The Body Wall and Musculature of the Marine Triclad Palombiella stephensoni (Palombi, 1938): Part One: General Tissue Structure as seen with the Light Microscope
Materials and Methods
Materials and Methods
Specimens were collected from Island Bay and the western shore of Lyall Bay, Wellington, New Zealand. At low water the rock pools and rocky shore of the lower littoral zone were harvested for their green sea-lettuce (Ulva lactuca). This was transported to the laboratory and placed in large, shallow enamel trays with sufficient sea water to cover most of the sea weed. The Ulva was then examined and any Palombiella present were drawn into a pipette and squirted into bowls of sea water. The worms were kept alive and healthy in the laboratory in glass bowls 6 ins. in diameter and 2 ins. high. The water in the bowls was changed every two days, and the worms were fed finely chopped beef liver and muscle twice a week: for every 40 worms, 1 gm. of beef and liver mixture was placed in the bowls. After ½ hour the excess food in the bowls was discarded and the water was changed.
The worms proved difficult to narcotize. Substances commonly used for anaes-thetisation of flat worms such as carbon dioxide, ether, coal gas and alcohol gave poor results and caused the worms to contract markedly. A 50:50 mixture of ground chloral hydrate and menthol crystals sprinkled onto the surface of the water finally gave the best results. However, the worms frequently extruded the pharynx, and some discharge from genital openings took place.
The specimens were fixed in buff erred 4% formaldehyde using the buffer mixture of Pease (Pease, 1964, p. 52);in Carnoy's 3:1 ethanol: acetic acid; in Lilies' ethanol-acetic acid -formaldhyde (Pearse, 1960); and in picroformaldehyde (4% formaldehyde 75cc. sat. aq. picric acid 25cc. calcium chloride 1gm.). In all cases fixation was for 18 hours after which the animals were washed and stored in 70% tertiary butyl alcohol.
Whole mounts stained in acetic acid-alum-carmine and in the copper sulphate-silver nitrate nerve fibre stain of Betchaku (1960) were prepared after bleaching of pigment with 100 Vols. hydrogen peroxide.
For paraffin embedding the animals were dehydrated in 95%, and in two baths of 100% TBA for one hour each. They were then infiltrated in a 50:50 mixture of TBA: paraffin wax for two hours in a paraffin oven, and finally placed in pure paraffin (M.P.54 C.) for two hours prior to embedding. Embedding troughs made from brass blocks and adhesive cellophane tape (Wigglesworth, 1959) were used, and the embedding process was carried out with the aid of a binocular microscope at a magnification of 16x. Serial transverse and sagittal sections were cut at 5µ thickness, extended on a water bath, and placed on standard microscope slides without the use of adhesive. Sections were pressed onto slides after the method of Gray (1953). When treated in this manner they adhered vigourously to slides and withstood treatment by most reagents. Sections were subjected to the following staining procedures: 1. Heidenhain's iron haematoxylin with a counter stain of Orange G; 2. Delafield's haematoxylin with Van Gieson's picrofuchsin mixture as counterstain; 3. The triple stain of Delafield's haematoxylin, eosin Y, and fast green FCF (Wineera, 1968), and a variation employing Heidenhain's haematoxylin in place of Delafield's; 4. Mallory's triple stain (Gray, 1953); 5. The sulphuric acidhaematoxylin stain for basement membranes (McManus and Mowry, 1952); 6. Gordon and Sweet's silver stain for reticulin (Pearse, 1960); 7. The orcinol-new fuchsin stain of Fullmer and Lillie for elastic fibres (Pearse, 1960); 8. Betchaku's (1960) copper sulphate-silver nitrate stain for nervous tissue; 9. The Falg technique (Gurr, 1965). Stained slides were dehydrated through an alcohol series, cleared in xylol, and mounted in D.P.X.
The embedding process of Pease (1964) was used for methacrylate embedding. Sections were cut at 0.2µ and 0.5µ on a sledge microtome; were mounted on slides, and stained using Mallory's triple stain.