A protocol is presented for formulation of a blood-free artificial diet to feed Anopheles mosquitoes in captivity. This diet has a similar performance to vertebrate blood and triggers oogenesis and egg maturation and produces viable adult progeny.
Malaria parasites such as meeted by mosquitoes that play a major role on the spread of the disease. So far, it has been impossible to rear malaria mosquitoes without blood. And to overcome this limitation, our protocol describes a blood substitute diet that is able to support mosquito breeding in the insectory.
The use of blood free diet is highly advantageous over blood. A blood free diet does not have the ethical constraints as the use of human blood, or experimental animals. Replacing animals on experimentation is part of our three R policy.
Replace, reduce, refine. Not using blood reduces costs, and the logistics associated with collection, storage and maintenancey of red blood. The use of artificial diets can facilitate testing of anti plasmodium molecules.
Until now, we have tested our diets using different anopheles species. In all of them, the diet was well engorged by the females and allowed egg production and laying. We believe this diet has potential use for the rearing of other mosquito species.
Actually, we are now testing it for aedes, a vector of many diseases such as dengue fever, zika virus, or yellow fever. Maintain anopheles coluzzii yaonde strain mosquitoes in a room at 26 degrees Celsius, 75%humidity, and under a 12 hour to 12 hour light dark cycle. House mosquitoes using standard insectory conditions in a single cage to guarantee mating.
Use a plastic pipette to collect mosquito pupae into a small water container. Place the container inside a mosquito cage to let adult mosquitoes emerge and mate. Provide 10%glucose feeding solution in the cage.
Three days after emergence, use an aspirator to collect the necessary number of females from the stock cage into a paper cup. To distinguish, females are larger and males have a wider and feathery proboscis. One day before feeding trails, remove the 10%glucose feeding solution.
The next day, prepare artificial liquid diets under sterile conditions in the laminar flow cabinet according to the manuscript. Add all ingredients to a plastic tube. Mix all ingredients thoroughly and filter using a 0.45 micron microfilter.
Fill a sterile 1 milliliter syringe equipped with a 27 gauge half inch needle with 100 microliters of one milligram per milliliter heparin. Then, anesthetize six to eight week old CD one female mice with ketamine and xylazine using the intra peritoneal route. Evaluate if the mouse displays any muscle reaction and response to different physical stimuli.
Perform the cardiac puncture. Then collect blood from the mouse into a micro tube and maintain blood at 37 degrees Celsius in a water bath. Next, collect approximately 30 female mosquitoes from the stock cage using an aspirator.
Transfer the female mosquitoes to 500 milliliter paper cups and cover with a fine mosquito net mesh so they cannot escape. Stretch parafilm membrane across the mouth of the glass feeder to contain the meal. Apply a glass bell artificial feeding apparatus connected to plastic tubes to the top of each cup.
Provide a constant water flow to the cylindrical tubing and feeder so the temperature within is kept at approximate 37.5 degrees Celsius. Apply one milliliter of prewarmed liquid diet at 37 degrees Celsius or fresh mouse blood into a glass feeder. Feed the mosquitoes for 60 minutes in the dark in 26 degrees Celsius.
After artificial feeding, cold anesthetize the mosquitoes at 20 degrees Celsius for 30 seconds. Then, place the mosquitoes in a refrigerated Petri dish. Record the number of fully engorged female mosquitoes.
Separate 30 fully engorged females and put them on a new cage. Now, place a humidified filter paper at the bottom of each cage. Keep the mosquitoes at 26 degrees Celsius, 75%humidity, and under a 12 hour to 12 hour light dark cycle with 10%glucose ad libitum.
At 96 hours and 120 hours post feeding, count the eggs with the help of a handheld magnifying glass. Flood the filter paper with distilled water to collect the eggs into trays filled with distilled water. Feed the larvae daily with approximately 13 milligrams of ground fish food per tray.
Remove dead pupae and larvae with a plastic pipette daily. When all pupae have developed into adults, count the number of adult males and females, register the dates of hatching and death, and calculate the mortality rates. To test for longevity, collect 15 adult males and 15 adult females from the F1 generation of each diet group into a paper cup.
Feed adults with 10%glucose solution ad libitum. Use tweezers or a brush to remove the dead adults daily. Maintain the mosquitoes at the same temperature, humidity, light cycle conditions and sugar feeding regime.
Register the death dates and calculate the longevity. To measure wing length, cold anesthetize five day old F1 male and female adult mosquitoes from each diet group at 20 degrees Celsius for 90 seconds. Under a stereoscope, gently grasp the thorax of each mosquito with forceps, and place them ventral side up.
Collect both wings using a scalpel, and place them on a clean microscope slide containing a dried drop of mounting medium for further measurement using a graduated eyepiece. Measure the wing length with a stereoscope using a micrometer. In this study, the performance of female anopheles mosquitoes fed on the formulated rich artificial meal and mosquitoes fed on the initial liquid diet, or a fresh blood meal was compared.
The number of engorged female mosquitoes fed with rich artificial meal at 89%was significantly higher than the number of engorged females fed on blood at 56%The F1 generation of mosquitoes fed on either the blood or the rich artificial meal had comparable mortality and survival rates. Variability was higher in the blood fed mosquitoes relative to mosquitoes fed with the rich artificial meal. In terms of adult body size, F1 anopheles mosquitoes fed with rich artificial meal was within the expected range, and was similar to blood fed insectory mosquitoes.
In my opinion, it is very important to assemble the feeders properly with the parafilm to avoid it's rupture. If the membrane is not well attached to the glass feeder, you can lose the meal and probably lose some of the mosquitoes as they can be covered by the meal and die. We would like to perform a dual choice attraction assay, for example using an olfactometer so we could actually evaluate if our females are more attracted to the artificial diet or to the blood.
Also, we are now lyophilizing the artificial diet and studying it's stability among different temperatures a long time. Besides the obvious improvements to stability and storage that we've been working on, the long term use of the diet on mosquito fitness and physiology should be investigated. I believe that producing the anopheles without blood will facilitate immensely vector research and implementation of control tools that depend of a large number of mosquitoes.
