Contents

Color

To use filters and stains effectively you must understand a bit of color theory. Because microscopy uses both additive and subtractive color mixtures, color can be confusing.

Subtractive color mixtures occur when particular colors are subtracted (absorbed) from light. This occurs when a light passes through a colored filter or a stained specimen. Components of the light that are not the color of the filter are absorbed by the filter; light the color of the filter is transmitted through the filter. Using filters with transmitted light results in a subtractive color mix. A look at the figure shows how subtractive color mixes work.

Red, yellow, and blue are the subtractive primaries. All other colors can be created by subtracting these primaries. An orange filter subtracts the blue component from white light; a green filter subtracts the red component from white light; and a violet filter subtracts the yellow component from white light. Subtracting red, yellow, and blue components from white light leaves black.

There is another way to think about subtractive color when mixing colored liquids. (If colored stains react chemically with one another or with a part of the specimen, this may not hold true.) Mixtures of the primaries give all other colors. Mixing red and yellow liquid creates an orange liquid, mixing blue and yellow creates green, and mixing blue and red creates violet. A balanced mixing of all three primaries leaves black; no light passes through.

Additive color mixtures (are achieved by adding one light to another, and obey different rules than subtractive mixtures. Red, blue, and green are the additive primaries. All other colors of light can be created by combining these primaries. If we shine a red light and a green light at a spot on a screen, the two lights will combine to create a yellow spot. Similarly, green and blue lights combine to create cyan, and red and blue lights combine to create magenta. White light results from a balanced combination of red, green, and blue.

A reflected light (shining down onto the specimen from above) will add its color to any light that is already being transmitted upward. If a white light is shining down, a tint of the transmitted light's color will result. That is, the color of the resulting light will be the color of the transmitted light, but it will be brighter and less saturated.

It is possible to predict the color of light that will reach your eye through the microscope. Start with the color of the transmitted light source, which will probably appear white to your eye. This light may, in fact, be quite a different color from light that your eye percieves to be white under other circumstances. Every light source has a color temperature, measured in degrees kelvin (K). Solid substances, such as lamp filaments, glow red at low temperatures, blue at higher temperatures. You should try to find out the color temperature of your microscope illuminator; this is particularly important if you take color photographs through your microscope. While your eye can adjust to differences in color temperature, film cannot. We will discuss this more fully in the section on photomicrography.

First, use the subtractive color chart to determine the color of light that will reach the specimen through any filters. Subtract the color of the (possibly stained) specimen from this color. Finally, use the additive color chart to add any colors reflected from the specimen by any reflected light shining down at the stage. The result of all these additions and subtractions is the color that reaches the eye.

Several things can complicate this procedure. The balance of the intensity of the reflected light to the transmitted light varies. The specimen may be multicolored. Reflected light from above may travel through colored, semitransparent parts of the specimen before being reflected back upward from the object's (possibly colored) interior. With Rheinberg illumination, different colors of light will pass through different parts of the specimen. With Rheinberg, darkfield, and oblique illuminations, some light will be reflected from the specimen that would ordinarily be transmitted through it.

Even with these complications, predicting the result of a color setup becomes possible after some practice; one becomes more sensitive to lighting conditions and more experimental with light. You should build illumination setups slowly, asking yourself, "How will the next component affect the color of light reaching my eye? How will the component affect contrast? What parts of the specimen will it bring out or suppress?" Experiment; excellent illumination setups will soon result.

The resolution in achromatic optical systems, which are usually corrected for red and blue, can be improved by using filters that allow only red and blue light to reach the eye. For instance, a red filter with a blue stained specimen or vice versa works well, or a blue and red stained specimen with white light passing through it works well. Another solution is to use a strongly monochromatic filter, which allows only one color of light to pass; green is often used. In either case, all light passing through the lens focuses at the same point. This reduction of chromatic aberration improves resolution.

Complementary colors lie exactly opposite each other on the color wheel. Examples of subtractive complements are red-and-green, yellow-and-violet, and blue-and-orange. If a filter is a subtractive complement of the specimen and transmitted light is used, value contrast between the specimen and background will be increased; value contrast between structures within the specimen will be reduced.

If the filter is the same color as the specimen and transmitted light is used, value contrast between the specimen and background will be reduced; value contrast between structures within the specimen will be increased. Resolution will also be improved because the effect will be much like that of a strongly monochromatic filter.

Contents

Microtechnique: Temporary Slide Preparation

Slides for general use can be kept in 70% alcohol. Ordinary rubbing alcohol, which is very inexpensive, is the best thing to use. The slides can be removed and dried with a cloth as needed.

The most common way of making a temporary slide is to place the specimen on the slide in a liquid and then place a cover slip on top of the specimen. Grasp the cover slip by the edges, set one edge of the slip on the slide near the drop, and slowly lower the cover slip using a teasing needle or toothpick.

For non-biological objects, choice of a mounting liquid is often determined by the difference of refractive index between the mounting medium and the specimen. The more difference between the refractive indices, the more strongly the specimen will be outlined against the background.

For biological specimens, it is usually best to choose a liquid that will not interfere with the structure of the specimen. Tissues from most animals do best in a weak salt solution. The amount of salt varies with different animals, usually between 6% and 10%. The solution is made by weighing the distilled water and salt on a small scale, then combing the proper amounts. It is not convenient to mix a new solution for each specimen. Just keep a couple of strengths on hand and test which works best for the type of specimen. Most botanicals do fine in distilled water. Water dwelling organisms require the water in which they have been living.

Living organisms like protozoans, tiny fresh water worms, and water fleas require room to move about, and the water must be kept from evaporating from the slide. A couple of approaches are used. Using either, organisms can live a long time on a slide. A special well slide can be used. These slides have a depression that is just big enough to hold a drop of water. The sequence of actions needed to make a hanging drop preparation with one of these slides is illustrated in the figure. Use a medicine dropper or inoculating loop to place the drop containing living organisms on a cover slip. Make a ring of Vaseline around the depression on the slide. Lay the slide face down on the cover slip, placing the depression over the drop, and flip the slide right side up quickly, so that the drop does not slide to the side of the depression. If an oil objective is to be used, try focusing on the edge of the drop before trying to look at other areas. Do not try to focus the oil immersion objective deeply into the drop, or you will break the cover slip and possibly scratch the objective. For smaller microscopic animals, a well slide is not necessary. Just place a drop of the culture inside a ring of vaseline and lower a cover slip onto the slide, pressing down lightly. The vaseline gives just enough shimming so that the tiny creatures are not crushed. They will be able to swim about, but will still be nicely flattened between the slide and the cover slip. Any time vaseline is used, the slide will require cleaning with detergent before being stored in alcohol.

The smear is another useful technique. Smears are used on both temporary and permanent slides that require the examination of liquids or the examination of objects dispersed in liquids. The figure illustrates the required action. A drop is placed near the end of one slide, and a second slide is used to pull the drop along the length of the first slide. The angle at which the slide is held influences the thickness of the smear. Do not make the common mistake of pushing the drop with the slide. This will crush the structures in the specimen.

Blood is one of the most commonly smeared substances. Old sources recommend pricking a finger to get a blood sample, a practice which is unnecessary for the hobby microscopist, and now highly dangerous if a needle should be shared or incorrectly cleaned. Unless you have proper and qualified supervision, work with blood from beef that you obtain from the supermarket.

WARNING: Any work with blood should be done under adult supervision. If you feel you must prick a finger, never use a needle that anyone has used in the past. Dangerous diseases can be spread by used needles.

We often need to draw a liquid under a cover slip while observing the reaction of the specimen. With a dropper, place a drop of liquid so that it touches one side of the cover slip while applying a little piece of blotting paper to the other side. The new liquid will be pulled under the cover slip, replacing the liquid that is absorbed by the blotting paper. An example of the use of this technique would be drawing iodine under a cover slip to watch its reaction with starch. As the iodine comes into contact with the starch particles they are turned blue or purple. This is a common test to determine whether an object contains starch particles.

Staining Temporary Mounts

Vital staining (also called supra staining) can be used with living specimens. The stain can be fed to or injected into the organism, or the organism can be immersed in a solution containing the stain. The idea is to color the specimen without killing it. Immersion is often used with microscopic organisms, which tend to be colorless. The technique is important for such specimens if you do not have access to sophisticated contrasting optics.

Neutral red is particularly valuable owing to its low toxicity and strong coloring power. Some organisms can even be raised for generations in very dilute concentrations of this stain. Higher concentrations of this or any other vital stain will kill the organism. In addition to neutral red, other vital stains include Janus green B, eosin Y, brilliant cresyl blue, nigrosin, Bismarck brown, crystal violet, and methylene blue. Dilute India ink will darken the water around organisms, allowing them to stand out brightly. The food coloring FD&C blue, available under different brand names in any grocery store, also provides an interesting negative stain. The water is turned blue while the organisms swimming about in it remain clear. Use of other food colorings as vital stains is also worth some experimentation if you have problems obtaining chemicals.

The liquid into which a vital stain is dissolved is sometimes toxic to organisms. If you are having this trouble with one of your stains, try the following procedure. Apply a thin smear of the stain to a slide and let it air dry before adding the liquid culture. The stain, if it is water soluble, will dissolve into the culture medium. A similar technique can be used to mix a stain with larger amounts of culture. Let some of the stain dry in the bottom of a container, and add the liquid culture after the stain dries.

The entire range of biological stains can be used for temporary mounts of dead material. We will discuss staining in more detail in the discussion of permanent slide preparation.

Permanent Slide Preparation of Biological Materials

WARNING: Several chemicals used in permanent slide preparation are toxic or explosive. If you are not an adult, get permission and supervision before working with any of these chemicals. Gloves, aprons, and eye protection should be worn to keep these chemicals off the skin; the area for their use should be either very well ventilated or furnished with a fume hood. Sparks and flames should be kept away from many chemicals. All chemicals should be stored properly. Formaldehyde is a positive carcinogen. Xylene and toluene are also suspected. Benzene is strongly linked with leukemia and liver damage, and, since substitutes exist, should not be used at all. Some biological stains are also toxic. Get to know all of your chemicals, and learn to handle and store them safely.

To make permanent slides of biological objects requires a collection of chemicals. Thousands of recipes have been developed for specialized purposes, but it is not convenient for most microscopists to keep hundreds of chemicals on hand. The following description uses only a limited number of chemicals, and will work for most specimens.

A minimal set of chemicals for an adult will include formaldehyde or (preferably) glutaraldehyde, absolute alcohol (isopropyl is acceptable, ethyl is better), distilled water, xylene or toluene, a set of biological stains, mounting resin, and some more commonly available household chemicals.

Younger microscopists can get satisfactory results with a set of safer chemicals that includes glycerin, rubbing alcohol, distilled water, some of the less toxic stains (crystal violet, India ink, methylene blue, and drawing inks in different colors), and mounting resin. All of these can be collected with a trip to the drugstore, art supply store, and grocery store.

Biology supply houses in the United States absolutely refuse to sell chemicals to individuals of any age, but in many areas local chemical suppliers will do so. They may carry formaldehyde, absolute alcohol, acetone, xylene, and toluene. Paint stores and art supply stores sometimes carry acetone, xylene, toluene, and turpentine. Chemicals from paint stores will not meet laboratory standards, but usually work well enough when better chemicals are not available. Local suppliers will probably sell in batches larger than desired, the smallest available size often being four liters, or about a gallon. Formaldehyde and glutaraldehyde are expensive in such large quantities.

The steps in making permanent slide preparations of biological objects involve fixing, dehydrating, embedding and sectioning (often optional), staining (often optional), clearing, and mounting.

Fixing

To fix is to kill living tissues while minimally disrupting their shape and structure. A solution of formaldehyde is a common fixing agent. One part of 40% formaldehyde solution (this is the standard strength of commercial formaldehyde solution) is mixed with four parts water. Glutaraldehyde is usually used in solutions of about 6%. Depending on the stains to be used, the acidity of the solution can be modified by adding various concentrations of acids or bases. Buffers are also commonly used. This will be discussed further in the section on staining. Many other special purpose fixatives are available, but we will confine our discussion to general methods that work on most materials.

The amount of time that an object should be left in fixative solution depends on its dimensions and fixative used. For paper thin slices ten minutes may be enough. If materials are about .5 cm thickness, 24 hours should be sufficient. In any case, leave the material in the solution until the fixative can penetrate throughout.

Formaldehyde and glutaraldehyde, being toxic, require careful handling. For younger microscopists, a safer substitute is dilute alcohol, which does a fair job on many specimens, although some disruption of cell contents will occur.

To fix with alcohol, simply begin dehydrating as described in the next section, starting with a 10% solution. By the time you work up to a 50% solution, the specimen should be thoroughly fixed.

Anyone without access to biological fixatives can order preserved materials from suppliers. This eliminates the fixing step. (Considerations that prevent sales of chemicals to individuals do not seem to prevent sale of objects treated with chemicals.) These preserved materials can be dissected to obtain samples for microscope slides. A collateral benefit is that dissection is a fascinating activity in itself. Dissection is easier if materials injected with color are ordered, but the injected chemicals may interfere with the desired stain. Plain preserved materials are inexpensive and better for permanent slide preparations.

Dehydration

After the material is fixed, it must be dehydrated. This does not mean setting the specimen aside to dry, which would be a serious mistake. When the specimen is dehydrated, all of the water in it is replaced by another chemical. This will usually be alcohol, acetone, or glycerin. The classic dehydration method involves soaking the material in increasingly stronger solutions of alcohol, culminating in several changes of nearly absolute (100%) alcohol. The number of steps depends on the delicacy and type of the tissues being dehydrated. For general use the following schedule of alcohol works well: 10%, 20%, 50%, 95%, 100%. (Acetone can be substituted for alcohol with this method of dehydration.)

To hydrate a specimen, simply reverse the steps, starting at 100% alcohol and soaking the specimen in weaker and weaker solutions of alcohol, finishing with distilled water. Moving in series from alcohol to water is necessary only for whole objects. Cut sections can be brought directly into water. Many recipes require dehydration and hydration several times; it is best to keep small containers of these alcohols or acetones premixed with distilled water and ready for use.

Glycerine can be used to dehydrate specimens in a single step. Place the material in a solution of one part glycerin (available in most drugstores) and six to ten parts distilled water. Let the solution set for several days in a watch glass or beaker until the water has evaporated and the glycerin returns to its normal viscosity. A glycerin mount, which will be described later, can then be used. If absolute alcohol is available, it can be used to remove the glycerin from the specimen and the usual dry mount can be used. Embedding, discussed in the next section, can only be done if the glycerin can be removed.

If a specimen is to be stained and also to be both dehydrated and mounted in glycerine, staining should take place before dehydration. Most stains are unstable in glycerin, which remains liquid. Staining before dehydration gives most of the stain that is going to leach into the glycerin time to do so before the specimen is mounted in fresh glycerin.

Embedding, Sectioning, and Pasting

The science most closely associated with embedding and sectioning is Histology. Histologists seek to understand the structure, function, and microanatomy of cells and tissues of animals and plants. The best way to gain an understanding of most plant and animal tissues is to slice them into extremely thin sections. One can build a mental image of a three-dimensional structure by looking at sections taken from it. Sections from as few as three planes are needed to show the entire structure of stems. Asymmetrical organs and tissues from animals require more sections.

Mechanical stress from sectioning will destroy microscopic structures of soft tissues, so they must be reinforced by being embedded in a material that will support them mechanically. Embedding is unnecessary for harder tissues. For instance, sections can be taken from many plants, like carrots, and from animal cartilage without embedding. For these, careful hand cuts made with a razor blade will do. For most tissues, however, embedding works best.

The practice of embedding, being time consuming, separates the serious microscopists from the casual. Nevertheless, it is a valuable method that can be carried out inexpensively, even in a home laboratory. Specimens that have been embedded, sectioned, and stained rarely require fancy illumination setups; standard brightfield illumination is preferred.

Paraffin is the most commonly used embedding material. Specially prepared embedding paraffin can be obtained from biology suppliers. Paraffin from the corner grocery store can even be used if the specialized product is unavailable.

The dehydrated specimen should soak in xylene just before embedding. Xylene is a good solvent for paraffin and removes any alcohol (from the dehydration step), which is not a good paraffin solvent. The xylene helps the wax to more thoroughly infiltrate the specimen. A good grade of water free turpentine may be substituted for xylene. Some microscopists soak the specimen in a room temperature solution of xylene and wax for several hours to ensure that the wax completely penetrates the specimen.

Next, the specimen is soaked in molten paraffin. The wax must not be overheated. Sixty degrees centigrade is about right. If a proper heating cabinet is not available, a toaster oven, door ajar, can be used. Another makeshift is to melt the wax in an egg poacher or double boiler with the cover removed and on very low heat. The water in the lower container must never be allowed to come to a boil. Use a thermometer often to test the temperature when using these methods. Overheated wax will ruin the specimen. Some microscopists place a heat lamp into the top of a beaker that has paraffin in the bottom. The lamp is lowered incrementally until most of the wax in the beaker is molten, but the wax at the bottom is still solid.

The specimen must be left in the paraffin long enough to be soaked through completely. This may take several hours for a cube with five millimeter sides. The paraffin should be changed several times to prevent the clearing agent in the specimen from contaminating it. While sectioning you may occasionally encounter some clearing agent on the knife. This means that either you have not soaked the specimen long enough or the paraffin has not been changed often enough.

After soaking, the specimen and wax are poured into a small container. Some quick work with a toothpick will orient the specimen in the container before the wax solidifies. Some workers prefer to pour a bit of wax into the container, orient the specimen, top up the container with wax, and remelt all of the wax in the container with a hot tool.

Choice of a container is personal. Specialized embedding frames are no longer easy to find. A shot glass or the like will do. A few microscopists still like to fold little paper boxes to hold the poured wax.

Coat the inside of the container with glycerin to allow quick release of the wax when it solidifies. Cooling the container before pouring also helps. After the surface of the wax has hardened, the container should be plunged into ice water. Quick cooling helps to prevent crystallization of the wax, but if the block is quinched before the surface has hardened, deformation of the specimen will occur. Be sure the wax has thoroughly hardened before trying to free it from the container.

A microtome, if available, is used to cut the specimen into very thin slices. After the wax has solidified, it is trimmed into a cube that surrounds the specimen. Usually, the dimensions of the cube are about one half inch on each side, although the dimensions of the specimen within the block should be much smaller.

The trimmed wax block is mounted either directly on the microtome stage or on a wooden block that can be clamped inside a clamp type stage. A spatula (an artist's palette knife works well) can be heated and used to press a bit of wax onto the wooden block or stage. Then the spatula is reheated and placed between this newly waxed base and the wax block with the specimen. The wax block is pressed toward the base, and the hot spatula is slipped from between the two. A strong welded bond results because both wax surfaces melt concurrently.

After the mounted block has cooled, it is trimmed to a smaller size, so that only a little wax surrounds the specimen. A specimen of only five millimeters to a side will still fill the visual field at 40X magnification, so there is no need for the final wax block to be more than seven millimeters to a side along the plane of the cut. Specimens for viewing at higher magnifications can be much smaller. Bear in mind that the smaller the sections are, the less problem you will have with curling of sections and with problems caused by the knife not being at a perfect angle for the cut.

Microtomes come in many shapes, sizes, and price ranges. We will concentrate on the least expensive, a hand microtome, which will do for most purposes. The straight razors that come with these microtomes are a pain to keep really sharp. You can also use a new utility knife blade. Cut as shown in the illustration, pushing or slicing with the thumbs while steadying the microtome with the fingers. If sections of the same thickness are desired, each section must be cut at the same speed. It is also best that the cut be made in one smooth movement. This is another reason to keep the size of the specimen block very small. A drop of alcohol spread along the blade will help to make the sections cut more smoothly. Clean the blade often.

If a hand microtome is unavailable, a nut and bolt can be substituted after the nut face has been carefully smoothed with emery paper. The wax block in this case should be trimmed not into a cube but into a cylinder with a diameter approximately the size of the bolt, and is attached to the bolt tip with a drop of hot wax.

Sections can also be cut from the wax block using nothing but a single edge razor blade. Although slightly thicker and more uneven than microtome cuts, sections cut this way can be informative and beautiful.

Thinly spread some egg white (albumin) on a clean slide with a finger tip. (Fresh egg whites can be used. Dried albumin is available from biology suppliers. Several brands of low cholesterol egg substitute, sold by grocery stores in small cartons, are 98% egg white, and work very well.) Some workers prefer to mix the albumin with equal amounts of glycerin. Sodium salicylate can be used as a preservative.

The best thickness of the sections depends on the material and the kinds of structures to be visualized. With very thin sections the curling will be so slight that the sections can simply be pressed into the albumin with a finger tip. More curling will occur with thicker sections. Another thing that affects curling is room temperature. When too much curling occurs, try heating the room or setting the microtome near a heater. If the sections still curl too much, special warming tables are made for relaxing them. There are also some makeshift substitutes.

A warm water bath can be used to uncurl the sections. The sections are hydrated at room temperature. Distilled water is heated to about 40 degrees centigrade and the sections are dropped in. The sections will immediately relax and float on the water's surface. Insert one end of an albumin coated slide into the water next to a section, hold the section to the slide with a dissecting needle or toothpick, and lift the slide so that the section is lifted with it. Gently press the specimen to the slide, and set the slide aside to let the water evaporate. (If glycerin is mixed with the albumin, the slide will not appear to be dry even after all of the water is gone.)

Another method is to place the hydrated sections on the albumin coated slide, then put enough water on the slide to float the sections. Quick bursts from a small flame can be used for heat. If a Bunsen burner is not available, a cigarette lighter can be used. If the wax should begin to melt, quickly remove the slide from the heat and let the slide cool for a few moments.

After the sections are flattened and the slide is completely dry, flame the slide from beneath to coagulate the albumin. The wax can be allowed to melt a bit at this point, but do not apply so much heat that the specimen is damaged. Wipe any accumulated carbon from the bottom of the slide.

Finally, dissolve the paraffin from the sections (decreation) by immersing the slide in xylene. With very thin sections this only takes a few minutes. The xylene does not dissolve the bond between the specimen and the albumin, but the wax floats away.

(Some brands of embedding paraffin contain polymers that are birefringent. These polymers will interfere with polarized viewing unless all of the embedding medium is carefully removed with a solvent for the polymer. If polarized viewing is contemplated, common paraffin is preferable for embedding in the absence of such a solvent. Sections for polarized viewing should not be pasted with albumin. Unpasted sections will stay on the slide if it is handled very carefully. The sections should also be as thin as possible to avoid overlap of interference color phenomena.)

We can now summarize embedding and sectioning:
(1) Dehydrate specimen;
(2) Soak specimen in xylene or turpentine to remove dehydrating agent;
(3) Soak specimen in paraffin, pour, and cool;
(4) Cut sections;
(5) Hydrate sections;
(6) Relax sections if necessary;
(7) Paste sections;
(8) Dehydrate sections;
(9) Decreate (remove wax)

Staining Permanent Preparations

The dyes used for staining organic specimens are usually salts. Salts are ionic compounds -- compounds composed of ions. An ion is formed when an atom gains or loses an electron, so ions are electrically charged. A salt possesses both a positively charged and a negatively charged ion. A colored ion is called a chromophore. Salts with a negative chromophore are called acidic dyes; salts with a positive chromophore are called basic dyes. Most stains are either basic dyes or acidic dyes. Either kind of dye can be any color. For example, safranin is a basic red dye, and eosin is an acidic red dye.

Organisms are composed of one or more cells. The cell constituents are negatively charged and attract positive ions. Because basic dyes have a positive chromophore, the cell's atoms combine with it and become colored. Acidic dyes do not combine with most cell constituents, so the cell is not stained. However, acidic dyes will form a deposit around a cell or color the background. Such negative stains give an enhanced view of the shape of the cell.

Mordant dyes are neither basic nor acidic. They require a mordant, a substance that is an intermediary between the dye and the specimen. The mordant dye binds to the mordant, and the mordant binds to the specimen. Common mordant dyes are cresyl violet and hematoxylin. Common mordants are metallic salts, acids and bases. (Iodine is also a common mordant, but should only be used for temporary preparations.)

Most biological stains will form crystals if a drop is placed on a slide and simply allowed to air dry; we would expect this of salts. Biological preparations are therefore never allowed to air dry through the entire process of staining, dehydrating, and clearing. If they were, the stain would crystallize. Amorphous substances are solids that do not have a crystalline structure. The mounting resin that will eventually surround the specimen will air dry, but, being amorphous, it discourages the stain and other parts of the preparation from forming more crystals than may already exist.

If sections that were embedded and pasted to a slide are being used, flood them with alcohol and then hydrate them before staining. The alcohol removes any xylene introduced during decreation of wax from sections. (Do not hydrate if the stain is dissolved in alcohol. Usually this is not the case.)

With sectioned materials, you will work by applying the chemicals to the pasted section on the slide, or by immersing the slide in the chemicals. A spot plate is handy for material that has not been sectioned. This is a plate that has a number of depressions about one centimeter in diameter. The specimen can be easily moved from one depression to another with a small soft haired artist's brush. A dropper is used to put stains and chemicals into the depressions.

After the stain is applied to the specimen, excess stain can be removed (known variously in the literature as destaining, decolorization, or differentiation) by moving the specimen through several changes of a solvent for the stain. The common destaining agents are alcohol or an acetone-alcohol mix. For basic dyes acid alcohol speeds the process. Acid alcohol is 95% ethyl alcohol that contains 2.5% nitric acid. (A mixture of any alcohol and household acid often works well enough. You may want to experiment with the citric acid in lemon juice or the acetic acid in vinegar.) For acidic dyes basic alcohol speeds the process. A few drops of ammonium hydroxide (household ammonia) to an ounce of alcohol can be used. For destaining mordant dyes, more of the mordant is normally used. Acid alcohol, basic alcohol, and mordant solutions should never be so strong that the specimen is damaged.

Some structures and types of cells are more easily decolorized than others. Counterstaining takes advantage of this. An initial stain is applied and destained; then a second stain of a contrasting color is applied. Structures that destain easily become the color of the second stain; those that do not remain the color of the first stain.

Basic dyes stain selectively depending on the conditions of acidity. Very few cellular structures stain if the dye is mixed with an acidic solution. If the dye is mixed with a basic solution, many more cellular structures will stain. Similarly, if the specimen has been fixed in an acidic solution fewer structures will stain in basic dye than if the specimen has been fixed in a basic solution. Acidity of liquids can be tested with pH test paper. When pH is below 4, only the ground substance that surrounds cells and connective tissue fibers stains well. When pH is raised to 5, DNA and RNA begin to stain. When pH is raised to 10 or above, proteins begin to stain. It is informative to stain similar specimens at each of these pH levels.

If you have access to chemicals, thousands of recipes for stains are available, many of them specialized to demonstrate particular structures of particular specimens. Consult a library for books on histology, histochemistry, cytology, microbiology techniques, and works on microscopy that stress staining and slide preparation techniques. Do not blindly follow a recipe when staining; inspect the specimen under the microscope.

At least a few good stains are available to anyone. Gentian violet can be found in drugstores. It is also known as crystal violet (a more chemically pure varient), and basic violet 3. A few drugstores carry methylene blue, also known as basic blue 9. Gentian violet and methylene blue are basic stains. India ink, strained through filter paper and diluted with water is a good negative stain. Some colored drawing inks and liquid clothing dyes will also work. You will have to experiment with different brands; failures are easy to recognize by the grainy or spotty effect that they have on the specimen when viewed under magnification.

After staining, the specimen must again be dehydrated. Because the sections are thin, the dehydration goes quickly.

Clearing and Mounting

Clearing is normally done just before mounting. Turpentine, toluene, xylene, and clove oil can be used for this. The choice depends on the mounting resin to be used. Clearing is done to increase the transparency of the specimen and to remove the alcohol left in the specimen during the final dehydration. Alcohol is not a good solvent for any of the common mounting resins.

Because Canada balsam and damar varnish are already dissolved in turpentine, it is a good clearant for them. Xylol and clove oil are also good with these mounting resins. Clove oil should only be used if you wish to increase the transparency of the specimen. For acrylic mounting, toluene or xylene are good clearants; most acrylics are already dissolved in either xylene or toluene. It is best not to use turpentine or clove oil with acrylic resins. Acrylic does not polymerize properly in the presence of oil.

Mounting in resin is the last part of a permanent slide's preparation. If a small object is being mounted, it is simply placed into a drop of liquid resin on the slide and covered with a cover slip.

The choice of a permanent mounting medium involves trade-offs. Canada balsam is the classic natural resin for mounting. Damar varnish, another natural resin, forms a tougher film but has a yellow tinge. This is not a problem unless the mount is extraordinarily thick. Both of these resins become brittle and more yellow with age, losing their adhesive qualities. This process is slow, however, and usually does not occur before the specimen itself deteriorates and the stains fade. Many microscopists have replaced the natural resins with acrylic resin dissolved in toluene or xylene. Acrylic resin does not yellow and retains adhesion better than the traditional balsam and varnish mediums. However, some brands of acrylic medium, as they come from the supplier, tend to trap air bubbles. Thinned more with toluene or xylene they do not trap bubbles, but suck bubbles under the cover slip as they dry. Canada balsam and damar varnish can be used at a thin consistency without sucking air bubbles under the cover slip.

Canada balsam and acrylic medium are available from microscope suppliers. Good grades of damar varnish and damar crystals are available from artist's supply sources. Do not try to use acrylic emulsion, available from art supply sources, for mounting. It contains water and will ruin the specimen.

If the specimen was dehydrated in glycerin, and no absolute alcohol or acetone is available to remove all of it, a glycerin mount must be used. This is an old fashioned and somewhat tricky technique, but no clearing step is necessary before mounting.

One way to mount in glycerin is to place the specimen in a drop of glycerin on a large cover slip. Place a smaller cover slip over the specimen. Put enough mounting resin on a slide to seal the entire large cover slip and place the slide over the large cover slip. This will seal in the smaller cover slip and specimen.

Another technique is to place the specimen on a slide, in glycerin, under a cover slip; then very carefully and completely seal the edges of the cover slip to the slide with a fast drying resin. A pointed artist's brush can be used. Several coats may have to be applied to the line of sealant.

Recipe summaries

The general summary for permanent slides is:
(1) Hand cut sections (optional);
(2) Fix;
(3) Stain;
(4) Dehydrate;
(5) Clear;
(6) Mount.

If embedding is used, the following, more complex procedure can be used:
(1) Fix;
(2) Dehydrate specimen;
(3) Soak specimen in xylene or turpentine to remove dehydrating agent;
(4) Soak specimen in paraffin;
(5) Cut sections;
(6) Hydrate sections;
(7) Relax sections;
(8) Paste sections;
(9) Dehydrate sections;
(10) Decreate (remove wax);
(11) Hydrate;
(12) Stain;
(13) Dehydrate;
(14) Clear;
(15) Mount cover slip with resin.
(Note that steps 2 - 10 repeat the steps discussed under Embedding, Sectioning, and Pasting.)

Although it takes only a few minutes each day to transfer objects from one chemical to the next, expect to spend several days preparing a set of tissue sections. As new objects are found, they may be started along the process while other objects are moved to more advanced stages. The liquids can be used several times before becoming so contaminated as to be useless.

Moving very small objects can be tricky. One way of doing this is to tie them into a small piece of nylon cloth, and move this bag through the different containers of chemicals. A good collection of small vials and test tubes with stoppers is inexpensive and handy to have. The end of the string used to tie the nylon bag can be left outside stoppered test tubes.

Frozen sections

Advanced methods of making frozen sections have been developed. Sections prepared in this way can be as good as traditionally fixed materials. These methods require quick freezing of specimens to extremely low temperatures using specialized equipment including freezing microtomes. Electron microscopists also use many advanced freezing techniques. Frozen sections made in a more basic laboratory or at home are inferior, but if time must be compressed they are useful.

Optionally embed the object in gelatin or sandwich it between pieces of raw potato or carrot. Place the object in the freezing compartment of a refrigerator, or into a container with dry ice. Because the cell contents are disrupted less with quicker freezing, dry ice is a better choice if available. When frozen, use a razor blade and quickly slice thin shavings from the object. You do not have time to place the object in a common microtome. Put the object back into the freezer if it should begin to melt.

The sections can be examined immediately using a drop of water and a cover slip. For a permanent preparation, drop the shavings into fixative, paste, stain, dehydrate, clear, and mount.

An improved method for frozen sections is to use drops of ethyl chloride to freeze the object to a block and to keep the object frozen while the block is being fitted to a microtome. The ethyl chloride is dripped onto the block as sections are cut.

WARNING: Ethyl chloride is dangerous if breathed; use it in a well ventilated space, and use eye and skin protection. It is probably best used only under laboratory conditions, and not at all by young people.

Slides and Cover Slips

The figure shows why a drop of oil is used between the oil immersion objective and the cover slip. Light travels at different velocities through different mediums. A medium's refractive index is the speed of light traveling through a vacuum divided by the speed of light traveling through the medium. Light bends most when it travels between mediums that have very different refractive indices. Air has a much different refractive index than glass, while oil has a refractive index that is about the same as glass. The illustration shows how light is lost through refraction when it moves from the cover slip into the air, and how the light continues to move in an approximately straight line when emerging into oil.

All of the mounting media discussed also have refractive indices that are significantly closer to glass than to air. If the slide is oiled to the condenser, even less light will be lost; light will no longer bounce around between the condenser lens and the slide. Type A oil is used between the cover slip and the objective. The more viscous type B oil is used between the slide and condenser. (Check with the microscope manufacturer before oiling the slide to the condenser. There are a few economy microscopes that can be damaged by this practice.) Think of the slide, mounting medium, cover slip, and oil as an integral part of the optical system.

The required cover slip thickness is recorded on the objective's engraving, as illustrated in figure 1.10. Most oil immersion objectives are designed to work with coverslips of .17 mm thickness. Here are the average thicknesses of cover slips as they come from suppliers:
#1: .13 to .17mm
#1 1/2 .16 to .19mm
#2 .19 to .25mm.

Because number 1 1/2 coverslips average between .16 and .19 mm in thickness, they are best for objectives that require .17mm thickness. Oddly, many suppliers no longer carry this thickness. When the correct size cannot be found, the thinner slip is usually the best choice.

Just as objectives are set for a coverslip thickness, condensers are set for a slide thickness. This thickness is usually not marked on the condenser. Focusing condensers can adjust to different slides easily.

Experiments can be made with lenses and magnifiers beneath fixed focus condensers. You can also try slides of several thicknesses if the microscope has a fixed focus condenser, and stick with the kind that works best for each illumination setup. It is worthwhile to experiment with doubled slides oiled together, which raise the specimen in relationship to the condenser's focal plane. The specimen can be placed between two large cover slips without using a slide at all to bring it very close to the condenser. The thickness that works best for darkfield, Rheinberg, and offset illuminations may not be the one that works best for other illuminations. Whether the slide is oiled to the condenser also makes a difference.

Micromanipulation

When it is necessary to see both sides of an object, it can be placed in liquid between two cover glasses. This cover glass sandwich is placed on the slide for viewing, then flipped. For high magnifications the sandwich can be oiled to the slide and the slide oiled to the condenser.

Devices for working with small objects are available. These include manipulators, syringes, drills, hammers, and other tools attached to gearing that allow very fine movements. In fact, it is possible to gear controllable movements that are smaller than the limit of resolution of any light microscope.

A common micromanipulation experiment for beginning microscopists is to cut apart hydras or planarian worms. Both of these organisms can regenerate whole organisms from cuttings. A small utility knife or razor blade can be used for this. Hydra parts can also be pushed together on a needle to form double headed hydras and other grafted monsters.

4: Records

The microscope can be used entirely for pleasure, but any scientific or educational use requires careful record keeping and measurement. Because the microscope is a visual tool, you must take notes, at least in part, graphically. Drawing, photomicrography, and video photomicrography can be used to record the image.

Drawing

Before rejecting the idea of drawing out of hand, be advised that the kind of drawing being discussed is not an artistic feat. A knowledge of perspective is rarely required. A closer analogy is drawing a map or making a diagram.

Drawing has some inherent advantages over photography through the microscope. A single drawing can depict structures that appear at a number of different depths within the specimen -- most of which would be out of focus in a photograph. Structures seen under different illumination setups can be included in a single drawing. Things of importance can be stressed in a drawing that might appear only vaguely in a photograph. Many microorganisms are so simple that it will only take a few seconds to complete a drawing of them.

The necessary materials are paper and any writing tool. Optional materials might also include a soft pencil, compass or circle template, a set of colored pencils, and a technical fountain pen.

First draw a circle about four inches in diameter to represent the visual field as seen through the eyepiece. Then, with faint lines, draw the outlines of the structures that you see. With a slightly heavier hand, go over the outlines, correcting them to more accurately correspond with the specimen. Shading is not usually necessary. You may sometimes want to go over the outlines in ink and make some loose color indications with colored pencil.

Find a book or article that explains the structures in your drawing. Add labels and arrows naming the structures as explained in the reading. Write a few lines near the drawing explaining what was seen. Make revisions in the drawing based on the reading. Could you see all of the structures that the article mentioned? If not, try different illuminations and stains. Try searching for other specimens that will display specific structures better. Do not be discouraged if you are never able to see everything shown in an article's illustrations. The researches who wrote the article may have had access to an electron microscope, a specialized microscope, or specialized techniques.

Camera lucida attachments are available to fit many microscopes. These devices allow the eye to see both the specimen and a superimposed view of the drawing paper. Drawing with one of these devices is a simple matter of drawing around the superimposed outlines with a pencil. An eyepiece reticle with a large grid pattern covering the entire field of view can also be used as an aid, and is more economical. Each square of the field of view can be reproduced in pencil on grid paper.

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