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.
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.
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 information1,2. Insects detect visual information using a pair of compound eyes and, in some cases, one to three dorsally-placed simple eyes called ocelli3,4,5.
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 predators6,7,8,9,10. 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-sizes11,12. 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., references13,14,15,16,17). 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.
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.
2. Quantifying Facet Numbers and Diameters
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.
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 also gives detailed overview images but additionally allows acquisition of high magnification and high resolution images. Particular regions of interest in the eye can be examined in detail and variations in lens shape can be identified (Figure 2). SEM images are especially useful for resolving ants with small eyes and ocelli. The cornea replicas provide information on the shape, size, and number of lenses in each eye (Figure 3). Semi-thin sections imaged using LM techniques allow investigation of the gross internal anatomy of the eye (Figure 4 and Figure 5); this includes the thickness of the lens, diameter of the crystalline cone, presence of a crystalline cone tract, shape, width, and length of the rhabdom, mapping the dorsal rim area, and location of the primary and secondary pigment cells. This technique can be nicely complemented by ultra-thin sections imaged using TEM, which allows for determining the ultrastructure especially, the microvillar orientation (Figure 4) and the quantifying smaller structures (e.g., width of the constricted crystalline cone tract, Figure 5).
Figure 1: Z-stack photomicrographs of the three castes of the Australian sugar ant, Camponotus consobrinus. This provides an overview of the layout of the visual system in all three castes. Adapted from reference20. Scale bar = 1 mm. Please click here to view a larger version of this figure.
Figure 2: Scanning electron micrographs of the ant visual system demonstrating the imaging capabilities of this technique. Top row shows different eye positions and eye sizes in: (A) Myrmecia nigriceps; (B) Opisthopsis pictus; and (C) Amblyopone australis (note the very small eyes, white arrow). Images acquired at high magnification showing: (D) the three simple eyes in workers of Myrmecia nigriceps; different sized compound eye in (E) Rhytidoponera metallica (note the different shaped ommatidia in different regions of the compound eye in yellow), (F) Amblyopone australis, (G) Myrmecia pyriformis, (H) Orectognathus clarki, and (I) Pheidole species. Scale bars = 1 mm (A-C), 100 µm (D-H), 10 µm (I). Please click here to view a larger version of this figure.
Figure 3: Cornea replicas of ant eye and ocelli. (A) Replica of the compound eye of a worker of Myrmecia nigriceps. The convex replica was flattened by making incisions. The inset indicates posterior (p), anterior (a), and dorsal (d), ventral (v) axes. (B) Replica of the ocelli of worker of Myrmecia tarsata. Scale bars = 0.5 mm (A), 10 µm (B). Please click here to view a larger version of this figure.
Figure 4: LM and EM images of rhabdom cross-sections. (A) Cross-section of distal rhabdoms in Myrmecia nigriceps stained in toluidine blue can be used to distinguish rhabdoms that are circular or rectangular in shape. Transmission electron micrographs show: (B) multiple orientations of microvilli in the circular rhabdom and (C) microvilli oriented in two opposite directions in the rectangular shaped rhabdom. (D) Using light microscopy, the long axis of the rectangular rhabdoms are mapped to show a fan-like organization in the dorsal region of the eye in a queen of Camponotus consobrinus; inset indicates posterior (p), anterior (a), and lateral (l), medial (m) axes. Panel D adapted from reference20. Scale bars = 10 µm (A), 1 µm (B-C), 100 µm (D). Please click here to view a larger version of this figure.
Figure 5: LM and EM images of an ommatidium in a light-adapted eye of Myrmecia tarsata. (A) Longitudinal section of an ommatidium showing the cornea (C), crystalline cone (CC), cone tract (ct), rhabdom (Rh) and primary pigment cells (PPC). (B) Dashed rectangular box in panel A from a different section viewed under a TEM to quantify the narrow width of the cone tract. Adapted from reference21. Scale bars = 10 µm. Please click here to view a larger version of this figure.
Figure 6: Common problems with semi-thin and ultra-thin sections (fixation and infiltration, cutting and staining). (A) Poor fixation of tissue due to inadequate penetration (arrow) in a semi-thin section of Pheidole species; (B) ripping during sectioning in Iridomyrmex calvus (semi-thin); (C) perfect staining (left) and over staining (right) with toluidine blue in Myrmecia croslandi; (D) pigment (circle) and tissue ripping during sectioning (arrows) due to poor matching of the resin and tissue density (resin too soft). Folding of the section (asterisks), can happen when collecting sections from the knife boat; (E) poor contrast due to insufficient staining (compare to inset), lead citrate crystals (white arrows) from exposure to CO2, and vertical knife mark (black arrows); (F) holes in the tissue (white arrows) caused by poor fixation in a Melophorus hirsutus compound eye; (G) resin too soft, leads to chitter when sectioning seen as vertical ripples in the section; (H) section too thick (~100 nm) resulted in dark image with poor contrast, contaminated distilled water lead to bacteria and particulate matter scattered throughout the section (white arrows) in Pheidole species. Scale bar = 25 µm (A-B), 10 µm (C-H). Please click here to view a larger version of this figure.
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 insects, albeit with slight modifications in the protocol (e.g., increasing duration of fixation and infiltration in thicker tissues). Slightly modified protocols have been used to characterize the visual systems of a variety of insects including cicadas22, flies14, bees23, wasps24, butterflies25, and moths26. Although most of the techniques outlined here have been in use for some time, this article takes the opportunity to bring them together in the context of studying the ant's optical system and compare alternative techniques and describe common pitfalls.
There are many imaging techniques currently available that have overlapping applications and it can be difficult to assess which technique is appropriate for the task at hand. A relevant example here is choosing a technique for overview imaging. The external morphology of the head and eye and relative positioning of the optical system on the head can be done using SEM or photomicrography. The strengths and weaknesses of these techniques have been reviewed27, however, there are some special considerations when imaging eyes. When imaging the relative positioning and size of the eyes, both techniques have their advantages and disadvantages. SEM images lack color information and hence where pigmentation is relevant photomicrography is better. However, SEM images can illustrate fine structures such as inter-ommatidial hairs and facet boundaries in greater detail and even reveal surface features not visible under photomicrography techniques (e.g., ocellar lenses, surface sculpturing of compound eye lenses). SEM is a versatile technique when it comes to exploratory imaging and identifying features of interest because it can operate on a large range of specimen sizes while still retaining very high resolution throughout this range. However, it is not as widely accessible as a dissection microscope and requires a higher level of expertise. There is often no single way of obtaining the information one requires. In such a scenario, it is useful to consider what is available and where it is most important to invest resources.
Nail-polish replicas of the cornea have proven to be most useful in obtaining the most accurate measure of facet numbers and facet diameters. This has now been used in a variety of insects11,22,28,29. While the quality of the images acquired from an SEM is far superior, the curvature of the eye prevents accurate measurements of the whole facet array. Mapping the facet size and facet distribution should also be feasible from scans acquired from micro-computed tomography5.
In both the LM and TEM techniques, it often is difficult to know whether the sample has been prepared and processed well until the final stage of imaging. To avoid complications, it is important to establish good practices such as maintaining clean working spaces and tools, preparing fresh solutions regularly, and thoroughly filtering water. Contaminants that are invisible to the naked eye can ruin EM samples. For this reason, it can be useful to wipe down surfaces and instruments using a solvent, such as ethanol or acetone, and a non-lint producing wipe. This is most relevant when sectioning, staining EM sections, and when preparing SEM samples. Similarly, distilled water sources can present problems and introduce contaminants so it is always best to check filters, change them regularly, and always use freshly filtered water (do not store). Most fixatives, stains, and embedding materials cannot be stored indefinitely and it is important to label all solutions with the date of preparation. It is important to take a systematic approach and set aside enough time to carry out protocols without interruptions.
Adapting techniques to different species is always a matter of trial and error. When working within Formicidae, the main differences lie in the size of the animal and the muscle mass within the head. Ants with more musculature in their head will typically take longer to fix. With very large ants, it is best to remove the mandibular muscles, trachea, and mandibular glands, while ensuring minimum interference with the neural tissue. In small ants and those with few mandibular muscles, it is possible to achieve adequate fixation by just removing the mandibles and exposing the clypeal region. In these cases, small holes using minutiae pins can be made on the head to improve fixation.
It is important to note that environmental conditions can also affect preparations. Hot and humid environments (especially field stations in the tropics) can prove to be a challenge during the infiltration stage. Warm conditions can lead resins to partially polymerize prematurely resulting in the unused resin becoming increasingly more viscous. In this case, the best option is to store the resin in small, single use, containers in the fridge or freezer. Cooling fixatives can be helpful to counter faster tissue decay in warm conditions. However, cooled solutions will disperse more slowly which means that treatment times should be extended to ensure proper penetration.
With these cautions in mind, investigation into the optical system of ants and other insects can prove very rewarding. Studying the visual system allows us to estimate the size of visual fields, interommatidial angles, optical sensitivity, and sampling resolutions. Understanding the anatomy of the eye informs our understanding and interpretation of animal behavior. For example, anatomy allows us to make predictions on the visual capabilities of animals such as whether they are diurnal or nocturnal, which may not have been previously documented. Given the current knowledge about the visual system of handful of ants, we hope our methods will inspire biologists and myrmecologists to investigate the compound eye and ocelli in ants to further our understanding.
The authors have nothing to disclose.
We are grateful to Jochen Zeil, Paul Cooper and Birgit Greiner for sharing their knowledge in insect anatomy and for demonstrating several of the techniques we have described here. We are grateful to the talented and supportive staff at the Centre for Advanced Microscopy at ANU and The Microscopy Unit at MQU. This work was supported by a graduate scholarship to FRE and grants from the Australian Research Council (DE120100019, FT140100221, DP150101172).
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 |