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Revised edition, 15 June 1994
R. B. Halliday

The following announcement was broadcast over the entomology and systematics electronic bulletin boards :

I am interested in hearing of people's experiences with specimens mounted on microscope slides. I refer mainly to mites, but insects or insect parts are also relevant. Specifically, what are the advantages and disadvantages of different mounting media, such as Euparal, Canada Balsam, gum/chloral media such as Hoyer's, Berlese's etc., and others?
How good is the optical quality of these media for your specimens?
How easy or difficult is it to prepare and mount specimens?
How easy or difficult is it to remove a specimen from a slide for re-mounting?
How successful are these media for long-term preservation of important specimens such as types and vouchers?
Are old specimens on slides in your collections still in good condition?
If specimens have deteriorated, what is the nature of the problem?

If responses are sufficient and useful, I will compile a summary and post it.

Thank you for your interest.

Bruce Halliday. 3 May 1994.

Dr. R. B. Halliday
Principal Research Scientist (Acarology) International Fax 61-6-2464000
CSIRO Division of Entomology Local Fax (06) 2464000
GPO Box 1700 Telephone 2464085
Canberra ACT 2601 Internet

Responses were received from 29 people. These are referred to below as Contributors 1-29. Their lightly edited responses are roughly sorted into groups under headings that refer to the different media mentioned.

Contributor number 7

I have made many slides of various invertebrates including mites. Having used a number of different mounting media my choice is still Euparal (Diaphane is nice too). Properly prepared specimens mounted in Euparal do not crystalize, fog, shrink, crack or do anything but last in good condition for a very very very long time. Slides I made using other media have done all of the previously listed things, and none of them have lasted as long as the Euparal slides. If I am going to go through all the trouble to mount something I usually want it to last.

Contributor number 11

Standard resin-based media are pretty much useless for detailed observation of mites because of refractive properties.

Contributor number 13

I worked for several years as a technician mounting shed skins of mosquito larvae and pupae. We experimented widely with mounting media and decided that Euparol had the best optical qualities. My experience with it was that it was quite pleassant to work with in terms of odor and ease of clearing. It can be dissolved in alcohol, if I remember correctly. On several occasions we had to remount specimens and while a nuisance, it is relatively simple to remove the cover slip and Euparal by dissolving in alcohol. It does not seem to crack the way old balsam-mounted slides do. As far as I know, most of the mosquito people use it now.

Contributor number 18.

Euparal is the classic example of a medium miscible in alcohol, and on which most variations are built. These are usually Gum Sandarac based media. Gum mastic and venice terpentine based media have also been used.

The advantages of these media are that specimens can be transfered directly from alcohol. The amount of dehydration (and thus the strength of alcohol the specimen must be processed through before mounting) is dependent on individual mounting media. More permanent than water based media, but not really good over the long haul (>20 years). Some media (e.g. euparal vert) also incorporate staining action with the clearing action of the media.

Their disadvantages are that these mountants still go bad over time. Type specimens should never be prepared in an alcohol miscible medium. They are really an intermediate solution for specimens that are too delicate to withstand the processing involved for permanent mountants. Their optical quality is good, in some cases excellent.

Ease of preparation is very simple and ease of remounting is good, but they are not viable for long term storage of types or vouchers. My euparal slides of chigger mites were made in 1987, and they're starting to polymerize and craze around the edges. They might, I say might, make it to the next millenium before they're toast.

Contributor number 24.

Euparal (refractive index 1.483). This medium was not used at all on mites. This was probably because of its labor intensive procedures and the delicateness of mites. Constant handling of the specimens would render mites to lose setae, fragmentation may occur. Preparation is difficult, remounting is unnecessary. Sclerotized large size Mesostigmata, Oribatida and Ixodida could be mounted in Euparal, I suppose. I prefer this medium for sciarid mounts but I have not ried it on mites.

Contributor number 26.

I've mounted a couple of thousand mosquito larvae pupae and exuviae in euparal, pretty much at my boss's insistance. It seems to be a good medium, not affected by atmospheric water once dry, like Hoyer's is. Euparal dries clear too. Be careful that the company doesn't send you Euparal vert, which is a green colored medium that some histologists use.

Contributor number 28.

An advantage of Euparal is the ability to stain a specimen before mounting, for example with carmine borax or clorazol black. The disadvantage is that specimens must be dehydrated prior to mounting, and this takes a lot of time.


Contributor number 1.

After more than 20 years in entomology I have found that the best medium for slide mounting is still the "good ol' " canada balsam, though the processes to go through sometimes appear longer and more difficult than some contemporary ones (euparal for example). In our Museum, all Coccids, aphids, and black-flies are prepared with canada balsam. Some of our slides of coccids are now more than 70 years old. Some slides of Pselaphidae are also more than 70 years old and contain types of new species. In these old slides the canada balsam has become brownish but this is not a real discomfort. The genitalia of my clerids are associated with the specimens in a small cell filed with canada balsam. If necessary a drop of xylol (or benzene) is added and the piece comes loose in minutes and can be observed again under any angle with a stereomicroscope. Some will object that for these preparations you have to cope with chemicals that are not really good for health. True, but when precautions are taken it is not a real danger, and except for people that prepare specimens all day long with these chemicals (for whom very special precautions must be taken), it cannot be worse than what we breathe in large towns at rush hours!

Contributor number 3

Histological mountants such as Canada balsam require excessive dehydration steps through alcohol and xylene to prepare the specimen for mounting.

Contributor number 4

Canada Balsam becomes reddish and the chitin inside becomes transparent with time, you spend too a long time doing dehydration through alcohols.

Contributor number 8

Many of our specimens remain in excellent condition after many years in Canada balsam; however, if the slides have been in contact with PDB (e.g. genitalic mounts in with pinned specimens), they turn completely black and opaque! We face a very time intensive re-preparation project. Beware!

Contributor number 16.

Canada balsam is crummy. Fairly easy to use, no clearing agent, dark to begin with and darkens more with age. Tough to re-mount, but mounts are permanent (if you can handle the darkening). Mounting dry specimens is OK (e.g. live), but I've had more trouble with wet specimens. I've never been able to soak off a cover slip for re-mounting. Balsam is permanent, but darkens with age, which I find annoying, especially if your phase-scope light source is not terribly powerful.

Contributor number 18.

Canada Balsam is an example of a mounting medium that is not miscible with water or alcohol. The advantages of these media are that they are permanent. That is, if specimens are properly prepared, they should, in theory, last forever. I have never personally viewed a resin mount older than ca. 120 years; but these specimens showed no appreciable sign of age. De rigueur for type specimens. (I have known systematists who have been asked to remount poorly mounted types in canada balsam as a condition for borrowing the types from a museum.)

The disadvantages of these media are that they are slightly more expensive. They are also more time consuming because a full dehydration and clearing/dealchoholization protocol is required. Sometimes (in the United States) it is difficult to find a GOOD media vendor with a high quality, reliable product (although they are out there). These media take a little more care and experience. The other advantage to things like canada balsam is that it really does make the lab smell wonderful while your slides are drying. Their optical quality is excellent, but beware, some stains are very sensitive to the acidity of the medium. One can usually take care of this by keeping some marble chips in the canada balsam bottle. Beware of "neutral" canada balsams: they usually aren't. Acid media tend to make hematoxylins fade over time. Most stain problems have been worked out though, and most of the common staining protocols are optimized for canada balsam or gum damar.

Their preparation is more difficult than other media, but only because it takes more time to prepare the specimen for mounting. The process itself is rather simple, but there are slight variations depending on the nature of the specimen. For example, protozoans can go from life to a permanent mount in about 8 hours. Leeches take a couple of days. Some tapeworms and flukes might take up to a week or so. (Not constant attention, of course. Just changing fluids on a regular schedule.). Their ease of remount is excellent.

I think they are the only option for types or vouchers (long term). I have never had a canada balsam slide go bad. In some cases, the mountant has yellowed over time. That is usually an indication that the resin was over-heated at some time, either by me, or more likely during the refinement process. Find a good dependable, reputable supplier and stick with it. I've only seen two gum damar slides that were older than 20 years, and both looked fine (they were types). Damar doesn't yellow, but it can be a little tricky to get it ready for use as a mountant.

Contributor number 22.

I only use balsam - except for rapid determinations. But it is not possible to make good mounts from older material. My best slides are from specimens I have collected and stretched within a few hours; they can then wait their turn in 60 per cent alcohol. But that early stretching and piercing of the membranes is essential for good quality slides (of aphids). Balsam is easy to handle, once you accept the discipline.

Contributor number 24.

During the 30s and 40s, Canada Balsam (refractive index 1.535) was used to mount oribatid mites. Mounts I've seen here were thick, not well prepared, and mites were not cleared. It seemed like they were collected and went directly to the mounting medium. Some darkening around coverslips and refraction was unsatisfactory. Preparation is intensive. It will be interesting to know how optimally cleared mites on thin mounts will appear under phase-contrast microscope examination.

Contributor number 26.

I mounted a couple of thousand Ceratopoginidae in blasam or phenol-balsam (Wirth & Marston recipes), and I happen to like balsam. It is permanent, and I would say that it's a responsible choice for mounting type material. It can be a little tricky optically, especially if color of specimen is important, since balsam has that nice amber yellow color.

Contributor number 27

Specimens mounted in balsam may in my experience be recovered by soaking in alcohol or xylene, and may then be remounted with good results.

Contributor number 28.

An advantage of Canada balsam is the ability to stain a specimen before mounting, for example with carmine borax or clorazol black. The disadvantage is that specimens must be dehydrated prior to mounting, and this takes a lot of time.

Contributor number 3

Histological mountants such as Permount require excessive dehydration steps through alcohol and xylene to prepare the specimen for mounting -- and the current formulation of Permount is a disgrace! (I have attended workshops on how to rescue specimens from Permount and re-mount them in Canada balsam!).

Contributor number 18.

Many of the synthetic permanent media (eg PERMOUNT) are not in fact, permanent. Permount has gone through several formulations because the earlier formulas tended to polymerize and craze over time (> 5 years).

Contributor number 20.

Do not use PERMOUNT! I used it 20 years ago for preparations of lepidopteran genitalia, and in the intervening years, the medium crystalized. The slides cannot be used. I saw the same result from a large collection of lepidopteran genitalia recently donated to the USNM. Fortunately, the PERMOUNT is soluble in xylene and similar chemicals. I spend a few hours each week remounting all my previous slides. Fortunately, the specimens seem unharmed by the crystalization, but it is a nuisance to do all the work twice.

Contributor number 25.

I'm very interested in mounting media, especially after my disastrous results with Permount.

Contributor number 26.

Permount is okay, although I've only used it for histological slides and a couple of times used it for insect larvae when I'd run out of anything else and I needed to get stuff mounted quickly.


Contributor number 3

The "standard" entomological media (Hoyer's, DeFaure's) seem to be designed as narcotizing agents, with little thought as to their effect as long-term preservatives.

Contributor number 4

Hoyer's mounting problems are well known.
Contributor number 7

I do like using fast water soluable mountants for disposable slides (maximum life a month because they shrink and squash the specimens. To do quick id checks on mites they work fine. You move the specimen from ethanol directly into the mountant and pop the cover slip on it. Just don't expect them to last.

Contributor number 10

I have used Hoyer's mounting medium for several hundred slides of meiotic chromosome squashes for miscellaneous plants. My work was all done in Phoenix, Arizona, an exceedingly arid environment for 10 months of the year. After a few months the Hoyer's would begin to dry from the edges of the coverslip and eventually the entire preparation was destroyed by the crystallization. We found that we could carefully wipe away excess Hoyer's that oozed from the coverslip edge during squashing (we used a moist towelette) and then, when the glass dried, seal the edge with a clear lacquer (we used fingernail polish). This extended the life of most of the slides for several years. In some the seal was likely flawed and crystallization occurred anyhow. I did not find the medium to be satisfactory for long term storage but used it because it was easy to use with aqueous material.

Contributor number 11

Gum-arabic media remain the best for observation, at least of those I have seen/tried.

Contributor number 16.

Gum/choral hydrate media are good at clearing (macerating) mite specimens, but can't clear heavy sclerotization. They are easy to use, easy to make fine mounts, mounts can be soaked open later for remounting. Questionable for long-term storage. The optical quality of Hoyer's is excellent, unless contaminated by non-miscible liquid. Mounting in Hoyer's is easy. Specimens can be mounted live, removed from water, alcohol, or other polar solvents and dumped directly into the Hoyer's, or taken from lactophenol, rinsed in water and mounted. Hoyers takes time to soak off the coverslip for re-mounting, but if you're patient it comes right off. Then the trouble is finding the specimen, especially if it's small and well cleared!

Despite a bad reputation, I've seen specimens mounted in Hoyer's which are 40+ years old and look perfect (of course I've also seen fairly bad ones...). Crystalization and bubbling seem to be the worst offenders. If the coverslip is completely filled with good-quality Hoyer's though, and is completely sealed with 2 coats of Glyptol, it should last indefinitely. Use a round coverslip and turntable for sealing. If it does deteriorate, it's simple to soak it off and re-mount.

Contributor number 17.

Hoyer's solution is great for mites, but the glycerine collapses the cells of certain species of mosses. It works fine for delicate tissues but needs luting. Nothing really works all the time when sealing slides.

Contributor number 18.

The ORIGINAL Hoyer's medium was a gum mastic based media. Most of the commercially available Hoyer's media are really Hoyer's variation of BERLESE's media.

Hoyer's medium, Doetschman's medium, Berlese's medium, etc. are primarily based on chloral hydrate (clearing and preservative), glycerol, water, and gum arabic. They tend to over clear and dry out ("craze" or powder) over time. They are great for a project with lots of specimens IF you don't need them to last, don't want to deposit them in a museum, and don't have time for a more permanent preparation. Often difficult in the United States because chloral hydrate is a controlled substance. They can be bought commercially, but hard to justify the price, given the temporary nature of the mount.

The advantage of these media is that specimens can be transferred to them directly without dehydration or clearing. Specimens are usually taken directly from water or glycerin. These types of mounts are quick and easy to make, and they usually have some type of clearing action on their own. By clearing, I mean the process of rendering the specimen transparent (increasing the specimen's refractive index).

Their disadvantage is that over time the clearing action of the media produces a specimen with a refractive index = to that of the mounting media (i.e., your specimen disappears). Water-based media tend to dry out rather quickly (>1 year). Your specimen is toast, and you can't reclaim it for remounting. This process can be delayed by ringing the slip with a sealant (venician terpentine, gold size, etc.), but ringing only slows the process--it doesn't solve the problem. These media are not suitable for long term storage of types or vouchers. I have never had a slide last longer than 2 or 3 years. If you make a lot of slides, it isn't worth the effort to go back and do the required biennial curation. Their optical quality is good initially, all downhill over time. Preparation is very simple, as long as you can drop a coverslip without creating bubbles. Ease of remounting is good.

Contributor number 22.

I have mounted thrips for 30 years or more. I started with hoyers and Berlese mountants because of the higher refractive index. However, I know of a collection of many thousand slides that has been undergoing a long and expensive process of remounting, since the slides all dried out and the specimens were often pulled to pieces in the process.

I used Berlese's medium for a year with patchy results. At best, it was wonderful; but with a 90 per cent error rate. One publication I produced points out that a complex mathematical analysis of variation in one species group led to the computer identifying two 'species' - regardless of where the material had been collected. Reinspection showed this to be 'specimens in balsam' vs 'specimens in Berlese'.

Berlese's medium causes various strange distortions in antennal segments and head shape. Two overseas colleagues use it - you can tell from their illustrations! All aphid material in Berlese's, 200,000 slides or so, is having to be remounted because the berlese is going black.

Contributor number 23.

We still use the methods using Faure's medium as modified by Massoud in 1968 (for Collembola). Unfortunately the specimens deteriorate with age and often after ten or so years are virtually useless for Collembola. The specimens can be remounted but this often results in only slight improvement. I have never found a good, simple, easliy learned method for Collembola. Mites pose much less of a problem.

Contributor number 24.

My mite collection is 98% mounted in gum arabic/chloral/glycerine medium (modified Hoyer's). It is so far the best in ease of preparation and practicality. Mites could be directly mounted in Hoyer's from 75% ethyl alcohol, clearing solution (Nesbitt's) or water. Optical quality is excellent (refractive indes 1.47). Remounting is fast and easy. The main disadvantage of this medium however is that under tropical conditions, the medium softens after a few hours at room temperature; it is also not permanent. Our present collection shows varying degrees of deterioration, from discoloration to air incursions from without and within coverslips resembling serpentine leaf mines. When leaf mines are extensive, moat-like condition around specimens exists.

Contributor number 26.

I've used several media for mounting specimens on slides. Probably the worst to use is Hoyer's. It is prety much a question of when, not of if, your colleciton will deteriorate. Ringing slides very well with red glyptal will keep them longer, but in the end, it's curtains. Lots of acarologists like to use Hoyer's because it is easy to use, can be thinned readily, and has a clearing action such that even uncleared mites, if very small, can be mounted, and in a little while, they're clear. I've made Hoyer's once or twice and it's no picnic. If you have impure gum arabic, or granular as opposed to flaked gum arabic, you're up a proverbial creek. Hoyer's dries almost perfectly transparent and it's very easy to see morphological characters.

Contributor number 27

For Collembola, Hoyer's is an excellent medium in the short term, because of the contrast between its refractive index and that of cuticle; it is far superior to Balsam in this respect. I doubt that the advantage is as great for mites. In the long term Hoyer's is unsatisfactory, and unpredictably so; some 40 year old preparations are still good, but many younger ones are no longer usable. I suspect variation in the quality of the medium is responsible, at least in part. The commonest fault is intrusion of air under the coverslip; I am not sure that ringing can always be counted on to prevent this. Specimens are easily remounted by soaking the slide in water.

Contributor number 28.

The preparation of Collembola for mounting in Marc Andre's Hoyers medium takes little time. Specimens preserved in alcohol or formalin can be used. The disadvantage of this medium is the appearance of crystals that destroy the specimen, and the need to seal the slide.

Contributor number 29.

HOYER'S - I have followed the procedure as outlined in "Baker, E.W. and Wharton, G.W. 1952. An Introduction to Acarology." All my mite specimens are mounted in this medium and rung with Gurr's glyceel. I have found this method to be very satisfactory, although some of my earliest slides (1960) have dried out with air penetrating the medium. However, it is easy to remount specimens -- which I have done on a few occasions -- since Hoyer's is water soluble.


Contributor number 7

I did like using low concentrations of formalin as a mounting medium (for nematodes and worm like things) when I could get my hands on nail-polish that actually would seal the coverslips. This is not the case anymore, there is still some type of plumbers sealant (red smelly stuff) that works ok.


Contributor number 7

I now prefer to use glycerol - it doesn't evaporate as readily and clears the specimens nicely.


Contributor number 3

I have been having some favourable short-term results (ca. 4 yrs.) with methyl cellulose (though you run the risk of mold contamination) and the CMCP series of mountants (polyvinyl lactophenol based, lost when Turtox Biological became extinct, but recently resurrected in an entomological monograph.


Contributor number 2

I have mounted many many wings, wing bases and wing articulations of beetles, and have experimented with all sorts of media. I found that lines of fusion, nifty little details etc. were totally obscured if any medium was used.

My recipe is quite simple. 1. Dissect parts. 2. Stick press-stick (it's rubber stuff used to stick up posters - not permanent) onto slide. 3. Stick parts onto press-stick.

Advantages - you can see all of the features, move the specimen around to see all angles etc. and it doesn't take forever to remove the parts for SEM'ing or whatever. Disadvantages - if carelessly used the press-stick may rip extremely thin parts like the apical regions of wings.

Kukalova-Peck and Lawrence simply squirt relaxed beetle wings with alcohol, spread onto a slide and let dry. The wing stays in an outstretched position quite well. I personally have had problems with very large wings, but some press-stick (tiny bits) under the wing apex and wing base stretches out the wing beautifully.

Quite frankly I don't know why anyone uses mounting media - nasty smelling, messy, a pain to reverse and it obscures all the nice details.


Contributor number 3

I have received some complaints about excessive clearing of the specimens in CMCP but, being water-based, CMCP allows for simple stains to be added to emphasize the structures (I personally prefer "CMC-S" -- saffranin added to the mountant to fix, stain and mount in one step!).

Contributor number 11

Master's Chemical Co. (Des Palines, Illinois) markets media called CMC and CMCP mountants. They don't require dehydration, yet do not seem subject to the deterioration problems of gum-arabic media, and don't seem to need ringing. The resolution is good, but perhaps not quite as good as the latter. I haven't tried the whole range of viscosities and refractivities available, but it's promising. I know nothing about the ability to remount from this. Details of the use of these media can be found in Beckett and Lewis 1982, Transactions of the American Microscopical Society, 101 : 96-99.

Contributor number 24.

CMC, CMCP compounds (refractive index 1.38 to 1.41). Relatively easy to prepare. Optical quality is good. Shrivelling of specimens was observed a few days after mounting and minute debris attached to legs do not separate during preparation compared to Hoyer's. Specimens mounted on CMC, CMCP products could be remounted since the medium is water miscible. A permanent mount. A satisfactory medium for Diptera mounts.


Contributor number 4

I would strongly recommend all of you to use as a mounting medium (microscopic slide or drop for larger pieces after dissection) dimethyl hydantoin formaldehyde resin (DMHF or DMHFR). DMHF is a syntehtic resin, you don't need to dehydrate, it has the same refraction index as glass and never reddens.
Contributor number 6

I have used DMHF and it's wonderful. It even allows you to carry on a dissection of the parts included some years later (a small amount of water is enough).

These are two useful references on DMHF, the first of them includes many useful others.

Bameul, F. 1990. Le DMHF: un excellent milieu de montage en entomologie. L'Entomologiste, 46(5): 233-239.

Steedman, H. F. 1958. Dimethyl Hydantoin Formaldehyde: a new water-soluble resin for use as a mounting medium. Q. Jl. microsc. Sci., 99(4): 451-452.


Contributor number 5

Taxonomic study of copepods is done on mounted animals, on slides containing the whole animal or dissected parts. We are interested in observe minute setae, pores, sensillae, hyaline frills, etc. Animals are cleared and decolorised in lactic acid for about 20 minutes. They are then transferred to a solution of 30% lactic acid and 70% glicerin. If a dense solution is needed (sometimes a dense solution helps in putting animals into the correct position) you can add glicerin-gelatine to this solution. The optical qualities are very good, after inmersion in lactic acid the specimens are completely transparent. Transparent objects are observed in a phase- contrast microscope, or interference contrast. The slides are sealed with paraffin. You can open the slides easily every time for re-mounting. For long-term preservation we seal the outer margins of the paraffin slides additionaly with Eukitt. (long-term means more than 10 years) We have 25 year old slides, they are ok !


Contributor number 12

Many mycologists have used mountants based on polyvinyl alcohol such as polyvinyl-lacto-glycerol (PVLG). PVLG is a colorless mountant that becomes permanent after drying (usually at 60 C for a few days). Specimens last years without substantial degradation (although some fading of color is evident). I have used this mountant with a wide variety of fungal and plant structures with excellent results. I see no reason why it can't be used for insects.

Because PVLG dries slowly, specimens can be moved around for proper positioning. However, after drying the slides are quite permanent and resistant to all sorts of abuse. PVLG can be mixed with various stains (I use Malzer's reagent - an iodine based stain) with excellent results. Thus staining and mounting can be achieved in one action. On the downside, PVLG is quite acidic (about pH 5.0 I think) and some structures tend to distort in acidic media.

Recipe for PVLG:

Distilled H2O   100 ml
Lactic acid   100 ml
Glycerol   10 ml
Polyvinyl alcohol (PVA) 16.6 g

The PVA should be 50-75% hydrolyside and have a viscosity of 20-25 centipoise in a 4% aqueous solution at 20 C. The PVA takes several hours to dissolve in a hot water bath. The mountant can be stored for 1 to 2 years (use a dark bottle).

Contributor number 23.

The P.V.L.P. mounts we used in the 1950s (for Collembola) proved diasterous. They shrunk greatly after about 5 years.

Contributor number 24.

Polyviol 17. This compound is a mixture of high grade polyvinyl alcohols, ethanol, distilled water and lactic acid. Being used by aphid workers in Sweden, specimens mounted 20 years ago looked like newly prepared. The disadvantage is that the appendages e.g., legs, palpi shrivel after a few weeks. Idiosoma usually is not affected by shrivelling or twisting. Like CMC compounds, the dirt and debris do not come off the integument during preparation. I think that fine tuning the clearing process will eliminate the shriveling effect. More tests are needed for this medium.

Contributor number 28.

The advantage of polyvinyl lactophenol is the ability to make preparations from fresh specimens, or those preserved in alcohol or formalin. Also, the medium gradually clears the specimen. The specimen can be removed by submerging it in hot water. Preparations take little time to prepare and do not require sealing.

Contributor number 29.

PVA - The procedure I have used is from "Salmon, J.T. 1954. A new polyvinyl alcohol mounting medium. The Microscope 10(3): 66-67." My specimens were distorted and not cleared enough for proper identification. Today, the mounts seem to be in the same condition as when they were made! I stopped using PVA in 1973.


Contributor number 17.

Details of the use of Lactophenol Gel in botany can be found in Taxon 32: 618-620, 1983.

Contributor number 19.

We phycologists use Karo (registered trade mark) corn syrup to make permanent mounts of algae. Woolies sells it and so does Coles. The leftovers are good for Pecan pies.


SWANN'S - I adopted the method as described by "Rusek, J. 1974. Die Praparation von Kleininsekten. Mikrokosmos, Heft 1 Januar 1974: 10-12." Since 1974, I have been using this procedure with slides sealed with Canada balsam. I have had good success with this method. Specimens are well cleared and characters need for identification are observable. All my specimens in Swann's are still in good condition. Like Hoyer's, deteriorated specimens in Swann's could be remounted easily. Since Canada balsam dissolves in xylene, care must be exercised when cleaning slides with this solvent.


Contributor number 1.

I have some experience with "easy" media using glucose and other chemicals that I do not remember. The process was quite easy and quick, but meant that every year you had to "feed" the slides with a tiny drop of distilled water. Even taking care, more than half of my slides are crystallised now and the specimens (mouth parts of larval diptera) are lost (fortunately there were no types in these).

Contributor number 21.

I dabble in slides occasionally and have used all the commercially available stuff because most of the recipes, for Hoyer's for instance are time-consuming and seem to leave out just enough information for me to make a big mess of it. I have used all the products that Bioquip sells: Euparol, UVA, Permount etc. Most of them seem to work ok but I'm not looking at long-term archival type use, just mounts for mite ID in class, etc. I suspect most of what's out there works for me when fresh but I've had trouble with shelf-life so my criteria is quite different from yours.

Contributor number 23.

Over the 45 years I have been working with Collembola I have tried about every mounting medium including Karo Corn Syrup. I have a problem which is unusual in that I have to use undergraduate students for most of the mounting and often have to mount very large samples. This means that some of the methods suggested by others which are difficult to learn or require great preparation are really not usable. With one exception none of the mounting methods I have used is entirely satisfactory. The only method which produces long term survival in excellent condition is using the technique developed by R.F. Wilkey P.O. Box 185 Bluffton IN 46714. Unfortunately we have found this method very difficult to use and also it produces only cleared specimens. The mounts are in Balsam or Euparol.


Contributor number 7

Problems that I have run into in making slides include:

(1) Having the right tools to move the specimens without breaking or losing it.
 Probes/scalpels made for eye surgery are often small enough to use.
 Paint brushes with only 2-3 hairs can be used to pick up small objects.
 Make your own probes by sticking an eyelash on to a thin wooden stick or use very thin wire to make a loop and attach as above.
 Inverted microscopes with low power objectives (2X is great). You can use trays etc on them with low power to see what you are doing and track the object you are moving.
 These neat slides that have removable multi-chamber plastic sides that make them into mini-trays. They were made to be used with blood, but work great for lots of microscopic work. Fisher and Carolina Biological carry them.
(2) Moving small objects from dish to dish to slide.
 Some just vanish, using the above equipment helps a lot.
 Use a mounting medium that doesn't have the same refractivew index as the object you want to mount, or they really do disapear. I had this happen once with mollusc radula (teeth), I thought I was going crazy. I finally found the radula under phase contrast. I then added some Eosin to the mounting media to stain the background and found all the rest of my lost specimens. If the specimens are in formalin or ethanol you will need to move them through a number of fluids till they are ready for mountant. The multi-chambered slides are great for this purpose. We were even able to used them with glycerin and 90% ethanol on slide warming trays to clear the mites before making slides.
(3) Specimens always end up in the wrong place on the slide. I mark each slide with a small circle one side using a lab pen. Then the specimen can be placed in the middle of the circle on the opposite side of the slide (all pens run in one or the other of the chemicals we use so use the opposite side of the slide). This makes finding the specimens easier and it also means you don't have to move the microscope adjustments as much between each slide. Also if you send your slides off to an expert for help, you will not get much help if your specimens are not centered!
(4) Specimens end up with the wrong end up. For really small mites this is a big problem, you can't see through them and you can really mess them up tring to remove the coverslip and turn them over. I started mounting some small Prostigmata between two coverslips and using removable tape to anchor the coverslips to a microscope slide. Wrong side up, just turn the mounted specimen and coverslips over. It is not as good as making a good slide to begin with, but when you are doing lots and lots of slides it saves time and specimens.
Contributor number 11

I have a little trick for making useable whole mounts of thick specimens. I use a #8 tungsten-carbide dental burr (any dental supply house has them) to make a small pit in a standard glass slide. A Dremel-tool with a matching micro-drill press works fine, or I sometimes use a full size drill press, to the same effect. Various depths can be cut, and the bottom can be left rough or it can be polished with various easily-obtained "rouges". Leaving it rough is fine, since the bottom of the pit is out of the range of the higher objectives anyway. I often mount two specimens in a pit, one dorsal, one ventral. In such slides, even if the medium deteriorates or shrinks (which is less a problem since only the pit is deep) the specimen survives and can be remounted. It's a great technique for teaching collections, and for a quick-access collection. I keep most of my material in alcohol, but have 1 or 2 slides of this type. I can see it being a good way to keep parts of paratype series for general loan.


Contributor number 8

I would like to encourage you to include information on the best ways to produce and afix slide labels, including paper types, lick'em stick'em vs. peel and stick, Elmers glue vs. glue stick, wet the slide & the label vs. just wetting the label, available software to produce 1" x 1" labels. I have some information on producing labels using Word Perfect and an HP Laserjet.

Archived 30 March 2001 by Acarology List-owner