An efficient protocol for maintaining and amplifying entomopathogenic nematode is essential for understanding the molecular basis of nematode pathogenicity and host anti-nematode immunity. This protocol enables us to produce a high number of symbiotic and axenic Heterorhabditis bacteriophora and Steinernema carpocapsae IJs using insect host, Galleria mellonella. Begin by covering the Petri dish with a piece of filter paper and add approximately 10 to 15 Galleria mellonella larvae.
Using a pipette, dispense two milliliters of water containing about 25 to 50 infective juveniles per 10 microliters of suspension onto the wax worms. And store the Petri dish in a cabinet at room temperature. Depending on the moisture of the filter paper, add one to two milliliters of water every two days.
Wax worms infected with infective juveniles usually die within 48 hours. Prepare White's water traps approximately 10 days after the wax worms are infected with the infective juveniles. Carefully transfer the dead insects onto the unsoaked part of the filter paper.
Use tap water to fill the bottom Petri dish and place a cap of a 15 milliliter tube as a spacer for ventilation. Use tap water instead of demineralized or deionized water to avoid nematode aggregation, and ensure that the water level in the water trap reaches approximately half the height of the Petri dish. When the water in the small Petri dish turns cloudy due to nematodes, use a pipette to move the new generation of infective juveniles into a T25 or a T75 cell culture flask.
Later, add tap water up to 40%volume or until the appropriate density is reached. And store the cell culture flask horizontally to avoid nematode congestion. Repeat transferring of infective juveniles into cell culture flasks and the addition of water until the infective juveniles stop emerging from the insect carcasses in approximately three to five days.
Using an inoculation loop, scrape several flakes of a frozen culture of Photorhabdus temperata Ret16 onto a MacConkey plate and streak for single colonies. Incubate the plate for two to three days at 28 degrees Celsius. After incubation, inoculate 10 milliliters of LB broth with a colony on MacConkey agar in a 50 milliliter tube and incubate the culture overnight at 28 degrees Celsius in a shaking incubator.
After incubation, transfer 100 microliters of overnight culture from a 50 milliliter tube into a new microcentrifuge tube, then wash 100 the overnight culture with 900 microliters of single strength PBS by centrifusion, in a 1.5 milliliter microcentrifuge tube, at 17, 900 times g. Decant the supernatant. Next, dilute the culture 10 times in single strength PBS and leave the tube on ice.
Now, immerse the wax worms into a 70%ethanol solution. After drying the insects with a paper towel, place them in a 50 milliliter tube. Place the tube on ice for 20 minutes to immobilize the wax worms.
Use filter paper to cover a Petri dish's top and bottom halves. Use the Petri dish lid covered with filter paper as a base support for injection. Moist in the filter paper of the bottom Petri dish and transfer it on ice to help the injected wax worms recover.
Pipette 50 microliters of ice cold bacteria on a piece of Parafilm then prepare a 22 gauge refiner needle syringe and press the plunger gently to remove the air at the tip of the needle. Keeping a wax worm close to the posterior end under the stereoscope. Inject bacterial culture into the dorsal side of the thorax.
Preferably at the junction between two segments just underneath the cuticle parallel to the wax worm to minimize internal damage and pay attention to how the larval body starts bulging. Next, transfer the injected insects to the recovery Petri dish. And once all the insects are injected and placed in the recovery dish, place the Petri dish in the dark.
Wax worms succumb two days after injection and appear brick-red after approximately three to four days. However, brown-colored insects indicate unsuccessful bacterial infection. At seven days post-infection, transfer the insects with the characteristic brick-red color onto a fresh filter paper lined Petri dish, and repeat the production of symbiotic nematode infective juveniles.
To begin surface sterilization, collect sufficient symbiotic, Heterorhabditis bacteriophora and candidate axenic infective juveniles by centrifugation. To make a pellet in a 1.5 milliliter centrifuge tube, add 500 microliters of 5%bleach to the pellet in each microcentrifuge tube and invert the tube. After incubating for 10 minutes, centrifuge at 17, 900 times g for one minute and remove the supernatant.
Then wash the pellet with one milliliter of sterile water. Centrifuge it, remove the supernatant and repeat this procedure four more times. The graph shows the results of assessing the status of Heterorhabditis bacteriophora nematodes that have gone through axenization.
Heterorhabditis bacteriophora from the stock culture carried symbiotic Photorhabdus luminescens cells. However, nematodes devoid of their associated Photorhabdus luminescens bacteria were axenic. To maintain the reproducibility of your experiment, always streak freshly from bacterial stock culture from a fresh colony and take out the overnight culture before it reaches stationary face.
Only use the brick-red Galleria wax worm for White's water trap and discard the rest. After this procedure, the nematodes can be used to investigate the interaction with insect immune system. This technique paves the way for researcher to explore the molecular basis of nematode parasitism and host anti-nematode immunity.
