Microinjection

BI379 at MBL Lab

Microinjection can be used to deliver any solution into insects. In our lab, its most common applications are in RNAi, CRISPR, and drug treatments. This protocol will not cover the preparation of those reagents, only the injection process. We will cover injection of beetle (e.g. Tribolium or Gnathocerus) larvae, and adults, juveniles and eggs of true bugs (e.g. Oncopeltus or Jadera).

General considerations

Insect care

Ethically, we have a duty of care to the animals involved in these experiments to minimize their discomfort. Take good care of the insects in experiments. Be sure you follow through and collect the data you are aiming for, including with adequate replicates to ensure a statistically conclusive answer. Importantly, it also follows that experiments should not needlessly use more individuals than necessary.

Injection necessarily involves wounding the animals. Most insects are quite robust, and tolerate injections well. To encourage survival:

• be sure your reagents are properly prepared (e.g. Cas9/sgRNAs)
• clean your work surface
• clean the tools you'll use to handle the insects (paint brushes or forceps)
• keep the injection needles clean

Sterilizing needles and other materials is not typically necessary. Although it should be possible to autoclave injection needles and to filter-sterilize injection solutions if anyone ever wants to try that.

Injection buffer

The solution you inject will depend on your experiment. However, injection solutions should not cause osmotic shock in the animal. (Recall from BI163 that, for example, injecting pure water would cause cells near the injection site to explode, as water rushes into them via osmosis.) We typically prepare reagents for injection in a solution of a physiological saline buffer. Different recipes exist. Below is the recipe for a 50X stock from Hughes and Kaufman (2000) that is suitable for milkweeb bugs and many other insects.

1. Prepare 0.5 M KCl using distilled water (37.275 g/L)
2. Prepare 1 M Na₂HPO₄ (141.96 g/L)
3. Prepare 1 M NaH₂PO₄ (119.98 g/L)
4. Use these solutions to prepare 0.1 M sodium phosphate buffer (pH 7.6 at 25°C)
   a. Combine 8.5 mL of 1 M Na₂HPO₄ with 1.5 mL of 1 M NaH₂PO₄ to obtain 10 mL of 0.1 M sodium phosphate buffer
   b. Check the pH with an indicator strip, and adjust accordingly.
5. Prepare 50X injection buffer (1 mL) by mixing:
   a. 50 μL sodium phosphate buffer (0.1 M)
   b. 500 μL KCl (0.5 M)
   c. 450 μL distilled water

The injection set-up

In general, you will need to inject insects under a dissecting microscope with stereoscopic vision. Ergonomically, it's best to sit, with your feet on the floor, and the scope adjusted to the height of your eyes so you can sit comfortably. Place the needle or micromanipulator on the side of your dominant hand.

Depending on your application, you may inject with needle using a syringe, a hand-pumped pressure source, or a regulator connected to compressed air or an air compressor.

Pulling injection needles

Large insects can be injected using a hand-held syringe and needle - the same type of needles used in medicine. This works for adult Oncopeltus, but survival is higher with a smaller needle.

For most applications, you will need to make your own injection needles from borosilicate glass capillary tubes. Capillaries are stored in the insectary (Arey 301A) above the microscopes.

To create injection needles, use the Sutter Instruments P-97 micropipette puller in the back corner of the Neuro lab (Olin 3rd floor), beside the window.

This machine heats and pulls the capillaries under precise conditions to produce precise needle shapes. Different programs store parameters for different needle shapes. You can also enter your own parameters to change needle shapes.

Pulling needles

• Turn on the Sutter P-97
• Enter a program number (See the section below for existing programs). If necessary modify the parameters. It may be necessary to adjust the heat based on a current ramp value. If you haven't done a ramp test recently (within a week or so), follow the instructions in the section below to find the ramp value.
  
• Lift the front sash
  
• Release each catch on the sliding bars
• Place a capillary into the groove of the sliding right-hand bar. Use the thumb and middle finger of your left hand to draw the two bars together.
  
• Slide the capillary through the wholes in the filament chamber. (Be careful not to touch filament!) Use your left index finger to steady the capillary in the groove on the left bar. Center the capillary, then use your right hand to tighten the knobs on the lefta and right bars.
  
• Press the PULL button.
  
• Discard the first capillary/needles in the glass waste. They are often slightly different shapes.
• Load another capillary and press PULL.
  
• Loose the knobs on the bar and remove the needle.
• Store needles on a large Petri dish with two rows of clay to immobilize the needles.
  
• Repeat this process until you have plenty of needles.

Video demonstration of pulling microinjection needles

Ramp test

• Turn on the Sutter P-97
• Enter any program number
• Hit the CLR button
• Press 0 - Do not clear all parameters!
• Press 1 to select the ramp test
• Lift the front sash
• Release each catch on the sliding bars
• Slide a capillary into the groove. Be careful not to touch filament!
• Press the PULL button. The machine will slowly increase the heat. Eventually it will display a ramp value.
• Record this value
• Discard the capillary in the glass waste.

Ramp values are typically around 720.

Needle parameters

If you are optimizing needle shapes for a new application, it may be helpful to consult Sutter Instrument's "pipette cookbook".

Be aware that the "ramp" value below must be obtained by first running a ramp test.

Program 33: Tribolium larvae and juvenile Oncopeltus and Jadera

heat = ramp+15 pull = 90 vel = 100 time = 250

Program 34: Oncopeltus and Jadera eggs

heat = ramp+15 pull = 90 vel = 100 time = 160

Loading an injection needle

Needles can be loaded from either the front or back. Back-loading is often easier, but may introduce bubbles and requires a larger volume of solution. Front-loading is not possible when using a regulator with compressed gas or an air compressor. When possible, front-loading typically produces better results.

• Remove the small silicon gasket from the needle holder
• Insert the back end of the needle through the cap on the holder and the gasket
  
• Screw the cap back onto the end of the needle holder
  
• Use the micromanipulator to position the needle in the center of the microscope view field.
• Break the tip off gently using clean forceps. Do so at the point where the pulled glass bends naturally.

Back-loading

• Remove the needle from the holder.
• Using an extra-thin pipette tip load 2-5 μL of injection solution into the back end of the tube. Exit the needle
• use 5 mL of air to
• Return it to the micro-injector.

Front-loading

• Pipette up 0.5-5 μL of injection solution in a regular tip.
• Gently remove the tip from the pipetter using an unscrewing motion. Try not to let the solution be expelled from the tip.
• Brace the large end of the pipette tip on the microscope stage using clay - do not cover the end entirely of the solution will be expelled from the tip - so that the pipette tip is near the tip of the needle. Do not touch the tip of the needle!
• While looking through the microscope, use the micromanipulator to move the needle tip so that it touches the solution inside the pipette tip. Capillarity will start to draw the solution into the needle. However, you will need to apply negative pressure to draw the solution in. Do not draw in air.

Once the needle is loaded you should proceed quickly to injections. Do not leave a bright or hot microscope light on the needle, as this may cause the solution to dry and clog the needle tip.

At this point the procedure diverges for different insects and life stages.

Using the air compressor

The air compressor is actually one of the simplest and best ways to provide consistent pressure for injections. While it can be loud, it is safe and easy to use.

• Be sure the air compressor's power cord is plugged into a wall outlet
• Check that the out-going air line is properly connected to the regulator
• Flip the on/off switch up to the "on" position. The machine will make noise for a few minutes while the tank pressurizes. - It's now ready to use!
• Adjust the delivery pressure at the regulator.
  

Do not adjust the red line pressure knob on the air compressor!

• When you're finished injecting, flip the on/off switch down into the "off" position
• Pull the pressure release valve - It will make a loud hiss as the air escapes.

Injecting beetle larvae

Beetle larvae can be injected for RNAi or drug treatments examining development of adult phenotypes.

Before loading the needle, collect enough prepupal larvae for a treatment (25 to 30). Beetle larvae are easy to inject once you learn to select the right stage. They should be prepupae: large larvae that are preparing to molt into pupae and do not move much.

Individuals can be collected from the stock and kept in a petri dish overnight at room temperature.

At this stage, the insect has undergone apolysis, when the external cuticle and underlying epidermis have released their connections.

Don't confuse prepupae with sick or dead larvae! They should look large and healthy. If a larva has any signs of blackness, it is likely diseased and should be euthanized immediately.

Be sure you have a large healthy stock of beetles before you plan to make injections.

• Place a strip of 3M/Scotch Brand double-sided tape on a glass slide. Using your thumb remove most of the "stick" to the tape.
• Using a paint brush or vacuum aspirator pen (a modified pooter) arrange the larvae on the tape. The slide should be vertical. Larvae should lie on their side, facing up about 30˚ and away from the needle.
• It is often helpful to place the slide on a small object, such as a petri dish or the lid of a pipette tip box.
• Arrange the slide and micromanipulator stand, so that the needle comes into the view of the microscope directly from the side (your dominant hand side) and has a 45° vertical angle down, slightly above the first larva.
• Focus the microscope on the larva, but include the needle in your view.
• If you're using a regulator, start with a pressure of 30 psi - Right now that's a guess!
  
• Holding the slide firmly with one hand, use the micromanipulator x-axis knob to extend the needle and puncture the cuticle of the larva. Aim for the posterior edge of one of the dorsal abdominal tergites. Ideally, make your injection site off the midline to avoid puncturing the dorsal blood vessel. Do not go too deep. The cuticle should deform as the needle tip first contacts it, but spring back as the needle enters the body.
• Depress the foot pedal or apply pressure to the syringe. Add only enough solution so that when you pull the need out you can clearly see the colored solution beneath the cuticle.
• While still holding the slide steady, retract the needle by reversing the x-axis knob.
• Occasionally, there may be back-flow of hemolymph into the needle, this can cause it to clog. If necessary, expel some solution to clear the needle. If nothing comes out, the needle will usually unclog after puncturing the next egg. If not, you can snip off the needle tip with clean forceps. - But doing so will enlarge the needle's opening, so do this only sparingly, or you will need to replace the needle.
• Once all the larvae have been injected, count them, record information in your notebook and in the online experimental log.
• On the slide, use a Sharpie to record key information: your name or initials, the date, treatment number, the dsRNA injected and anything else that's helpful.
• Place the slide up-side down over a small petri dish bottom, so that the larvae are suspended by the tape over the small dish. Place the small Petri dish in a large Petri dish and cover it. As the larvae molt into pupae they should fall into the dish.
  • Occasionally larvae are not stuck strongly to the tape. That's fine! Larvae can actually be coaxed off the slide using a small brush, but be careful not to injure them!
  • Pupae do not eat, so no flour is necessary. However, it may be helpful to place a damp paper towel or KimWipe in the outer dish to increase humidity.
  • If any pupae remain on the tape after molting, use a small brush to gently coax them off.
• Pupae should molt into adults after about a week. New adults should be provided with food or they will eat the remaining pupae!
  • Alternatively, pupae can be moved into individual 1.5 ml microfuge tubes where they will eclose into adult beetles in isolation.

Injecting adult true bugs

Adult Oncopeltus or Jadera can be injected for RNAi or drug treatments examining gametogenesis, or females can be injected to examine effects on their embryos. Be sure to have large, health stocks of these bugs before planning these experiments. It can be helpful to have individual cages that are staged.

Most experiments will require young adults. Bugs will not be reproductively capable for 1-3 days after molting to adulthood. Do not use sickly, dirty or moldy bugs. Females and males of these species can also be identified and separated in the fifth instar.

Injections of adult bugs can be done using a hand-held syringe or a pulled capillary needle attached to a hand-held pump for pressure. While it's possible to use a compressed gas source, this is not really necessary.

• To anesthetize a bug, place it on the CO2 pad. If the bug poops, mop it up with a KimWipe to keep the bug clean. Inject one bug at a time.
• The bug should be on its back, facing away from the needle.
• Focus the microscope on the bug, but include the needle in your view.
• With your non-dominant hand, gently brace the bug's head. This should prevent it from sliding away as the needle contacts it.
• If you're using a micromanipulator, arrange the CO2 pad and micromanipulator stand, so that the needle comes into the view of the microscope directly from the side (your dominant hand side) and has a 30° vertical angle down. Use the micromanipulator x-axis knob to extend the needle and puncture the cuticle of the bug.
• Position the needle tip between abdominal sternites, just above the ventral midline in the middle of the abdomen. Be sure to inject in a posterior-to-anterior direction.
• Do not go too deep. The cuticle should deform as the needle tip first contacts it, but spring back as the needle enters the body.
• Apply pressure to the syringe. Add only enough solution so that when you pull the need out you can clearly see the colored solution beneath the cuticle. An adult bug will take 1-3 μL depending on its size.
• While still holding the bug steady, retract the needle.
• Occasionally, there may be back-flow of hemolymph into the needle, this can cause it to clog. If necessary, expel some solution to clear the needle. If nothing comes out, the needle will usually unclog after puncturing the next bug. If not, you can snip off the end of a glass needle with clean forceps. - But doing so will enlarge the needle's opening, so do this only sparingly, or you will need to replace the needle.
• After injecting a bug, move it to a clean Petri dish containing a wet paper towel.
• Once all the bugs have been injected, count them, record information in your notebook and in the online experimental log.
• Move the injected bugs to a new cage. Label the cage with the words "Experiment" and key information: your name or initials, the date, treatment number, the dsRNA, etc. injected and anything else that's helpful.
• If you are collecting eggs from these bugs, check the cage regularly and remove eggs to a Petri dish where they can develop or hatch.

Injecting juvenile true bugs

Oncopeltus or Jadera can be injected at the fourth instar and beyond to examine development of adult phenotypes. Third instars and younger bugs do not survive injection well. Be sure to have large, health stocks of these bugs before planning these experiments. It can be helpful to have individual cages that are staged.

For RNAi experiments investigating adult development, we have traditionally injected 4th instars. However, if you obtain mild phenotypes, it has been shared that injecting the same solution in the 4th and 5th instar can increase the effect of gene knock-down.

Injections of juvenile bugs should be done using a micromanipulator with either a hand-held pump or a regulator attached to air or nitrogen cylinder or an air compressor.

• To anesthetize a bug, place it on the CO2 pad. If the bug poops, mop it up with a KimWipe to keep the bug clean. Inject a small number of bugs on the pad at a time (1-10, although a treatment should include 25-30 bugs).
• Arrange bugs on their back, facing away from the needle.
• Focus the microscope on the bugs, but include the needle in your view.
• With your non-dominant hand, gently brace the bug's head. This should prevent it from sliding away as the needle contacts it.
• Arrange the CO2 pad and micromanipulator stand, so that the needle comes into the view of the microscope directly from the side (your dominant hand side) and has a 30-45° vertical angle down. Use the micromanipulator x-axis knob to extend the needle and puncture the cuticle of the bug.
• Position the needle tip between abdominal sternites, just above the ventral midline in the middle of the abdomen. Injection work best at about a 30˚ angle to the body - slightly anterior-facing, rather than perpendicular.
• Do not go too deep. The cuticle should deform as the needle tip first contacts it, but spring back as the needle enters the body.
• Tab the foot pedal or apply pressure to the syringe. Add only enough solution so that when you pull the need out you can clearly see the colored solution beneath the cuticle.
• While still holding the bug steady, retract the needle. If the bug is stuck to the needle, gently sweep it free with a clean paint brush.
• Occasionally, there may be back-flow of hemolymph into the needle, this can cause it to clog. If necessary, expel some solution to clear the needle. If nothing comes out, the needle will usually unclog after puncturing the next bug. If not, you can snip off the end of a glass needle with clean forceps. - But doing so will enlarge the needle's opening, so do this only sparingly, or you will need to replace the needle.
• After injecting the bugs on the CO2 pad, gently move them to a clean Petri dish containing a wet paper towel.
• Once all the bugs have been injected, count them, record information in your notebook and in the online experimental log.
• Move the injected bugs to a new cage. Label the cage with the words "Experiment" and key information: your name or initials, the date, treatment number, the dsRNA, etc. injected and anything else that's helpful.
• If you are phenotyping adults, check the cage regularly and remove new adults to image them, collect RNA, etc.

Injecting eggs of Oncopeltus or Jadera

Oncopeltus or Jadera eggs can be injected to study embryonic development, using RNAi, CRISPR or other treatments, or for genetic manipulation using CRISPR. Be sure to have a large number of healthy breeding adults before planning these experiments. It can be helpful to have individual cages that are staged.

Injections of insect eggs must be done using a micromanipulator and regulator attached to an air or nitrogen cylinder or an air compressor. With practice, survival rates are about 30%.

Special thanks to Derek Hernandez '21 and Asha Sidhu '21 for optimizing this protocol and for sharing photographs!

Preparing the arena

A small agarose mold is used to hold the egg for injection. We have 3D-printed casting trays sized for Oncopeltus and Jadera eggs. The original design for Oncopeltus comes from Reding & Pick (2020), and was modified for Jadera by Derek Hernandez and Tim Stonesifer.

• Add 25 mg of agarose to 30 mL of 1X TAE buffer in a 150 ml flask. (Do not use the molecular-grade agarose or the TBE buffer used for agarose gel electrophoresis!)
• Microwave the mixture for 55 seconds.
• Place the casting tray in a Tupperware-style food container.
• Once the agarose is fully dissolved, fill the casting tray with the agarose. Then cover the container.
• Place the container at -20˚C for 5-10 minutes. This allows the gel to set quickly.
  • Alternatively, arenas can be stored in a sealed container at 4˚C for several weeks.
• Open the container and let the tray come to room temperature for a minute or two.
• Use a scalpel blade to remove the gel arena. Cut around the arena and then carefully lift it out.
• Place the arena in the center of a large petri dish cover
• Cut it in half. Each half can be used in a separate round of injection.

Setting up staged embryo collections

Most egg injections are targeting embryos of a particular stage - usually before cell membranes have formed. Check the embryology of your species and the goals of your experiment to be sure what window you need. True bugs lay eggs at any time of day, but many insects favor the early morning for egg-laying. Oncopeltus embryos should be injected at 2–8 hours after egg laying (Reding & Pick 2020).

• Place egg-laying adults in a clean cage or Petri dish with food, water and (for Oncopeltus) a loose cotton ball or (for Jadera) torn egg cartons.
• Record the time the adults are placed in the cage.
  • For Oncopeltus, place adults in a clean cage without cotton, the day before you plan to collect eggs. Later, add the cotton to start the clock. Females will be eager to lay eggs. (Fertilization and embryonic development occur just before egg laying, regardless of the behavior of males.)
• Come back at the maximum time window. Any eggs in the cage have been laid within that time window.

For example, if adults are provided with a new cage at 8:00 am, and eggs are removed at 2:00 pm, then the embryos will be 2-8 hr AEL at 4:00 pm.

The clock is ticking, so get injecting!

Loading the arena

• Collect Oncopeltus eggs from cotton balls by gently teasing the cotton apart by hand. Let the eggs drop onto a clean piece of white paper.
• Collect Jadera eggs from the bottom of cages. Adults often drop then in the seed dishes or directly below the egg cartons.
• Using a fine paint brush, place each egg on top of the arena. About 20-30 eggs can fit in each lane. It is important to get the eggs into the lane rather than just being laid on top of the lanes since this will reduce their rotation during injection.

Egg injections

For CRISPR, be sure to prepare a fresh Cas9/sgRNA solution.

• Arrange the arena and micromanipulator stand, so that the needle comes into the view of the microscope directly from the side (your dominant hand side) and has a 45° vertical angle down.
• Focus the microscope on the eggs, but include the needle in your view.
• Arrange the lanes perpendicular to the needle, so that the needle will puncture the eggs from the side, and the egg will push against the walls of the lane rather than against another egg.
• Start with a pressure of 20 psi and keep the needle's tip diameter minimal. Change the needle or reduce pressure down to 10 psi at anytime if solution leaks continuously from the tip.
  
• Ensure that there is no air at the tip of the needle.
• Align the needle to minimize side-to-side movement of the needle. It should only move in and out (rotating the x-axis knob only). For further alignment, use your left hand to move the petri dish with the arena around. You want to aim for the center of the egg on a side angle, this reduces the rotation of the egg and allows pressure from the needle to be focused at the point, of injection.
• Rotate the knob to puncture the egg. This may take several tries. Sometimes, moving the petri dish with the left-hand helps.
• Tap the pedal to release the injection solution. Adjust the regulator pressure to allow sufficient solution to be delivered. You must continue to look down the microscope to determine this. You do not want a fully green egg; the egg is dead if this occurs. You want a small green spot within the egg. Once the spot is identified, immediately rotate the knob to pull the needle out of the egg.
  • Sometimes, capillary action alone will release the injection solution into the egg. If so, then it's done with injection. Simply remove the needle from the egg before any yolk is taken up.
• Occasionally, there may be back-flow of yolk into the needle, this can cause it to clog. If necessary, expel some solution to clear the needle. If nothing comes out, the needle will usually unclog after puncturing the next bug. If not, you can snip off the end of a glass needle with clean forceps. - But doing so will enlarge the needle's opening, so do this only sparingly, or you will need to replace the needle.
• Repeat this process for all of the eggs.
  
• Once all the eggs have been injected, count them, record information in your notebook and in the online experimental log. You can remove any eggs that are deflated; these will not survive and can be eliminated from determinations of embryo survival from this point on.
• Carefully lift the arena into a petri dish with a wet paper towel. This prevents the arena ands eggs from drying out.
• Label the dish with key information: your name or initials, the date, treatment number, the target gene, etc. and anything else that's helpful.
• Oncopeltus eggs take about 10 days to hatch at room temperature. On day 7, the petri dish should be covered with KimWipes to prevent hatchlings from escaping. A small dish of seeds should also be added.
• Hatchlings should be removed to a new dish or cage, following regular culture protocols. After 8 days, checked constantly because hungry hatchlings will eat their siblings.
• Successfully injected hatchlings may continue to carry the dye from the injection solution for some time.

These third and fourth instar Oncopeltus were injected as newly laid eggs. (photo: Derek Hernandez '21)

Injecting eggs of Vanessa

• Take an injection needle (a pulled capillary tube made of borosilicate glass).
• Mount the needle on the micro-injector.
• Use the micromanipulator to position the needle in the center of the microscope view field.
• Break the tip off gently using clean forceps. Do so at the point where the pulled glass bends naturally.
• Remove the needle and back-load it with 1 μL of injection mix. Then return it to the micro-injector.
• Press the foot pedal to release a small amount of fluid.
• Move the lid with one hand so the needle is at a 45° vertical angle, slightly above the egg top. Focus the microscope so the egg top is sharp.
• Start with a pressure of 20 psi and keep the injection diameter minimal. Change the needle or reduce pressure down to 10 psi at anytime during the injection if expelled droplets become more than twice the size of the micropyle.
• Holding the egg-covered lid firmly with one hand, use the coarse knob to puncture the upper side of the egg. Do not go very deep beyond the chorion. Release a minimal volume of sgRNA/Cas9 solution. Backflow of yolk will occasionally occur, but is often expelled by repeated bursts.
• While still holding the egg dish stedily, retract the needle and inject outside of the egg to expel any yolk. If nothing comes out, the needle will usually unclog after puncturing the next egg. Try to achieve consistency and decent speed.
• Upon piercing the egg, only add enough solution so that when you pull the need out you can clearly see the red solution inside the egg.
• Once all the eggs have been injected, count them, record information on the lid, such as the name of the person making the injections, the date, the age range of the embryos, and the target sgRNA.
• Snap the lid to a matching cup
• Place the cups in a closed plastic conatiner with a wet paper towel to avoid desiccation during the next 24 hrs. Include a control batch of uninjected eggs in a similarly prepared dish.
• Open the tupperware after 24 hrs to prevent condensation.

Care of Vanessa after injections

• Embryos will develop at a temperature-dependent rate, hatching after about 3 days at 28°C to about 4 days at 22°C. In the morning of the hatching day, press small crumbs of fresh artifical larval diet onto the side and lid of the cup. Hatchlings should be found feeding on the diet within 24 hrs.
• Record the number of hatchlings. Typically 30-40% of injected eggs survive for WntA sgRNA/Cas9 injections made by an experimented operator. Low rates of survival may indicate lethal phenotypes, in which case you can inject older eggs, or with a more dilute sgRNA/Cas9 solution in order to generate mosaic embryos with fewer editted cells.
• Leave the larvae at room temperature (22-25°C). Check for excess moisture daily. Temperatures of 24-25°C (75°F) provide an optimal trade-off between developmental time and survival, with a developmental time of 30-36 days. Pinholes in the lid may be necessary to let excess moisture out of the cup.
• Examine the larvae every two days.
• Add small amounts of fresh diet as needed.
• Remove individuals to seperate cups once they reach the third instar.
• Dispose of dead larvae as biohazardous waste.
• Hang pupae in a dedicated cage. Spray them with water every 1-2 days. Record the number of pupae. Note any abnormalities.
• After emergence, allow 1 day for wings to fully to dry, then freeze all G₀ adults at -20°C for >24hrs.
• Pinning is not necessary for this class and phenotypic analysis can be done on wings that have been detached from the insect body and stored in glassine envelopes.

---

BI379 at MBL Lab