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: December 01, 2023
Last Modified: May 20, 2024
Protocol Integer ID: 91954
Funders Acknowledgement:
National Science Foundation
Grant ID: IOS-1911723
Abstract
The collection, fixing, and immunohistochemical staining of Aedes aegypti embryos is challenging in comparison to D. melanogaster since the vitelline membrane of Ae. aegypti must be manually removed. Herein we report on an improvement for the methods to prepare Ae. aegypti embryos. The adapted protocol increases the throughput capacity of embryos by an individual user, with experienced users able to process an average of 100-150 embryos per hour. The protocol provides high-quality intact embryos that can be used for morphological, immunohistochemical, and in situ RNA hybridization studies. Critical to the success of the optimized protocol is the selection and description of the tools required, and differing approaches for younger and older embryos.
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Safety warnings
This protocol contains hazardous chemicals including formaldehyde, heptane, methanol, and animal blood. Appropriate PPE and institutional-specific safety precautions should be taken. (PPE was occasionally removed during production of the videos to improve visualization.)
Before start
This protocol was developed using the Liverpool strain of Ae. aegypti reared under optimal conditions at 27°C, 80% RH and a 12:12 (L:D) photoperiod. Larvae were reared in 14 x 8 x 5 “ containers and fed a slurry of Tetramin fish food (Spectrum Brands Pet, LLC, Blacksburg, VA) ad libitum. Adults were provided with raisins as a sugar source.
Preparation of an aspirator
Preparation of an aspirator
Cut both ends from a 10 mL serological pipette. Save the end with the cotton plug.
Trim the tapered tip and remove any sharp edges.
Whittle the thinner plastic end of the serological pipette where the cotton plug is.
Fix two layers of chiffon fabric as a blocking filter by inserting the shaved end of the pipette tip into the pipette.
Attach a length of tygon tubing for mouth aspiration.
Preparation of an egging chamber
Preparation of an egging chamber
Remove the threads from the cap and tube of a 50 mL conical tube.
Create an entry port for mosquitoes by drilling a hole that is slightly larger than the outer diameter of a 10 mL serological pipette into the 50 mL conical tube.
Use a strip of laboratory tape to create a removable closure over the entry port.
Place a second smaller section of tape on the back of the tape closure so that when the mosquito entry port is closed, there is no sticky side of the tape facing the inside of the 50 mL conical tube.
Electrosharpen a tungsten wire
Electrosharpen a tungsten wire
Cut a 3-4 cm length of 0.005” tungsten wire.
Secure the tungsten wire in an inoculating loop holder.
Connect the anode (negative terminal) of a 9V battery to a 1.0M solution of KOH or NaOH.
Connect the cathode (positive terminal) of the 9V battery to the metallic end of the inoculating loop holder.
Repeatedly dip the end of the tungsten wire into the 1.0M solution of KOH (or NaOH) to electrosharpen the tungsten wire.
Collection of embryos
Collection of embryos
1h
Place 250-300 Ae. aegypti pupae into a cage at a sex ratio ranging from 1:1 to 2:1 (F:M).
Provide the mosquitoes with a fresh blood meal three to five days post adult emergence.
Four days post blood meal, use the aspirator to transfer 20-50 females into an egging chamber lined with three overlapping strips of wetted Whatman #1 filter paper.
Darken the conical tube by covering the tube with aluminum foil. Allow the mosquitoes to lay eggs undisturbed for 01:00:00.
1h
Transfer the egg paper to a plastic petri dish lid lined with two layers of wetted Whatman #1 filter paper and close the lid with the bottom of the dish.
Incubate the embryos at 27 °C for the desired duration of time. Make sure the embryos are positioned on a slight incline so that they are not submerged in water.
Fixation of embryos
Fixation of embryos
36m
Prepare the following solutions for fixing embryos: 20 mL 50% bleach (Final NaClO = 3%), 10 mL PEM-F (prepared fresh), 10 mL MeOH, 10 mL heptane.
Set up an egg basket (i.e., 100 µm cell strainer) in a 9-cm-diam Petri dish bottom and fill the dish bottom with distilled water.
Dislodge the embryos from the edges of the filter papers with a fine tipped paintbrush.
Rinse embryos into the egg basket with distilled water.
Dechorionate embryos in 50% bleach (3% NaClO) for 00:00:30.
30s
Generously rinse embryos in distilled water three times.
Rinse embryos from the egg basket into a scintillation vial.
Remove excess water from the scintillation vial and add 10 mL of PEM-F.
Incubate embryos in PEM-F at 60 °C for 00:30:00.
30m
Remove the PEM-F and add 10 mL of heptane.
Transfer embryos to a -70 °C freezer for 00:05:00.
5m
Add 10 mL methanol and shake the vial under running hot (60 °C) water for 00:00:30.
30s
Remove the upper heptane layer and most of the methanol from the embryos.
Add an additional 10 mL methanol to the scintillation vial.
Transfer embryos to a 1.5 mL tube.
Wash embryos with three changes of 1 mL of 100% methanol. Embryos can be stored in methanol at -20 °C. Alternatively proceed with the rehydration step.
Removal of the vitelline membrane (“embryo peeling”)
Removal of the vitelline membrane (“embryo peeling”)
30s
Rehydrate the embryos using a methanol/PBS(P) dilution series (1 mL and 10 min each): 75%, 50%, 25%, 100% PBS(P), PT.
Replace the final rehydration with a new 1 mL aliquot of PT and store embryos Overnight at 4 °C prior to peeling.
30s
Wet a 1”x2” square of Whatman #1 filter paper with PBS(P) and place on a smooth glass surface.
Add a glass coverslip so that it slightly overlaps onto the filter paper.
Flood the underneath of the coverslip with PBS(P) so that a complete “water bridge” is formed. Wick off any excess PBS(P).
Affix an equal-length strip of double-faced tape to the edge (1-2 mm overlapping) of a plastic coverslip.
Trim the other edge of the double-faced tape to leave a clean edge.
Peel back a corner of the tape backing and place the prepared transfer tape aside.
Add 50-100 fixed embryos along the edge of the glass coverslide overlapping the wetted filter paper.
Gently brush embryos to the edge glass coverslip.
Use a fine tipped paintbrush to position the posterior (thinner end) of the embryo towards the edge of the glass slide and the anterior (wider end) away from the glass slide.
Continue to align embryos tightly against each other until all embryos have been aligned.
Remove the PBS from underneath the glass coverslip by pressing paper toweling onto the filter paper.
Remove the glass coverslip and gently move the line of embryos towards the edge of the glass plate by sliding the paper across the glass surface without lifting.
Transfer the embryos to the double faced tape so that the anterior of the embryos slightly overhangs the outer edge of the tape. Gently press on the backing of the double faced tape to ensure that the embryos are affixed to the tape.
Remove the tape backing and place the tape into the well of a glass well slide (embryo side up).
Flood the well with PBS(P). Add PBS(P) as needed to prevent the embryos from drying out.
Peeling older embryos (>20h AEL at 27°C)
Peeling older embryos (>20h AEL at 27°C)
Make a small incision at the anterior of each embryo using a fine stab knife.
Carefully slip the anterior end of the vitelline membrane off of the embryos by gently inserting the tungsten wire underneath the membrane at the hole/seam and pushing forward with a very slight upward pressure. The main pressure of the tungsten wire should be forward with a very slight upward pressure.
Remove the anterior end of the vitelline membranes from all of the embryos.
Rinse the loose vitelline membrane fragments from the slide with a gentle stream of PBS(P), then refill the reservoir with clean PBS(P).
Using the blunt Roboz probe, gently push on the posterior end of the embryo until it extrudes out of the membrane.
Move and press any extruded embryos retaining a serosal cuticle into the sticky surface of the double faced tape.
Tear the serosal cuticle at the anterior end of the embryo.
Extrude the embryo from the serosal cuticle by pushing on the posterior end of the embryo with the blunt side of the tungsten wire probe.
Collect embryos into a pile and transfer into a 1.5 mL tube using a Pasteur pipette.
Peeling younger embryos (<20h AEL at 27°C)
Peeling younger embryos (<20h AEL at 27°C)
Tear a small opening in the posterior of the embryo using the tungsten wire probe.
Insert the tungsten wire probe into the posterior opening of the vitelline membrane and carefully tear upwards in a stepwise fashion until the entire vitelline membrane has been dissected along the sagittal plane.
Press the edges of the vitelline membrane into the tape adhesive to free the embryo.
Collect embryos into a pile and transfer into a 1.5 mL tube using a Pasteur pipette.
Store all peeled embryos at 4 °C and process for staining within one day for best results.
Protocol references
1. Clemons A, Haugen M, Flannery E,Kast K, Jacowski C, Severson D, Duman-Scheel M. 2010. Fixation and Preparation of Developing Tissues from Aedes aegypti. Cold Spring Harb Protoc. doi:10.1101/pdb.prot5508.
2. Schember I, Reid W, Halfon MS. 2023. Conserved and novel enhancers regulate expression of Aedes aegypti single-minded in the embryonic ventral midline. bioRxiv 2023.08.01.551414; doi: https://doi.org/10.1101/2023.08.01.551414.