Fixation and preservation of Invertebrates
Download as PPTX, PDF14 likes17,490 views
1. Fixation involves coagulating cell contents to prevent degradation, using formalin for most invertebrates. 2. Preservation methods include storing in alcohol, sorting solution, or drying specimens. Alcohol is the preferred long-term fluid for both vertebrates and invertebrates. 3. Invertebrates can be killed via freezing, liquid chemicals in killing jars, or solid cyanide poisons which are highly dangerous. Freezing is now preferred for safety and specimen well-being.
Fixation and preservation of Invertebrates
- 1. M.Raj Associate Professor Dept. of Zoology Fixation and Preservation of Invertebrates
- 2. Fixation The fixation of biological specimens involves the coagulation of cell contents into insoluble substances with the purpose to prevent autolysis and the degradation of tissue. Fixation coagulates and stabilizes protein in specimens so that they do not distort or deteriorate during preservation, study and storage. A good fixation is generally achieved in a brief amount of time (hours to days) and as soon as the animal is collected. Formalin is generally the preferred fluid for fixation and is widely used. Invertebrates typically require only 4% formalin. In fish, amphibians, reptiles, birds, and mammals, the ratio of formalin to carcass must be at least 12 to 1 to
- 3. Preservation Preservation is any process that serves to keep the dead body of an organism from decay, in part or in whole, presumably to be studied later. Both vertebrates and invertebrates can be preserved in fluid or as dry specimens. The archival preservation fluid that has been used the longest and is generally preferred is alcohol. The standard is 70-75% ethyl alcohol or ethanol 40-50% isopropyl alcohol is used on some animal taxa. It tends to make them brittle over time. For this reason, one needs to buffer it with a few drops of glycerin and a pinch of calcium carbonate. "Sorting solution" (1.5 parts propylene phenoxetol, 5.0 parts propylene glycol, 10.0 parts full strength Formaldehyde, & 83.5 parts distilled water) has been used successfully for the long-tem storage of some taxa.
- 4. Invertebrate Killing methods Killing Jars Liquid killing agents are ethyl acetate, ether, chloroform, and ammonia water. Ethyl acetate is most widely used. All of these chemicals are extremely volatile and flammable and should never be used near fire. Solid killing agents are the cyanides potassium cyanide, sodium cyanide, or calcium cyanide. They are dangerous, rapid acting poisons with no known antidote and hence are to be handled with extreme care. Absorbent material- Plaster of Paris/Cotton
- 5. Invertebrate Killing methods Freezing Due to environmental and safety concerns, and the well-being of the specimen also, these methods have been done away with and are today being replaced by freezing. After the specimens have been collected, they can be transported home or to the lab in a plastic zip-lock bag or paper envelopes. The specimens are then carefully placed into a portion of the freezer where they will not be damaged. Leaving invertebrates in the freezer for prolonged periods of time however may damage the specimen. They are to be freezed only long enough to render them immobile.
- 6. Preservation of Invertebrates Dry Method Micropaleantology slide Amoebas and their relatives, including foraminifera and radiolaria, belong to the subphylum Sarcodina. Both groups produce skeletal casts or shells, casts of radiolaria are siliciceous (silica), those of the forams are calcarious (calcite). Foraminifera and radiolaria
- 7. Preservation of Invertebrates Dry Method Sponges Wear hand gloves Put under clear, running water rinse and squeeze to drain out excess water For removing the smell, the sponge is then placed in a container of alcohol with a lid for 48 hours. Before drying large specimens, labels are attached by means of string, threaded through the body of the sponge. Dry it in the sun When thoroughly dried, sponges are kept in small cardboard boxes supported with tissue paper. Paradichlorobenzene or naphthalene flakes may be added to dry containers.
- 8. Preservation of Invertebrates Dry Method Stony corals First, the animals using the coral as a home must be removed. This is achieved by soaking the coral in fresh water, as the animals are saltwater animals and can not survive for more than three days in fresh water. Then, the coral are to be soaked in two parts bleach and one part water for 48 hours. For preserving the coral specimen, it should be soaked in alcohol and let dry, but most e.g. brain coral, star coral, staghorn coral, elkhorn coral, and pillar coral. Coral: a hard stony substance secreted by certain marine coelenterates as an external skeleton, typically forming large reefs in warm seas
- 9. Preservation of Invertebrates Dry Method Horny Coral Specimens are dipped in neutral formalin for 15 minutes and then dried in a warm but shaded place. Prior to drying branches are so arranged that they will take up the least amount of space in a museum tray. After the specimens are thoroughly dried up they are kept in light proof boxes with a few crystals of paradichlorobenzene. e.g. Gorgonia
- 10. Preservation of Invertebrates Dry Method Dry shells of Mollusca The first step is to clear the shell of any dead tissue, which is a cause of bad odor. Most shells can be cleaned by coating them on the inside and outside or filling it completely with a dry solution of - three parts salt, one part baking soda and freezing them in zipper bags. An alternative to the process of cleaning dead tissue is to boil the shells.
- 11. Preservation of Invertebrates Dry Method Dry shells of Mollusca After removing the dead tissue, the shells are sorted out into sturdy and fragile shells. Sturdy shells are soaked in a container having four cups of water mixed with four cups of bleach and one tablespoon of dishwashing liquid. They are periodically scrubbed with a toothbrush to remove any seaweed, algae, barnacles and debris attached to them. Fragile and bivalve seashells are washed in mild dish detergent. After the seashell is clean, it is removed from the solution, rinsed well, and dried gently with paper towel. They are then air dried or sun dried. Some collectors like to use baby oil on a well-dried shell to bring out its luster.
- 12. Preservation of Invertebrates Dry Method Echinoderms (asteroids and ophiuroids) Clean in a mild detergent solution that is mostly water and very little detergent and dry in the sun. Soak the starfish immediately in rubbing alcohol for 48 hours and forego cleaning it. It is then allowed to dry in the sun. Soak it in formalin, which is 1 part formaldehyde and 5 parts water, and then dry it properly While drying it is necessary to put weight on every arm of the starfish, or else they will curl up
- 13. Preservation of Invertebrates Insect Preservation Relaxing methods Flat plastic container with an airtight lid makes an ideal relaxing dish. Base lined with moist cotton wool or synthetic sponge saturated with water and covered with a blotting paper. Phenol crystal/few drops of dettol added to prevent fungal growth A sheet of blotting paper is also placed on the inside of the lid to avoid condensed water to fall over the specimens. The dry insects are kept in the relaxing dish and the dish sealed and left at least for a day. Relaxing Chamber Alternatively robust insects (e.g.most beetles) can be quickly relaxed by dropping them into near boiling water. Small specimens will soften in a few seconds, whereas large ones require a minute or two. Large moths and butterflies can also be relaxed by injecting a 10% solution of ammonia or hot water into the thorax.
- 14. Preservation of Invertebrates Insect Preservation Insect Pins English: The English pins are 18 to 30 mm long and quite stout. These can be handled well with pinning forces and preferred for staging or double mounting. Continental: The continental pins came in three sizes a. 35mm long (numbers 000,00,0 and 1 to 7 ) b. 38 mm long (numbers 8 to 10 ) and c. 50mm long ( numbers 11 and 12). These are good for direct pinning. Mintuten nadein: The Minuten nadein are very fine, short ( numbers 10, 15, 20 or 22 mm) with or without heads and used for only double mounting of very small insects.
- 15. Preservation of Invertebrates Insect Preservation Mounting Insects Pinning Blocks: It is important that all the specimens and labels are placed at a uniform height on the pins. Setting/Spreading: Moths, butterflies, lacewings and dragonflies set with both pairs of wings spread, whereas Grasshoppers, cockroaches, mantids, stick insects and occasionally bees are set with only one pair of wings extended.
- 16. Preservation of Invertebrates Insect Preservation Mounting large insects Insects longer than around 8 mm are mounted by direct pinning. The body part through which the pin is placed differs in some orders of insects (Page 139)
- 17. Preservation of Invertebrates Insect Preservation Mounting small insects Card points: Small bugs, wasps and flies Card Platforms: Small insects, particularly certain beetles and parasitic wasps Minuten pins: This method is also called double mounting or staging. Gelatine capsules Glueing to pins
- 18. Preservation of Invertebrates Wet method
- 19. Preservation of Invertebrates Wet method
- 20. Preservation of Invertebrates Parasites Arthropods preserved in 70% alcohol in glass vials with screw caps lined with anti-evaporation inserts made of plastic. - Mites, ticks, lice, fleas, louse flies, and bugs can be dropped straight into alcohol. - Nest fly larvae should be killed by immersion in boiling water prior to storage in alcohol or, better yet, immersed for 24 hours in KAAD solution (1 part kerosene, 10 parts 95% alcohol, 2 parts acetic acid, 14 parts dioxane). If specimens are to be stored in alcohol for a long time (years), 5-10% glycerin should be added to the alcohol to prevent hardening of specimens To prevent small vials from drying out in long-term storage, they can be placed in an inverted position between layers of cotton in a larger jar of alcohol, so long as the caps are not loosened in the process. This ensures that the fluid in the vials will be the last to evaporate should they go unchecked. If specimens are to be mounted on slides, they should first be washed several times with 70% alcohol to remove the glycerol.