Name: crispr/cas9 gene editing to make conditional mutants of human malaria parasite p. falciparum
Uploaded: 2018-09-18T12:30:00.000+00:00
Duration: 9 min 25 s
Description: Watch this Scientific Journal Video about CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum at JoVE.com

We thank Muthugapatti Kandasamy at the University of Georgia (UGA) Biomedical Microscopy Core for technical assistance and Jose-Juan Lopez-Rubio for sharing the pUF1-Cas9 and pL6 plasmids. This work was supported by ARCS Foundation awards to D.W.C. and to H.M.K., UGA startup funds to V.M., grants from the March of Dimes Foundation (Basil O&#39;Connor Starter Scholar Research Award) to V.M., and US National Institutes of Health grants (R00AI099156 and R01AI130139) to V.M. and (T32AI060546) to H.M.K.

Continuous culture of P. falciparum requires the use of human RBCs, and we utilized commercially purchased units of blood that were stripped of all identifiers and anonymized. The Institutional Review Board and the Office of Biosafety at the University of Georgia reviewed our protocols and approved all protocols used in our lab.
1. Choosing a gRNA Sequence
<ol>
	<li>Go to CHOPCHOP (http://chopchop.cbu.uib.no/) and select &#39;Fasta Target&#39;. Under &#39;Target&#39;, paste the 200 base pairs from the 3&#8217; end of the open reading frame (ORF) of a gene and 200 base pairs from the start of the gene&#8217;s 3&#8217;-UTR. Under &#39;In&#39;, select &#39;P. falciparum&#39; (3D7 v3.0), and select &#39;CRISPR/Cas9&#39; under &#39;Using&#39;. Next, click &#39;Find Target Sites&#39;.</li>
	<li>Select a gRNA sequence from the options presented, giving preference to the most efficient gRNA that is closest to the site of modification and that has the fewest off-target sites. 
	NOTE: Potential gRNA sequences are identified because they are immediately upstream of a Protospacer Adjacent Motif (PAM), which is required for recruitment of Cas9 to DNA. The sequence that is cloned into pMK-U6, the vector that drives gRNA expression, is the 20 bases immediately upstream of the PAM. The PAM specific for S. pyogenes Cas9 is the nucleotide sequence NGG and should not be included in the sequence that is cloned into pMK-U6. 
	NOTE: CHOPCHOP visually ranks the gRNA sequences, displaying the best options in green, the less ideal options in amber, and the worst options in red. CHOPCHOP gives each gRNA sequence an efficiency score that is calculated using the most up-to-date parameters found in the literature, and they predict off-target sites that could be recognized by the gRNA. Two or three gRNA sequences may need to be attempted to find the gRNA best suited to a particular gene.</li>
	<li>Purchase the gRNA sequence and its reverse-complement as Polyacrylamide Gel Electrophoresis-purified oligos. The gRNA sequence used to target PfHSP70x can be found in Figure 1B. 
	NOTE: This oligo should include 15 base pairs homologous to the gRNA-expressing plasmid, which are necessary for sequence and ligation-independent cloning (SLIC) into the pMK-U6 vector16.</li>
</ol>
2. Cloning the gRNA Sequence into pMK-U6
<ol>
	<li>Digest the pMK-U6 with BtgZI.
	<ol>
		<li>Digest 10 &#956;g of pMK-U6 with 5 &#956;L of BtgZI enzyme (5,000 units/mL) for 3 h at 60 &#176;C. Follow the enzyme manufacturer&#8217;s protocol for reaction conditions.</li>
		<li>After the 3 h incubation period, add an additional 3 &#956;L of BtgZI to the reaction to ensure complete digestion of the plasmid. Digest for an additional 3 h, still following the manufacturer&#8217;s instructions for ensuring the correct reaction conditions.</li>
		<li>To purify the digested pMK-U6 from the reaction, use a column-based PCR cleanup kit according to the manufacturer&#8217;s instructions.</li>
		<li>Separate the digested DNA using a 0.7% agarose gel and extract the 4,200-base pair band.</li>
	</ol>
	</li>
	<li>Anneal the oligos containing the gRNA sequence.
	<ol>
		<li>Reconstitute the PAGE-purified oligos to a concentration of 100 &#956;M using nuclease-free water.</li>
		<li>Combine 10 &#956;L of each oligo with 2.2 &#956;L of 10x buffer 2 (see Table of Materials). Ensure that the total reaction volume is 22.2 &#956;L.</li>
		<li>Run the gRNA annealing program in a thermocycler: Step 1: 95 &#176;C, 10 min; step 2: 95 &#176;C, 1 s, with a reduction in temperature of 0.6 &#176;C/cycle; step 3: Go to step 2, 16 times; step 4: 85 &#176;C, 1 min; step 5: 85 &#176;C, 1 s, with a reduction in temperature of 0.6&#176;C/cycle; step 6: Go to step 5, 16 times; step 7: 75 &#176;C, 1 min; step 8: 75 &#176;C, 1 s, with a reduction in temperature of 0.6 &#176;C/cycle; step 9: Go to step 8, 16 times. Steps 10 to 21: Repeat the procedure used in Steps 4&#8211;9 until the temperature reaches 25 &#176;C; step 22: 25 &#176;C, 1 min.</li>
	</ol>
	</li>
	<li>Insert the annealed gRNA oligos into the BtgZI-digested and gel-purified pMK-U6 plasmid.
	<ol>
		<li>Combine 100 ng of digested pMK-U6 with 1 &#956;L of 10x buffer 2.1 and 3 &#956;L of annealed gRNA oligos. Increase the volume to 9.5 &#956;L with nuclease-free water.</li>
		<li>Add 0.5 &#956;L of T4 polymerase and incubate the reaction at room temperature for 2.5 min.</li>
		<li>Move the reaction to ice and incubate for 10 min.</li>
		<li>Immediately transform 5 &#956;L of the reaction into competent E. coli according to the bacteria supplier&#8217;s instructions. Plate the bacteria on Lysogeny Broth (LB) agar plates containing 100 &#956;g/mL Ampicillin.</li>
		<li>Allow the transformed bacteria to grow at 37 &#176;C overnight, then select colonies and extract DNA with a commercially available plasmid miniprep kit.</li>
	</ol>
	</li>
</ol>
3. Designing Homology Regions of the Repair Template
<ol>
	<li>Design shield mutations within the homology repair template to prevent re-cutting of the DNA that is integrated into the genome. 
	NOTE: A shield mutation typically consists of introducing a silent mutation to alter the PAM so that Cas9 will not induce a break in the repair template. The PAM required for the Cas9 used in this protocol is the nucleotide sequence &#8220;NGG&#8221;, where &#8220;N&#8221; is any nucleotide. If possible, change one of the G nucleotides to an A, C, or T.
	<ol>
		<li>If the PAM cannot be silently mutated, introduce at least 2 silent mutations into the 6 base pairs directly adjacent to the PAM7,8. 
		NOTE: These mutations will prevent recognition of the repair template by the gRNA and prevent re-cutting of the repaired locus by the Cas9/gRNA complex. The shield mutations can be introduced into the homology region by amplifying the DNA with primers that contain the mutation.</li>
	</ol>
	</li>
	<li>Amplify the ORF homology region for the repair template.
	<ol>
		<li>Using PCR, amplify 800 base pairs from the 3&#8217; end of the target gene&#8217;s ORF. Design the primers that will be used to exclude the stop codon from this amplicon.</li>
		<li>Design the primers for insertion of this amplicon into the pHA-glmS that has been digested with SacII and AfeI through either a DNA ligation reaction or SLIC16.</li>
	</ol>
	</li>
	<li>Amplify the 3&#8217;-UTR homology region for the repair template.
	<ol>
		<li>Using PCR, amplify the 800 base pairs immediately following the stop codon of the target gene. Design primers for insertion of this amplicon into the pHA-glmS that has been digested with HindIII and NheI through either a DNA ligation reaction or SLIC16. 
		NOTE: The high AT content of the P. falciparum genome can make amplification of regions such as UTRs difficult. An alternative approach to using PCR is synthesizing the homology regions.</li>
	</ol>
	</li>
</ol>
4. Cloning Homology Regions into the Repair Plasmid
<ol>
	<li>Insert the ORF homology region into the pHA-glmS.
	<ol>
		<li>Digest the pHA-glmS with SacII and AfeI, according to the enzyme manufacturer&#8217;s instructions. Insert the ORF homology region PCR product into the digested plasmid using SLIC16.</li>
		<li>Transform into competent E. coli as performed in steps 2.3.4 and 2.3.5.</li>
	</ol>
	</li>
	<li>Insert the 3&#8217;-UTR homology region into a pHA-glmS plasmid that already contains the ORF homology region (see step 4.1).
	<ol>
		<li>Digest the plasmid with HindIII and NheI according to the enzyme manufacturer&#8217;s instructions. Insert the 3&#8217;-UTR homology region amplicon into the digested plasmid using SLIC16.</li>
		<li>Transform into competent E. coli and extract the plasmid DNA (steps 2.3.4 and 2.3.5).</li>
	</ol>
	</li>
</ol>
5. Precipitating DNA for Transfection
<ol>
	<li>Add 40 &#956;g each of pMK-U6, pUF1-Cas9, and pHA-glmS DNA (for a total of 120 &#956;g of DNA) into a sterile 1.5 mL microcentrifuge tube.</li>
	<li>Add 1/10th the volume of DNA of 3 M sodium acetate in water (pH 5.2) to the tube and mix it well using a vortex (e.g., if the volume in step 5.1 was 100 &#956;L, add 10 &#956;L of sodium acetate).</li>
	<li>Add 2.5 times the volume of 100% ethanol to the tube and mix it well using a vortex for at least 30 s (e.g., if the volume in 5.1 was 100 &#956;L, add 250 &#956;L of 100% ethanol).</li>
	<li>Place the tube on ice or at -20 &#176;C for 30 min.</li>
	<li>Centrifuge the tube at 18,300 x g for 30 min at 4 &#176;C.</li>
	<li>Carefully remove the supernatant from the tube. Do not disturb the pellet.</li>
	<li>Add 3 times the volume of 70% ethanol to the tube and mix it briefly using a vortex (e.g., if the volume in 5.1 was 100 &#956;L, add 300 &#956;L of 70% ethanol).</li>
	<li>Centrifuge the tube at 18,300 x g for 30 min at 4 &#176;C. 
	NOTE: This step should be performed under sterile conditions in a biological safety cabinet.</li>
	<li>Carefully remove the supernatant from the tube. Do not disturb the pellet. Leave the tube open and allow the pellet to air-dry for 15 min.</li>
	<li>Store the precipitated DNA at -20 &#176;C until it is needed for transfection.</li>
</ol>
6. Isolating Human RBCs from Whole Blood in Preparation for Transfection
<ol>
	<li>Aliquot fresh blood into sterile 50 mL conical tubes (approximately 25 mL per tube).</li>
	<li>Centrifuge the tubes at 1,088 x g for 12 min, with centrifuge brakes set to 4.</li>
	<li>Aspirate off the supernatant and buffy coat. Resuspend the RBC pellet with an equal volume of incomplete RPMI. 
	Note: Incomplete RPMI is prepared by supplementing RPMI 1640 with 10.32 &#956;M thymidine, 110.2 &#956;M hypoxanthine, 1 mM sodium pyruvate, 30 mM sodium bicarbonate, 5 mM HEPES, 11.1 mM glucose, and 0.02% (v/v) gentamicin.</li>
	<li>Repeat steps 6.2&#8211;6.3 twice. After the last wash, resuspend the RBCs in an equal volume of incomplete RPMI and store the tubes at 4 &#176;C.</li>
</ol>
7. Transfecting RBCs with the CRISPR/Cas9 Plasmids (To Be Done Aseptically)
NOTE: P. falciparum cultures are maintained as described in other reports17. Maintain all the cultures at 37 &#176;C under 3% O2, 3% CO2, and 94% N2 unless stated otherwise. Whenever blood is used in this protocol, it is referring to the pure red blood cells prepared in step 6. The blood used should not be older than 6 weeks, as there is typically a decrease in parasite proliferation in older blood. The following steps describe pre-loading RBCs with DNA and adding a parasite culture to the transfected cells. Other established transfection protocols are compatible with transfecting these constructs18,19.
<ol>
	<li>Prepare a 1x cytomix buffer in water (120 mM KCl, 0.15 mM CaCl2, 2 mM EGTA, 5 mM MgCl2, 10 mM K2HPO4, 25 mM HEPES, pH 7.6). Filter-sterilize the buffer using a 0.22 &#956;m filter.</li>
	<li>Add 380 &#956;L of the 1x cytomix to the DNA precipitated in step 5, and vortex to dissolve. Allow the DNA to dissolve in the 1x cytomix for 10 minutes, vortexing every 3 min for 10 s.</li>
	<li>In a sterile 15 mL conical tube, combine 300 &#956;L of RBCs (50% hematocrit, from step 6) in the incomplete RPMI with 4 mL of 1x cytomix.</li>
	<li>Centrifuge the RBCs from step 7.3 at 870 x g for 3 min, and then remove the supernatant from the RBC pellet.</li>
	<li>Resuspend the RBC pellet with the DNA/cytomix mixture from step 6.2 and transfer to a 0.2 cm electroporation cuvette.</li>
	<li>Electroporate the RBCs using the following conditions: 0.32 kV, 925 &#956;F, capacitance set to &#8220;High Cap&#8221;, and resistance set to &#8220;Infinite&#8221;.</li>
	<li>Following electroporation, transfer the contents from the cuvette to a 15 mL conical containing 5 mL of complete RPMI (cRPMI). Centrifuge the tube at 870 x g for 3 min at 20 &#176;C, and then decant the supernatant. 
	NOTE: cRPMI is prepared through the same method as incomplete RPMI with the addition of 0.25% (w/v) lipid-rich bovine serum albumin.</li>
	<li>Resuspend the pellet in 4 mL of cRPMI and transfer to one well in a 6-well tissue culture plate. Add 400 &#956;L of a high-schizont culture (7&#8211;10% schizont parasitemia is ideal) to the transfected RBCs. 
	NOTE: Parasitemia is defined as the percentage of parasite-infected RBCs.</li>
	<li>The next day, wash the culture with 4 mL of cRPMI. Centrifuge the culture at 870 x g for 3 min and aspirate the supernatant. Resuspend the culture in 4 mL of cRPMI.</li>
	<li>48 h after completing step 7.6, wash the culture with 4 mL of cRPMI. Then resuspend the culture in cRPMI containing 1 &#956;M DSM1 to select for the Cas9 plasmid.</li>
	<li>Continue washing the cultures each day with cRPMI until parasites are no longer visible by blood smear. After this point, replace the culture medium with fresh cRPMI plus 1 &#956;M DSM1 every 48 h.
	<ol>
		<li>To make a blood smear, pipette 150 &#956;L of culture into a 0.6 mL centrifuge tube. Pellet the cells by centrifugation at 1,700 x g for 30 s.</li>
		<li>Aspirate off the supernatant. Use a pipette to transfer the pelleted cells to a glass slide. Using a second glass slide held at a 45&#176; angle to the first slide, smear the blood droplet. Stain the slide using a commercially available staining kit according to the manufacturer&#8217;s protocol.</li>
		<li>View the parasites using a 100X oil immersion objective.</li>
	</ol>
	</li>
	<li>Beginning 5 days post-transfection (step 7.6), remove 2 mL of the culture with RBCs resuspended in the culture medium. Add back 2 mL of fresh medium (cRPMI plus 1 &#956;M DSM1) and blood at 2% hematocrit. Add fresh blood in this manner once a week until parasites reappear, as determined by thin blood smear (step 7.11). 
	NOTE: If integration is successful, parasites generally reappear in the culture by one-month post-transfection.</li>
	<li>Once parasites reemerge, remove DSM1 drug pressure. Alternatively, remove drug pressure after parasites have been cloned out.</li>
</ol>
8. Checking Parasites for Integration of the Repair Template
<ol>
	<li>When parasites are visible again by thin blood smear, isolate DNA from the culture using an appropriate kit.</li>
	<li>Use PCR to amplify the modified region of the genome to determine if the targeted locus has been successfully altered and if the unmodified wild-type locus (indicative of wild-type parasites) is detectable.
	<ol>
		<li>To detect parasites that have integrated the repair template, use a forward primer that sits at the beginning of the ORF, outside of the cloned homology region. Use a reverse primer that sits in the 3&#8217;-UTR. 
		NOTE: As this amplification includes the sequences of the HA tags and glmS ribozyme, amplicons from integrated parasites will be longer than the same region amplified in wild-type parasites.</li>
	</ol>
	</li>
</ol>
9. Cloning Parasites by Limiting Dilution
<ol>
	<li>Perform serial dilutions of the parasite culture from step 7.13 to achieve a final concentration of 0.5 parasites/200 &#956;L. Add 200 &#956;L of the diluted culture to the wells of a 96-well tissue culture plate. 
	NOTE: Because parasitemia is defined as the percentage of infected RBCs and the hematocrit is also a defined number (2%), the number of parasites per unit volume is easily inferred.
	<ol>
		<li>Prepare 1 mL of culture in cRPMI at 5% parasitemia and 2% hematocrit (at these parasitemia and hematocrit levels, the culture contains 1 x 107 parasites/mL).</li>
		<li>Dilute this culture 1:100 with cRPMI. Dilute again 1:100 with cRPMI.</li>
		<li>Dilute 1:400. Perform this dilution by adding 62.5 &#956;L of culture to 25 mL of cRPMI and 1 mL of blood. This dilution results in the desired concentration of 0.5 parasites/200 &#956;L.</li>
	</ol>
	</li>
	<li>Maintain the cloning plate until parasites are detectable in the wells.
	<ol>
		<li>Every 48 h, replace the medium in the 96-well plate with fresh medium.</li>
		<li>Once a week, starting 5 days after beginning the cloning plate (step 9.1), remove 100 &#956;L from each well and add back 100 &#956;L of fresh medium + blood (2% hematocrit).</li>
	</ol>
	</li>
	<li>Identify any wells containing parasites.
	<ol>
		<li>Place the 96-well plate at a 45&#176; angle for approximately 20 min, allowing the blood to settle at an angle within the plate.</li>
		<li>Place the 96-well plate on a light box. Notice that the wells containing parasites contain media that is yellow in color, compared to the pink media of parasite-free wells, due to acidification of the medium by the parasites.</li>
		<li>Using a serological pipette, move the contents of the parasite-containing wells to a 24-well tissue culture plate to allow expansion of the parasitemia.</li>
		<li>Using PCR analysis as described in step 8, check these clonal parasite lines for correct integration.</li>
	</ol>
	</li>
</ol>
10. Knockdown of the Protein by Treating Parasites with Glucosamine and Confirmation via Western Blot Analysis
<ol>
	<li>Prepare a 0.5 M GlcN (glucosamine) stock solution, which can be stored at -20&#176; C.</li>
	<li>Add GlcN to the glmS parasite cultures and allow them to grow in the presence of GlcN. 
	NOTE: The final concentration and timing of GlcN treatment depends on the experiment and parasite line. GlcN can impact parasite growth, so the parental parasite strain should be exposed to a range of GlcN concentrations to determine its sensitivity to the compound. Often, a concentration of 2.0-7.5 mM GlcN is used13,14,20.</li>
	<li>Isolate protein samples from the GlcN-treated parasites13.</li>
	<li>Use protein samples for western blot analysis to detect reductions in the protein13.
	<ol>
		<li>Use an anti-HA antibody according to the manufacturer&#8217;s instructions to detect the HA-glmS-tagged protein. Compare the HA band to a loading control, such as PfEF1&#945;.</li>
	</ol>
	</li>
</ol>

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J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High+efficiency+transfection+of+Plasmodium+berghei+facilitates+novel+selection+procedures.">High efficiency transfection of Plasmodium berghei facilitates novel selection procedures.</a> Molecular and Biochemical Parasitology. 145 (1), 60-70 (2006).</li><li>Counihan, N. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plasmodium+falciparum+parasites+deploy+RhopH2+into+the+host+erythrocyte+to+obtain+nutrients,+grow+and+replicate.">Plasmodium falciparum parasites deploy RhopH2 into the host erythrocyte to obtain nutrients, grow and replicate.</a> eLife. 6, (2017).</li><li>Klemba, M., Beatty, W., Gluzman, I., Goldberg, D. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Trafficking+of+plasmepsin+II+to+the+food+vacuole+of+the+malaria+parasite+Plasmodium+falciparum.">Trafficking of plasmepsin II to the food vacuole of the malaria parasite Plasmodium falciparum.</a> Journal of Cell Biology. 164 (1), 47-56 (2004).</li><li>Labun, K., Montague, T. G., Gagnon, J. A., Thyme, S. B., Valen, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CHOPCHOP+v2:+a+web+tool+for+the+next+generation+of+CRISPR+genome+engineering.">CHOPCHOP v2: a web tool for the next generation of CRISPR genome engineering.</a> Nucleic Acids Resesarch. 44 (W1), W272-W276 (2016).</li><li>Peng, D., Tarleton, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=EuPaGDT:+a+web+tool+tailored+to+design+CRISPR+guide+RNAs+for+eukaryotic+pathogens.">EuPaGDT: a web tool tailored to design CRISPR guide RNAs for eukaryotic pathogens.</a> Microbial Genomes. 1 (4), e000033 (2015).</li><li>Shen, B., Brown, K. M., Lee, T. D., Sibley, L. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+Gene+Disruption+in+Diverse+Strains+of+Toxoplasma+gondii+Using+CRISPR/CAS9.">Efficient Gene Disruption in Diverse Strains of Toxoplasma gondii Using CRISPR/CAS9.</a> mBio. 5 (3), (2014).</li><li>Janssen, B. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR/Cas9-mediated+gene+modification+and+gene+knock+out+in+the+human-infective+parasite+Trichomonas+vaginalis.">CRISPR/Cas9-mediated gene modification and gene knock out in the human-infective parasite Trichomonas vaginalis.</a> Scientific Reports. 8 (1), 270 (2018).</li><li>Spillman, N. J., Beck, J. R., Ganesan, S. M., Niles, J. C., Goldberg, D. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+chaperonin+TRiC+forms+an+oligomeric+complex+in+the+malaria+parasite+cytosol.">The chaperonin TRiC forms an oligomeric complex in the malaria parasite cytosol.</a> Cellular Microbiology. 19 (6), (2017).</li><li>Brancucci, N. M. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lysophosphatidylcholine+Regulates+Sexual+Stage+Differentiation+in+the+Human+Malaria+Parasite+Plasmodium+falciparum.">Lysophosphatidylcholine Regulates Sexual Stage Differentiation in the Human Malaria Parasite Plasmodium falciparum.</a> Cell. , (2017).</li><li>Ng, C. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR-Cas9-modified+pfmdr1+protects+Plasmodium+falciparum+asexual+blood+stages+and+gametocytes+against+a+class+of+piperazine-containing+compounds+but+potentiates+artemisinin-based+combination+therapy+partner+drugs.">CRISPR-Cas9-modified pfmdr1 protects Plasmodium falciparum asexual blood stages and gametocytes against a class of piperazine-containing compounds but potentiates artemisinin-based combination therapy partner drugs.</a> Molecular Microbiology. 101 (3), 381-393 (2016).</li><li>Lim, M. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=UDP-galactose+and+acetyl-CoA+transporters+as+Plasmodium+multidrug+resistance+genes.">UDP-galactose and acetyl-CoA transporters as Plasmodium multidrug resistance genes.</a> Nature Microbiology. , (2016).</li><li>Adjalley, S. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+assessment+of+Plasmodium+falciparum+sexual+development+reveals+potent+transmission+blocking+activity+by+methylene+blue.">Quantitative assessment of Plasmodium falciparum sexual development reveals potent transmission blocking activity by methylene blue.</a> Proceedings of the National Academy of Sciences USA. 108 (47), E1214-E1223 (2011).</li><li>Sidik, S. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Genome-wide+CRISPR+Screen+in+Toxoplasma+Identifies+Essential+Apicomplexan+Genes.">A Genome-wide CRISPR Screen in Toxoplasma Identifies Essential Apicomplexan Genes.</a> Cell. 166 (6), 1423-1435 (2016).</li><li>Amberg-Johnson, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Small+molecule+inhibition+of+apicomplexan+FtsH1+disrupts+plastid+biogenesis+in+human+pathogens.">Small molecule inhibition of apicomplexan FtsH1 disrupts plastid biogenesis in human pathogens.</a> elife. 6, (2017).</li><li>Crawford, E. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plasmid-free+CRISPR/Cas9+genome+editing+in+Plasmodium+falciparum+confirms+mutations+conferring+resistance+to+the+dihydroisoquinolone+clinical+candidate+SJ733.">Plasmid-free CRISPR/Cas9 genome editing in Plasmodium falciparum confirms mutations conferring resistance to the dihydroisoquinolone clinical candidate SJ733.</a> Public Library of Science One. 12 (5), e0178163 (2017).</li></ol>

The authors have nothing to disclose.

The implementation of CRISPR/Cas9 in P. falciparum has both increased the efficiency of and decreased the amount of time needed for modifying the parasite&#39;s genome, compared to previous methods of genetic manipulation. This comprehensive protocol outlines the steps necessary for generating conditional mutants using CRISPR/Cas9 in P. falciparum. While the method here is geared specifically for the generation of HA-glmS mutants, this strategy can be adapted for a variety of needs, including t...

Malaria is a devastating disease caused by protozoan parasites of the genus Plasmodium. P. falciparum, the most deadly human malaria parasite, causes approximately 445,000 deaths per year, mostly in children under the age of five1. Plasmodium parasites have an intricate life cycle involving a mosquito vector and a vertebrate host. Humans first become infected when an infected mosquito takes a blood meal. Then, the parasite invades the liver where it grows, develops, and divides for approximately one week.After this process, the parasites are released into the bloodstream, where they undergo asexual replication in red blood cells (RBCs). Growth of the parasites within the RBCs are directly responsible for the clinical symptoms associated with malaria2.
Until recently, production of transgenic P. falciparum was a laborious process, involving several rounds of drug selection that took many months and had a high failure rate. This time-consuming procedure relieson the generation of random DNA breaks in the region of interest and the endogenous ability of the parasite to mend its genome though homologous repair3,4,5,6. Recently, Clustered Regularly Interspaced Palindromic Repeat/Cas9 (CRISPR/Cas9) genome editing has been successfully utilized in P. falciparum7,8. The introduction of this new technology in malaria research has been critical for advancing understanding of the biology of these deadly Plasmodium parasites. CRISPR/Cas9 allows for specific targeting of genes through guide RNAs (gRNAs) that are homologous to the gene of interest. The gRNA/Cas9 complex recognizes the gene through the gRNA, and Cas9 then introduces a double-strand break, forcing the initiation of repair mechanisms in the organism9,10. Because P. falciparum lacks the machinery to repair DNA breaks through non-homologous end joining, it utilizes homologous recombination mechanisms and integrates transfected homologous DNA templates to repair the Cas9/gRNA-induced double-strand break11,12.
Here, we present a protocol for the generation of conditional knockdown mutants in P. falciparum using CRISPR/Cas9 genome editing. The protocol demonstrates usage of the glmS ribozyme to conditionally knockdown protein levels of PfHsp70x (PF3D7_0831700), a chaperone exported by P. falciparum into host RBCs13,14. The glmS ribozyme is activated by treatment with glucosamine (which is converted to glucosamine-6-phosphate in cells) to cleave its associated mRNA, leading to a reduction in the protein14. This protocol can be easily adapted to utilize other conditional knockdown tools, such as destabilization domains or RNA aptamers4,5,15. Our protocol details the generation of a repair plasmid consisting of a hemagglutinin (HA) tag and glmS ribozyme coding sequence flanked by sequences that are homologous to the PfHsp70x open reading frame (ORF) and 3&#39;-UTR. We also describe the generation of a second plasmid to drive expression of the gRNA. These two plasmids, along with a third that drives expression of Cas9, are transfected into RBCs and used to modify the genome of P. falciparum parasites. Finally, we describe a polymerase chain reaction (PCR)-based technique to verify integration of the tag and glmS ribozyme. This protocol is highly adaptable for the modification or complete knockout of any P. falciparum genes, enhancing our ability to generate new insights into the biology of the malaria parasite.

<table><tbody><tr><td>Gene Pulser Xcell Electroporator</td><td>Bio-Rad</td><td>1652660</td><td></td></tr><tr><td>Gene Pulser Xcell Electroporator</td><td>Bio-Rad</td><td>165-2086</td><td>We buy the ones that are individually wrapped</td></tr><tr><td>Sodium Acetate</td><td>Sigma-Aldrich</td><td>S2889-250g</td><td></td></tr><tr><td>DSM1</td><td>Gift from Akhil Vaidya lab</td><td>Ganesan &lt;em&gt;et al.&lt;/em&gt; Mol. Biochem. Parasitol. 2011 177:29-34</td><td></td></tr><tr><td>TPP Tissue Culture 6 Well Plates</td><td>MIDSCI</td><td>TP92006</td><td></td></tr><tr><td>TPP 100 mm Tissue Culture Dishes (12 mL Plate)</td><td>MIDSCI</td><td>TP93100</td><td></td></tr><tr><td>TPP Tissue Culture 96 Well Plates</td><td>MIDSCI</td><td>TP92096</td><td></td></tr><tr><td>TPP Tissue Culture 24 Well Plates</td><td>MIDSCI</td><td>TP92024</td><td></td></tr><tr><td>NEBuffer 2</td><td>New England Biolabs</td><td>#B7002S</td><td></td></tr><tr><td>NEBuffer 2.1</td><td>New England Biolabs</td><td>#B7202S</td><td></td></tr><tr><td>BtgZI</td><td>New England Biolabs</td><td>#R0703L</td><td></td></tr><tr><td>SacII</td><td>New England Biolabs</td><td>#R0157L</td><td></td></tr><tr><td>HindIII-HF</td><td>New England Biolabs</td><td>#R3104S</td><td></td></tr><tr><td>Afe1</td><td>New England Biolabs</td><td>#R0652S</td><td></td></tr><tr><td>Nhe1-HF</td><td>New England Biolabs</td><td>#R3131L</td><td></td></tr><tr><td>T4 DNA Polymerase</td><td>New England Biolabs</td><td>#M0203S</td><td></td></tr><tr><td>500 mL Steritop bottle top filter unit</td><td>Millipore</td><td>SCGPU10RE</td><td>You can use any size that fits your needs</td></tr><tr><td>EGTA</td><td>Sigma</td><td>E4378-100G</td><td></td></tr><tr><td>KCl</td><td>Sigma-Aldrich</td><td>P9333-500g</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>C7902-500g</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>M8266-100g</td><td></td></tr><tr><td>K&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Fisher</td><td>P288-500</td><td></td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H4034-500g</td><td></td></tr><tr><td>pMK-U6</td><td>Generated by the Muralidharan Lab</td><td>n/a</td><td></td></tr><tr><td>pHA-glmS</td><td>Generated by the Muralidharan Lab</td><td>n/a</td><td></td></tr><tr><td>pUF1-Cas9</td><td>Gift from the Jose-Juan Lopez-Rubio Lab</td><td>Ghorbal &lt;em&gt;et al.&lt;/em&gt; Nature Biotech 2014</td><td></td></tr><tr><td>Glucose</td><td>Sigma-Aldrich</td><td>G7021-1KG</td><td></td></tr><tr><td>Sodium bicarbonate</td><td>Sigma-Aldrich</td><td>S5761-500G</td><td></td></tr><tr><td>Sodium pyruvate</td><td>Sigma-Aldrich</td><td>P5280-100G</td><td></td></tr><tr><td>Hypoxanthine</td><td>Sigma-Aldrich</td><td>H9636-25g</td><td></td></tr><tr><td>Gentamicin Reagent</td><td>Gibco</td><td>15710-064</td><td></td></tr><tr><td>Thymidine</td><td>Sigma-Aldrich</td><td>T1895-1G</td><td></td></tr><tr><td>PL6-eGFP BSD</td><td>Generated by the Muralidharan Lab</td><td></td><td></td></tr><tr><td>Puf1-cas9 eGFP gRNA</td><td>Generated by the Muralidharan Lab</td><td></td><td></td></tr><tr><td>NucleoSpin Gel and PCR Clean-up</td><td>Macherey-Nagel</td><td>740609.250</td><td></td></tr><tr><td>Albumax I</td><td>Life Technologies</td><td>N/A</td><td>You will want to try a few batches to find out what the parasites will grow in best</td></tr><tr><td>Human Red Blood Cells</td><td>Interstate Blood Bank, Inc</td><td>Email or call them directly for ordering</td><td>We typically use O+ blood</td></tr><tr><td>3D7 parasite line</td><td>Available upon request</td><td>N/A</td><td></td></tr><tr><td>Lysogeny Broth (LB)</td><td>Fisher</td><td>BP1426-2</td><td>You can make your own, it is not necessary to use exactly this</td></tr><tr><td>Ampicilin</td><td>Fisher</td><td>BP1760-25</td><td>We make a 1000X stock at 100mg/ml in water and store in the -20C</td></tr><tr><td>Ampicilin</td><td>Clonetech</td><td>R050A</td><td></td></tr><tr><td>Anti-EF1alpha</td><td>Dr. Daniel Goldberg&#039;s Lab</td><td>Washington University in St. Louis</td><td>You can use your preferred loading control for western blots. This is just the one we use in our laboratory</td></tr><tr><td>Rat Anti-HA Clone 3F10, monoclonal</td><td>Made by Roche, sold by Sigma</td><td>11867423001</td><td>You can use your preferred anti-HA antibody</td></tr><tr><td>0.6 mL tubes</td><td>Fisher</td><td>AB0350</td><td></td></tr><tr><td>Fisher HealthCare* PROTOCOL* Hema 3* Manual Staining System (Fixative+Solution I and II)</td><td>Fisher</td><td>22-122-911</td><td>You can also use giemsa stain</td></tr><tr><td>Fisherfines Premium Frosted Microscope Slides - Size: 3 x 1 in.</td><td>Fisher</td><td>12-544-3</td><td></td></tr></tbody></table>

crispr/cas9 gene editing to make conditional mutants of human malaria parasite p. falciparum

Malaria is a significant cause of morbidity and mortality worldwide. This disease, which primarily affects those living in tropical and subtropical regions, is caused by infection with Plasmodium parasites. The development of more effective drugs to combat malaria can be accelerated by improving our understanding of the biology of this complex parasite. Genetic manipulation of these parasites is key to understanding their biology; however, historically the genome of P. falciparum has been difficult to manipulate. Recently, CRISPR/Cas9 genome editing has been utilized in malaria parasites, allowing for easier protein tagging, generation of conditional protein knockdowns, and deletion of genes. CRISPR/Cas9 genome editing has proven to be a powerful tool for advancing the field of malaria research. Here, we describe a CRISPR/Cas9 method for generating glmS-based conditional knockdown mutants in P. falciparum. This method is highly adaptable to other types of genetic manipulations, including protein tagging and gene knockouts.

A schematic of the plasmids used in this method as well as an example of a shield mutation are shown in Figure 1. As an example of how to identify mutant parasites after transfection, results from PCRs for checking integration of the HA-glmS construct are shown in Figure 2. A representative image of a cloning plate is shown in Figure 3 to demonstrate the color change of the medium in the pre...

Watch this Scientific Journal Video about CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum at JoVE.com

CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

We describe a method for generating glmS-based conditional knockdown mutants in Plasmodium falciparum using CRISPR/Cas9 genome editing.

CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

Malaria is a significant cause of morbidity and mortality worldwide. This disease, which primarily affects those living in tropical and subtropical ...

crisprcas9-gene-editing-to-make-conditional-mutants-human-malaria

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Genetics

9.9K Views.  University of Georgia. We describe a method for generating glmS-based conditional knockdown mutants in Plasmodium falciparum using CRISPR/Cas9 genome editing.

Video: CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

QTL Mapping and CRISPR/Cas9 Editing to Identify a Drug Resistance Gene in Toxoplasma gondii

Scientific knowledge is intrinsically linked to available technologies and methods. This article will present two methods that allowed for the identification and verification of a drug resistance gene in the Apicomplexan parasite Toxoplasma gondii, the method of Quantitative Trait Locus (QTL) mapping using a Whole Genome Sequence (WGS) -based genetic map and the method of Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas9 -based gene editing. The approach of QTL mapping allows one to test if there is a correlation between a genomic region(s) and a phenotype. Two datasets are required to run a QTL scan, a genetic map based on the progeny of a recombinant cross and a quantifiable phenotype assessed in each of the progeny of that cross. These datasets are then formatted to be compatible with R/qtl software that generates a QTL scan to identify significant loci correlated with the phenotype. Although this can greatly narrow the search window of possible candidates, QTLs span regions containing a number of genes from which the causal gene needs to be identified. Having WGS of the progeny was critical to identify the causal drug resistance mutation at the gene level. Once identified, the candidate mutation can be verified by genetic manipulation of drug sensitive parasites. The most facile and efficient method to genetically modify T. gondii is the CRISPR/Cas9 system. This system comprised of just 2 components both encoded on a single plasmid, a single guide RNA (gRNA) containing a 20 bp sequence complementary to the genomic target and the Cas9 endonuclease that generates a double-strand DNA break (DSB) at the target, repair of which allows for insertion or deletion of sequences around the break site. This article provides detailed protocols to use CRISPR/Cas9 based genome editing tools to verify the gene responsible for sinefungin resistance and to construct transgenic parasites.

QTL Mapping and CRISPR/Cas9 Editing to Identify a Drug Resistance Gene in Toxoplasma gondii

Details are presented on how QTL mapping with a whole genome sequence based genetic map can be used to identify a drug resistance gene in Toxoplasma gondii and how this can be verified with the CRISPR/Cas9 system that efficiently edits a genomic target, in this case the drug resistance gene.

Scientific knowledge is intrinsically linked to available technologies and methods. This article will present two methods that allowed for the ...

Selection-dependent and Independent Generation of CRISPR/Cas9-mediated Gene Knockouts in Mammalian Cells

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide RNA (sgRNA) that confers specificity, the Cas9 protein cleaves both DNA strands at the targeted locus. The DNA break can trigger either non-homologous end joining (NHEJ) or homology directed repair (HDR). NHEJ can introduce small deletions or insertions which lead to frame-shift mutations, while HDR allows for larger and more precise perturbations. Here, we present protocols for generating knockout cell lines by coupling established CRISPR/Cas9 methods with two options for downstream selection/screening. The NHEJ approach uses a single sgRNA cut site and selection-independent screening, where protein production is assessed by dot immunoblot in a high-throughput manner. The HDR approach uses two sgRNA cut sites that span the gene of interest. Together with a provided HDR template, this method can achieve deletion of tens of kb, aided by the inserted selectable resistance marker. The appropriate applications and advantages of each method are discussed.

Recent advances in the ability to genetically manipulate somatic cell lines hold great potential for basic and applied research. Here, we present two approaches for CRISPR/Cas9 generated knockout production and screening in mammalian cell lines, with and without the use of selectable markers.

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide ...

Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has quickly become a commonplace amongst researchers pursuing a wide variety of biological questions. Users most often employ the Cas9 protein derived from Streptococcus pyogenes in a complex with an easily reprogrammed guide RNA (gRNA). These components are introduced into cells, and through a base pairing with a complementary region of the double-stranded DNA (dsDNA) genome, the enzyme cleaves both strands to generate a double-strand break (DSB). Subsequent repair leads to either random insertion or deletion events (indels) or the incorporation of experimenter-provided DNA at the site of the break.
The use of a purified single-guide RNA and Cas9 protein, preassembled to form an RNP and delivered directly to cells, is a potent approach for achieving highly efficient gene editing. RNP editing particularly enhances the rate of gene insertion, an outcome that is often challenging to achieve. Compared to the delivery via a plasmid, the shorter persistence of the Cas9 RNP within the cell leads to fewer off-target events. 
Despite its advantages, many casual users of CRISPR gene editing are less familiar with this technique. To lower the barrier to entry, we outline detailed protocols for implementing the RNP strategy in a range of contexts, highlighting its distinct benefits and diverse applications. We cover editing in two types of primary human cells, T cells and hematopoietic stem/progenitor cells (HSPCs). We also show how Cas9 RNP editing enables the facile genetic manipulation of entire organisms, including the classic model roundworm Caenorhabditis elegans and the more recently introduced model crustacean, Parhyale hawaiensis.

Utilizing a preassembled Cas9 ribonucleoprotein complex (RNP) is a powerful method for precise, efficient genome editing. Here, we highlight its utility across a broad range of cells and organisms, including primary human cells and both classic and emerging model organisms.

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has ...

Introducing a Gene Knockout Directly Into the Amastigote Stage of Trypanosoma cruzi Using the CRISPR/Cas9 System

Trypanosoma cruzi is a pathogenic protozoan parasite that causes Chagas&#8217; disease mainly in Latin America. In order to identify a novel drug target against T. cruzi, it is important to validate the essentiality of the target gene in the mammalian stage of the parasite, the amastigote. Amastigotes of T. cruzi replicate inside the host cell; thus, it is difficult to conduct a knockout experiment without going through other developmental stages. Recently, our group reported a growth condition in which the amastigote can replicate axenically for up to 10 days without losing its amastigote-like properties. By using this temporal axenic amastigote culture, we successfully introduced gRNAs directly into the Cas9-expressing amastigote to cause gene knockouts and analyzed their phenotypes exclusively in the amastigote stage. In this report, we describe a detailed protocol to produce in vitro derived extracellular amastigotes, and to utilize the axenic culture in a CRISPR/Cas9-mediated knockout experiment. The growth phenotype of knockout amastigotes can be evaluated either by cell counts of the axenic culture, or by replication of intracellular amastigote after host cell invasion. This method bypasses the parasite stage differentiation normally involved in producing a transgenic or a knockout amastigote. Utilization of the temporal axenic amastigote culture has the potential to expand the experimental freedom of stage-specific studies in T. cruzi.

Introducing a Gene Knockout Directly Into the Amastigote Stage of Trypanosoma cruzi Using the CRISPR/Cas9 System

Here, we describe a protocol to introduce a gene knockout into the extracellular amastigote of Trypanosoma cruzi, using the CRISPR/Cas9 system. The growth phenotype can be followed up either by cell counting of axenic amastigote culture or by proliferation of intracellular amastigotes after host cell invasion.

Trypanosoma cruzi is a pathogenic protozoan parasite that causes Chagas&#8217; disease mainly in Latin America. In order to identify a novel drug target ...

Generation of Defined Genomic Modifications Using CRISPR-CAS9 in Human Pluripotent Stem Cells

Human pluripotent stem cells offer a powerful system to study gene function and model specific mutations relevant to disease. The generation of precise heterozygous genetic modifications is challenging due to CRISPR-CAS9 mediated indel formation in the second allele. Here, we demonstrate a protocol to help overcome this difficulty by using two repair templates in which only one expresses the desired sequence change, while both templates contain silent mutations to prevent re-cutting and indel formation. This methodology is most advantageous for gene editing coding regions of DNA to generate isogenic control and mutant human stem cell lines for studying human disease and biology. In addition, optimization of transfection and screening methodologies have been performed to reduce labor and cost of a gene editing experiment. Overall, this protocol is widely applicable to many genome editing projects utilizing the human pluripotent stem cell model.

This protocol provides a method to facilitate the generation of defined heterozygous or homozygous nucleotide changes using CRISPR-CAS9 in human pluripotent stem cells.

Human pluripotent stem cells offer a powerful system to study gene function and model specific mutations relevant to disease. The generation of precise ...

Introducing Point Mutations into Human Pluripotent Stem Cells Using Seamless Genome Editing

Custom designed endonucleases, such as RNA-guided Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)-Cas9, enable efficient genome editing in mammalian cells. Here we describe detailed procedures to seamlessly genome edit the hepatocyte nuclear factor 4 alpha (HNF4&#945;) locus as an example in human pluripotent stem cells. Combining a piggyBac-based donor plasmid and the CRISPR-Cas9 nickase mutant in a two-step genetic selection, we demonstrate correct and efficient targeting of the HNF4&#945; locus.

Here, we describe a detailed method for seamless gene editing in human pluripotent stem cells using a piggyBac-based donor plasmid and the Cas9 nickase mutant. Two point mutations were introduced into exon 8 of the hepatocyte nuclear factor 4 alpha (HNF4&#945;) locus in human embryonic stem cells (hESCs).

Custom designed endonucleases, such as RNA-guided Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)-Cas9, enable efficient genome editing ...

Sand Fly (Phlebotomus papatasi) Embryo Microinjection for CRISPR/Cas9 Mutagenesis

Sand flies are the natural vectors for Leishmania species, protozoan parasites producing a broad spectrum of symptoms ranging from cutaneous lesions to visceral pathology. Deciphering the nature of the vector/parasite interactions is of primary importance for better understanding of Leishmania transmission to their hosts. Among the parameters controlling the sand fly vector competence (i.e. their ability to carry and transmit pathogens), parameters intrinsic to these insects were shown to play a key role. Insect immune response, for example, impacts sand fly vector competence to Leishmania. The study of such parameters has been limited by the lack of methods of gene expression modification adapted for use in these non-model organisms. Gene downregulation by small interfering RNA (siRNA) is possible, but in addition to being technically challenging, the silencing leads to only a partial loss of function, which cannot be transmitted from generation to generation. Targeted mutagenesis by CRISPR/Cas9 technology was recently adapted to the Phlebotomus papatasi sand fly. This technique leads to the generation of transmissible mutations in a specifically chosen locus, allowing to study the genes of interest. The CRISPR/Cas9 system relies on the induction of targeted double-strand DNA breaks, later repaired by either Non-Homologous End Joining (NHEJ) or by Homology Driven Repair (HDR). NHEJ consists of a simple closure of the break and frequently leads to small insertion/deletion events. In contrast, HDR uses the presence of a donor DNA molecule sharing homology with the target DNA as a template for repair. Here, we present a sand fly embryo microinjection method for targeted mutagenesis by CRISPR/Cas9 using NHEJ, which is the only genome modification technique adapted to sand fly vectors to date.

Sand Fly Phlebotomus papatasi Embryo Microinjection for CRISPR/Cas9 Mutagenesis

This protocol details the steps of CRISPR/Cas9 targeted mutagenesis in sand flies: embryo collection, injection, insect rearing, and identification as well as selection of mutations of interest.

Sand flies are the natural vectors for Leishmania species, protozoan parasites producing a broad spectrum of symptoms ranging from cutaneous lesions to ...

CRISPR/Cas9 Editing of the C. elegans rbm-3.2 Gene using the dpy-10 Co-CRISPR Screening Marker and Assembled Ribonucleoprotein Complexes.

The bacterial Clustered Regularly Interspaced Short Palindromic Repeat (CRISPR)/Streptococcus pyogenes CRISPR-associated protein (Cas) system has been harnessed by researchers to study important biologically relevant problems. The unparalleled power of the CRISPR/Cas genome editing method allows researchers to precisely edit any locus of their choosing, thereby facilitating an increased understanding of gene function. Several methods for editing the C. elegans genome by CRISPR/Cas9 have been described previously. Here, we discuss and demonstrate a method which utilizes in vitro assembled ribonucleoprotein complexes and the dpy-10 co-CRISPR marker for screening. Specifically, in this article, we go through the step-by-step process of introducing premature stop codons into the C. elegans rbm-3.2 gene by homology-directed repair using this method of CRISPR/Cas9 editing. This relatively simple editing method can be used to study the function of any gene of interest and allows for the generation of homozygous-edited C. elegans by CRISPR/Cas9 editing&#160;in less than two weeks.

CRISPR/Cas9 Editing of the C. elegans rbm-3.2 Gene using the dpy-10 Co-CRISPR Screening Marker and Assembled Ribonucleoprotein Complexes.

The goal of this protocol is to enable seamless CRISPR/Cas9 editing of the C. elegans genome using assembled ribonucleoprotein complexes and the dpy-10 co-CRISPR marker for screening. This protocol can be used to make a variety of genetic modifications in C. elegans including insertions, deletions, gene replacements and codon substitutions.

The bacterial Clustered Regularly Interspaced Short Palindromic Repeat (CRISPR)/Streptococcus pyogenes CRISPR-associated protein (Cas) system has been ...

Genome Editing in the Yellow Fever Mosquito Aedes aegypti using CRISPR-Cas9

The emergence of the clustered, regularly interspersed, short palindromic repeats (CRISPR)-Cas9 technology has revolutionized the genetic engineering field and opened the doors for precise genome editing in multiple species, including non-model organisms. In the mosquito Aedes aegypti, loss-of-function mutations and DNA insertions have been accomplished with this technology. Here, we describe a detailed protocol for genome editing through embryonic microinjection in the mosquito A. aegypti using the CRISPR-Cas9 technology, focusing on both the generation of gene knockout and knockin lines. In this protocol, quartz needles are filled with a mixture of guide RNA, recombinant Cas9, and a plasmid containing a DNA cassette encoding a gene for a fluorescent marker, if gene knockin is desired. Embryos at the preblastoderm stage are lined up onto a strip of double-sided sticky tape placed onto a coverslip, which is subsequently mounted onto a glass slide. With the help of a microinjector, the needles are inserted gently into the posterior end of the embryos and a small volume of the CRISPR mixture is dispensed. When the embryos are hatched, the larvae are checked under the fluorescent scope, and the pupae are sex-sorted and separated in different cages. Once the adults emerge, these are reciprocally crossed with wild-type individuals, blood-fed, and placed for egg laying. Once these eggs are hatched, the fluorescent larvae collected represent individuals with stable insertion of the DNA cassette into their genome. These larvae are then grown to the adult stage, outcrossed to wild-type individuals, and then further assessed through molecular techniques to confirm that the exact sequence of the DNA cassette is present at the desired site of the mosquito genome. Homozygous lines can also be obtained by following the provided pipeline of crossing schema and molecular screening of the mutations.

Here, we describe a detailed protocol for genome editing through embryonic microinjection in the mosquito A. aegypti using the CRISPR-Cas9 technology.

The emergence of the clustered, regularly interspersed, short palindromic repeats (CRISPR)-Cas9 technology has revolutionized the genetic engineering ...

Preparing and Injecting Embryos of Culex Mosquitoes to Generate Null Mutations using CRISPR/Cas9

Culex mosquitoes are the major vectors of several diseases that negatively impact human and animal health including West Nile virus and diseases caused by filarial nematodes such as canine heartworm and elephantasis. Recently, CRISPR/Cas9 genome editing has been used to induce site-directed mutations by injecting a Cas9 protein that has been complexed with a guide RNA (gRNA) into freshly laid embryos of several insect species, including mosquitoes that belong to the genera Anopheles and Aedes. Manipulating and injecting Culex mosquitoes is slightly more difficult as these mosquitoes lay their eggs upright in rafts rather than individually like other species of mosquitoes. Here we describe how to design gRNAs, complex them with Cas9 protein, induce female mosquitoes of Culex pipiens to lay eggs, and how to prepare and inject newly laid embryos for microinjection with Cas9/gRNA. We also describe how to rear and screen injected mosquitoes for the desired mutation. The representative results demonstrate that this technique can be used to induce site-directed mutations in the genome of Culex mosquitoes and, with slight modifications, can be used to generate null-mutants in other mosquito species as well.&#160;

Preparing and Injecting Embryos of Culex Mosquitoes to Generate Null Mutations using CRISPR/Cas9

CRISPR/Cas9 is increasingly used to characterize gene function in non-model organisms. This protocol describes how to generate knock-out lines of Culex pipiens, from preparing injection mixes, to obtaining and injecting mosquito embryos, as well as how to rear, cross, and screen injected mosquitoes and their progeny for desired mutations.

Culex mosquitoes are the major vectors of several diseases that negatively impact human and animal health including West Nile virus and diseases caused by ...

CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

Summary

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Chapters in this video

QTL Mapping and CRISPR/Cas9 Editing to Identify a Drug Resistance Gene in Toxoplasma gondii

Selection-dependent and Independent Generation of CRISPR/Cas9-mediated Gene Knockouts in Mammalian Cells

Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms

Introducing a Gene Knockout Directly Into the Amastigote Stage of Trypanosoma cruzi Using the CRISPR/Cas9 System

Generation of Defined Genomic Modifications Using CRISPR-CAS9 in Human Pluripotent Stem Cells

Introducing Point Mutations into Human Pluripotent Stem Cells Using Seamless Genome Editing

Sand Fly (Phlebotomus papatasi) Embryo Microinjection for CRISPR/Cas9 Mutagenesis

CRISPR/Cas9 Editing of the C. elegans rbm-3.2 Gene using the dpy-10 Co-CRISPR Screening Marker and Assembled Ribonucleoprotein Complexes.

Genome Editing in the Yellow Fever Mosquito Aedes aegypti using CRISPR-Cas9

Preparing and Injecting Embryos of Culex Mosquitoes to Generate Null Mutations using CRISPR/Cas9