This work was supported in part by JSPS KAKENHI Grant Number 18K15141 to Y.T.

NOTE: An overview of the entire experimental flow is depicted in Figure 1.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59962/59962fig01.jpg" /> 
Figure 1: Overview of the knockout experiment using EA.&#160;Tissue culture-derived trypomastigotes are harvested and differentiated into EA. gRNA is transfected into Cas9-expressing amastigotes by electroporation, and growth phenotype of the knockout amastigote is evaluated either by axenic replication or by intracellular replication after host cell invasion. <a href="https://www.jove.com/files/ftp_upload/59962/59962fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
1. Parasite Culture Preparations
<ol>
	<li>Tulahuen strain of Trypanosoma cruzi&#160;is used throughout this report. Maintain epimastigotes of T. cruzi in LIT medium (10% FCS, see Supplemental Table 1). Securely close the cap and keep the culture flask at 28&#160;&#176;C.</li>
	<li>Generate a transgenic strain of T. cruzi that harbors the Cas9 endonuclease. Examples of the expression plasmids that contain Cas9 coding sequence and G418 (neor) selection marker can be found in Lander et al.4 and Peng et al.5
	<ol>
		<li>Transfect the above plasmid into epimastigote by electroporation, using the kit listed in the Table of Materials. For 1 cuvette, spin down 2 x 107 cells, discard the supernatant, and resuspend with 100 &#181;L of electroporation buffer containing the provided supplement solution. 
		NOTE: The electroporation buffer can be substituted with EM buffer24 (3:1 mixture of cytomix25 and phosphate-sucrose buffer).</li>
		<li>Add 20-40 &#181;g of plasmid and transfer the mixture into a 2 mm gap electroporation cuvette. Apply the pulse with an electroporation device (see Table of Materials), using the X-14 program26.</li>
		<li>Transfer the cuvette contents into a T-25 flask containing 5 mL of LIT medium (10% FCS). Incubate the flask at 28&#160;&#176;C for 24 h.</li>
		<li>Add G418 to a final concentration of 250 &#181;g/mL and continue incubation at 28 &#730;C. It takes about 1 week to kill off the non-transfected epimastigotes. Once the G418-resistant population starts to recover, passage the culture once or twice a week to avoid saturation and to dilute out the dead cells. Establishment of a stable cell line usually takes total of 4 weeks. 
		NOTE: If constitutive expression of Cas9 is a burden for the cell5, make the expression specific to the amastigote stage by conjugating the 3&#8217;-UTR of amastin gene immediately downstream of the Cas9 open reading frame27. The detailed description of the amastigote-specific Cas9 expression plasmid can be found in Takagi et al.22</li>
	</ol>
	</li>
	<li>Establish a host-parasite co-culture of Cas9-expressing T. cruzi and mammalian host cells. In this report, use 3T3-Swiss Albino fibroblast cell as a host.
	<ol>
		<li>Differentiate T. cruzi epimastigotes into metacyclic trypomastigotes. Determine the cell density of the epimastigote culture using a hemocytometer and collect 5 x 107 cells by centrifugation for 15 min at 2,100 x g. Discard the supernatant and resuspend the cells with 10 mL of RPMI medium. Incubate the parasite at 28 &#176;C for 1 week28.</li>
		<li>Collect metacyclic trypomastigotes in RPMI medium. Carefully tilt the flask and pipet out the solution without disturbing the parasites adhered to the bottom surface. Transfer the medium to a conical tube and centrifuge for 15 min at 2,100 x g. Discard the supernatant and resuspend the parasite with 5 mL of DMEM (10% FCS). 
		NOTE: The parasite population is a mixture of metacyclic trypomastigotes, epimastigotes, and some intermediate forms. Although not necessary, you can isolate trypomastigote by DEAE ion exchange chromatography29. Alternatively, epimastigotes can be eliminated by incubating the parasites with active serum to subject them to complement lysis30.</li>
		<li>Seed 3T3-Swiss Albino fibroblast cell to 60-70% confluency, or about 1.7-2.0 x 106 cells in 5 mL of DMEM (10% FCS) in T-25 culture flask. Remove growth medium and apply the parasite mixture from step 1.3.2. Incubate for 24 h at 37 &#730;C under 5% CO2 in a humidified incubator to establish infection.</li>
		<li>Remove the parasites remaining outside of the host 3T3 cells by washing the co-culture with DMEM (10% FCS) twice.</li>
		<li>Once the co-culture is saturated, passage amastigote-infected host cells by trypsinization. Aspirate the medium and rinse the culture once with PBS. Apply 1 mL of 0.05% trypsin solution to cover the entire culture surface, and incubate for few minutes at room temperature until the attached host cells become loose. Detach the cells from the flask surface by flushing 3 mL of DMEM (10% FCS) over the cells.</li>
		<li>Transfer detached host cells into a conical tube, and centrifuge for 3 min at 300 x g. Aspirate the supernatant and resuspend the cells with 3 mL of fresh DMEM (10% FCS). This step helps to eliminate remaining epimastigotes. Transfer all into a clean T-75 flask containing 9 mL of DMEM (10% FCS). Continue culturing until trypomastigote is released into culture supernatant.</li>
		<li>Maintain the co-culture by passaging with trypsinization twice a week. Once 70-80% of the host population become infected, regularly add uninfected host 3T3 cells at the ratio of 5:1 (carryover:fresh), in order to avoid culture deterioration. Trypomastigote egresses continuously if the balance between host cells and T. cruzi is properly maintained.</li>
	</ol>
	</li>
</ol>
2. Differentiation of Trypomastigotes into EA
<ol>
	<li>On the day before this experiment, remove the growth medium of host-parasite co-culture and add fresh DMEM (10% FCS) to wash away EA and trypomastigotes that had already been released from the host cells in previous days. Regular experiments require at least two T-75 flasks of confluent co-culture.</li>
	<li>Collect culture supernatant into a conical tube to harvest freshly emerged trypomastigotes. Check the sample under a microscope (10x or 20x objective lens) for the quality. If there is host cell debris, briefly centrifuge the sample and transfer the supernatant into a new tube. If there are significant number of EAs, isolate trypomastigotes by the following swim-out procedure.
	<ol>
		<li>Spin down the mixture of trypomastigotes and amastigotes for 15 min at 2,100 x g. Discard most of the supernatant, leaving 0.5-1.0 mL of medium in the tube. 
		NOTE: Reducing the volume is optional, but it makes the following step easier.</li>
		<li>Incubate the pellet at 37 &#730;C for 1-2 h, allowing active trypomastigotes to swim out of the pellet (Figure 2).</li>
		<li>Transfer the supernatant containing trypomastigotes to a 1.5 mL microcentrifuge tube.</li>
	</ol>
	</li>
	<li>Centrifuge the conical tube for 15 min at 2,100 x g to collect trypomastigotes. If the swim-out procedure was performed above, centrifuge the 1.5 mL tube for 2 min at 4,000 x g to pellet the trypomastigote. Discard the supernatant.</li>
	<li>Resuspend the pellet with 5 mL of DMEM buffered with 20 mM MES (pH 5.0), supplemented with 0.4% BSA19. Transfer the parasite to a T-25 culture flask. Leave the cap loose. The cell density must be around or below 1 x 107 cells/mL, since oversaturation increases the chance of cell death. 
	NOTE: The color of the DMEM must be yellow, not orange. If the original DMEM had high buffering capacity, 20 mM MES (pH 5.0) is not enough to lower the pH. The pH of the medium must be adjusted by addition of HCl in that case. The acidic medium can be kept at 4 &#730;C, but no longer than 1 month.</li>
	<li>Incubate the culture flask at 37 &#176;C under 5% CO2 in a humidified incubator. Around 95% of parasites differentiate into amastigotes after 24 h.</li>
</ol>
3. Electroporation of EA
<ol>
	<li>Prepare gRNA for electroporation. This can be done by in vitro transcription, or simply by purchasing synthetic RNA oligonucleotides from a manufacturer. In this report, crRNA and tracrRNA from Integrated DNA Technologies, Inc. are used.</li>
	<li>Centrifuge the culture of EAs for 15 min at 2,100 x g. Discard supernatant.</li>
	<li>Resuspend the pellet with electroporation buffer containing provided supplement solution to the final cell density of 1 x 108 cells/mL. 
	NOTE: EM buffer causes more cell deaths compared to the electroporation buffer listed in Table of Materials; thus, it is not recommended for amastigote transfection (Supplemental Figure 1).</li>
	<li>Aliquot 100 &#181;L of resuspended parasites (1 x 107 cells) into 1.5 mL microcentrifuge tubes. Add 5-10 &#181;g of gRNA and gently mix by pipetting.</li>
	<li>Transfer the mixture to a 2 mm gap electroporation cuvette. Apply the pulse with the electroporation device, using X-14 program.</li>
	<li>Transfer the cuvette contents into a T-25 flask containing 5 mL of pre-warmed LIT medium (10% FCS). Leave the cap loose and incubate the flask at 37 &#176;C under 5% CO2.</li>
	<li>Monitor the cell growth either by continuation of axenic culturing (section 4) or as intracellular amastigotes after host cell infection (section 5).</li>
</ol>
4. Monitoring the Growth of Knockout Cells as Axenic Amastigotes
<ol>
	<li>EAs settle at the bottom of the culture medium, so gently shake the flask to resuspend them into the solution. Washing the flask surface by pipetting helps, as some cells are adhered to the flask.</li>
	<li>Mix 1 &#181;L of propidium iodide (PI) solution (20 &#181;g/mL) with 20 &#181;L of amastigote culture. 
	NOTE: Do not leave the culture flask outside of incubator for longer than necessary. Temperature is one of factors that enables axenic proliferation22.</li>
	<li>Apply the sample onto hemocytometer and observe under a fluorescence microscope. PI is permeant to damaged cell membrane but is excluded from live cells. Count the number of viable amastigotes that are not stained by PI (ex/em 570 nm/602 nm).</li>
</ol>
5. Monitoring the Growth of Knockout Cells as Intracellular Amastigotes
<ol>
	<li>Seed host 3T3 cells in a 12-well plate with DMEM (10% FCS). Adjust the cell density to 70-80% confluency, or about 3 x 105 cells per well. 
	NOTE: Since amastigotes are not motile, covering most of the growth surface by the host cells improves infection efficiency.</li>
	<li>Collect knockout amastigotes from step 3.6 by centrifugation one day after electroporation. Discard the supernatant, and resuspend the parasite with 2 mL of DMEM (10% FCS).</li>
	<li>Remove medium from the host cell culture and apply resuspended amastigotes. Multiplicity of infection should be 20 or higher. Incubate the plate at 37 &#176;C under 5% CO2 for 2 days to allow amastigotes to establish infection. 
	NOTE: Infection period can be 1 day, depending on the purpose.</li>
	<li>Wash away EAs remained outside of the host cells twice with DMEM (10% FCS).</li>
	<li>Add fresh DMEM (10% FCS) to the host-parasite co-culture and continue the incubation at 37 &#176;C for additional 2 days.</li>
	<li>To evaluate infection efficiency, visualize nuclei of the host cells and intracellular amastigote. 
	NOTE: Nuclei tend to be overlapped in saturated co-culture. Re-plating the cells at lower cell density (such as 1:5 dilution) prior to fixation and staining helps to count nuclei more easily.
	<ol>
		<li>Remove culture medium and apply 1 mL of 10% formalin solution in PBS to fix the cells. Incubate for 10 min at room temperature.</li>
		<li>Replace the formalin solution with 1 mL of PBS containing 1 &#181;g/mL Hoechst 33342 and 0.1% Triton X-100. Incubate for 5 min at room temperature.</li>
		<li>Remove the Hoechst solution and rinse the cells once with PBS. Add 1 mL of fresh PBS.</li>
	</ol>
	</li>
	<li>Observe under fluorescence microscope and identify host cell nuclei that are associated with smaller parasite nuclei (Figure 4). Host cells that contain more than 2 amastigotes should be considered as infected, not to include nonproductive initial infection or unwashed EAs.</li>
</ol>

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G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparative+studies+of+the+isolation+of+metacyclic+trypomastigotes+of+Trypanosoma+cruzi+by+DEAE+ion+exchange+chromatography.">Comparative studies of the isolation of metacyclic trypomastigotes of Trypanosoma cruzi by DEAE ion exchange chromatography.</a> Zhonghua Minguo wei sheng wu ji mian yi xue za zhi (Chinese Journal of Microbiology and Immunology). 17 (3), 146-152 (1984).</li><li>Nogueira, N., Bianco, C., Cohn, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Studies+on+the+selective+lysis+and+purification+of+Trypanosoma+cruzi.">Studies on the selective lysis and purification of Trypanosoma cruzi.</a> The Journal of Experimental Medicine. 142 (1), 224-229 (1975).</li><li>Minning, T. A., Weatherly, D. B., Atwood, J., Orlando, R., Tarleton, R. L., Tarleton, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+steady-state+transcriptome+of+the+four+major+life-cycle+stages+of+Trypanosoma+cruzi.">The steady-state transcriptome of the four major life-cycle stages of Trypanosoma cruzi.</a> BMC Genomics. 10, 370 (2009).</li><li>Rico, E., Jeacock, L., Kov&#225;&#345;ov&#225;, J., Horn, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inducible+high-efficiency+CRISPR-Cas9-targeted+gene+editing+and+precision+base+editing+in+African+trypanosomes.">Inducible high-efficiency CRISPR-Cas9-targeted gene editing and precision base editing in African trypanosomes.</a> Scientific Reports. 8 (1), 7960 (2018).</li><li>Fu, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-frequency+off-target+mutagenesis+induced+by+CRISPR-Cas+nucleases+in+human+cells.">High-frequency off-target mutagenesis induced by CRISPR-Cas nucleases in human cells.</a> Nature Biotechnology. 31 (9), 822-826 (2013).</li><li>Kim, S., Kim, D., Cho, S. W., Kim, J., Kim, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Highly+efficient+RNA-guided+genome+editing+in+human+cells+via+delivery+of+purified+Cas9+ribonucleoproteins.">Highly efficient RNA-guided genome editing in human cells via delivery of purified Cas9 ribonucleoproteins.</a> Genome Research. 24 (6), 1012-1019 (2014).</li><li>Chiurillo, M. A., Lander, N., Bertolini, M. S., Storey, M., Vercesi, A. E., Docampo, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Different+roles+of+mitochondrial+calcium+uniporter+complex+subunits+in+growth+and+infectivity+of+Trypanosoma+cruzi.">Different roles of mitochondrial calcium uniporter complex subunits in growth and infectivity of Trypanosoma cruzi.</a> mBio. 8 (3), e00574-e00617 (2017).</li></ol>

The authors have no conflict of interest to disclose.

We demonstrated that the axenic culture of T. cruzi amastigotes can be utilized in CRISPR/Cas9-mediated gene knockout, by electroporating gRNA directly into Cas9-expressing EA. This way, the essentiality of the target gene specifically in amastigote stage can be evaluated without going through other developmental stages.
Another beneficial aspect of amastigote transfection is the convenience in testing for a large number of target genes. Once the co-culture of Cas9-expressing T. c...

Trypanosoma cruzi is the causative agent of Chagas&#8217; disease, which is prevalent mainly in Latin America1. T. cruzi has distinctive life cycle stages as it travels between an insect vector and a mammalian host2. T. cruzi replicates as an epimastigote in the midgut of a blood-sucking triatomine bug and differentiates into an infectious metacyclic trypomastigote in its hindgut before being deposited on a human or animal host. Once the trypomastigote gets into the host body through the bite site or through a mucous membrane, the parasite invades a host cell and transforms into a flagella-less round form called an amastigote. The amastigote replicates within the host cell and eventually differentiates into trypomastigote, which bursts out of the host cell and enters the blood stream to infect another host cell.
Since currently available chemotherapeutic agents, benznidazole and nifurtimox, cause adverse side effects and are ineffective in the chronic phase of the disease3, it is of a great interest to identify novel drug targets against T. cruzi. In recent years, the CRISPR/Cas9 system has become a powerful tool to effectively perform gene knockout in T. cruzi, either by transfection of separate or single plasmid(s) containing gRNA and Cas94, by stable expression of Cas9 and subsequent introduction of gRNA5,6,7 or transcription template of gRNA8, or by electroporation of the pre-formed gRNA/Cas9 RNP complex7,9. This technological advancement is highly anticipated to accelerate the drug target research in Chagas&#8217; disease.
To proceed with the drug development, it is crucial to validate the essentiality of the target gene or efficacy of drug candidate compounds in the amastigote of T. cruzi, as it is the replication stage of the parasite in the mammalian host. However, this is a challenging task, because amastigotes cannot be directly manipulated due to the presence of an obstructive host cell. In Leishmania, a closely related protozoan parasite to T. cruzi, an axenic amastigote culturing method was developed and has been utilized in drug screening assays10,11,12,13. Although there are some discrepancies in susceptibility to compounds between axenic amastigotes and intracellular amastigotes14, the ability to maintain the axenic culture nonetheless provides valuable experimental tools to study the basic biology of the clinically relevant stage of Leishmania15,16. In the case of T. cruzi, literatures regarding the presence of naturally occurring extracellular amastigotes (EA)17 and in vitro production of EA17,18,19 date back to decades ago. In addition, EA is known to have an infectious capability20, albeit less than that of trypomastigote, and the mechanism of amastigote host invasion has been elucidated in recent years (reviewed by Bonfim-Melo et al.21). However, unlike Leishmania, EA had not been utilized as an experimental tool in T. cruzi, primarily because EA had been regarded as an obligate intracellular parasite, and thus had not been considered as &#8220;replicative form&#8221; in a practical sense.
Recently, our group proposed to utilize EA of T. cruzi as a temporal axenic culture22. Amastigotes of T. cruzi Tulahuen strain can replicate free of host cells in LIT medium at 37 &#176;C for up to 10 days without major deterioration or loss of amastigote-like properties. During the host-free growth period, EA was successfully utilized for exogenous gene expression by conventional electroporation, drug titration assay with trypanocidal compounds, and CRISPR/Cas9-mediated knockout followed by growth phenotype monitoring. In this report, we describe the detailed protocol to produce in vitro derived EA and to utilize the axenic amastigote in knockout experiments.

<table>
	<tbody>
		<tr>
			<td>20% formalin solution</td>
			<td>FUJIFILM Wako Pure Chemical</td>
			<td>068-03863</td>
			<td>fixing cells</td>
		</tr>
		<tr>
			<td>25 cm2 double seal cap culture flask</td>
			<td>AGC Techno Glass</td>
			<td>3100-025</td>
			<td></td>
		</tr>
		<tr>
			<td>75 cm2 double seal cap culture flask</td>
			<td>AGC Techno Glass</td>
			<td>3110-075</td>
			<td></td>
		</tr>
		<tr>
			<td>All-in One Fluorescence Microscope</td>
			<td>Keyence</td>
			<td>BZ-X710</td>
			<td></td>
		</tr>
		<tr>
			<td>Alt-R CRISPR-Cas9 crRNA (for Control)</td>
			<td>IDT</td>
			<td>custom made</td>
			<td>target sequence = GGACGGCACCTTCATCTACAAGG</td>
		</tr>
		<tr>
			<td>Alt-R CRISPR-Cas9 crRNA (for TcCGM1)</td>
			<td>IDT</td>
			<td>custom made</td>
			<td>target sequence = TAGCCGCGATGGAGAGTTTATGG</td>
		</tr>
		<tr>
			<td>Alt-R CRISPR-Cas9 crRNA (for TcPAR1)</td>
			<td>IDT</td>
			<td>custom made</td>
			<td>target sequence = CGTGGAGAACGCCATTGCCACGG</td>
		</tr>
		<tr>
			<td>Alt-R CRISPR-Cas9 tracrRNA</td>
			<td>IDT</td>
			<td>1072532</td>
			<td>to anneal with crRNA</td>
		</tr>
		<tr>
			<td>Amaxa Nucleofector device</td>
			<td>LONZA</td>
			<td>AAN-1001</td>
			<td>electroporation</td>
		</tr>
		<tr>
			<td>Basic Parasite Nucleofector Kit 2</td>
			<td>LONZA</td>
			<td>VMI-1021</td>
			<td>electroporation</td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Sigma-Aldrich</td>
			<td>A3294</td>
			<td>component of the medium for in-vitro amastigogenesis</td>
		</tr>
		<tr>
			<td>Burker-Turk disposable hemocytometer</td>
			<td>Watson</td>
			<td>177-212C</td>
			<td>cell counting</td>
		</tr>
		<tr>
			<td>Coster 12-well Clear TC-Treated Multiple Well Plates</td>
			<td>Corning</td>
			<td>3513</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>FUJIFILM Wako Pure Chemical</td>
			<td>044-29765</td>
			<td>culture medium</td>
		</tr>
		<tr>
			<td>Fetal bovine serum, Defined</td>
			<td>Hyclone</td>
			<td>SH30070.03</td>
			<td>heat-inactivate before use</td>
		</tr>
		<tr>
			<td>G-418 Sulfate Solution</td>
			<td>FUJIFILM Wako Pure Chemical</td>
			<td>077-06433</td>
			<td>selection of transformant</td>
		</tr>
		<tr>
			<td>Hemin chloride</td>
			<td>Sigma-Aldrich</td>
			<td>H-5533</td>
			<td>component of LIT medium</td>
		</tr>
		<tr>
			<td>Hoechst 33342</td>
			<td>Thermo Fisher Scientific</td>
			<td>H3570</td>
			<td>staining of nuclei</td>
		</tr>
		<tr>
			<td>Liver infusion broth, Difco</td>
			<td>Becton Dickinson</td>
			<td>226920</td>
			<td>component of LIT medium</td>
		</tr>
		<tr>
			<td>MES</td>
			<td>FUJIFILM Wako Pure Chemical</td>
			<td>349-01623</td>
			<td>component of the medium for in-vitro amastigogenesis</td>
		</tr>
		<tr>
			<td>PBS (&#8211;)</td>
			<td>FUJIFILM Wako Pure Chemical</td>
			<td>166-23555</td>
			<td></td>
		</tr>
		<tr>
			<td>Propidium Iodide</td>
			<td>Sigma-Aldrich</td>
			<td>P4864-10ML</td>
			<td>staining of dead cells</td>
		</tr>
		<tr>
			<td>RPMI 1646</td>
			<td>Sigma-Aldrich</td>
			<td>R8758</td>
			<td>medium for metacyclogenesis</td>
		</tr>
	</tbody>
</table>

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.

Isolation of trypomastigotes by the swim-out procedure
To harvest fresh trypomastigotes from contaminating old EAs by swim-out procedure, cell pellets need to be incubated at least for 1 h. Incubating the pellets for more than 2 h does not significantly increase the number of trypomastigotes swimming in the solution (Figure 2B). In this particular experiment, the percentage of trypomastigote in the initial mixture was 38%, and the percentage after...

Watch this Scientific Journal Video about Introducing a Gene Knockout Directly Into the Amastigote Stage of Trypanosoma cruzi Using the CRISPR/Cas9 System at JoVE.com

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.

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 ...

introducing-gene-knockout-directly-into-amastigote-stage-trypanosoma

Research

JoVE Journal

Genetics

8.9K Views.  National Institute of Advanced Industrial Science and Technology (AIST). 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.

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

Using a Fluorescent PCR-capillary Gel Electrophoresis Technique to Genotype CRISPR/Cas9-mediated Knockout Mutants in a High-throughput Format

The development of programmable genome-editing tools has facilitated the use of reverse genetics to understand the roles specific genomic sequences play in the functioning of cells and whole organisms. This cause has been tremendously aided by the recent introduction of the CRISPR/Cas9 system-a versatile tool that allows researchers to manipulate the genome and transcriptome in order to, among other things, knock out, knock down, or knock in genes in a targeted manner. For the purpose of knocking out a gene, CRISPR/Cas9-mediated double-strand breaks recruit the non-homologous end-joining DNA repair pathway to introduce the frameshift-causing insertion or deletion of nucleotides at the break site. However, an individual guide RNA may cause undesirable off-target effects, and to rule these out, the use of multiple guide RNAs is necessary. This multiplicity of targets also means that a high-volume screening of clones is required, which in turn begs the use of an efficient high-throughput technique to genotype the knockout clones. Current genotyping techniques either suffer from inherent limitations or incur high cost, hence rendering them unsuitable for high-throughput purposes. Here, we detail the protocol for using fluorescent PCR, which uses genomic DNA from crude cell lysate as a template, and then resolving the PCR fragments via capillary gel electrophoresis. This technique is accurate enough to differentiate one base-pair difference between fragments and hence is adequate in indicating the presence or absence of a frameshift in the coding sequence of the targeted gene. This precise knowledge effectively precludes the need for a confirmatory sequencing step and allows users to save time and cost in the process. Moreover, this technique has proven to be versatile in genotyping various mammalian cells of various tissue origins targeted by guide RNAs against numerous genes, as shown here and elsewhere.

The genotyping technique described here, which couples fluorescent polymerase chain reaction (PCR) to capillary gel electrophoresis, allows for high-throughput genotyping of nuclease-mediated knockout clones. It circumvents limitations faced by other genotyping techniques and is more cost effective than sequencing methods.

The development of programmable genome-editing tools has facilitated the use of reverse genetics to understand the roles specific genomic sequences play ...

A Standard Methodology to Examine On-site Mutagenicity As a Function of Point Mutation Repair Catalyzed by CRISPR/Cas9 and SsODN in Human Cells

Combinatorial gene editing using CRISPR/Cas9 and single-stranded oligonucleotides is an effective strategy for the correction of single-base point mutations, which often are responsible for a variety of human inherited disorders. Using a well-established cell-based model system, the point mutation of a single-copy mutant eGFP gene integrated into HCT116 cells has been repaired using this combinatorial approach. The analysis of corrected and uncorrected cells reveals both the precision of gene editing and the development of genetic lesions, when indels are created in uncorrected cells in the DNA sequence surrounding the target site. Here, the specific methodology used to analyze this combinatorial approach to the gene editing of a point mutation, coupled with a detailed experimental strategy to measuring indel formation at the target site, is outlined. This protocol outlines a foundational approach and workflow for investigations aimed at developing CRISPR/Cas9-based gene editing for human therapy. The conclusion of this work is that on-site mutagenesis takes place as a result of CRISPR/Cas9 activity during the process of point mutation repair. This work puts in place a standardized methodology to identify the degree of mutagenesis, which should be an important and critical aspect of any approach destined for clinical implementation.

This protocol outlines the workflow of a CRISPR/Cas9-based gene editing system for the repair of point mutations in mammalian cells. Here, we use a combinatorial approach to gene editing with a detailed follow-on experimental strategy for measuring indel formation at the target site&#8212;in essence, analyzing onsite mutagenesis.

Combinatorial gene editing using CRISPR/Cas9 and single-stranded oligonucleotides is an effective strategy for the correction of single-base point ...

Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing

The complete genome of the channel catfish, Ictalurus punctatus, has been sequenced, leading to greater opportunities for studying channel catfish gene function. Gene knockout has been used to study these gene functions in vivo. The clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 (CRISPR/Cas9) system is a powerful tool used to edit genomic DNA sequences to alter gene function. While the traditional approach has been to introduce CRISPR/Cas9 mRNA into the single cell embryos through microinjection, this can be a slow and inefficient process in catfish. Here, a detailed protocol for microinjection of channel catfish embryos with CRISPR/Cas9 protein is described. Briefly, eggs and sperm were collected and then artificial fertilization performed. Fertilized eggs were transferred to a Petri dish containing Holtfreter's solution. Injection volume was calibrated and then guide RNAs/Cas9 targeting the toll/interleukin 1 receptor domain-containing adapter molecule (TICAM 1) gene and rhamnose binding lectin (RBL) gene were microinjected into the yolk of one-cell embryos. The gene knockout was successful as indels were confirmed by DNA sequencing. The predicted protein sequence alterations due to these mutations included frameshift and truncated protein due to premature stop codons.

Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing

A simple and efficient microinjection protocol for gene editing in channel catfish embryos using the CRISPR/Cas9 system is presented. In this protocol, guide RNAs and Cas9 protein were microinjected into the yolk of one-cell embryos. This protocol has been validated by knocking out two channel catfish immune-related genes.

The complete genome of the channel catfish, Ictalurus punctatus, has been sequenced, leading to greater opportunities for studying channel catfish gene ...

A Protocol for the Production of Integrase-deficient Lentiviral Vectors for CRISPR/Cas9-mediated Gene Knockout in Dividing Cells

Lentiviral vectors are an ideal choice for delivering gene-editing components to cells due to their capacity for stably transducing a broad range of cells and mediating high levels of gene expression. However, their ability to integrate into the host cell genome enhances the risk of insertional mutagenicity and thus raises safety concerns and limits their usage in clinical settings. Further, the persistent expression of gene-editing components delivered by these integration-competent lentiviral vectors (ICLVs) increases the probability of promiscuous gene targeting. As an alternative, a new generation of integrase-deficient lentiviral vectors (IDLVs) has been developed that addresses many of these concerns. Here the production protocol of a new and improved IDLV platform for CRISPR-mediated gene editing and list the steps involved in the purification and concentration of such vectors is described and their transduction and gene-editing efficiency using HEK-293T cells was demonstrated. This protocol is easily scalable and can be used to generate high titer IDLVs that are capable of transducing cells in vitro and in vivo. Moreover, this protocol can be easily adapted for the production of ICLVs.

We describe the production strategy of integrase-deficient lentiviral vectors (IDLVs) as vehicles for delivering CRISPR/Cas9 to cells. With an ability to mediate quick and robust gene editing in cells, IDLVs present a safer and equally effective vector platform for gene delivery compared to integrase-competent vectors.

Lentiviral vectors are an ideal choice for delivering gene-editing components to cells due to their capacity for stably transducing a broad range of cells ...

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development of a customizable platform to rapidly generate gene modifications in a wide variety of organisms, including zebrafish. CRISPR-based genome editing uses a single guide RNA (sgRNA) to target a CRISPR-associated (Cas) endonuclease to a genomic DNA (gDNA) target of interest, where the Cas endonuclease generates a double-strand break (DSB). Repair of DSBs by error-prone mechanisms lead to insertions and/or deletions (indels). This can cause frameshift mutations that often introduce a premature stop codon within the coding sequence, thus creating a protein-null allele. CRISPR-based genome engineering requires only a few molecular components and is easily introduced into zebrafish embryos by microinjection. This protocol describes the methods used to generate CRISPR reagents for zebrafish microinjection and to identify fish exhibiting germline transmission of CRISPR-modified genes. These methods include in vitro transcription of sgRNAs, microinjection of CRISPR reagents, identification of indels induced at the target site using a PCR-based method called a heteroduplex mobility assay (HMA), and characterization of the indels using both a low throughput and a powerful next-generation sequencing (NGS)-based approach that can analyze multiple PCR products collected from heterozygous fish. This protocol is streamlined to minimize both the number of fish required and the types of equipment needed to perform the analyses. Furthermore, this protocol is designed to be amenable for use by laboratory personal of all levels of experience including undergraduates, enabling this powerful tool to be economically employed by any research group interested in performing CRISPR-based genomic modification in zebrafish.

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Targeted genome editing in the model system Danio rerio (zebrafish) has been greatly facilitated by the emergence of CRISPR-based approaches. Herein, we describe a streamlined, robust protocol for generation and identification of CRISPR-derived nonsense alleles that incorporates the heteroduplex mobility assay and identification of mutations using next-generation sequencing.

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development ...

Highly Efficient Gene Disruption of Murine and Human Hematopoietic Progenitor Cells by CRISPR/Cas9

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. Complete gene ablation in HSCs required the generation of knockout mice from which HSCs could be isolated, and gene ablation in primary human HSCs was not possible. Viral transduction could be used for knock-down approaches, but these suffered from variable efficacy. In general, genetic manipulation of human and mouse hematopoietic cells was hampered by low efficiencies and extensive time and cost commitments. Recently, CRISPR/Cas9 has dramatically expanded the ability to engineer the DNA of mammalian cells. However, the application of CRISPR/Cas9 to hematopoietic cells has been challenging, mainly due to their low transfection efficiencies, the toxicity of plasmid-based approaches and the slow turnaround time of virus-based protocols.
A rapid method to perform CRISPR/Cas9-mediated gene editing in murine and human hematopoietic stem and progenitor cells with knockout efficiencies of up to 90% is provided in this article. This approach utilizes a ribonucleoprotein (RNP) delivery strategy with a streamlined three-day workflow. The use of Cas9-sgRNA RNP allows for a hit-and-run approach, introducing no exogenous DNA sequences in the genome of edited cells and reducing off-target effects. The RNP-based method is fast and straightforward: it does not require cloning of sgRNAs, virus preparation or specific sgRNA chemical modification. With this protocol, scientists should be able to successfully generate knockouts of a gene of interest in primary hematopoietic cells within a week, including downtimes for oligonucleotide synthesis. This approach will allow a much broader group of users to adapt this protocol for their needs.

A protocol for fast CRISPR/Cas9-mediated gene disruption in mouse and human primary hematopoietic cells is described in this article. Cas9-sgRNA ribonucleoproteins are introduced via electroporation with sgRNAs generated through in vitro transcription and commercial Cas9. High editing efficiencies are achieved with limited time and financial cost.

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. ...

Endogenous Protein Tagging in Human Induced Pluripotent Stem Cells Using CRISPR/Cas9

A protocol is presented for generating human induced pluripotent stem cells (hiPSCs) that express endogenous proteins fused to in-frame&#160;N- or C-terminal fluorescent tags. The prokaryotic CRISPR/Cas9 system (clustered regularly interspaced short palindromic repeats/CRISPR-associated 9) may be used to introduce large exogenous sequences into genomic loci via homology directed repair (HDR). To achieve the desired knock-in, this protocol employs the ribonucleoprotein (RNP)-based approach where wild type Streptococcus pyogenes Cas9 protein, synthetic 2-part guide RNA (gRNA), and a donor template plasmid are delivered to the cells via electroporation. Putatively edited cells expressing the fluorescently tagged proteins are enriched by fluorescence activated cell sorting (FACS). Clonal lines are then generated and can be analyzed for precise editing outcomes. By introducing the fluorescent tag at the genomic locus of the gene of interest, the resulting subcellular localization and dynamics of the fusion protein can be studied under endogenous regulatory control, a key improvement over conventional overexpression systems. The use of hiPSCs as a model system for gene tagging provides the opportunity to study the tagged proteins in diploid, nontransformed cells. Since hiPSCs can be differentiated into multiple cell types, this approach provides the opportunity to create and study tagged proteins in a variety of isogenic cellular contexts.

Described here is a protocol for tagging endogenously expressed proteins with fluorescent tags in human induced pluripotent stem cells using CRISPR/Cas9. Putatively edited cells are enriched by fluorescence activated cell sorting and clonal cell lines are generated.

A protocol is presented for generating human induced pluripotent stem cells (hiPSCs) that express endogenous proteins fused to in-frame&#160;N- or ...

CRISPR/Cas9 Ribonucleoprotein-mediated Precise Gene Editing by Tube Electroporation

Gene editing nucleases, represented by CRISPR-associated protein 9 (Cas9), are becoming mainstream tools in biomedical research. Successful delivery of CRISPR/Cas9 elements into the target cells by transfection is a prerequisite for efficient gene editing. This protocol demonstrates that tube electroporation (TE) machine-mediated delivery of CRISPR/Cas9 ribonucleoprotein (RNP), along with single-stranded oligodeoxynucleotide (ssODN) donor templates to different types of mammalian cells, leads to robust precise gene editing events. First, TE was applied to deliver CRISPR/Cas9 RNP and ssODNs to induce disease-causing mutations in the interleukin 2 receptor subunit gamma (IL2RG) gene and sepiapterin reductase (SPR) gene in rabbit fibroblast cells. Precise mutation rates of 3.57%-20% were achieved as determined by bacterial TA cloning sequencing. The same strategy was then used in human iPSCs on several clinically relevant genes including epidermal growth factor receptor (EGFR), myosin binding protein C, cardiac (Mybpc3), and hemoglobin subunit beta (HBB). Consistently, highly precise mutation rates were achieved (11.65%-37.92%) as determined by deep sequencing (DeepSeq). The present work demonstrates that tube electroporation of CRISPR/Cas9 RNP represents an efficient transfection protocol for gene editing in mammalian cells.

Presented here is a protocol for efficient CRISPR/Cas9 ribonucleoprotein-mediated gene editing in mammalian cells using tube electroporation.

Gene editing nucleases, represented by CRISPR-associated protein 9 (Cas9), are becoming mainstream tools in biomedical research. Successful delivery of ...

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 ...

Investigation of the Transcriptional Role of a RUNX1 Intronic Silencer by CRISPR/Cas9 Ribonucleoprotein in Acute Myeloid Leukemia Cells

The bulk of the human genome (~98%) is comprised of non-coding sequences. Cis-regulatory elements (CREs) are non-coding DNA sequences that contain binding sites for transcriptional regulators to modulate gene expression. Alterations of CREs have been implicated in various diseases including cancer. While promoters and enhancers have been the primary CREs for studying gene regulation, very little is known about the role of silencer, which is another type of CRE that mediates gene repression. Originally identified as an adaptive immunity system in prokaryotes, CRISPR/Cas9 has been exploited to be a powerful tool for eukaryotic genome editing. Here, we present the use of this technique to delete an intronic silencer in the human RUNX1 gene and investigate the impacts on gene expression in OCI-AML3 leukemic cells. Our approach relies on electroporation-mediated delivery of two preassembled Cas9/guide RNA (gRNA) ribonucleoprotein (RNP) complexes to create two double-strand breaks (DSBs) that flank the silencer. Deletions can be readily screened by fragment analysis. Expression analyses of different mRNAs transcribed from alternative promoters help evaluate promoter-dependent effects. This strategy can be used to study other CREs and is particularly suitable for hematopoietic cells, which are often difficult to transfect with plasmid-based methods. The use of a plasmid- and virus-free strategy allows simple and fast assessments of gene regulatory functions.

Investigation of the Transcriptional Role of a RUNX1 Intronic Silencer by CRISPR/Cas9 Ribonucleoprotein in Acute Myeloid Leukemia Cells

Direct delivery of preassembled Cas9/guide RNA ribonucleoprotein complexes is a fast and efficient means for genome editing in hematopoietic cells. Here, we utilize this approach to delete a RUNX1 intronic silencer and examine the transcriptional responses in OCI-AML3 leukemic cells.

The bulk of the human genome (~98%) is comprised of non-coding sequences. Cis-regulatory elements (CREs) are non-coding DNA sequences that contain binding ...