Silverizing 'Forams'

by Richard L. Howey, Wyoming, US


A few months ago, I began trying to find a method to demonstrate the silverline system in ciliates that would be simpler than the traditional methods and more accessible to the amateur microscopist. I have not yet had much success for several reasons:

1) The results have been mixed and only partly predictable.

2) Paramecium is by no means the best organism to use to try to develop such a technique. For starters, one has to contend with trichocyst discharge. Also, there are other organisms, such as, Euplotes and Blepharisma which are far better for demonstrating the silverline system. Unfortunately, at this time, I only had cultures of Paramecium and highly contractiles such as, Spirostomum, Stentor, and Lacrymaria.

3) I got sidetracked. As often happens, I will be working on a particular project and suddenly get a hare-brained idea about applying an aspect of it to something completely different. In this case, it was forams. I was fortunate enough, during this time, to be given a substantial number of fossil forams, all of which had been cleaned and concentrated. I was examining some under my stereo microscope and experiencing a bit of frustration about observing much detail, since there was rather little contrast in many of the specimens. I started to wonder what would happen if I applied my silver staining technique to foram samples. I should mention that these samples also contained the occasional sponge spicule, Dentalium shell, echinoderm fragment, radiolarian, snail shell, etc.
Let me describe the technique and how it led me to try yet another staining technique, and then I'll tell you about the results.

Silver staining technique

1) Place a small sample of foram and other shells at the edge of a small Petri dish.

2) Add enough 1% silver nitrate solution to fully cover the sample.

3) Take a dissecting needle or other small implement and push the specimens under the surface of the silver nitrate. Many of these specimens, but forams in particular, tend to float because of the air trapped in their minute chambers. This process may take several minutes and even then not all of the specimens will submerge. Don't worry about it. Just make sure that the majority are well-covered by the silver solution.

4) Let the sample soak in the silver nitrate for 10 minutes or 48 hours if you like. The time doesn't seem to make much difference, which is an advantage if you are in a hurry.

5) Now, this step may seem rather bizarre, but stay with me. Many of the silver techniques for protozoa involve exposing the sample to intense sun for some hours, or using ultraviolet light for some minutes and/or the use of a developing agent such as one would employ in photography.

What I use is a solution of vitamin C made by taking a standard 1000 mg. tablet and dissolving it in 60 c.c. of distilled water. (In one old bottle of solution there is even some mold growing, but it doesn't affect the efficacy of the solution). Where I got this idea, I have no idea, but it works.

Add a quantity of the vitamin C solution about equal to the amount of silver nitrate solution in the Petri dish.

In a matter of seconds, things start happening. Vitamin C being ascorbic acid, does, of course, produce an acidic solution which starts to attack the calcium carbonate in the foram shells, as evidenced by the formation of bubbles. However, not to worry; we're only going to leave the solution on for about 2 minutes and the slight etching that takes place will actually improve the visibility of the detail in the shells. A brownish, then grayish-silver precipitate begins to form first around some of the shells and then on the surface of the solution, blocking visibility of the shells. Take a toothpick and agitate the solution to distribute it as evenly as possible.

After about 2 minutes, take a micro-pipet and remove the solution. When most of the solution has been removed, add some 91% ethyl or isopropyl alcohol to this dish. (This can be obtained at your local drugstore.) Set the dish aside, uncovered, and allow the alcohol to evaporate. Put it in a place away from dust and away from flame or sparks as it is very flammable. In a few hours, examine your sample and you will discover that you have some rather elegantly silverized foram shells. They won't, however, look silvery, more like a flat, matte bronze, like your baby shoes your parents had preserved. (Do parents still do that?)

These specimens should be examined at about 40x with a stereo microscope using oblique illumination for best results. You will discover that with many specimens, the contrast has been significantly improved and, in addition, the results are often quite aesthetically pleasing; certainly, more so, than the results produced by the standard green food coloring that micro-paleontologists frequently use.

While I was developing this technique, I had a bottle of concentrated nigrosin sitting on my lab table. Nigrosin is a dye that, used in diluted form, can sometimes quite vividly reveal the surface structure of the envelope or pellicle of ciliates. They have to be, however, ciliates which will dry in a relatively undistorted and intact state, such as, Paramecium. The nigrosin deposits a thin film on the surface and can reveal remarkable detail.

While I was waiting for some specimens to soak a bit in the silver solution, I began to wonder what would happen if I used some of my concentrated nigrosin on some of the foram shells. I put some forams in a small Petri dish, added enough nigrosin to cover them, pushed them down with a tooth pick, and set it aside uncovered to let it dry. Since this is an aqueous solution, the drying takes considerable time.

The next day when I examined the sample, I was both surprised and pleased. Some of the specimens looked almost as though they had been coated with a dark metal film, while others had a bluish tint. The specimens tend to clump, held together by the nigrosin film. Using a pair of micro-forceps, you can gently pry out the specimens which you wish to separate and mount. [See the end of this article for a note on micro-forceps.] Sometimes a bit of the film will cling to the specimen, but this can usually be easily removed with a micro-needle after mounting. [See the end of this article for a note on micro-needles.] Again, the best way to observe these is with a stereo microscope at 20x to 60x using oblique illumination. At 60x to 80x, one can clearly notice a reflective patina on both the silverized and nigrosin specimens.

A few words about mounting. Scientific supply houses sell special slides for paleontology. There are two common types with several variations. The slides are made of a sturdy cardboard stock. The first type has a circular depression cut in the center 18 mm. in diameter and usually 1 mm. deep. The surface of the slide is white, but the bottom of the depression is black. This type is particularly advantageous for mounting sizable single micro-fossils or for creating interesting and aesthetically pleasing arrangements of medium-sized specimens.
The second type has a large rectangular depression cut in it approximately 22 mm. by 66 mm. The depression is black, but there are white lines forming a grid which divides the area into 60 numbered squares, each approximately 5 mm. by 5 m. These are made in two depths—1 mm. and 2 mm. The 1 mm. depth is adequate for most foram mounts and considerably cheaper than the slides with a 2 mm. depth.

I select the specimens which I wish to mount from small Petri dishes using a stereo microscope and micro-forceps. I transfer a few more than 60 specimens to the slide, since inevitably a few get broken or go flying off the slide during the process of mounting, never to be seen again. Using another small Petri dish, I place a fair-sized drop of Elmer's glue in it and one or two drops (ideally a drop and a half) of distilled water. Stir thoroughly with a flat toothpick and you are ready to begin mounting. I use two micro-needles to do the actual mounting. The first needle—I put a slight bend in this one near the tip—is the one I dip in the glue. The second one I use to nudge the specimen into the square in which I wish to mount it. Once in position, just slightly off-center in the square, I moisten the first needle with a tiny drop of glue and place it in the center of the square. With the second needle, I then gently position the specimen on top of the glue droplet. With a bit of practice, you will be able to judge the right amount to firmly hold the specimen in place, but not so much as to get it on the surface of the specimen.

Silver stained specimens need to be handled very carefully, as the coating of silver is very thin and rough handling with the forceps or the needles can scrape off portions of the coating.

Nigrosin stained specimens pose a different set of problems. As mentioned above, the shells tend to clump together and must be carefully separated. Bits of the nigrosin film may cling to a foram even after separation. Remove what you can easily with your micro-forceps and, for the moment, don't worry about the rest. Position your specimen as before and place it on the glue. Nigrosin is readily soluble in water and the diluted glue will dissolve any stray bits of the nigrosin film around the base of the shell. Nigrosin is a deep purplish-black and the slide background is black, so with a bit of care and luck, the stray bits of nigrosin film are absorbed undectectably into the background. If, however, there are still unwanted small projections of the film for the surface of the shell, these can almost always be effectively removed by a quick and careful flick with a micro-needle.

Once the mounting is completed, it is necessary to protect the slides from dust. There are four simple methods:

1) In the U.S., Ward's scientific supply company sells, in addition to the paleo-slides, aluminum slide holders, which unfortunately are rather expensive—currently $72 per 100 for the 1 mm. slides and even more for the 2 mm. ones. I cover the paleo-slide with a 1 mm. glass slide and carefully insert them into the holder. This is quick and easy and allows for the removal of the slide to examine the specimens directly or repair the slide if a shell or two happens to come unstuck.

2) This method is a bit more effort, but the results are quite acceptable and much cheaper. Purchase a box of 24 mm. x 50 mm. coverglasses. These are just lightly larger than the rectangular cell in the paleo-slide. When you use one, you must insure that it is as clean and dust-free as possible. I store mine in a small jar of 91% alcohol, remove one with a pair of forceps, wipe it gently with a lint free lab wipe, and then dust it with canned air. I use the same glue for the coverglass as I do for mounting the forams, only this time, undiluted. Use a flat toothpick or other convenient implement and apply a thin smooth layer of the glue around the edge of the cell. Carefully position the coverglass and lower it with a dissecting needle and then press it firmly into place around the edges with the needle. This method is slightly less convenient, since one must take great care to make sure the coverglass is spotless and if a shell comes loose, then one must pry off the coverglass, remount the shell, carefully remove the old glue and put on a new coverglass. Again, this method requires a bit more effort, but it is substantially less expensive.

3) This method is somewhat simpler than method #2, but less satisfactory. Instead of a coverglass, us a thin slide, preferably with a frosted end upon which you can write a code number and relevant data. If you use a clear slide, you can, of course, use a paper label for that information; I prefer the self-stick labels. With this method, you must make certain that the slide is as clean as possible, for once you glue it to the paleo-slide, attempts to remove it will effective destroy your preparation.

4) A somewhat less permanent method, which does allow one to salvage the slide if necessary, utilizes tape. Cut a 1" x 3" piece of posterboard or suitable equivalent and set the paleo-slide on this. Cover the paleo-slide with a clean, thin glass slide. Cut two 3" long strips of tape that are just wide enough to overlap the top and bottom neatly and tape each side.

These techniques are straightforward, easy, and quite fun to experiment with. In fact, right now I am trying out a modification of the silver staining technique on radiolaria and I hope to soon have an essay on that. Nigrosin, for a variety of reasons, is not appropriate for radiolaria or so I think at the moment. Maybe one of you will prove me wrong. I am especially pleased with the results of the silver staining, but sometimes the nigrosin is equally impressive. In any case, to my mind, both techniques are preferable to any of the colorizing techniques. I tried Alizarin Red S which is a specific stain for calcium and the results were both disappointing and rather garish.

I would be most interested in hearing from anyone who tries out these techniques and learning what sorts of results you achieved.

A Note on Micro-Forceps

One can spend an outrageous amount of money on micro-forceps. Some of the Swiss and German ones cost $20 to $30 each and they're quite good, but not excellent. A scientific supply house, with which I deal regularly, advertised "micro-forceps" for $1.40 each. I was skeptical, but ordered a few to try out. The forceps are made in Pakistan and, in fact, Pakistan makes quite a few different kinds of stainless steel surgical instruments, most of which are of quite reasonable quality. The forceps are not, however, suitable for handling forams, but with a bit of modification, they work splendidly for handling even quite delicate shells.

I bought a pocket diamond hone of the sort that fishermen use. It was less than $10. It has a thin groove down the center for sharpening fishhooks. I press the forceps closed and holding them at a 30 to 40 degree angle, draw them from left to right 10 to 15 times on one side, then turn them over and repeat the process.

I then remove any scraps or rough edges on the tips by rubbing them very gently on the hone. I check the tips under the stereo microscope to be sure that they are sharp and close properly. Sometimes, but not often, I have to repeat the entire process. Almost always, even with expensive micro-forceps, one tine is thicker than the other. By using the procedure described above, one can get sharp enough points so that that difference no longer matters.

A Note on Micro-Needles

Micro-needles can be easily made by using the small sizes of insect pins such as size 000 or even the "minuten Naeldeln" available from biological supply houses. Take some wooden match sticks from which the heads have been removed or, preferably, some 3inch lengths of very thin dowel. Now comes the tricky part; you need to make a small hole in the end of the stick or dowel. Position it securely in a small vise or clamp, so that if it splits, your hands are well out of the way and you don't end up skewering one of your fingers. Using a pair of wire cutters, snip the head off of the insect pin and insert that end into the hole into the dowel into which you have already placed a drop of household glue. An ordinary wood glue is quite satisfactory and avoids the risks of epoxy and superglues. I make up 8 or 10 such needles at a time and find a wide variety of uses for them.

Comments to the author Richard Howey welcomed.

Editor's notes:
The author's other articles on-line can be found by typing in 'Howey' in the search engine of the Article Library, link below.


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