Mar 16, 2024
Short-Term Freezing of Anopheles stephensi (Mosquito) Larvae
- Michael C. Young11,
- Jackson Watkins1,
- Gabriela Ramirez1,2,
- Emily N. allichotte1,
- MaKala Herndon1,
- Elisha Xiao-Kim3,
- Madison Stoltz1,
- Maria Alexandra Marquez3,
- Gaukhar Iskakova3,
- Jennifer Barfield3,
- Karen M. Dobos1,
- Gregory D. Ebel1
- 1Department of Microbiology, Immunology and Pathology, College of Veterinary Medicine and Biomedical Sciences, Colorado State University, Fort Collins, CO;
- 2Cellular and Molecular Biology, Colorado State University, Fort Collins, CO;
- 3Department of Biomedical Sciences, Animal Reproduction and Biotechnology Laboratory, Colorado State University, Fort Collins, CO, USA
Protocol Citation: Michael C. Young1, Jackson Watkins, Gabriela Ramirez, Emily N. allichotte, MaKala Herndon, Elisha Xiao-Kim, Madison Stoltz, Maria Alexandra Marquez, Gaukhar Iskakova, Jennifer Barfield, Karen M. Dobos, Gregory D. Ebel 2024. Short-Term Freezing of Anopheles stephensi (Mosquito) Larvae. protocols.io https://dx.doi.org/10.17504/protocols.io.j8nlk8b5dl5r/v1
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: March 06, 2024
Last Modified: March 16, 2024
Protocol Integer ID: 96801
Keywords: cryopreservation method for mosquito larvae, term freezing of anopheles stephensi, robust method of cryopreservation, cryopreservation method, term maintenance of mosquito colony, cryopreservation, larvae mosquito insectary, mosquito larvae, mosquito colony, anopheles stephensi, larvae, mosquito, freezing, term freezing, including silkworm, other insect, honey bee, lack of preservation procedure, preservation procedure, fruit fly
Funders Acknowledgements:
NIH/NIAID
Grant ID: HSN272201700018I, task order 75N93020F0002
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Abstract
Mosquito insectaries must be maintained around the clock without interruption, due to the lack of preservation procedures. Long-term maintenance of mosquito colonies can lead to population bottlenecks, laboratory adaptation, loss of genetic
markers, etc., leading to colonies that are attenuated and different from their wildtype counterparts. Cryopreservation is an essential tool to combat these challenges that has been developed in other insects, including silkworms, fruit flies, and honey bees. Unfortunately, there is no robust method of cryopreservation for mosquitoes. This protocol details the short-term freezing of Anopheles stephensi (Mosquito) larvae; an initial step towards establishing a cryopreservation method for mosquito larvae.
Attachments
pytpb8vuf.pdf
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Materials
- Mosquito eggs on egg paper
- Insectary chamber, capable of programable temperature, light, and humidity
- Distilled water
- Fish food
- Small hatch bin (a solid, polycarbonate or greater strength container, that can hold water to a depth of 5 or more cm, with dimensions approximately 10 cm x 20 cm)
- Two (or more) Large Hatch bins (dimensions approximately 20 cm x 40 cm)
- Plastic transfer pipettes
- Strainer
- Gloves
- Small (2 oz) disposable cups
- Insect rearing cage with nylon mesh screen and port
- Scissors
- Circulating water pump
- Glass insect feeder
- Parafilm
- Filter paper
- Defibrinated calf blood
- Microscope slides
- Olympus SZX16 Advanced Stereo Microscope (or similar)
- Penetrating cryoprotective agents (see examples, section III.A
- Non-penetrating cryoprotective agents (we use 0.5 M Trehalose)
- Square culture (petri) plates
- FalconTM Cell Strainers, 70 um (or similar)
- Thermoelectric Laboratory Cold Plate (AHP-1200CPV or similar)
- Kim wipes
Phase I: Establish and maintain mosquito colony
2w 0d 11h 15m
Phenotypic Analysis of developing mosquitoes (Figure 1)
- Larvae develop from eggs to first instar larvae (L1) around 2-3 days after transferring egg paper.
- L1 develop into L2/L3 over the next 4 days (approximately). During that time, they show the
ability to hatch, feed and further develop. L4 transition into pupa and emerge as adults
approximately 2 days after pupation
An. stephensi life stages showing hatch and maturation rate data collection points
Husbandry (modified from MR4 Methods in Anopheles Research. MR4 Methods in
Anopheles Research Laboratory Manual, BEI Resources, 2015)
1. Hatch eggs
- Obtain egg paper with referenced eggs from BEI or other qualified source.
- Take clean small hatch bin, label with date, and fill with ~500 mL of distilled water. Dispense water slowly to avoid bubbles (eggs will hatch asynchronously if bubbles are present).
- Put on gloves and place the egg paper into the water.
- Add a pinch of fish food and cover the bin.
- Place bin in insect chamber and maintain under controlled conditions of temperature (28 °C ), relative humidity (70%) and light (12:12 L:D diurnal cycle).
2. Larvae splitting and feeding
- Slowly fill a large hatch bin with water. Tap or distilled water can be used.
- Move larvae into a larger bin using a strainer, roughly 24 hours after eggs are hatched.
- Make sure to thoroughly rinse out the small bin where eggs were hatched to ensure all larvae are transferred to the large bin.
- For the following days, make sure that bins are not over saturated with larvae (~60% saturation), or else the larvae will stop maturing; split larvae between current bin and new bin as much as necessary.
- Add food and cover bin to any new bins prepared.
- Changing the water may be necessary to avoid mold and other microbial growth; Use strainers and transfer larvae into a fresh bin if necessary.
3. Collecting pupae
- Pour an ounce of tap water into a clear, plastic, disposable 2oz (about 59.15 ml) cup.
- Use a transfer pipette or a small mesh-made-scoop to collect the pupae into the cup.
- Transfer the cup into an insect rearing cage by placing cup through top port. Add sugar and water cups through the port.
- Males will emerge first, so it is important to pick up pupae every day for several days (~120:00:00 ) to ensure females emerging from pupae are also picked.
5d
4. Blood feeding mosquitoes (glass feeder protocol)
- The optimal time to blood feed mosquitoes is 3-5 days post-eclosion.
- Remove sugar and water 12-16 hrs prior to blood feed.
Note
The colony will significantly die off if starved for more than 24 hrs.
- Fill water basin with tap water and attach heating element/pump. Turn on and pre-warm water to 37 °C – 42 °C .
- Prepare glass feeders. Use scissors to cut the length of parafilm or hog gut sufficient to cover mouth of glass feeder.
Note
Hog gut is often stored frozen. Allow time to thaw before use or thaw in tap water.
- Briefly turn off the water pump and use plastic tubing to connect all feeders in a circuit that leads back into the water bath. Wet each end of each piece of plastic tubing before use to ensure easier fit. Turn pump back on and ensure A) a steady flow of water around circuit and B) no dripping. Do not, however, place feeders on top of rearing cages yet, as mosquitoes may begin to drink and fill up on water.
Note
If hog gut is used, be careful not to let it dry out. A petri dish with wet paper towels may be used for this purpose. Do not place prepared glass feeders in petri dish with standing water as the water may enter the feeder through the membrane and dilute the blood.
- Briefly turn off the water pump and use plastic tubing to connect all feeders in a circuit that leads back into the water bath. (Wet each end of each piece of plastic tubing before use to ensure easier fit.) Turn pump back on and ensure
- a steady flow of water around circuit and
- no dripping. Do not, however, place feeders on top of cartons yet, as mosquitoes may begin to drink and fill up on water.
- Use transfer pipette to measure appropriate amount of blood into well of glass feeder. 1 mL -2 mL per feeder is more than enough.
- Place feeders on top of cartons and begin feeding. Optional: tear off a long strip of tape to hold glass feeder firmly in place against roof of mosquito carton.
- Visually inspect mosquitoes to see if they are feeding. Look for females clustered around feeder with bright red full bellies. Ensure that the temperature of the water is approximately 37 °C . Some variability in temperature is permissible but note that parafilm may melt/deform if bath is run too hot.
- Allow mosquitoes to feed for up to 01:00:00 . Return all mosquito cartons to insectary and disassemble water bath apparatus.
- Let glass feeder and membrane soak in 10% bleach solution for at least 00:15:00 . Dispose of membrane and rubber band in trash. Wash glass feeder and let dry.
- Eggs are laid roughly 48 hrs after the blood meal. Collect eggs onto filter paper and continue the process to maintain colony.
1h 15m
C. Obtaining and counting larvae
At 16- 18 hrs post-egg hatching, remove An. stephensi larvae hatch bin.
Transfer a small amount of water from the hatching habitat to a clean bin lid.
Cut the tip of a plastic bulb pipette with scissors, approximately 1cm above the tip, to widen the inlet area of the pipette.
Remove 7-20 larvae at a time by pipetting, transfer to water on clean bin lid, and count the larvae collected into the water drops. Continue quickly, until all larvae are counted, or the number needed for experiments are collected.
Transfer to a clean bin lid containing hatch bin water.
After larvae are counted, transfer them back to the original habitat, along with the water from the new hatch bin.
D. Phenotypic Characterization of Larvae at each Instar by Advanced Stereo
Microscope (Larval Microscopy)
Cut the tip of a plastic bulb pipette with scissors, approximately 1cm above the tip, to widen the inlet area of the pipette.
Remove 5-7 larvae from their hatch bin, and transfer in one drop to a glass microscope slide.
Analyze larvae using an Olympus SZX16 Advanced Stereo Microscope, at variable magnification settings and determine (a tabulation might help):
- Microscope settings
- Photographic and descriptive phenotypic characterization of larvae at each instar:
- Number of larvae at first instar (L1)
- Characteristics of L1 larvae and on through pupation
Phase 2: Freezing and Recovery of L1 Larvae
5h
Note
for each of these initial conditions, 150-400 larvae are used per condition. Maturation rate
(L1-pupae) is determined for evaluation during initial experiments.
A. Cryoprotectant Selection
Prepare cryoprotectants (or Cryoprotective Agents, CPAs) in nano pure water Room temperature and determine toxicity of cryoprotectants to the L1 larvae. Potential cryoprotectants are listed below.
Potential Cryoprotectants
- 1.5 Molarity (M) ethylene glycol (EG),
- 1.5 Molarity (M) methanol (ML),
- 1.5 Molarity (M) dimethyl sulfoxide (DMSO), or
- 1.5 Molarity (M) methyl acetamide or methyl formamide (MA or MF)
B. Determination of the Toxicity of Cryoprotectants
Collect 50 or more L1 larvae from the insectary bin using a transfer pipette with a blunt end and transfer onto a filter dish.
Lift filter basket with larvae out of the water, blot the sides of the basket to remove excess water, and submerge the basket into a Petri dish containing 1.5 Molarity (M) EG, 1.5 Molarity (M) ML, 1.5 Molarity (M) DMSO, or 1.5 Molarity (M) MA/MF, or water alone (control) and hold for 1, 2, 3, 4, 5, 6, or 08:00:00 at Room temperature (22 °C -25 °C ).
8h
After the appropriate exposure time, lift the basket out of the respective CPA dish and rinse larva by immersion in tap water while they remain in the filter basket.
Repeat step three two more times, or empirically based on loss of glossy sheen on the L1 larvae.
Culture the L1 larvae in water with fish food (as described in Step 2.1) and assess for survival based morphological changes from L1-L4 larvae.
Determine the percent survival (Percent survival = # of pupae collected/initial egg count) for each time point for a given CPA solution.
Record a minimum of three replicates for each CPA to assess toxicity.
C. Freezing larvae
In this example, a mixture of 7 Molarity (M) methyl formamide (MF) in 0.5 Molarity (M) of trehalose was chosen as the optimum CPA. Further, our studies with newly emerged versus late L1 stage larvae supported using 14–24-hour old L1 larvae for freezing. A diagram of the protocol for this process is provided (Figure 2).
Diagram of L1 larvae freezing process
Prepare the CPA solution of 7 Molarity (M) MF in 0.5 Molarity (M) trehalose in a petri dish and place dish on cooling table set at 28 °C .
Place 14-20 hr emerged L1 larvae in clean water. Count larvae.
Transfer larvae to a nylon mesh basket submerged in clean water that is in a square culture dish set on the cooling table. Use ~ 200 L1 larvae per condition group for optimum data sets.
Begin slow cooling on the cooling plate from 28 °C to 0 °C over 00:30:00 .
30m
Lift the basket from the water, blot dry the sides of the mesh basket to remove excess water, and transfer the basket to the Petri dish containing the CPA solution cooled to 0 °C . Hold for 00:05:00 at 0 °C
5m
Begin rapid cooling of L1 Larvae on cooling table, and cool to -15 °C using a rate of 0.65-0.7 degrees / min.
Leave L1 larvae on cooling table (10 min – 2 hr), or transfer to -20 °C freezer (longer time periods) until warming.
D. Warming larvaae
The current warming method is described below. We note that additional warming trials may need to be conducted for further optimization; viability using this method and maintaining freezing conditions for 00:10:00 is 60%.
10m
Remove the petri dish from the cooling table, and hold on a shelf in the refrigerator at 4 °C for 00:10:00 .
10m
Remove the mesh basket and submerge into a petri dish containing water held at 4 °C and incubate for an additional 00:05:00 .
5m
After this time, collect the L1 larvae using a clipped transfer pipette and transfer them to a square dish with water held at 28 °C for 1-02:00:00 . Perform initial viability assessment by watching for movement (including mid-gut movement) using a dissecting microscope.
2h
Return L1 larvae to husbandry conditions for recovery and development through life stages (Figure 1).
Determine recovery and analysis of viability as described below.
E. Viability studies
Determine the number of pupae that emerge from each freezing incubation time point; calculate the percent recovery as done for percent survival, based on number of L1 larvae from each time point.
Determine the percentage of recovered pupae that survive and develop into adults.
Determine the ratio of males to females (sexual differentiation) in adult mosquitoes.
Perform phenotypic similarity analyses (via microscopy; section I.C. of this protocol)
Determine the number of eggs subsequently produced from adults maturing after recovery from cryopreservation.
Determine the number of larvae (F2) hatching from eggs produced by adults that matured after recovery from freezing to confirm the colony has returned to healthy, normal husbandry conditions.