Preserving Plants - By: The Royal Botanic Gardens & Domain Trust

http://www.rbgsyd.nsw.gov.au/plant_info/identifying_plants/processing_plant_specimens/Preserving_plant_specimens

Pressing and drying Techniques for pressing and drying specimens have been established for many years. There are minor variations in recommended methods, but they are essentially the same worldwide. The best specimens are plants that are pressed as soon as possible after collection, before wilting and shrivelling. Most plants may be kept in sealed containers such as plastic bags for up to a day if it is inconvenient to press immediately. However, some plants show such rapid wilting, particularly of the flowers, that such delays are best avoided. Flowers with a lot of nectar may go mouldy very quickly if excess nectar is not shaken off before pressing. Specimens are pressed flat and dried between sheets of absorbent blotters or semi-absorbent paper such as newspaper. Papers with a glossy surface should be avoided because they are not absorbent enough to aid drying. The plant should be carefully laid out between the drying sheets, as their form at this stage largely determines their ultimate appearance. The flowers should be spread out with the petals carefully arranged, wilted leaves should be straightened and unnecessary shoots of excessively twiggy shrubs may be cut away. Large flowers (e.g. Nymphaea) or inflorescences (e.g. Telopea) are best cut in half lengthways before pressing. Large and/or succulent fruit is often best preserved by cutting both longitudinal and transectional (from different fruit) sections from them and drying these. Care is necessary to ensure that the maximum amount of useful information s preserved. Sheets of thick, preferably smooth-sided, centre-corrugated cardboard (such as used in cardboard carton sides), placed between the drying folders will assist air circulation through the press. These are particularly necessary when using a forced circulation of warm air. If such cardboard is not available, additional sheets of newspaper or wooden board (e.g. plywood) may be used to absorb moisture from succulent specimens. When plants are uneven in thickness, e.g. where flowers are borne on thick twigs or arise from a thick bulbous base, sheets of spongy plastic foam (polyurethane or similar) about 1 cm thick, placed between the newspaper folders, help to distribute pressure evenly across the specimen. If foam sheets are not available, several thicknesses of folded newspaper may be used. Care must be taken to ensure 'damp spots' do not develop in the press. When using foam sheets it is advisable to circulate warm air around the press or change the drying papers more frequently. Specimens are best pressed with moderate pressure, preferably in an arrangement that will permit as free a circulation of air as possible. This can be achieved by strapping the pile together in a press, i.e. between frames made, for example, from sheets of heavy (non-bending) cardboard, hardboard, plywood, pegboard or, best of all, a lattice of wood or weldmesh (see picture). Supplies of suitable materials can usually be obtained from packaging and cardboard manufacturers, who will cut materials to suitable sizes, or from hardware or building suppliers. The press frames should be the same size as or a little larger thatn the drying paprs. Amateur collectors often press small numbers of specimens by placing books or other weights on the pile of specimens, but this is not recommended as specimens quickly go mouldy without air circulation. The papers should be checked for dampness and changed when necessary. As the number of changes required will vary with the original succulence/water content of the plants and with the weather conditions, no exact guide can be given. Most plants should dry in less than ten days. Foir the first few days the paper should be changed daily, but after that time the frequency of changes needed depends on conditions and relative humidity levels. In tropical and wet conditions daily changes will be necessary throughout the drying period, but in drier conditions the last one or two changes need only be given at intervals of three or four days. Used paper should be discarded, or thoroughly dried again before re-use. When in the field for an extended time, drying can be aided by placing the pressed plants in a warm, sunny position during the day. In reasonably dry climates, drying is aided by securing the presses to the roof rack of the vehicle whilst driving in dry daytime conditions. If available, a hot-air fan directing air around the press will assist drying. Drying cabinets with a forced circulation of warm air are used in large herbaria to shorten drying time and to lessen the need to change drying papers, but are not necessary for small quantities of specimens. A few species regularly turn black on drying, but in general, brownish or blackish colours in the completed specimens, or the growth of mould, indicate that drying was too slow, often because the papers were not changed frequently enough in the early stages of drying. Microwave ovens Small numbers of specimens can be dried using a microwave oven. The technique recommended in the literature is to place the specimens between unprinted absorbent paper, for example butcher's paper, not newspaper, which is unsuitable because the chemicals present in the ink may cause a fire. The specimens should be put in a special press which should be of a microwave-safe material (wood, acrylic or polycarbonate sheeting e.g. plexiglass or perspex, NO metal components). If such a press is not available, sheets of cardboard can be placed above and below the specimens and then weighted down. Drying time depends on the power of your oven. In most cases drying is accomplished by irradiating at maximum power for 1-2 minutes per specimen, although it is often a case of trial and error. It is best to process no more than 10-12 specimens of average thickness per batch. Specimens are usually dried after the moisture that characteristically appears on the glass door has disappeared. If the specimen is damp when taken out of the oven, allow it to stand before re-radiating as moisture continues to evaporate from the specimen for some time. Care must be taken not to irradiate the specimens for too long. It should be noted that microwave treatment damages seeds and the cellular structure of the plants which may reduce the long-term value of the specimens. Alternative drying techniques Silica gel/other desiccants & freeze drying Alternative methods of drying plant specimens have been used for some time, but are mostly restricted to special purpose collections. The main alternatives are freeze-drying and drying in a desiccant powder such as desiccant silica gel. In general these techniques are used where it is essential to preserve the shape of a delicate plant of organ of the plant such as the flower. Freeze-drying has also been used to preserve the chemical composition of a plant as accurately as possible for later study. Disadvantages and special conservation problems of specimens dried in these manners are that they are particularly susceptible to damage. The dried parts are fragile, lack support and often catch on packing materials. They must, therefore, be packed especially carefully and stored in small boxes or tubes with some appropriate packing material that does not snag and break small projections. Acid-free tissue paper is often used. Drying in desiccant silica gel crystals or powder can also have the disadvantage that it is difficult to remove all traces of the silica gel after drying. Special preservation and processing techniques Wet or spirit collections Very fleshy or delicate structures, including small algae and orchid flowers, are best preserved in an air-tight glass or plastic jar with a liquid preservative rather than by drying. The type of preservative used should be clearly labelled in the jar. Such material is often referred to as a spirit collection or wet collection. Most material can be satisfactorily preserved in 70% ethyl alcohol (or 70% methylated spirit or denatured alcohol) with 30% water. Your pharmacist can make this up for you and it will keep indefinitely in a tightly stoppered bottle. Colours will fade quickly in spirit, however, so it is a good idea to keep comprehensive notes and photographs. Small algae Microscopic algae are often collected in a jar and in the water in which they were found. If the algae are to be stored for more than 2-3 days, a preservative needs to be used. Traditionally this has been the extremely toxic formalin - a small amount can be added to the water to make a 5% final solution, and the container labelled. This must not be sent through the post or by courier. Thee are some other equally toxic options, for example propylene phenoxytol, but none should be sent through the post. A safer option is to add sufficient concentrated alcohol or methylated spirits  to the water containing the algae to make a final solution of 70% alcohol. This treatment dilutes the algae making them difficult to find, so if they can be concentrated somehow first (e.g. by filtering) they can be stored in much less liquid. Another option is to fix the algae in formalin (or something similar) first, and then prepare a microscope glass slide with a permanent water-soluble mounting medium. See also Collecting & examining freshwater algae Some plants and certain climatic conditions require the use of specialised processing treatments. This is a brief summary: Succulent plants Very succulent plants e.g. cacti, many species of Ficus ('figs') and mistletoes drop their leaves entirely upon drying or remain alive for an excessively long period in the press. This is overcome by killing the plant before pressing, either by freezing the specimen for a few hours, dipping it in boiling water for a few minutes, or by using a microwave oven. The correct time in a microwave oven depends on the type of oven and the specimen itself, but is usually about 2 minutees. Succulent material is 'done' when it has a flaccid, water-soaked appearance. When the cell tissue has been killed (by freezing, scalding or radiation) the specimen will still require special attention until it has dried completely. The papers must be changed at least daily for the first few days, and complete drying in the case of cacti may take more than a month. An alternative technique is to place collected succulent material in 70% alcohol, as this preserves its original shape. Bulky specimens Very bulky objects (e.g. Banksia spikes, thistle heads) may be cut or sawn lengthwise before pressing. Orchids Orchids require particular care when pressing due to their delicate flowers. The flowers (at least one) should be spread out evenly so that the flower parts face the paper surface without creases or folds (never allow the parts to fold up or stick together). Alternatively, cut off each organ of a flower (three sepals, two lateral petals, a lip petal and the column attached to ovary) and spread these parts on the same piece of paper and then press. A superior method is to preserve the specimen as a spirit collection. Water plants These should be carefully laid on a sheet of paper, excess water removed, then pressed and dried in the normal way. Very soft water plants may require special treatment such as being floated onto a sheet of paper immersed in water and then dried (as is usual for marine algae) or preerving in alcohol or formalin solutions). See also Collecting & examining freshwater algae Large algae These can be kept damp for a day or so, but it is preferable to dry specimens immediately. If very soft or filamentous, such plants may be best arranged on the mounting sheet while in a dish of shallow water. The mounting sheet is placed first into the dish and specimens on the sheet then gently slid from the water. Because such specimens tend to adhere to the drying papers they are best pressed between a mounting sheet (to which the underside of the specimen may remain permanently attached) and a sheet of adhesion-resistant material (e.g. muslin) to prevent the top of the specimen adhering to the drying papers. Tropical conditions Under humid, tropical and coastal conditions special methods must be adopted to prevent rapid mould growth before the specimens can be placed in drying cabinets. Placing the entire bundle of drying papers and specimens in a plastic bag and adding a small quantity of ethyl alcohol (enough to saturate with vapour) is a method commonly adopted. This sometimes called the Schweinfurth method, after an Austrian botanist who collected extensively in tropical areas. Such methods alter specimen colours and should be avoided unless conditions make them essential. Mounting Mounting specimens prevents most fragile material from fragmenting and prevents specimens becoming separated from their labels. If the plant collection is a long-term project, specimens should be mounted on sheets of archival (permanent) cardboard or paper with archival-quality fixing media. These include stitching with cotton thread, dental floss, nickel-plated copper wire (for heavier specimens), narrow strips of archival paper, linen tape, or by using an archival adhesive such as methyl cellulose adhesive. A range of archival material is available from S& M Supply Company Pty Ltd. Dental floss can be used for bulky specimens by puncturing the sheet on either side of the specimen, threading the floss through and tying ends together in a simple reef knot. Another alternative is a clear, long-lasting 3M tape (Y8440) which is available as a special order from 3M ('Scotch brand') and their distributors. This tape has been in use in some Australian herbaria for approximately 15 years with good success. The use of tape is faster than most adhesives, and is easier to remove (by cutting and peeling from the specimen) if the specimen needs to be examined more thoroughly. Ordinary sticky tapes are unsuitable as the adhesive breaks down, becoming tacky and detached after a few years. One disadvantage of mounting specimens is that it can make parts of the specimen inaccessible for examination, so it is essential that this be borne in mind during specimen arrangement and mounting. For example, easily reversible mounting media should be used, specimens should be strapped to the sheet, rather than glued all over, and the specimen should be carefully arranged before it is attached so that it shows all features. Full-size herbarium mounting sheets are usually about 43 cm long x 28 cm wide. The plant name and accompanying field notes should be transcribed on a permanent label stuck to one corner of the herbarium sheet (the bottom right-hand corner being the most common) or, sometimes, annotations may be written directly on the sheet or card. Example specimen sheets from the NSW National Herbarium are illustrated in the diagrams. Cards 20 cm x 13 cm are a suitable size for personal reference sets of identified specimens but are unsuitable for research collections, Note: mounted specimens should not be placed in microwave ovens - adhesives often melt, and tape may ignite. Small pieces of material which may have become separated from the specimen (e.g. seeds) can be placed in small plastic bags and pinned to the sheet. Long-term preservation and storage The long-term preservation of dry plant specimens is largely dependent on protection from insect attack. Specimens collected by Linnaeus in the eighteenth century, and by Banks and Solander on the Endeavour voyage in 1788, are still excellently preserved. Pests and their control A range of pests attack dried plant material. The most common pests are insects and fungi, though rodents and other large animals can cause damage in poor storage conditions. Insects eat the material, the paper surrounding the material, and the adhesives and mounting media. Such insect pests range from psocids (book lice), which attack mainly the softer parts such as flowers and soft fruits, to tobacco beetles and carpet beetles, which can bore holes through the toughest of specimens. Many insects are particularly sensitive to relative humidity levels and do not thrive at levels below 50%. The most common and acceptable specimen treatments for insect control are: Freezing Freezing the specimens is the technique least dangerous to human health, and is very simple. The specimens must be frozen to -18oC or colder and kept at that temperature for at least 48 hours. In practice, when specimens are frozen in domestic deep-freezers in bulk and/or in boxes, it is necessary to freeze them for 72 hours (3 days and 3 nights) to ensure that the centres of thick specimens and specimens in the middle of large bundles are reduced to a low enough temperature for long enough time to kill all pests. Bundles of specimens should be sealed in plastic bags to avoid moisture condensing on the sheets as they thaw, or alternatively, dry air should be circulated around the parcel in a desiccating cabinet during re-warming. Microwave Specimens may also be treated in a microwave oven to kill any animal life present on them. Microwave treatment is a fast method for small numbers of specimens. The technique is similar to microwave drying of specimens except that a press is not essential for already dry material, and times may be reduced from those required for drying. No absolute guidelines can be given as it is best to use trial and error testing for each set of circumstances and different types of microwave, but times of 1-2 minutes per dried plant specimen should be adequate. Poisoning A traditional method of insect control was to poison the specimens with a chemical to make them unpalatable or deadly to pests. However, this is not recommended due to obvious health hazards. Domestic spray-type insecticide is of limited effectiveness and, to avoid staining, should not be sprayed directly on mounted sheets. Sprays may kill surface insects but, for instance, would not penetrate to insects living near the centre of a Banksiainfructescence or 'cone'. Many spray insecticides are now regarded as possibly detrimental to human health, so health and safety should be carefully considered before these are used. It is essential that specimens that have been poisoned be so identified, both to warn users of the health risks involved and to avoid misleading any later chemical research using the specimens. Insect deterrents A number of chemicals have been used or proposed for use as insect deterrents. Of these naphthalene (commonly found as 'moth balls') is probably the most commonly used in herbaria because of its reputation for reasonable effectiveness in insect control, coupled with low toxicity to humans. It should be noted, however, that naphthalene is poisonous if ingested, naphthalene dust can cause eye health problems for people with contact lenses, and chronic exposure is believed to be implicated in the formation of cataracts. Thee are also reports of naphtha vapour causing allergies and headaches and of possible carcinogenic effects at very high concentrations. Naphthalene in commercial quantities is most commonly available in flake or chip form. If left loose in containers/boxes it is more readily inhaled or ingested and is more likely to case problems to people with contact lenses than is naphthalene in block or ball form or naphtha flakes or chips encased in porous bags or boxes. If naphthalene is used as an insect deterrent the levels around specimens must be maintained at a steady, level to ensure effective insect control. Because of the exposure limits for humans this is best done by storing specimens in boxes or in a sealed cupboard. Fungal pests Fungal (mould) attack is mainly a danger to damp specimens, either through incomplete drying during specimen preparation, or to collections that become wet later through flood, other water damage or improper storage conditions. Properly dried plant specimens will not suffer from fungal attack if stored in the correct conditions (see recommendations below) though freeze-dried fungal bodies such as mushrooms have been reported to be very susceptible to mould growth. Specimens with sugary exudations or large quantities of nectar are also particularly attractive to fungi, and need special care during drying to ensure that they dry fast enough to prevent mould growth. If fungus grows on the specimens these can be brushed with alcohol or methylated spirits (denatured alcohol). However, this may alter the specimen unacceptably for chemical and other investigative research, and only kills the fungus present on the specimen; it does not correct the problems that allowed the fungus to develop. Specimens treated for fungal attack should be clearly annotated as such, including date and treatment given. Storage Dried and pressed plant specimens can be stored in cardboard or plastic boxes, or tied in bundles in light-weight cardboard folders placed in 'pigeon holes'. Alternatively, they can be placed in protective plastic jackets and displayed in ring folders which is recommended if they are to be frequently handled, such as for a reference collection. Filing Specimens should be filed in a systematic order if a relatively permanent collection is being made. The major groups, i.e. ferns and fern allies, cycads, conifers, dicotyledons and monocotyledons, are best kept separately or according to some classification scheme, such as that given in a flora or handbook. Similarly, the genera within each family and the species within each genus may be filed alphabetically or following some such classification.

Dermestid Beetle Colonies From a Professional - Courtesy of: Carla Brauer, Owner of dermestidarium.com

Care and Maintenance of a

Dermestid Beetle Colony

The Basics:

Dermestes maculatus, the Dermestid beetle, is a flesh-eating scavenger found nearly everywhere in the world. These small, dull black beetles and their hairy larvae can be found in nature feasting on the dried or decomposing remains of animals. Because of their ability to clean flesh from even the most delicate of bones, colonies are often kept by museums, forensic scientists and taxidermists, where these small creatures are enlisted as helpers in skeletal preparation. 
     
The life span of a Dermestid beetle is around four to five months. Adult beetles lay their eggs on or near a food source, and incredibly tiny larvae will emerge from the eggs about four days later. The larvae are the most voracious flesh eaters in a colony, and grow through seven to nine instars over the course of about a month and a half. With each new life stage, the larvae must shed their exoskeleton; new keepers of colonies will often worry about finding a lot of "dead" larvae that are in fact just the shed exoskeletons of maturing insects.

The larvae will then burrow into some available material – they seem to enjoy styrofoam, but will bore into meat, corrugated cardboard, or their own bedding – and emerge as beetles a week later. After a month, the female beetles (which are slightly larger than the males) will begin laying eggs, completing their life cycle.

With the right care and conditions, a Dermestid colony can multiply quite quickly, and the more the merrier as far as skeletal preparation is concerned!

Obtaining a Colony:

Although Dermestid beetles are a naturally occurring species, the best way to start keeping them is to buy a small starter colony from someone who has a healthy, large colony of their own. It is entirely possible to gather and breed wild Dermestids; however, you are taking the risk of accidentally bringing in beetle pests like mites or other insects/diseases that may wind up wiping out your contained colony entirely. Never introduce wild Dermestids (or any other insects) to an existing healthy colony for exactly these reasons.

I recommend Kodiak Bones & Bugs in Alaska as a trusted source of healthy starter colonies. You can also look on taxidermy forums or live insect keeping forums for people who have starter colonies available or can recommend places to look. 

Beetle Housing:

Dermestid beetles are simple creatures, and quite easy to please. The main goal in housing a colony is to keep the beetles contained, while allowing ample ventilation and enough space for your specimens to fit. Keeping your beetles where they belong is incredibly important. Escaped beetles have the potential to become pests that can eat your books, taxidermy mounts, carpets, and generally make a pain of themselves. With how quickly they reproduce, this is not a chance you want to take.

Glass aquariums and hard plastic storage containers can make good Dermestid housing, as they cannot climb the slick sides to get out. Note that Dermestids have been known to eat through Rubbermaid tubs, but the Sterlite brand seems to work perfectly. Make sure that you have a tight fitting lid, because at high temperatures, beetles are capable of flying; you want to keep the beetles in and other bugs out, while allowing plenty of ventilation and airflow. Cutting holes in sides of plastic tubs and covering them with fine screen is ideal. Screening the tops of lids seems like the obvious choice, but flies of all sizes will sit on top of the screen and lay eggs that can fall through remarkably small holes and infiltrate your colony. Broken chest freezers also make excellent housing for larger colonies, provided they have adequate ventilation. Beetles work best in the dark, so covering clear glass or plastic enclosures is a good idea.

As you can probably imagine, your beetle area will smell! While you can take steps to reduce the smell coming from your colony, you are basically keeping slightly warm rotting flesh around, and there's no way of avoiding a certain amount of stink. Keeping your colony outdoors is not ideal, as you risk too much temperature fluctuation and the potential for scavengers and insect pests to break into your colony. Beetle colonies are best kept in sheds, workshops, barns, or other areas with good ventilation and not much foot traffic. On the bright side, they don't smell nearly as bad as maceration, the alternative method for getting impeccably clean bones!

Bedding Materials:

An inch or two of bedding should be provided for a beetle colony. This can be shredded paper, cut up cardboard, wood shavings (except cedar, which contains a natural insecticide), or, my personal preference, Carefresh animal bedding, available at any pet store. If you are using corrugated cardboard, I recommend freezing it for at least 72 hours before putting it in your colony to ensure that any insects hiding in the corrugated gaps are killed. 

        
Odd as it sounds, styrofoam is your beetles' best friend. Add a large chunk of styrofoam to your colony, and you will see them quickly burrow their way in to pupate. While they will pupate in other materials, this seems to be their favorite. Over time, the styrofoam will hollow out completely and be reduced to small shredded bits that mix with their bedding.

The beetles will add to the bedding layer with their own frass, a powdery waste material, and chewed up bits of bedding materials and styrofoam. Allowing the bedding and frass mix to build up is good for the beetles – it gives them insulation from both overly warm and cold temperatures, as well a place to burrow and nest. When the beetles aren't eating, you'll find them buried in their bedding. Keep an eye on the bedding/frass layer, stir it occasionally, and make sure that it stays loose and dry. Too much moisture can lead to mold or promote mites, a potentially toxic situation for your colony. If you do need to remove some bedding or frass, it’s safe to assume that eggs and small larvae are stowing away in it no matter how carefully you check, so it’s a good idea to store it in the freezer until trash day to prevent runaway beetles.

Food & Water:

As you can probably guess, Dermestid beetles are 100% carnivorous. They survive on meat, and, while the beetles can go long periods without food, if you want to increase your colony size, you should have food freely available to them. You can give them freezer burned meat scraps or hot dogs when you don't have a specimen for them to clean. 

           
The most important thing to remember when feeding your colony is to avoid any possibility of introducing other insects or eggs. If there is even a small chance that what you're going to feed your colony has had a fly land on it and lay eggs, put it in the freezer for at least 72 hours to ensure that they are dead. Then defrost the meat in a sealed container and give it straight to your colony. 

The most common Dermestid keeping advice is to only give your beetles dried meat. This is what they typically eat in the wild, and preventing excess moisture in your colony is good practice for avoiding mold or damp situations that promote mites. However, I have found that my beetle colony eagerly accepts fresh and moist meat, and in fact seems to prefer it. If you would like to dry specimens prior to feeding your colony, you can set up a screened area with a fan, or set the specimen on a shelf in the freezer for a week or so.

           
Ideally, avoid giving your colony large pieces of meat that they can't finish in several days. This will help prevent excessive rotting, mold and moisture. Any bone specimens the beetles are to clean should be skinned and have as much tissue as possible (including eyes and brains) removed prior to being put in with the beetles. 

           
Every so often, when I'm not in a hurry, I will leave some brain in a skull for the beetles to eat. It's a feeding and egg laying frenzy every time! Brains seem to be their favorite food, and are a great way to motivate them to increase their population. Feeding small skulls (birds, rodents, etc.) with brains left in is a great way to increase the population of smaller colonies. Just be aware that the smell of rotting brain is a special kind of awful, so try not to give them more than they can finish quickly.

            
Beetles don't need much in the way of water. A folded up paper towel dampened weekly is enough to keep them going. Thirsty beetles and larvae will flock to the paper towel and have their fill. Remember that moisture in the bedding is bad, so the towels should be moist enough for them to drink, but not so wet that they don't dry within a few hours.

Temperature:

The temperature of your colony is incredibly important. The beetles work most efficiently at around 75-80 degrees Fahrenheit. While lower temperatures won't necessarily kill your colony, you will notice that they move much slower. Any warmer than 80ºF, and your beetles are capable of flight. No one wants to open a colony of flying Dermestids. 

          
The beetles conveniently thrive right around room temperature, and often all that is needed is some insulation around their enclosure and a reptile heating pad underneath in cooler weather. If that's not enough to keep your beetles in the proper temperature range, a heat lamp can be used. However, beetles do their best work in the dark, so this is less desirable.

Removal of Specimens:

When the beetles have finished cleaning the flesh from a skeletal specimen, it's time to remove it from the colony. When you do, you'll notice beetles and larvae hiding in every crack and crevice. My method for saving as many beetles as possible is to gently shake the skull and tap it against the side of the enclosure to loosen and fling out as many as I can. Holding a bright light up to the bone can help motivate them to emerge seeking darker pastures. 

          
No matter how sure you are that you've removed all Dermestids from a specimen, don't take any chances. You can kill any stowaways by immersing the bones immediately in hot water or putting it in the freezer for at least 72 hours. If you will be degreasing the bones, putting them directly into your degreasing bath should be enough to kill any remaining beetles. 

Good luck! :)

With proper care, your Dermestid colony will be ready to take on large projects like bear and deer skulls before you know it. These fascinating little creatures are a wonderful tool for anyone who regularly finds themselves with bones to clean. Their ability to clean soft tissue from even the tiniest and most delicate of bones will have give you excellent results that few other cleaning methods can equal.

Courtesy of a local shop owner:

Owner: Carla Brauer

503.560.9121 – Email: carla@dermestidarium.com

dermestidarium.com

{You can find the link to her website in my recommended sellers list!} 

Wet Preserved Reptiles and Amphibians - By: Richard E. Etheridge(1996). Modified/Updated By: The owner of this blog

Before reading: I do not recommend using alcohol as a preservative, from my research and from what I have been told by many professionals is that formalin is the only way to go. Please see the MSDS for 10% formalin and take all of the required precautions prior to even attempting preservation with formalin. Please note that this was written in 1996 and is not completely up to date; I have modified what he has written for this reason. If you would like to read the original document please see: http://www.lsa.umich.edu/ummz/herps/collections/herp-prep.asp

PLEASE NOTE: Do not remove any animal from the wild or pick up anything dead that you find with out checking with your area/states/government Department of Fish&Wildlife to see if it's legal to have in your possession. Any animal that is endangered or protected under the law is illegal to pick up and you should not touch it without risking getting a large fine.

Information written by: Richard E. Etheridge (adapted for the WWW and updated by M. O'Brien and G. Schneider, May 1996)

Most of the larger museums and universities that maintain preserved collections of reptiles and amphibians have curators trained in the approved methods of preparing and maintaining an alcoholic collection. On the other hand, many individuals with a non-professional interest in natural history have the inclination and opportunity to obtain and preserve herpetological specimens but lack knowledge of the proper techniques. It is the purpose of this article to be of help to these persons, for even small collections casually assembled may be of great usefulness if the specimens are adequately labeled, well preserved, and fixed in a standard position.

Steps for the preservation of specimens for sclentific study are as follows:

1. Injection and slitting. Liquid preservatives must be introduced into the body cavity, limbs and tail, either by hypodermic injection or through slits.
2. Fixing. While the specimens are still relaxed, they should be arranged in trays so that they will harden in the proper position.
3. Labeling. Each specimen should be accompanied by certain data, either attached directly or entered in a notebook with a number corresponding to a numbered tag tied to the specimen.
4. Storage. After specimens have been fixed in the proper position, they should be stored in liquid preservative for at least several days, after which they may be allowed to remain in the liquid, or transferred to plastic bags for temporary storage.

Preserving Solutions

Formalin: If at all possible, formalin should be used for injecting and fixing specimens. Formalin is the commercial name of a solution of formaldehyde gas (CH20) in water. This is sold only online. In Latin American countries, formalin may be purchased in many drugstores under the name "Formol" or "Formolina". Formalin must be diluted with water before it is used as a preservative. A strength of 10% formalin is best for most purposes. If the original strength is 40%, it should be mixed at a ratio af nine parts water to one part formalin. Issues with using formalin over other preservatives are: it is a carcinogen, it is not easily obtained, there are many precautions you have to take, it can be expensive. The positive to using formalin is that specimens almost never decay in it. Its other principal disadvantages are: it has a very irritating odor, it is very poisonous and may cause skin irritation or rash, it has a tendency to make specimens become brittle if the solution is too strong(depending on the type of animal), and tends to fade out certain colors rapidly, and it must be stored in rustproof containers. (Buffering of the 10% solution is recommended as formalin is slightly acidic. One buffering system that may be used is a mixture of monobasic and dibasic Sodium Phosphate, at 13 gm/gallon [Monobasic] and 24 gm/gallon [Dibasic]).

Alcohol: I DO NOT RECOMMEND THIS FORM OF "PRESERVATION" WHAT SO EVER. This "preservative" is easily attainable, but not recommended by myself(owner/writer of blog). For injection and fixing it should be used at full strength you need to use 95% or higher. For storage of reptiles it should be used in the proportion of 3 parts 95% alcohol to 1 part water. Alcohol which has been stored in open containers loses its strength rapidly due to evaporation. Strength may be tested with an alcoholometer. Specimens which have been fixed in alcohol should be carefully watched for signs of rotting(this is because alcohol is not a good preservative, it may preserve the skin but its innards are definitely going to rot). 

Preparation: If specimens are to be made permanently immune to decomposition, it is necessary that liquid preservative be introduced into the body cavity, limbs and tail within as short a time as possible after the animals have been killed. This may be accomplished either by injection (with a hypodermic syringe) or by making deep cuts with a sharp scalpel, razor blade or scissors(I personally recommend using a syringe). The most satisfactory way is by injection. A ten or twenty cc. syringe with a needle lock and several needles (guages 18 to 26) will serve to inject most specimens.

Frogs and Toads: Injection should be made through the belly, directly into the body cavity. If the body is puffed with air, it should be deflated by gently squeezing with the fingers. Very small frogs require only a few drops of preservative; frogs two or three inches long only a few cc. Introduce only enough preservative required to make the specimen look natural--it should not look bloated. It is not necessary to inject the legs of any but the largest frogs. If equipment for injection is not available, a single slit may be made in the abdomen, to one side of the midline. The slit should be deep enough to allow free access of the preservative into the body cavity.

Salamanders: Most salamanders do not require injection or slitting. If your specimens look "caved in" a small amount of preservative may be injected into the body cavity, or a single slit made in the abdomen to permit preservative.

Tadpoles: Tadpoles and small salamander larvae should always be preserved in 10% formalin, never in alcohol. Simply drop the tadpoles into formalin while they are still alive. Be sure there is enough preservative to cover them and avoid overcrowding. After 24 hours all the liquid should be drained off and replaced with fresh formalin.

Lizards: Injection should be made through the belly directly into the body cavity. Care should be taken not to use too much, or the body will become unnaturally distended. A series of slits should be made in the under side of the tail with a sharp scalpel or razor blade. The slits should be from 1/8 to 1/4 inch long and about 1/4 inch apart, and should extend from the base of the tail to the tip. Very large lizards must be injected or slit in the thicker portions of the limbs and neck. If space does not permit preservation of very large lizards whole, they may be skinned out, except for the head. To skin a large lizard, make a cut down the belly from the neck to the base of the tail. Work the skin loose from the body, pulling the skin of the arms and legs inside out as far as the wrists and ankles. Do not attempt to skin out the head, hands, feet or tail. Sever the wrists, ankles, neck and base of the tail, and remove the carcass. The skin should then he placed directly into preservative. If possible, one hemipenis of male lizards should be everted. This can be accomplished hy injecting preservative into the base of the tail (before slitting) and at the same time applying pressure with the thumb just behind the anus (Fig. 1 A).

Snakes: Make a series of injections an inch or two apart(sometimes 3") through the belly into the body cavity. Begin just behind the head and continue the injections to the anus. If a syringe is not available, a series of slits must be made in the belly. For most snakes the slits should be about an inch apart and an inch long; smaller slits closer together for very small snakes. Just as in lizards, a series of slits must be made in the under side of the tail and one hemipenis everted in males (Fig. 1B). Very large snakes may be skinned out, leaving the head and tail attached. To skin a snake make a single, long cut in the belly, just to one side of the midline, beginning about an inch behind the head and continuing to about an inch in front of the anus. Do not cut through the anal plate. Work the skin loose from the body, but do not attempt to remove the skin from the head or tail. Sever the body an inch behind the head and an inch in front of the anus, and (after recording the stomach contents, number of eggs, embryos, etc.) discard the carcass. Put a strip of cloth on the inner side of the skin and roll it up, beginning at the head. Tie the roll with a piece of string and put it directly into preservative.

Alligators and Crocodiles: Small individuals may be preserved just as lizards. Larger individuals should be skinned out with the head attached, rolled up and placed directly into preservative.

Turtles: Preservative should be injected into the body cavity just in front of each of the four limbs, between the carapace and plastron. Use a long needle and continue injections until the head and · limbs are forced out of the shell. If a syringe is not available, make deep cuts into the body cavity just in front of each leg. Limbs, neck and tail should be injected or slit, as in large lizards.

Below are some old practices and are not required, you can pose your animal however you want. It is required that you check if the animal you want to preserve is legal. 

Specimens should be injected (or slit) and tagged as soon as possible after they are dead and fixing should immediately follow. If you are unable to preserve at this time, store your animal in a plastic bag or plastic container and in your freezer(this helps keep your specimen from decaying prior to preservation). Individuals may be placed close together on the tray but should not touch each other. The tray should be covered to prevent evaporation. Most amphibians will harden in a few hours, reptiles in 10 or 12 hours. Large lizards, frogs and turtles may take a little longer.

Snakes: Small snakes may be coiled flat in the tray if the coil does not exceed three and one half inches in its outside diameter. The head should be inside as in Fig. 2 C. Larger snakes should be coiled in a jar and covered with preservative. If the snake has been injected it may be coiled with the belly down, tail at the bottom and head on top as in Fig. 2D. If slits are used, it should be coiled with the belly up, head on the bottom and tail on top. Tall, narrow bottles should be avoided; quart and pint sizes are best. Snakes too large to coil in a gallon jar should be skinned.

Lizards: Place the lizard belly down, with arms, legs and tail extended. If the tail is very long it may be bent around the side of the body (Fig. 3 A). Attenuate, limbless lizards may be coiled like snakes.

Turtles: Most important in fixing turtles is that the head and neck be extended and the mouth propped open with a bit of wood or cork. The limbs should also be extended if possible.

Salamanders: Belly down, arms, legs and tail extended as in Fig. 3 B. A salamander tail will often twitch back and forth long after the animal seems to be dead. Ten or fifteen minutes after they have been laid out check to be sure the tail is still straight. Large specimens, 10 or more inches in length, may be coiled like snakes.

Frogs and Toads: Place the frog belly down, arms and legs extended as in Fig. 4 A. The fingers and toes should be separated and extended, especially if they are webbed. The inner margin of the forelimb from the elbow to the tip of the fourth finger should form a straight line. The sole of the foot may be up or down, whichever seems most natural (down in treefrogs and up in most other frogs and toads).

I do recommend the following

Storage: After the specimens have been injected or slit, tagged(not required unless your fish and wildlife department directs you to do so), and fixed, they should be put directly into preservative. If they are to be transferred later to plastic bags for temporary storage or to be shipped they should first be allowed to remain completely immersed in preservative for at least 48 hours if formalin is used, or a week if alcohol is used. The longer they are allowed to stay in preservative, the better. They should be loose and completely covered with plenty of liquid. Specimens which have been hardened in trays should also be allowed to soak in preservative for a day or two before being shipped or placed in plastic bags for storage. If space is no problem, preserved specimens are best kept in glass containers. Bail-top jars with a glass top and rubber gasket are best. Fruit jars with a metal screwtop lid may be used but should be carefully watched for rust and evaporation. Glass jars with polyethylene lids and liners are more commonly used in collections, since the lids form a tight seal and are easier to obtain than the traditional bail-top jars. Metal containers should be used only for temporary storage unless coated on the inside with paraffin, "Bakvar", or some other rustproof material.

Specimens should be loosely packed and completely covered with liquid. Containers must be periodically checked for evaporation and refilled if necessary. At the first sign of decomposition the affected specimen should be removed and thrown away, or deep cuts made into the body cavity and placed alone in a large container with plenty of fresh preservative. A green spot on the abdomen of a snake or lizard indicates a rotten gall bladder which should be cut out. Any specimen that floats in the preservative contains air or other gases and is not properly preserved. It should be squeezed or slit to permit the gases to escape and the preservative to enter.

When in your personal possession: 
Most people recommend storing your specimens in 70% isopropyl alcohol because it's cheap and readily available. There are other holding liquids but it all varies on personal opinion and what you find works best for you.

Preserving Insects With Hand Sanitzer

Highly recommended - Watch Sam Droege's youtube video on how to preserve insects with hand sanitzer! I recommend drying any bug prior to putting your specimen into the hand santizer from personal experience just like Sam says to do in his video. I'm not exactly sure if this works with animal specimens, but I have seen it been done and I'm sure you should dry/dehydrate the animal you plan on using prior to attempting this process. Note that in the video he says to not use any fresh material(insect or animals) and also take not that if your specimen of choice is light weight, it will float indefinitely, if it's a heavier specimen it may sink over time and you need to be gentle with it! Good luck to those of you that try this:)

Instructions: Watch the video

Materials Needed:

1. Hand-sanitizer, try to buy some that doesn't have any bubble's in it

2. Plastic or clear glass container with air tight lid

3. Dried specimens or specimens that have been in alcohol, make sure these have been dried or in alcohol prior to this process. You can buy pre-dried bugs on Etsy and on Ebay for decent prices

4. Pipette/Syringe - you can find disposable pipette's on Etsy.com for under a dollar!

5. Paper clip/Thin wire/Tweezers

6. Super Glue

7. Patience

Another good reference (http://www.slideshare.net/sdroege/how-preserve-insect-specimens-in-hand-sanitizer)

• 1. How To Preserve Insect Specimens in Hand Sanitizer
  By DejenMengis, Carl White, Moriah Browning, Denise Williams, Sarah Fisher
• 2. Purpose and History
  Hand sanitizer is a gelled alcohol and can be used to create very cool insect specimen displays that, unlike pinned specimens, can be handled by children and the public
  Specimens will appear to float in air inside the vials and do not sink or move despite any amount of handling (Sam Droege has kept a vial in his pocket for 2 months without any shifting, for example)
  This technique was shown to us by Wayne White, BCE, of American Pest who has used hand sanitizer to preserve and display bed bug specimens
• 3. Pour hand sanitizer into an empty vial
• 4. Choose a dried, alcohol- or glycol- preserved specimen
• 5. Which Specimens to Use?
  Specimens that have been in alcohol, glycol, or dried work well
  Specimens that are freshly killed appear to dissolve the gel in the sanitizer for some reason (so dry out a specimen for 1-2 weeks then soak in 70% alcohol for 24 hours, then soak in hand sanitizer for 24 hours prior to displaying)
• 6. Move the specimen toward the bottom of the vial with a probe 
• 7. Bubble Removal: Water Bath Method
  At this point, there are probably many air bubbles in the vial. The next series of slides demonstrates how to remove these bubbles
  Air also exists inside specimens and needs to be removed or bubbles will gradually migrate outside the specimen over time
• 8. Pour an inch of water into a pot
• 9. Place the vial or vials into the pot, don’t forget that hand sanitizer will burn if exposed to an open flame! (if you are using a plastic vial, it will melt. So for this method you will need a glass vial)
• 10. Boil the vial in the water for 10 minutes or until most of the bubbles are gone
• 11. Carefully, take the vial out of the water
• 12. Use a pipette with a bulb to remove any remaining bubbles (you can do this instead of boiling)
• 13. Top off the vial with more hand sanitizer (Fill to the top, it's okay if it overflows when you put on the lid)
• 14. Position the specimen as preferred Even dried specimens become flexible (You can use a paper clip or tweezers to move your specimen)
• 15. Tips…
  Thoroughly clean vials before use
  You can add labels to vials that “float”
  You can add things like beads, dried flowers, sand etc. that will also float in place…and makes good “clean” fun for kids to make their own insect dioramas. OK, some adults like this too
  Always be sure to overfill with hand sanitizer and if possible permanently seal the vials to eliminate bubbles
• 16. Alternative Bubble Removal: Vacuum 
  Works reasonably well but….
• Specimens and hand sanitizer tend to bubble out of the vial more easily as air bubbles expand under vacuum
• 17. Not as efficient as heating in removing all the bubbles in the vial
• 18. Vacuum pumps are expensive

Also see these helpful links:
 http://www.entsoc.org/PDF/Pubs/Periodicals/AE/AE-2004/winter/Musings.pdf
http://thedragonflywoman.com/2011/02/21/hand-sanitizer-preservation/

Wet Specimen Preservation {Liquid Preservation}

So you're interested in preserving a cool critter or a dead pet, but you can't seem to find the right information. I am not a professional in this subject field, but from what I've researched and learned this is what I do know, take this information and experiment with different ways to preserve. The process that I have decided to share with you is just the basics, everyone has their own way of doing it and some have done this for many years and have figured out what works best for them. 

Facts and History - ( info listed below can be found at: http://www.amazon.com/Fluid-Preservation-Comprehensive-John-Simmons/dp/1442229659/ref=sr_1_3?s=books&ie=UTF8&qid=1420749967&sr=1-3&keywords=wet+preservation )

Fluid preservation refers to specimens and objects that are preserved in fluids, most commonly alcohol and formaldehyde, but also glycerin, mineral oil, acids, glycols, and a host of other chemicals that protect the specimen from deterioration. Some of the oldest natural history specimens in the world are preserved in fluid. 

Despite the fact that fluid preservation has been practiced for more than 350 years, this is the only handbook that summarize all that is known about this complex and often confusing topic. Fluid Preservation: A Comprehensive Reference covers the history and techniques of fluid preservation and how to care for fluid preserved specimens in collections. 

Although most fluid-preserved specimens are found in natural history and medical museums, it is not at all uncommon to find them in art museums, history museums, and science centers or even in the homes of collectors. In addition to animals, plants, and anatomical specimens, fluid preserved collections include some minerals and fossils and many other objects. 

A book called(referenced in the last paragraph) called, Fluid Preservation is an essential reference for: 

• Natural history curators
• Natural history collections managers
• Conservators
• Medical and anatomical museum collections managers and curators
• Art and history museum staff who have fluid preserved specimens and objects in their care (e.g., works by Damien Hirst)
• Private collectors
• Researchers using museum collections as sources of DNA, isotopes, etc.
• Health and safety professionals
• Exhibit planners and designers
• Museum facilities planners and managers
• People interested in the history of science
• People interested in the history of natural history museums
• Museum studies students

Getting Started:

Step 1: DO YOUR RESEARCH! This blog will supply you with a lot of information but not everything, find out which process works best for you! Quick tip: Rubbing Alcohol is a natural solution, but simply placing a dead animal in this will not preserve it properly. You have to fix the tissues with formalin prior to this and go through a long process. Alcohol will not preserve the organs and the insides of your specimen will begin to rot over time.

Step 2: Get all of the right chemicals and take the proper precautions prior to preservation.

WARNING: The chemicals that I recommend for this process contain carcinogens, which are also known to cause cancer. Read the MSDS on any chemical you buy prior to purchase!

Step 3: Pick your animal, make sure it has been frozen almost immediately after death or within the first few days, you can't preserve rotten animals. Also, make sure you have a jar big enough for your critter! ATTENTION: Before picking up road kill or picking up any animal that isn't domestic, be sure to read your state's (and government) laws and regulations prior to picking it up. Many animals are protected and endangered, picking up or preserving a protected animal or bird, is in many cases illegal!! You can be charged and fined for having it in your possession. Depending on where you are if you find a dead cat or dog that isn't fetal, it is also illegal to obtain. Fetal animals are considered organs therefore are legal to have (read more on this below). Also, never pick up or touch a dead animal with your bare hands/without gloves! You don't know what diseases it may have, if you suspect it of a disease I would avoid it.

Below is a great resource with state by state laws:

www.thegreenwolf.com/animal-parts-laws

Make sure to read up on wildlife laws, and make sure you read about the Migratory Bird Act (which makes illegal the possession or sale of birds except European starlings, English house sparrows, and captive-bred or legally taken game birds and pet birds), CITES (an international treaty dealing with protected species), and the Dog and Cat Protection Act (which makes illegal the purchase or sale of any cat or dog specimen with the exception of bones or internal organs, unless mounted by a taxidermist for the original owner). Roadkill laws vary from state to state. 

Precautions and Material List:

1. READ THE MSDS!!!!!!
2. Your area of choice for this process should have A LOT of ventilation and air flow in the room! Breathing in this chemical can cause dizziness and you could potentially faint!
3. Buy a MSA rated respirator mask with cartridges that ventilate out toxic dust and chemicals
4. Bins for your chemical baths, don't use these for anything but a specific chemical, label as poisonous and write warning if you think anyone may get into them.
5. Syringes, you will use this to inject the chemical into your animal.
6. Heavy duty and disposable gloves. You will use the heavy duty rubber gloves(no fabric!) during any formalin process, use the disposable gloves when touching any dead animal!
7. A freezer, you should keep all of your animals frozen when they aren't going through the preservation process! Rotten animals won't preserve properly
8. Plastic bags, to store your animals in, if you put several animals in one bag they could potentially freeze together!
9. A designated work bench/table
10. Jars to store your specimens in after preservation
11. The right amount of each chemical for the process, see below in the next segment.

The chemical's I use and my preferred process:

Please note that everyone has a different process! First read the precautions and material list and gather the correct materials before you attempt preservation.

10% Buffered Formalin: 
This chemical is 96% water and 4% Formaldehyde. YOU MUST READ THE ATTACHED MSDS BEFORE YOU PURCHASE! Click on the link below to read the MSDS. Formaldehyde is a carcinogen, meaning it is known to cause cancer, 10% buffered formalin is a lot safer than most preservatives, just make sure you read into it(See MSDS) prior to purchasing it.

Shttp://www.sigmaaldrich.com/MSDS/MSDS/DisplayMSDSPage.do?country=US&language=en&productNumber=HT501128&brand=SIGMA&PageToGoToURL=http%3A%2F%2Fwww.sigmaaldrich.com%2Fcatalog%2Fproduct%2Fsigma%2Fht501128%3Flang%3Den

There is no need to mix your own solution to make Formalin, it's too hard, too smelly and the amount of money you save by mixing your own solution is quite small, unless your making many gallons. You won't need a lot of Formalin, as a little does go a long way. Injecting takes a little formalin, I recommend not using old formalin when injecting, and you should have a “bath for soaking” that can be reused many many times. ALWAYS rinse in distilled water, small animals really only need a rinse, larger ones a few hours soak. There is no need for them to sit in water for a few days unless you have a very rare or expensive specimen. Technically you should have a few different alcohol solution baths from 40%-70% in 10% increments. Each with a separate soak of a few days. Again, not really necessary unless you have a special piece. If you are doing something very fleshy, like a fetal animal or let's say a hairless cat, a stepped soak in a 40/50% alf solution is good before plopping it in a 70% isopropyl solution(70% isopropyl is what you can store your specimen in after the process, some store in formalin but that can be spendy as you should replace it's holding solution every 3-6 months). This helps from shrinkage and dehydration. There is a great product you can use if you get an old dehydrated specimen that will in some cases bring it back to original. Jacob Cain, owner of Death Isn't The End(recommended seller), uses a chemical in his respiration work, its called Decon 90. If used, you must go through the whole preservation process again, most of the time. Another great resource, is a new published book on the history and how too’s for wet preservation. It's really spendy but its been a good book for to use if you're serious about taking up this hobby. The name of this book is - Fluid Preservation: A Comprehensive Reference by: John E. Simmons. This book is priced at $90.25 new. The book focuses on recipes new and old and also in restorative work. 


also see these helpful links for wet specimens: 
http://www.taxidermy.net/forums/IndustryArticles/02/k/02985396A5.html
http://conservation.myspecies.info/node/33
http://www.ou.edu/research/electron/bmz5364/prepare.htm