydia elicits a great burden on global public health. C.&#160;trachomatis is the leading bacterial cause of ...

Identification of Novel Genes Associated with Alginate Production in Pseudomonas aeruginosa Using Mini-himar1 Mariner Transposon-mediated Mutagenesis

Pseudomonas aeruginosa is a Gram-negative, environmental bacterium with versatile metabolic capabilities. P. aeruginosa is an opportunistic bacterial pathogen which establishes chronic pulmonary infections in patients with cystic fibrosis (CF). The overproduction of a capsular polysaccharide called alginate, also known as mucoidy, promotes the formation of mucoid biofilms which are more resistant than planktonic cells to antibiotic chemotherapy and host defenses. Additionally, the conversion from the nonmucoid to mucoid phenotype is a clinical marker for the onset of chronic infection in CF. Alginate overproduction by P. aeruginosa is an endergonic process which heavily taxes cellular energy. Therefore, alginate production is highly regulated in P. aeruginosa. To better understand alginate regulation, we describe a protocol using the mini-himar1 transposon mutagenesis for the identification of novel alginate regulators in a prototypic strain PAO1. The procedure consists of two basic steps. First, we transferred the mini-himar1 transposon (pFAC) from host E. coli SM10/&#955;pir into recipient P. aeruginosa PAO1 via biparental conjugation to create a high-density insertion mutant library, which were selected on Pseudomonas isolation agar plates supplemented with gentamycin. Secondly, we screened and isolated the mucoid colonies to map the insertion site through inverse PCR using DNA primers pointing outward from the gentamycin cassette and DNA sequencing. Using this protocol, we have identified two novel alginate regulators, mucE (PA4033) and kinB (PA5484), in strain PAO1 with a wild-type mucA encoding the anti-sigma factor MucA for the master alginate regulator AlgU (AlgT, &#963;22). This high-throughput mutagenesis protocol can be modified for the identification of other virulence-related genes causing change in colony morphology.

Identification of Novel Genes Associated with Alginate Production in Pseudomonas aeruginosa Using Mini-himar1 Mariner Transposon-mediated Mutagenesis

Here we describe a protocol using the mini-himar1 mariner transposon-mediated mutagenesis for generating a high-density insertion mutant library to screen, isolate and identify novel alginate regulators in the prototypic Pseudomonas aeruginosa strain PAO1.

Pseudomonas aeruginosa is a Gram-negative, environmental bacterium with versatile metabolic capabilities. P. aeruginosa is an opportunistic bacterial ...

Identification of Rare Bacterial Pathogens by 16S rRNA Gene Sequencing and MALDI-TOF MS

There are a number of rare and, therefore, insufficiently described bacterial pathogens which are reported to cause severe infections especially in immunocompromised patients. In most cases only few data, mostly published as case reports, are available which investigate the role of such pathogens as an infectious agent. Therefore, in order to clarify the pathogenic character of such microorganisms, it is necessary to conduct epidemiologic studies which include large numbers of these bacteria. The methods used in such a surveillance study have to meet the following criteria: the identification of the strains has to be accurate according to the valid nomenclature, they should be easy to handle (robustness), economical in routine diagnostics and they have to generate comparable results among different laboratories. Generally, there are three strategies for identifying bacterial strains in a routine setting: 1) phenotypic identification characterizing the biochemical and metabolic properties of the bacteria, 2) molecular techniques such as 16S rRNA gene sequencing and 3) mass spectrometry as a novel proteome based approach. Since mass spectrometry and molecular approaches are the most promising tools for identifying a large variety of bacterial species, these two methods are described. Advances, limitations and potential problems when using these techniques are discussed.

Matrix-assisted laser desorption/ionization time of flight mass spectrometry (MALDI-TOF MS) and molecular techniques (16S rRNA gene sequencing) permit the identification of rare bacterial pathogens in routine diagnostics. The goal of this protocol lies in the combination of both techniques which leads to more accurate and reliable data.

There are a number of rare and, therefore, insufficiently described bacterial pathogens which are reported to cause severe infections especially in ...

A Simple and Efficient Approach to Construct Mutant Vaccinia Virus Vectors

The CRISPR-associated endonuclease Cas9 can edit particular genomic loci directed by a single guide RNA (gRNA). The CRISPR/Cas9 system has been successfully employed for editing genomes of various organisms. Here we describe a protocol for editing the vaccinia virus (VV) genome in the cytoplasm of VV-infected CV-1 cells using the RNA-guided Cas9. RNA-guided Cas9 induces double-stranded DNA breaks facilitating homologous recombination efficiently and specifically in the targeted site of VV and a transgene can be incorporated into these sites by homologous recombination. By using a site-specific homologous vector with transgene(s), a N1L gene-deleted VV with the red fluorescence protein (RFP) gene incorporated in this region was generated with a successful recombination efficiency 10 times greater than that obtained from the conventional homologous recombination method. This protocol demonstrates successful use of RNA-guided Cas9 system to generate mutant VVs with enhanced efficiency.

Vaccinia virus (VV) has been widely used in biomedical research and the improvement of human health. This article describes a simple, highly efficient method to edit the VV genome using a CRISPR-Cas9 system.

The CRISPR-associated endonuclease Cas9 can edit particular genomic loci directed by a single guide RNA (gRNA). The CRISPR/Cas9 system has been ...

A Method to Assess Fc-mediated Effector Functions Induced by Influenza Hemagglutinin Specific Antibodies

Antibodies play a crucial role in coupling the innate and adaptive immune responses against viral pathogens through their antigen binding domains and Fc-regions. Here, we describe how to measure the activation of Fc effector functions by monoclonal antibodies targeting the influenza virus hemagglutinin with the use of a genetically engineered Jurkat cell line expressing an activating type 1 Fc-Fc&#947;R. Using this method, the contribution of specific Fc-Fc&#947;R interactions conferred by immunoglobulins can be determined using an in vitro assay.

We describe a method to measure the activation of Fc-mediated effector functions by antibodies that target the influenza virus hemagglutinin. This assay can also be adapted to assess the ability of monoclonal antibodies or polyclonal sera targeting other viral surface glycoproteins to induce Fc-mediated immunity.

Antibodies play a crucial role in coupling the innate and adaptive immune responses against viral pathogens through their antigen binding domains and ...