Studying Cryptosporidium Infection in 3D Tissue-derived Human Organoid Culture Systems by Microinjection

All tissue handling and resection was performed under Institutional Review Board (IRB) approved protocols with patient consent.

1. Preparation of C. parvum Oocysts for Injection

NOTE: Cryptosporidium oocysts were purchased from a commercial source (see the Table of Materials). These oocysts are produced in calves and are stored in phosphate-buffered saline (PBS) with antibiotics. They can be stored for about 3 months at 4 °C and should never be frozen. We normally use oocysts within one month. Organoids can be infected with either intact oocysts, or sporozoites may be isolated from excysted oocysts and used to infect organoids if it is important not to have oocysts carryover from the original inoculum.

1. Prepare Cryptosporidium oocysts for infecting cells (Figure 1A).
   1. Keep oocysts on ice throughout all manipulations until they are added to the organoids.
   2. Calculate the number of oocysts needed for a full six-well plate of organoids (usually about 5 x 10⁵–2.5 x 10⁵ for the plate). Count the numbers of oocysts in a hemocytometer to verify the quantity and transfer to a centrifuge tube.
      NOTE: To aid in visualization, oocysts may be mixed 1:1 with an oocyst-specific fluorescent antibody (see the Table of Materials) before being loaded onto the hemocytometer. The fluorophore-labeled oocysts can then be easily visualized and enumerated using a fluorescence microscope. We suggest injecting about 100–1,000 oocysts/organoid. In general, 1,000–2,000 organoids can be grown in a six-well plate.
   3. Bring the volume of the oocysts suspension up to 900 µL with PBS. Add 100 µL of sodium hypochlorite (e.g., Clorox) bleach (at 4 °C). Incubate for 10 min on ice.
   4. Centrifuge for 3 min in a microcentrifuge at 8,000 x g at 4 °C. Orient the tubes in the centrifuge with the cap opening facing inward. The pellet can be hard to see so knowing where the parasites have pelleted in the tube is essential.
   5. Remove the supernatant with a pipette being careful to avoid the pellet. Add 1 mL of Dulbecco's modified Eagle's medium (DMEM) and vortex to mix.
   6. Centrifuge for 3 min in a microcentrifuge at 8,000 x g at 4 °C.
   7. Repeat washes with DMEM two more times.
   8. Prepare expansion medium (OME) or differentiation organoid medium (OMD) to which taurocholate has been added to a final concentration of 0.5% (w/v) (See Table of Materials). Taurocholate should always be prepared and added fresh.
      NOTE: We have successfully used 0.5% taurocholate in our infection assays where the inoculum is intact oocysts, and saw improved rates of infection without deleterious effects on the host cells. However, taurocholate may have unanticipated effects on cells, and lower concentrations have been used successfully in infection assays¹⁶.
   9. Resuspend oocysts in 100 μL of organoid culture medium supplemented with 0.5% (w/v) sodium taurocholate. Count oocysts again as described in step 1.1.2.
   10. Add Fast Green dye to the suspension in order to visualize injection.
   11. Fill micro-loader tips (see the Table of Materials) with the oocyst suspension and use it to fill pulled capillaries.
       CAUTION: The whole procedure should be done in a tissue culture hood with level-2 safety protocols. Use of masks is recommended as Cryptosporidium oocysts can also be infectious when airborne.

2. 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 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 remaining oocysts and shells, filter the suspension through a 3 µm filter (Figure 1B). Use a 47 mm filter holder apparatus fitted with 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 cold room.
   8. Add 7.5 mL of the sporozoite suspension to the filter assembly and allow 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 buffer⁵ (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.

3. In Vitro Culture of Human Intestinal and Lung Organoids for Microinjection

1. Culture intestinal organoids under expansion and differentiation media conditions.
   NOTE: The details of intestinal and lung organoid propagation have been previously described in other articles^8,13 (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 expansion medium are preferable to those in grown differentiation media as there is less debris accumulation than that seen in organoids grown in 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 (expansion media) on top and refresh every day.
      NOTE: For lung organoids, we do not have separate expansion and differentiation media.
   2. To split and plate organoids for microinjection, remove media from the 6-well plate containing human organoids and 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 and lung 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. Lung organoids have a thicker outer membrane and therefore require stronger shearing with the glass pipette as compared to intestinal organoids. Moreover, they have a slower growth rate than intestinal organoids (up to 14 days between each passaging).
   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 the 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 for 5–6 additional days before injecting the parasites.
      NOTE: For expansion of organoids, it is recommended to plate the organoids densely. For microinjection, 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. Expansion of lung organoids is done is a similar manner but using lung organoid specific media components (Table 1)⁸ .

4. 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 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.

5. Immunofluorescence Staining of Organoids

1. Collect organoids (1–2 x 24 wells) with a P1000 pipette into a 15 mL tube containing cold F12+++.
2. Pellet organoids at 300 x g for 2 min, remove the supernatant without disrupting the pellet, and gently pipet the pellet loose into the remaining volume.
3. Add 5 mL of 2% paraformaldehyde in PBS. To prevent the organoids from sticking to the wall do not invert the tube. Allow the organoids to settle to the bottom of the tube and fix at 4 °C overnight or 1 h at room temperature.
4. Remove the fixative and add 10 mL of permeabilization buffer (0.2% Triton in PBS).
5. Rotate the tube at room temperature for 20 min (this ensures that all the organoids remain in suspension).
6. Pellet the organoids at 300 x g for 2 min, and then discard the supernatant.
7. Resuspend the organoids gently in 500 µL of blocking solution (See the Table of Materials) and transfer to a 2 mL microcentrifuge tube.
8. Incubate for 20 min at room temperature on a shaker. Allow organoids to settle to the bottom of the tube by gravity. Replace the blocking solution with primary antibody solution (See Table of Materials) and incubate for 1–2 h at room temperature or overnight at 4 °C.
9. Wash 3x with PBS containing 0.1% Tween. Let the organoids settle every time and remove the supernatant.
10. Add secondary antibody solution (See the Table of Materials) and incubate for 2 h at room temperature.
11. Wash 3x with PBS containing 0.1% Tween. Leave 50 μL of PBS after the third wash.
12. Mount on the slide by pipetting the organoids suspended in 50 μL of PBS on the slide. Remove excess PBS, add a drop of mounting agent (See the Table of Materials) and add the coverslip on top. Seal the sides with nail polish and let it dry.

6. Isolation of Oocysts from Organoids

1. Isolate newly formed oocysts from the organoid lumen.
   NOTE: Oocysts are isolated from the organoids by immunomagnetic separation using an oocyst isolation kit (see the Table of Materials) with the modifications described below. The isolated oocysts can then be analyzed by immunofluorescence and electron microscopy.
   1. Start with differentiated organoids that have been maintained in OMD for 5–7 days and that are uninfected, infected for 1 day and infected for 5 days. Use the first two as negative controls.
      NOTE: We have found that differentiated organoids produce more oocysts than lung or expanding intestinal organoids¹⁰.
   2. Collect organoids into 15 mL centrifuge tubes. Centrifuge the organoids for 20 min at 3,000 x g and 10 °C.
      NOTE: This high speed is needed to make sure no oocysts are lost out of any organoids that may be broken.
   3. Remove the organoid media and replace it with 5 mL of water.
   4. Disrupt the organoids by repeated vigorous pipetting with a fire-polished glass Pasteur pipette.
   5. If clumps are visible, transfer the organoid suspension to a glass dounce homogenizer, and homogenize until organoids are well disrupted. The dounce homogenizer will not affect the oocysts.
   6. Once there are no visible clumps, add 5 mL of buffer A from the oocyst isolation kit. Mix and then add 120 µL of the magnetic beads coated with anti-oocyst IgM.
   7. Incubate the cell suspension and magnetic beads for 2 h at room temperature with continuous mixing on a rocker platform.
   8. At the end of the incubation, place the tubes containing cells and beads on a magnetic separation rack designed for 15 mL tubes.
   9. Rotate the tubes in magnetic separation rack manually for 3 min. The beads will adhere to the side of the tube next to the magnet.
   10. Carefully, with a 10 mL pipette, remove the supernatant from the beads. Resuspend the beads in 450 µL of Buffer B and transfer to a 1.5 mL microcentrifuge tube.
       NOTE: Keep the supernatant until the isolation of the oocysts is confirmed.
   11. To collect any remaining beads and oocysts, wash the 15 mL tube with 450 µL of Buffer B and add this wash to the magnetic beads in the microcentrifuge tube.
   12. Repeat step 6.1.11 one more time. All beads and captured oocysts should now be transferred to the microcentrifuge tube.
   13. Place the microcentrifuge tube on a magnetic separation rack designed for holding microcentrifuge tubes.
   14. Rotate the tube in the magnetic separation rack by hand for 3 min.
   15. Carefully remove the supernatant with a pipette into a new tube.
       NOTE: Keep the supernatant until the isolation of the oocysts is confirmed.
   16. Remove the microcentrifuge tube containing magnetic beads and oocysts from the magnetic separation rack.
   17. Add 100 µL of 0.1 N HCl to magnetic beads to elute the oocysts off the beads. Vortex for 30 s.
       NOTE: Vortexer should be set to slightly less than maximum speed.
   18. Incubate the beads in 0.1 N HCl for 10 min at room temperature.
   19. Vortex again. Then place the tube back on magnetic separation rack. Wait for beads to adhere to the side of the tube and then transfer the supernatant to a new microcentrifuge tube.
   20. Repeat steps 6.1.17 through 6.1.19 and combine the second eluate with the first elution.
   21. Neutralize the eluate with 20 µL of 1 N NaOH, or another neutralizing buffer such as 1 M Tris, pH 8.
   22. To count oocysts, take 10 µL of the eluate, combine it with 10 µL of oocyst-specific antibody (See Table of Materials) and count fluorescent oocysts on a hemocytometer.
       NOTE: Isolated oocysts can be stored at 4 °C or used immediately for immunofluorescence or electron microscopy imaging.