The first rule for studying protists is to observe them alive, if at all possible, and there are many organisms which will challenge your ingenuity.  Some ciliates are so fast, such as the aptly named Urocentrum turbo, that one has to find ways to slow them down to observe any sort of morphological detail and, that may ultimately mean killing, fixing, and staining the organisms.  Other ciliates and flagellates are so small that examining their structure requires studying them near the limits of light microscopy under oil immersion at a magnification of 1000.  Even an extremely small movement by the organism can transport it out of your field of view and trying to track it down again at 1000x is usually hopeless.  There are also some parasitic protists that require killing, fixing, and staining in order to differentiate the organism from the surrounding tissue.

 However, to begin with, let’s assume we have a nice cooperative Amoeba proteus, 500 microns in size, active, and willing to show us both structure and behavior.   And, at this point, the first three rules are: observe, observe, observe.  Next, take notes!  Often you’ll think to yourself: “Oh, no problem, I’ll remember that.” Ten years later, you’ll wish you’d taken notes.  Next, make drawings!  They don’t have to be professional; they need to be your own personal archive that can trigger your memory, so that you can go back years later and say “Oh, yes, that’s Amoeba proteus and there’s that peculiar macronucleus.”  Take lots of notes; make lots of drawings.  They may seem wrong or simplistic or exaggerated to you later, but they’ll keep getting better.  I flunked stick-figure drawing in the third grade, but I still keep making sketches, because they preserve for me important information, even if no one else can decipher it.

 The next step, if you’re fortunate enough to have a camera system for your microscope, is to take some photomicrographs.  I’m a total ignoramus when it comes to using film cameras and, so if you want to know about those mysteries, you’ll have to get in touch with Wim van Egmond, Jan Parmentier, Dave Walker, Roland Mortimer, or Maurice Smith (to mention just a few of the fine photographers who contribute to Micscape) and try to get them to reveal their secrets. I still haven’t figured out how to take off the lens cap.  However, when I retire next year, I’ll learn how to do it.  In the meantime, I’m lazy and I’m fortunate to have a video camera for my microscope system and, so when I encounter an unknown organism or a bit of odd behavior in a known organism, I try to capture it on video tape.  I realize that the resolution is not as good, but the fact that I can sometimes capture interesting and bizarre behavior seems to be a fair trade off.  However, as  wonderful as photos and videos are, they are no substitute for drawings.  The hand-eye coordination which you are forced to develop means that you observe differently and, you greatly enhance the likelihood of seeing detail, which you might well have missed had you not made the drawing.  No matter how much equipment you have, it’s useless unless you also employ the very highest technology—your own brain.

 Earlier I mentioned the problem of rapid locomotion and what a problem that can be.  Early microscopists tried a number of techniques to slow protists down and achieved varying degrees of success.  Bits of cotton fiber scattered though a drop of water was a popular mechanical method and, if you could get a good distribution, it could be effective for Paramecium-sized organisms, but for small fast ciliates and small flagellates—forget it.  The basic idea is to create miniature cages, which ideally will also have a lot of bacteria to attract the feeding Paramecia; however, even in these tiny enclosures Paramecia can dart back and forth quickly enough to make it extremely difficult to observe much detail.  Another early technique was to use some kind of non-toxic thickener which could be added to the water to increase the viscosity, thus slowing down the organisms.  A popular such substance in the 19th Century was quince mucilage—I’m not sure I even know what a quince looks like.  However, today the same basic idea is still used. The most popular substance is methyl cellulose or some commercial variant thereof.  The commercial versions work well for some organisms, but for others, they are rather too thin.  I prefer to make my own solutions from the “powder” (actually it’s more like finely shredded cotton) with differing amounts of water, so that I can have solutions of different viscosity. [Note: I use artesian water to make up these solutions, since distilled water contains no mineral salts and can be toxic to some micro-organisms.  One can buy a gallon of artesian or spring water for less than a dollar at a local supermarket.] Clearly, if the solution is too thin, it will be useless in slowing the organisms down; if it is too thick, the organisms may be distorted as they attempt to move through this medium and, in some cases, it may even cause them to lyse (break apart).  I keep 3 or 4 solutions of differing viscosities and this allows me to cover a wide range of organisms.  Making one’s own solutions does require a bit of patience as methyl cellulose is rather slow to dissolve and if you stir or shake it very much, myriads of small bubbles will form and you will need to wait (perhaps several days) for these to either dissolve or rise to the top.  One disadvantage of methyl cellulose is that, since it consists of lots and lots of cellulose fibers, not all of which dissolve, it can create problems if you are doing polarizing or Nomarski differential interference microscopy, since such fibers are birefringent.  On the whole, however, methyl cellulose is a valuable tool and has the very large advantage of not being toxic to the organisms.

 Another valuable technique for enhancing contrast for studying micro-organisms is the use of vital stains.  Since I have discussed this in a previous brief article, I won’t repeat that discussion here. A significant advantage of these techniques is that the stains are relatively non-toxic to the organisms, allowing one to study various aspects of their morphology while they are still alive.

 A wide range of chemicals has been used by microscopists to “anesthetize” micro-organisms.  Some of these compounds were used with a fairly good understanding of the possible biochemical reactions and effects; but, in many instances, with early microscopists and more recent amateurs, there was a large element of trial and error.  Curiosity is a fascinating, but sometime dangerous, even deadly, aspect of the human psyche.  There are some individuals who just can’t seem to resist the temptation: “Gee, I wonder what would happen if I mix this compound with that compound?”  Well, maybe nothing drastic, but, on the other hand, you might get an explosion or produce a deadly gas or ignite your curtains—so, don’t mix things randomly; always be aware of the potential safety hazards.  Some of the older books on microtechnique don’t contain specific warnings about dangerous substances or reactions, either because the seriousness of the hazards was not known at the time or because the author, rather naively, assumed that the person using the procedures would already be aware of any potential dangers or would be under the supervision of someone who would.

 The major protists that swim rapidly enough to be a problem are a number of ciliates and certain flagellates.  Two chemicals that can be useful are either nickel sulfate in a 0.5% to 1.5% solution (experiment!), or a 0.5% to 1.5% solution of potassium iodide.  These substances affect the mechanisms that are involved in ciliary action and, in the right concentration for a particular organism (trial and error), can be quite effective.

 Occasionally magnesium sulfate (Epsom salts) is effective with ciliates and here is a chance to experiment extensively, since different organisms vary widely in their tolerance to this substance.  For example, I have found that Lacrymaria olor has a very high tolerance for this salt.  Some organisms—Stentor, Spirostomum, Vorticella, to mention just a few—are highly contractile and this makes them especially difficult to study.  In professional scientific circles, a substance is regarded as a true anesthetic only if its effects are completely reversible.  The substances which may accomplish this feat with protists are virtually unobtainable by the amateur and consist almost exclusively of legally restricted and expensive compounds.  However, for those of you who are determined to experiment, certain over the counter medications or leftover prescription drugs which you have sitting around, just might provide some interesting results.  Take notes and let us all know if you succeed.  For example, neosynephrine has been successfully used as an anesthetic for certain micro-invertebrates.  A 10% solution of grain alcohol (dilute down a bit of vodka) has proven effective with a variety of micro-critters.  Ingenuity and lots of patience will likely yield results—not likely a Nobel Prize—but who cares, we do all this stuff because we love it, not for the fame or fortune. (Please send all contributions to the Howey Microscope Fund).

 Some of the organisms we’re interested in, can only be effectively examined when they have been properly killed, fixed and/or stained.  Sometimes this can be accomplished with a single reagent, but, in general, more techniques are  required to obtain satisfactory results.  Interestingly, there are a few relatively simple and straightforward techniques that sometimes produce very good results with some organisms. Let me mention two such techniques:

 1) Make up a solution which consists of ½ undiluted commercial formaldehyde [CAUTION: Very nasty stuff! Carcinogenic.] and ½ absolute ethyl alcohol [CAUTION: Removes water from tissue almost immediately.  If ingested, can be fatal!] Absolute ethyl alcohol is difficult to obtain and in many countries requires a government permit, but often one can buy 95% ethyl alcohol in a liquor store or you can try substituting 91% or 99% (when you can find it) isopropyl alcohol or 95 methyl (wood) alcohol (methanol).  All of these above substances are potentially dangerous and must be handled with care.  Even skin contact and fumes are toxic, so proper precautions must be taken.

 So, with all this toxic stuff in a mixture, it’s no surprise that it’s going to kill micro-beasties and, in some instances, quickly enough to fix them with relatively little distortion.  Using this mixture , one can sometimes achieve very good fixation of flagellates and ciliates, and the precise reasons for this are not well understood.

 2) This method involves less dangerous, but still somewhat toxic substances.  Copper salts have long be used in very dilute solutions, to try to control algae blooms in ponds and lakes, especially those that serve as reservoir supplies of drinking water.  These sudden population explosions of organisms, such as the flagellate Synura, can significantly affect the taste and odor of the water.

 I have gotten the best results so far, using a 1%-2% solution of copper acetate.  This will kill a variety of organisms with minimal distortion, but must be followed up with a fixative, such as formaldehyde (5%-10%) within a minute or two.  There are some combinations of copper salts and fixative agents which are said to produce a very delicate fixation, but I have gotten the most desirable results by using the copper salt first and then following it almost immediately with a fixative.  You can try using the copper acetate and the formaldehyde together, and the formaldehyde will coagulate the proteins (which is the purpose of fixation), but it also contains methyl alcohol which increases the possibility for distortion.

 Another very simple technique which sometimes works very well for rotifers, occasionally works well for protists too—boiling water.  This then needs to be followed immediately by a fixative.  To use this method, one does have to have a fairly abundant supply of the organisms.  Put the organisms into a small amount of water in a heat-resistant dish and allow them time to accommodate and extend if they are contractile, then quickly flood the dish with boiling water.  With luck, you will obtain some nicely extended specimens.

 Protists have so many intriguing little tricks that they are a constant challenge to our inventiveness.  Think for a moment about the problem of surfaces.  We tend to think of amoebae as protoplasm-filled bags, rather like a balloon half full of water and with a smooth, constantly changing surface.  However, some amoebae, such as Difflugia, secrete a sticky sort of mucous, thus building a protective housing around themselves, by cementing sand grains and/or other bits of detritus around themselves.  Certain species seem quite meticulous and select sand grains that are remarkably uniform in shape and size, whereas other species are quite content with gathering sand grains of various shapes and sizes along with the occasional diatom.  Euglypha, and some other related amoebae, secrete silica (glass) plates or scales to form lovely protective housings.  Arcella and its relatives secrete a chitinous hemisphere, somewhat shaped like a low-form dome, and other species have spines and projections.  Clathrulina elegans builds a hollow sphere on a stalk through which it extends its axiopoda.  Foraminifera are ingenious architects forming thousands of intricately chambered shells, but the supreme architects are the Radiolarians and Acantharians which create spherical fantasies out of glass and are the envy of any aspiring architect or neophyte deity.  And, although there has been a lot of taxonomic shuffling in the last 25 years or so, these organisms all used to be classified as types of amoebae.  However, what interests us here is what we need to do to look at the surfaces of these organisms.

 Let’s start with the easy ones.  Buy a vial of radiolarians from a biological supply house.  The chances are 90% that they will be Eocene fossil specimens from Barbados, washed and cleaned and free of extraneous material  for less than $15 per vial.  It doesn’t get any better than this!  Many of these shells have very delicate, intricately sculpted spines and many of the specimens will be damaged, but this is unavoidable, unless you have the means to dredge your own samples or the laboratory facilities and reagents to process core samples collected by a major research institute and, even then, you probably won’t get any better results.

 A minute drop of these shells placed on a clean slide can provide hours of delight.  If you are fortunate enough to have access to a really fine stereo dissecting microscope with 100 or more magnifications, then this is where your observations should begin.  We need always to remember that these are 3-dimensional objects and the dissecting microscope vividly reminds us of that fact.

 Under the compound microscope, we confront a problem of contrast.  The high silica content gives these shells a refractive index very near that of glass and so, a very advantageous and aesthetically pleasing way to observe these shells, is by using Rheinberg illumination.  If you have a mounting medium with a high refractive index, you can let a drop containing the shells dry on a fresh slide, place a drop of mounting medium on it, cover slip it, and you will have a permanent preparation for your collection. The same procedures will work for diatoms and again, some biological supply houses sell small vials of diatoms.  You can also buy diatomaceous earth (it is sometimes used in aquarium filters and pet shops sell boxes of the powder).  You must, however, handle this material very carefully and avoid inhaling any of the dust or allowing skin contact, especially with eyes and mucous membranes, since this material is essentially ground glass.  Take a minute amount on the end of a flat toothpick and place it in a small drop of distilled water on a slide, using the toothpick to spread it out as evenly as possible.  Let dry and mount.  Extraordinarily beautiful exhibition slides of radiolaria and diatoms have been and are being made, but unless you have endless patience and a remarkably steady hand, I don’t recommend such an undertaking. Of the amateur microscopists who have attempted this feat, 72% have ended up in mental asylums, another 48% have required an average of 15 years of Freudian therapy at enormous cost, another 37% sold their microscopes and took up the accordion, and the final 17% had to have lobotomies.  (I know, I know, statistics is not one of my strong points.)

 Forams are less taxing and, with a moderate amount of patience and practice, you can produce some very respectable and attractive dry mounts.  For your own observations, you should make mounts using several specimens of the same species, so that you can see it from a variety of perspectives.  If it is a flat type of shell, mount specimens so that you can see both sides and then mount one or two upright to show a top view as well. I also always look for broken specimens which might provide me a glimpse into the wonderful maze of chambers which these extraordinary creatures create.  If you can’t find broken ones, sometimes, if the shell is not too impossibly tiny, you can get one to stick to your finger tip and then rub it very gently on a fine whetstone. Then, put it on a slide in a drop of water along with a very tiny, very, very dilute drop of acetic acid for just a minute or two and then transfer it back to tap water.  The acid attacks the calcareous material and will help loosen and rid you of the fine, annoying powder you have produced by grinding.  But, if you leave it too long., it will attack the shell.  I suggest the transfer to tap water, because most tap water is slightly alkaline and this will help stop the action of the acid.  If you’re not sure about your tap water, add a tiny (and I do mean tiny) bit of baking or washing soda to the water.  Then, rinse in distilled water.  You will likely discover that you have either ground the shell too much or not enough and will have to start over.  However, after a few tries, you will either get some nice results or abandon the procedure altogether.  For some of the large forms like Nummilites, which can be several millimeters in diameter, this grinding technique can give you a splendid glimpse of the complexity of the interior architecture of Forams.

 If you find a freshwater sample which you have collected and which has abundant shelled amoebae, it is a good idea to divide the sample in half and add pond or artesian water to both, but add food, such as, boiled wheat grains, only to one.  With the second one, add water regularly to counteract evaporation and, after weeks or even month, you will have a nice supply of relatively clean amoebae shells on the bottom of the dish or jar.  Take a pipet and remove some of the sediment, place it in a Petri dish, add some clean water and then with a micro-pipet, you can isolate a quantity of the shells onto a slide and examine their surface structure in detail.

 Trying to look at the surface detail of ciliates presents quite a different set of problems.  The outer membrane or pellicle is often sculpted in a distinctive way in a particular species.  After observing what I can in live specimens, I almost always use a simple technique which is sometimes very effective.  I say “sometimes”, because it will only work with ciliates which can go through drying out on a slide without completely disintegrating, in other words, they have to have a certain type of relatively tough pellicle.

 It does work with Paramecia.  So, take a drop of a rich culture, place it on a slide, add a drop of 1% to 2% Nigrosin (a very interesting purplish black stain), mix it up thoroughly using a toothpick and set it aside to dry.  You will find a considerable number of specimens which have dried in a relatively undistorted state and, the Nigrosin particles will have deposited themselves in such a manner, that you can get a clear idea of the surface structure.  In using this technique, it is important to have an abundant supply of the organisms in question, since with some genera, you may find that only 2 or 3 specimens on a slide produce the desired result.  Some ciliates are not suitable at all for this treatment and so it is important to keep careful records, so that you know whether or not you have had success or failure with this organism before.  Unfortunately, there are few parameters for predicting which organisms can be treated successfully and which cannot—trial and error, which makes it all the more important that we share the results of our experiments in places like Micscape.  It is easy to make assumptions that later turn out to be questionable.  It would seem natural to think that highly contractile organisms would not be amenable to this technique, but it turns out that, in a qualified way, a few are.  There will be considerable distortion in most cases, but occasionally, you will get a specimen that shows you just what you wanted to know.

   If you wish to observe trichocysts, a drop of blue ink or 1% methylene blue will stain them, but a drop of 1% tannic acid (or some leftover tea) will cause the trichocysts to discharge without staining them.

 A tiny amount of powdered carmine added to a small amount of culture will demonstrate the food vacuoles of a variety of ciliates and present you with a quite colorful display. This will leave the organisms alive and you can observe the feeding and digestive processes.  Vital staining with Neutral Red will also allow you to observe the digestive process, since this compound is a pH indicator, as well as a stain, and will change color as the pH within the vacuoles shifts.

 Cilia and flagella are often visible using phase contrast, but there are several special stains for demonstrating them, and these solutions are best purchased from a biological supply house.

 Many micro-algae secrete a mucilaginous envelope which, using ordinary contrast techniques, is virtually invisible.  A simply, effective, and therefore exciting technique, is to take a drop of a rich algal sample and thoroughly mix a drop of India ink with it on a slide.  If there is a mucilaginous envelope, it will stand out in a stunning fashion against the dark background of the ink.

 There is a wonderful old book: Bolles’ Lee Microtomist’s Vade Mecum which went through many editions.  It is full of helpful tips and techniques, many of which are useful to the amateur.  Most modern references are highly technical, specialized, and require the use of instrumentation and/or chemical reagents rarely available to the amateur.  What we need now is a modern Microtomist’s Vade Mecum focussed on the needs of the modern amateur.  Any volunteers?

All comments to the author Richard Howey are welcomed.