CR Scientific: Articles: How to Make Your Own Slides for the Microscope

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Prepare Your Own Microscope Slides
by Christian Thorsten

Synopsis: Some methods for making permanent mounts are examined, with focus on commonly-available materials.

Introduction: Making permanent slides is a useful skill for the teacher or the aspiring scientist, veterinarian, or physician. The experimenter should have access to methods that don't require expensive equipment (rotary microtomes, etc) or the more hazardous reagents. This article will focus on such simplified, reduced-hazard methods, although formalin is mentioned because it is so useful and difficult to substitute. The reagents used may have some flammability hazard, but their toxicity is unremarkable (with the possible exception of formalin). Compared to osmium tetroxide and perhaps a few other fixatives used in the professional lab, however, even formalin is no cause for alarm as long as safety precautions are followed.

Before we continue, let's examine some important definitions:
Hydrophilic ("loves water"): mixes well with water or water-based liquids. Includes alcohol, glycerin, vinegar, lactic acid, formalin, and polyethylene glycol.
Hydrophobic ("avoids water"): does not mix with water or water-based liquids. Examples: xylene, benzene, kerosene, heptane, vegetable oil, mineral oil, etc. Longer-chain alcohols are hydrophobic; butanol and pentanol, for example, are only partly soluble in water. Octanol is essentially insoluble in water.

Hydrophobic molecules are sometimes called "oil-soluble". There are a couple of oil-soluble stains which, predictably, are used to stain fats and fatty deposits in cells.
Amphipathic: refers to compounds that have both hydrophobic and hydrophilic functional groups in the same molecule. Some of these are fully soluble in both water and oils, a feature that has practical importance for slide-making.

Materials: Because it is useful for the student to experiment and learn the uses and inherent limitations of various microscopy reagents, one should acquire a variety at the outset. These may include:
Corn Syrup (preferably high-fructose; it won't crystallize)
Glycerol (glycerine)
Acetic Acid, 5% (vinegar)
Acetone
Alcohol, denatured (usually contains methanol, a poison)
Alcohol, ethyl, 95% (neutral grain spirits U.S.P.)
Alcohol, isopropyl (2-propanol; "IPA"), 70% and 91%
Biological stains (crystal violet, safranin O, hematoxylin, methylene blue, etc)
Buffers (e.g., phophoric acid + phosphate; citric acid + citrate; "Tris"; etc) (optional)
Formalin (30-40% formaldehyde solution; poisonous)
Gum Arabic (used to make "Apathy's Syrup", a mounting medium)
Lacquer, clear gloss (the kind used for wood surfaces)
Lactic Acid
Paraffin wax (used mainly for embedding and sectioning, covered elsewhere)
PEG (polyethylene glycol) 300 or 400 (both are liquids down to about 5°C)
PEG-4000: a solid at room temp; melts at about 54°C (129°F)
PEG-6000: a solid at room temp; melts at roughly 60°C (140°F)
Nail polish (clear / colorless)
Xylene ("Xylol")

Methods:

1. Specimen choice: The easiest type of specimen to use is that which does not require thin-sectioning (we'll save that for another article). Possible choices include:
• Algae and other organisms (from a pond or puddle)
• Blood, smeared out very thinly with a needle or coverslip and allowed to air-dry
• Bread mold
• Cheek cell scrapings (i.e., epithelial cells)
• Nasal swabs
• Protozoa (from a pond or from hay infusion)
• Saliva
• Tooth scrapings (i.e., tartar)
• Urine sediments (obtained by centrifugation)
• Yogurt (the home of several benign species of bacteria)

2. Fixing: Specimens are treated with a fixative to kill the cells and preserve them from auto-degradation and breakdown by bacteria / mold / yeasts.
To accomplish this we might use one of the following fixatives:
• Alcohol, 90-96%
• Acetic acid, 5% (vinegar)
• Formalin diluted 1:10 (1 part formalin + 9 parts distilled water).
• Lactic acid diluted 1:10 (1 part lactic acid + 9 parts distilled water)
• Copper acetate or copper sulfate, a 2 to 5% solution in distilled water
More advanced practitioners use chromic acid, lead salts, uranium salts, picric acid, or other materials; however, the above-listed ones should suffice in many applications. Copper is mildly toxic but has enough in common with the so-called "toxic heavy metals" that it makes a decent fixative-- while being nowhere near as toxic as mercuric chloride or osmium tetroxide.

Depending on the thickness of the specimen and the type of fixative, the time required may range from as little as 15 minutes to as much as 2 days. Acetic and lactic acids will ruin the specimen if left in contact for too long; they hydrolyze proteins and will dissolve red blood cells. Alcohol fixation takes up to 12 hours; formalin may require 24 to 48. This writer has found heavy metals to work in as little as 15 to 20 minutes, especially when dissolved in 5% acetic acid.

After the desired time has elapsed, the excess fixative is removed by soaking the slide in tap water and then in distilled water.

3. Staining: Since most stains are water-based, this is the time to stain the specimen (i.e., after fixing but before dehydrating). Stain helps visualize certain features that might otherwise be colorless and transparent.

The desired stain formula is applied to the fixed specimen. This can be accomplished by immersing the whole slide in stain, but it is more economical to place a few drops of the stain on the specimen and let it sit for some period of time (anywhere from 15 minutes to several hours) (fig. 1). The staining time depends on (A.) the stain being used, (B.) the type of specimen being stained, (C.) how the specimen was "fixed", and (D.) the desired stain intensity in the final specimen.

Staining can be finicky and is often dependent on pH and other factors. Sometimes a pH buffer is used to ensure uniformity; the choice of buffer depends on the optimal pH for the stain being used.

3a. Destaining. After the desired amount of staining time has elapsed, the specimen is washed gently: first in tap water, then in distilled water, and finally in 70% isopropyl alcohol ("IPA").

Figure 1. Applying stain directly to the slide is advisable when only one or two slides are being prepared at a time.
Shown here: the "Five-Minute Food Color Stain" being applied to some epithelial cells on a slide. Staining time: approx. 15-20 minutes.

4. Dehydration: After treating the specimen in 70% IPA, it is then soaked in 91% IPA for 20 minutes to an hour. After draining off the excess 91% IPA, the specimen is treated with a few drops of 100% (i.e., anhydrous) isopropanol for 20 minutes to an hour. The excess is then poured off and the specimen allowed to dry at room temperature.

An alternate route to dehydration might be treatment with successively more concentrated solution of glycerin, PEG, or acetone. All these of these have some dehydrating action. PEG and glycerin can also be used as actual mountants in the final step.

Note that complete dehydration is not necessary if the specimen will be mounted in a hydrophilic medium (e.g., glycerin, PEG, or corn syrup). Stopping the step 4 treatment at 91% IPA is acceptable.

5. "Clearing": The dehydrated specimen is covered with a few drops of xylene and let stand for 20 minutes.

This step is skipped if a hydrophilic mountant is used.

6. Mounting: One drop of mountant is applied to a clean coverslip, and a couple drops are applied to the specimen. The coverslip is pressed slowly but firmly down on the specimen.

The best mountants typically have refractive index between 1.45 and 1.55.

Clear nail polish or clear wood-lacquer can be used as mountants (others have suggested nail polish previously. When shopping for microscopy mountants, one begins to scan store shelves for any kind of resin or varnish that can solidify and might have a reasonably high refractive index.) Both nail polish and wood-lacquer eventually harden and produce a good, permanent mount. Water-clear epoxy is another choice; it is miscible with alcohol while still wet. It is also the most difficult to remove once it is set.

Instead of lacquer or nail polish, one might use a non-setting, hydrophilic mountant. Corn syrup, PEG-300, and glycerin are three examples. In this case, the outside edges of the coverslip are sealed with several coats of lacquer, nail polish remover, or epoxy to seal in the liquid. Alternatively, a higher-weight PEG can be used (e.g., PEG-6000); it can be melted on a slide heater above 60°C, the coverslip pressed into place, and the specimen allowed to solidify at room temperature. In this case it is wise to choose a PEG that will remain solid even if left in an unventilated car in the summertime (120+ °F); after all, one might want to show the slide collection to a colleague or friend.

A traditional, hydophilic mounting medium is Apathy's Syrup (Apathy's gum-syrup). The basic recipe is mentioned in Locquin and Langeron (1983), but it goes back at least a hundred years. The original formula used sucrose, but this tends to crystallize over time, causing destruction of the specimen. High fructose ("levulose") corn syrup, on the other hand, doesn't crystallize.

Gum arabic.................................10 g

High fructose corn syrup.............10 g

Water.........................................10 mL

Phenol or camphor....................trace (as a preservative)

When ready to use, dilute 1:3 with water. Place on the specimen. Allow to evaporate until highly viscous, then cover specimen with a coverslip. The formulation will eventually dry completely at the edges of the cover-slip.

Depending on the mountant and the specimen, it may be advisable to add some kind of preservative to the mounting medium (e.g., sodium benzoate, iodoform, camphor, thymol, formalin, or phenol). This is more likely to be necessary when using hydrophilic mounting media in specimens that have not been dehydrated.

7. Labeling: This is just as important as the other steps, because it's all too easy to forget just what procedure you followed. If something works well, write it down. If something doesn't work well, write it down. It is difficult to fit all the information on slide labels, although a fine enough pen and a steady hand can accomplish a great deal (figure 2). The author uses adhesive paper labels and writes on them with a 0.20 mm drafting pen. After the ink is dry they are coated with two or three successive coats of clear gloss, fast-drying polyurethane varnish.

Figure 2. A slide made with the above steps. Notice the small arrow affixed to the coverslip. This points out the most interesting feature so that it can be located quickly. The arrow was positioned using forceps and 100x magnification.

8. Storage: After all that work, keep your slides in a slide box so they don't get broken. Keep prepared slides out of the light and away from extremes of heat or cold. (Glass and various mounting media have different coefficients of thermal expansion. Under certain conditions this can lead to cracked coverslips and ruined specimens).
Study the prepared slides over time to see which formulae and methods are the most color-fast and otherwise durable. Certain mounting media, for example, are acidic (Canada balsam comes to mind) and will lead to unwanted changes over prolonged periods. A properly dehydrated specimen mounted in a neutral, hydrophobic medium should last for at least a few years and may last for decades.

References:

Locquin, M. and Langeron, M. Handbook of Microscopy. London: Butterworths, 1983.

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