Anopheles Egg Collection, Disinfection, and Hatching
- 1Entomology Branch, Division of Parasitic Diseases and Malaria, Centers for Disease Control and Prevention (CDC), Atlanta, Georgia 30333, USA
- 2CDC Foundation, Atlanta, Georgia 30308, USA
- ↵4Correspondence: mbenedict{at}cdc.gov
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↵3 These authors contributed equally to this work.
Abstract
Gravid (i.e., with fully developed eggs), mated Anopheles females typically lay their eggs directly on water ∼48–72 h after a blood meal. Unlike some other mosquito species, Anopheles eggs cannot be desiccated and stored for long durations, and, hence, colonies must be reared continuously. In this protocol, we discuss methods for egg collection, including individual and en masse oviposition; egg disinfection to avoid the transmission of infectious agents to the next generation; and egg hatching for colony maintenance or experimentation. We also include optional methods for estimating life history traits such as fecundity, fertility, and larval mortality rates from egg counts.
MATERIALS
Reagents
Anopheles mosquitoes in a cage (gravid, mated females that have been housed with males for several days)
H2O (refer to the Water Types section in the Introduction: Considerations for Rearing and Maintaining Anopheles in the Laboratory [Leite et al. 2023a])
Hot H2O (>70°C)
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Ideally, an insectary would have an InSinkErator or comparable instant hot H2O tap installed. An electric tea pot is a convenient alternative.
Household bleach solution (1%–2%, v/v, in H2O)
Larval food (2%, w/v, brewers or baker's yeast or 25 mg of ground koi food)
Roccal-D and bleach disinfectant solution (1%–2%, v/v, of household bleach solution plus 0.7% Roccal-D) (optional; see Step 10)
Sugar H2O (10%, w/v)
Equipment
Camera (see Step 18)
Container with a secure lid (any container that can hold ∼50 mL of H2O)
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The lid should have an opening covered with a fine mesh. Alternatively, the container can be covered with fine mesh and an elastic band.
Cotton balls
Dissecting microscope
Dissecting needle (e.g., Fisherbrand 08-960A)
Elastic bands
Environmental chamber or room with temperature, humidity, and light controls
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Maintain mosquitoes in an environmental chamber at ∼27°C, 80% relative humidity, and a 12 h light:12 h dark cycle, with or without a sunrise and sunset period.
Filter paper (e.g., Whatman Grade 3 Qualitative, 90-mm-diameter circle, Sigma-Aldrich WHA1003090)
Fine mesh (any commercially available mosquito netting)
Forceps
Image software program (e.g., egg counter [Mollahosseini et al. 2012] or ImageJ).
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ImageJ is free software that allows the researcher to count eggs one by one using the multipoint tool that adds a number every time a mouse click is made on the image.
Mechanical counter (two-place denominator; optional, see Step 18)
Pencil (optional; see Step 18)
Petri dish (90-mm-diameter)
Rearing pan with lid (lid is optional though strongly recommended; e.g., US Plastics 52051)
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See Introduction: Considerations for Rearing and Maintaining Anopheles in the Laboratory (Leite et al. 2023a) for more information
Scissors
Sealed container (see Step 14)
Sponge cloth (optional; see Step 17)
Squat, wide-opening plastic cups (∼250-mL)
Stainless-steel forceps
Suction device to collect adult mosquitoes (e.g., Fulton RHM 200 aspirator)
Vacuum filtration system
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A Nalgene 500-mL vacuum filtration system can be adapted to this method by breaking off the upper chamber and removing the membrane. Connect it to a vacuum system with a trap, which is composed of an Erlenmeyer filtration flask capped with a two-holed stopper. Insert the tube from the Nalgene filter into one hole and connect the tube to vacuum source to the side of the flask. Use a blue grinding pestle or similar object to plug the second hole to allow for easy release of the vacuum pressure.
Wash bottle (500-mL; e.g., Sigma-Aldrich Z423106)
METHOD
Egg Collection
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Mated female mosquitoes are typically ready to lay eggs 48–72 h after taking a blood meal, with some variability among species. Gravid (with fully developed eggs), mated Anopheles females lay individual eggs directly on the water or damp filter paper in the laboratory, mostly at night (dark cycle). En masse oviposition and hatching are the most used methods to maintain a mosquito colony in the insectary, whereas individual ovipositions are more common when few females are available or for experimental activities such as genetic crosses or when life history traits are being determined.
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1. Provide an oviposition substrate to gravid female Anopheles mosquitoes.
Single-Mosquito Oviposition
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i. Line the wall of a 50-mL container with a strip of filter paper and fill with H2O up to 0.5–1 cm above the bottom of the filter paper (Fig. 1A).
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ii. Cover the opening of the container with fine mesh secured with an elastic band, cotton ball, or other similar item.
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iii. With scissors, create a small hole on the mesh (one that can easily be covered with a cotton ball or a piece of tape once the mosquito is in the cup).
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iv. Using an aspirator, remove a female from the cage, carefully transfer it into the container through the small hole in the mesh, and plug the container with a cotton ball.
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v. Let the females oviposit overnight (dark cycle, ∼12 h)
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This step does not have to occur during the night. A laboratory may have the dark cycle occurring during the actual day, which will also stimulate females to oviposit.
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For one night only, providing sugar H2O is usually not necessary.
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vi. On the following day, examine the container(s) for the presence of eggs. With an aspirator, remove the female from tubes containing eggs. Allow the remaining females an opportunity oviposit another night and provide them sugar H2O.
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If checking fecundity or fertility parameters, allow eggs to hatch (generally ∼2–3 d after oviposition occurs) and continue to Step 17; otherwise, proceed to Protocol: Anopheles Larval Rearing (Leite et al. 2023b) for information on how to rear larvae.
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Types of containers for egg collection. (A) Plastic cup lined with filter paper used for individual females. The cup is filled with enough H2O to ensure the filter paper remains wet. Mesh at the top of the container allows the female to have access to cotton wetted with sugar H2O if necessary. (B,C) To collect eggs en masse, a cup with H2O and lined with filter paper can be placed in a cage with gravid and mated female mosquitoes or (D,E) a filter paper can be placed on top of sponges soaked in H2O contained in a Petri dish. (F–I) Alternatively, a 9-cm-diameter round filter paper can be cut from the edge to the center to create a funnel. The funnel is then placed on top of a cup filled with H2O touching the paper on which females will land and oviposit.
En Masse Oviposition
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vii. Line the sides of a labeled plastic cup (∼250-mL) with filter paper (Fig. 1B,C) and fill to a depth of ∼1.0 cm of H2O such that the paper is halfway covered with H2O.
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Do not leave gaps in the paper because eggs will stick to many plastics and dry out.
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viii. Place the cup into a cage with gravid females.
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ix. Allow the females to oviposit “overnight” (dark cycle, ∼12 h).
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This step does not have to occur during the night. A laboratory may have the dark cycle occurring during the day, which will also make females oviposit.
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x. Remove the egg cup from the cage. Take care that no adult mosquitoes escape during this step, by puffing briskly down the cage sleeve as the egg dish is being pulled out.
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xi. Cover the egg cup with a Petri dish and proceed to Step 2.
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Egg Disinfection
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Although it is possible to disinfect eggs collected from individual females, this method is typically used only for en masse collections. Disinfecting eggs with bleach is a key step to keeping healthy Anopheline colonies for many generations, as it can inhibit the growth of harmful microbes (especially microsporidia) and prevent their transfer between generations. Use the Roccal-D solution along with bleach solution when a colony infection is suspected, as bleach solution alone does not prevent infection.
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2. Ensure that all the necessary materials (Fig. 2A) are clean and available before starting.
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3. Remove any dead adult mosquitoes from the oviposition cup(s) or container(s) with clean forceps and wipe the adults from the forceps with a clean cotton ball. Use a fresh cotton ball for each oviposition cup.
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4. Kill any eggs remaining on the forceps by submerging the forceps in hot H2O (>70°C) before and after removing the adults from oviposition cups to prevent colony contamination. If possible, use another clean pair of forceps for each cup.
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5. Turn on the vacuum filtration system at low suction.
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6. Center a fresh filter paper on the platform with the vacuum applied.
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7. Wet the filter paper with purified H2O using a wash bottle. Increase the vacuum suction until the filter paper disk forms a depression.
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It may be necessary to press the paper down gently to close gaps and achieve suction.
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8. Slowly pour H2O and eggs onto the center of the disk, taking care that the eggs do not spill outside the depression. Rinse the sides of the oviposition cup with purified H2O via the wash bottle until the desired number of eggs are on the filter paper (Fig. 2B).
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9. Release the vacuum.
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10. Add the bleach solution (or, if necessary, Roccal-D and bleach disinfectant solution) to the depression (Fig. 2C). Allow the eggs to sit in the bleach for 30 sec.
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11. Apply the vacuum to drain the liquid completely.
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12. Release the vacuum and rinse the eggs by adding purified H2O to the entire depression. Allow eggs to sit for 3–5 sec.
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13. Apply suction and drain the H2O completely.
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14. Rinse the eggs a third time with H2O if disinfecting with the Roccal-D and bleach disinfectant solution.
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15. Remove the damp egg paper and place it in a labeled sealed container large enough to fit the filter paper (Fig. 2D). Secure the lid. Place in an environmental chamber at the standard rearing conditions.
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16. Store the eggs for up to 24 h.
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The 24 h holding time will allow embryos to develop completely before hatching and promotes simultaneous hatching when the eggs are transferred to H2O. Longer storage periods can reduce embryo viability; however, A. gambiae eggs have been stored at 4°C and 48% ± 2% relative humidity for up to 5 d without affecting egg hatching rates, larval development and survival, and adult sex ratio (Mazigo et al. 2019). Trials with other species and strains are recommended before using these procedures.
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Egg disinfection and hatching. (A) Components of an egg bleaching station. (B) Rinsing eggs from the oviposition cup onto the filter paper using the vacuum filtration system. (C) Floating eggs in a bleach solution for disinfection. (D) Clean eggs on filter paper placed in a holding container for storage before hatching. (E) Setup for hatching on sponge cloth. A filter paper containing eggs is placed on top of a raised bed of sponge cloth, allowing the eggs to stay moist but contained and the larvae to wriggle off. (F) Example of an infertile egg (egg 1) and fertile eggs that are either unhatched (egg 2), hatched (egg 3 and 4), or in the process of hatching (larva eclosing, egg 4). The open operculum is observed in eggs 3 and 4.
Egg Hatching for Colony Maintenance or Experimentation
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Larval eclosion, or egg hatching, takes place after embryogenesis is complete, which normally occurs 2–3 d postoviposition (Clements 1992; Yaro et al. 2006). The duration of embryonic development can vary by species and other external factors, such as the presence of certain chemical components in the H2O and temperature, in particular, during early embryonic development (Cardozo Gonçalvez de Carvalho et al. 2002; Yaro et al. 2006; Impoinvil et al. 2007). Agitation-induced egg hatching has also been investigated in A. gambiae, and it has been shown that early hatchers produce smaller larvae than those that hatch later (Yaro et al. 2006).
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17. Provide a rearing pan containing H2O for the eggs to hatch and a food source for the newly emerged larvae (2% brewers or baker's yeast or 25 mg ground koi food).
General En Masse Hatching for Colony Maintenance
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i. Place a larval rearing pan filled with 500 mL (∼1–1.5 cm in depth) of rearing H2O and food on the shelf where the larvae will be reared. If yeast is used, add 1 mL of 2% brewer's or baker's yeast/500 mL of H2O.
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Place the pan on the shelf before adding eggs. If a pan with H2O and eggs is moved, many eggs will be stranded on the sides of the pan where they will dry out and die unless they are washed back onto the water using a wash bottle.
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ii. Wash the eggs off the filter paper with H2O into the larval pan (see Protocol: Anopheles Larval Rearing [Leite et al. 2023b]).
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iii. Cover the pan with a lid.
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Continue to Step 18 if counting eggs; otherwise, proceed to Protocol: Anopheles Larval Rearing (Leite et al. 2023b) for information on how to rear larvae.
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Hatching on a Sponge Platform
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This method is suitable for separating eggshells from larvae and is especially useful when the number of eggs, hatching rate, or timing needs to be analyzed.
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iv. Place a larval rearing pan filled with 500 mL (∼1–1.5 cm in depth) of rearing H2O on the shelf where the larvae will be reared.
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v. Carefully place a filter paper containing eggs (bleached or unbleached) on a raised platform consisting of a stack of sponge cloth disks slightly higher than the H2O level (Fig. 2E).
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This will provide enough moisture for larvae to hatch while preventing the eggs from spreading across the H2O. The larvae will wriggle off the paper. It is important to use a smooth sponge cloth that prevents larvae from crawling underneath the filter paper rather than swimming away.
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vi. (Optional) After hatching is complete, collect the paper with the eggshells and unhatched eggs without disturbing the larvae for the tracking of parameters to estimate several life-history traits of the colony. To assess these parameters, proceed with Step 18; otherwise, proceed to Protocol: Anopheles Larval Rearing (Leite et al. 2023b) for information on how to rear larvae.
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Counting Eggs
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18. If needed for experimental data, count the total number of eggs and determine hatching rate to assess fecundity and/or fertility (see Steps 19 and 20). Perform egg counts with one of the following options.
Manual Counting
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i. Lay the egg paper(s) on a smooth nonporous surface such as a Petri dish under a dissecting microscope.
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ii. (Optional) To make counting easier, draw sections on the filter paper with a pencil in a pattern that makes it possible to ensure all eggs are counted once.
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iii. Move the filter paper to each cluster of eggs, pricking them with a dissecting needle if necessary to determine whether the egg has hatched (see Fig. 2F for examples of unfertilized, fertilized/unhatched, and fertilized/hatched eggs).
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iv. Record the status of the egg hatch of the desired number of eggs on the denominator.
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A mechanical counter can be used when the number of eggs is high.
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Automated Counting
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This method of counting is not recommended if hatch rate needs to be assessed because hatching rate cannot be reliably determined from a photograph.
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v. Lay the filter paper on a flat, evenly illuminated nonporous surface.
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This step will help to better identify eggs from other debris when counting.
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vi. Take a photo with a camera, filling the photograph with the eggs but not missing any and ensuring good focus over the entire field.
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Although a microscope camera can be used, a cell phone camera is adequate. Whichever option is chosen, perform preliminary tests to ensure all eggs are in focus.
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vii. Use an image software program such as ImageJ to determine the total number of eggs.
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Estimating Life-History Traits
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19. Estimate fecundity, which is the number of hatched eggs per female, by dividing the total number of hatched eggs laid on a filter paper by the number of gravid females in the cage.
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To determine fertile, infertile, hatched, or unhatched, see Steps 20 and 21 for more details. This estimate should be considered a population estimate because it does not consider whether all females may not have mated or blood fed.
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20. Determine fertility rate by dividing the number of fertile (hatched or unhatched) eggs on the filter paper by the total number of eggs laid.
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Because some fertile eggs may not have hatched by the time the eggs are analyzed, distinguishing between infertile and fertile/unhatched eggs is essential for a good estimation of this parameter. Use the following visual cues to determine egg fertility:
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Color: Fertile eggs are dark brown (Fig. 2F, eggs 2–4), whereas infertile eggs are lighter in color (Fig. 2F, egg 1).
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Bursting: If the egg was fertile and the larva hatched, a visibly loosened cap (the operculum) on the anterior pole of the egg is usually observed (Fig. 2F, eggs 3 and 4). If uncertain, prod the egg with a dissecting needle.
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Contents: If the cap is not loose and the color of the egg is not pale or is uncertain, apply slight pressure on the egg with a dissecting needle to observe the contents. If the egg is fertile, a larva will be present (Fig. 2F, egg 4), whereas a watery yolk will come out of infertile eggs.
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Some mosquito strains may have a delayed hatching. Thus, if the fertility rate at a particular time point is needed, unhatched but fertile eggs should be considered separately from fertile and hatched eggs for the calculation.
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21. To estimate larval mortality at a time point, count the total number of larvae alive at a particular larval instar and divide by the total number of eggs that hatched.
This calculation will provide the number of larvae that eclosed but died afterward. Thus, larval mortality at a particular time point can be determined. Early-instar larval mortality may be difficult to assess because dead larvae may be eaten by other larvae and decay rapidly; therefore, determining later-instar larval mortality with this method is recommended.
ACKNOWLEDGMENTS
We thank all members of the MR4 Vector Activity (Malaria Research and Reference Reagent Resource Center, BEI Resources) and Target Malaria Teams who contributed to this protocol by providing technical information on rearing methods, some of the images, and general insectary support. In particular, we thank Catherine Steele, Ilsse Ortega, and Katelyn Cavender. We also thank Michael Aidoo and Audrey Lenhart for proofreading this manual and providing valuable feedback.
Footnotes
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From the Mosquitoes collection, edited by Laura B. Duvall and Benjamin J. Matthews.
