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

Immunology and Infection

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We describe protocols to prepare oocysts and purify sporozoites for studying infection of human intestinal and airway organoids by Cryptosporidium parvum. We demonstrate the procedures for microinjection of parasites into the intestinal organoid lumen and immunostaining of organoids. Finally, we describe the isolation of generated oocysts from the organoids.

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Dutta, D., Heo, I., O'Connor, R. Studying Cryptosporidium Infection in 3D Tissue-derived Human Organoid Culture Systems by Microinjection. J. Vis. Exp. (151), e59610, doi:10.3791/59610 (2019).


Cryptosporidium parvum is one of the major causes of human diarrheal disease. To understand the pathology of the parasite and develop efficient drugs, an in vitro culture system that recapitulates the conditions in the host is needed. Organoids, which closely resemble the tissues of their origin, are ideal for studying host-parasite interactions. Organoids are three-dimensional (3D) tissue-derived structures which are derived from adult stem cells and grow in culture for extended periods of time without undergoing any genetic aberration or transformation. They have well defined polarity with both apical and basolateral surfaces. Organoids have various applications in drug testing, bio banking, and disease modeling and host-microbe interaction studies. Here we present a step-by-step protocol of how to prepare the oocysts and sporozoites of Cryptosporidium for infecting human intestinal and airway organoids. We then demonstrate how microinjection can be used to inject the microbes into the organoid lumen. There are three major methods by which organoids can be used for host-microbe interaction studies—microinjection, mechanical shearing and plating, and by making monolayers. Microinjection enables maintenance of the 3D structure and allows for precise control of parasite volumes and direct apical side contact for the microbes. We provide details for optimal growth of organoids for either imaging or oocyst production. Finally, we also demonstrate how the newly generated oocysts can be isolated from the organoid for further downstream processing and analysis.


Development of drugs or vaccines for treatment and prevention of Cryptosporidium infection has been hindered by the lack of in vitro systems that precisely mimic the in vivo situation in humans1,2. Many of the currently available systems either only allow short term infection (<5 days) or do not support the complete life cycle of the parasite3,4. Other systems which enable the complete development of the parasite are based on immortalized cell lines or cancer cell lines which do not faithfully recapitulate the physiological situation in humans5,6,7. Organoids or ‘mini-organs’ are 3D tissue derived structures which are grown in an extracellular matrix supplemented with various tissue specific growth factors. Organoids have been developed from various organs and tissues. They are genetically stable and recapitulate most functions of the organs of their origin, and can be maintained in culture for extended periods of time. We have developed a method for infecting human intestinal and lung organoids with Cryptosporidium that provides an accurate in vitro model for the study of host-parasite interactions relevant to intestinal and respiratory cryptosporidiosis8,9,10,11,12,13. In contrast to other published culture models, the organoid system is representative of real-life host parasite interactions, allows for completion of the life cycle so that all stages of the parasite life cycle can be studied, and maintains parasite propagation for up to 28 days10.

Cryptosporidium parvum is an apicomplexan parasite that infects the epithelium of the respiratory and intestinal tracts, causing prolonged diarrheal disease. The resistant environmental stage is the oocyst, found in contaminated food and water14. Once ingested or inhaled, the oocyst excysts and releases four sporozoites that attach to epithelial cells. Sporozoites glide on hosts cells and engage host cell receptors, but the parasite does not fully invade the cell, and appears to induce the host cell to engulf it15. The parasite, which is internalized within an intracellular but extracytoplasmic compartment, remains at the apical surface of the cell, replicating within a parasitophorous vacuole. It undergoes two rounds of asexual reproduction—a process called merogony. During merogony, type I meronts develop which contain eight merozoites that are released to invade new cells. These merozoites invade new cells to develop into type II meronts containing four merozoites. These merozoites, when released, infect cells and develop into macrogamonts and microgamonts. Microgametes are released and fertilize the macrogametes producing zygotes that mature into oocysts. Mature oocysts are subsequently released into the lumen. Oocysts are either thin-walled which immediately excyst to reinfect the epithelium, or thick walled which are released into the environment to infect the next host14. All stages of the Cryptosporidium life cycle have been identified in the organoid culture system previously developed by our group10.

Since human organoids faithfully replicate human tissues9,11,13, and support all replicative stages of Cryptosporidium10, they are the ideal tissue culture system to study Cryptosporidium biology and host-parasite interactions. Here we describe the procedures for infecting organoids with both Cryptosporidium oocysts and excysted sporozoites, and isolating the new oocysts produced in this tissue culture system.

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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.
    The details of intestinal and lung organoid propagation have been previously described in other articles8,13 (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.
    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.

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Representative Results

The protocols presented here result in the efficient purification of oocysts and sporozoites (Figure 1A) ready for microinjection. The excystation protocol results in the release of sporozoites from approximately 70–80% of the oocysts, therefore it is essential to filter out the remaining oocysts and shells through a 3 µm filter. Filtration results in almost 100% sporozoite purification (Figure 1B). Furthermore, addition of a green dye helps ensure injection of all organoids and allows visualization of injected organoids for at least for 24 h after injection (Figure 2B).

These protocols for preparation of oocysts and sporozoites are straightforward and have been used for many years, so it is expected that the treated oocysts and purified sporozoites will be viable and infectious. However, in our studies, we used scanning electron microscopy to ensure that the excystation process did not damage the sporozoites or oocysts (Figure 2A)10. Injection of equal amounts of oocysts into the organoid lumen can be visually confirmed by simple microscopic imaging (Figure 2C). A portion of infected organoids should be set up to verify parasite propagation by quantitative PCR as we have described10.

Progress through the parasite life cycle can be visualized by collection of infected organoids at different time points post infection and analysis by transmission electron microscopy or by immunofluorescence combined with 4′,6-diamidino-2-phenylindole (DAPI) staining of parasite nuclei10. For example, antibodies to merozoite surface antigens, such as gp40 and gp1517 can be used to identify meront stages; type I meronts will have 8 nuclei and type II meronts, 4 nuclei10. Recently, a panel of monoclonal antibodies specific to trophozoites, merozoites, type I versus II meronts, and macrogamonts has become available18. These antibodies would also be very effective in marking progress of the parasite through its various life cycle stages in the organoids.

Immunofluorescence assays can also be used to explore which cell types are infected by Cryptosporidium. This was especially important to look at in the airway organoids as very little is known about respiratory cryptosporidiosis, and the exact host cell for the parasite was not known. We conducted immunofluorescence assays on Cryptosporidium-infected organoids, co-localizing CC10, a marker for club cells and found that Cryptosporidium infected both CC10- negative and positive cells (Figure 3). These results were corroborated by TEMs in which we observed Cryptosporidium infecting secretory and non-secretory cells in the airway organoids10.

After differentiated organoids have been infected for five days, there should be significant numbers of oocysts being produced. In our hands, infection of organoids from one six-well plate yielded about 4000 oocysts, which could be easily identified and counted on a hemocytometer by labeling with an oocyst-specific antibody. The presence of four sporozoites in the oocysts could be confirmed by drying down a portion of the oocysts onto an adhesive slide, fixing with methanol and combining DAPI staining with oocyst specific antibody (Figure 4). Verification of production of thick walled oocysts could be done by TEM analysis10.

Figure 1
Figure 1: Preparation and purification of Cryptosporidium oocysts and sporozoites. (A) Schematic representation of the method used for oocyst and sporozoite preparation for infection. (B) Image showing in vitro excystation of oocysts. Filtration of unexcysted oocyts and shells gives a purified solution of sporozoites. Scale bar = 10 µm. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Microinjection of oocysts into the organoid lumen. This figure has been modified from Heo et al.10. (A) scanning electron microscopy (SEM) images of oocysts and sporozoites. (B) Image showing oocyst-injected organoids. The green dye helps visualize the injection of each organoid and persists over at least 24 h. (C) Image of an organoid injected with oocysts. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Immunofluorescence image of Cryptosporidium-infected airway organoid. Mucin is labeled with anti-mucin 5 antibody (red) in the lumen of the organoid, club cells are labeled with anti-CC10 (yellow), Cryptosporidium is detected with oocyst-specific antibody (green), and cell nuclei are stained with DAPI (blue). Panel B is an enlargement of the area indicated in the square in panel A. Please click here to view a larger version of this figure.

Figure 4
Figure 4: Immunofluorescence image of oocyst isolated from differentiated intestinal organoids. Oocyst wall is labeled with oocyst-specific antibody in green and the four sporozoite nuclei are visualized with DAPI (blue) Please click here to view a larger version of this figure.

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Culture of Cryptosporidium parasites in intestinal and airway organoids provides an accurate model to study host-parasite interactions10 but also has many other applications. For example, current methods of selecting and propagating genetically modified Cryptosporidium parasites require passage in mice19 which does not allow isolation of parasites that have modifications essential for in vivo infection. Organoid culture of Cryptosporidium provides an alternative to this procedure. However, we have noted that electroporated sporozoites clump together and block the micropipette. For the purpose of selecting genetically modified parasites, organoids can be grown on collagen coated transwells in a two-dimensional format under differentiation conditions to allow infection with transfected sporozoites and consequently the selection of the genetically modified oocysts. The transwells allow access to both the apical and basolateral surfaces and are stable for extended periods of time.

Currently, we culture organoids in a two-dimensional format for high throughput screening of drugs for cancer tissue-derived organoids (unpublished data). This method of organoid culture can also be adapted for testing of anti-Cryptosporidium drugs using the genetically-modified luciferase tagged Cryptosporidium strains19. Moreover, even though the infection is not tightly synchronized, infection of organoids with sporozoites provides sufficient synchronization of the life cycle that drugs can be tested for their efficacy against specific life cycle stages.

Organoid co-culture systems are now being developed taking into account some other aspects of the host system such as microbiota and immune cells20. Thus, the ability to dissect interactions between the parasite and host cells, immune cells and microbiota will soon be possible in vitro. Genetic manipulation of Cryptosporidium is also now possible19, and the combination of fluorescent reporter strains of Cryptosporidium and organoid culture will provide the tools for single cell sequencing of infected cells, and even more specifically single cell sequencing of cells infected with specific stages of the parasite.

The success of the experiments described here is highly dependent on the viability and infectivity of the oocysts. Different batches of Cryptosporidium oocysts can vary widely in excystation rates and ability to infect host cells. Sufficient yields of sporozoites is dependent on good excystation rates and the excystation rate is not always correlated to infectivity. If low levels of infection or poor excystation are observed with a particular batch of oocysts, time and effort may be saved by obtaining a new lot of oocysts rather than attempting to increase oocyst numbers, or lengthening incubation times.

Organoid culture media should be refreshed every alternate day. Use of earlier passages of organoid cultures is advisable. It is important to thaw a new vial of organoids if organoids start to differentiate in later passages as health of organoid cultures vastly determine viability of the parasite. After infection, organoid media should be refreshed every day to avoid accumulation of toxic substances in the media.

Organoid culture of Cryptosporidium is limited in that the parasite cannot be propagated indefinitely, and the infection peters out after three passages over 28 days10. Microinjection of sufficient organoids for mouse experiments such as we have described can be time-consuming and physically taxing. Nevertheless, to date, no other method enables the complete life cycle in an in vitro system completely representative of human infection, nor has any culture system been described that allows exploration of the host-pathogen interactions important for respiratory infection. Organoid culture of Cryptosporidium provides a powerful new tool that opens up avenues of exploration into host-parasite interactions not previously possible for Cryptosporidium.

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The authors have nothing to disclose.


We are grateful to Deborah A. Schaefer from the School of Animal and Comparative Biomedical Sciences, College of Agriculture and Life Sciences, University of Arizona, Tucson, AZ, USA for helping us with oocyst production and analysis. We also thank Franceschi Microscopy and Imaging Center and D.L. Mullendore at Washington State University for TEM preparation and imaging of isolated organoid oocysts.

D.D. is the recipient of a VENI grant from the Netherlands Organization for Scientific Research (NWO-ALW, 016.Veni.171.015). I.H. is the recipient of a VENI grant from the Netherlands Organization for Scientific Research (NWO-ALW, 863.14.002) and was supported by Marie Curie fellowships from the European Commission (Proposal 330571 FP7-PEOPLE-2012-IIF). The research leading to these results has received funding from the European Research Council under ERC Advanced Grant Agreement no. 67013 and from NIH NIAIH under R21 AT009174 to RMO. This work is part of the Oncode Institute, which is partly financed by the Dutch Cancer Society and was funded by a grant from the Dutch Cancer Society.


Name Company Catalog Number Comments
Basement membrane extract (extracellular matrix) amsbio 3533-010-02  
Crypt-a-Glo antibody (Oocyst specific antibody) Waterborne, Inc A400FLR-1X Final Concentration = Use 2-3 drops/slide
Crypto-Grab IgM coated Magnetic beads Waterborne, Inc IMS400-20  
Dynamag 15 rack Thermofisher Scientific 12301D  
Dynamag 2 rack Thermofisher Scientific 12321D  
EMD Millipore Isopore Polycarbonate Membrane Filters- 3µm EMD-Millipore TSTP02500  
Fast green dye SIGMA F7252-5G    
Femtojet 4i Microinjector Eppendorf 5252000013  
Glass capillaries of 1 mm diameter WPI TW100F-4  
Matrigel (extracellular matrix) Corning 356237  
Microfuge tube 1.5 mL Eppendorf T9661-1000EA  
Micro-loader tips Eppendorf 612-7933  
Micropipette puller P-97 Shutter instrument P-97  
Normal donkey Serum Bio-Rad C06SB  
Penstrep Gibco 15140-122  
Sodium hypoclorite (use 5%) Clorox 50371478  
Super stick slides Waterborne, Inc S100-3  
Swinnex-25 47 mm Polycarbonate filter holder EMD-Millipore SX0002500  
Taurocholic acid sodium salt hydrate SIGMA T4009-5G  
Tween-20 Merck 8221840500  
Vectashield mounting agent Vector Labs H-1000  
Vortex Genie 2 Scientific industries, Inc SI0236  
Adv+++ (DMEM+Penstrep+Glutamax+Hepes)     Final amount
DMEM Invitrogen 12634-010 500 mL
Penstrep Gibco 15140-122 5 mL of stock in 500 mL DMEM
Glutamax Gibco 35050038 5 mL of stock in 500 mL DMEM
Hepes Gibco 15630056 5 mL of stock in 500 mL DMEM
INTESTINAL ORGANOID MEDIA-OME (Expansion media)     Final concentration
A83-01 Tocris 2939-50mg 0.5 µM
Adv+++     make up to 100 mL
B27 Invitrogen 17504044 1x
EGF Peprotech AF-100-15 50 ng/mL
Gastrin Tocris 3006-1mg 10 nM
NAC Sigma A9125-25G 1.25 mM
NIC Sigma N0636-100G 10 mM
Noggin CM In house*   10%
P38 inhibitor (SB202190) Sigma S7076-25 mg 10 µM
PGE2 Tocris 2296/10 10 nM
Primocin InvivoGen ant-pm-1 1 mL/500 mL media
RSpoI CM In house*   20%
Wnt3a CM In house*   50%
In house* - cell lines will be provided upon request      
INTESTINAL ORGANOID MEDIA-OMD (Differentiation media)     To differentiate organoids, expanding small intestinal organoids were grown in a Wnt-rich medium for six to seven days after splitting, and then grown in a differentiation medium (withdrawal of Wnt, nicotinamide, SB202190, in a differentiation medium (withdrawal of Wnt, nicotinamide, SB202190, prostaglandin E2 from a Wnt-rich medium or OME)
LUNG ORGANOID MEDIA- LOM (Differentiation media)     Final concentration
Adv+++     make up to 100 mL
ALK-I A83-01 Tocris 2939-50mg 500 nM
B27 Invitrogen 17504044 0.0763888889
FGF-10 Peprotech 100-26 100 ng/mL
FGF-7 Peprotech 100-19 25 ng/mL
N-Acetylcysteine Sigma A9125-25G 1.25 mM
Nicotinamide Sigma N0636-100G 5 mM
Noggin UPE U-Protein Express Contact company directly 10%
p38 MAPK-I Sigma S7076-25 mg 1 µM
Primocin InvivoGen ant-pm-1 1:500
RhoKI Y-27632 Abmole Bioscience M1817_100 mg 2.5 µm
Rspo UPE U-Protein Express Contact company directly 10%
Reducing buffer (for resuspension of oocysts and sporozoites for injection)     Final concentration
L-Glutathione reduced Sigma G4251-10MG 0.5 μg/μL of OME/OMD/LOM
Betaine Sigma 61962 0.5 μg/μL of OME/OMD/LOM
L-Cysteine Sigma 168149-2.5G 0.5 μg/μL of OME/OMD/LOM
Linoleic acid Sigma L1376-10MG 6.8 μg/mL of OME/OMD /LOM
Taurine Sigma T0625-10MG 0.5 μg/μL of OME/OMD/LOM
Blocking buffer (for immunoflourescence staining)     Final concentration
Donkey/Goat serum Bio-Rad C06SB 2%
PBS Thermo-Fisher 70011044 Make up to 100 mL
Tween 20 Merck P1379 0.1%
List of Antibodies used      
Alexa 568 goat anti-rabbit Invitrogen A-11011 Dilution-1:500; RRID: AB_143157
Crypt-a-Glo Comprehensive Kit- Fluorescein-labeled antibody Crypto-Glo Waterborne, Inc A400FLK Dilution- 1:200
Crypta-Grab IMS Beads- Magnetic beads coated in monoclonal antibody reactive Waterborne, Inc IMS400-20 Dilution-1:500
DAPI Thermo Fisher Scientific D1306 Dilution-1:1,000; RRID : AB_2629482
Phalloidin-Alexa 674 Invitrogen A22287 Dilution-1:1,000; RRID: AB_2620155
Rabbit anti-gp15 antibody generated by R. M. O’Connor (co-author). Upon request Upon request Dilution-1:500
Sporo-Glo Waterborne, Inc A600FLR-1X Dilution- 1:200



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