Yeast cells in the hemocytometer. The grid is clearly visible.

Yeast cell suspension applied to the chamber. Notice that some of the cell suspension has gone into the overflow area.

One counting chambers has grids of different sizes. Consult the manual to find out the size.

Do not count cells on the top and right lines. Here it's necessary to count the in the big square because there are too few cells in individual small squares.

Counting chamber seen from the side.

Grid layout of the Neubauer Improved hemocytometer.

Purpose of the hemocytometer

The hemocytometer (or haemocytometer or counting chamber) is a specimen slide which is used to determine the concentration of cells in a liquid sample. It is frequently used to determine the concentration of blood cells (hence the name “hemo-”) but also the concentration of sperm cells in a sample. The cover glass, which is placed on the sample, does not simply float on the liquid, but is held in place at a specified height (usually 0.1mm). Additionally, a grid is etched into the glass of the hemocytometer. This grid, an arrangement of squares of different sizes, allows for an easy counting of cells. This way it is possible to determine the number of cells in a specified volume.

Preparing the sample

The fluid containing the cells must be appropriately prepared before applying it to the hemocytometer.

• Proper mixing: The fluid should be a homogenous suspension. Cells that stick together in clumps are difficult to count and they are not evenly distributed.
• Appropriate concentration: The concentration of the cells should neither be too high or too low. If the concentration is too high, then the cells overlap and are difficult to count. A low concentration of only a few cells per square results in a higher statistical error and it is then necessary to count more squares (which takes time). Suspensions that have a too high concentration should be diluted 1:10, 1:100 and 1:1000. A 1:10 dilution can be made by taking 1 part of the sample and mixing it with 9 parts water (or better saline of correct concentration to prevent bursting of the cells). The dilution must later be considered when calculating the final concentration.

Counting the cells

• Counting cells that are on a line: Cells that are on the line of a grid require special attention. Cells that touch the top and right lines of a square should not be counted, cells on the bottom and left side should be counted.
• Number of squares to count: The lower the concentration, the more squares should be counted. Otherwise one introduces statistical errors. How many squares? To find out one could calculate the cell concentration per ml based on the numbers obtained from 2 different squares. If the final result is very different, then this can be an indication of sampling error.

Calculating the cell density

Here it is necessary to do some simple math. The following numbers are needed: number of cells counted in a square, area of the square, height of the sample, dilution factor. The objective is to find the number of cells in 1ml of original solution.

• Step 1 – Averaging: If one did not count all of the cells in a large square (1mmx1mm) then it is necessary to average the results first before proceeding. For the purpose of this example, I use an average cell count of 123.456 cells.
• Step 2 – Computing the volume: It is necessary to determine the volume represented by the square. The width and height of the square (e.g. 0.25mm x 0.25mm) must be multiplied by the height of the sample (often printed on the hemocytometer, in this example it is 0.1mm): v = 0.25mm x 0.25mm x 0.1mm = 0.00625mm³ = 0.00625ul (where ul is microliters).
• Step 3 – Calculating the number of cells in 1 ml: if there are 123.456 cells in 0.00625ul, then how many cells are there in 1ml (=1000ul)? We do simple direct proportion:

  123.456cells/0.00625ul = X/1000ul
  (123.456cells*1000ul)/0.00625ul = X (the ul cancel out)
  X = 19 752 960 cells

• Step 4 – Correcting for dilution: If the sample was diluted before counting, then this must be taking into consideration as well. We assume that the sample was diluted 1:10. The final result is therefore 19 752 960 cells x 10 = 197 529 600 cells in 1 ml. That a lot of cells.

Things to watch out for

• Type of counting chambers: There are different types of counting chambers available, with different grid sizes. One counting chamber also has grids of different sizes. Take care that that you know the grid size and height (read the instruction manual) otherwise you’ll make calculation errors.
• Use the provided cover glasses: They are thicker than the standard 0.15mm cover glasses. They are therefore less flexible and the surface tension of the fluid will not deform them. This way the height of the fluid is standardized.
• Moving cells: Moving cells (such as sperm cells) are difficult to count. These cells must first be immobilized.
• Objective The hemocytometer is much thicker than a regular slide. Be careful that you do not crash the objective into the hemocytometer when focusing.

Ranunculus pollen mounted in air with cover glass.

Ranunculus pollen mounted in water.

Ranunculus pollen mounted in Euparal. The pollen grains started to shrink.

Ranunculus pollen mounted in clear nail polish. The pollen grains show signs of significant shrinkage.

The mounting medium can have a significant effect both on the image quality and on the specimen itself. I tried a little experiment by observing pollen from a plant (in this case the buttercup, Ranunculus), mounted in five different ways:

• Air-mounted, with no cover glass
• Air-mounted, with a cover glass
• Mounted in water (temporary mount)
• Mounted in Euparal medium (permanent mount)
• Mounted in nail polish (permanent mount)

All observations were made using a 20x achromatic objective.

Results

The images on the right show that the mounting method has a significant impact on the way that the pollen grains appeared. The results can be summarized as follows:

• Air-mounted specimens show the least details. The pollen grains show a thick dark fringe, which covers much of the details. This is due to the large difference in refractive index between the pollen grains and the surrounding air. Opening the condenser diaphragm reduces the dark fringes, but also lowers contrast and depth of field. The cover glass presses the pollen against the slide, so that more of them are in focus. Otherwise the cover glass did not seem to make much difference.
• The water-mounted sample provides a much better image. The dark fringes are now gone, due to the similar refractive index of the pollen and the medium. The pollen appear spherical, because the water causes them to swell up.
• Pollen mounted in Euparal started to shrink and therefore appear smaller in size. Kinks and folds are also visible. These artifacts are produced because the (non-water based) Euparal has withdrawn moisture from the pollen.
• Clear nail polish showed a similar, but more pronounced effect as Euparal. The deformations of the pollen are very clearly visible. Evidently the solvent of the nail polish also removed significant amounts of water from the specimen. The nail polish itself lost some of its volume during drying and started to shrink as well. Air bubbles also became visible in the nail polish. Irregular drying of the mounting medium and a change in the shape of the mounting medium during drying can lead to shear-forces, which may distort the shape of the specimen.

What about Glycerin Gelatin (glycerol gelatin, jelly)?

Glycerin Gelatin is a water-based mounting medium. Glycerin Gelatin according to Kisser is one of several Glycerin Gelatin variations. It is a common medium for mounting pollen. Due to its water-based nature it does not cause the pollen to shrink. I’ll add a picture of this, when I have some of this mounting medium available. An alternative water-based mounting medium is fructose syrup. Both Glycerin Jelly and fructose syrup do not dry completely and therefore require a sealing of the sides of the cover slip with nail polish (but the pollen do not touch the nail polish).

Lessons learned

What can we learn from these observations?

• First, permanently mounting a specimen is not only important for slide storage. The mounting medium significantly influences the transparency, resolution and shape of the specimen.
• Second, the choice of the mounting medium depends on the type of specimen to be observed and on the type of microscopic technique to be used. For phase-contrast work the refractive index of the mounting medium should be different from the refractive index of the specimen. For bright-field work the refractive indexes should be similar. Large differences in refractive index can lead to the dark fringes as seen in the air-mounted specimens.

Some philosophy

So which mounting medium now results in pollen grains with a “true” or “correct” shape? The problem now is: what is the “correct” shape? Biological specimens may change their appearance depending on the environment. After a rain shower, the pollen may have a more roundish appearance, after having osmotically absorbed much liquid. Pollen that has dried in the air may resemble more the shape of the Euparal and nail polish samples. The choice of the mounting medium may therefore even include these considerations.

External Links, References

• An introduction to pollen analysis
• Aqueous Mounting Medium Protocols
• Making and Using Aqueous Mounting Media

Requirements of the specimen

When microscoping with children, I recomend the

• Not too abstract: The specimen should not be too abstract for the children. I mean, YOU may be interested in the circuitry of computer electronics parts under the microscope (and they DO look interesting), but for kids I’d suggest something more tangible.
• Flat: A flat object makes it easier to adjust the depth of field. Most of the object will then be in focus.
• Contrast: A high contrast makes it easier to see structures and details.

Specimens to look at

• Safe: This is self-explanatory. Do not use organisms or substances that are hazardous.
• Rocks: Collect some smooth rocks, wash and clean them in running water. Either observe the rocks while they are wet (and still shiny) or make them shiny by polishing them with a drop of oil. Shiny rocks have more contrast and simply look better than dull ones.
• Leaves: They are flat and transparent. They can be observed both with the light source from the top and from the bottom.
• Insects: This can be problematic. The insects should be dead, otherwise they are too difficult to observe, moving around all the time. Be aware that catching insects (such as butterflies) may not be allowed, as some of them are protected.
• Foods: Cornflakes, cut open fruits, seeds, mushrooms can make very educational samples. Place the cut surface is horizontally under the stereo microscope, and you won’t have a depth of field problem.
• Money: Count the scratches on the coins! The highly reflective surface of the coins make them an easy specimen.
• Pictures: This is the first specimen that we use in school when teaching the students how to use the stereo microscope. Printed pictures are made of many dots, which can be observed. Many kids did not know this. This way the children learn how to focus properly and how to change magnification. Later we give them specimens with a thickness.
• Own fingers: Here it is important to instuct the children to rest their fingers on the platform of the microscope. Many children will attempt to view their fingers by holding them mid-air beneath the objective. It is nearly impossible to find a proper focus this way.
• Own handwriting: This is a good possibility to estimate size and magnification.
• Textiles: stretch them flat and observe how they look different in epi- and trans- illumination.

Specimens not to look at

• Dust: some kids may have a dust allergy (mites), but it depends on the type of dust.
• Body fluids (blood): for hygienic reasons. And they are not interesting anyway at a low magnification.
• Spoiled food: fungal spores are not healthy to breath in, and bacteria on food are not good…
• Anything else which can be considered dangerous

Why bacteria are difficult to see

Bacteria are difficult to see with a bright-field compound microscope for several reasons:

• They are small: In order to see their shape, it is necessary to use a magnification of about 400x to 1000x. The optics must be good in order to resolve them properly at this magnification.
• Difficult to focus: At a high magnification, the bacterial cells will float in and out of focus, especially if the layer of water between the cover glass and the slide is too thick.
• They are transparent: Bacteria will show their color only if they are present in a colony. Individual cells present on the slide are clear. Regular bright-field optics will only show the bacteria if one closes the condenser iris diaphragm. This is due to the difference in the refractive index between the water and the bacterial cells.
• Difficult to recognize: An untrained eye may have problems differentiating bacteria from small dust and dirt which is present on the slide. Some bacteria also form clumps and therefore it is difficult to see the individual cells.

Research organizations and advances amateurs use phase contrast optics to see bacteria. This system converts the differences of the refractive index of the bacteria into brightness. The transparent bacteria can then be seen dark on bright background. In bright-field, closing the condenser iris diaphragm will also make the bacteria appear darker, but at the same time one also introduces artifacts (“fringes”) around the individual cells. One possibility is to stain the bacteria, but in this case there fixing and staining process may introduce artifacts.

What is a safe source of bacteria? For recreational or educational purposes, one should never use spoiled food or (heaven forbid!) use bacteria obtained from the human body and grown on agar plates. The risks involved are simply not worth it, especially when working with students. Other sources, such as soil or humus have other disadvantages. The impurities make it difficult to keep bacteria from other particles apart, especially if one uses bright-field optics. Rather I recommend the use of yogurt. It should be possible to see small circular cells (cocci), which may also occur in pairs. It is also possible to scratch some bacterial cells off from certain kinds of cheese. Brevibacterium can be found on Limburger cheese, for example. One has to be aware that some cheeses use a combination of bacteria and fungi, however, and that the larger fungal cells may outweigh the bacteria.

In summary, there are easier (and maybe also more interesting) specimens to observe than bacteria. I you want to see individual cells, then I do recommend that you start out with yeast suspensions. These eukaryotic cells are much larger and can be more easily identified.

The general procedure of making a wet mount

1. Place a drop of water on the center of the slide. It is also possible to first place the specimen on the slide, but small specimens usually separate more easily from the tweezers or needle if dipped into the drop of water.
2. Place the specimen into the drop of water and if the specimen floats, add another drop of water on top of it. This reduces the possibilities of air bubbles forming.
3. Carefully lower the cover glass so that it touches with one side the drop of water. The cover slip should form an angle of about 45 degrees with the slide. Touch the cover glass on the sides only to prevent finger prints. Alternatively, use tweezers to hold the cover glass.
4. Then lower the cover slip completely. Placing the cover slip at an angle prevents the formation of air-bubbles.
5. Remove excess water with a filter paper or tissue paper

Possible problems of making a wet mount

• The cover glass floats and moves: This is due to too much water. Remove water with the help of a tissue paper. Under no circumstances should there be water droplets on top of the cover glass. This water may get into contact with the objectives.
• The liquid streams and does not settle: This could be due to evaporation. Add more water between coverslip and slide.
• Air bubbles start to become visible: If bubbles were not present before and start to form, then this could be an indication of oxygen production due to photosynthesis. This depends on the oxygen saturation of the water and the amount of photosynthetic algae present.
• Air bubbles are present: Often the cover glass was not lowered from the side at an angle, but placed horizontally on the water drop. It may also be that the the specimen is hydrophobic (fatty) and /or fluffy. In this case, the the water may have problems reaching all of the areas of the speciemen and there is much air caught by the fine structures. Wet the specimen briefly in alcohol and then transfer directly from the alcohol to water. Alternatively you can try to break the surface tension of the water by adding a small amount of surfactant, such as soap or shampoo. Be aware that alcohol or soap may have adverse effects on living organisms.

There are several methods of preparing pollen grains, each one offers advantages and disadvantages. I can not give a general rule, it simply depends on the goal of the investigation and on the sample investigated. Pollen from wind-pollinated plants taken from a dry environment are probably best left in a dry condition, and not mixed with a water-based mountant, which may cause them to swell (depends on the osmotic potential of the medium, however). On the other hand, the obtained image quality and resolution may not be satisfactory in such a dry mount. It is a compromise, in which several factors have to be taken into consideration. A microscopy enthusiast, who does not need the slides for identification purposes, will again set different standards (such as avoidance of toxic solvents). People who want to publish their results, in turn, may have to rely on the preparatory technique which is customary in their field of research, for reasons of comparison. I recommend that the different methods are tried out.

Mounting techniques

Glycerol wet mount: Place a small drop of glycerol on a clean slide and tap the anthers of the plant so that the pollen falls into the glycerol. If necessary, carefully separate large chunks of pollen grains by stirring. Place a cover slip on top and seal the sides of the cover slip with nail polish. Use a very small amount of glycerol to make sure that the nail polish has enough area to stick the coverslip to the slide. Glycerol wet mounted slides can be stored for months if there is no leakage. The glycerol will withdraw water from the pollen. If the pollen is not dry, then there is a possibility of the pollen to shrink.

Air mounts (dry mounts): In this case, no liquid mounting medium is used. A cover slip is placed on top of the pollen grains and sealed on the side, either with nail polish or with tape. Nail polish may flow very quickly between cover slip and slide, so it may be best to use a nail polish of low viscosity (by letting some solvent evaporate first).

Glycerol jelly (according to Kisser): This is a very popular mounting medium for pollen. It is phenol-free (antiseptic additive) and therefore non-hazardous. It contains 10g of gelatin, 35ml distilled water and 30ml of glycerol (glycerin). After mounting, the sides of the cover slip need to be sealed. Due to the lack of an antiseptic, it is also necessary to work in a sterile manner, otherwise there is the risk of fungal growth in the medium. Maybe it is a good idea to treat the pollen grains first in alcohol to reduce the chance of fungal contamination by spores. Alternatively, one could experiment by increasing the concentration of glycerol.

Non-water-based mounting media: Euparal is a mounting medium which is not water based. Specimens which are present in alcohol can be directly transferred to Euparal. Place a pollen suspension on the slide and let the alcohol evaporate. Before mounting pollen in Euparal, I recommend that the pollen are first washed in alcohol and then compared to the original shape. Does washing in alcohol result in an unacceptable shrinking of the pollen or unacceptable loss of pigments? If not, then mounting the pollen in Euparal may be an alternative.

Reading materal

I found the following article: Marvels of pollen shown by your microscope (Popular Science, September 1939)
(The article recommends the use of organic solvents (such as xylol/xylene and others) to remove oil from the pollen. I do not recommend this due to health reasons, especially when preparing samples for educational purposes. Still, it gives a nice overview of the topic.)

Generally, mounting media for permanent slides can be categorized into water-based and organic solvent based mounting media. While many water-based mounting media for permanent slides solidify and hold the specimen firmly in place, some others remain in a liquid state. In this latter case, it is necessary to prevent the liquid from flowing out by sealing the four sides of the cover slip. Nail polish can be used for this. In the following paragraphs, I’d like to give you an overview of the different types of mounting media.

Water-insoluble mounting media that solidify

Euparal: This mounting medium was invented in 1904 by Prof. G. Gilson, Professor of Zoology at Louvain University, Belgium. It contains the substances sandarac, eucalyptol, paraldehyde, camphor, and phenyl salicylate. Euparal possesses a nice odor (but don’t smell it anyway), due to the natural oils that are included. Euparal is commonly used to mount histological specimens and insects. One big advantage of Euparal is, that the specimens can be transferred directly from the alcohol in which they are stored. Do not embed specimens which contain water, this may result in a clouding of the mounting medium.

Summary: Advantages of Euparal include the possibility to directly transfer specimens from alcohol to Euparal without the need of toxic solvents. A disadvantage is the relatively long drying time of a few days.

Canada Balsam: This is a natural mounting medium obtained from the e balsam fir tree (Abies balsamea). The optical properties are nearly identical with those of glass. For this reason, Canada Balsam was used for many years as a kit to hold optical lenses in place. Meanwhile, synthetic lens kits have replaced Canada Balsam, it is still used as a mounting medium for microscopy, however. Canada Balsam has the advantage that its optical properties do not deteriorate with age. Permanent slides mounted with Canada Balsam have been stored for a century and are still useful.

The disadvantage of Canada balsam is, that the specimen must be placed into xylene (toxic!) before embedding. Wet specimens must first be dehydrated in alcohol and then transferred to xylene. Transferring specimens directly from alcohol to Canada balsam won’t work, because the alcohol won’t dissolve the Canada balsam.

Summary: The advantage of Canada balsam is the long storage ability of the slides. Other, modern, mounting media may have a similar storage ability, but with Canada balsam there is historic experience. A disadvantage is the need for toxic solvents when preparing the specimen. Apparently, it is also not very cheap to obtain.

Eukitt and other resin-based media: Eukitt is a very fast drying general-purpose resin-based mounting medium. Eukitt will solidify within about 20 minutes. The specimens must be free of water and placed first in alcohol and then in xylene prior to mounting. The use of xylene is a disadvantage, as it is harmful when inhaled. Eukitt itself can also be diluted by xylene to adjust it viscosity.

Besides Eukitt, a range of other resin-based mounting media are commercially available, such as Diatex, Entellan, Malinol, Rhenohistol and Depex. They differ in their refractive index. All of these mounting media require the specimen to be first dehydrated in alcohol and then transferred to xylene. Some of these resins shrink significantly during the drying process.

Summary: The advantage of Eukitt is that it is a fast drying mounting medium. The disadvantage is the need for toxic solvents to prepare the specimen.

Clear nail polish: Nail polish can be used to seal the sides of the coverslip when using aqueous mounting media. It can also be used directly as a mounting medium. The specimens must first be dehydrated in alcohol and can then be directly mounted (without xylene) in nail polish.

Summary: The advantage of nail polish is, that it is readily available and that it avoids the use of toxic organic solvents to treat the specimens. One disadvantage is, that it seems to shrink a lot when making very thick mounts (such as whole insects).

Water-insoluble mounting media that remain liquid

While it is possible to use various oils (immersion oil and paraffin oil) as a mounting medium, they are generally not used to make permanent slides. The specimen must be dehydrated with alcohol and then transferred to xylene so that the liquid mounting medium (the oil) is able to reach all the parts of the specimen. I can imagine that it is this xylene which causes a problem with the sealing of the cover slip, by preventing hardening of the nail polish used for sealing.

Water-soluble mounting media that solidify

Glycerol jelly: This is a water-based (aqueous) mounting medium. There are several variations to the recipe, fine tuned for specific mounting applications. The classical recipe according to Kaiser (1880) includes Phenol as an antiseptic, so it hazardous for the use in schools and at home. The handling of this mounting medium, is also not too easy. The bottle with the solid glycerol jelly must first be warmed in a water bath to make it liquid. Do not make it too hot, otherwise it will not solidify any more. The specimen is submerged in the warm jelly and the cover glass is placed on top. Bubbles are a problem with this medium. The edges of the cover glass now must be sealed with nail polish to prevent drying out.

Glycerol jelly is one of the most difficult mounting mediums to use, but sometimes there is no other satisfactory alternative to an aqueous mounting medium. Water-based mounting media are useful for making permanent mounts of water organisms, algae, protozoa, etc. Glycerol jelly according to Kisser (not Kaiser) is commonly used to preserve pollen samples. Treating some specimens with organic solvent-based mounting media would cause them to shrink or change their shape in other unacceptable ways. Solvent-base media may also dissolve some of the pigments, such as chlorophyll, from the specimen, which does not happen when using aqueous media such as glycerol jelly.

Summary: The advantage of Glycerol jelly is that it s water-based and that this avoids the need of alcohol dehydration (which possibly deforms the specimens), and other toxic organic solvents. Some specimens can only be satisfactorily mounted in Glycerol jelly. It also does not shrink. The disadvantages include the need for a potentially toxic antiseptic in the jelly, the difficulty of mounting the specimens and the need to seal the cover slip with nail polish.

Water-soluble mounting media that remain liquid

Glycerol: It is possible to make a permanent mounts by embedding the specimen either in pure liquid glycerol or a specified glycerol-water mixture. The glycerol-water mixture can be adjusted to an appropriate refractive index. Adding more water lowers the refractive index. It is also possible to use pure water alone (for some delicate algae, for example).

Algae and other water organisms can be embedded this way. Algae that are embedded in pure glycerol may shrink because the glycerol withdraws water from the cells. If the algae shrink too much, then the glycerol should be more diluted with water. A high concentration of glycerol should be maintained, however, otherwise there is a risk of fungal growth in the medium.

Making liquid permanent slides is somewhat more advanced. The drop of glycerol must be very small so that it will not touch the sides of the cover slip. On all sides, there should be a few mm of air between the sides of the cover slip and the glycerol. The sides of the cover slip are then sealed with nail polish two or three times to prevent glycerol from leaking out. Here it is very important that the glass surfaces are completely clean and have not been in contact with glycerol, otherwise the nail polish will not hold.

Summary: The advantage of glycerol is, that fungi and algae do not shrink as much as with other mounting media. It is also not necessary to treat the specimens with alcohol or organic solvents, which may introduce artifacts and remove pigments. The disadvantage is, that it is difficult to prepare slides that are truly permanent in nature. A proper sealing of the cover slip corners is absolutely necessary if one wants to store the slides over extended periods.

Toxicity: Solvent-based mounting media (such as Eukitt and Canada Balsam) require the specimen to be in xylene prior to embedding. This substance is toxic. Other mounting media, such as Glycerol jelly, may contain hazardous antiseptics. This aspect of toxicity is something to consider when making permanent mounts either as a hobby or for educational purposes in schools. One should ask oneself, if one should not use other alternatives.

Refractive index: The correct refractive index (RI) of the mounting medium can be critical for seeing details of the structure. If one uses phase contrast microscopy, then the RI of the mounting medium should be very different from the RI of the specimen. For regular bright-field work with pigmented specimens, the RI should be the same. In an ideal world, the mounting medium should be matched with the type of specimen. For amateur or educational work, this may be of less relevancy, however. Some high-end microscope objectives are calibrated to be used for a specific RI of the mounting medium, otherwise the resolution is reduced.

Compatibility with specimen: Specimes which are kept in water should be transferred into a water-based mounting medium. Transferring them into a solvent-based mounting medium may result in a clouding of the resin. Likewise, specimens which are kept in alcohol should be transferred to xylene and then embedded in a solvent-containing mounting medium. Euparal allows the specimen to be present in alcohol.

Pigment stability: Some mounting media cause a fading of pigments and stains over time. If pigment stability is of relevancy, then one should use mounting media which do not react with the pigments of the specimen. In some cases a fading of pigments is desirable, however. This brightens the specimen and makes it more easy to observe.

Shrinkage: Some mounting media shrink when they dry. The effect is particularly noticeable when thick specimens (e.g. whole insects) are embedded. Non-water based mounting media are known to do this. Glycerol jelly, which is water-based, does not shrink, however.

Durability: How long should the permanent slides be stored? Non-solidifying mounting media may not hold the specimen in place very well and there is the risk of running out if not sealed properly. Other mounting media may start to crystallize over the years. Still others may adversely react with the pigments of the specimens. Canada balsam is known for its good durability.

Cost: Some mounting media (such as Canada Balsam) are quite expensive. Others can be made in the kitchen from readily available materials (Glycerol jelly).

Ease of use: Here we have to consider two aspects, the preparation of the specimen prior to mounting and the actual mounting process. Some mounting media require the specimens to be dehydrated and fixed before mounting (for resin-based media). This can be a time consuming process. During the mounting process, some media are more prone to form air bubbles (Glycerol jelly).

Link to the article: Microb hunting with your Microscope (Popular Science, Sept 1934)

• Common household chemicals: this includes Iodine, for example. They are very readily available.
• Substances used mostly for microscopy: Methylene blue, Hematoxyline, and Eosine belong to this group.
• Commercial substances: they are sold by companies specializing in microscopic chemicals.

The article also provides a step-by-step guide on how to stain a blood sample (don’t do this in schools due to danger of infection).

Link to the article: Help Your Microscope with Stains and Reagents (Popular Science, March 1937)

• Observing dust samples: Students should collect house-dust and bring it to class to be observed under the stereo or compound microscope. Careful, some people may be allergic to dust!
• Observing sand and soil samples: Students should collect sand and soil samples to be observed under the stereo microscope.
• Observing textile fibers: Observing various fibers obtained from clothing (cotton, polyester, nylon etc.). Different colors and textures become visible under the microscope.
• Which printer is the best? Students bring in print-outs of different pictures on different types of paper. The printing resolution can be observed under the stereo microscope.
• Observing water life: A large jar is filled with pond water and a little soil. Algae and other organisms will (hopefully) develop over the course of a few weeks. Do not let the water rot!
• Fungi from cheese: Camembert, Brie, etc. contain edible molds (not hazardous) and can be used. Much safer than rotting food and observing the molds.
• Vegetables and fruits: The teacher cuts the tomatoes and mushrooms in various ways, they can be observed under the stereo microscope. Do not eat the food afterward, you never know what chemicals were left behind on the microscope by previous classes…..
• Hair samples: Each student donates one hair and then they have to match them with the hair left behind on the “crime site”. This is a playful approach into forensics and gives the observation some purpose. Maybe a competition between different groups is also a nice idea. The teacher may have to prepare a set of permanent slides with some hair samples.
• Coins: Coins collect many scratches (and dirt) over the years. How can the scratches be quantified? Is it possible to predict the age of a coin by looking at the number of scratches? The year is imprinted in the coin.
• Observing human cheek cells: This is a classic, really. Using a cotton swab, some epithelium cells from the inside of the mouth are collected and transferred to a microscopic slide.

Things NOT to observe – Some specimens or samples should not be observed in a classroom setting:

• Spoiled food material: they contain hazardous bacteria and fungi. Spores are unhealthy to breath in.
• Body parts: Samples taken from wounds (pus etc).
• Blood samples or other body fluids.
• Urine: Some students (often boys…) may be interested in observing their own urine. Fresh urine should be free of microorganisms (unless there is an infection) and it is not an interesting sample to be observed.
• Animal wastes: Excrements of animals are prone to contain parasites and are a clear health hazard.
• Polluted water Water from polluted rivers, lakes may contain toxic substances and harmful microorganisms. Leave stuff like this to university-level students, who (should) know appropriate safety procedures.

Materials: A large glass jar, fresh and unfertilized garden soil, water, hot plate, celophane foil

Method:

• Fill the glass jar with a few centimeters of the garden soil.
• Add non-chlorinated tap water to the soil and fill the jar with the water (3/4 full).
• Boil the soil-water mixture for about 30 min. This will kill off bacteria in the soil and will extract nutrients from the soil. Bacterial spores may survive the boiling, as they are heat-resistant. This is not a problem, though. These bacteria will serve as a food for other microorganisms later on.
• Cool the water to room temperature and let the soil settle to the bottom of the glass jar. Do not filter the soil away. The soil will continue to supply nutrients and will act as a buffer.
• Add a small amount of pond water which contains algae. Do not add too many algae. You may want to scrape off some algae from rocks or take a few algal filaments floating in a pond.
• Cover the jar with celophane foil. This will allow for gas exchange and prevent dirt and dust falling into the water. It also reduces evaporation.
• Wait a few weeks for the algae and ciliates to develop. With a bit of luck, paramecia will grow and form white clouds in the water. The color of the water may also change, an indicator for algal growth.
• Store the jar in a bright place but not in direct sunlight.
• Using a pipette, extract some of the microorganisms to be observed under the microscope.

Troubleshooting:

• Microorganisms do not form: This is probably due to the fact that there were none or not enough in the pond water which was added.
• The water starts to smell bad: This may be due to the system becoming anaerobic. Make sure that enough oxygen is able to enter the water. Paramecia and other ciliates are probably dead by now…..

Method:

1. Cut the potato in half and scrape a little of the potato onto the microscope glass slide. This can be done either with a knife or with the fingernails. There should not be any large potato pieces on the glass.
2. Place a small drop of water on the “potato juice” and then place the glass cover slip on top.
3. Observe using the microscope. The starch grains will be visible as oval structures.
4. Now dilute a small amount of iodine in some water. The water should only turn slightly yellow. Place a drop of the dilute iodine next to the glass cover glass, so that some of the solution is able to flow between the cover glass and the slide.
5. You should be able to see how the starch grains change color. The iodine will react with the starch and turn it blue-black.
6. Alternatively, you can observe the starch grains in dark field or in polarized light (without adding iodine): Darkfield Microscopy | Simple Polarization Microscopy | Potato Starch Grains

Method:

1. Make sure that the specimen in completely dry. You may first place the specimen in alcohol to withdraw water, and then let the alcohol evaporate. Note, that this procedure may deform the specimen, however.
2. Stick a piece of the double-sided tape on the slide. The tape should have about the same size of the cover slip, or be slightly smaller.
3. Using the knife (not suitable for children!), cut out a square in the center part of the tape and discard this piece of tape. You should now have a square “frame” of double sided tape on the microscope slide.
4. Place the specimen into the center, it is now surrounded by the tape. The specimen should not be thicker than the thickness of the tape.
5. Place a cover slip on the tape and carefully (!) press the glass against the tape. The tape will hold the cover glass in place. You should not apply pressure to the center part of the glass slide, or it may break. You could roll a round pencil over the cover glass to press it against the tape.
6. Observe using low magnification. The specimen is not embedded in a mounting medium with an appropriate refractive index. The resolution of the image will therefore be lower at higher magnifications.

Suitable objects for dry mounting:

• Wings of insects
• Whole small insects
• Scales of butterfly wings
• Sand or soil particles
• Dust samples
• Dried skin, dandruff
• Different types of paper, etc.

A specimen for compound microscopy must fulfill several criteria:

• It must be sufficiently thin.
• It should not be too dark (too heavily pigmented).
• If it is not pigmented at all, then it should possess a different refractive index compared to its surrounding medium, otherwise the structure is invisible.
• It should possess sufficient color contrast.

What should one do if the specimens do not fulfill the above criteria? It depends on the type of specimen.

• Thin and strongly pigmented specimen: bleaching. Depending on the type of specimen, different bleaching methods can be used. It is also possible to remove some pigments (such as chlorophyll of plants) by immersing the specimen in alcohol.
• Thin specimen with low contrast: staining. Selective stains react differently with different parts of the specimens. Certain DNA stains (careful, potentially carcinogenic!) interact with the DNA and make nuclei visible. Other stains interact with other substances. Here it is necessary to consult a catalog to determine the right stain for the task.
• Thin specimen with low contrast: observing in phase contrast. Phase contrast microscopy is an optical method in increase contrast. A prerequisite is, that the specimen possesses a different refractive index than the surrounding medium, which is the case most of the time.
• Thick and soft specimen: squeezing. The specimen can be squeezed between the slide and the cover glass. One example of this method is the observation of various fruits, such as a soft kiwi.
• Thick and soft specimen: hardening followed by microtoming. Soft specimens (ripe fruits, soft leaves etc.) are often difficult to cut into thin sections. They have to be hardened first. Plant materials can be hardened by placing them into alcohol for a few days. This removes water and makes the object easier to cut into small slices. Be careful again, this method is not suitable for children, due to the sharp tools involved. Also note, that the removal of water by the alcohol may cause the specimen to shrink.
• Thick and hard specimen: softening. Certain specimens can be softened by boiling them. Alternatively, certain chemicals also achieve the same effect. The soft specimen can then be squeezed between the slide and cover glass before microscopic observation.
• Thick and hard specimen: grinding them thin. This method is sometimes used when observing rocks and other hard substances which can not be softened. Specialized tools are required.
• Thick and hard specimen: use stereo-microscopes. The easiest way is to observe them with a stereo microscope using epi-illumination (light from the top).

Materials: microscopic slides, cover glass, a soft kiwi, tissue paper.

Method:

1. Take a small piece of the soft part of a kiwi (not the seed and not the white center) and place it between the slide and the cover glass.
2. Carefully tap against the cover glass with a hard object, such as a pen. This is to test if the kiwi is actually compressible (or if it is not ripe enough). A hard kiwi may result in a broken cover glass.
3. Place a small piece of tissue paper on top of the cover glass and carefully and gently press down on the cover glass using your fingers (provided that the fruit is soft enough). Excess kiwi juice will be absorbed by the tissue paper. Be careful: do not move the cover glass horizontally. A thin, green kiwi film should have formed between slide and cover glass.
4. Observe under the microscope as normal.

Note: The image on the right shows some (unidentified) structures found in a kiwi fruit. The final picture is a stack of 12 images, processed with the program Combine ZP. Stacking of the images increases its depth of field. Without stacking, some of the green bubbles would not be in focus.

Fructose syrup is a water-based mounting medium, which is suitable for a wide variety of specimens. It is safe to use and it is easy and cheap to make. Spills can be easily washed out with water. One disadvantage is that the color of the specimens may fade and that some stains will loose intensity over time. This is due to the low pH of the medium. Fructose syrup is not suitable for making slides that last for many years, but is should be sufficient for classroom usage, where students would like to re-examine their specimens over and over again over a period of time. The medium will not completely solidify, so it is necessary to seal the cover glass at the side.

Materials: distilled water, fructose, dropper bottle or other container, optionally nail polish / nail varnish.

Method for making fructose syrup:

1. Fill several grams of fructose into the dropper bottle.
2. Using a marker, mark the level of the fructose on the glass bottle.
3. Using the dropper, add distilled water to the fructose. The fructose will dissolve and the volume will decrease. Add more water to maintain the total volume level.
4. Store the bottle for several days in a warm place, or use a warm water bath. It takes this time for all of the fructose to dissolve. At the end, you should have a clear, sticky liquid. It is then ready for use.

Method for using fructose syrup:

1. The specimen to be mounted (eg. a small insect, some plant sections etc.) must be first placed into water. In most cases, fresh material is already stored in water. It could, however, be that due to previous processing or storage the specimens are soaked in alcohol or other organic solvents. This solvent must be removed first. If the specimens were stored in alcohol, then slowly transfer them into distilled water by placing them gradually into more and more dilute alcohol. If you transfer the specimen directly from concentrated alcohol into pure water, then there is the danger that the specimen changes its shape.
2. Place a drop of the mounting medium on the slide, then place the specimen (not wet) into the drop. Place another drop of mounting medium on top of the specimen. The specimen is now surrounded by the medium from top and bottom. Finally, place a cover glass on top of the mounting medium.
3. Store the slide for a few days horizontally. Some water will evaporate, but the syrup will not solidify completely. If you store the slide for a long time (in a dry environment), then the fructose may start to crystallize out. You can then observe the specimen under the microscope.
4. Optional (careful, organic solvents involved!): Seal the corners of the cover glass with some nail polish (nail varnish). This will prevent the syrup from flowing out and will prevent moisture exchange. The slide should be stable for a few months.

Materials: milk, water, slide, cover slip

Method:

1. Dilute one drop of milk in about 5ml of water.
2. Place one drop of the dilute milk on the microscope slide and place a cover slip on top of it.
3. Observe under the microscope in bright field and in dark field. The microscopic fat droplets of the milk can be seen moving randomly. This is Brownian motion.
4. Change the temperature by placing the slide into the refrigerator or by warming it up. Observe again. The Brownian motion is temperature dependent.

Materials: a small amount of yogurt, water, hot plate, ink from a fountain pen or a marker, alcohol

Method 1:

1. Suspend a small amount of yogurt (tip of a knife) in a few ml of water.
2. Spread a drop of this suspension on a slide and let it dry completely at room temperature. Be patent here, do not accelerate the drying process by heating the slide.
3. Briefly place the dry (!) slide on the hot plate with the bacteria facing the top. If you “boil” the bacteria, then they may pop open and lose their shape, and will not accept the stain. So make sure that all the water is gone before heat-fixing.
4. Remove the slide. You should be just able to place the slide on your palm without burning yourself. If it hurts (or if you burn yourself) then the slide was heated too much and you have to retry and place a new suspension on the slide to dry. In this case the bacteria were burned and may have lost their shape. The heat treatment fixes the bacteria to the slide, so that they will not be washed off. In microbiological labs, the heat fixing process is usually conducted with a gas burner, but this may be too dangerous for schools.
5. Place a drop of ink on the specimen and wait for about 10 minutes. Carefully rinse the ink off by slowly pouring water or alcohol (depending on ink) over the slide. Continue this washing step until no more ink is given off, but do not over-wash. Also do not pour the washing liquid directly over the bacteria, but rather let it flow over it.
6. Let the slide dry.
7. If the bacteria are observed with oil immersion, then it is not necessary to place a cover glass on top of the sample. Instead place a drop of oil directly on the stained bacteria. This is only for experienced students, there is the danger that the wrong objectives are rotated into the oil….
8. The safest method would be to use water and a cover glass and to start observation with the low magnification objectives (In this case, of course, it is not necessary to let the slide dry after the washing).

Method 2: this method is easier and does not need a heat-fixing step.

1. Take a knife tip of yogurt and directly add 1-2 drops of water-based blue fountain pen ink. Do not use calligraphy ink. This type of ink is composed of suspended ink particles which can not be taken up by the bacteria.
2. There is no need for a washing step. The bacteria will accumulate the ink and will become darker than the surrounding medium.
3. Take a small drop and place on the slide for microscopic investigation. The drop has to be sufficiently small to form a very thin film between the slide and cover glass.
4. You should be able to see blue clusters of bacteria. Individual bacteria are probably too small to show a blue stain, but the diffraction pattern should make them visible. Some clusters may not have taken up the ink. The ability to take up the ink may be an indicator if the bacteria are still alive.

About the ink:

• Different types of inks contain different substances that may be more or less suitable for staining. I recommend you to experiment. The teacher could also try to dissolve some black or blue marker ink in some alcohol and then use this solution for staining. Inks used for calligraphy will most certainly not work. They contain suspended particles (carbon?) which are not able to enter the cells. Also be careful when using commercial stains. Some of them are designed to stain DNA and this is then not suitable for the use in schools (carcinogenic) – read the instructions that accompany the stain.
• If you use ink which is soluble in alcohol, then you may need to include a (brief) washing step with alcohol to remove excess ink. Experiment first.

Troubleshooting:

Problem: The unstained bacteria are not visible.
Solution: They are transparent, close the condenser aperture diaphragm all the way. You will then see diffraction patterns around the bacteria.

Problem: You see a blue mass but not individual cells.
Solution 1: The suspension was to dense. Dilute the suspension with more water, or if you directly observe the yogurt, make the drop smaller.
Solution 2: Remove more ink by rinsing longer.

• Electricity: Students should not be allowed to exchange light bulbs or perform other maintenance work. When plugging in the microscope, make sure that the main power is switched off and that the dimmer (light control) is set to “low”.
• Handling the Microscope: Hold the microscope with one hand on the arm (of the microscope) and use your other hand to support the base. Keep the cable wrapped around the microscope.
• Growth of bacteria: A simple rule – don’t grow bacteria in a school setting. Even if you work with a defined bacterial culture there are the chances of contamination and you never know what you are growing. Also note that there are certain laws that may apply for the growth of bacteria in areas that are not designated as microbiological labs. If you want to stain and bacteria then use safe sources, such as yogurt.
• To be continued….

When the glue has dried completely, carefully peel off the glue. It should separate easily from the leaf. The leaf has left an impression on the glue.

Cut the glue into shape using scissors and observe it with the microscope. If the glue is still water soluble after drying, then do not immerse the glue into water. The contrast is low, it is necessary to close the condenser aperture diaphragm.

Materials: Leaf of a plant, white wood glue (PVC glue etc., water soluble), slides, scissors.

Method:

1. Evenly spread a drop of water soluble wood glue on the bottom side of a leaf (the stomata are located on the bottom side).
2. Wait several hours or overnight for the glue to dry.
3. Carefully peel off the glue. It has become transparent.
4. Use scissors to cut the glue into shape and observe under the microscope. The leaf epidermis cells have left an impression on the glue, which can be observed.

Troubleshooting:

Problem: The glue does not want to separate from the leaf
Solution: Spread the glue on an even section of the leaf underside. Some leaves may have microscopic hair, which have become attached to the glue.

Problem: Nothing can be seen.
Solution: The contrast of this specimen is very low. You have to close the condenser aperture diaphragm to increase contrast.

Problem: The resolution is low.
Solution: This is due to the fact that the specimen (the dried glue) is not embedded in water and a cover glass is missing. Either make a permanent mount in with a non water based mounting medium or try to use glue which is not water soluble anymore after it has dried.

Issues to consider:

• Do not cover the whole underside of the leaf with glue, this will block gas exchange and may harm the plant.
• Do not use glue with organic solvents (acetone, alcohols etc.). This will possibly harm the leaf and these solvents withdraw water from the cells and dissolve the cell membrane. Or: try it anyway, maybe it still works… Take care that the glue does not contain solvents that are harmful when inhaled.

Things to try (I never tried them, success not guaranteed!):

• Spread the glue at night (do not turn on the lights) and compare the shape of the stomata with those during day. The stomata of the “daytime glue” should be open, the stomata of the “night time glue” should be closed.
• Compare the size and shape of the leaf epidermis cells of different plants.
• Does the size of the leaf have an effect on the number of stomata, on their shape?
• Approximately how many epidermis cells are there to one pair of guard cells?

Wikipedia explanation of stomata and guard cells.

Carefully tear off the layer of red cells. Remove the thick part of the onion (where the cut was made) and only observe the thin layer. Many cells will probably break open during this process and be useless, we only need a few intact cells.

The top image shows the cells before plasmolysis. The cells are filled with a red pigment and appear pink. The bottom image shows the same cells after the addition of saturated salt water. Intact cells will lose much of the water due to osmosis. The concentration of the pigment rises resulting in a darker color. The shape of the cell wall remains unaffected.

Materials: kitchen knife, red onions, salt, tap water, microscopic slides, cover slips

Method – Obtaining a single layer of red onion cells.
For this experiment, we can not use the onion skin which is found between the layers of the onion. We need a single layer of pigmented cells. These cells, however, do not separate easily.

1. We need a thin layer of cells of the red part of the onion. It is not possible to directly cut a single cell layer, so we need to use the “peeling method” to obtain a single layer of cells. Obtain a small piece of onion about (1cm x 1cm). The onion layer is about 2mm thick.
2. With the red side of the onion facing you, cut beneath the red layer, about half way into the onion. This cut does not have to be very thin. There will be about 1mm of onion between the knife and the red pigmented layer.
3. Press the onion firmly against the knife with your thumb.
4. Now tear off or peel away the red part of the onion. The red layer will become thin. Some red pigment may be released from broken cells.
5. Cut away and discard the thick part of the onion (the place where the initial cut was placed).
6. Observe the remaining cells (the thin, peeled part) under the microscope (using a glass slide, water and cover slip, of course.
7. Only consider those cells that are filled with the red pigment. White cells are broken and have lost the red pigment.

Method – Plasmolysis.

1. Make a saturated solution of salt-water
2. Using a pipette, add one drop of this solution to the specimen. The salt water should flow beneath the cover slip. There should be no need to remove the cover slip to add the salt water
3. If there is too much water beneath the cover slip, then the salt water will not flow between the cover slip and the slide. In this case use tissue paper to withdraw water from one side of the cover slip while adding the salt solution at the other side.
4. Observe what happens to the red pigment inside the cells.

Explanation: Water from the cells moves to the surrounding salt water. The shape of the cells does not change, the cell wall maintains the cell shape. The cell content (the red part of the cell) starts to shrivel up. At the same time it is possible to see that the intensity of the red pigment increases because it becomes more concentrated as water is removed (the red pigment is not able to move out of the cell). The process can be reversed when the salt water is removed and when distilled water is added.

Materials: Slide Projector (an overhead projector will not work!), slide frames with glass, onion, ruler, marker.

Method:

1. Cut out a piece of onion skin of about 1 cm² size. The onion skin is the membrane which can be found between the layers of the onion.
2. Using a ruler, draw a 1 cm long line on the inside glass of the projector slide. This is our internal reference.
3. Place the onion skin flat into the slide.
4. Project the skin on the wall and measure the length of 10 cells using a ruler. The students then calculate an average. Also measure the length of the 1cm reference line.
5. We now can calculate the magnification by dividing the length of the projected line by the original size: Magnification = length of projected line / length of original line E.g. if the projected line is 30 cm, then the magnification is 30 cm / 1 cm = 30x.
6. Now divide the size of the projected cell by the magnification to obtain the real cell size.

Materials: A hand full of hay, a large beaker, pond water, some milk

Method:

1. Take a hand full of dried grass or hay (free from pesticides or herbicides) and cut the grass into smaller pieces
2. Place the cut grass into the beaker and about 0.5-1 liter of water.
3. Add 1-2 drops of milk. The water will turn slightly turbid. The milk is food for the bacteria and they will start to reproduce. The ciliates feed on the bacteria and will also reproduce.
4. Let the beaker stand open for several days, protected from direct sunlight as this may result in overheating and the heat will reduce the oxygen concentration. Do make sure that the beaker receives sufficient light, though. Photosynthetic algae present in the pond water will produce oxygen.
5. Keep adding 1-2 drops of milk when the turbidity disappears. Bubble some air through the water at regular intervals (using an air-pump from an aquarium) or agitate the water a bit to enrich it with oxygen.
6. Replace the evaporated water.
7. Take some sample from the surface of the water for microscopic investigation.

Troubleshooting:

Problem: The water starts to smell.
Solution: This is normal. Bacteria are starting to decompose the hay and the added food. If bubbles start to appear though, then this is an indication that methane is formed anaerobically. This should not be and indicates that there is not enough oxygen in the water.

Problem: There are many bacteria but too few protozoa in the water.
Solution: Probably there was overfeeding. Add less milk and less hay. The bacteria multiplied too quickly and the protozoa could not keep up.

Problem: Nothing much seems to happen after a few days
Solution: Did you use chlorinated tap-water? Was the hay treated chemically?

Materials: Fresh pond water, wheat grains, glass beakers

Method 1:

1. Pour some pond water containing ciliates into the beakers and place 1-2 wheat grains into the water.
2. Wait for 2-3 days. The wheat grains will start to decompose and will seem to form a slimy layer around it. There should be thousands of ciliates in this slime. We have established a small food chain. Bacteria will break down the wheat grain. Paramecia will feed on the bacteria and reproduce.

Troubleshooting:

Problem: No paramecia have formed.
Solution: There were probably none in the original water sample. Paramecia and other ciliates can be found on the ground of ponds, in the slimy surface of rocks, etc. Include some of this material as well.
Solution: Did you use a complete wheat grain (with seed coat)? If you use rice or other polished cereals, then there are not enough nutrients available. The seed coat contains DNA and proteins (phosphates and nitrogen compounds) which are used by the bacteria.

Vitamin C (ascorbic acid) in polarized light.

Citric acid (citrate) in polarized light.

Materials: Any one of these substances: Vitamin C, Salycilic acid (the active component in Aspririn), citric acid, tartaric acid or table salt, destilled water, pure alcohol, a set of liniar polarizing filters

Method: for Vitamin C, tartarc acid, table salt

1. Dissolve A knife-tip of the substance in a few milliliters of destilled water. Shake until all of the substance is dissolved
2. Evenly spread the solution on a clean slide
3. Place the slide on a warm, but not hot plate or rest the slide for several hours (or over night) for the water to evaporate. It is possible to follow the formation of the crystals.

Troubleshooting Vitamin C

Problem: As the water evaporates, it starts to retract and does not form a nice thin layer of crystals on the slide. The crystals are thick and dense.
Solution: This is due to the surface tension of the water. Carefully spread the water all the way to the corners of the slide. The water should actually contact all the corners. The water is then not capable of retracting as it evaporates. The addition of surface tension reducing substnces (such as soap) may impair the crystalization process.

Method for Aspirin

1. Dissolve A knife-tip of the substance in a few milliliters of pure alcohol. Shake until all of the substance is dissolved.
2. Evenly spread the solution on a clean slide
3. Place the slide horizontally on a table and wait until the alcohol has evaporated. This should only take a few minutes. Crystal formation can be observed.
4. Careful: do not ingest! Only use very small amounts.

Method for Citric acid

1. Place a few grains of crystal on a microscopic slide and put a cover slip on top.
2. Carefully place the slide on a hot plate. The crystal will melt and will spread between the slide and the cover-slip. Do not overheat. The substance should not start to smoke or change its color.
3. Remove the slide and rest over night. Crystal formation can take several hours.
4. Alternatively, the melt can be spread out over the slide without a cover slip. Crystal formation can then be easily initiated by carefully touching the (cold) melt. Dust, crystal parts still sticking on the the fingers will initiate the crystalization.

Troubleshooting Citric Acid
Problem: Crystals do not form
Solution: This is quite possible, but not the rule. If both slide and cover slip are too clean, then there is not place for crystal formation to start. Use more citric acid the next time, so that some of the substance flows out beneath the cover slip. This can then be a place to initiate crystal formation by carefully scratching the substance with a sharp object. It could also be that the citric acid was overheated.

Problem: There are many bubbles between cover slip and slide
Solution: Bubbles are difficult to avoid completely, and in many cases they make the specimen more interesting (and beautiful) to observe. It is possible to melt the crystals first, and then press the cover slip on top of the melt. This also reduces the bubbles.

Materials: Onion, tap water, alcohol, fountain pen ink, several small beakers or film containers.

Method:

1. Using a sharp knife, cut out about 1 cm² of onion material. This may be done by the teacher to reduce the risk of injury.
2. The individual layers of the onion are separated by a thin skin. Peel this skin off using your fingernails or tweezers. The skin can be found on the inside part of each layer.
3. Place the skin into pure alcohol for about 30 seconds. This procedure removes water from the cells.
4. Place the skin into the ink. The ink, which is water-based, will enter the cells and strain the onion skin deep blue.
5. Using tweezers, transfer the skin into pure water and rinse it for about 30 seconds or until no more ink is given off. The skin will still have retained its blue color.
6. We now start the washing steps. We have to remove excess ink from the cells. Transfer the skin into pure alcohol for about 30 seconds. Some of the ink will be removed staining the alcohol slightly blue.
7. The skin is transferred into pure water for about 30 seconds.
8. The skin is carefully spread on a microscope slide and a cover slip is placed on top. Excess water is removed with filter paper
9. If the cytoplasm of the cells is still too dark, then it is necessary to repeat the washing steps 6 and 7 for a second time.

Troubleshooting:

Problem: The cells are too blue and too dark.
Solution: The cells were insufficiently washed. Prolong the washing times or introduce another washing cycle.

Problem: The nuclei are not stained, or not stained enough.
Solution: There could be several reasons for this.

• The onion was not placed into the alcohol. Therefore the water inside the cell was not removed and the ink could not enter the cell.
• The onion was not submerged long enough in the ink or the onion was not fully covered by ink on all sides
• The onion was washed too intensively. This is the most probable cause. If the onion was washed twice, then the washing step should be conducted only once.
• It could also be that the washing times were too long. If the nuclei are stained lightly blue, then this indicates that the staining procedure is in principle working, but that too much of the ink was removed.

This picture shows the tip of a maple leaf. Note that not all leaves can be processed this way.

You may be interested in the “Virtual Microscope”, which allows you to zoom into the leaf veins: Virtual microscope: maple leaf skeleton

Materials: Maple leaves, hot plate, cooking pot, eating plates, small but stiff brush or toothbrush

Method:

1. Let the leaves simmer for 1-2 hours. Periodically check the leaves by carefully rubbing them between your fingers. They should start to feel slimy and you should be able to rub off some of the surface plant tissue.
2. Carefully lift out the leaves. They are now very delicate and they tear easily. Put one leaf on one dish each.
3. Add a bit of water to the leaf on the dish. Use the brush to carfully remove the soft plant tissue of the leaf. The brush presses the leaf against the plate. This gives the leaf stability. Use the fingers of the other hand to prevent the leaf from moving while brushing. The leaf veins start to appear. Carefully turn the leaf around and remove the plant tissue on the other side as well. The water of the dish starts to accumulate plant tissue and should be exchanged periodically.
4. You now have a delicate network of leaf veins on the plate. Lift it out and place it flat on tissue paper to remove most of the liquid. Press the leaf veins between layers of tissue paper and a book. Otherwise there is the danger that the leaf will warp during the drying process.
5. Observe the leaf veins using a stereo microscope. They can also be observed using a compound microscope using a low magnification. Alternatively it is possible to scan the leaf veins with a flat-bed scanner.
6. Make a quality check. Observe any soft leaf material that has not been removed. Observe any tears and breaks in the leaf veins that were caused by brushing too forcefully.

Alternative method:

• Press the leaf between two books.
• Place the leaved into a solution of washing soda (pH 11 – don’t let children do this!) until they become pulpy and the soft material starts to come off.
• Rinse the leaves and brush off the soft material with a soft brush.

The Efficient Method: Do an Internet search for “skeleton leaves” and buy some ready made ones…

Other Ideas:

• Students may also attempt to remove the soft tissue directly under the stereo microscope. In this case the leaf should be placed in a petri dish.
• The cleaned leaf veins can be brightened by washing them in pure alcohol. This removes remains of the chlorophyll. The alcohol also removes water and the network of veins will shrink. Wash the veins in pure water after the alcohol treatment to restore the original size.
• The network of veins can be scanned using a flatbed scanner using high resolution. This also visualizes small structures. A dark background gives a nice contrast.

Troubleshooting:

Question: It is not possible to remove the soft tissue of the leaf.
Answer: Some leaves can be boiled for hours and still not macerate. Oak leaves are completely unsuitable for this preparatory technique. Try out a variety of different leafs. Alternatively, the leaf may not have been boiled long enough.