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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.


Introduction
Host range determines the extent of a parasites' prevalence. Some parasites have very specific host requirements that limit the area from which they are found, others are generalists. One such generalist is Toxoplasma gondii (T. gondii). This parasite is found worldwide as it can infect all mammals and many birds. Humans are also susceptible and it is estimated that approximately 1/3 of the global population has been infected. Fortunately, a robust immune response normally controls the growth of the parasite, but in situations where the immune system is compromised the parasite can grow unchecked and cause diseases, often encephalic. Also, this parasite can cause congenital diseases if previously uninfected women are infected during pregnancy as they lack immune memory to quickly limit the spread of the parasite. Additionally, there is a burden of ocular toxoplasmosis that can result in vision loss 1 . For these reasons T. gondii has become a focus of study, and due to the many molecular methods developed for its study, a model for Apicomplexan parasites. Two methods that will be discussed here are Quantitative Trait Locus (QTL) mapping and Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas9 gene editing. QTL mapping and CRISPR/Cas9 editing are, respectively, forward and reverse genetic approaches that have been used in previous studies to identify and/or characterize T. gondii virulence genes. Here, these methods are combined to identify and confirm the function of a sinefungin resistance (SNF r ) gene, TgME49_290860, and its orthologs (annotated as SNR1) 2 .
Although T. gondii can infect a wide variety of intermediate hosts, it must pass through and infect the intestinal epithelial cells of a felid to complete the life cycle. Cats are the definitive hosts of the parasite where the sexual stages are produced and genetic recombination via meiosis occurs. To conduct a QTL study one must create a genetic cross, and in the case of Toxoplasma this means passing two different parasite strains that differ in a phenotypic trait, the parental stains, through a cat to produce recombinant progeny 3 . Before being fed to cats, the parental strains are made resistant to separate drugs to allow for more efficient identification of recombinants by double drug selection of the progeny 4 . Three drugs have been used in T. gondii for this purpose; fluorodeoxyribose (FUDR) for which uracil phosphoribosyl transferase (UPRT) is the resistance gene 5 , adenosine arabinoside (ARA) for which Adenosine Kinase (AK) is the resistance gene 6 , and sinefungin (SNF) for which the resistance gene was unknown 7 . Several genetic crosses have been created for T. gondii, but only the 24 progeny of the ME49-FUDR r X VAND-SNF r cross were genotyped using whole genome sequencing (WGS) rendered some of the progeny sinefungin resistant.
In order to identify the causal single nucleotide polymorphism (SNP) in the SNF r progeny, several computational based open source resources can be used to analyze the data. To create the genetic map for the ME49-FUDR r X VAND-SNF r cross the REDHORSE software suite 9 was developed that uses WGS alignments of the parents and progeny to accurately detect the genomic positions of genetic crossovers. This mapping information can then be combined with the phenotypic data (SNF r in the progeny) to format a dataset compatible with the 'qtl' package 10 in the R statistical programming software where a QTL scan can be run to reveal significant loci correlated with the phenotype. To identify the causal SNP located within the QTL locus, the WGS reads of the progeny can be individually aligned to the sinefungin sensitive VAND genome using the Bowtie 2 alignment program 11 from which SNPs can be called using the VarScan mpileup2snp variant caller program 12 . Using these SNPs, the QTL locus can then be scanned for polymorphisms that are present in the SNF r but not in the SNF s progeny. With the causal SNP identified in the coding region of a gene, genetic modification of the candidate SNF r gene can be performed in a SNF s strain to verify the drug resistance function.
The CRISPR/Cas9 genome editing system was recently established in Toxoplasma 13 , which added important tools for the exploration of the complex biology of this parasite, particularly for genetic studies in non-laboratory adapted strains. Because of the highly active Non-Homologous End Joining (NHEJ) activity in WT Toxoplasma cells, targeted genome modification is difficult to achieve since exogenously introduced DNA is randomly integrated to the genome at extremely high frequency 14 . To increase the success rate of locus specific modification, different approaches have been employed to increase the efficiency of homologous recombination and/or decrease the NHEJ activity 15,16 . One of these approaches is the CRISPR/Cas9 system. Compared to other methods, the CRISPR/Cas9 system is efficient at introducing site-specific modifications and is easy to design 13,17,18 . In addition, it can be used in any Toxoplasma strain without extra modification to the parasite 13,19 .
The CRISPR/Cas9 system originated from the adaptive immune system of Streptococcus pyogenes, which uses it to defend the invasion of mobile genetic elements such as phages 20,21,22 . This system utilizes the RNA-guided DNA endonuclease enzyme Cas9 to introduce doublestrand DNA break (DSB) into the target, which is subsequently repaired either by the error-prone NHEJ to inactivate target genes through short indel mutations, or by homology directed recombination to alter the target locus exactly as designed 23,24 . The target specificity is determined by a small RNA molecule named single guide RNA (gRNA), which contains an individually designed 20 nt sequence that has 100% homology to the target DNA 22 . The gRNA molecule also contains signatures recognized by Cas9 that guide the nuclease to the target site, which includes a special Protospacer Adjacent Motif (PAM, sequence is 'NGG') 25,26 . Therefore, the gRNA molecule and the PAM sequence work together to determine the Cas9 cleavage site in the genome. One can easily change the gRNA sequence to target different sites for cleavage.
When the CRISPR/Cas9 system was first developed in Toxoplasma, a single plasmid expressing the Cas9 nuclease and the gRNA molecule was used to introduce DSB at the targeting site 13,17 . It has been shown that the CRISPR/Cas9 system drastically increases the efficiency of sitespecific genome modification, not only by homologous recombination, but also non-homologous integration of exogenous DNA 13 . It does this in wildtype strains that contain NHEJ activity. Therefore, this system can be used in essentially any Toxoplasma strain for efficient genome editing. In a typical experiment, the target specific CRISPR plasmid and the DNA fragment used to modify the target are co-transfected into parasites. If the DNA fragment used to modify the target contains homologous sequences to the target locus, homologous recombination can be used to repair the DSB introduced by CRISPR/Cas9 to allow precise modification of the target. On the other hand, if the introduced DNA fragment does not contain homologous sequence, it can still be integrated into the CRISPR/Cas9 targeting site. The latter is often used to disrupt genes by insertion of selectable markers or to complement mutants at loci that allow negative selection 13 . Here the SNR1 locus serves as an example to show how CRISPR/Cas9 can be used for gene disruption and the generation of transgenic parasites.

Verification of Identified Hits by CRISPR/Cas9-mediated Gene Inactivation.
NOTE: To confirm the causal SNP identified by QTL mapping and WGS SNP analysis, the corresponding genetic changes need to be made in a WT SNF s background and the resulting phenotype examined. In the case of sinefungin resistance, the responsible mutation results in inactivation of the SNR1 gene by early termination 2 . Therefore, disruption of SNR1 can be used for confirmation. Here a detailed protocol is provided for using CRISPR/Cas9 induced indel mutations to disrupt SNR1, to demonstrate its involvement in sinefungin resistance (Figure 4).  4. Filter the parasite solution into a new conical tube through a membrane with the pore size of 3 µm to remove HFF cells and cellular debris. 5. Pellet the filtered parasites by centrifugation at 400 x g for 10 min. 6. Pour off the supernatant and resuspend the pelleted parasites with 10 mL cytomix buffer, take a 10 µL aliquot to determine the parasite concentration with a hemocytometer and pellet the rest by centrifugation at 400 × g for 10 min. 7. Remove supernatant and resuspend the pellet in cytomix buffer to obtain a density of 4 x 10 7 parasites/mL. 8. In a 4 mm gap cuvette, mix 250-300 µL parasite solution (1-1.3 x 10 7 parasites) with 7.5 µg CRISPR plasmid, 6 µL ATP (100mM), and 6 µL glutathione (GSH) (250 mM). 9. Electroporate the parasites 35 . Include a separate electroporation as a negative control in which a CRISPR plasmid targets elsewhere, such as the UPRT targeting CRIPSR plasmid. NOTE: The electroporation settings depend on the device. For the electroporator listed in the Materials use the 4 mm gap cuvettes. The following protocol is recommended: 1,700 V, 176 µs of pulse length, 2 pulses with 100 ms interval.
3. Determine the frequency of sinefungin resistance after CRISPR targeting.
1. Seed HFF cells to coverslips placed in 24-well plates 3-4 d before electroporation so that they are confluent by the time to perform transfection in 4.2.
1. Trypsinize the HFF cells from one confluent T25 flask and subsequently add 12 mL D10 medium and mix well. 2. Add a coverslip to each well of a 24-well plate and aliquot 500 µL HFF cell solution to each well. Incubate the plate at 37 °C for 3 4 d to let the cells grow until confluent. NOTE: Cells from one T25 flask are enough to seed one 24-well plate. 3. Add 50 µL of electroporated parasite solution from step 4.2.9 into one well of 24-well plate containing a coverslip seeded with confluent HFF cells. Transfer the rest of electroporated parasite solution into a T25 flask seeded with confluent HFF cells.
2. Assess the efficiency of transfection. 1. Grow the cells in the 24-well plate for 24 h and subsequently fix the cells (host cells and parasites) with 500 µl 4% formaldehyde to check the expression of Cas9-GFP by immunofluorescent assay (IFA).

Probe the cells with an anti-GFP antibody and a Toxoplasma specific antibody (such as anti-TgALD) to label total parasites.
See the antibody product sheet for the recommended dilution (usually, 1:1,000 is sufficient for IFAs). If the antibodies are unconjugated, use two different secondary antibodies conjugated with fluorescent dyes to label the primary antibodies. 3. Observe the labeled cells on a fluorescent microscope with filters appropriate for the secondary fluorescent dyes. Obtain transfection efficiency by dividing the number of GFP positive parasites by the number of total parasites. NOTE: The two antibodies used for IFA should come from two different host species, for example using the combination of mouse anti-GFP and rabbit anti-TgALD.
3. Determine the frequency of sinefungin resistance in transfected parasites. 1. For the transfected parasites cultured in T25 flasks, grow them for 2-3 d until natural egress. Then collect the parasites, blunt needle lyse host cells to release intracellular parasites, and purify them by filtration through membranes with pore size of 3 µm. Count the parasites with a hemocytometer to estimate the density. 2. Add purified parasites into 6-well plates seeded with confluent HFF cells: for the first row (3 wells), add 200 parasites/well and grow them in regular D10 medium (2 mL/well); for the second row, add 5000 parasites/well and grow them in D10 medium containing 0.3 µM sinefungin. 3. Put the plates in a 5% CO 2 incubator and grow the parasites for at 37 °C 8-10 d without disturbance to allow plaques to form. Fix the samples with 70% ethanol (2 mL/well) and stain the monolayer with 0.1% crystal violet (2 mL/well). Wash the wells with water to visualize the plaques (clear zones) formed by parasite growth. NOTE: The negative control should be processed the same way side by side.

Representative Results
This article outlines in detail several methods that can be used in succession to identify a gene responsible for drug resistance (Figure 1). The 24 progeny of the ME49-FUDR r X VAND-SNF r cross were assessed for resistance to the drug sinefungin as described in Protocol 1. Using the genetic map and the SNF r phenotype of the progeny, a QTL scan was run in R/qtl -Protocol 2 (Figure 2). This resulted in one significant peak on chromosome IX spanning approximately 1 Mbp. It is in this region that the causal mutation is located.
To identify the causal mutation, WGS reads from the progeny were aligned to the VAND-SNF s reference genome using Bowtie2 -Protocol 3.1, SNPs were called using VarScan mpileup2snp -Protocol 3.2, and the QTL locus in the VAND genome was identified using MUMmer -Protocol 3.3. Progeny SNPs within the QTL locus were extracted and scanned for a pattern where the SNF r progeny have a SNP and the SNF s do not, which is feasible because progeny SNPs were obtained by comparison to the VAND-SNF s reference genome (Figure 3). Only one SNP matched this pattern that results in an early stop codon in a putative amino acid transporter gene named SNR1.
Confirmation that SNR1 is the SNF r resistance gene was performed using the CRISPR/Cas9 system. A new CRISPR/Cas9 plasmid engineered for T. gondii gene editing was made containing a gRNA targeting the SNR1 gene near the SNF r mutation identified in the progeny -Protocol 4 ( Figure 4A). The SNR1 targeting CRISPR/Cas9 plasmid was electroporated into a SNF s WT parasite strain and resistant mutants were obtained when cultured in 0.3 µM sinefungin. No SNF r parasites were obtained when electroporated with the UPRT targeting CRISPR/Cas9 plasmid.
Several SNF r CRISPR mutants were cloned and the region around the SNR1 gRNA target was sequenced. Each mutant had an indel that disrupted the coding sequence of the SNR1 gene ( Figure 4B). This method can also be used to insert a targeting construct into the SNR1 locus via NHEJ (Figure 5), or by HR (Figure 6) -Supplemental File 2.

Discussion
These protocols present several methods that when combined allow for the identification of a drug resistance gene in T. gondii. Two in particular were integral to the project, the relatively seasoned method of QTL mapping and the recently developed method of CRISPR/Cas9 gene editing. Lander and Botstein published their influential paper in 1989 demonstrating QTL mapping which correlates genetic loci with phenotypes 36 . More recently in 2012, Dounda and Charpentier described the CRISPR/Cas9 editing system in Streptococcus pyogenes 22 that was quickly adapted as a genetic tool in many different models, including T. gondii 13,17 . Both methods were useful here, where QTL mapping defined the locus containing the drug resistance mutation that was ultimately identified using WGS based SNP detection, and CRISPR/Cas9 editing provided the means to confirm SNR1 is the sinefungin drug resistance gene.
The ME49-FUDR r X VAND-SNF r cross 8 was originally developed to interrogate a virulence phenotype 19 , but the parental strains also happened to have an additional phenotypic difference for which the causal gene was not known, sinefungin resistance in the VAND parent. This highlights one benefit of crosses in that they can be repurposed when additional phenotypic differences in the parents are observed. This was the case for another Toxoplasma cross, type 2 x type 3, where several genes involved in virulence were found by mapping multiple phenotypes 37,38,39,40 .
To date, four different T. gondii crosses have been described and used to map genes responsible for phenotypes 8,37,41,42 , all of which have the potential to be reused to map new phenotypes for which the genetic basis is unknown. Along these lines, the genomes for 62 T. gondii strains representing the known global genetic diversity have been sequenced 43 . New crosses could be made from this pool for strains that differ in interesting phenotypes. Having extolled the advantages of QTL mapping, it needs to be said that generating a cross isn't a minor undertaking. There are other methods that can be used to identify causal genes. One powerful technique uses chemical mutagenesis to create mutants that can be screened for phenotypes. To find the causal gene, mutants can either be complemented with cosmid libraries 44 or genome resequencing methods can be used to find the causal mutation 45 . For more on this, see two JoVE articles by Coleman et al. and Walwyn et al. that outline these approaches 46,47 .
Many of the steps leading to the identification of the causal SNF r mutation (Protocols 2 and 3) rely on computational methods conducted with software that is freely available for academic use. Detailed commands for each step are provided and when run with the proper files will allow the user to recreate the datasets necessary to find the causal SNF r SNP. Keep in mind that some of the syntax in the commands refer to filenames or directory structure ($PATH) that can be modified to the user preference. Although the commands given here certainly do not exhaust the ways one can analyze a cross with QTL analysis and WGS based SNP identification, they are comprehensive enough to repeat the experiment described in this article and should allow the user to become more familiar with how these approaches are utilized in a step by step fashion.
, and make gene knockouts 16,19,50,51,52,53,54 in T. gondii, promising to be a useful tool for many other studies in the future.
The discovery that SNR1 inactivation leads to sinefungin resistance makes the SNR1 locus a promising site for transgene insertion or genetic complementation. To maximize the success of using CRISPR/Cas9 mediated gene targeting to direct the integration of transgene into the SNR1 locus, the following aspects should be considered during the experimental design. First, a selection marker is recommended to be included in the transgenic construct to increase the efficiency of strain construction. If a selection marker is included, both positive and negative selection can be used, resulting in almost 100% of the doubly selected parasites being transgenic with the GOI integrated at the SNR1 locus. In contrast, if the transgenic construct does not contain additional selectable traits and relies on negative selection at the SNR1 locus, the efficiency of successful transgenesis largely depends on the efficiency of cotransfection of the CRISPR plasmid and the transgenic construct.
Second, although CRISPR/Cas9 mediated site-specific integration of a non-homologous DNA fragment is frequently used for complementation and transgenesis, it should be noted that the orientation of insertion cannot be guaranteed in such cases as either direction is possible. This may pose problems to some applications. For example, to complement a mutant with different gene alleles, it is difficult to ensure that all alleles are inserted in the same orientation, and discrepancies in orientation may cause expression differences. For this type of application, DNA constructs with sequences homologous to the SNR1 locus are recommended, which will drive the proper integration orientation (Figure 6).
Third, during transfection, the ratio between the CRISPR plasmid and the transgenic DNA molecule is critical for successful transgenic strain construction. This ratio needs to be adjusted according to the selection strategies. The following guidelines are recommended: 1) If the transgenic construct contains a drug resistant marker and the corresponding drug is the only selection used to generate transgenic parasites, i.e. sinefungin is not used, the suggested molar ratio between the transgenic construct and the CRISPR plasmid is 1:5. Using more CRISPR plasmid in this case increases the likelihood that parasites receiving the transgenic construct will also receive the CRISPR plasmid, therefore the drug resistant parasites are more likely to have the marker inserted at the CRISPR targeting site. If the ratio is reversed, most parasites that receive the transgenic construct won't get the CRISPR plasmid. As a consequence, the vast majority of drug resistant parasites obtain the transgenic construct through random integration not associated with CRISPR/CAS9 mediated site-specific insertion. 2) If the transgenic construct contains a drug resistant marker and the corresponding drug is used along with sinefungin to select transgenic parasites, the suggested molar ratio between the transgenic construct and the CRISPR plasmid is 1:1. This strategy provides the highest efficiency of transgenic strain construction.
3) If the transgenic construct does not contain a selectable maker, relying on negative selection by sinefungin alone to obtain transgenic parasites, the suggested molar ratio between the transgenic construct and the CRISPR plasmid is 5:1. The rationale for this design is the same as in the first guideline above. Since both pyrimethamine and sinefungin were used for selection in Protocol 5, the ratio between DHFR* mini gene and the SNR1 targeting CRISPR plasmid was set as 1:1.
Taken together, the methods outlined here have a level of detail that was not possible to convey in the original publication that identified SNR1 2 . These protocols; specifically, the command line syntax, sequential layout of the programs utilized, and the use of CRISPR/Cas9 should aid future endeavors to identify new genes responsible for phenotypes.

Disclosures
The authors declare that they have no competing financial interests.