A possible health hazard: Biofilm on the underside of a bathtub stopper.

Much less has been written about the precautions that one should take when dealing with the organisms themselves. It is now my objective to address some precautionary measures when dealing with organisms that are to be microscoped.

Amateur microscopy certainly can not be considered a high-risk hobby, especially when one looks at ready-made permanent slides. Here the organisms in question safely killed and embedded in mounting medium. The issue starts to look a little different when one engages in collecting, concentrating and possibly even growing microorganisms for microscopic observation. Safety issues like this are not only relevant to amateur microscopists, but also for teachers who want to conduct basic microbiological and microscopic work in a school laboratory. In this case the organisms are alive and depending on the type of organism, they may pose a possible health hazard.

It it not, and can not be the intention of this article to give a detailed overview of the aseptic procedures used in a microbiology laboratory. I am not going to address the growing of bacterial colonies on agar petri dishes or the the making of a nutrient broth for the enrichment of bacteria. I am also not going to address the proper use of an inoculation loop and a Bunsen burner for sterile colony transfer. These laboratory methods are, in my opinion, too specific for the majority of amateur microscopists and require a properly equipped lab and appropriate training. Such issues can also not be covered in the little space available. The growth of (unknown) bacteria on agar plates or liquid culture medium also poses a potential health hazard, as the bacterial densities can be extremely high, and I generally would be cautious when working with nutrient media. There are also legal issues associated with these methods, as the legislation of some countries only permit the enrichment and growth of bacteria for certified laboratories. As a matter of fact, the growth of unknown bacteria isolated form the environment even requires the application of aspetic methods of an elevated biohazard level. Readers who are interested in these methods should consult introductory microbiology books, which cover these aspects in detail.

Rather, I would like to place a focus on the methods that are relevant for microscopists. In particular, I would like to address the making of a hay infusion, the observation of pond water as well as the observation of molds and other fungi.

Aseptic technique

The term aseptic technique refers to a medical or laboratory procedure that is performed under sterile conditions. The aseptic technique fulfils several objectives. First, the technique should protect the sample under investigation from contamination. This is of particular importance when culturing microorganisms, as fast-growing contaminants may possibly grow faster than the microorganism that one is interested in. The sample may thus quickly become overgrown by unwanted microorganisms.

While still working in a microbiology lab, I was told that a student working towards his diploma thesis accidentally sub-cultured a contaminant for several months. All of the experimental tests were performed on this contaminant and at the end of the thesis work the obtained data of several months was considered worthless. A quick check of the microorganism under the microscope would have quickly revealed the mix-up. For those of you who were wondering: Luckily I was not the unfortunate student.

Second, the aseptic technique should protect oneself from infection by potentially pathogenic microorganisms. The procedure therefore includes measures that prevent the inhalation of microorganism containing aerosols as well as the prevention of skin contact and ingestion.
Last, the environment and other people should be protected as well. Proper disposal of petri dishes and microorganism-containing sample materials is therefore necessary and often also required by law.

Risk Assessment

The dangers of contacting an infection depend on several aspects:

• Immune status of the person: The weaker the immune system of the person, the higher the chance of contacting an infection. For this reason, only handle unknown bacteria if you are healthy and have no immune system problems.
• Infectivity of the organism: Some pathogens can be infective at a low concentration, others require a higher concentration. Keep the concentration of the microorganisms low.
• Density of the organism: The higher the density, the higher the chance that a critical level of the microorganism is reached to cause infection. Just as above, keep the density of the microorganisms low and only grow them if it is not possible to observe them from natural samples.
• Mode of transmission: Different pathogens prefer a different method of transmission. Certain pathogens, for example, are transmitted over the air, others over water and still others over food. Others require insect vectors for transmission.

Most microorganisms are harmless, but one never knows what substances they are producing when grown at a higher concentration. Certain Cyanobacteria, for example, are known to cause eye irritations or allergies.

Hay infusion issues

A hay infusion is a culture medium which is commonly used to grow protists, such as the well-known Paramecium, for microscopic observation. Hay infusions have been popular since the beginning days of microscopy and are still a popular way of obtaining protozoa for educational uses in schools and universities.
There are two ways in which a hay infusion can be made. A handful of hay is boiled with water to extract nutrients, which serve as a food source for the microorganisms. The obtained culture medium must then be inoculated with the microorganisms that one wants to enrich. Pond water containing ciliates, for example, can be used. Generally this procedure is not recommended, as heat-resistant spored of potentially pathogenic bacteria can survive the boiling process. Alternatively one can try to enrich the microorganisms that can be naturally found on the hay. In this case the hay-water mixture is not boiled, but simply left standing for 24-48 hours. A thin iridescent bio-film will start to form on the water surface. This film is teeming with bacteria. In the presence of ciliates, the number of bacteria may decrease over time, and a progression of different organisms can be observed. Be aware that some countries have laws that regulate the use of hay infusions (and growth ob bacteria in general) for educational purposes.

One should be aware that unknown (and therefore potentially pathogenic) microorganisms may also start to grow in the hay infusion. The boiling process does not necessarily kill all of the microorganisms present on the hay. It is not uncommon to find heat-resistant spores of Bacillus and Clostridium on the hay. After the cooling of the infusion, these spores may start to germinate giving rise to live, possibly pathogenic, bacteria. The fact is, that you simply do not know what you are growing and appropriate safety precautions should be taken.

It is not possible to determine the pathogenicity of bacteria by microscopic observation. A range of biochemical and genetic tests are necessary. The enthusiast microscopist should therefore treat such a hay infusion with utmost care. Do not ingest the hay infusion, avoid skin contact (especially if there are open wounds), do not inhale the aerosols and prevent spills. Generally avoid contact of the liquid with mucous membranes, including the eye. Also make sure that the hay is clean and has not been in contact with excrements of animals. You may otherwise enrich bacteria from the animal’s digestive system. If a spillage or skin contact has occurred, then use 70% ethanol for disinfection (mix 7 parts of alcohol with 3 parts of water). A higher concentration of alcohol may actually have a lower disinfection efficiency.

Do not simply flush the hay infusion down the toilet. This may cause aerosol formation. Add chlorine bleach to the infusion and allow the substance to work for a few hours. Some people may be concerned that the bleach will then also find its way into the waste water, which is not very environmentally friendly. I would agree, but have no solution to this issue. Be aware that the addition of 70% ethanol to the infusion will dramatically lower the concentration of the alcohol. You can add concentrated alcohol to the infusion but this is a cost issue (and the glass jar may not be large enough).

Pond water safety

Even pond water may be the source of some unexpected surprises. I recently introduced mosquito larvae into my household this way. The mosquitoes caused me quite some irritation at night. Be aware that keeping a jar of standing water may even be illegal in countries with Malaria, which can be spread by certain mosquitos.
Other issues relate to the water quality of the pond water, may or may not be very high. Decomposing animals close to the sampling site can give rise to microorganisms that one does not want to have in the household.

Ponds which are clean enough for swimming should not be problematic, there are rare cases, where people did get infected by certain protozoa, however. Water samples from ponds which are rich in (potentially irritating) Cyanobacteria or eutrophicated should be handled with more care. Some ponds may be close to agricultural areas and there is the possibility for manure to run into the ponds.

Molds

Molds can be easily grown by treating an appropriate substrate (such as bread) with a soil-water suspension. Fungi will start to grow and release spores into the air. These spores may not only contaminate other types of food in the household, but may also be responsible for allergic reactions when inhaled. Many types of mold also produce potent toxins, which are capable of causing severe health problems.

Prevent the spreading of spores by keeping the container with the mold closed and avoid air currents which may distribute the spores.
If you want to investigate molds for educational purposes, then I would suggest that you try to first use edible molds, as can be found on foods, such as cheeses.

Biofilms

Biofilms are composed of microorganisms that stick together and to a surface. They can often be found on objects that are moist. The slimy covering of rocks in a pond are an example. Biofilms that harbor bacteria from human sources (e.g. bathroom stoppers) may pose a possible health hazard, also because the bacterial density can be quite high. Harmful microorganisms can also be found on other places in the household, such as dishwashers. What does this have to do with microscopy? Microscopy enthusiasts should establish clear procedures when taking samples from these sources to prevent contact.

General Advice

Here is some general advice when handling samples that contain microorganisms.

• Open wounds: Do not handle microorganism-containing media if you have open wounds or cuts in your skin. Intact skin can be considered as a very effective physical barrier against infection and open wounds can be problematic.
• Disinfection: Disinfect hands and surfaces with 70% ethanol. More concentrated ethanol may actually work less efficiently in killing microorganisms.
• Disposal: Autoclave the used culture medium at 120°C for 30 minutes. This should also be able to kill spores. If you do not have an autoclave available, then cover the petri-dishes or culture medium with chlorine bleach. Allow sufficient time for these substances to work. When you add bleach, be aware that this is a corrosive substance when concentrated. Eye and skin contact must really be avoided. Also be aware that liquid bleach becomes more diluted when you add it to liquid culture medium, losing its efficiency.
• Avoid aerosolization: Some microorganisms spread over air. Avoid spillage of the culture medium and carefully add the disinfectant to the medium before disposal, avoiding splattering of the liquid.
• Keep bacterial counts low: Make sure that the sample (such as a hay infusion) contains many ciliates that consume the bacteria. Keep the level of nutrients low to avoid too many bacteria from forming and ensure that the medium has sufficient oxygen supply for the ciliates to grow.
• Do not use polluted water: Dirty and polluted water can contain contains many bacteria and a lower count of the more interesting ciliates. If the water sample was isolated from a stream that came in contact with household waste water, then it may be possible that pathogenic enterobacteria are present.
• Do not decompose food: Some teachers like to decompose food to demonstrate the spoiling process to children. Be aware that Clostridium perfringens may be found on spoiled meat or poultry. This bacterium can cause food-borne illnesses. Personally, I would not use microorganisms from spoiled food for educational microscopy. I would resort to much safer and easily available bacteria and fungi. These can be isolated from fresh cheese, or example.
  Do not culture bacteria obtained from humans: In particular, do not inoculate growth medium with bacteria from the skin. You may be growing Staphylococcus, otherwise.
• Keep petri dishes closed and sealed: This minimizes the risk of accidentally touching the agar surface, which may be covered by bacterial colonies. Generally speaking, I do not recommend the growth of unknown bacteria in petri dishes by people who do not have basic microbiological training in aseptic technique. The bacterial concentrations are simply too high to be safe.

What is the take-home message? A good portion of common sense and basic hygienics will greatly reduce the possibility of you catching an infection and will hopefully keep you healthy.

For someone who wants to observe ready-made permanent slides or an occasional pond water sample, a fully equipped home laboratory may not be necessary and somewhat of an overkill. In this case it is sufficient to find a reasonably dust-free place to store and operate the microscope. The microscope can then be unpacked as required. For someone wants to prepare slides, perform microtoming and staining procedures, the issue may be somewhat different and space as well as equipment requirements are higher. As so often the case, it depends very much on the type of work that needs to be done.

The advantages of a dedicated lab can be summarized in a few points:

• Safe working environment – You need to protect family members, furniture and your own health from the chemicals that you use.
• Convenience and comfort – A dedicated work place does not require you to pack and unpack the chemicals and equipment that you use.
• Equipment safety – Microscopes should not be moved around too much – there is the danger that you drop them on your toes. This may hurt your microscope…
• Specimen quality – A proper work place makes it easier to produce (nearly) dust-free specimens. There is also less hassle.
• Fun – It’s simply more fun to work in an environment which has been designed accordingly. After all, it’s a hobby.

Be cautious about growing bacteria

There are several legal issues that you must be aware of if you intend to furnish a “wet” laboratory for microbiological work. If you want to grow (unidentified) bacteria in Petri dishes and culture medium, then you are already working in an elevated Biohazard Level 2 (out of 4 levels). You simply do not know if you are growing a pathogen or not. Even Level 1 laboratories must adhere to certain safety standards and decontamination procedures. Level 2 is even more stringent.

Now, what does this mean for the amateur microscopist? The answer is: do not enrich and grow unidentified bacteria. Even the enrichment and growth of bacteria that belong to the lowest Biohazard Level (level 1), such as E. coli and B. subtilis, may not be permitted, because a home is (legally) not considered a laboratory. And how do you want to obtain these known microorganisms? Cell culture collections such as the DSMZ (Deutsche Sammlung für Mikroorganismen und Zellkulturen) in Germany or the ATCC (American Type Culture Collection) may not even send samples to private individuals. Microbiological work may be prohibited even in school laboratories, because they do not possess the appropriate license to conduct microbiological work. They generally also do not possess the appropriate equipment in order to conduct safe work. The legal situation may differ from country to country, naturally, but I would not take the risk. Proper microbiological work also requires you to use a gas Bunsen burner, an additional hazard source.

As a side note: properly observing bacteria requires you to use a phase contrast microscope, something that not all amateur microscopists have available. Personally I also think that there are more interesting samples to observe than bacteria.

Microorganisms to observe

The amateur microscopist should not despair, there are many safe microorganisms, including bacteria that can be observed. My advice: go for microorganisms that can be found growing on fresh food:

• Joghurt - This is a good source of Lactobacillus delbrueckii subsp. bulgaricus and Streptococcus salivarius subsp. thermophilus.
• Cheese - Roquefort cheese, including other blue cheeses, can serve as a source for molds. Camembert cheese is a source for the moulds Penicillium candidum and Penicillium camemberti.
• Pond water samples and water from a home aquarium - These are good sources for a wide variety of ciliates, water fleas and algae. What about safety? Can you take a swim in the pond? Be aware that keeping pond water samples for extended periods of time in a jar may result in the water to turn foul. Unfriendly microorganisms may start to grow and I would be more cautious.
• Yeast - Also safe. Can be grown in a petri dish.

The requirements of setting up a microscopy work place

• Place for the microscope - The scope should have its own place and ideally it should not be necessary to pack and unpack the instrument. The table should be extremely stable to minimize vibrations. It should be easily cleanable with water to remove dust. There should be drawers for storing microscopic tools, slides and mounting media.
• Place for chemicals - You need a safe place to store the chemicals. You must be able to lock away the substances to protect them from kids. The place should also allow for containment and easy cleaning, in case there are spills. I once dropped a small bottle of iodine solution on our wood floor. The top layer of the wood floor had to be polished away because the solution ate its way into the wood, staining it red.
• The “WAF” - This one is often overlooked: the “Woman Acceptance Factor”. I once got into trouble because I wanted to store fly maggots and earth worms for dissection in the kitchen refrigerator. I did not even dare to ask if it is OK to modify the living room to accommodate a work bench for the microscope. The living room cupboards are also taboo for chemicals, also due to safety considerations.
• Dust-free environment - Often a difficult thing to achieve. Electronic equipment likes to attract dust due to static electricity. This dust can be quite interesting to observe under the microscope, but in most cases it is a serious nuisance, greatly decreasing the quality of microscopic images.
• A place for storing water samples - Pond water samples should not be stored in direct sunlight. This may cause overheating and (if there are few algae in the sample) a reduction in oxygen. The water can turn foul.
• Running water and sink - This is needed for cleaning the equipment and for disposing (permitted) solutions. Note, that some wastes must be collected and disposed separately.
• Work bench - You need some space for staining and preparing the slides. Some stains can be very aggressive and will irreversibly stain wood and other organic materials. Make sure that the work bench is easily cleanable.
• Ventilation - You need fresh air if you work with volatile solvents such as alcohol.

Equipment of a microbiology lab

Some amateurs (or teachers) may be interested in growing safe microorganisms such as yeast. It still needs to be mentioned that contaminations of the culture medium can be a health hazard. For people who want to equip a wet lab, the following equipment is necessary. You may also want to read the post: What accessories should be bought?.

• An autoclave - This is a pressure cooker. Used for sterilizing equipment and nutrient media. It is also used to kill off microorganisms on petri dishes before they are discarded.
• An incubator - This device allows for the control of the temperature. Petridishes with microorganisms can be placed into the incubator. This one is not always necessary. If the room temperature is too low, microorganisms may simply take longer to grow.
• Flowing water and a sink - Used for cleaning and washing. This one is pretty self-explanatory.
• Gas - The gas flame is used for sterilization and to minimize the risk of contamination when making the agar plates. It is also used to heat-fix the microorganisms on the slide.
• A shaker - This one is only needed if one intends to grow microorganisms in liquid medium. The shaking ensures that the liquid medium is supplied with oxygen from the air.
• Inoculation loop - For picking up colonies of microorganisms
• Nutrient media and agar - They supply the food to the microorganisms. The agar is used to solidify the medium.
• Petridishes - It contains the agar nutrient media.
• Parafilm - For sealing off the petri dishes.
• Various stains and reagents - These are used for fixing and staining the specimens.
• Miscellaneous - Materials such as gloves, alcohol for disinfection etc. are also needed

Measure the length of 1mm using a ruler or caliper. In this case, 1mm is magnified to 108.5mm.

Then measure the size of the structure on paper. In this case, we look at stomates from the bottom of a leaf. The guard cells are 3.6mm long.

• Place a ruler on the stage and take a picture. A full unit (1 mm) should be visible. Transparent ruler are better, otherwise it’s not possible to see the markings. Take a digital photograph of the ruler.
• Print the micrograph of the ruler.
• Take a picture of the specimen. Make sure that you use the same magnification.
• Print the picture of the specimen and be sure that the size of the picture is the same as the size of the picture of the ruler. Do not change the size of the print out.
• Now it’s time for a little math. Use the ruler and measure out the size of the 1mm on the print out. Measure it out in mm. Let’s call this “r”.
• Measure out the size of the structure that you want to determine. This is “s”. Make sure that you use the same units (mm)!
• The real size of the structure in mm can be calculated as follows: size = (1mm * s) / r

Let’s use the example in the pictures on the left:
size = (1mm * 3.6mm) / 108.5mm
size = 0.03mm = 30 micrometers

The air bubbles possess a different refractive index than the surrounding medium (water). This makes the bubbles appear to have a thick dark border. The shape of the bubble focuses the light in such a way that the center of the bubble appears bright.

Samples that are prone to form air bubbles

Not all specimens are the same. Some specimens can be the cause for more air bubbles than others. This depends on a variety of factors. The following characteristics may result in more bubbles:

• Large sheet-like specimens (e.g. onion skin): These specimens may catch air bubbles underneath them and prevent them from escaping. Push out the air bubbles before adding a cover slip.
• Specimens with many fine hair: The hair catch much air and prevent the water from reaching all the parts of the specimen. The surface tension of the water is too high, and the water therefore does not “flow” into all parts of the specimen. This is comparable to the “Lotus Effect”, where the water does not wet the surface of the lotus leaf.
• Fatty and hydrophobic specimens: These too do not accept water well, especially if the surface area of the specimen is large (many fine hair, etc). It may help to treat the specimen in alcohol or an alcohol-water mixture to remove the fatty surface.
• Porous specimens: The pores of the specimen may be filled with air, which can be difficult to remove. The cells of plant stems, the vascular tissue, for example, are able to hold air. It is possible to remove the air by placing the specimen into a vacuum while it is submerged in the fixing solution. Aspirators (eductor-jet pumps) can be mounted to a water tap to produce a vacuum.

Why air bubbles should generally be avoided

Some air bubbles are certainly tolerable and unless one wants to produce high-quality pictures it is often not worth the effort to make a completely bubble-free specimen. It is easily possible to simply move the slide and observe a different part of the specimen. Generally, air bubbles should be avoided, especially by beginning microscopists, who may have a problem distinguishing bubbles from the real specimen. The reasons why air bubbles can be problematic are:

• Bubbles hinder the free movement of organisms, such as ciliates
• The bubbles cause optical artifacts at the place where the air meets the water. The air bubble appears to be surrounded by a dark ring. This dark ring covers some parts of the specimen and makes observation more difficult.
• The microscope optics are designed to give optimum resolution for a specimen which is surrounded by water. If the bubble is large and the specimen completely surrounded by air, then the resolution is lower.

Are there cases when air bubbles are beneficial?

Under some rare circumstances, air bubbles can even be beneficial. The bubbles can serve as a source of oxygen for some organisms, such as paramecia and other ciliates. It is possible to see them collect around the bubbles. Air bubbles are also easily viewable and can therefore help beginners to more easily find the correct focus. Naturally, the bubbles should not be confused with the actual specimen, something that beginners sometimes do because the bubbles are so prominent and can be seen even if the specimen itself is not in focus.

How to minimize air bubbles in wet mounts

Needless to say, the preferred method depends on the characteristics of the specimen. Try out the following:

• Cover slip placement: Lower the cover slip on the water droplet with an angle. This permits air to escape on one side.
• Water placement: If the specimen is not fully submerged in the water droplet, add another droplet on top of the specimen before lowering the cover slip.
• Immersion oil: Use a medium other than water. Try immersion oil, which is hydrophobic. Some specimens prefer water, others oil.
• Break the surface tension: Add a small amount of detergent, such as soap. This will break the surface tension of the water. The water will therefore adhere better to some specimens, thus preventing bubbles. The soap may also harm some water organisms, however.
• Apply a vacuum: This speeds up the movement of the fixing solution or water into the specimen.
• Dehydrate the specimen: Place the specimen into alcohol. Some specimens will shrink and lose water and air. By placing the specimen into water again, the specimen will take up the water.
• Remove oil and fat: Wash the specimen in alcohol.
• Add water: If the air bubble is large and reaches the side of the cover glass, you can add more water from the side of the cover glass.

A few days ago I ordered a microtome. Here is a video showing you the different parts: Parts of a Microtome

Now it’s time to test the device. The first sample is a carrot. It can be cut into the right shape to fit into the specimen holder of the microtome and it is sufficiently solid to allow for easy cutting, but not too hard. Carrots can also be used to hold other specimens. In this case the “carrot cylinder” is cut in half and the specimen can be inserted between the carrot halves. The carrot acts as a support.

What is a wet mount?

In a wet mount, the specimen is suspended in a drop of liquid (usually water) located between slide and cover glass. The water refractive index of the water improves the image quality and also supports the specimen. In contrast to permanently mounted slides, wet mounts can not be stored over extended time periods, as the water evaporates. For this reason, a wet mount is sometimes also referred to as a “temporary mount” to contrast it from the “permanent mounts”, which can be stored over longer times. The permanently mounted slides use a solidifying mounting medium, which holds the cover glass in place. The naming can be a bit problematic, because it is also possible to make wet mounts that can store over extended time periods. These are special cases, however.

Different types of wet mounts

Wet mounts can be made using several different kinds of liquids. Water, immersion oil and glycerin (glycerol) can be used, with water probably being the most commonly used. The source of the water is quite important, especially when observing living specimens. If you use water with a wrong osmotic potential (ie. too much or too little salt and mineral content), then there is the danger of damaging the specimen. A too high salt content can result in the specimen to lose too much water. Too low a salt content, and the specimen may swell and burst.

• Using water from the natural habitat of the organism: In the case of water organisms, such as algae or ciliates, the liquid water should come directly from the sample. In this case the organism is immersed in its own natural environment. The microscopist uses a dropper to place a drop of pond water directly on the microscope slide.
• Using 0.9% salt water: In some cases water from the natural habitat may not be available. This is the case when observing bacteria or molds grown on petri-dishes. Yoghurt bacteria, for example, need to be diluted a lot before being able to observe them, otherwise they are too dense to be observed as single cells. In this case it is necessary to mix some salt (NaCl) into some water to ensure an optimal osmotic potential. This “physiological saline”, as it is called, can be made by dissolving 9 grams of table salt (NaCl) in 1 liter of water (or 0.9g Nacl in 100ml of water).
• Using tap water: If one wants to observe non-living specimens, such as dust samples, sand grains, or thin section cuts of plant material, then it is also possible to use regular tap water. These specimens are not osmotically sensitive. If the specimen is observed without water, in a dry condition, then the resolution and image quality may not be sufficiently high. I advise you to try out both to see the difference. The following post includes images of pollen grains mounted in air and water, for comparison: The effect of the mounting medium on specimen and image quality
• Using immersion oil: Some wet mounts are not made with water, but by using immersion oil. Immersion oil is usually placed on top of the cover glass. In this case the specimen does not get into contact with the oil. It is also possible to submerge the specimen in the oil, however. Heat-fixed bacteria can be observed directly by placing a drop of immersion oil on the specimen, without cover glass. The oil-immersion objective is then rotated directly into the oil for observation. It goes without saying, that this procedure can only be used for specimens that do not contain water (and are, therefore, not living). It also only works for specimens that stick to the glass slide – there is no cover glas. If you need to observe these specimens with a lower magnification (ie. no immersion objective), then one needs to use a cover glass, of course. Other specimens, such as synthetic textile fibers, are hydrophobic in nature, and do not like to be mixed with water. They tend to float on top of the water drop and this can be cause for air bubbles. In this case I also recommend to use immersion oil and a cover glass to keep the sample flat.
• Pure glycerin or glycerin-water mixtures: Glycerin has a strong tendency to withdraw water from the sample. For this reason it also acts as a preservative. On the down side, the glycerin may therefore cause the specimen to shrink and deform. Especially algae and other water organisms are sensitive to dehydration. Other specimens, such as sectioned or microtomed plant material are not as sensitive. The reason why glycerin is used is because of its high refractive index. This may be necessary to see certain structures. If a lower refractive index is needed, then one should mix some water into the glycerin. It is possible to seal the glycerin mount by applying nail polish to the sides of the cover glass. This will hold the cover glass in place for longer time periods. This is then an example of a wet mount, which was made into a permanent mount.

Advantages and disadvantage of a wet mount

Compared to permanently mounted slides, wet mounts do have certain advantages:

• Quick preparation: specimen fixation, dehydration and staining are not necessary (but possible, if required). For this reason, wet mounts are the first kind of mounts that students learn to make.
• Few artifacts: If there is no chemical and physical processing of the specimens before observation (no fixation), there are little artifacts and the specimens appear in their natural condition.
• Living and moving: It is possible to observe living and moving organisms. It is also possible to observe certain processes of life, such as feeding, cell division etc. (for water-based mounts)
• Natural colors: The colors are natural and not faded. The colors of permanently mounted specimens may fade over time.

Disadvantages of wet mounts include:

• Movement: The advantage of observing movement can also be a disadvantage. Due to the movement of the organisms it may be more difficult to take pictures or to make drawings. There is a solution to this problem: one can slow down ciliates and other protozoa by adding a solution such as ProtoSlo, which increases the viscosity of the water.
• Evaporation: The heat of the lamp causes the water to evaporate more quickly. More water must be added under the cover glass from time to time.
• Focus: Some organisms may swim vertically in the water and therefore move in and out of focus. Here it is important not to use too much or too little water. Too little water may squeeze the specimen between cover glass and slide.
• Storage: Wet mounts can not be stored over a longer time.

Materials and Method

For making a wet mount you need these materials:

• Microscope slides
• Cover glasses
• The specimen to be observed: make sure that the specimen is sufficiently small and thin. Thick specimens must either be cut (microtomed) into sections, be squeezed or torn apart.
• Water: take care that the osmotic potential of the water is compatible with the specimen. For example, do not use fresh water with marine specimens, and vice versa. Use pond water (and not tap water) for observing pond organisms.
• Droppers, pipette: these are for transferring the water
• Tweezers: for handling the specimen, the cover glass and for adding water

If the specimen is already in water (algae, ciliates etc.) then you can proceed the following way:

1. Place a small drop of sample fluid (containing the specimen) in the center of the microscope slide.
2. Hold the cover glass on one side with the help of tweezers. Lower the cover glass onto the water drop at an angle.
3. Then slowly lower the cover glass into the liquid. This will minimize disturbing air bubbles.
4. Remove excess water with filter paper or tissue paper. The cover glass should not float freely. The surface tension of the water should hold it in place. Alternatively you can add more water using a pipette or tweezers.

If the specimen is not in water:

1. Place a small drop of water (without specimen) in the center of the microscope slide.
2. Place the specimen into the water.
3. Add some more water on top of the specimen and make sure that the specimen is completely submerged. Otherwise there is the possibility for air bubbles forming between cover glass and specimen. The remaining steps are the same as above.
4. Hold the cover glass on one side with the help of tweezers. Lower the cover glass onto the water drop at an angle.
5. Then slowly lower the cover glass into the liquid. This will minimize disturbing air bubbles.
6. Remove excess water with filter paper or tissue paper. The cover glass should not float freely. The surface tension of the water should hold it in place. Alternatively you can add more water using a pipette or tweezers.

If you are using a dry specimen (dust, insect parts, etc.), then place a small drop of tap water

How to prevent drying out

The heat of the microscope light will evaporate the water relatively quickly. There are several possibilities to counteract this:

• Keep adding more water from the side of the cover glass. Surface tension will pull the water in.
• Seal the sides of the cover glass with a thick layer of Vaseline (petroleum jelly). Press the cover glass against the slide so that the vaseline is able to seal off the water from the outside.
• Use nail polish to seal off the cover glass. This is used when making wet mounts with glycerin. Keep the glycerin drop very small. The nail polish will not stick to those parts of the cover glass and slide which came into contact with the glycerin.
• Use slides that have an indentation (concave) and are therefore able to hold more fluid. This only works for some samples because the liquid layer may be to thick. These slides are more expensive.
• Use two additional cover glasses to support a third cover glass left and right. These two cover glasses serve as a distance holder for the third cover glass. This way the third cover glass does not float freely on the liquid but is held in place by the two supporting glasses. More fluid can be stored in a stable manner.

Characteristics of a chemical fixative

A good fixing agent should fulfill several criteria:

• It must kill the specimen quickly: But be careful, some chemical fixing agents are toxic and are also harmful to the health of a person.
• It must preserve the structures of the specimen, without introducing deformations or other artifacts. Insects may pull together their appendages, making them more difficult to see. The structures should then be sufficiently stable to withstand the dehydration and mounting.
• It must enter the specimen well to react with all parts: This can be problematic with some specimens. Make sure that the specimen is sufficiently small. Alternatively it is possible to puncture the specimen (insects) so that the fixing agent can enter more easily. Some specimens may contain air bubbles which prevent the fixing agent to reach all parts. In this case it may be necessary to apply a vacuum to remove the air.

Types of fixing agents

Chemical fixing agents can be categorized into the following 4 groups:

• Alcohol and acetic acid: This combination denatures proteins. The alcohol also removes some lipids. This is probably the preferred fixing agent for hobbyists, because it is less toxic than some other fixatives.
• Aldehydes (such as formaldehyde – toxic!): these react with amino groups in the specimen.
• Oxidation agents: these react with lipids.
• Tanning agents: react with proteins and with amino groups.

The choice of the fixing agent must be carefully matched with the specimen. Some fixing agents (eg. alcohol) may result in the shrinking of the specimen and therefore introduces artifacts. Sometimes it may be necessary to gradually increase the concentration of the fixing agent in order to prevent the formation of artifacts, but this depends much on the type of specimen used. I can not give general advice here, and recommend that one consults specific laboratory manuals.

Using alcohol

For the hobbyist who wants to prepare a slide every now and then, keeping a whole set of different chemical fixatives is probably an overkill (and not healthy either). I keep a small bottle of 96% rubbing alcohol on my shelf, into which I drop the specimens, usually small insects, as they arrive. They will store nearly indefinitely in this solution. When For making permanent slides, I directly transfer them into Euparal mounting medium.

Pure alcohol (ethanol) is also suitable for fixing and storing plant specimens, without cell contents. The alcohol has the tendency to shrink the cytoplasm, but does not affect the cell walls. The alcohol also hardens the plant material, making it easier to cut with a microtome (which often removes the cell contents anyway).

Alcohol/acetic acid solution

Acetic acid (acetate) compensates the shrinking effect of the alcohol. The Carnoy Clarke solution uses 3 parts 92% rubbing alcohol mixed with one part pure acetic acid. The correct alcohol:acetate ratio should be fine-tuned experimentally. If the cytoplasm still shrinks too much, the recipe according to Farmer may be tried out (2:1 alcohol:acetate ratio). Fixing should take place for about 24 hours.

After fixing

There are two more steps necessary: the fixing agent has to be removed (washing) and the specimen has to be dehydrated. Several fixing agents are water-based and this water has be be removed before mounting them in a non-water based mounting medium. Dehydration is not necessary when mounting in a water-based mounting medium such as glycerin gelatin. Dehydration is commonly done by placing the specimen in successively higher concentrations of ethanol. Afterwards the specimen is transferred into a solvent which is compatible to the mounting medium. Some mounting media require the specimen to be submerged in xylene (toxic). Other mounting media are able to directly accept the specimen from the alcohol (Euparal). If one sees a clouding of the slide, then this can be an indication that there was still some water in the specimen.

Heat-fixing of bacteria

Bacteria are treated differently. They must not only be killed, but also physically fixed to the glass slide. Otherwise they will be washed off during the staining process. This method also works with cells collected from the inside of the cheek and water samples.

• Place a bacterial suspension on the slide and let dry. Dry gently, dry completely but do not heat, otherwise the cells may pop open.
• Pull the glass slide through the flame of a Bunsen burner (1-2 times). The specimen should not come into contact with the flame (specimen on top, flame on the bottom). This step is called “heat fixing”. It kills of the bacteria and binds them to the glass slide much like an egg to a frying pan. The glass slide should be so hot that you are just able to hold it in the palm of your hands without causing burns. Heat the slide too much and you end up burning the bacteria 8and destroying their structure).
• The bacteria can now be stained. Place a drop of the staining solution on the cold slide. Rinse off with water and dry it in air. Do not dry-wipe, you will remove the fixed bacteria. You can then observe the bacteria directly in oil immersion even without a cover glass. Place the immersion oil directly on the fixed and stained bacteria.

Counting chamber: This one is called the Neubauer improved. There are other standards with different grids available as well.

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.

There are a variety of different factors that determine the achievable resolution. Some of these factors can not be actively influenced by the microscopist, others can. Some of the factors play a larger role, others a smaller one. In the following post, I want to summarize some of these factors.

Objective-related factors

• Correction of lens errors: In contrast to achromatic objectives, apochromatic objectives focus more colors of the spectrum to one point. This results in a sharper image.
• The numerical aperture of the objective: This value is printed on the objective. The higher the value, the higher the resolution. The numerical aperture is a dimension less value which represents the cone of light that can be caught by the objective.

Lighting system

• General color of light: The shorter the wavelength, the higher the resolution. If your microscope uses halogen or tungsten lamps (instead of LEDs), then the color of the light will shift towards the red end of the spectrum with increasing age. This will reduce the resolution. The color of the light also changes with its intensity. If you turn up the light to maximum intensity, then the color of the light will be more towards the blue end of the spectrum (shorter wavelength and higher resolution). LEDs do not change their color with age or brightness.
• Light spectrum (color range): The color range may also impact on resolution. In the case of monochromatic light, chromatic aberration does not play a role and the light can be focused on one point.

Specimen-related factors

• The correct thickness of the cover glass: The correct cover glass thickness is extremely important for high numerical-aperture objectives. For other objectives, the effect may not be noticeable.
• The correct refractive index of the cover glass: This is something that you do not have to worry about, this is the task of the cover glass manufacturer.
• The correct refractive index of the mounting medium: This one should be as close to the refractive index of glass as possible.
• Thickness of the mounting medium: the thinner the better.
• The presence of immersion oil: Objectives that carry the label “OIL” need the correct immersion oil for best resolution.

Adjustments of the microscope

• The correct condenser diaphragm setting: This setting must match the numerical aperture of the microscope in use.
• The correct setting of the correction collar: Some objectives have a correction collar (a turnable ring) to adjust to the cover glass thickness. Most objectives do not have one, however.

Maintenance-related factors

• The cleanness of the optical parts: Dust and dirt generally decrease image quality and are a big annoyance, especially if one uses dark-field microscopy.

Stability of the photomicrographic system

• Moving objects: Moving cells naturally cause a blurring when long exposure times are used. This decreases resolution of the moving object.
• Stability: A shaky photographic system generally decreases resolution of the image.

The checlkist: how to obtain the best image quality

• Use new light bulbs and turn up the light. This will reduce the wavelength of the light. Alternatively, use a blue filter.
• Use cover glasses of the correct thickness and make sure that the mounting medium has a refractive index which is close to the refractive index of glass.
• Adjust the condenser aperture diaphragm to the numerical aperture of the objective
• If you use oil immersion, make sure that the oil has the correct refractive index
• Use fresh light bulbs (low in red light, high in blue light)
• Keep the microscope free of dust
• Make sure that the objectives, eye pieces are clean

Types of cover glasses

Cover glasses come in all sorts of different sizes. I already wrote a post about cover glass size: Microscope Slides and Cover Glasses. In this post, we’ll now have a look at the importance of cover glass thicknesses. The table gives a summary of available thicknesses:

 

Why cover glass thickness is important

Most microscope objectives have the optimum cover glass thickness engraved into them. For most objectives this is 0.17mm. Read the following post for more information on the engravings: About the numbers on the Objective. The correct cover glass thickness is important to achieve the best resolution with a given objective. But do not go out to buy the more expensive 0.17mm cover glasses, get the thinner and cheaper ones (will be explained below).

Generally speaking, the higher the numeric aperture of the objective, the more serious the loss in resolution if the wrong cover glass thickness is used. For some high-aperture objectives, a cover glass thickness of only a few micrometers can significantly reduce resolution. Therefore, some more advanced objectives possess a correction collar. This is an adjustment ring which can be turned to adjust the objective to the actual cover glass thickness which is in use.

Importance of the mounting medium

The optimum cover glass thickness of many objectives is 0.17mm. Now, why is it that the most commonly available cover glasses are of category 1 (0.13-0.16mm), which is thinner than the calculated optimum? The answer is a bit more complex: The thickness of the cover glass is not the only parameter which is important. The specimen is embedded in mounting medium. The thickness of this medium must be added to the thickness of the cover glass. A specimen which is located deep in the medium will have a larger “effective” cover glass thickness than a specimen which is located right beneath the cover glass. A calculated (ideal) cover glass thickness 0.17mm is therefore a good compromise, even if the “real” cover glass is thinner. And yes, the refractive index of the mounting medium also plays a role.

How to determine the thickness of a cover glass

Cheap cover glasses which are used for uncritical routine observations will show a statistical spread of different thicknesses. There are also assorted cover glasses available that show a much more narrow spread of thicknesses. Some people buy cheap cover glasses (with a larger spread) and then manually measure their thickness using a caliper to sort them. Is it worth the effort? When using low-magnification objectives with a low numeric aperture, the difference in cover glass thickness may not even be noticeable and the more expensive pre-selected cover glasses may only be necessary for specific applications where a high resolution is necessary and the objectives do not possess a correction collar. One should not forget that the thickness and refractive index of the mounting medium also has an impact on the resolution, and mounting medium thickness may be much more difficult to standardize.

Ranunculus pollen mounted in air, no cover glass.

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

Field diaphragm is wide open. Reflections from the side of the tube are very strong.

Field diaphragm is half open. The reflections are less.

Field diaphragm is closed. Only the direct light is able to reach the camera.

Taking a picture of the tube with a webcam. Any camera with a small lens would also have done the job.

Advantages of Köhler illumination for photography

Köhler illumination increases the contrast of a photomicrograph because it reduces stray light and glare caused by reflections inside the microscope. On the right side you can see images taken through a trinocular head with a web cam. The more that the field diaphragm is closed, the less the reflections coming from the side of the tube. The bright spot in the center is the light which comes directly (unreflected) from the light source. In order to see a picture, it would be necessary to remove the lens from the webcam and project the image directly on the sensor of the webcam. In this case, the lens was left on to be able to see the side of the tube.

For more background info on Köhler illumination, you may be interested in the following two posts:

• Advantages of Koehler Illumination
• Adjusting Koehler Illumination

Different filters (patch stops) printed on overhead foil. The blue filter on the left is a commercial blue glass filter, on the bottom: the condenser with the 2 centering screws.

In a previous post, I already mentioned the making of patch stops from cardboard. For some background information (and more pictures) try these articles:

• Oblique Illumination
• Increasing Contrast using Optical Methods

Making Patch stops for dark-field illumination.

• Measure the diameter of the filter holder of your condenser.
• Using a program, such as PowerPoint or OpenOffice Impress to draw a circle, fill-color white, of the same diameter as the filter holder. You can adjust the size of the circle in the context menu.
• Draw a smaller black circle into the center. Copy-paste both circles and then change the size of the inner smaller circle. You want to make several filters to find the one that works best.
• Print the filters on overhead foil. Print with a laser printer. The overhead foils for laser printers are more heat resistant.
• Cut out the filters with a scissor
• Take a black marker and darken the black inner circle.
• For microscopy work, take two of these filters and place them on top of each other. This ensures that the central circle is completely black.
• Place the filter into the filter holder, completely open the condenser aperture diaphragm and the field diaphram (should you have one).
• Try out the different objectives and find the suitable filter/objective combination.

Making patch stops of oblique illumunation

The method is very similar to making patch stops for dark filed. In this case, light is only allowed to hit the specimen from one side only. This will produce a relief-like image.

• Draw a black and a white circle of the diameter of the condenser filter holder.
• Overlap the two circles, so that the white circle covers part of the black circle. The white circle should not reach the center of the black circle.
• Cut out and proceed as described for making a dark field patch stop.
• Again it is necessary to experiment to find the appropriate filter/objective combination.

Making Rheinberg filters

Maybe you want to show yellow specimens on a blue background. Take the dimensions of the dark-field patch stop and color the center yellow and the periphery blue (color printer!). You have to use intensive colors to achieve an effect. Try different color combinations.

Cocci in packets

Cocci in pairs and in packets

Short rods

Rods, slightly curved

About phase contrast

Bacteria are transparent and therefore difficult to see using regular bright-field microscopy. The bacterial cells will appear just as bright as the surounding medium and there is no color contrast. Phase contrast optics provides a solution. Phase contrast optics convert the differences in optical density (i.e. the refractive index) of the bacterial cells into different shades of brightness. The optics achieves this by interference of the light which passes through the specimen (the bacteria) with the light that goes around the medium. Phase contrast optics therefore work only if the cells have a different refractive index compared to the medium.

How the bacteria were prepared

The bacteria were grown in pure culture in an appropriate microbiology laboratory. A colony was then suspended in saline (salt water) of right concentration and then microscoped with a 1000x magnification in oil immersion (using a 100x oil objective).

If one takes too much liquid, then the cells start to float in and out of focus and it is not easily possible to capture the shape of the individual cells. A similar problem can occur if the cells are much smaller than the film of liquid between the slide and cover slip. The evaporation of the liquid from the edges of the cover slip will cause a constant movement of the cells and make it difficult to take a steady picture. In this case it is necessary to heat-fix the bacteria. A colony was then suspended in saline and dried at room temperature. The slide was briefly pulled through the flame of a bunsen burner, with the bacteria on the opposite side of the the flame. This heating process fixed the bacteria to the glass slide. Immersion oil was then directly applied to the slide and the bacteria were observed without cover glass. One disadvantage of heat fixing is, that during the drying process the bacteria may aggregate (as the volume of liquid decreases) and it may become more difficult to see individual cells.

About the photographs

The pictures were taken on analog B/W film and then digitized with a camera and an adapter (see the following post for more info on the set-up: Digitizing photographic slides with a digital camera ). The negative was then inverted and the contrast levels adjusted. The soft, slightly blurry appearance of the pictures shows that we are already at the limits of the resolution. The images were not sharpened. Notice the bright halo around the bacterial cells. This is typical for phase contrast microscopy.

Slide duplicator attachment: The left duplicator is mounted instead of the camera objective (via T2 adapter ring). The right one is attached to the existing objective (via filter threading)

Both systems compared.

The slide/film holder is the same in both cases.

Digitized slide showing vitamin C.

Digitized slide showing vitamin C.

There are several ways to digitize the slides:

• Using a slide or film scanner: This is the method of choice if you want to retain image quality. These devices are connected over USB to a computer. On the down side, scanning takes a long time and a film scanner is also not cheap. Some better slide scanners have a dust removal system.
• Use a flat-bed scanner: This is possible, if the resolution of the scanner is high and if there is a background lighting. Some flat bed scanners come with an appropriate slide holder. I found this system too time consuming, however.
• Get the slides scanned by a company: I did this once, it was expensive, but the quality was good. This is probably suitable for a smaller number of slides
• Photographing slides with a dedicated slide duplicator: This duplicator is directly mounted on the camera, instead of the existing objective. There is a slide/film holder attached. The slide duplicator that I initially tried was designed to reproduce 36mm slides again on 36mm analog systems (or digital cameras with a large sensor – the “full-format” systems). My digital camera’s sensor is smaller than film size. As a consequence it was not possible to fit the whole slide on the image and I always had added magnification. The objective allowed me to zoom in, but not zoom out (what I would have needed.) There are objectives like this that are specifically made for digital SLR cameras with a smaller sensor. So watch out if you get one of these devices.
• Photographing with a duplicator in front of the objective: This system is mounted in front of the camera’s existing objective. It contains extra lens elements to magnify the slide. This is the system that I used, and it worked well. The adapter is screwed into the filter threading of the camera’s original objective, so be careful that they are compatible (or use an extra adapter ring). One possible problem may be, that there are now many lens elements between the slide and the camera’s sensor. The image quality may suffer because of this. For my purposes, this was perfectly fine. Considering the generally low resolution of microscopic images, the quality loss was negligible. This duplicator also allows me to zoom in. This way I can take overlapping pictures of the slide and assemble them (“stitch” them) using panorama software. This way it is possible to reproduce the slide with an extremely high total resolution – but it’s time consuming (and it’s questionable if the slide / microscopic image has the necessary resolution in the first place.)

About exposure time

It’s very important to rest the camera body as well as the objective (whatever system is used) solidly on a stable surface. The objective should not be able to vibrate in relation to the camera body. If both are stable, then the optimum exposure time (to minimize vibrations) should not be too critical. Because I am in no hurry, I set the exposure to about 2 sec. The whole system will have vibrated out (and be steady) for the most part of the exposure. Long exposure times are more important when the camera is mounted on a microscope. In this case the effects of vibrations are much more evident. To minimize vibrations even more, I use the mirror-lock up feature of my camera.

About white balance

My camera allows me to adjust a custom white balance. I first take a blank picture of the white screen (the adapter system has a white screen) of the and use this as a reference image. The camera will then automatically adjust the white balance of all images that are taken. If your camera does not allow for the use of a reference image, then you should set the white balance manually based on the actual light source used. It’s not a good idea to use auto-white balance, as there is a color drift. Depending on the algorithm used, the camera may assume that the brightest spot on the image represents white (or a shade of grey, if darker), which may not be the case.

Potato starch grains in dark field

Potato starch grains in bright field.

Here I would like to show you two images of potato starch grains taken with different optical contrasting methods. The top image was taken in dark field, the bottom one in bright field. The purple or red structures are the starch grains of the potato (Solanum tuberosum). This is a nice example on how the addition of a simple field-stop filter can result in drastically different images. The contrast of the images was adjusted and both images were sharpened slightly. Image stacking was not necessary. The starch grains of potatoes are also called amyloplasts, they are found inside the cells of the potato tuber. Starch is a polysaccaride, made of long chains of glucose molecules. The glucose was originally produced by the leaves of the potato plant. Starch can be present in the form of either amylose or amylopectin. It is not water soluble and therefore suitable for storage.

Why talk about drawing microscopic images, if it is now possible to record the images using digital cameras? Drawing is not an old-fashioned or outdated method, rather it complements the possibilities of photographic documentation.

Advantages of Drawing Microscopic Images over Photography

• Combining different focus levels into one picture: Especially high-magnification images suffer from a low depth of field. A drawing is able to combine the different focus levels. It is now also possible to use image stacking software to combine different (digital) photographs from different focus levels into one final image.
• Removing artifacts: Dust and dirt do not have to be included in a drawing, but they are automatically part of a photograph.
• It is possible to draw a “typical” structure: The artist is able to look at several different specimens and then produce a final, typical drawing of the specimen.
• Emphasizing: The artist is able to emphasize different structures of the specimen, and to ignore others. This becomes useful if the drawing is to be used for identification purposes. This way a drawing can aid an inexperienced viewer. A photograph is often more complex with unnecessary details.
• Training of observation: Drawing takes practice and requires careful observation. These two aspects are trained.
• Same style: For publication purposes, it may be an advantage to show different microscopic specimens in the same style and size. Artists can use the same drawing style even for vastly different specimens. It is then possible to arrange the drawings on the same page next to each other without causing too much visual confusion.

Drawing Techniques

• Drawing without technical aid: For right-handed people, look with the left eye through the eyepiece of the microscope and with the right eye at a white drawing surface. You may need to adjust the angle of the drawing surface (placed right of the microscope) appropriately. With a bit of practice, your brain will combine the microscopic image and the white sheet of paper into one single image. You can then trace the image onto the paper.
• Drawing tubes: These devices can be installed beneath the microscope head. It will direct the image into a tube and project it directly on the table to be traced.
• Using a small mirror: A small mirror is mounted in front of the eye piece to project the image onto the drawing surface. The image can then be traced.

Impression of a leaf epidermis on white wood glue, oblique illumination. The color levels of the left image were adjusted to use the maximum contrast range. The right image shows the original color.

• Enhancing contrast: Photo editing software (such as Adobe Photoshop or GIMP) contain functions that enhance the contrast of an image. Find the menu point “Auto Levels” or simply “Levels”. This tool will make the darkest part of the image black (even if it was not black before) and the brightest part white. The resulting image will have the same information content, of course, but it may be easier to see the different structures. The photomicrograph will also not have its original color distribution anymore. This may be desired if the original picture has a red color tint due to the lamp of the microscope.
• Sharpening: Photomicrographs can be sharpened. This process results in aesthetically more pleasing images (if not overdone) but it too will not increase the information content of the image. The software enhances the contrast of the edges that it finds. An over-sharpening of photomicrographs results in so-called artifacts. The background noise (random color fluctuations) of the image is increased as well and structures that are not relevant may become more pronounced.
• Increasing depth of field: It is in the nature of compound microscopes to possess a limited depth of field. This can be an advantage, because it allows the observer to “slice-through” the different layer of a sample. By turning the fine-focus knob, it is possible to observe the different depths of a sample. When making photomicrographs, this may be a disadvantage, however. There are software packages available (see the links page) which are able to combine several photomicrographs (each on taken with a different part of the specimen in focus) into one final image. This process is called image stacking. The quality of the final photomicrograph depends both on the number of different images processed and if the focus of the images was sufficiently close together. See a stack of six separate photomicrographs of a Kiwi fruit.

Impression of a leaf epidermis on white wood glue. The stomata are clearly visible. Left: oblique illumination; Right: regular brightfield illumination. Oblique illumination gives the appearance of a 3-D surface structure.

Left: Home-made cardboard patch stops for oblique illumination. Notice the off-center hole. Top right: filter holder of the condenser; Bottom right: Commercial dark field patch stop for comparison.

Leaf Stomata impression in glue. The light appears to shine from the left, with one side illuminated and the other side in shadow.

Rotating the patch stop results in an image with different lights and shadows. The contrast of both images was digitally enhanced to increase the effect.

Oblique illumination only allows light to hit the specimen from the side. The main light beam is not able to reach the objective. This can be achieved by placing a patch stop into the filter holder of the condenser. These filters can be made of dark cardboard or other suitable heat-resistant material. The patch stop contains an off-center hole. The main light beam from the microscope lamp is not able to reach the objective. The specimen is illuminated from the side. This results in the image to appear 3D.

The best size and shape of the patch stop filter hole is best determined by experimentation. In any case, the hole should not approach the center of the filter, otherwise the main light beam from the lamp is capable of directly entering the objective, which weakens the effect.

A darkfield filter (patch stop) placed into the filter holder of the condenser. To the left and the right are the centering screws.

Potato starch grains. Left: darkfield image; Center: Brightfield, inverted colors; Right: Brightfield; The comparison shows that a darkfield image is not simply an inverted version of a brightfield image. Darkfield images have more sharply defined corners.

Maize. Left: darkfield image; Center: Brightfield, inverted colors; Right: Brightfield; The darkfield image possesses less contrast due to the opened aperture diaphragm and a different color representation.

To achieve a darkfield image, it is necessary to place a dark field filter (a “patch stop”) into the filter holder of the condenser. This filter prevents light of the lamp to directly enter the objective (therefore the background appears dark). The specimen will be illuminated from the side and will scatter some of the light to enter the objective. The specimen will appear bright on dark background.

It can be compared to dust floating in the air with sun shining in from the side through a window. The dust is illuminated by the sun and appears bright on dark background.

There are two possibilities to achieve a darkfield image:

• By using specialized darkfield condensers: This is the best but also the most expensive solution.
• By using a darkfield filter (a “patch stop”) which is placed into the filter holder of the condenser. It is possible to make the patch stop out of cardboard or a tin can using a cutting knife and scissors.

Advantages of darkfield microscopy:

• It is a simple procedure which can be used on live transparent specimens, specimens which normally need to be stained (and therefore killed).
• The images appear spectacular and are visually impressive.
• Darkfield microscopy even allows for the visualization of objects that are below (!) the resolution of the microscope. These objects will appear as bright spots on a dark background. It is not possible to see the shape of these objects, however.

Some possible disadvantages of darkfield microscopy:

• Darkfield microscopy is very sensitive to dirt and dust located in the light path.
• It is not suitable for all specimens. If the refractive index of a transparent specimen is similar to the surrounding medium, then the specimen light will pass right through the specimen and it will not be scattered into the objective.
• The intensity of the illumination system must be high so see the specimen properly.
• It is necessary to open the condenser aperture diaphragm, and this limits the effective use of the diaphragm.
• One patch stop is generally sufficient for low magnification work, but at a higher magnification the quality of the image drops. It may be necessary to experiment with different patch stop sizes for the different objectives.

The condenser aperture diaphragm can be controlled with a small horizontal lever (top). Left and right are the condenser centering screws. They are needed for adjusting Koehler illumination. Behind the left centering screw you can see the condenser focus knob.

Here the condenser aperture diaphragm is set to a value of 0.25, which is the recommended value for the objective in use. The depth of field is low, the resolution high, the contrast is low.

Here the condenser aperture diaphragm is set to a value of 0.1, which is the closed position. The depth of field and contrast are both high. The image appears crisp, but resolution is lower.

An improper setting of the condenser aperture diaphragm (especially at higher magnifications) can be the cause of much frustration both for teachers and students.

• Students may attempt to find the focus with the condenser aperture diaphragm all the way open. This is difficult if the sample is very thin or weakly stained or the microscope is not equipped with parfocal objectives. Remember, an open condenser aperture diaphragm results in a low depth of field.
• Students may not see anything at all when working with high magnifications because the image is too dark. In this case the diaphragm is closed too much. The diaphragm should not be used to control the amount of light, but for some specimens or magnifications there may simply be no way around this especially if the lamp is not very powerful.

Many beginners are place an overly strong emphasis on magnification. Many think that they are able to see more at a higher magnification. But especially at higher magnifications the role of the condenser diaphragm becomes more important.

I recommend the following steps:

• Instruct the students to completely close the condenser aperture diaphragm when starting to use the microscope.
• They should then rotate the low power objective (4x) into position and find the focus with the coarse focus knob. The larger depth of field and higher contrast makes it easier for the students to focus the specimen.
• When switching to a higher magnification, the students should start to gradually open the condenser aperture diaphragm, to observe the differences in image quality. At the same time they have to adjust the light intensity with the dimmer to prevent glare.
• Students should be made aware that the condenser aperture diaphragm should be adjusted to the numerical aperture value which is printed on the objective. Opening the diaphragm further will not increase image quality, but may result in glare.
• If the sample is thick, strongly stained or pigmented then the diaphragm has to be opened to allow more light to pass through the specimen. As a consequence, the depth of field becomes smaller. It is then necessary to use the fine focus adjustment knob to focus through the different layers of the specimen.

The Koehler diaphragm is centered and in focus. The adjustment is correct.

The Koehler diaphragm is centered but out of focus. Raise or lower the condenser to focus the diaphragm.

The Koehler diaphragm is off-center. Turn the centering screws on the condenser to move the aperture into the center.

Koehler illumination was developed by August Köhler (1866-1948). This illumination principle greatly enhances the quality of the microscopic images (especially photographs). The illumination principle offers the following advantages:

• It illuminates the specimen uniformly without the need of a diffuser.
• It only illuminates the part of the specimen which is actually observed (at a higher magnifications a smaller section of the specimen). This reduces the heating of the specimen.
• It reduces internal reflections. This improves the contrast in photomicrographs.

The Koehler illumination must be adjusted before observation:

1. Rotate a low power objective (eg. 4x or 10x) into position. This will increase the field of view.
2. Insert a slide with a specimen and focus it.
3. Adjust the field iris diaphragm (the diaphragm of the light source) in such a way that its edges become visible. The field of view is reduced this way, only a small round part of the specimen is visible.
4. Raise or lower the condenser (not the stage!) and bring the edges of the field iris diaphragm (not the condenser aperture diaphragm) into focus. The focus of the specimen is not changed. Now both the edge of the iris diaphragm and and the specimen should be in focus. If the height of the condenser is not properly adjusted, then dust of the lamp will come into focus and disturb the image.
5. There are two condenser centering screws/knobs at the side of the condenser. Turn these knobs to bring the field into the center of view.
6. Now you can open the field diaphragm and start regular microscopic observation.
7. When doing photographic work, open the field diaphragm only as far as necessary. Opening it further will increase internal light reflections and result in a lower contrast. You need to observe the edges of the field diaphragm through the camera viewfinder. It may also be necessary to refocus the specimen when looking through the camera.

When the polarizing filters are turned into a crossed position, then they will not allow light to go through. This is the position used for microscopy.

When the polarizing filters are turned into a parallel position, then they will allow light to go through.

Place one polarizing filter on top of the light source, and the other one on top of the specimen.

Polarization microscopy of crystals is an aesthetically rewarding experience. Obtain two linear polarizing filters. Make sure that the two filters will not let light go through if crossed. Many polarizing filters sold in photography stores are circular polarizing and they will not work. It is best to test the filters first, or to buy polarizing filters from a school supplies company.

Place one filter on top of the light source and the other filter on top of the specimen, beneath the objective. Then rotate the filter of the light source into a crossed position. Be careful – The filter changes the focal distance and focus. Be careful of not smashing the objective into the filter when refocusing. For safety, only use this system with the low power objectives.

There are a wide range of different samples that can be viewed under polarized light:

• Various crystals
• Potato starch grains
• House dust: many components of dust are de-polarizing the light and these components will appear bright on dark background, similar to dark-field illumination.
• Transparent materials (plastics) that contain tensions. The tensions turn the plane of polarization of light and will result in colorful images.

It is possible to purchase dedicated polarization optics. These optics are tension free and will deliver a completely dark image when used with crossed polarization filters. Regular achromatic bright field objectives (as commonly used in schools) are not tension free and there may be a slight background illumination even when the filters are completely crossed. For practical purposes, this is of no relevance.

Vascular tissue of a pumpkin plant.

Here is a list of common mistakes which I observed over the years:

• Viewing specimens without a cover slip: The objectives are designed to be used with a cover slip. If no cover slip is used (or no water beneath the cover slip and the slide), then the focal distance will change and the quality of the image is reduced as well.
• Using immersion oil with a non-immersion objective: Lower image quality and dirty optics are the consequence. The oil, if not properly cleaned, will start to accumulate dust and image quality may decrease to the extent that no image is visible at all. Use an alcohol:ether mixture and lens paper to clean the objectives, but make sure that the solvent does not contact the lens too long. Otherwise the lens kit holding the lens in place may start to become soft.
• Using the coarse focus with higher magnification objectives: This may result in crashing the objective into the slide. Spring-loaded objectives offer a level of security here.
• Turning the fine focus adjustment for a long time to find a focus: This too may result in crashing the (high-power) objective into the slide. Instruct the students to restart their observation with the low power objective.
• Using the iris diaphragm as a means to control the amount of light: The iris diaphragm of the condenser is there to regulate resolution and contrast, but not to regulate the amount of light. At high magnifications it may be necessary to open the diaphragm to produce a brighter image, but the students should first use the dimmer to control the light.
• Switching the microscope on and off with the dimmer set to the highest light intensity: The lamp is heated up quickly. It is better to slowly increase the light intensity with the dimmer.
• Starting to observe with a high magnification objective: This is a common thing to observe. Students should start with the lower magnifications first. This allows them to select the area of interest of the specimen.
• Using thick, non-translucent specimens: For specimens of these types, it is better to use a stereo (binocular-) microscope.
• Using oil-immersion objectives without oil: This changes the focal distance of the objective and results in a low quality image. Students may then turn the focus knob to the extent of crashing the slide into the objective.
• Moving the microscope with the lamp switched on: This may result in a lower lamp lifetime. Move the microscope only when the lamp is cold.

• Optical techniques: The use of phase contrast is a very popular technique to increase contrast in research labs, but it is probably too expensive to be used in schools. Phase contrast optics transform transparent objects into a black-white image, depending on their refractive index.
• Staining techniques: Transparent specimens, such as bacteria, can be heat-mounted on the slide and then stained with specific chemicals.
• Use of filters: Colored filters can be used to enhance the contrast of certain objects. If the object already possesses a certain color, then a filter with a complimentary color will result in the specimen to appear darker.
• Use of dark-field illumination: A dark-field ring can be placed into the filter holder of the condenser. Specimens will then appear bright on dark background. This system does not simply invert the colors, but makes specimens with a refractive index different from the medium visible.