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Protozoan Houses
by
Richard L. Howey, Wyoming, US
Images by Jan Parmentier, Holland
Houses, shells, test, loricas, gelatinous envelopes, mucilaginous domes—these are names of enclosure strategies for a variety of types of protozoa. If one wonders about the function of such devices, the immediate answer that suggests itself is, of course, protection and, indeed, that is often an aspect. However, it is certainly not the only one. Some writers have suggested that buoyancy and feeding are also factors in the development of such structures.
There are also specialized membranes in the protozoa, such as, those in the Sporozoa or the special membranes secreted in a variety of protozoa for encystment, but those will not be considered here. I hope to make those a topic for a future discussion.
These "houses" show a wonderful variety of form and structure. Indeed, the contemplation of their intricacy can provide many hours of aesthetic pleasure. In some instances, the details of these structures can provide important taxonomic information. To obtain such information may, however, be a test of one's powers of observation, one's equipment, and one's patience. In some cases, such detail is beyond the reach of optical microscopy, but, with many forms, optical microscopy can certainly provide sufficient data to determine the genus.
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The amoebas have probably the most spectacular variety of protective strategies. The foraminifera secrete shells with a startling variety of forms. Some look like miniature ammonites while others look like bits of coral. What they all have in common is that each shell possesses tiny holes or pores through which delicate filaments of protoplasm project. The radiolaria too secrete shells, but they tend to have a radial symmetry. The shells of forams are largely composed of calcium carbonate, whereas the shells of radiolaria are predominately silica. Radiolaria often look like tiny space capsules bristling with special projections designed to take in information about the environment, feed, and withdraw when getting adverse signals. Arcella's shell looks like a tiny dome, smooth in some species and stippled in others. Difflugia builds its house from materials at hand. Some species cement tiny sand grains together to form a pattern consistent for that particular species. Some species are less fussy and will incorporate sand grains, diatoms, desmids, and bits of debris into the shell construction. Such shells may have the virtue of providing some camouflage. Euglypha secretes shells which consist of plates, sometimes having spines at the edges. These can be very elegant indeed and look like tiny pieces of sculpture. Some flagellates and ciliates have protective collars or sheaths as well.
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For a first venture, I suggest looking at some samples of marine beach sand. If at all possible, use a low power dissecting stereo microscope. Place a small quantity of the sand in a small plastic Petri dish. Using a dissecting needle, one can sort the sand and separate out any foram shells. Forams have tiny chambers inside the cell which give some of them the appearance of miniature Nautilus shells. Sorting requires considerable patience. Some sand samples will prove to have virtually no forams, while others will provide a rich variety. Some locations provide a delightful wealth of specimens; others are virtual deserts. Whenever you hear about friends traveling, if only a few miles up the coast or to some exotic location halfway across the world, ask them to bring you a couple of small vials of sand. As a gentle, but not so subtle reminder, provide them the vials. Some friends of mine brought me some samples from a beach near Cancun, Mexico, apparently from an area with heavy surf. There were some very nicely polished specimens of forams resulting from the wave action. I gather that there was also a coral reef not too far off the coast, for there were also bits of mollusc shells, tiny pieces of corals, sea urchin spines, and other calcareous treasures. Often such sand samples, even if they don't provide forams, do contain small jewels from other organisms.
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When you have isolated some good specimens of forams, I suggest soaking them in mineral oil, castor oil, or immersion oil before mounting them on a slide with a cover glass for observation with the compound microscope. The oil penetrates through the pores into the chambers and improves the visibility of the inner detail. If the specimens are fairly large, they should be mounted in a depression slide or a slide with a cell and then covered.
Small vials of forams and radiolaria can often be purchased for a nominal cost from biological supply companies. If one does not feel confident about making one's own preparations, such suppliers also usually sell prepared slides of both forams and radiolaria. As mentioned above the shells of forams are composed mostly of calcium carbonate, whereas those of radiolaria consist largely of silica. Radiolaria are difficult to collect and prepare, so it is well worth buying a vial or two of "radiolarian ooze" from a supplier. Radiolaria shells are among the most beautiful of all microscopic objects. They are also difficult to study, for several reasons. Because they are made of silica, their composition is very nearly that of glass, and they, like diatoms, are difficult to resolve optically in order to see the finer details. There are a number of approaches which can be used to improve the image. Different types of contrast can produce spectacular results. Such contrast can be achieved by means ranging from inexpensive home-made devices to very expensive systems specially designed to produce high contrast images. Here I shall provide a brief overview of some of the techniques which are useful in observing micro-shells.
1) Darkfield—Darkfield illumination can provide spectacular images and make visible very small shells that one might easily overlook. This technique also has the virtue of producing quasi-3-dimensional images that are most striking. Darkfield can be achieved in a number of ways. Some of the older microscopes wer metal darkfield stops that can be placed in a filter holder below the condenser. If you have a condenser with a filter holder, but no darkfield stops, it is easy to make your own. With a little experimentation and a basic textbook, one can cut stops of the appropriate size for particular objectives out of black construction paper and mount them on clear plastic disks cut to fit the filter holder.
If you have a turret type phase condenser, this can be used to achieve a reasonably good darkfield. You simply rotate the condenser until you find the phase ring which provides you with a good dark field and the brightest contrast in the specimens.
Naturally, the best results will be obtained with the use of a specially designed darkfield condenser. Such condensers are available for virtually all good quality microscopes and are well worth the investment. Using such a condenser is rather messy, since one must put a drop of immersion oil on the top lens of the condenser and raise it until it makes contact with the bottom of the slide. This maneuver requires a combination of care and luck, because it is crucial to avoid the formation of any air bubbles in the immersion oil. Such bubbles will cause the light to flare dramatically obscuring the specimens. The preparation of the specimens is also critical and here too one must avoid air bubbles and the slide and cover glass must be scrupulously clean as bits of lint or dust will also stand out. The slides and cover glasses can be rinsed in alcohol, wiped with a clean, soft, lint-free cloth, and then dusted with a jet of air from a can of compressed air, such as that used for cleaning camera lenses or computer parts.
After observation, the slide must be removed from the stage with care to avoid getting the oil onto the stage. First, lower the condenser so that it is no longer in contact with the bottom of the slide. Next, using a fingernail, tilt the slide and lift it at an angle so that it does not come in contact with the stage, taking care not to let the slide touch the objective. If you wish to observe another preparation, you may need to add another small drop of immersion oil to the top lens of the condenser. If you are finished with your session, take a cotton swab or a piece of lens paper and remove the excess oil. Finally, using a special lens cleaner formulated for microscope and camera lenses, moisten a cotton swab with a small drop of the lens cleaner and with a circular motion gently remove the last vestiges of the oil. Remove the last residue of the lens cleaner with another dry cotton swab or two and then remove any cotton lint with compressed air.
This sounds like a rather tedious procedure, but with a bit of practice it can be accomplished quickly and efficiently and the results are well worth the trouble.
2) Oblique illumination—Oblique illumination is essentially a technique which involves side-lighting. Again, this can be achieved with a bit of experimentation by cutting crescent-shaped stops out of heavy black construction paper or by decentering the condenser, if one's microscope is equipped with centering screws for the condenser. An alternative method, if one has a phase condenser, is to slowly rotate the turret with the phase rings until one obtains oblique illumination. This technique is especially desirable, since if one gets everything properly "misaligned", one can produce pseudo-Nomarksi images of excellent contrast.
3) Rheinberg contrast—This method is fundamentally a modification of darkfield illumination and can be achieved relatively easily and inexpensively. This technique is sometimes disparaged by "purists" who argue that while it produces "pretty images", it does not provide any additional information of scientific importance. My own feeling is that an image that is aesthetically pleasing may very well serve to increase a general interest in microscopy and, further, I am convinced that sometimes the use of color contrast can lead the observer to notice detail that might otherwise escape notice. The psychological aspects of perception are by no means insignificant when applied to microscopic images Julius Rheinberg, a British microscopist of the late 19th and early 20th Centuries, used a variety of different colored transparent disks to create some quite wonderful effects. Disks of several types can be made. The stops are mounted on a circle of transparent clear or colored plastic. Green and red, blue and yellow are frequently used contrast combinations. One can use sheets of colored plastic from an office supply shop or preferably one can purchase a booklet containing an assortment of sheets from which one can cut the filters and stops. Edmund Scientific sells such books of filters. One will also require some sheets of clear plastic upon which one can mount the smaller colored disks as shown in the accompanying diagram.
Type 1 has a darkfield center stop mounted on a colored disk.
Type 2 has a colored center stop mounted on a clear disk.
Type 3 has a center stop of one color and then the outer disk is divided into quadrants with two opposite quadrants being a second color and the remaining two opposing quadrants of a third color.
Type 4 has a colored center stop mounted on a colored disk of a contrasting color.
Type 5 has three colors. The center stop is one color, half of the outside disk is a second color and the other half is a third color.
Type 4 is relatively easy to make, produces beautiful images, and is probably the most common type.
Color saturation is important for Rheinberg illumination to work properly. If necessary one can use two layers of a given color to increase the color intensity, thus enhancing the contrast.
4) Polarized light—One usually employs polarization for the observation of crystals, but since a number of protozoan shells are calcareous, this technique is also useful here. In fact, one can place a specimen slide on the stage, cross the polars so that the background field is dark, and then scan the slide. Calcareous material will stand out vividly.
If one is fortunate enough to have a dedicated polarizing microscope, then one can experiment with the various compensators to achieve additional color contrast effects. However, one can easily make one's own compensators either by cutting sheets of mica or by cutting a piece of plastic from a small plastic Petri dish. Differences in the thickness of the layers of the mica or plastic will produce different color effects. Trial and error will permit one to find the most pleasing combinations. Compensators must be placed between the two polars (i.e., between the polarizer and the analyzer). If one does not have access to sheets of mica and does not have the tools for cutting the plastic Petri dish, there is a simple technique that works for some specimens. One can take a very small Petri dish bottom and place the specimens in it. This works best if the specimens are dry and, since they are not covered, one should use only low power objectives to avoid damaging any lenses. The Petri dish is placed on the stage and the plastic of the dish will act as a compensator.
5) Nomarski Differential Interference Contrast—This technique gives superb contrast for certain types of specimens and allows for optical sections. However, the equipment is very expensive and requires very careful adjustment. Nomarski DIC is especially valuable for examining delicate membranous houses, such as those found in certain collar flagellates or in some species of Stentor.
I strongly recommend trying out a variety of contrast methods when looking at such protozoan "houses".
Comments to the author Richard Howey are welcomed.
Editor's note:
Look in the Article Library 'Techniques' section for articles discussing darkfield, oblique illumination and use of polarised light.
Use the keyword 'Foraminifera' in the Library Search page to show Micscape articles on the collection of foraminifera. Or use 'protozoa' in a library search to locate related Micscape articles and images.
Published in December 1998 Micscape Magazine.
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