A Technique to Develop a Parasite Infection in Intestinal Organoids via Microinjection

1. In Vitro Purification of Sporozoites from C. parvum Oocysts

1. Purify sporozoites from C. parvum oocysts after bleaching and washing out the bleach as described above.
   1. Transfer the oocysts to a 15 mL tube. Resuspend oocysts in room temperature excystation medium (0.75% w/v sodium taurocholate in DMEM) to obtain 1 x 10⁷ oocysts/mL. The addition of taurocholate improves the excystation rate of the oocysts, improving sporozoite yield.
   2. Incubate oocyst suspension at 37 °C for 1–1.5 h.
   3. Check the sample microscopically for the extent of excystation; 60–80% excystation is reasonable for good recovery of sporozoites. If the level of excystation is low, incubate longer (another 30 min to 1 h).
   4. Determine the percent excystation relative to the number of starting oocysts. Excystation is calculated as:
      % excystation = [1 – (number of intact oocysts/number of oocysts at start)] x 100
   5. Wash cells to remove excystation reagents by adding 14 mL of PBS or medium, mixing, and recovering cells (intact oocysts, oocyst shells, and sporozoites) by centrifugation at 3,400 x g for 20 min to recover sporozoites. Aspirate carefully to avoid losing cells.
   6. Resuspend the sporozoite pellet in 1–2 mL of DMEM to obtain 3 x 10⁷ oocysts/mL (based on the number of starting oocysts).
   7. To remove the remaining oocysts and shells, filter the suspension through a 3 µm filter (Figure 1B). Use a 47 mm filter holder apparatus fitted with a polycarbonate filter (3 µm pore size) attached to a 10 mL syringe barrel. Place the filter holder apparatus on top of a 15 mL tube. Place the assembly in an ice bucket or in a cold room.
   8. Add 7.5 mL of the sporozoite suspension to the filter assembly and allow it to filter through by gravity. Wash through with another 7.5 mL of DMEM.
      NOTE: To ensure success in sporozoite isolation fresh oocysts and good excystation are critical. If there are too many unexcysted oocysts, the suspension will not flow through by gravity. Applying pressure on the syringe can force unexcysted oocysts through. Microinjection of sporozoites is more challenging than that of oocysts because sporozoites may clump and block the capillary. To avoid this, we recommend making a wider capillary tip when injecting organoids with sporozoites. To achieve sufficient levels of infection, 2–4 times the number of sporozoites need to be injected into each organoid as compared to organoids infected with oocysts.
   9. Centrifuge the filtered sporozoite suspension at 3,400 x g using a swinging bucket rotor for 20 min to pellet sporozoites.
   10. Resuspend in 50–100 μL of OME or OMD organoid culture medium (see the Table of Materials) supplemented with 0.05% (w/v) Fast Green dye and L-glutathione, betaine, L-cysteine, linoleic acid and taurine-containing reducing buffer (see the Table of Materials).
       NOTE: Incubating oocysts for too long may result in the lysis of sporozoites and poor recovery and therefore should be avoided.

2. In Vitro Culture of Human Intestinal for Microinjection

1. Culture intestinal organoids under expansion and differentiation media conditions.
   NOTE: The details of intestinal propagation have been previously described in other articles (see Table of Materials for media recipes). Here, we briefly describe the organoid culture method with specific reference to optimization for Cryptosporidium injection and growth. We have found that for imaging of parasites in organoids, organoids grown in an expansion medium are preferable to those grown in differentiation media as there is less debris accumulation than that seen in organoids grown in a differentiation medium. However, if the goal is to isolate oocysts, organoids grown in differentiation media produce far higher numbers of oocysts.
   1. Maintain organoids in 3D cultures in extracellular matrix (see the Table of Materials) at 37 °C. Add OME (organoid expansion media) on top and refresh every day.
   2. To split and plate organoids for microinjection, remove media from the 6-well plate containing human organoids add F12+++ (See the Table of Materials) to the well, and break up the matrix by pipetting with a 1 mL pipette tip several times. Collect cells into a 15 mL tube (2 mL of F12+++ per tube is enough for further procedures).
   3. Add 10–12 mL of F12+++ into another 15 mL tube and place a fire-polished glass pipet into the medium, pipette up and down 3 times to break up the human intestinal organoids.
      NOTE: Use a long glass pipet (20–30 cm) and fire-polish it briefly. Do not make the opening (1 mm diameter) very small because organoids can be damaged. Make the tip of the pipet smooth by briefly fire-polishing it. Break organoids into smaller pieces of ~50 µm.
   4. Add F12+++ up to 5–7 mL and centrifuge at 350 x g for 5 min.
      NOTE: The centrifugation speed in this step is higher than normal in order to make a good cell pellet that is well separated from the extracellular matrix (see Table of Materials). We have observed that compared to mouse small intestine organoids, human small intestine organoids are harder to disrupt.
   5. Remove as much medium as possible without disturbing the cells, then resuspend the pellet with matrix maintained at 4 °C; 200–300 µL of matrix per well of a six-well plate is required. Organoids should be split one in three to maintain a fairly high cell density.
   6. Plate the organoids in matrix droplets of about 5–10 µL each in the well of a six-well plate. Incubate for 20–30 min at 37 °C and then add expansion medium (OME) on top.
   7. Change the medium every 2–3 days.
      NOTE: In about 5–7 days, organoids growing in EM reach a size of 100–200 μm and are ready for injection.
   8. To differentiate the organoids, after 5–6 days in EM, change the media to differentiation media (DM) conditions and keep it for 5–6 additional days before injecting the parasites.
      NOTE: For expansion of organoids, it is recommended to plate the organoids densely. For microinjection, the use of a six-well plate is recommended with organoids plated at a lesser density. For example, plate organoids from three wells of a six-well plate into a full six-well plate for microinjections. Matrix should be maintained at -20 °C for long-term storage and thawed at 4 °C or on ice before use.

3. Microinjection of Oocysts/Sporozoites into the Organoid Lumen

1. Microinject parasites into the apical side of the 3D organoid (Figure 2).
   1. Prepare glass capillaries of 1 mm diameter using a micropipette puller.
      NOTE: Settings used on the micropipette puller (See Table of Materials) are Heat = 663, Pull = 100, Velocity = 200, Time = 40 ms. Settings will need to be adjusted according to the user instructions for a particular machine.
   2. Cut the tip of the capillary with forceps. The size/diameter of the capillary end measures about 9–12 μm; this enables easy flow of oocysts (4–5 μm size).
   3. Fill capillaries with oocyst or sporozoite suspension using micro-loader tips.
   4. Load the oocyst-filled capillary onto a microinjector.
   5. Microinject 100–200 nL suspension into each organoid under an inverted microscope at 5x magnification, keeping the pressure constant. After microinjection, refresh media with OME or OMD (organoid differentiation medium) every day and maintain the plate at 37 °C.
      NOTE: We do not use a micromanipulator for microinjection. Use of the same capillary is recommended for the entire experiment to ensure equal injection in every sample.