Microinjection Method for Anopheles gambiae Embryos

Summary

Microinjection techniques are essential to introduce exogenous genes into the genomes of mosquitoes. This protocol explains a method used by the James laboratory to microinject DNA constructs into Anopheles gambiae embryos to generate transformed mosquitoes.

Abstract

Embryo microinjection techniques are essential for many molecular and genetic studies of insect species. They provide a means to introduce exogenous DNA fragments encoding genes of interest as well as favorable traits into the insect germline in a stable and heritable manner. The resulting transgenic strains can be studied for phenotypic changes resulting from the expression of the integrated DNA to answer basic questions or used in practical applications. Although the technology is straightforward, it requires of the investigator patience and practice to achieve a level of skill that maximizes efficiency. Shown here is a method for microinjection of embryos of the African malaria mosquito, Anopheles gambiae. The objective is to deliver by microinjection exogenous DNA to the embryo so that it can be taken up in the developing germline (pole) cells. Expression from the injected DNA of transposases, integrases, recombinases, or other nucleases (for example CRISPR-associated proteins, Cas) can trigger events that lead to its covalent insertion into chromosomes. Transgenic An. gambiae generated from these technologies have been used for basic studies of immune system components, genes involved in blood-feeding, and elements of the olfactory system. In addition, these techniques have been used to produce An. gambiae strains with traits that may help control the transmission of malaria parasites.

Introduction

Microinjection techniques have been used to experimentally manipulate organisms since the early 1900s¹. Microinjection has been used to study both basic biological functions and/or introduce important changes in the biology of a desired organism. The microinjection technique has been of particular interest to vector biologists and has been widely used to manipulate vector genomes^2-11. Transgenesis experiments in arthropod vectors often aim to make vectors less efficient at transmitting pathogens by either enacting changes that decrease a vector's fitness or increase refractoriness to the pathogens they transmit. Mosquitoes transmit a variety of human pathogens and have a significant impact on morbidity and mortality worldwide. The Anopheles genus of mosquitoes transmits the human malaria parasitic pathogens, Plasmodium spp. Genetic engineering experiments with Anopheles have aimed to better understand the biology and reduce the vectorial capacity of these mosquitoes in efforts to develop novel malaria elimination strategies.

The mosquito vectors that contribute the most malaria infections worldwide are in the Anopheles gambiae species complex. However, the majority of successful transgenesis experiments have been performed on the malaria vector of the Indian subcontinent, Anopheles stephensi. While plenty of laboratory-adapted Anopheles gambiae strains exist, the number of transgenic Anopheles gambiae spp. lines reported in the literature does not compare to that of Anopheles stephensi. It is thought that the Anopheles gambiae embryo is more difficult to inject and achieve successful transgenesis than Anopheles stephensi, although the reasons for these differences are unknown. This protocol describes a method that has been proven to be consistently successful in achieving transgenesis of Anopheles gambiae embryos via microinjection. The protocol is based on a method previously developed by Hervé Bossin and Mark Benedict¹² with some additional details and alterations added that have been found to increase the efficiency of transgenesis.

Protocol

1. Preparing mosquitoes for microinjection

1. Seed a cage¹³ (~5000 cm³) with ~100 male and 200-300 female 1-2 day adult post-eclosion mosquitoes and allow them to mate for 2 days.
2. After the mating period, provide mosquitoes a blood meal using either 2 mL of blood with an artificial feeding device or live anesthetized animals depending on insectary practices¹⁴. The following day provide mosquitoes a second blood meal to ensure that all mated females have had an opportunity to feed and that partially-fed mosquitoes have become fully-engorged.
3. Utilize the mosquitoes for embryo collections 2-4 days after the second blood meal.
   ​NOTE: Larval nutrition is important for adult robustness and reproductive fitness. See Benedict et al. (2020) for information about how to rear healthy insects¹⁴. No significant differences in egg laying have been observed when mosquitoes are fed on an artificial feeder or live anesthetized animals. Use whichever method is routinely used for Anopheles gambiae in the insectary.

2. Embryo preparation

1. Use an aspirator to place 20-30 females in a transparent 50 mL conical tube that has been cut (with a band saw) to be open at both ends and is covered with latex dental film at one end and nylon mesh and filter paper at the other (Figure 1).
   NOTE: The mosquitoes will deposit their eggs onto the filter paper and the nylon mesh will keep the filter paper securely in place. Mosquito eggs are cylindrical in shape, ~500 µm in length, ~200 µm in diameter at their widest point, and tapering at the anterior and posterior ends (Figure 2).
2. Place the mosquito-filled tube in a small (60 mm x 15 mm) Petri dish filled with double-distilled water (ddH₂O). Place the tube and dish in an incubator at 28 °C for 45 min (Figure 3).
3. Remove the tube from the incubator and insert the tube into an empty cage. Gently tap the tube to allow mosquitoes to fly out and remove the tube from the cage once all mosquitoes have flown out.
4. When the tube is free of mosquitoes, unscrew the bottom ring, remove the nylon, and use forceps to carefully remove the filter paper with the eggs from the tube. Place the filter paper in a plastic Petri dish (100 mm x 15 mm) containing a layer of filter paper moistened with ddH₂O.
5. Observe the eggs. Eggs that have aged to the point where they are light gray in color (Figure 4) are ready for alignment.
   1. If eggs are still white, return them to the incubator and check the color every 5 min. White eggs are fragile, rupture easily, and do not survive well after the injection process resulting in low hatching.
   2. Eggs that are dark gray or black have aged too much. Do not use them.

3. Embryo alignment

1. Cut a piece of nylon blotting membrane (2 cm x 1 cm) with a razor blade, making sure that the edge is neatly trimmed.
   NOTE: If the edge is not straight, the eggs will not stay affixed to the membrane properly during the injection process.
2. Put a membrane on a glass slide and cover the membrane with a piece of filter paper (2 cm x 2 cm), leaving ~1 mm of the membrane filter uncovered (Figure 5).
   NOTE: Ensure microscope slides are clean before use and use clean forceps to manipulate the membrane filter.
3. Wet the filter paper with deionized H₂O as eggs will die if desiccated for prolonged periods of time.
   NOTE: Do not put too much water on the paper because the embryos will move during the injection process, but make sure that there is enough to prevent the embryo from drying (Figure 6A, B). Excess water can be removed by absorbing it with a piece of filter paper.
4. Gently transfer 30-50 embryos to the edge of the membrane with a paintbrush (size #0). Use the brush to align the eggs vertically along the membrane by gently rolling them over on the slide so that the ventral side (convex) is facing upward.
5. Orient all the eggs in the same direction so that when the eggs are observed under the microinjection microscope, the posterior end (more pointed, Figure 6A, B) is in a down position and forms an angle of ~15° with the needle (Figure 6C). Line the entire edge of the membrane with eggs (30-50) and place the slide under the microscope for injection.
   NOTE: Choose the eggs carefully. Discard those that are white (too young or undeveloped) or dark gray (too old and will break the needle), and the abnormal ones (Figure 7). Make sure the filter paper remains moist at all times.

4. DNA preparation

1. Purify plasmid DNAs for injection, at the minimum, with an endotoxin-free DNA plasmid maxiprep kit following the manufacturer's protocols.
2. Resuspend the DNA in PCR-grade water and centrifuge at 17,100 x g in a refrigerated centrifuge for 30 min at 4 °C. Then, transfer the supernatant to a new, clean 1.5 mL conical tube. Carefully micro-pipette to avoid transferring undesired particulate matter from the column.
3. Perform a second precipitation of DNA using one-tenth volume of 3M sodium acetate and a two-fold volume of 100% ethanol. 
4. Resuspend the DNA in 300-500 µL of PCR-grade water for a final plasmid concentration of 1000 ng/µL. Mix the plasmid cocktail in the injection buffer for a final plasmid concentration of 300-400 ng/µL and make 10-20 µL aliquots.
   ​NOTE: DNA can be stored at -20 °C. DNA microinjection aliquots can be stored at -20 °C, or at -80 °C if using RNA components. Do not exceed 800 ng/µL of DNA in the injection mix because viscosity will cause the needles to clog.

5. Needle preparation

1. Make the needles by pulling 10 cm quartz capillaries using a programmable micropipette puller.
2. Examine the needle under microscope to ensure that the tip is formed well. Ensure that the needle setting on the programmable puller yields a tip that starts to taper ~0.5 mm from the terminal end and ends in a fine point (see Figure 6). Needles that break easily usually have too long of a taper.
   NOTE: Conditions might differ depending on the needle puller (The authors will make available on request the settings for the programmable puller they use. Journal restrictions prevent us from citing a specific brand). Needles can be pulled by hand, but this requires a skill-set not available in most laboratories.

6. Embryo microinjection

1. Use a micro-loader tip to fill the needles with 2 µL of DNA mixture. Insert the needle into the needle holder and connect the automated pressure pump tubing (Figure 8).
2. Important: Align the needle so that it makes an angle of 15° with the plane of the slide (Figure 8).
3. Open the needle tip by carefully touching the first egg and inject it by inserting the tip of the needle ≤ 10 µm in the posterior pole. A successful injection will lead to a small movement of the cytoplasm within the egg.
4. Use the microscope coaxial stage controls to move to the next egg to continue injection.
   1. To ensure that the needle tip remains open and has not clogged, press the Inject button before entering another embryo and visualize the small droplet at the opening of the needle.
   2. If the needle gets clogged, press the Clear button to clear the needle and repeat the droplet visualization test. Adjust the pressure as needed if the needle tip opening gets slightly bigger to ensure that the size of the droplet stays small.
   3. Inject ~40-50 eggs with one needle.
      NOTE: Make sure the filter paper stays moist at all times. Keep sufficient back-pressure on the needle to keep it cleared. A needle that cannot be unclogged must be replaced with a new needle. Embryo placement, needle insertion and injection are made easier with microscopes that have coaxial controls for horizontal movement of the stage (Figure 9).
5. After injections are complete, rinse the eggs off into a glass container lined with filter paper and filled with 50 mL of deionized H₂O (Figure 10).
   NOTE: Embryos will start hatching 2 days post-injection and may take as long as 3-5 days. Hatched first instar larvae must be transferred immediately (check 2 times daily) to a clean container with water and food.

Results

A representative example of the application of the microinjection protocol described can be found in Carballar-Lejarazú et al⁵. The intent here was to insert an autonomous gene-drive system into the germline of a laboratory strain, G3, of An. gambiae. The system was designed to target the cardinal ortholog locus (Agcd) on the third chromosome in this species, which encodes a heme peroxidase that catalyzes the conversion of 3-hydroxykynurenine to xanthommatin, the last st...

Discussion

With the increased availability of precise and flexible genetic engineering technologies such as CRISPR/Cas9, transgenic organisms can be developed in a more straightforward and stable way than previously possible. These tools have allowed researchers to create transgenic strains of mosquito vectors that are very close to achieving the desired properties of either refractoriness to pathogens (population modification) or heritable sterility (population suppression). However, to develop the most safe and stable genetically...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
10x Microinjection Buffer	-	-	1 mM NaHPO4 buffer, pH 6.8, 50 mM KCl
Blotting membrane (Zeta-Probe GT Genomic Tested Blotting Membrane)	Bio-Rad		Neatly and straightly cut into 2x1 cm piece
Conical tubes 50 ml (disposable centrifuge tube, polypropylene)	Fisher Brand		Ends cut
De-ionized or double-distilled water (ddH20)	Mili-Q		In a wash bottle
Dissecting microscope	Leica	Leica MZ12	For embryo alignment
Forceps			No. 5 size
Glass container	Pyrex	No. 3140	125 x 65
Glass slide	Fisher Brand	No. 12-549-3	75x26 mm
Incubator	Barnsted Lab-line	Model No. 150	28 °C
KCl			50 mM
Latex dental film	Crosstex International	No. 19302
Microinjector	Sutter Instrument		XenoWorks Digital Microinjector
Microloader Pipette tips	Eppendorf		20 µL microloader epT.I.P.S.
Micromanipulator	Sutter Instrument		XenoWorks Micromanipulator
Micropipette	Rainin		20 µL
Micropipette puller	Sutter Instrument	Sutter P-2000 micropipette puller
Microscope	Leica	DM 1000 LED or M165 FC	For microinjection
Minimum fiber filter paper	Fisher Brand	No. 05-714-4	Chromatography Paper, Thick
Mosquitoes	MR4, BEI Resources		Anopheles gambiae, mated adult females, blood-fed 4-5 days post-eclosion
NaHPO4 buffer			1 mM, ph 6.8
Nylon mesh
Paint brush	Blick	No. 05831-7040	Fine, size 4/0
Petri dish			Plastic, (60x15 mm, 90x15 mm)
Sodium acetate			3M
Quartz glass capillaries	Sutter Instrument	No. QF100-70-10	With filament, 1 mm OD,  ID 0.7 10 cm length
Water PCR grade	Roche	No. 03315843001