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://jove.unicartagenaproxy.elogim.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 cm&lt;sup&gt;2&lt;/sup&gt; double seal cap culture flask</td><td>AGC Techno Glass</td><td>3100-025</td><td></td></tr><tr><td>75 cm&lt;sup&gt;2&lt;/sup&gt; 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 (&amp;ndash;)</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

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 ...

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 ...

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. ...

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.

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.

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

Development of Knock-Out Muscle Cell Lines using Lentivirus-Mediated CRISPR/Cas9 Gene Editing

One important application of clustered regulatory interspaced short palindromic repeats (CRISPR)/Cas 9 is the development of knock-out cell lines, specifically to study the function of new genes/proteins associated with a disease, identified during the genetic diagnosis. For the development of such cell lines, two major issues have to be untangled: insertion of the CRISPR tools (the Cas9 and the guide RNA) with high efficiency into the chosen cells, and restriction of the Cas9 activity to the specific deletion of the chosen gene. The protocol described here is dedicated to the insertion of the CRISPR tools in difficult to transfect cells, such as muscle cells. This protocol is based on the use of lentiviruses, produced with plasmids publicly available, for which all the cloning steps are described to target a gene of interest. The control of Cas9 activity has been performed using an adaptation of a previously described system called KamiCas9, in which the transduction of the cells with a lentivirus encoding a guide RNA targeting the Cas9 allows the progressive abolition of Cas9 expression. This protocol has been applied to the development of a RYR1-knock out human muscle cell line, which has been further characterized at the protein and functional level, to confirm the knockout of this important calcium channel involved in muscle intracellular calcium release and in excitation-contraction coupling. The procedure described here can easily be applied to other genes in muscle cells or in other difficult to transfect cells and produce valuable tools to study these genes in human cells.

The protocol describes how to generate knock-out myoblasts using CRISPR/Cas9, starting from the design of guide-RNAs to the cellular cloning and characterization of the knock-out clones.

One important application of clustered regulatory interspaced short palindromic repeats (CRISPR)/Cas 9 is the development of knock-out cell lines, ...

Embryo Microinjection and Knockout Mutant Identification of CRISPR/Cas9 Genome-Edited Helicoverpa Armigera (H&#252;bner)

The cotton bollworm, Helicoverpa armigera, is one of the most destructive pests in the world. A combination of molecular genetics, physiology, functional genomics, and behavioral studies has made H. armigera a model species in Lepidoptera Noctuidae. To study the in vivo functions of and interactions between different genes, clustered regularly interspaced short palindromic repeats (CRISPR)/ associated protein 9 (Cas9) genome editing technology is a convenient and effective method used for performing functional genomic studies. In this study, we provide a step-by-step systematic method to complete gene knockout in H. armigera using the CRISPR/Cas9 system. The design and synthesis of guide RNA (gRNA) are described in detail. Then, the subsequent steps consisting of gene-specific primer design for guide RNA (gRNA) creation, embryo collection, microinjection, insect rearing, and mutant detection are summarized. Finally, troubleshooting advice and notes are provided to improve the efficiency of gene editing. Our method will serve as a reference for the application of CRISPR/Cas9 genome editing in H. armigera as well as other Lepidopteran moths.

Embryo Microinjection and Knockout Mutant Identification of CRISPR/Cas9 Genome-Edited Helicoverpa Armigera H&#252;bner

Presented here is a protocol of Helicoverpa armigera (H&#252;bner) embryo microinjection and knockout mutant identification created by CRISPR/Cas9 genome editing. Mutant insects enable further research of gene function and interaction among different genes in vivo.

The cotton bollworm, Helicoverpa armigera, is one of the most destructive pests in the world. A combination of molecular genetics, physiology, functional ...

Behavior

A Protocol for Multiple Gene Knockout in Mouse Small Intestinal Organoids Using a CRISPR-concatemer

CRISPR/Cas9 technology has greatly improved the feasibility and speed of loss-of-function studies that are essential in understanding gene function. In higher eukaryotes, paralogous genes can mask a potential phenotype by compensating the loss of a gene, thus limiting the information that can be obtained from genetic studies relying on single gene knockouts. We have developed a novel, rapid cloning method for guide RNA (gRNA) concatemers in order to create multi-gene knockouts following a single round of transfection in mouse small intestinal organoids. Our strategy allows for the concatemerization of up to four individual gRNAs into a single vector by performing a single Golden Gate shuffling reaction with annealed gRNA oligos and a pre-designed retroviral vector. This allows either the simultaneous knockout of up to four different genes, or increased knockout efficiency following the targeting of one gene by multiple gRNAs. In this protocol, we show in detail how to efficiently clone multiple gRNAs into the retroviral CRISPR-concatemer vector and how to achieve highly efficient electroporation in intestinal organoids. As an example, we show that simultaneous knockout of two pairs of genes encoding negative regulators of the Wnt signaling pathway (Axin1/2 and Rnf43/Znrf3) renders intestinal organoids resistant to the withdrawal of key growth factors.

This protocol describes the steps for cloning multiple single guide RNAs into one guide RNA concatemer vector, which is of particular use in creating multi-gene knockouts using CRISPR/Cas9 technology. The generation of double knockouts in intestinal organoids is shown as a possible application of this method.

CRISPR/Cas9 technology has greatly improved the feasibility and speed of loss-of-function studies that are essential in understanding gene function. In ...

Bioengineering

Designing, Packaging, and Delivery of High Titer CRISPR Retro and Lentiviruses via Stereotaxic Injection

Replication defective lentiviruses or retroviruses are capable of stably integrating transgenes into the genome of an infected host cell. This technique has been widely used to encode fluorescent proteins, opto- or chemo-genetic controllers of cell activity, or heterologous expression of human genes in model organisms. These viruses have also successfully been used to deliver recombinases to relevant target sites in transgenic animals, or even deliver small hairpin or micro RNAs in order to manipulate gene expression. While these techniques have been fruitful, they rely on transgenic animals (recombinases) or frequently lack high efficacy and specificity (shRNA/miRNA). In contrast, the CRISPR/Cas system uses an exogenous Cas nuclease which targets specific sites in an organism's genome via an exogenous guide RNA in order to induce double stranded breaks in DNA. These breaks are then repaired by non-homologous end joining (NHEJ), producing insertion and deletion (indel) mutations that can result in deleterious missense or nonsense mutations. This manuscript provides detailed methods for the design, production, injection, and validation of single lenti/retro virus particles that can stably transduce neurons to express a fluorescent reporter, Cas9, and sgRNAs to knockout genes in a model organism.

The CRISPR/Cas9 system offers the potential to make targeted genome editing accessible and affordable to the scientific community. This protocol is intended to demonstrate how to create viruses that will knockout a gene of interest using the CRISPR/Cas9 system, and then inject them stereotaxically into the adult mouse brain.

Replication defective lentiviruses or retroviruses are capable of stably integrating transgenes into the genome of an infected host cell. This technique ...

Neuroscience

Electroporation-Based CRISPR-Cas9-Mediated Gene Knockout in THP-1 Cells and Single-Cell Clone Isolation

The human acute monocytic leukemia (AML) THP-1 cell line is widely used as a model to study the functions of human monocyte-derived macrophages, including their interplay with significant human pathogens such as the human immunodeficiency virus (HIV). Compared to other immortalized cell lines of myeloid origin, THP-1 cells retain many intact inflammatory signaling pathways and display phenotypic characteristics that more closely resemble those of primary monocytes, including the ability to differentiate into macrophages when treated with phorbol-12-myristate 13-acetate (PMA). The use of CRISPR-Cas9 technology to engineer THP-1 cells through targeted gene knockout (KO) provides a powerful approach to better characterize immune-related mechanisms, including virus-host interactions. This article describes a protocol for efficient CRISPR-Cas9-based engineering using electroporation to deliver pre-assembled Cas9:sgRNA ribonucleoproteins into the cell nucleus. Using multiple sgRNAs targeting the same locus at slightly different positions results in the deletion of large DNA fragments, thereby increasing editing efficiency, as assessed by the T7 endonuclease I assay. CRISPR-Cas9-mediated editing at the genetic level was validated by Sanger sequencing followed by Inference of CRISPR Edits (ICE) analysis. Protein depletion was confirmed by immunoblotting coupled with a functional assay. Using this protocol, up to 100% indels in the targeted locus and a decrease of over 95% in protein expression were achieved. The high editing efficiency makes it convenient to isolate single-cell clones by limiting dilution.

The THP-1 cell line is widely used as a model to investigate the functions of human monocytes/macrophages across various biology-related research areas. This article describes a protocol for efficient CRISPR-Cas9-based engineering and single-cell clone isolation, enabling the production of robust and reproducible phenotypic data.

The human acute monocytic leukemia (AML) THP-1 cell line is widely used as a model to study the functions of human monocyte-derived macrophages, including ...

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

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Electroporation-Based CRISPR-Cas9-Mediated Gene Knockout in THP-1 Cells and Single-Cell Clone Isolation