The World Health Organization estimates that one in four people on this planet is infected with parasitic worms known as helminths. We aim to understand how the protective immune response against helminths is initiated, regulated, and executed. Helminths survive within the mammalian hosts for decades, at least long enough to reproduce, even when faced with a strong immune response.
We are beginning to understand now how these parasites manipulate, evade, and suppress their host's immune response to extend their survival. A challenge studying Strongyloides Ratti infection is the special migration of this parasite, which is predominantly through the tissue to the head and intestine. By quantifying migrating larvae in the head and intestine, we can dissect different immune effectors at different sites.
Our group established that mast cells and basal fields are completely dispensable for tissue migration, but are essential for ejection of S.ratti from the intestine. By contrast, neutral fields do not play an important role in intestinal immunity, but are essential in killing migrating larvae in the tissue. During my study of sex differences in S.ratti infection, I observed that both male and female mice had similar numbers of migrating larvae in their tissue.
However, I found a higher parasite number in the intestine of male mice compared to female mice. This suggests that intestinal immunity varies between the biological sexes. To begin, prepare the Baermann apparatus.
Using a clamp, close the bottom of the hose at an oblique angle and fill the apparatus with lukewarm water until the sieve is submerged. Place a tissue wipe in the sieve and fill it with the feces charcoal mixture. Then, switch on the light positioned directly behind the Baermann apparatus and allow viable iL3 to migrate actively through the wipe.
After 30 minutes, briefly open the clamp to collect the settled larvae into a 50 milliliter tube, minimizing the volume as much as possible. Then, fill the 50 milliliter tube with PBS containing 1%penicillin streptomycin. Let the larvae settle at the bottom of the tube by gravity at four degrees Celsius for 30 minutes before removing the supernatant.
After the third wash, re-suspend the pellet in 30-50 milliliters of PBS penicillin streptomycin depending on pellet size. Immediately before pipetting, agitate the solution as the iL3 larvae settle quickly and transfer one microliter drop of the solution onto a microscopy slide. Inspect the droplets under an inverted microscope at 40 times magnification.
Ensure the iL3 larvae are moving vividly. To begin, prepare 1000 or 2000 iL3 larvae in PBS containing 1%penicillin streptomycin using the Baermann apparatus. Once the iL3 larvae settle by gravity, use a 0.5 milliliters syringe to aspirate the supernatant as completely as possible, leaving approximately 30 microliters in the tube.
Resuspend the iL3 suspension by flicking the tube or using a 0.5 milliliter syringe and aspirate the remaining suspension into the syringe. Now pick up the mouse by the scruff and secure one hind foot. Subcutaneously, inject the entire solution containing the iL3 larvae into the foot pad at a flat angle and then slowly retract the syringe.
Spray the abdomen and neck of the euthanized, infected mouse with a commercial disinfectant. Using scissors, cut the skin over the abdomen and pull it back to reveal the anterior abdominal wall. After opening the peritoneal cavity, cut the diaphragm and open the ribs to both sides to expose the lungs in the pleural cavity.
Then, collect the lung lobes into a 24-well plate marked with lines drawn or printed at approximately four millimeters and filled with one milliliter of tap water per well. Now, cut the entire lung into six pieces, each approximately 0.75 centimeters by 0.75 centimeters. After cutting off the head of the mouse, remove the skin and fur using fingers, minimizing the removal of muscle tissue.
Dissect the complete head into four quarters by cutting behind the eyes and longitudinally through the center of the head. Place the anterior and posterior quadrants into a six-well plate containing two milliliters of tap water. After incubation, swirl the plates one final time, then remove the remaining tissue parts from the wells using forceps and discard them.
Count all the larvae in the wells under a 40 times inverted microscope. Follow the lines on the well bottoms to ensure a complete count. After soaking the abdominal region in a commercial disinfectant, cut the skin over the abdomen with scissors and pull it back to expose the anterior abdominal wall.
Make a midline incision to open the peritoneal cavity. Using scissors, cut between the stomach and the proximal duodenum as well as between the colon and the anus to detach the entire intestine. Then, using fingers, gently pull out the intestine and place it into a Petri dish containing tap water.
Cut the intestine sections open longitudinally and vigorously shake them in tap water for at least 10 seconds to wash out feces and mucus. Transfer the cleaned intestinal sections into 50-milliliter tubes containing 20 milliliters of tap water and incubate. After three hours of incubation, remove the intestinal tissue from the water.
Place the tubes vertically at room temperature and allow the parasites to sediment by gravity for 30 minutes. Aspirate the supernatant until approximately five milliliters of water remains in the tube. Refill the tube with tap water to a total volume of 25 milliliters.
When the suspension's visibility is clear, aspirate the supernatant until approximately five milliliters of water remain. Transfer this liquid into two wells per mouse intestine in a six-well plate marked with lines. Count the adult female parasites under an inverted microscope with a 40 times magnification by moving along the lines drawn on the well bottoms.
Larvae were first detected in the head on day one post-infection, showing uniform distribution, with a small fraction also retrieved from the lungs. The number of larvae in the head increased significantly on day two post-infection, with an average of 174 larvae in the head and 17 larvae in the lungs. Larvae numbers in the head decreased markedly by day three post-infection, with no larvae retrieved from the head by day four.
Parasites were detected in the intestine from day four post-infection, with the majority localized in the duodenum and the first two-thirds of the jejunum persisting until day six. By day seven post-infection, most adult parasites were localized in the cecum, where they persisted until day nine, before their numbers began to decline significantly by day 11. No parasitic adults were retrieved from the intestine on day 14 post-infection, and total intestinal parasite numbers dropped to zero by this time point.
To begin, hold rats infected with Strongyloides ratti in cages with several layers of cellulose and minimal litter at days five and 12 post-infection. On days 6 to 8 and 13 to 15 post-infection, transfer the rats to fresh cages and collect all fecal pellets from the old cages. Pool the collected feces from one cage into a single 50-milliliter tube.
Take the pre-soaked activated charcoal and wash it before use. Mix the feces with pre-soaked activated charcoal in equal amounts. Arrange the mixture diagonally to form a gradient and cover it with an additional layer of activated charcoal to reach a final ratio of 1 to 2.
Then, cover the glass beaker with a transparent film, ensuring that air holes are made to allow proper air circulation.
