Techniques for Investigating the Anatomy of the Ant Visual System

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.

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The overall goal of this suite of microscopy techniques is to obtain detailed anatomical information of the ant visual system. This method can help address key questions in the field of visual ecology, to estimate various visual parameters, such as optical sensitivity, visual acuity, and size of the visual field. The main advantage of this technique is that it gives the comprehensive overview of a given visual system.

It can also be adapted to use with many different species. To make the cornea replicas, use ants preserved in ethanol or mounted on a pin, or in Plasticine. If the head is relatively large, the remaining body parts can be removed.

To begin, quickly spread fast-drying, colorless nail polish over the eye, using a toothpick, without scratching the eye. Let the polish set at room temperature. Then, use a fine insect pin to gently lift the replica from the head capsule surrounding the eye.

And using forceps grasp the part of the replica that covers the head, but not the eye. Next, place the replica on a glass slide. There, use a razor blade to carefully trim excess material around the eye, while securing the replica.

Before placing a cover slip over the replica, it may be necessary to make three to four fine partial radial slices to help flatten it. When placing the cover slip, document the replica's orientation on the slide. Then, seal the cover slip, using small dabs of nail polish on the corners, making certain that the liquid does not touch the replica.

The slide is then ready to image using a compound microscope. To begin, anesthetize the ant and place it in a petri dish. If dye needs to be adapted for bright or dot conditions this should be done for a few hours before the dissection.

If the dot conditions seem to be maintained, the dissection should be carried out in the red light. First, remove the antennae using forceps. Then, remove the mouth parts using a sharp razor blade.

In large specimens, cut through the anterior part of the eye. In a small specimen, cut as close to the eyes as possible, without tugging on the brain, as this may tear out the retina. Next, orient the head and make a transverse incision, that removes the ventral portion of the head, which may include part of the ventral eye.

Then, detach the head capsule with a coronal incision, just posterior to the compound eye. In this example, the transverse and coronal incisions are made simultaneously. Finally, transfer the dissected head capsule with the compound eyes to ice cold fixative.

It's important to perform these steps quickly, to prevent tissue from degrading, so the tissue should go into the fixative within two minutes of the first incision. Following the text protocol, prepare the specimen and drain the excess resin. Then, place the specimen on the block in a vertical position, and if needed, use a little excess resin to secure it there.

Label the block with paper labels embedded in the resin, or attached to the block. Then, cure the resin in a 60 degree Celsius oven, for 12 to 14 hours. Once cured, load the specimen into a clean envelope.

Next, mount the block onto a microtome chuck. Before sectioning, first make certain that the specimen's orientation is appropriate for the sectioning plane. Under a dissecting microscope, trim the resin block using a razor blade.

Then, remount the chuck onto the microtome arm, and angle the specimen. Next, fill the knife boat with distilled water using a syringe with a 0.45 micron filter. Load the water up to the edge of the knife.

In some places the meniscus may be convex. Then, drain the boat until the meniscus is very slightly concave, but still reaches the edge of the knife. Now, slowly and carefully bring the knife towards the specimen, and align the block to the knife.

Make frequent checks via the ocular. When the knife is close but not yet cutting the specimen start cranking the microtome wheel to complete the approach. Then, when approaching the region of interest in the block adjust the section collecting thickness.

In this case, a diamond knife is used to make one micron sections. Then, from the filter equipped syringe place a series of small droplets of distilled water on a slide. To collect a section of interest, pick it up using an eyelash tool.

And carefully, float the collected section onto a water droplet. Continue loading as many sections onto each slide as can fit. When a slide gets fully loaded transfer it to a hot plate at 60 degrees Celsius to evaporate the water, and adhere the sections to the slide.

This takes about 15 to 20 minutes. Next, use another filter equipped syringe to deposit a drop of toluidine blue dye onto the slide. Place the drop on one end of the slide and spread it using the edge of a needle without touching or scraping the sections.

Then, place the slide on the hot plate for 20 to 60 seconds. Afterwards, spray the slide with distilled water from a wash bottle, and use the hot plate to dry it off. Later, check the slides under the compound microscope and then image them.

The described techniques enable a detailed study of the simple and compound eyes of ants. Imaging the dorsal view of the head using Z-Stack photomicrography provides an overview of the layout of the visual system useful for making gross measurements. SEM imaging provides the detail needed to resolve small eyes and the detail needed to identify the shape of each lens.

Semi-thin sections imaged using light microscopy show the internal anatomy of an eye. Lens, crystalline cone, and rhabdomere dimensions can be measured at this resolution and magnification. Pigment cells, cone tract presence and rhabdomere shape distributions can also be studied.

Ultra-thin sections imaged by TEM provide ultrastructure information, such as microvillar orientation, and for example, the width of the constricted crystalline cone tract. While attempting this technique, it's really important to be patient, to use clean instruments, and to use freshly made solutions. Following these experiments, additional behavioral or physiological experiments can then be carried out.

These experiments will inform us about the functional visual abilities of the animal, relative to the eye anatomy. Using these techniques allows us to explore the diversity of visual systems, and how they have evolved to suit different lifestyles and different ecologies, in different species of ants. Though this method can provide insight into the ant visual system, it can also be used to study the visual system of other insects, as well as, other nervous tissues.

Don't forget that working with fixatives and resins is extremely hazardous and precautions such as, working in the fume hood and wearing nitrile gloves, should always be taken.

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Journal (Environment)

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Techniques for Investigating the Anatomy of the Ant Visual System

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