Genetic Manipulation in Δku80 Strains for Functional Genomic Analysis of Toxoplasma gondii

1. Gene Targeting to Delete a Gene of Interest

This protocol is designed for the efficient generation of a mutant strain that has a targeted gene deletion at a defined genetic locus. The previously described T. gondii hypoxanthine-xanthine-guanine phosphoribosyltransferase (HXGPRT) selectable marker is used in this protocol for safe and reliable marker insertion following mycophenolic acid and xanthine (MPA + X) selection^14-16. This protocol uses a validated and cost-effective method for constructing DNA targeting molecules based on yeast recombinational cloning^17,18. Several alternative protocols are commercially available for constructing recombinant targeting molecules by using bacterial recombinational cloning methods, in vitro recombinational cloning methods, or fusion PCR. The protocol described below provides the highest overall efficiency and reliability of recovering cloned parasites possessing a targeted gene deletion in Δku80 strains of T. gondii.

1.1 Preparation of targeting DNA

1. Access ToxoDB⁶ (http://www.toxodb.org) to obtain genomic sequences for the gene of interest (GOI).

Note: Genomic sequences should be obtained from the Type I, II or III genome sequence in the ToxoDB database that is closest to the strain being manipulated. For example: GT1 for RH and ME49 for Pru.

2. Design specific overlap primers that amplify the 5' and 3' genomic targeting flanks of the gene of interest (GOI) incorporating a 33 bp overlap with the yeast shuttle vector pRS416 (or alternatively pRS426) and incorporate a 33 bp overlap with the selectable marker (HXGPRT) (Figures 1A-B). Add a unique restriction enzyme site in each primer at both ends of the 5' and 3' genomic targeting flanks, placed between the 33 bp overlap and the T. gondii-specific primer sequence(s) (Figures 1A-B).

Note: A greater than 30 bp overlap is required for efficient yeast recombination cloning. The 5' and 3' genomic targeting flanks are designed to amplify specific DNA fragments between 800 and 1,400 bp that define a targeted deletion. Be aware that shorter flanks will significantly reduce targeting efficiency in the Δku80Δhxgprt background².

3. PCR amplify the 5' target flank using primers F1 and R1, and PCR amplify the 3' target flank using primer F2 and R2 (Figures 1A-C) from genomic DNA³. Separately, PCR amplify the ~2 Kbp HXPGRT gene including the associated 5' and 3' dhfr flanking regions of the HXGPRT cDNA pminiHXGPRT cassette¹⁴ using primers HXF and HXR (Figures 1B-C).

Note: Amplify target flanks using genomic DNA from the parental strain to be transfected.

Note: Place the HXGPRT marker in the forward orientation relative to the 5' and 3' DNA targeting flanks to facilitate subsequent efficient genetic manipulations, such as the targeted removal of HXGPRT from the disrupted locus.

4. Verify correct PCR product sizes by agarose gel electrophoresis and estimate the DNA fragment concentration using agarose gel standards or another method for determining DNA concentration.
5. Generate competent yeast aliquots as described in Gietz and Schiestly¹⁷.
6. Combine 50 ng of 5' genomic targeting flank, 50 ng of 3' genomic targeting flank, 100 ng of HXGPRT selection marker, and 50 ng of linearized shuttle vector with sterile H₂O to achieve a final volume of 10 – 20 μl. Transform competent yeast with the three PCR products and yeast-E. coli shuttle vector for recombinational cloning using the protocol described by Gietz and Schiestly¹⁷. Plate transformed yeast on uracil-minus minimal medium agar plates and incubate at 30 °C for 2 – 3 days.

Note: The ordered assembly of the plasmid, the 5' target flank, the HXGPRT marker, and the 3' target flank is facilitated by the engineered 33 bp overlap between the DNA fragments (Figures 1A-B) and is mediated by homologous recombination in yeast.

7. Harvest yeast by adding 2 ml of 2x YPAD to the uracil-minus plates and gently scraping to dislodge colonies without disrupting the agar. Centrifuge the yeast solution for 5 min at 5,000 x g and discard the supernatant.
8. Resuspend the yeast pellet in 250 μl of a resuspension buffer containing RNaseA and add 50 – 100 μl of acid-washed glass beads to the solution. Vortex for 5 min at the highest speed to smash the yeast cells.
9. Allow the glass beads to settle to the bottom of the tube and transfer the supernatant to a 2 ml Eppendorf tube.
10. Isolate yeast DNA using a miniprep DNA isolation kit, resuspending in a final volume of ~100 μl.
11. Mix 2 μl yeast DNA (~50 ng) with 40 μl of DH10B electroporation competent E. coli kept on ice.
12. Transfer the solution to a chilled 2 mm gap electroporation cuvette and electroporate the bacterial cells at 2.4 kV and 129 Ω.
13. Rescue cells by adding 0.8 ml SOC broth to each cuvette and transfer solution to a 14 ml snap-cap Falcon tube. Incubate cells at 37 °C on a roller drum for 40 – 60 min.
14. Plate E. coli on 2XYT + ampicillin (AMP) plates and incubate the plates at 37 °C overnight.
15. Select ~6 – 8 single colonies and grow in 3 ml 2XYT + AMP for ~16 hr at 37 °C.
16. Prepare a 30% glycerol freezer stock of each E. coli clone.
17. Isolate plasmid pΔGOI from E. coli clones using a miniprep kit, resuspending in a final volume of ~100 μl.
18. Validate the pΔGOI using restriction enzyme digests to measure the DNA plasmid size and verify the expected pΔGOI DNA banding patterns.
19. Finalize validation of pΔGOI by DNA sequencing of the 5' and 3' genomic targeting flanks to verify 100% DNA sequence based on genome sequence data in http://www.ToxoDB.org.

Note: Near perfect sequence homology is essential for maximizing gene targeting efficiency³ since each base pair difference constitutes a corresponding cut in the length of perfect homology.

Note: The genome sequence of the type II Δku80 strain based on the Pru parental strain is not available at this time. This work uses the type II ME49 genome sequence as the surrogate genome for the type II Δku80 strain. Based on sequence data at a number of genetic loci, it is estimated that the ME49 genome and the Pru genome exhibit single nucleotide polymorphisms at a frequency of ~1 per 10,000 nucleotides, or less.

20. Generate >200 μg of pΔGOI stock by inoculating a ~250 ml 2XYT + AMP overnight culture with glycerol stocks from a validated E. coli miniprep clone.
21. Isolate pΔGOI plasmid DNA from the large culture using a maxiprep DNA isolation kit, resuspending in a final volume of ~1,000 μl.
22. Repeat restriction enzyme digests, or DNA sequencing of the pΔGOI maxiprep DNA to verify the correct targeting DNA molecule prior to transfection.
23. Linearize ~15 μg pΔGOI at the 5' end using the unique restriction enzyme X (RE.X) digest site built into the 5' target flank (Figures 1B).

Note: Gene targeting in T. gondii requires a minimum of ~10 μg of targeting DNA to obtain an efficient frequency of targeting events at the gene locus of interest².

24. Validate linearization of pΔGOI and determine the DNA concentration by agarose gel electrophoresis or an alternative method.

Note: If restriction enzyme digestion at the 5' flank does not yield complete linearization, the designed 3' unique RE.Y digest site can be used as an alternative site for linearization of pΔGOI.

Note: A completely linearized targeting DNA molecule is essential for successful gene targeting by homologous recombination in the Δku80 strains.

25. Neutralize the restriction enzyme by incubating at 68 °C for 15 – 20 min.
26. Prepare at least 15 μg of linearized plasmid in 100 μl of sterile H₂O. Store linearized pΔGOI at -20 °C until the time of transfection.

1.2 Preparation of T. gondii parasites, transfection, selection, subcloning, validation, archival and maintenance of genetically manipulated strains

General methods for the culture and manipulation of T. gondii in human foreskin fibroblast (HFF) cells are described^19-21. The Δku80Δhxgprt strains replicate normally in parasite infection medium (Eagles Minimal Essential Medium (EMEM) growth medium supplemented with 1% fetal bovine serum (FBS) and 1% Antimycotic-Antibiotic diluted from a 100x stock)^1,2. All work with Toxoplasma gondii must be performed using biosafety level 2 procedures. The T. gondii RHΔku80² parental strain was generated from the RHΔhxgprt strain from the Roos Lab¹⁴. The PruΔku80¹ parental strain was generated from PruΔhxgprt (Prugniaud strain BSG-4²²) that contains a stably integrated CAT selectable marker and a green fluorescent protein (GFP) reporter under the control of the bradyzoite stage specific promoter LDH2.

1. Inoculate a 25 cm² flask containing confluent HFF cells with 1 x 10⁶ viable type I RH Δku80Δhxgprt parasites, or inoculate two 25 cm² flasks with 2 x 10⁶ viable type II Prugniaud (Pru) Δku80Δhxgprt parasites.

Note: Viable parasites are defined as extracellular parasites that have freshly lysed out.

Note: This scale provides a sufficient number of viable parasites for three transfection experiments.

2. Inspect the Δku80Δhxgprt infected cultures ~68 – 72 hr post-infection. Verify the presence of viable parasites and ~90 – 95% lysis of HFF cells to plan the precise timing of transfection.

Note: The isolation of freshly egressed parasites is essential for the success of this protocol because low parasite viability will abolish gene-targeting efficiency.

3. Agitate viable parasites into solution by closing the plug-seal lid on the 25 cm² flask and vigorously shake the flask back and forth to agitate the parasites off of the growth surface without splashing liquid onto the inside of the lid or the neck of the flask.

Note: Parasite harvest does not require a syringe-needle release protocol for the type I or type II Δku80 strains.

4. Transfer the parasite solution to a 10 ml syringe attached to a filter holder containing a 3 μm membrane and place the filter unit on top of an open 15 ml screw-cap tube. Syringe filter the parasites into the 15 ml screw-cap tube.

Note: Filtering the parasites through the membrane removes infected cells and cellular debris.

5. Determine the parasite concentration (tachyzoite forms per ml) using a hemocytometer.

Note: Optional: Using the parasite solution, set up a plaque forming unit (PFU) assay²¹ that will be read 7 – 8 days later to determine adequate viability of the transfected parasites (PFU to tachyzoite ratio of ≥0.2).

6. Pellet parasites for 7 min at 1400 x g and aspirate supernatant to ~0.4 ml without disturbing the parasite pellet. Spin for 2 min at 1,400 x g and aspirate remaining supernatant to ~0.01 ml without disturbing the parasite pellet.
7. Resuspend the parasite pellet by flicking the bottom of the tube and immediately add cytomix²³ buffer (1.33x concentration) to the parasite pellet to obtain a parasite concentration of 4 – 5 x 10⁷ parasites/ml cytomix.

Note: Omit the addition of ATP and glutathione to cytomix since these additions do not improve the efficiency of gene targeting.

8. Thaw the 100 μl aliquot of the linearized pΔGOI from protocol 1.1 (step 26).
9. Transfer 0.3 ml of the parasite cytomix solution (~1.3 – 1.6 x 10⁷ parasites) to the 100 μl of linearized pΔGOI from protocol 1.1 (step 26), and immediately transfer the entire 0.4 ml parasite + pΔGOI mix to a chilled 2 mm gap electroporation cuvette.
10. Electroporate the parasites at 1.4 kV and 24 Ω.
11. Rest the transfection cuvette at room temperature for ~5 min then transfer the entire contents of the transfection cuvette into a 150 cm² flask containing confluent HFF cells with 30 ml infection medium and incubate the infected culture overnight.
12. Begin selection in mycophenolic acid + xanthine (MPA + X) ~20 hr post-transfection (Figure 2A) by replacing the infection medium with MPA + X selection medium (MPA (25 μg/ml) and xanthine (50 μg/ml) in infection medium).

Note: Do not disturb the incubating MPA + X selection flask.

13. Estimate the total number of PFUs in the MPA + X selection flask ~8 days post-transfection by visually inspecting the monolayer. Verify the presence of developing PFUs and healthy zones of infection by light microscopy.

Note: If plaques are not visible by day 8, maintain the selection and monitor for signs of infection since mutant strains may have a reduced replication rate.

Note: The Δku80Δhxgprt parasite strains have an extremely low background of undesired nontargeted events, which allows for the successful isolation of mutant strains with quite severe, but not lethal growth defects. If a gene is essential and it cannot be deleted, the primary transfection, or subsequently passed parasite population, may reveal a phenotype where the parasites cease to replicate during the selection as the population resolves to targeted knockouts.

14. Shake the flask as in step 3 to dislodge extracellular parasites from the monolayer [after visible plaques have developed] to generate a parasite solution. Transfer ~0.5 ml of the parasite solution from the MPA + X selection flask to a 25 cm² MPA + X selection flask containing confluent HFF cells (designate this culture pass 1A).
15. Dislodge parasites in the original 150 cm² MPA + X selection flask ~3 days later and transfer ~0.5 ml parasite solution to a second 25 cm² MPA + X selection flask containing confluent HFF cells (designate this culture pass 1B).
16. Allow zones of infection to develop in pass 1A and/or pass 1B flasks for ~5 days. Visually verify by light microscopy that pass 1A and 1B flasks contain a minimum of 25 viable PFUs with healthy zones of infection.
17. Choose either the pass 1A or pass 1B flask for continued passage in MPA + X selection medium in 25 cm² flasks. Maintain passage under MPA + X selection.

Note: Parasites must be cultured for a sufficient number of generations to delete nonintegrated persisting episomes from the population prior to subcloning. Do not perform subcloning prior to 20 days post-transfection for type I RH Δku80Δhxgprt, or prior to 25 days post-transfection for type II Pru Δku80Δhxgprt parasites to allow sufficient time to dilute nonintegrated episomes^1,2.

18. Subclone parasites in a 96-well tray containing confluent HFF cells with MPA + X selection medium. Prepare one tray at a concentration of 1 parasite/well and a second tray at a concentration of 2 parasites/well.

Note: Subclone parasites that have recently lysed (~90 – 95% of the HFF cells in the 25 cm² flask) to ensure that the parasites have high viability.

Note: The optimal time frame post-transfection for subcloning was established by measuring how rapidly successfully targeted parasites arise at a high frequency in the selected population¹.

19. Continue to passage the primary population of transfected parasites in MPA + X selection medium using a weekly passage schedule of infecting a 25 cm² HFF flask with 10 μl of the parasite solution.

Note: If clones carrying the targeted gene deletion (knockouts) are not obtained from the first subcloning in step 18, resubclone the parasites from the continuously maintained population ~10 – 12 weeks post-transfection.

Note: One of the reasons a gene deletion is not obtained arises from the episomal persistence of some pΔGOI plasmids. Episomal persistence is determined by the sequences contained in the 5' and the 3' genomic targeting flanks and does not appear to arise from the HXGPRT genetic element. Approximately 5 – 10% of targeting plasmids exhibit significant episomal persistence that necessitates a later time of subcloning to allow the parasite population a sufficient number of generation times to dilute the persisting episomes and resolve the population to primarily stable integrants.

20. Score the 96-well tray 6 – 7 days post-subcloning (type I parasites) or 7 – 8 days post-subcloning (type II parasites) for wells that contain a single PFU, identified as a single zone of infection in light microscopy at either 40X or 60X power. Mark a small dot on the 96-well tray lid to designate the location of the PFU in the well.
21. Mix the contents of each well containing a single PFU using a pipette set at 50 μl (200 μl tip) by directing fluid flow over the PFU to disperse parasites in the well.

Note: Mixing the well accelerates parasite lysis of the HFF monolayer. Type I RH strains will lyse the well ~4 days after mixing, type II Pru parasites will lyse the well ~5 days after mixing.

22. Select a dozen lysed wells (clones) and scratch across the bottom of each of these lysed wells using a 0.5 – 10 μl pipette tip while simultaneously drawing-up 6 μl of parasite solution. Transfer the 6 μl of parasite solution to a well in a 24-well tray containing confluent HFF cells in 1 ml MPA + X selection medium.

Note: It is essential to scratch the bottom of the well to keep the opening bore of the pipette tip in constant contact with parasites residing at the bottom of the well.

23. Monitor the 24-well tray for lysis of the HFF cells.

Note: Type I RH parasites will typically lyse the well in the 24-well format in ~4 days. Type II Pru parasites will typically lyse the well in ~5 days.

24. Transfer 2 μl of viable parasite solution scratched from the bottom of each well in the 24-well format to the corresponding well of a new 24-well tray containing confluent HFF cells and 1 ml MPA+X selection medium per well.

Note: All parasites clones and lines can be continuously maintained by passing 2 μl of viable parasite solution every 7 days in this 24-well tray format.

25. Verify that parasites were successfully transferred to a new 24-well tray by visually inspecting with light microscopy ~18 hr post-passage.
26. Harvest parasites from the lysed wells of the 24-well tray from step 23 – 24 using either a 1 ml or 10 ml pipette and transfer the parasite solution to an Eppendorf tube.
27. Pellet the parasites at 1,400 x g for 7 min, rinse once in 1 ml phosphate buffered saline (PBS), and pellet again at 1,400 x g for 3 min.
28. Aspirate PBS from the pellet, then add 200 μl of PBS to the pellet to resuspend the parasites. Freeze the parasite solution at -80 °C until DNA isolation.
29. Isolate parasite DNA from each clone using a tissue DNA isolation minikit.
30. Validate the targeted gene deletion (knockout) by PCR using validation primers (Figures 2A-C). Test for the absence of the functional coding region of the gene of interest and using the upstream CxF and downstream CxR genomic DNA primers (Figures 2A-B) test for the presence of PCR products that span the genomic targeting flanks and HXGPRT to demonstrate correct 5' and 3' targeted integration of the HXGPRT gene into the deleted gene locus.

Note: Primers for the validation must be unique to the gene of interest or a false positive result can be obtained. Primer design can be validated on http://www.toxodb.org by blasting primer sequences to the genome(s) to verify primers are unique.

31. Passage parasite clones on a weekly schedule as described in step 24. Once the targeted deletion has been validated, continuing selection in MPA+X is optional.
32. Archive parasite clones by preparing freezer stocks. Pellet extracellular parasites from lysed HFF cells in a 25 cm² culture flask, remove media and gently resuspend parasite pellet in cell culture freezing medium at a parasite concentration >4 x 10⁷ parasites per ml. Transfer aliquots to cryovials and store parasites indefinitely in liquid nitrogen, or at -80 °C.

2. Deletion of HXGPRT

This protocol is designed for removing the HXGPRT marker from its integration site in the genome of a genetically manipulated Δku80 strain. Removal of HXGPRT allows for marker recovery and the generation of strains with multiple genetic manipulations using only selections based on HXGPRT^1-3. While the protocol detailed below describes the method to re-target a locus to delete HXGPRT, it should be noted that removal of HXGPRT at the gene locus of interest also permits simultaneous re-integration of the wild-type gene (complementation), re-integration of a mutant gene, as well as re-integration of a tagged gene (N- or C-terminal GFP, HA tag, etc,), allowing for a variety of genetic manipulations. The mechanisms of action²⁴ and targeting protocols using 6-thioxanthine selections have been previously described^1-3,15.

2.1 Delete HXGPRT

1. Excise the HXGPRT selectable marker from pΔGOI by digesting with RE.Z to cut at the unique restriction enzyme sites flanking the HXGPRT (Figure 1B).
2. Verify complete digestion of DNA by agarose gel electrophoresis. Isolate the larger of the two bands from the agarose gel.

Note: The larger of the two bands contains the plasmid along with the 5' and 3' DNA target flanks, but not the HXGPRT gene.

3. Place the agarose section containing the larger DNA band in a spin column laying the gel flat on the membrane. Spin the column at 13,000 x g for 4 min at RT. Add sterile H₂O to the flow-through to bring the volume to 100 μl.

Note: Other commercial methods are available to isolate DNA from agarose.

4. Concentrate DNA by EtOH precipitation, resuspending the DNA in a final volume of 18 μl sterile H₂O.
5. Mix 1 μl of concentrated DNA with 7 ml H₂O, 1 μl 10x ligation buffer and 1 μl T4 DNA ligase (5 units) and place the reaction at 4 °C overnight to generate pΔGOIc (pΔGOIclean) (Figure 3A).

Note: DNA fragments with RE.Z containing DNA ends can be designed and included at this step to create plasmids suitable for targeted re-integration of the wild-type gene (complementation), targeted re-integration of a mutant gene as well as targeted re-integration of a tagged gene (N- or C-terminal GFP, HA tag, etc.).

6. Dilute the reaction 2x with sterile H₂O prior to electroporation.
7. Transform pΔGOIc into E. coli as in protocol 1.1 to subclone, isolate, and validate the targeting plasmid. Then linearize pΔGOIc in preparation for transfection (see steps 1.1.11 to 1.1.26).
8. Repeat the parasite transfection protocol as outlined in protocol 1.2 (steps 1 to 11).

Note: Prior to conducting steps 1 to 8, verify that targeted removal of HXGPRT is feasible in the mutant strain. Infect a 150 cm² flask of confluent HFF cells with 1 x 10⁶ tachyzoites of the mutant strain using 6-thioxanthine (6TX) selection medium (200 μg/ml 6TX in infection medium) and place the flask in a PFU assay. Inspect the flask 8 days later to verify that no (or very few; <10) PFU are visible, which is essential to establish that the potential gene targeting efficiency will exceed any potential spontaneous reversion of the mutant strain to a phenotype with reduced HXGPRT expression (a 6TX resistance phenotype).

Note: Approximately 5 – 10 % of HXGPRT integration sites exhibit a significant and spontaneous frequency of 6TX resistance even though the HXGPRT marker is still integrated at the targeted site (see Representative Results section for explanation).

9. Begin 6TX selection ~20 hr post-transfection by changing the medium in the 150 cm² flask to 6TX selection medium.

Note: Do not disturb the flask for ~10 days post-transfection.

10. Inspect the flask for PFU formation at day 10 – 12 post-transfection (type I parasites) or day 10 – 16 post-transfection (type II parasites).

Note: Parasites being targeted for the deletion of HXGPRT will not begin to grow under 6TX selection until the HXGPRT gene and mRNA is deleted, and the residual HXGPRT protein is inactivated.

11. Shake the selection flask to create a parasite solution, and transfer 0.5 – 1.0 ml of the parasite solution to a new 25 cm² flask containing confluent HFF cells and 5 ml 6TX selection medium. Repeat the transfer from the primary 150 cm² to new 25 cm² flasks once a day for two more days.

Note: Multiple sampling increases the chances of capturing viable targeted parasites that have lost the HXGPRT gene.

12. Inspect the 25 cm² flasks ~5 days after infection to verify the presence of developing PFUs with zones of healthy replicating parasites. Pick one or two flasks containing zones of infection.

Note: Parasites deleted of the HXGPRT gene replicate at a normal growth rate in 6TX selection.

13. Continue to passage the parasites in 6TX selection medium.
14. Subclone the parasite population after 25 – 30 days of selection. Set up one 96-well tray with ~ 1 parasite/well and another with ~ 2 parasites/well.
15. Prepare parasite DNA from isolated clones and maintain clones in a 24-well format culture according to protocol 1.2 (steps 19 to 29).
16. Validate deletion of HXGPRT by PCR using the strategy outlined in Figures 3A-B.

3. C-terminal Tagging of Proteins

This protocol is designed for C-terminal tagging of proteins by targeting the tag for integration via double cross over homologous recombination at the genomic locus of the gene using MPA + X selection^2,4. This protocol works efficiently because the 5' dhfr sequence of the HXGPRT marker is a fully validated functional 3' untranslated region for other genes^2,4.

3.1 Direct C-terminal tagging of proteins at endogenous genetic loci

1. Create targeting pΔGOItag plasmid construct using yeast recombinational cloning (Figure 4) and methods described in protocol 1.1. The 5' genomic targeting flank contains the last 800 to 1,200 bp of coding region (or genomic DNA) of the GOI, except the termination codon is moved to a position 3' to the tag of choice (HA tag, Myc tag, His tag, etc.)
2. Create the targeted insertion of the C-terminal tag at the endogenous locus of the protein-coding gene by following the steps in protocol 1.1 and 1.2 using the strategy outlined (Figure 4).
3. Verify the insertion of the C-terminal tag at the endogenous gene locus using a PCR strategy.
4. Retarget the gene locus using methods described in protocol 1 and protocol 2 to delete the HXGPRT selectable marker to create a precisely regulated endogenous gene locus that expresses a tagged protein. This protocol also recovers the HXGPRT selectable marker that can be used again to target another locus in the tagged strain (see protocol 1).