Processing Embryo, Eggshell, and Fungal Culture for Scanning Electron Microscopy

We have standardized sample preparation protocols for imaging painted turtle embryos, rigid eggshells and fungal cultures using scanning electron microscopy. These comprehensive methods use subtle alterations to known protocols to process three delicate tissues allowing the differentiation between original structures and processing the artifacts. Demonstrating the procedure with me will be Jessica Gibbons, a graduate student from my laboratory.

Before collecting turtle eggs from a field site during nesting season, make four to six 0.25 centimeter holes along the sides of plastic boxes and on their lids to allow aeration. Before filling the boxes with a moist mixture of vermiculite and peat moss bedding mixture at a one to one ratio. At the nesting site, gently remove the soil to uncover the eggs and wipe the surface of each turtle egg with a diluted iodine tincture to protect against microbial contamination during incubation.

Place a maximum of eight to nine eggs per box in the same orientation and alignment as they were laid, keeping individual clutches separate from each other. When all of the eggs have been collected, manually bury the eggs half in the bedding and place the lidded boxes inside a 30 degree Celsius incubator for 10 to 17 days to obtain embryonic stages 12, 13 and 18 embryos respectively. Add distilled water to partially wet the bedding medium every other day to avoid dehydration, and to maintain the moisture level for normal embryo development.

At the appropriate stage of development, use pointed scissors to make a cut onto one side of the dorsal eggshell and yolk membrane together, vertical to the long axis of the egg. Next, cut the other side of the egg along the short axis, and use forceps to peel open the excised piece with the embryo side up. Cut the other lateral side of the eggshell and place the excised piece of shell into PBS.

Using a stereomicroscope and forceps, peel the embryo and yolk membrane from the eggshell. Use the forceps and micro scissors to remove the extra embryonic membranes, and use an embryo spoon to transfer the embryo into a Petri dish of fresh PBS. After washing away any blood and yolk, place up to three embryos per well of a 12-well plate, containing 4%paraformaldehyde at four degrees Celsius for two to three days.

When the samples appear white, use polystyrene inserts with polyester mesh bottoms to rinse the embryos with three five minute washes in fresh PBS per wash. After the last wash, dehydrate samples in an ascending ethanol series for one hour per concentration. After the second ethanol immersion, dry the embryos in a series of HMDS to ethanol concentrations for 20 minutes per immersion with the Petri dishes partially covered.

Then place the embryos in a final 100%HMDS solution, completely or partially covered in a fume hood overnight. Slow drying by gradually increasing the concentration of HDMS to ethanol is important for revealing an excellent surface structure in the embryos. The next morning, use double stick carbon conductive tape to carefully mount the completely dried samples on a standard aluminum pin stub.

When all of the samples have been mounted, introduce the samples into the sputter coater chamber to coat the specimens with a very thin film of gold for 60 to 120 seconds at a 35 milliamp sputter. Then securely fasten the set screws to mount the stubs onto the corresponding pour plates. Use the sample exchange tool to transfer the sample holder into the sample chamber of the scanning electron microscope for imaging.

To prepare eggshells for scanning electron microscopy, place the eggshells in distilled water for at least one hour after the embryo harvest to eliminate any yolk or albumin contamination. Next, air dry the cleaned eggshells on delicate anti-static wipes in the fume hood overnight. Store the dried eggshells in clean specimen bottles, labeled by number and embryo stage.

Then, mount, sputter coat and image the eggshell specimens as just demonstrated for the embryos. To establish slide fungal cultures, use a sterile scalpel to cut half to three quarter inch blocks of solid potato dextrose agar, and place the blocks on individual glass microscope slides. Place sterile toothpicks on the bottom of clean Petri dishes for raising slides off of them to create surface tension between the plate and the slide.

Place each slide on the toothpicks in the Petri dish. Next, use a sterile loop to transfer fungi from the specimen inoculum to each of the four lateral sides of an agar block. Add a few drops of sterile distilled water to the Petri dish around the slide to ensure moisture for the growing fungi.

Partially seal the plate with paraffin film and place the plate in the 30 degree Celsius incubator for the appropriate incubation period for the fungal species. At the end of the incubation, remove the slide from the Petri dish and use sterile forceps to transfer the tightly adhered agar block from the slide to a container of 3%glutaraldehyde for an overnight incubation at four degrees Celsius. The next morning, dehydrate the samples in an ascending ethanol series for 15 minutes per concentration.

After the 90%immersion, process the samples for final dehydration with two 30 minute changes in 100%ethanol to ensure a complete saturation before placing the dehydrated samples in the chamber of the critical point drying apparatus. Seal the chamber and open the valves to allow liquid carbon dioxide to vent in and to let the ethanol vent out until liquid carbon dioxide completely fills the chamber. Next, seal and slowly heat the chamber to achieve a critical point when the chamber pressure exceeds 1, 000 pounds of pressure per square inch, the temperature exceeds 31 degrees Celsius, and the liquid and gas space of carbon dioxide is in equilibrium.

Then slowly drain the carbon dioxide from the chamber and the sample as a gas, to avoid any effects of surface tension on the sample, and mount, plate, and image the fungal specimens as demonstrated. A lateral scanning electron micrograph view of a stage 12 painted turtle embryo reveals how the maxillary prominence extends beyond the mandibular and medially limits a well marked nasal pit. Five pharyngeal arches can also be observed.

In a stage 15 embryo, newly formed somites are visible at the posterior tail region of the embryo, and the forelimb buds point more cortally than ventrally. A well-defined outgrowth of a carapace ridge can also be visualized along the entire interlimb flank region of the embryo. At stage 18, a short feebly projecting snout has formed with a slightly upturned lower beak with a terminal hook that fits into the central notch of the upper beak.

The upper jaw exhibits a notched appearance, and a tiny egg tooth can be seen forming at the tip of the upper jaw. Ultrastructural analysis of the Chrysemys picta eggshell and shell membrane by scanning electron microscopy, reveals an outer calcarinus eggshell layer, firmly attached to the inner filamentous shell membrane. The outer surface of the eggshell consists of well-distinguished mineralized shell units made of globular spherical nodules arranged in groups and in between adjacent shell units, with a concentration of small rounded depressions or pores of various sizes.

Common methodologies cannot be generalized for all biological specimen to achieve a decent image quality using SEM. Unique alterations as demonstrated are essential for a specific tissue types. The chemical and air drying method could also be used for other organic samples, and the coupling slide culture technique could be used for all soft, robust microbial cultures.