Techniques for Investigating the Anatomy of the Ant Visual System

Summary

This article outlines a suite of techniques in light and electron microscopy to study the internal and external eye anatomy of insects. These include several traditional techniques optimized for work on ant eyes, detailed troubleshooting, and suggestions for optimization for different specimens and regions of interest.

Abstract

This article outlines a suite of techniques in light microscopy (LM) and electron microscopy (EM) which can be used to study the internal and external eye anatomy of insects. These include traditional histological techniques optimized for work on ant eyes and adapted to work in concert with other techniques such as transmission electron microscopy (TEM) and scanning electron microscopy (SEM). These techniques, although vastly useful, can be difficult for the novice microscopist, so great emphasis has been placed in this article on troubleshooting and optimization for different specimens. We provide information on imaging techniques for the entire specimen (photo-microscopy and SEM) and discuss their advantages and disadvantages. We highlight the technique used in determining lens diameters for the entire eye and discuss new techniques for improvement. Lastly, we discuss techniques involved in preparing samples for LM and TEM, sectioning, staining, and imaging these samples. We discuss the hurdles that one might come across when preparing samples and how best to navigate around them.

Introduction

Vision is an important sensory modality for most animals. Vision is especially crucial in the context of navigation for pinpointing goals, establishing and adhering to routes, and obtaining compass information^1,2. Insects detect visual information using a pair of compound eyes and, in some cases, one to three dorsally-placed simple eyes called ocelli^3,4,5.

The eyes of ants are of particular interest because, while ants are wonderfully diverse, they conserve some key characteristics across species. Despite dramatic variation in anatomy, size, and ecology, the vast majority of species are eusocial and live in colonies; as a result, different species face similar visual challenges in terms of navigating back and forth between a central place and resources. Across ants the same basic eye bauplan can be observed in animals ranging from 0.5-26 mm in body length, from exclusively diurnal to strictly nocturnal species, and from slow walking subterranean to leaping visual predators^6,7,8,9,10. All of these staggering differences in ecology and behavior give rise to innumerable permutations of the same basic eye structures to suit different environments, lifestyles, and body-sizes^11,12. As a consequence, studying the visual ecology of ants provides a veritable treasure trove of possibilities to the determined investigator.

Understanding the visual system of insects is essential in gaining an insight into their behavioral capabilities. This is apparent from integrative studies which nicely combine anatomy with ecology and behavior to a great success in a few insect groups (e.g., references^{13,14,15,16,17}). Though the field of ant navigation and ant behavior in general has been quite successful, very little emphasis has been placed on ant vision outside of a few selected species. Here, we will elaborate on the techniques involved in investigating eye design of ants. While we will focus on ants, these techniques can be applied, with slight modifications, to other insects, too.

Protocol

1. Specimen Preparation

NOTE: It is necessary to first understand the relative location of the compound eye and ocelli to each other and on the head. This can be achieved by acquiring images of the dorsal view of the head. For this, we recommend processing samples either for photomicrography or using SEM techniques. Below are steps involved in both processes.

1. Specimen collection
   1. Collect and store specimens directly into 70% ethanol. Collect different castes whenever possible.
   2. Label specimens with time, date, and place as well as any other relevant observations (e.g., collected while foraging, mating aggregation, nest inside a twig, etc.)
   3. Collect enough specimens to have multiple replicates in each treatment.
2. Photomicrography and Z-stacking
   1. Air dry specimens and mount them on triangular point cards, using water soluble glue, and then on an insect pin. For details, see reference¹⁸.
   2. Image using a high magnification stereomicroscope with a Z-stepper motor and a color camera.
   3. Use a diffuser to have uniform lighting for specimens.
   4. Capture images at different focal planes and save images in a lossless file format (such as tiff) and focus stack them using commercially available software.
3. Scanning electron micrographs
   NOTE: Thoroughly clean all tools and working surfaces with ethanol to avoid contamination of the specimen with dust and other particulates.
   1. Dehydrate specimens in ethanol overnight and air dry in a Petri dish.
   2. Use a sharp razor blade to separate the head from the remainder of the body.
   3. Mount the head at the required viewing angle (e.g., dorsal facing up) on aluminum stubs using conductive carbon tape or tabs. Cut the carbon tape into thin strips and fold it to support the head capsule.
   4. Use a sputter coater to apply gold to the surface of the specimen for 2 min at 20 mA with a rotary stage. The time and current may need to be adjusted depending on the instrument.
   5. Transfer specimens to a new aluminum stub with fresh carbon tape or tabs.
      NOTE: The uncoated carbon will provide a black background but transferring specimens can damage the gold coating.
   6. Check that the specimen orientation is still correct using a dissecting microscope and adjust as necessary with a pair of fine forceps or similar tool. Take care not to scratch the gold coating; handle as little as possible.
   7. Load specimens onto the SEM stub holder, making note of the position of stubs and specimens relative to each other.
      NOTE: Some SEMs are equipped with a stage camera but many are not and it can be difficult to locate small specimens at high magnification.
   8. Image the specimens. Use a low accelerating voltage to avoid charging and a small aperture for good depth of field.
      NOTE: Settings are best optimized in consultation with a technician specialized in the particular instrument being used.

2. Quantifying Facet Numbers and Diameters

1. Cornea replicas
   1. Use ants preserved in ethanol or mounted on a pin for this purpose (step 1.1.1).
   2. Mount the animal on an insect pin or on plasticine. If the head is relatively large, the remaining body parts can be removed.
   3. Use an insect pin or a fine toothpick to pick up a small drop of fast-drying colorless nail polish and quickly spread it over the eye. Ensure the pin does not scratch the eye. The nail polish should cover the entire eye and some of the surrounding head capsule.
      NOTE: It is important that the nail polish be of relatively uniform thickness across the eye.
   4. Leave the nail polish to set at room temperature.
   5. Once it is fully set, use a fine insect pin to gently lift the replica from the head capsule surrounding the eye.
   6. Use a fine pair of clean forceps to lift the replica, grasp the part of the replica that covers the head and not the eye.
   7. Ensure awareness of the orientation of the eye: the anterior, posterior, dorsal, and ventral region.
   8. Place the replica on a glass slide. Use a razor blade to trim the replica by carefully removing excess material around the eye. Use a needle or a pair of forceps to prevent the replica from moving.
   9. If the eye is very convex, use a razor blade to make 3-4 fine partial radial incisions around the edge to help 'flatten' the replica. If the eye is relatively 'flat', there is no need to make such incisions.
   10. Place a cover slip gently on the eye replica, ensuring that the orientation of the replica is known. Do not apply pressure as this can eliminate the corneal impression on the nail polish.
   11. Seal the coverslip using very little nail polish on four corners. If the nail polish flows between the cover slip and glass slides, it will damage the replica.
   12. Image the slide on a compound microscope.
       NOTE: If only some facets are in focus, then this suggests the eye replica is not flattened enough. Discard and start again from step 2.1.3.
   13. Import the image into freely available programs such as ImageJ/Fiji where the number of ommatidia and size of each ommatidia can be measured.
       NOTE: This method can be used to prepare replicas of ocelli, too. Since it is a single lens, we recommend keeping all the ocelli together in one replica.

3. Analyzing the Structure of the Eye

NOTE: To study the anatomy of the eye requires in most cases two complementary techniques of LM and TEM. The initial processing stages require similar techniques for both LM and TEM. The difference arises from the sectioning stage onwards. Processing samples requires the use of hazardous chemicals which must be handled with care and discarded responsibly. Use personal protective equipment, work in a fume hood, always read the safety data sheets(SDS), and carry out risk assessments before starting.

1. Dissection
   1. Anaesthetize specimens by cooling or by exposure to gaseous CO₂.
      1. CO₂ anesthesia is very fast (generally under 1 min) and care should be taken to avoid overexposure as this may result in death of the specimen. If using dry ice pellets (solid CO₂) avoid direct contact with specimens as this can cause cold burns.
      2. Cold anesthesia is slower; 4 °C is sufficient, and colder temperatures are not recommended. Establish an appropriate cooling time for the species. Large or cold resistant ants such as bull ants may require >10 min to become fully immobilized, while smaller species may only need 1-2 min. Excessive cooling will kill the specimen (avoid direct contact with ice). Specimens should preferably be held in small, foam-stoppered, plastic containers and placed in an icebox where they can be observed rather than in an electric refrigerator or freezer.
   2. Place specimen on a Petri dish, and adjust for viewing under a dissecting microscope. Working quickly is important to preserve anesthesia and to avoid degeneration of the tissue once incisions are made (this can happen within seconds).
   3. Remove antennae with forceps. If working with a stinging insect, it is advisable to amputate the gaster first to avoid being stung.
   4. Remove the mouth parts using a sharp razor blade; forceps may be used to hold the specimen down. Cut through the anterior part of the eye (large specimens) or as close to the eyes as possible (small specimens) without tugging on the brain as this may tear out the retina.
   5. Prepare to open the head capsule. Angle the specimen so that the first incision is pointing up; this may be done either under the dissecting microscope while holding the specimen in forceps or under visual control while holding the specimen between the thumb and fore finger.
   6. Make a transverse incision through the head to remove the ventral portion of the head; part of the ventral eye may be removed in large species to improve fixation and infiltration. The head should still be attached to the body at this point.
   7. Sever the head capsule from the body by making a coronal incision just posterior to the compound eye.
   8. Place the dissected head capsule with the compound eyes in ice cold fixative: 2.5% glutaraldehyde and 2% paraformaldehyde in phosphate buffer (pH 7.2-7.5).
      Caution: Fixative is corrosive and toxic; wear appropriate protection and work in a fume hood.
      NOTE: It is important to work quickly to arrest neural tissue degeneration. The dissection should be completed in 2 min or less (efficient dissection may require some practice).
      NOTE: If the eye needs to be adapted to bright or dark conditions, then first expose the animals to the required light condition for a few hours. Carry out the dissections in the respective light conditions. Dissections can be carried out under red lights to simulate darkness.
2. Specimen processing
   1. Keep the specimens in the fixative at room temperature with motion on an orbital shaker, for 2 h. Large specimens may require longer fixation times.
   2. Remove the fixative and dispose it appropriately. Wash specimens in room temperature phosphate buffer (3 times, 5 min each) on the shaker.
      NOTE: The phosphate buffer comprises of 8 g NaCl, 0.2 g KCl, 1.44 g Na₂HPO₄, and 0.244 g KH₂PO₄ in 1 L of distilled H₂O (pH 7.2).
   3. Remove the phosphate buffer and add 2% OsO_4. Place the specimen jar on the shaker in the fume hood for 1-2 h. This is a post-fixation step to fix fats and also provide contrast for TEM.
      NOTE: Osmium fixation times are subject to specimen size; as a rough rule of thumb, calculate 1 h of fixation per 1 mm³.
   4. Remove the osmium solution and dispose of it appropriately. Wash specimens in room temperature buffer (3 times, 5 min each) on the shaker.
   5. Dehydrate the specimens by placing them in increasing concentrations of ethanol or acetone; for example, 50, 70, 80, and 95% for 10 min each and finally 100% (2 times, 15 min each). Place the specimens on the shaker between solution changes.
      NOTE: If necessary, specimens may be stored in 70% ethanol overnight.
   6. Drain the ethanol and add 100% acetone. Leave it for 20 min on the shaker (skip this step if dehydrating it in acetone). Replace with fresh acetone and leave it for an additional 20 min.
   7. Infiltrate the tissues with resin using the following ratios of acetone to resin: 2:1 (3 h), 1:1 (overnight), 1:2 (4 h), and pure resin (overnight). At each step leave specimens on the shaker inside the fume hood, and cap the container for all but the last two steps.
      NOTE: Resin is too viscous to be drained so specimens must be moved to a new disposable container at each step.
   8. Prepare blocks to mount the samples. Blocks can be custom made by cutting acrylic glass into small rectangular blocks (1.5 x 0.5 x 0.3 cm). Blocks can also be made by pouring epoxy resin (there are many commercial kits available) into a silicon mold and then cure it in the oven for 12-14 h at 60 °C.
      Caution: Uncured (liquid) resin is carcinogenic and should be returned to the oven until fully hardened.
   9. Place the block vertically in the mold. Carefully take the specimens from the liquid resin, allow excess resin to drain, and place the specimen on the top of the block. A small amount of additional resin can be used to bind the specimen to the block.
   10. Label the block. Print the paper labels and embed it in the block or attach it to a block face.
   11. Keep the mold with the embedded blocks in the oven for 12-14 h at 60 °C.
   12. Store the specimen block in a clean envelope. This can be stored in this manner over several months to years.
   13. Leave the used containers, dirty gloves, and other contaminated equipment in the fume hood to allow acetone to evaporate completely (minimum 12 h).
   14. Cure resin in the oven before discarding disposable equipment or scraping resin off other items such as forceps.
3. Sectioning
   1. Observe the block under the dissecting microscope to ensure that the specimen orientation is appropriate for the sectioning plane.
   2. If the orientation is not appropriate, use a jeweler's saw to cut out the specimen and re-orient using fragments of set resin and fresh resin to re-seat the specimen. The head may also be split into two halves to section the two eyes separately. Cure resin again before proceeding.
   3. Mount the block on the removable microtome chuck. Remove the chuck and place on holder.
   4. Trim the resin block using a razor blade under the dissecting microscope.
      Caution: Do not do this when the chuck is mounted on the microtome arm as it can jolt the arm and damage the bearings.
   5. Remount the chuck onto the microtome arm and angle the specimen.
   6. Mount the knife on the holder at the appropriate angle (0° for glass knives, see manufacturer's instructions for diamond knives).
      NOTE: Glass knives can be made cheaply with a glass knife maker but must be replaced periodically as they lose their edge; high quality diamond knives can be purchased but are expensive, require special care, and are not suitable for beginners.
   7. Fill the knife boat with distilled water using a syringe equipped with a filter (0.45 µm pore size).
   8. Fill the boat until the water level reaches the edge of the knife; the meniscus may be convex in other areas of the boat.
   9. Drain the boat until the meniscus is very slightly concave but still reaches the edge of the knife. The level of the water can be adjusted at any point but always away from the edge of the knife.
   10. Carefully bring the knife towards the specimen and align the block to the knife. This is best done slowly, periodically checking the proximity of the knife through the eyepiece and from the side.
       NOTE: Check the instrument handbook for specific instructions as instruments vary.
   11. Set the section thickness on the microtome. Selecting the correct thickness will be dependent on specimen size, region of interest and the kind of knife being used.
   12. If using a glass knife select a higher setting (e.g., 4 µm), if a lot of material must be cut away before reaching the area of interest. If a diamond knife is being used or if the specimen is very small, 1-2 µm may be more appropriate.
   13. Start the "cutting" (cranking the microtome wheel) when the knife is close but not yet cutting into the specimen to perform the last part of the approach. Sections should start appearing within a few rotations; if not, stop and very carefully bring the knife a little closer.
   14. Adjust the section collecting thickness (1 µm for semi-thin sections) when approaching the region of interest.
   15. Collect any sections using an eyelash tool.
       NOTE: Eyelash tools can be made with an eyelash mounted onto a thin stick with nail polish.
   16. If a lot of material must be removed, allow the sections to accumulate and remove en masse by removing the knife and flushing it out with water. If using a glass knife, this may be an appropriate time to change to a fresh section of the knife or to a new knife.
   17. Place a series of small droplets of distilled water on a slide using a Pasteur pipette or ideally, the filter equipped syringe.
   18. Carefully float the collected section on the eyelash onto the water droplet.
   19. Collect sections like this until it is appropriate to check the depth of sectioning.
       NOTE: Although it can be tedious, it is always best to check often.
   20. Place the slide on a hotplate set to 60 °C. Allow all water to evaporate and the section to adhere to the slide.
   21. Dye sections with toluidine blue for 10-60 s (staining time will vary with section thickness). Dispense the dye with a syringe equipped with a filter (as above).
   22. Place a drop of the dye on one end of the slide and spread it using the side of the needle without touching or scraping the sections. Place the slide on the hotplate for approximately 10-20s.
   23. Rinse the slide by spraying it with distilled water in a wash bottle and place it on the hot plate to dry it.
   24. Check under the compound microscope and image them.
   25. Repeat until region of interest is reached.
   26. For ultra-thin sections: set the cutting thickness between 40-60 nm to collect TEM sections.
   27. Cut about 3-5 sections and check thickness using an interference color chart. Sections should reflect light grey when viewed at an angle.
       NOTE: Chloroform fumes can be pipetted over sections to relax any creases. This is most relevant to experienced users, and beginners need not worry. Too much chloroform released too close to the section can damage sections.
   28. Pick up a Formvar-coated, copper, slot grid held with forceps with the Formvar side up. Be careful not to puncture the Formvar coating.
   29. Dip the grid into the boat edgeways and away from the sections, then bring the grid up parallel to the surface under the sections. If necessary use the eyelash to guide the sections over the grid.
   30. Carefully dab around the tip of the forceps with filter paper to soak up water trapped between the arms. If this is not done, water tension can pull the grid up between the arms or make it stick to one side. This can result in contamination or mechanical damage.
   31. Carefully remove excess water from the grid itself by standing the grid edgeways on filter paper.
   32. Place the grid in a grid holder.
   33. Repeat until enough sections have been collected.
   34. Semi-thin sections can be imaged directly with any compound microscope equipped with a camera. Immersion oil can be placed directly onto the sections. Slides should be stored in a slide box to prevent discoloration.
4. Staining ultra-thin section for TEM contrast
   NOTE: The following steps should be carried out under cover as the dyes are light and CO₂ sensitive. Furthermore, EM dyes use heavy metals to produce contrast and are therefore hazardous substances. Appropriate care must be taken when handling these stains.
   1. Cover a few large Petri dishes with aluminum foil to block light. Work under these for the following steps. Partially uncover them to allow working space but place the covers back on as soon as possible.
   2. Cut a piece of wax film and carefully place five droplets of 6% saturated uranyl acetate on it using a Pasteur pipette.
      1. To prepare the solution, mix 2 g uranyl acetate with 100 mL 50% methanol in distilled water, and filter the solution before using. The solution cannot be stored and should be made fresh each time¹⁹.
   3. Carefully pick up the TEM grids with forceps and balance on top of dye droplets with the section side down. Leave for 25 min.
   4. Rinse grids individually by rapidly dipping them in and out of distilled water; progress through four different vials of distilled water.
   5. Place five droplets of lead citrate on a fresh piece of film. Arrange a few NaOH pellets around the dye droplets (this absorbs atmospheric CO₂ to prevent precipitation of lead carbonate).
      1. To make the lead citrate solution, prepare bi-distilled water by boiling distilled water for 0.5 h. Allow the water to cool and in a sealable container, add 0.3 g lead citrate to 100 mL of the bi-distilled water. Add 1 mL 10 M NaOH, seal the container tightly, and shake until dissolved¹⁹.
   6. Place the grids on the dye drops as described in 4.3 and cover. Stain for 5 min.
   7. Rinse in distilled water as before by dipping the grids in and out of distilled water 20 times. Progress through three vessels of distilled water.
   8. Soak up excess water with filter paper and allow the grids to dry in a grid box.
   9. Image in a TEM at low accelerating voltage.

Results

The methods described here enable detailed study of the simple and compound eyes of ants. Imaging the dorsal view of the head using Z-stack photomicrography techniques allows one to obtain an overview of the layout of the visual system (Figure 1). This is good preparation for dissections and to determine the required sectioning angle. This technique is also useful for taking measurements such as head width, eye length, and ocellar lens diameters. SEM imaging ...

Discussion

The suite of methods outlined above allow for an effective investigation into the optical system of ants and other insects. These techniques inform our understanding of sampling resolution, optical sensitivity, and potential polarization sensitivity of the eye being studied. This knowledge provides an important foundation for physiological and behavioral investigation into their visual capabilities. Furthermore, while the methods detailed here have focused on ant visual systems, these techniques can be used on other inse...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Ant	Myrmecia midas
Stereomicroscope	Leica M205 FA
Sputter coater	Pro Sci Tech
Ethanol	Sigma Aldrich
Petri dish	ProSciTech
Dissecting microscope	Leica MZ6
Insect Pin	ProSciTech
Colourless nail polish	Non branded: from any cosmetic store
Glass slide	ProSciTech
Razor blade	ProSciTech
Foreceps	ProSciTech
Cover slip	ProSciTech
Compound microscope	Leica DM5000 B
Glutaraldehyde	Sigma Aldrich
Paraformalydehyde	Sigma Aldrich
Potassium Chloride (KCl)	Sigma Aldrich
di-Sodium Hydrogen phosphate (Na2HPO4)	Sigma Aldrich
Potassium di-Hydrogen Phosphate (KH2PO4)	Sigma Aldrich
Sodium Chloride (NaCl)	Sigma Aldrich
Osmium tetroxide	Sigma Aldrich
Acetone	Sigma Aldrich
Araldite Epoxy Resin	Sigma Aldrich
Pasteur pipette	Sigma Aldrich
Toluidie Blue	Sigma Aldrich
Hotplate	Riechert HK120