Plasmodium: new tools advance genetic tractability

Massive efforts have been made to develop tools to study Plasmodium spp., and particularly P. falciparum, due to the impact these parasites have on society. Malaria causes severe morbidity and mortality that is sustained by the lack of an effective malaria vaccine and the ability of the parasite to develop drug resistance. In vitro cultivation and genetic manipulation have been available to study P. falciparum for decades, but these approaches have been inefficient and time-consuming. Conventional gene knockout in P. falciparum takes months to obtain a null mutant and relies on spontaneous single- or double-crossover recombination using plasmids containing homologous sequences to the target region. Since P. falciparum lacks the machinery for NHEJ, genome integration is not random and occurs mainly in the region of homology between parasite chromosomes and plasmid DNA maintained episomally. Unfortunately, integration is a stochastic event that occurs at very low frequency and thus gene disruption in P. falciparum requires 1–3 months of continuous culture, protracted on–off cycling of drug selection, and/or negative-selection procedures. Zinc finger nuclease technology (ZFN) has been successfully used in P. falciparum to improve the generation of knockout parasites (Straimer et al. Reference Straimer, Lee, Lee, Zeitler, Williams, Pearl, Zhang, Rebar, Gregory, Llinas, Urnov and Fidock2012; Singer et al. Reference Singer, Marshall, Heiss, Mair, Grimm, Mueller and Frischknecht2015; Veiga et al. Reference Veiga, Dhingra, Henrich, Straimer, Gnadig, Uhlemann, Martin, Lehane and Fidock2016), but is limited by its targeting capability, arduous design and implementation, and high cost. ZFN vectors are also often very large and thus not suitable for genome-wide screening.

Successful establishment of CRISPR/Cas9 technology to edit Plasmodium spp. genomes has provided a powerful tool to allow rapid and efficient genetic manipulation of these parasites. Several adaptations of CRISPR/Cas9 have been developed in Plasmodium spp. based on the use of either one- or two-vector strategies, depending on whether the donor DNA is in the same or a separate plasmid as Cas9 and the sgRNA, and if different selectable markers are used to maintain the plasmids in transformed parasites. Two seminal studies in 2014 independently reported adapting CRISPR/Cas9 technology to genetically manipulate P. falciparum using different approaches to express the sgRNA (Ghorbal et al. Reference Ghorbal, Gorman, Macpherson, Martins, Scherf and Lopez-Rubio2014; Wagner et al. Reference Wagner, Platt, Goldfless, Zhang and Niles2014). In the first study, Ghorbal et al. developed a two-plasmid system expressing Cas9, under the HSP86 promoter, in one vector and the sgRNA, driven by the U6 promoter, along with the HR donor DNA flanking a selectable marker hDHFR (human dihydrofolate reductase) in a second vector. After co-transfecting P. falciparum-infected erythrocytes with both vectors, drug selection for integration of hDHFR at the target locus was applied. The sgRNA/HR donor vector also carried the negative selectable marker yfcu (yeast cytosine deaminase and uridyl phosphoribosyl transferase) to subsequently eliminate parasites carrying copies of this plasmid. This study also found that transfection of linear HR donor plasmid DNA is a viable alternative to negative selection since linear DNA can mediate recombination but does not persist (Deitsch et al. Reference Deitsch, Driskill and Wellems2001). The authors used this system to disrupt a reporter transgene (EGFP) and an endogenous gene (PfKAHRP) with hDHFR, along with introducing a point mutation in two genes (PfORC1 and PfKELCH-13) without drug selection. A possible limitation of expressing sgRNAs from the U6 promoter is that guanosine is preferred at the sgRNA 5′ position for efficient RNA polymerase III transcription. Nevertheless, the authors reported that the P. falciparum U6 promoter was able to drive expression of gRNAs in the parasite without the functional requirement for the initial guanosine nucleotide, thus expanding in P. falciparum the ability of Cas9 to target any sequence with the -NGG PAM motif. We have similarly observed in T. gondii that expression from the U6 promoter seems to be free of this initial guanosine nucleotide restriction, having successfully used sgRNAs starting with one of the other three nucleotides, although efficiency has not been determined (Di Cristina et al. Reference Di Cristina, Dou, Lunghi, Kannan, Huynh, McGovern, Schultz, Schultz, Miller, Hayes, van der Linden, Emiliani, Bogyo, Besteiro, Coppens and Carruthers2017).

In the second study, Wagner et al. also used a two-plasmid system, wherein Cas9 and the sgRNA were placed in the same vector along with a BSD (blasticidin-S deaminase) selection marker. In this system, expression of the sgRNA was driven by the T7 promoter, thus requiring expression of the T7 RNA polymerase from a second plasmid. This second plasmid also carried the HR donor and a NEO (neomycin) resistance cassette to maintain this vector episomally. The authors demonstrated disruption of two individual genes (PfKAHRP and PfEBA-175) without integration of a selectable marker.

These studies were followed by several other reports utilizing similar strategies, including some with new innovations. The Ghorbal approach allowed Nacer et al. to identify PfVAP1 (virulence-associated protein 1) as a key factor involved in the P. falciparum cytoadherence (Nacer et al. Reference Nacer, Claes, Roberts, Scheidig-Benatar, Sakamoto, Ghorbal, Lopez-Rubio and Mattei2015). Using a similar general strategy Lu et al. introduced the use of a suicide vector approach (Lu et al. Reference Lu, Tong, Pan, Yang, Liu, Tan, Zhao, Qin and Chen2016). In this work, the authors designed a suicide vector encoding Cas9 nuclease, sgRNA and a drug selection marker along with a second plasmid carrying the HR donor for cotransfection. Parasites receiving only the Cas9/sgRNA suicide vector die because of inefficient repair of the DSB by MMEJ, whereas those receiving both plasmids survive via HR repair from the donor plasmid. This approach avoids the need for a selectable marker in the donor plasmid, thus freeing up space to introduce larger knock-in tags. The system also has the potential to efficiently mediate consecutive gene manipulations. In another adaptation of the method developed by Ghorbal and colleagues, Mogollon et al. (Reference Mogollon, van Pul, Imai, Ramesar, Chevalley-Maurel, de Roo, Veld, Kroeze, Franke-Fayard, Janse and Khan2016) generated marker-free P. falciparum fluorescent reporter lines by redesigning the vector carrying the sgRNA and HR donor to include a hybrid positive and negative selectable marker (hDHFR-yfcu) outside the homology arms of the HR donor. In this new approach, positive selection is applied initially to maintain the HR donor plasmid followed by negative selection to eliminate parasites containing the episomal plasmid. In very recent work, Bryant et al. used the Ghorbal system to functionally interrogated an important conserved genetic element of var genes, the var2csa intron, without the introduction of a drug-selectable marker (Bryant et al. Reference Bryant, Regnault, Scheidig-Benatar, Baumgarten, Guizetti and Scherf2017). Also, the Wagner approach based on the sgRNA expression driven by the T7 promoter was successfully used in two other studies to introduce point mutations in either PfMDR1 (Ng et al. Reference Ng, Siciliano, Lee, de Almeida, Corey, Bopp, Bertuccini, Wittlin, Kasdin, Le Bihan, Clozel, Winzeler, Alano and Fidock2016) or PfCARL (LaMonte et al. Reference LaMonte, Lim, Wree, Reimer, Nachon, Corey, Gedeck, Plouffe, Du, Figueroa, Yeung, Bifani and Winzeler2016) to identify residues involved in drug resistance.

Distinct from the two-plasmid CRISPR/Cas9 systems developed for P. falciparum, Zhang and colleagues developed a P. yoelii system based on one-vector carrying all the products required for cleavage and DNA repair (Zhang et al. Reference Zhang, Xiao, Jiang, Zhao, Li, Gao, Ling, Wei, Li, Lu, Su, Cui and Yuan2014). The authors used this system to generate targeted deletion, reporter knock-in, and nucleotide replacement in multiple genes with high efficiency and accuracy. In subsequent work, the same group improved this single-vector system by introducing the yfcu negative selection marker to remove the episomal vector in transfected parasites after the genomic editing was achieved (Zhang et al. Reference Zhang, Gao, Yang, Jiang, Li, Wang, Xiao, Su, Cui and Yuan2017a, Reference Zhang, Gao, Wang and Zhaob). The CRISPR/Cas9 system was also employed by Knuepfer et al. to introduce a rapamycin-inducible DiCre recombinase for conditional disruption of target genes engineered with flanking loxP sites (Knuepfer et al. Reference Knuepfer, Napiorkowska, van Ooij and Holder2017). DiCre-mediated gene deletion was also achieved using the Ghorbal CRISPR/Cas9 adaptation by Volz et al. (Reference Volz, Yap, Sisquella, Thompson, Lim, Whitehead, Chen, Lampe, Tham, Wilson, Nebl, Marapana, Triglia, Wong, Rogers and Cowman2016) to demonstrate the essential role of PfRh5/PfRipr/CyRPA complex during P. falciparum invasion of erythrocytes.

Very recently, P. falciparum transfection of a purified CRISPR/Cas9-guide RNA ribonucleoprotein complex and a 200-nucleotide single-stranded oligodeoxynucleotide (ssODN) repair template introduced drug resistance mutations without the use of plasmids or the need for cloning homologous recombination templates (Crawford et al. Reference Crawford, Quan, Horst, Ebert, Wu and DeRisi2017). This approach is ideally suited for introducing mutations that confer a fitness advantage under selection conditions. The efficacy of using ssODN as a repair template in P. falciparum may greatly simplify future genome editing of these parasites.

Toxoplasma gondii: higher throughput and genome-wide screens

Contrary to Plasmodium spp., T. gondii is an apicomplexan parasite offering relative ease of growth in culture and a wide array of genetic tools that include chemical or insertional mutagenesis, homologous gene replacement, conditional knockdown and tagging techniques (Wang et al. Reference Wang, Huang, Behnke, Chen, Shen and Zhu2016a). The availability of numerous selectable markers for generation of stable Toxoplasma strains (Roos et al. Reference Roos, Donald, Morrissette and Moulton1994, Reference Roos, Sullivan, Striepen, Bohne and Donald1997; Donald and Roos, Reference Donald and Roos1995, Reference Donald and Roos1998; Fox et al. Reference Fox, Belperron and Bzik1999, Reference Fox, Belperron and Bzik2001; Soete et al. Reference Soete, Hettman and Soldati1999; Wang et al. Reference Wang, Huang, Behnke, Chen, Shen and Zhu2016a) and the rapid loss of exogenous non-integrated DNA, with no detectable exogenous DNA 7 days post-transfection (Soldati and Boothroyd, Reference Soldati and Boothroyd1993; Black and Boothroyd, Reference Black and Boothroyd1998), makes the study of the biology of T. gondii by genetic manipulations more manageable. Integration of foreign DNA in T. gondii is relatively efficient, with recombination rates of ~0.1% without restriction enzyme-mediated integration (REMI) and ~5% with REMI (Black et al. Reference Black, Seeber, Soldati, Kim and Boothroyd1995; Roos et al. Reference Roos, Sullivan, Striepen, Bohne and Donald1997). The presence of a very active NHEJ pathway in T. gondii is a limitation for generating strains with homology-directed knockout or the precise insertion of tags or mutations because of a high prevalence of random DNA integration. Due to this feature, conventional generation of knock out strains for T. gondii is inefficient and requires homology flanks of 2–3 kbp (Donald and Roos, Reference Donald and Roos1994; Roos et al. Reference Roos, Sullivan, Striepen, Bohne and Donald1997; Zhang et al. Reference Zhang, Kim, Ma, Wittner, Tanowitz and Weiss1999; Craver and Knoll, Reference Craver and Knoll2007). To overcome this, T. gondii type I (e.g., RH) or type II (e.g., Prugniaud, Pru) strains were modified to disrupt NHEJ-mediated insertion by deleting one key component of this pathway, the gene encoding the Ku80 protein (Fox et al. Reference Fox, Ristuccia, Gigley and Bzik2009, Reference Fox, Falla, Rommereim, Tomita, Gigley, Mercier, Cesbron-Delauw, Weiss and Bzik2011; Huynh and Carruthers, Reference Huynh and Carruthers2009). The impact of these two strains, named RHΔku80 and PruΔku80, on understanding Toxoplasma biology has been substantial, allowing higher fidelity, rapid generation of knockout strains, and the introduction of epitope tags. Although the generation of Δku80 strains minimized the problem of random DNA integration, this approach, of course, limits the studies to these two strains currently. Moreover, since the Ku80 protein is involved in DNA repair, strains lacking this gene may be prone to accumulate genetic mutations after prolonged culture.

Introduction of CRISPR/Cas9 into the Toxoplasma field has revolutionized the capability to efficiently generate gene knockouts in any strain. Several CRISPR/Cas9-based approaches have been developed in different laboratories to inactivate selected gene function. For example, complete or partial deletion of the target sequence was obtained through double crossover triggered by a site-directed sgRNA/Cas9-mediated DSB and subsequent DNA repair using a donor DNA comprised a drug resistance expression cassette flanked by about 1 kbp of DNA homologous to the target locus (Shen et al. Reference Shen, Brown, Lee and Sibley2014a). This approach also substantially increased the throughput of gene deletions, exemplified by individual or sequential disruption of entire gene families (Shen et al. Reference Shen, Buguliskis, Lee and Sibley2014b). Gene inactivation was also obtained via ‘indels’ in the coding region generated by NHEJ repair of a sgRNA/Cas9-mediated DSB (Sidik et al. Reference Sidik, Hackett, Tran, Westwood and Lourido2014; Wang et al. Reference Wang, Huang, Li, Chen, Ning and Zhu2016b). A ‘clean’ knockout with complete gene deletion is desirable because it avoids the potential expression of a truncated protein and precludes homologous reinsertion of the gene or cDNA for genetic complementation. Nonetheless, template mediated complete gene deletion is less efficient in non-Δku80 strains because homologous recombination is active only when parasites are in the S/G2 phase and thus NHEJ is the prevalent form of DNA repair in T. gondii when extracellular, G0-phase parasites are used for transfection. To enhance HR vs. NHEJ events, Behnke and colleagues (Behnke et al. Reference Behnke, Khan, Lauron, Jimah, Wang, Tolia and Sibley2015) developed a new CRISPR vector that expresses Cas9 and two sgRNAs. The two sgRNAs direct Cas9 to generate two DSBs, one at each end of the target gene or locus. This tactic not only improves efficiency, but it also permits efficient disruption of large genes or tandem gene arrays. In our hands, the two-sgRNA approach allowed successful knock out of several genes individually including those encoding TgCPL (cathepsin protease L) (Di Cristina et al. Reference Di Cristina, Dou, Lunghi, Kannan, Huynh, McGovern, Schultz, Schultz, Miller, Hayes, van der Linden, Emiliani, Bogyo, Besteiro, Coppens and Carruthers2017) and TgASP1 (aspartic protease 1) (Di Cristina and Carruthers, unpublished) in the T. gondii type II strain ME49. In summary, CRISPR/Cas9 is opening up genetic manipulation of any T. gondii strain and allowing for large genetic disruptions to interrogate gene function.

Although CRISPR/Cas9 is versatile for any strain, applying it in a Δku80 strain provides the advantage of using short homology sequences, thereby permitting convenient and precise gene knockouts or knocking of tags or mutations. For example, Sidik and co-authors (Sidik et al. Reference Sidik, Hackett, Tran, Westwood and Lourido2014) introduced tags or mutations in a Δku80 strain using synthetic oligonucleotide repair templates with 40 bp of homology without the need for a selectable marker. This strategy is based on co-transfecting a Cas9 + sgRNA expression plasmid with a synthetic double strand oligonucleotide bearing the desired tag or mutation together with a silent mutation to eliminate the PAM site NGG beside the 20 bp sequence targeted by the gRNA. The efficiency of this approach was enhanced by fluorescence-activated cell sorting (FACS) parasites that received the Cas9/sgRNA vector, exploiting the fluorescence emitted by the GFP fused to the Cas9 protein. Only ~20–30% of parasites obtain the Cas9/sgRNA vector after electroporation, making it critical to remove the predominant fraction of non-transfected parasites to enrich the population with edited parasites. Since FACS is expensive and not available for all the laboratories, we developed a protocol that allows the enrichment of Cas9-expressing parasites with a Cas9/sgRNA vector bearing a bleomycin resistance gene (Di Cristina and Carruthers, unpublished). Parasites receiving this new plasmid, named pCas9/sgRNA/Bleo, are subjected to phleomycin treatment 24 h after transfection to eliminate parasites that have not received the plasmid and enrich for parasites expressing Cas9 and the sgRNA. Treatment one day after transfection ensures that bleomycin resistance is transiently expressed by the 20–30% of the population that incorporate the plasmid. Stable integration is not favoured due to the Cas9 toxicity, which works as a negative selection against vector integration. In our hands, this approach results in about 30–80% efficiency of edited or tagged parasites, depending on the impact to parasite fitness of the mutation introduced, allowing easy identification of single mutant clones. Beyond this variation of the approach, the reader is referred to an excellent how-to guide for using CRISPR/Cas9 for various applications in Toxoplasma that was published recently (Shen et al. Reference Shen, Brown, Long and Sibley2017).

High-throughput strategies to genetically engineer and screen large numbers of mutants or populations are powerful weapons in modern systems biology. To this end, the emergence of CRISPR/Cas9 has prompted new screening approaches using sgRNA libraries to perform genome-wide knockouts in a parasite population. Such approaches allow measuring the fitness contribution of every gene in the parasite genome. Exploiting the high rates of NHEJ in T. gondii, Sebastian Lourido's laboratory efficiently created frame-shift mutations and insertions at the DSBs generated by transfecting a Cas9-expressing RH strain with a library of sgRNAs containing 10 guides against each of the 8158 predicted T. gondii protein-coding genes (Sidik et al. Reference Sidik, Huet, Ganesan, Huynh, Wang, Nasamu, Thiru, Saeij, Carruthers, Niles and Lourido2016). The guide RNA library was cloned into the sgRNA expression vector and the integrated sgRNAs were exploited as barcodes to measure the contribution of each gene to parasite fitness. Generation of a Cas9-expressing RH strain was likely instrumental in obtaining high rates of gene disruption. Due to the toxicity of Cas9 expression in T. gondii, as observed for other microorganisms (Jiang et al. Reference Jiang, Brueggeman, Horken, Plucinak and Weeks2014; Peng et al. Reference Peng, Kurup, Yao, Minning and Tarleton2014), the authors developed a strategy to obtain strains of T. gondii stably expressing this nuclease by co-expressing a decoy sgRNA to prevent the detrimental effect to parasites by unintended Cas9 activity directed by endogenous RNAs. This work represents the first genome-wide functional analysis of an apicomplexan, thus providing broad-based functional information on T. gondii genes and their contributions to parasite fitness during infection of human fibroblasts. One initial limitation of this outstanding work is its restriction to the tachyzoite stage of T. gondii, the rapidly growing form responsible for the acute phase of the infection. Applying this approach to other life stages will require improvements to the transfection and integration efficiencies of strains that competently differentiate into other stages. The contribution to tachyzoite fitness of each predicted T. gondii protein-coding genes is now available (www.toxodb.org).

Recently, David Sibley's group developed an auxin-inducible degron (AID) tagging system for conditional protein depletion in T. gondii (Brown et al. Reference Brown, Long and Sibley2017; Long et al. Reference Long, Brown, Drewry, Anthony, Phan and Sibley2017). They exploited a new combination of CRISPR/Cas9-mediated gene editing and a plant-derived AID system to identify which cyclic GMP (cGMP)-dependent protein kinase G (PKG) isoforms are necessary for PKG-dependent cellular processes (Brown et al. Reference Brown, Long and Sibley2017) and to examine the roles of three apically localized calmodululin-like proteins (Long et al. Reference Long, Brown, Drewry, Anthony, Phan and Sibley2017). Adaptation of the AID system to T. gondii adds a powerful new tool to identify the consequences of rapidly down-regulating expression of cytosolic proteins to infer function.

CRISPR technology has been adapted to apicomplexan parasites relatively recently and thus has not been fully exploited and expanded. In mammals, evolution of this technology led to the development of tissue or time-specific promoters to restrict the genome editing to a precise cell type or developmental stage (Harrison et al. Reference Harrison, Jenkins, O'Connor-Giles and Wildonger2014; Ablain et al. Reference Ablain, Durand, Yang, Zhou and Zon2015; Bortesi and Fischer, Reference Bortesi and Fischer2015; Wang et al. Reference Wang, Li, Zhao, Li, Zhou and Liu2015; Yoshioka et al. Reference Yoshioka, Fujii, Ogawa, Sugiura and Naito2015; Lee et al. Reference Lee, Ng and Ingham2016; Xu et al. Reference Xu, Zhao, Gao, Xu and Han2017; Zhang et al. Reference Zhang, Gao, Wang and Zhao2017b). In the classic CRISPR technology, sgRNAs are usually transcribed under control of RNA polymerase III promoters to obtain transcripts devoid of both capping and poly(A) tails, thereby generating the correct 5′-end of the sgRNA and avoiding exportation of the sgRNA to the cytoplasm, respectively. Tissue/developmental-specific sgRNA expression requires using RNA polymerase II promoters active exclusively in the desired cell type or stage. To generate RNA polymerase II-driven functional sgRNAs, a strategy based on the use of ribozymes was developed by several laboratories (Yoshioka et al. Reference Yoshioka, Fujii, Ogawa, Sugiura and Naito2015; Ng and Dean, Reference Ng and Dean2017; Xu et al. Reference Xu, Zhao, Gao, Xu and Han2017; Zhang et al. Reference Zhang, Gao, Yang, Jiang, Li, Wang, Xiao, Su, Cui and Yuan2017a, Reference Zhang, Gao, Wang and Zhaob). Hammerhead or hepatitis delta virus ribozymes perform site-specific self-cleavage, resulting in mature sgRNAs with correct 5′- and 3′-ends. This ribozyme-flanked gRNA expression system can be exploited for the spatiotemporal expression of gRNA employing cell type or developmental-specific promoters. In Toxoplasma, this approach has not been investigated yet but may allow programmed gene inactivation in a stage-specific manner by exploiting promoters active at specific phases of the parasite life cycle. For example expressing the ribozyme-sgRNA and Cas9 under the control of a bradyzoite-specific promoter could allow stage-specific inactivation of all genes, even those that are essential for tachyzoites. This approach would allow assessing the role of essential genes during the chronic stage of T. gondii. Moreover, multiple sgRNAs linked with self-cleaving ribozymes could be simultaneously expressed from a single promoter to exert genome editing at different sites. Alternatively, several new approaches permit conditional expression of Cas9 via chemical or optical activation (Nihongaki et al. Reference Nihongaki, Kawano, Nakajima and Sato2015; Polstein and Gersbach, Reference Polstein and Gersbach2015; Wright et al. Reference Wright, Sternberg, Taylor, Staahl, Bardales, Kornfeld and Doudna2015; Zetsche et al. Reference Zetsche, Volz and Zhang2015; Liu et al. Reference Liu, Ramli, Woo, Wang, Zhao, Zhang, Yim, Chong, Gowher, Chua, Jung, Lee and Tan2016). In principle, it should also be possible to append a destabilization domain to Cas9 for ligand-dependent expression of Cas9 in any stage of the parasite.

Cryptosporidium: introducing a new genetic era

Cryptosporidium spp. causes severe diarrhoea in young children, with 10% mortality in such cases (Liu et al. Reference Liu, Johnson, Cousens, Perin, Scott, Lawn, Rudan, Campbell, Cibulskis and Li2012). Cryptosporidiosis also causes life-threatening chronic disease in immunocompromised individuals, including those afflicted by HIV/AIDS. Infections occur worldwide in association with oocyst contaminated water, with no vaccines available, and only a single drug (nitazoxanide) has been approved with limited benefit for malnourished children and immunocompromised patients (Amadi et al. Reference Amadi, Mwiya, Musuku, Watuka, Sianongo, Ayoub and Kelly2002, Reference Amadi, Mwiya, Sianongo, Payne, Watuka, Katubulushi and Kelly2009). Progress in understanding Cryptosporidium spp. biology and developing new treatments have been hindered by the limited tractability of the parasite, which includes a lack of systems for continuous culture, the absence of facile animal models, and the dearth of molecular genetic tools (Striepen, Reference Striepen2013; Checkley et al. Reference Checkley, White, Jaganath, Arrowood, Chalmers, Chen, Fayer, Griffiths, Guerrant, Hedstrom, Huston, Kotloff, Kang, Mead, Miller, Petri, Priest, Roos, Striepen, Thompson, Ward, Van Voorhis, Xiao, Zhu and Houpt2015). Cryptosporidium spp. cultures last a few days in vitro since parasites undergo one or two rounds of replication at most, limiting experiments to small numbers of parasites during a fraction of the life cycle. Species that infect humans cannot be easily studied in standard model hosts such as mice. Also, since Cryptosporidium spp. is intrinsically refractory to antifolate drugs, selection of genetically modified parasites using these drugs, as for Toxoplasma and Plasmodium spp., is not possible. Further hindering Cryptosporidium spp. genomic manipulation, transient transfection is 10 000-fold less efficient than that of T. gondii (Vinayak et al. Reference Vinayak, Pawlowic, Sateriale, Brooks, Studstill, Bar-Peled, Cipriano and Striepen2015). Genetic validation of potential drug targets for Cryptosporidium spp. is a key unmet need.

Toward this goal, CRISPR/Cas9 technology has proven again to be a powerful system even for Cryptosporidium spp. Boris Striepen's laboratory recently developed for the first time a protocol for transfecting C. parvum sporozoites in tissue culture and isolation of stable genetically modified parasites (Vinayak et al. Reference Vinayak, Pawlowic, Sateriale, Brooks, Studstill, Bar-Peled, Cipriano and Striepen2015). Notwithstanding a 10-fold optimization of transfection, the low efficiency still required the use of a highly sensitive nano-luciferase (nLuc) reporter. Sporozoite expression of nLuc was achieved using the strong enolase promoter, whilst the aminoglycoside antibiotic paromomycin was used as selection marker since it is effective in tissue culture and in immunocompromised mice (Theodos et al. Reference Theodos, Griffiths, D'Onfro, Fairfield and Tzipori1998). After electroporation, transfected sporozoites were directly introduced into the mouse intestine by surgery due to the low oral infection efficiency of this stage. Expression of sgRNA and the Cas9 nuclease was achieved using the C. parvum U6 and aldolase promoters, respectively. Since Cryptosporidium spp. lack NHEJ, similar to Plasmodium species, transgene integration is likely to require homologous recombination. Thus, in a series of elegant experiments, the Striepen group restored a dead version of the nLuc carrying a stop codon that ablated luciferase activity by using short double-stranded templates for repair. This demonstrated that genome editing through homologous recombination was also possible in the recalcitrant Cryptosporidium spp. The authors also achieved for the first time a gene knock out in this parasite by deleting the gene encoding thymidine kinase (TK). This genetic manipulation provided evidence of the non-essentiality of TK and its role as an alternative route for thymidine monophosphate synthesis, explaining why C. parvum tolerates high doses of antifolate drugs. The Striepen group also recently published a how-to guide for genetic manipulation of Cryptosporidium spp. that will be invaluable to the field (Pawlowic et al. Reference Pawlowic, Vinayak, Sateriale, Brooks and Striepen2017). In summary, an adaptation of CRISPR/Cas9 technology to this nearly intractable parasite allowed generation of the first Cryptosporidium knock out strain and, at the same time, deletion of the TK gene provided a new potential selection marker for genome manipulation. Overcoming such barriers for Cryptosporidium spp. opens new avenues to import the RNA- or protein-based regulatory strategies developed from other apicomplexans.