Cultivation of Heligmosomoides Polygyrus: An Immunomodulatory Nematode Parasite and its Secreted Products

NOTE: All procedures in this protocol are performed in accordance with guidelines set out by the United Kingdom Home Office and the University of Edinburgh Veterinary Services.

1. Infection of Mice by Gavage

1. Store Heligmosomoides polygyrus L3 larvae in distilled water for up to six months at 4 °C.
2. Before use, wash L3 larvae three times in distilled water: centrifuge at 300 x g for 10 min (with brake), remove all but 500 μl of water (to avoid disturbing pelleted L3 larvae) and resuspend the pellet each time.
3. For the third wash, add water to an exact volume (typically 40 ml) and aspirate 20 µl with a 200 µl tip cut to widen its aperture. Place two 20 µl samples on the surface of a 60 mm culture dish and count the L3 larvae (usually mobile, and best viewed under 50X magnification with a dissecting microscope). Resuspend the final pellet in distilled water to a concentration of 2,000 L3 larvae per ml.
4. For life cycle production, infect 8 week-old F1 (C57BL/6xCBA) mice with 400 H. polygyrus L3 larvae in 200 μl of distilled water by oral gavage (restrain mice in the upright position by the scruff of the neck and gently pass the blunt gavage needle through the mouth and esophagus into the stomach). Agitate thoroughly prior to each infection (larvae settle quickly in water) and aspirate 200 µl in a 1 ml syringe; use a dedicated gavage needle with a rounded end. For experimental infection of younger mice (6-8 weeks old), or other inbred strains (e.g. C57BL/6 or BALBc), infect mice with 200 L3 larvae.

2. Propagation and Maintenance of H. polygyrus

1. Place charcoal in the center of a large plastic tub and allow cold tap water to run over it for a minimum of 30 min (unwashed activated charcoal is toxic to L3 larvae). Drain the water from the tub and place the charcoal on two layers of absorbent paper, leaving it exposed to room air until completely dry.
2. If eggs are required, scrape feces first out of the colon with forceps (and scissors if necessary). If a large number of L3 larvae is needed, place mice on a wire grid and collect fecal pellets over several days.
3. Mix the feces with granulated charcoal at a ratio of at least a 1:1, to achieve a consistency just damp enough to adhere to filter paper. Smear a thin layer on the center of some dampened filter paper in a petri dish and place this in a humid box (add some damp paper towel and a dish of water) in the dark for 12-14 days.
4. Remove L3 larvae from day 7 onwards, and collect them on at least two occasions before the paper is discarded. The larvae form a ring around the edge of the filter paper; lift the filter paper out of the petri dish and rinse the larvae that are left of the plate (using a pipette and 5 ml of sterile water per plate) into a 50 ml tube.
5. Lift off the filter paper and harvest the remaining larvae left on the plate with distilled water into a 50 ml tube. Centrifuge the effluent solution at 300 x g for 10 min. Wash the larvae three times with distilled water and then store at 4 °C in up to 50 ml of distilled water until required.
   NOTE: Larvae remain viable and infective for at least 6 months.

3. Collection of Adult H. polygyrus Worms

1. Prepare the modified Baermann Apparatus in advance as shown in Figure 1.
2. Cull mice fourteen days after infection.
3. Wash the abdomen with 70% ethanol. Cut the skin over the abdomen and pull back to reveal the anterior abdominal wall. Make midline incision to enter the peritoneal cavity.
4. Remove the entire small and large intestine (from proximal duodenum to distal rectum). Place into a dry Petri dish.
5. Straighten the gut along its entire length; excise the feces-containing colon; place this into a separate dish for egg preparations later.
6. Excise the proximal 20 cm of small intestine that contains the adult worms –identified by the relatively thick wall of the duodenum and often a red appearance due to the intra-luminal worms. Place into a (100 mm diameter) Petri dish with 5 ml of Hanks’ Solution, warmed to 37 °C (two specimens per dish).
7. Open the worm-filled proximal gut portion longitudinally with scissors (round-ended scissors are best for this), and scrape down inside of gut lining with two glass slides to remove the worms. Then discard the clean gut wall.
8. Tip worms into small muslin bags, staple closed and secure with paperclips around the edge of the glass funnel (Figure 1).
9. Fill funnel with Hanks’ Solution and add approximately 4 Petri dishes of worms into each funnel.
10. Place apparatus in 37 °C incubator for 1-2 hr, gently agitating half way through to dislodge debris from the gut preparation that may occlude the muslin filter. Take care to avoid spillage of debris outside the muslin bag – this will cause contamination of the final HES preparation. Adult worms should have slowly migrated through the muslin cloth and settled at the bottom of the glass test tube. Carefully detach the test tube from the connecting rubber hose over the sink (taking care to avoid losing worms at this point).
11. Using a plastic pipette, transfer the worms into a 50 ml tube and wash six times with Hanks’ Solution (allow worms to settle with gravity, remove media with a stripette, add 40 ml of Hanks’ Solution and repeat five times).
    NOTE: worm culture must be kept sterile from this point onwards.
    1. Move to a laminar flow hood and wash another six times in sterile Hanks’ Solution supplemented with 100 U/ml penicillin and 100 µg/ml streptomycin, ready for in vitro culture.
12. Count the adult worms recovered by taking two samples of 20 µl taken up with a yellow tip cut to widen its aperture; expect approximately 50% of the quantity of inoculated larvae.

4. Setting Up Cultures for HES: Medium Preparation, Washing Adult Worms

1. Soak the worms from 3.11 in approximately 10 ml of RPMI supplemented with 10% Gentamycin for 20 min, leaving the tube resting at an angle to ensure worms are fully covered.
2. Perform this in a laminar flow hood: wash again six times with Hanks' Solution (supplemented with 5 U/ml penicillin and 5 μg/ml streptomycin).
3. Prepare H. polygrus media.
4. Maintaining sterility in a laminar flow hood, to 500 ml of RPMI1640, add 11.1 ml of 45% glucose (final concentration 1.2% as RPMI1640 contains 0.2% glucose), 5 ml of 100x Peniclllin-Streptomycin (final concentration 5 U/ml penicillin, 5 μg/ml streptomycin), 5 ml of L-glutamine (final concentration 2 mM), and 5 ml of Gentamycin (final concentration 1%). Do not add FCS.
5. Aliquot worms into vented T25 flasks, approx. 1,000 worms in 15 ml H. polygyrus media per flask, and place upright in 37 °C incubator (5% CO₂) for 3 weeks.

5. Preparation of HES

1. Collect HES-containing culture media from cultures at intervals of no longer than twice per week, setting aside the first collection after 24 hr of culture (due to high levels of LPS contamination and host proteins – can be processed separately or discarded). Keep each collection separate and clearly labeled with the date and batch number. Replace with an equal volume of H. polygyrus media on each occasion.
2. Centrifuge HES-containing media at 400 x g for 5 min. Then filter sterilize through 0.2 µm low-protein-binding filters into 50 ml tubes. Store in the -20 °C freezer clearly labeled with date of worm harvest and date of HES collection.
3. After 21 days of HES collection from culture, discard worms.
4. Pool 500-1,000 ml of HES supernatant (usually from frozen stock, and not including the first 24 hr collection) and concentrate over a 3,000 MWCO filter in the ultrafiltration device under nitrogen pressure.
   NOTE: Be very careful not to let the filter run dry.
   1. To set up the filter device, first wash the 3 kDa membrane shiny side down in a 1 liter beaker with distilled water for 3 x 20 min whilst stirring. Assemble the ultrafiltration device as per manufacturer’s instructions with filter membrane shiny side up. Place in cabinet at 4 °C and pass 50 ml of distilled water through before starting to concentrate pooled HES.
   2. Add each tube of HES into the filtration device as required (100-140 ml per day), until the volume is concentrated down to 2-5 ml.
5. In order to remove contaminants from the HES-containing culture media, add 50 ml of pyrogen-free PBS to the filtration device and and then concentrate down to approximately 2 ml. Repeat this step twice (150 ml of PBS in total). Transfer HES into a 15 ml tube, filter sterilize (with a 0.2 μm filter) in a laminar flow hood and measure protein concentration using a spectrophotometer (E₂₈₀ = 10) or by Bradford assay.
6. Aliquot, label with batch number and date, and freeze at -80 °C.
7. Perform a chromogenic LAL assay according to the manufacturer’s protocols on each batch of HES prior to use. If LPS levels are greater than 1 U LPS per 1 µg protein, consider not using this batch for in vivo experiments or in vitro cultures.
8. Process the HES collected at 24 hr separately in the same manner; it will contain LPS and some host proteins and, while not suitable for functional experiments, it is a useful source of individual molecules that may be isolated by monoclonal antibody affinity purification.