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.

1. Place a drop of water on the slide and then a small piece of the specimen into the water.
2. Carefully position the specimen in the center and place the cover glass on top, as if making a regular wet mount.
3. Using a soft round object, such as an eraser, carefully press down on the coverslip without horizontal movement, which would introduce shearing forces. This is the testing stage, to check if the specimen is sufficiently soft. The cover glass may break otherwise. This is, why you should use an eraser, to protect yourself.
4. If the sample is sufficiently soft, you can press down with more force. The sample should form a thin, almost transparent film between coverslip and slide. Again, do not introduce a horizontal movement.
5. Excess water should be soaked up with tissue paper. If some of the specimen starts to appear from beneath the cover glass, then you used too much specimen.
6. You can use any objective to observe under the microscope.

Samples that are too solid need to be softened first. Some plant material can be made softer by boiling, but this may not be enough to soften the cellulose of the cell walls. The cellulose of the cell walls can be made softer by heating with an acid, such as diluted HCl or acetate (careful, dangerous). Rinse the specimen after acid treatment with water and compress it.

Staining the samples should take place before squashing the specimen, as it is otherwise difficult for the stain to reach the cells.

Suitable specimens include soft fruits and fungi. Squashing may introduce artifacts. The cells are separated from each other and it is not possible to see the original place of the cells. If you want to see the arrangement of cells, as they occur naturally, then you need to resort to microtoming. The advantage of squashing is, that it is a fast and easy method to obtain very thin specimen samples.

Electron microscopic image of E. coli bacteria (10000x). Image credit: Eric Erbe, digital colorization by Christopher Pooley, both of USDA, ARS, EMU.

I do not know to what extent the news is also dominating in other parts of the world, but here in Europe the last few weeks saw one of the worst bacterial outbreaks since decades. The media, naturally, was (and still is) full with reports on this epidemic. The EHEC (enterohemorrhagic Escherichia coli) bacterium is a new E. coli strain which is not only highly infective, but can also cause a range of serious, life threatening conditions, including bloody diarrhea, kidney failure and neurological disorders. The bacterium is contracted over food, at least this is the suspected mode of transmission. Most infections occurred in Germany.

The general public scare resulted in a dramatic decrease in the purchase and consumption of fresh vegetables, even in those parts of Europe that were not affected by the outbreak. Consumers were (and are) simply afraid and this had a dramatic impact on the sales statistics of vegetables.

Microscopic observation of EHEC?

The public scare, emerging conspiracy theories (is the epidemic a biological warfare experiment?), as well as sometimes contradictory information (first Spanish cucumbers were the source of infection, later this was taken back), may motivate some people to take things into their own hands. Is it possible to test vegetables and other food, as well as one owns stool for the presence of EHEC, using microscopic observations? I have already read such questions in microscopy related forums and I think that this issue needs some clarification. After all, this question of microscopically testing food and body fluids for the presence of pathogenic (ie. disease causing) bacteria is not only limited to EHEC but was also asked in connection to Clostridium difficile and other bacteria. It is a question that reappears periodically and there seems to be a need for an explanation.

Here, for once, the answer is quite easy: It is not possible to use microscopic observations for identifying bacteria. I know that this may sound ironic, because, historically, microscopes were those devices that gave the field of microbiology and bacteriology a great push forward. In the following, I would like to outline a few points why it is not possible to use microscopes for testing for the presence of pathogenic bacteria.

Morphology

The shape of bacteria, their morphology, does not provide information on the danger of the organism. Bacteria are disease causing, if they possess so called “virulence factors” (this term has nothing to do with viruses). The virulence factors can either be toxins that are released, or structures on the surface of the cells that allow the bacteria to adhere to surfaces, such as the wall of the human intestine. These virulence factors can not be observed microscopically. Two bacteria that have exactly the same morphology, one can be pathogenic, the other one not, depending on whether they possess these factors or not. For diagnostic purposes morphology is pretty much irrelevant. To use an analogy: Just because two cars have the same shape does not mean that both of them are safe to drive, one of them could have a safety problem, which can not be seen from the outside. And just because your own car is a blue Ford does not mean that all blue cars are Fords. Just like morphology, the color of a car says nothing about its internal characteristics and model.

Density

The bacterial density on food is too low for microscopic observation. In the case of EHEC, eating about only one hundred bacterial cells are enough to cause an infection. If these bacteria are distributed over the whole food, then which part of the food should be microscoped? The chances are pretty good that there are thousands of other, non pathogenic, bacteria present as well, so how do you want to distinguish them? Generally, the bacteria have to be enriched by placing the suspected food sample into a growth medium, where they reproduce. The bacteria have to be cultured first, before they can be analyzed. Now this is something that I would definitely not recommend to do for safety reasons. It is even illegal for non authorized people to do this, after all, one does enrich potentially pathogenic organisms. Even if an enriched culture of bacteria is available, the problem would be the same as the problem mentioned above. The shape of a bacterium says nothing about its danger and its kind.

Testing for presence of bacteria

Some people may simply be interested to test for the presence of bacteria on their food, to assess whether it is safe for consumption or not. They simply want to check if “something is there”, disregarding if it is EHEC or not. If bacteria are present (regardless of the kind), then they would play it safe and not consume the food at all. No fancy tests would be needed in this case, and enrichment is also not necessary. Take a cotton swab, collect some bacteria from the surface of the vegetable, transfer them to a slide and observe them under the microscope. This should work? Or not?

There several false assumptions made. First, they assume that food is generally free of bacteria, which is not the case. The vast majority of them are not pathogenic, however. Bacteria are omnipresent and there are even more bacterial cells growing on and in our human body than we have body cells. They simply can not be avoided and are part of out natural environment. Unless one sterilizes the food, bacteria are certainly to be found.

The second false assumption is that often people think that most bacteria are pathogenic. Bacteria have a negative connotation, and are linked, in public perception, to the “three Ds”: dirt, disease and dismay. Bacteria truly need a lobby, and more positive PR, I may remark. Generally finding bacteria on food says nothing about the quality of the food product. It’s the type of bacteria that matter, and this, we know, can not be determined microscopically.

There is also a third false assumption. If one is not able to see bacteria under the microscope, it does not mean that none are present. Bacteria require more advanced optics (phase contrast) and a certain amount of skill to distinguish them from non-bacterial objects. They can be quite small as well. And the density of the bacteria can be quite low.

What do the labs then do? They selectively enrich the bacteria and analyze a range of biochemical, immunological and genetic parameters. At the end they make a nice false color a 3d image of EHEC using a scanning electron microscope so that they have something nice to show to the media. Pictures from a light microscope often do not look spectacular enough. Still, these pictures are not used for diagnosis or identification purposes.

What about Fluorescence Microscopy?

Now, it is possible to use labeled antibodies and mark the bacterial antigens to test for EHEC and other pathogens. This is a more advanced procedure and in this case it is not the bacterial morphology that is used for identification, but rather the ability of the antibody to interact with structures on the bacterial surface. I just wanted to briefly include this point for the sake of completion.

In summary

Many words, but simple message. One can observe bacteria microscopically as much as one wants, but one will not be able to assess their danger this way. Microscopic observation has no diagnostic relevance. One will not even be able to say if two bacteria of the same shape are genetically related or not. Without culturing the bacteria (don’t do this), chances are pretty good that the bacterial density is not even high enough for you to see anything under the microscope, unless your food sample was spoiled. But in this case you don’t even need a microscope to know that.

Questions? Comments?

There is a comment form below!

Yeast

You can either use fresh (wet) yeast or dried yeast. In either case, take a small amount and dissolve in a little bit of water, until the liquid becomes turbid. Use this suspension for microscopy. (read The hemocytometer (counting chamber) to sell how yeast cells look like in a counting chamber).

Yogurt

One of the more difficult specimens. Yogurt contains many bacteria, these are a bit difficult to see with bright-field microscopy. You can stain them (read ). Take a small sample (knife-tip) and dissolve in water. Then apply a drop to the slide, apply a cover glass, and observe under the microscope.

Cheese

Here you have to take the right kind of cheese. The toast-cheese (the one where each one is wrapped separately in plastic foil) won’t work. They don’t have any fungus growing on them (Do not let it rot, you may be growing poisonous fungi). I’m a cheese lover and I consider Camembert, Brie, Gorgonzola blue cheese not only good for eating but also a valuable source for the fungi Penicilium.

Honey

Some of them contain pollen. If the honey is turbid (opaque) then this may be due to sugar crystals or due to pollen. Clear honey won’t work.

Heat fixing

Heat fixing essentially “bakes” the cells to the glass slide much like a fried egg sticking to a frying pan. Heat fixing is absolutely essential before staining. Otherwise the staining procedure will wash away the cells.

• Take a cotton swab and rub the inside of your cheeks to collect epithelium cells.
• Smear these cells on a microscopy slide
• Completely air-dry the slide, without applying heat. This should not take long because there is not much liquid on the slide anyway. If you heat the slide before it is completely dry, then you end up “boiling apart” the cells. The vapor pressure inside the cells will burst them…
• Heat fix the dried slide by quickly pulling it through a Bunsen burner (2x), but in a way that the cells do not touch the flame. Pull it through the flame with the cells on top and the flame below. The slide should be pretty hot but you should still be capable of holding it in the palm of your hand without burning yourself. You should just be capable of holding the slide. Too high a temperature and you destroy the cells on the slide (and on your skin!). Too low a temperature and the cells will not stick to the glass slide.

Staining

Apply a drop of the stain (eg. methylene blue) to the heat fixed but cold specimen slide. Allow the stain to work for a few minutes and then carefully rinse away the stain with water. Do not apply the water stream to the cells directly. Allow the water to run over the cells from the top of the slide. Air dry the slide and observe in the microscope.

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

I would now like to show you a much better method of determining the size of microscopic structures. You do need a hemocytometer (counting chamber), however. These specialized slides are designed to determine the concentration of cells but they can also be used to determine size. The disadvantage is, that hemocytometers do cost quite a bit more than regular slides. There are different types of hemocytometers around, it does not matter which one you use, as long as you know the real-life size of the engraved squares.

In this case, we use the side length of one of the squares of the hemocytometer as a reference. These lines are very fine and therefore permit you to make very precise measurements and size calculations (in comparison to the picture of a ruler, see above link). The math is easy, but be careful that you use the same units.

• Step 1: Take a picture of a square of the hemocytomer with associated cells. There are squares of different sizes, so make sure that you know the dimensions of the square that you are looking at. Read this post for more information on the different square sizes of the Neubauer improved haemocytomerter: The hemocytometer (counting chamber). Make sure that one complete side length of a square is visible.
• Step 2: Print the micrograph. The square of the hemocytometer is out internal reference. You do not have to worry about the size of the print out. The larger the print out, the more precise the result, however.
• Step 3: Measure the length of the side of one square and the diameter of a cell. Use the same units (generally mm is appropriate).
• Step 4: Calculate the real-life size of a cell: You know the real-life side length of a square and the length of the square on the print out. You also know the diameter of the magnified cell. This data is enough for you to calculate the real-life size of the cell.
• Step 5: Divide the length of a square of the print-out with the real-life side length. This gives you the magnification on paper. This magnification has nothing to do with the magnification of the objective and eye piece. We’re talking about magnification on the paper.
• Step 6: Divide the size of the cell on paper with the magnification to obtain the real-life cell size. If you mix units, (cm, mm), then you won’t get the right result. You need to convert to the same units first. E.g. do not divide the square size in cm with the real life size in mm.

Things to watch out for:

• Do be careful when observing cells that have a vastly different refractive index to the surrounding. In this case the cells will appear to have a thick “wall” around them, which is actually nothing more than a diffraction pattern. This may make obscure the true size of the cell. Open the condenser aperture diaphragm to minimize this artifact.
• Counting chambers have squares of different sizes. Read the manual first so that you know the true size of the square that you are looking at.

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.

Compared to other mounting media, Euparal has a significant advantage: Specimens can be directly transferred from alcohol into Euparal. Other mounting media, such as Canada Balsam, require the specimen to be transferred to xylene (toxic) prior embedding.

Euparal was first described by G. Gilson (prof. of Zoology at Louvain University, Louvain, Belgium) in 1906, and is liked for its ease of use and stability.

I wanted to find out more about this mounting medium and conducted a quick Web search, only to find out that the information is scarce. I wanted to know more about the composition of Euparal, also because of safety considerations. Many non-aqueous mounting media contain toxic organic solvents, which I try to avoid.

After some time I was indeed able to find the original publication by G. Gilson [1], published in French. An article published in the Journal of the Royal Microscopical Society [2] summarized parts of this his and together with a translation software, I was able to extract some meaning of Gilson’s article.

Composition of Euparal

Gilson found out that Sandarac (or sandarach) is a suitable resin for mounting. It is obtained from Callitris quadrivalvis, a tree belonging to the cedar family. The resin has been previously used to make varnish and protective coatings for paintings. Alcohol could disolve the sandarac well, but the resulting medium was not suitavle for microscopic work. During the curing process the sandarac started to crystalize and crack.

Gilson then mixed the sandarac with camsal, a mixture of phenyl salicylate (salol) and camphor. The camsal was not a good solvent for the sandarac, but prevented the formations of crystals and cracks. He added either isobutylic or propylic alcohol to further dissolve the sandarac. Especially isobutylic alcohol was considered suitable, as it was commonly used to dehydrate microscopic specimens. The specimens could then be directly transferred into the mounting medium (containing the same solvent).

While this was aldready a step into the right direction, the added alcohol was a substantial disadvantage. Stained specimens could not be mounted with this medium, as the alcohol dissolved many the dyes which are commonly used in microscopy, such as eosin, safranin, methyl green were affected by the alcohol. Camsal alone was not able to sufficiently dissolve the resin.

Gilson, therefore, searched for replacements for the alcohol. He discovered that a combination of eucalyptol and paraldehyde was able to substitute for the alcohols, without harming pigmentation. He thus gave the mixture containing sandarc, salol, EUcalyptol and PARAldehyde the name Euparal.

• Sandarac: A resin which solidifies in air. Originally used as a varnish for furniture.
• Paraldehyde: Preservative and solvent.
• Eucalyptol: Solvent of Euparal. dominant portion of Eucalyptus globulus oil. eucalyptol is used as an insecticide.
• Phenyl salicylate (salol): Antiseptic substance, was introduced in 1886 under the name Salol, a desinfectant.
• Camphor: An antimicrobial substance, previously used for embalming. Obtained from the evergreen tree camphor laurel (Cinnamomum camphora).
• Camsal: A mixture (1:1) of camphor and Phenyl salicylate (salol).

Advantages of Euparal

Towards the end of the article, Gilson lists several advantages of Euparal. These advantages are now briefly summarized:

• It is possible to directly transfer specimens which were stored in 70% alcohol into Euparal for permanent mounting. It is not necessary to completely dehydrate the object by placing it into absolute alcohol.
• It has a low refractive index (1.481), which can be an advantage to observe certain structures. Other publications consider this low refractive index a disadvantage, however.
• Euparal can be colored green (“Euparal vert”) by adding some copper salt. This can further increase the contrast of specimens stained with hematoxylin.
• Euparal possesses reducing properties and therefore prevents oxidation of some dyes (such as hematoxylin).
• Last, Euparal possesses good fluidity, does not pull strings and handles easily.

I now would like to mention a few advantages of Euparal:

• Unlike other non-water-based mounting media, Euparal does not use the harmful solvent xylene
• Euparal cures relativley quickly
• And on the less serious side: Euparal smells nicely (but don’t inhale, nevertheless!!! Irritant and flammable!).

Euparal does possess some disadvnatages as well:

• Acid sensitive dyes do not keep well, when embedded in Euparal.
• Euparal does have the tendency to shrink a bit. This can introduce air bubbles.
• It is flammable and an irritant. Eye and skin contact must be avoided.

References

[1] Gilson, G. (1906). La Cellule, Vol. 23, pp. 425-432. http://www.archive.org/details/lacellule23lier
[2] Hebb, RG., ed., (1907). Journal of the Royal Microscopical Society. p.501. http://www.archive.org/details/journalofroyalmic1907roya

]]> http://www.microbehunter.com/2010/08/22/euparal-mounting-medium/feed/ 5 http://www.microbehunter.com/2010/08/05/fixing-specimens-for-making-permanent-slides/ http://www.microbehunter.com/2010/08/05/fixing-specimens-for-making-permanent-slides/#comments Thu, 05 Aug 2010 14:18:36 +0000 Oliver http://www.microbehunter.com/?p=2496 Before specimens can be processed for making permanent slides, they may need to be fixed. This step kills the specimen and preserves the structures. It also prepares the specimen for staining. There is no one single method to fix a specimen, too much depends on the nature of the specimen itself and on the subsequent preparation steps.


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.
]]> http://www.microbehunter.com/2010/08/05/fixing-specimens-for-making-permanent-slides/feed/ 5 http://www.microbehunter.com/2010/06/27/the-hemocytometer-counting-chamber/ http://www.microbehunter.com/2010/06/27/the-hemocytometer-counting-chamber/#comments Sun, 27 Jun 2010 08:35:24 +0000 Oliver http://www.microbehunter.com/?p=2459


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.
Disclaimer: This page is intended purely for educational purposes. Do not use this information for medical diagnosis. No guarantee is given for the correctness of the information published in this site.
]]> http://www.microbehunter.com/2010/06/27/the-hemocytometer-counting-chamber/feed/ 10 http://www.microbehunter.com/2010/05/13/the-effect-of-the-mounting-medium-on-image-quality/ http://www.microbehunter.com/2010/05/13/the-effect-of-the-mounting-medium-on-image-quality/#comments Thu, 13 May 2010 10:55:07 +0000 Oliver http://www.microbehunter.com/?p=2426

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
]]> http://www.microbehunter.com/2010/05/13/the-effect-of-the-mounting-medium-on-image-quality/feed/ 1 http://www.microbehunter.com/2010/02/08/stereo-microscope-projects/ http://www.microbehunter.com/2010/02/08/stereo-microscope-projects/#comments Mon, 08 Feb 2010 11:00:15 +0000 Oliver http://www.microbehunter.com/?p=1489 You’ve bought your kid a stereo microscope as a birthday present and now wonder what to look at. Or maybe you are teacher and want to give your class an introduction into (stereo) microscopy and need some specimens to look at (or maybe you bought yourself one, and now want to start out observing…)

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
]]> http://www.microbehunter.com/2010/02/08/stereo-microscope-projects/feed/ 0 http://www.microbehunter.com/2010/01/31/observing-bacteria-under-the-light-microscope/ http://www.microbehunter.com/2010/01/31/observing-bacteria-under-the-light-microscope/#comments Sun, 31 Jan 2010 11:00:08 +0000 Oliver http://microscopy.okim.info/?p=1403
Can one see bacteria using a compound microscope? The answer is a careful “yes, but”.
Generally speaking, it is theoretically and practically possible to see living and unstained bacteria with compound light microscopes, including those microscopes which are used for educational purposes in schools. There are several issues to consider, however.

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.

For pictures of bacteria in phase contrast read the following post: Bacteria in phase contrast

]]> http://www.microbehunter.com/2010/01/31/observing-bacteria-under-the-light-microscope/feed/ 0 http://www.microbehunter.com/2010/01/29/making-a-wet-mount-for-microscopy/ http://www.microbehunter.com/2010/01/29/making-a-wet-mount-for-microscopy/#comments Fri, 29 Jan 2010 11:00:59 +0000 Oliver http://microscopy.okim.info/?p=1402 A wet mount (or temporary mount) is one of the most common ways of observing specimens under the microscope. The sample to be viewed floats in a layer of water which is between the slide and the cover glass. The water performs an important optical function. Without it, the resolution is lower.

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.
]]> http://www.microbehunter.com/2010/01/29/making-a-wet-mount-for-microscopy/feed/ 0 http://www.microbehunter.com/2010/01/27/making-mounts-of-pollen-grains/ http://www.microbehunter.com/2010/01/27/making-mounts-of-pollen-grains/#comments Wed, 27 Jan 2010 11:00:14 +0000 Oliver http://www.microbehunter.com/?p=1480 Permanent slides of pollen grains can be used as a reference for identifying unknown pollen samples. It is therefore important, that the pollen grains remain in an authentic, natural shape. The preparation and mounting of the pollen can introduce artifacts: the pollen may lose some of its pigment, start to shrink and shrivel or absorb water and swell. A careful preparation is therefore necessary.

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.)

]]> http://www.microbehunter.com/2010/01/27/making-mounts-of-pollen-grains/feed/ 0 http://www.microbehunter.com/2010/01/23/an-overview-of-mounting-media-for-microscopy/ http://www.microbehunter.com/2010/01/23/an-overview-of-mounting-media-for-microscopy/#comments Sat, 23 Jan 2010 11:00:46 +0000 Oliver http://www.microbehunter.com/?p=1473
This post gives an overview of different water-based and non-water-based mounting media and their advantages and disadvantages.
Mounting media are needed for making permanent slides. The mounting medium holds the specimens in place between the cover slip and the slide. The choice of the right mounting medium is a separate topic all on its own. There are countless commercial and home-made mounting media available. Which ones should one use? In many cases the microscopist has no choice: some specimens simply require the use of a specific mounting medium, otherwise the structure that one wants to observe is not properly visible. Alternatively, not all specimen types are chemically compatible with the solvents of the medium.

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.

]]> http://www.microbehunter.com/2010/01/23/an-overview-of-mounting-media-for-microscopy/feed/ 2 http://www.microbehunter.com/2010/01/21/choosing-the-right-mounting-medium-for-making-permanent-slides/ http://www.microbehunter.com/2010/01/21/choosing-the-right-mounting-medium-for-making-permanent-slides/#comments Thu, 21 Jan 2010 11:00:18 +0000 Oliver http://www.microbehunter.com/?p=1478
Here I will give an overview of the different factors that may be used to decide on which mounting medium to choose.
There are numerous different mounting media available for making permanent slides. What factors determine the choice of the mounting medium? Here are some possible points to consider.

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).

]]> http://www.microbehunter.com/2010/01/21/choosing-the-right-mounting-medium-for-making-permanent-slides/feed/ 0 http://www.microbehunter.com/2010/01/15/staining-bacteria/ http://www.microbehunter.com/2010/01/15/staining-bacteria/#comments Fri, 15 Jan 2010 11:00:31 +0000 Oliver http://www.microbehunter.com/?p=1466 Here is yet another link to an article from Popular Science magazine. It deals with the isolation, fixing and staining of bacteria. I would not recommend the use of some of the solvents that they use (such as xylol) with children, however. They also describe a blood smear preparation, what I do not recommend for schools (it may not even be allowed in some countries). Still, the article gives a very nice introduction into several preparatory techniques. The article stretches over several pages, click the link at the end of the pages to continue reading. The fact that the article was published 75 years ago, in 1934, does not matter. The preparatory method stayed the same.

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

]]> http://www.microbehunter.com/2010/01/15/staining-bacteria/feed/ 0 http://www.microbehunter.com/2010/01/12/stains-and-reagents-for-microscopy/ http://www.microbehunter.com/2010/01/12/stains-and-reagents-for-microscopy/#comments Tue, 12 Jan 2010 13:30:32 +0000 Oliver http://www.microbehunter.com/?p=1461 I found an article in Popular Science Magazine (see link below) which gives a general overview of different stains that can be used in microscopy. The article divides the stains into three categories:
• 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)

]]> http://www.microbehunter.com/2010/01/12/stains-and-reagents-for-microscopy/feed/ 0 http://www.microbehunter.com/2009/02/19/introductory-microscopy-projects-for-schools/ http://www.microbehunter.com/2009/02/19/introductory-microscopy-projects-for-schools/#comments Thu, 19 Feb 2009 12:52:50 +0000 Oliver http://microscopy.okim.info/?p=1140
Are you looking for simple microscopy projects for classrooms? Here is a list of ideas. Do not forget about safety measures!

Here is a list of microscopy ideas that could be conducted with students and children:
• 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.
]]> http://www.microbehunter.com/2009/02/19/introductory-microscopy-projects-for-schools/feed/ 0 http://www.microbehunter.com/2009/01/26/making-a-soil-culture-for-growing-algae/ http://www.microbehunter.com/2009/01/26/making-a-soil-culture-for-growing-algae/#comments Mon, 26 Jan 2009 21:51:33 +0000 Oliver http://microscopy.okim.info/?p=1057 It may be necessary to grow large amounts of green algae (and other microorganisms) to be used for microscopic observations in schools. A soil culture allows you to enrich various types of algae.

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…..
]]> http://www.microbehunter.com/2009/01/26/making-a-soil-culture-for-growing-algae/feed/ 0 http://www.microbehunter.com/2009/01/18/observing-potato-starch-grains/ http://www.microbehunter.com/2009/01/18/observing-potato-starch-grains/#comments Sun, 18 Jan 2009 19:20:30 +0000 Oliver http://microscopy.okim.info/?p=966
Potato starch grains are an ideal for observation in polarized light and in dark-field. Sample preparation is simple and straight-forward.

Materials: a potato, kitchen knife, slides, cover slips, water, iodine.

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
]]> http://www.microbehunter.com/2009/01/18/observing-potato-starch-grains/feed/ 0 http://www.microbehunter.com/2009/01/08/dry-mounted-permanent-slides/ http://www.microbehunter.com/2009/01/08/dry-mounted-permanent-slides/#comments Thu, 08 Jan 2009 20:59:04 +0000 Oliver http://microscopy.okim.info/?p=911
Wings of insects, small insects and other small specimens do not have to be enclosed in a mounting-medium, they can also be dry-mounted. If they are completely dry, then they will also store for a long time.

Materials: microscope slide, cover glass, adhesive tape which sticks on both sides, sharp cutter knife.

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.
]]> http://www.microbehunter.com/2009/01/08/dry-mounted-permanent-slides/feed/ 2 http://www.microbehunter.com/2009/01/06/processing-specimens-for-microscopy/ http://www.microbehunter.com/2009/01/06/processing-specimens-for-microscopy/#comments Tue, 06 Jan 2009 18:55:49 +0000 Oliver http://microscopy.okim.info/?p=860 Not all microscopic specimens can be observed directly with a compound microscope, many of them need to be brought into a form which is suitable for observation. Different specimens have to be processed differently. This article gives an overview of different preparation methods.

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).
Not all of these methods are suitable for children and beginners. The purpose of this page is to give the reader an overview of possible methods.
]]> http://www.microbehunter.com/2009/01/06/processing-specimens-for-microscopy/feed/ 2 http://www.microbehunter.com/2009/01/05/kiwifruit/ http://www.microbehunter.com/2009/01/05/kiwifruit/#comments Mon, 05 Jan 2009 10:00:54 +0000 Oliver http://microscopy.okim.info/?p=826

Kiwi Fruit: stacked with the software Combine ZP.

Soft specimens can be observed by squashing a small sample between the slide and the cover glass. Here I would like to present: a Kiwi fruit

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.

]]> http://www.microbehunter.com/2009/01/05/kiwifruit/feed/ 0 http://www.microbehunter.com/2008/12/31/fructose-mounting-medium-for-permanent-slides/ http://www.microbehunter.com/2008/12/31/fructose-mounting-medium-for-permanent-slides/#comments Wed, 31 Dec 2008 15:56:44 +0000 Oliver http://microscopy.okim.info/?p=753 Many mounting media for making permanent microscope slides include organic solvents and are less suitable for the use in classrooms, at home and with children. In this article I would like to show you how to make fructose syrup to be used as a safe mounting medium.

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.
]]> http://www.microbehunter.com/2008/12/31/fructose-mounting-medium-for-permanent-slides/feed/ 0 http://www.microbehunter.com/2008/12/27/observing-brownian-motion/ http://www.microbehunter.com/2008/12/27/observing-brownian-motion/#comments Sat, 27 Dec 2008 20:55:15 +0000 Oliver http://microscopy.okim.info/?p=609
Brownian motion is the random movement of particles. It is possible to observe this movement under the microscope.

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.
]]> http://www.microbehunter.com/2008/12/27/observing-brownian-motion/feed/ 0 http://www.microbehunter.com/2008/12/27/staining-yogurt-bacteria/ http://www.microbehunter.com/2008/12/27/staining-yogurt-bacteria/#comments Sat, 27 Dec 2008 11:51:22 +0000 Oliver http://microscopy.okim.info/?p=679

The heat-fixed specimen can be stained using regular (non-toxic) ink. The ink is then carefully rinsed off with water.

School microscopes are often not equipped with phase contrast optics, which would be suitable for viewing bacteria. It is possible to see bacteria also in regular bright field, but the results are better if they are stained. Yogurt bacteria are safe for the use in schools.

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.

]]> http://www.microbehunter.com/2008/12/27/staining-yogurt-bacteria/feed/ 0 http://www.microbehunter.com/2008/12/27/some-safety-issues/ http://www.microbehunter.com/2008/12/27/some-safety-issues/#comments Sat, 27 Dec 2008 07:35:36 +0000 Oliver http://microscopy.okim.info/?p=676 Here are some safety issues to consider when doing microscopy lab work. This is not a comprehensive list.
• 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….
]]> http://www.microbehunter.com/2008/12/27/some-safety-issues/feed/ 0 http://www.microbehunter.com/2008/12/21/observing-leaf-stomata/ http://www.microbehunter.com/2008/12/21/observing-leaf-stomata/#comments Sun, 21 Dec 2008 21:27:24 +0000 Oliver http://microscopy.okim.info/?p=461

Evenly spread a thin layer of water soluble wood glue on the bottom side of a leaf.


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.

It is possible to observe the impression of leaf epidermis cells on white wood glue. The stomata and guard cells are easily visible. The regular shape of the stomata makes it an ideal specimen for practicing drawing.

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.

]]> http://www.microbehunter.com/2008/12/21/observing-leaf-stomata/feed/ 1 http://www.microbehunter.com/2008/12/16/observing-plasmolysis/ http://www.microbehunter.com/2008/12/16/observing-plasmolysis/#comments Tue, 16 Dec 2008 10:59:53 +0000 Oliver http://microscopy.okim.info/?p=238

Make a cut beneath the red layer and firmly press the red part of the onion against the edge of the knife, without cutting yourself...


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.

It is possible to observe the plasmolysis of cells under the microscope. When salt water is added to onion cells, then the cells will lose water due to osmosis, this can be observed.

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.

]]> http://www.microbehunter.com/2008/12/16/observing-plasmolysis/feed/ 0 http://www.microbehunter.com/2008/12/12/determine-cell-size-with-a-slide-projector/ http://www.microbehunter.com/2008/12/12/determine-cell-size-with-a-slide-projector/#comments Fri, 12 Dec 2008 22:35:36 +0000 Oliver http://www.okim.info/microscopy/?p=56 This is one of my favorite lab activities. Onion cells are visualized using a slide projector. Using an internal reference mark, the students can calculate the actual size of onion cells. It does not require the use of microscopic equipment and can be conducted in the normal classroom (lab not required).

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.
]]> http://www.microbehunter.com/2008/12/12/determine-cell-size-with-a-slide-projector/feed/ 0 http://www.microbehunter.com/2008/12/12/making-a-hay-infusion/ http://www.microbehunter.com/2008/12/12/making-a-hay-infusion/#comments Fri, 12 Dec 2008 22:20:18 +0000 Oliver http://www.okim.info/microscopy/?p=49 It is possible to enrich microorganisms such as ciliates by making a hay infusion.

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 (where there is oxygen) for microscopic investigation. If the water is agitated, then the microorganisms are (of course) not able to collect beneath the water surface.

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?

Safety issue: You are cultivating unknown microorganisms. Potentially harmful bacteria could also be in the sample. It is therefore important to observe the rules of hygenics. Use this method at your own risk.
]]> http://www.microbehunter.com/2008/12/12/making-a-hay-infusion/feed/ 0 http://www.microbehunter.com/2008/12/12/growing-paramecia/ http://www.microbehunter.com/2008/12/12/growing-paramecia/#comments Fri, 12 Dec 2008 22:19:30 +0000 Oliver http://www.okim.info/microscopy/?p=46 Paramecia are fresh-water ciliates that make excellent microscopic specimens. They are relatively large and therefore easily observable, even under low magnification. Pond water usually does not contain sufficiently high concentrations of them. For educational purposes it is necessary to enrich them.

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.

Safety issue: You are cultivating unknown microorganisms. Potentially harmful bacteria could also be in the sample. It is therefore important to observe the rules of hygenics. Use this method at your own risk.
]]> http://www.microbehunter.com/2008/12/12/growing-paramecia/feed/ 0 http://www.microbehunter.com/2008/12/12/growing-crystals/ http://www.microbehunter.com/2008/12/12/growing-crystals/#comments Fri, 12 Dec 2008 22:18:38 +0000 Oliver http://www.okim.info/microscopy/?p=44

Vitamin C (ascorbic acid) in polarized light.


Vitamin C (ascorbic acid) in polarized light.


Citric acid (citrate) in polarized light.

Crystals of organic substances make interesting microscopic specimens to be viewed under 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.

]]> http://www.microbehunter.com/2008/12/12/growing-crystals/feed/ 0 http://www.microbehunter.com/2008/12/12/staining-of-onion-cell-nuclei/ http://www.microbehunter.com/2008/12/12/staining-of-onion-cell-nuclei/#comments Fri, 12 Dec 2008 22:13:57 +0000 Oliver http://www.okim.info/microscopy/?p=39

The nuclei of onion cells stain blue.

This is a simple preparatory technique that allows students to observe the otherwise difficult to see nucleus of onion cells. There is no need to employ, possibly harmful, DNA staining chemicals.

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.
]]> http://www.microbehunter.com/2008/12/12/staining-of-onion-cell-nuclei/feed/ 0 http://www.microbehunter.com/2008/12/12/observing-leaf-veins/ http://www.microbehunter.com/2008/12/12/observing-leaf-veins/#comments Fri, 12 Dec 2008 21:34:36 +0000 Oliver http://www.okim.info/microscopy/?p=14

Maple leaf veins after the removal of the soft tissue. The leaf was dried and then scanned at high resolution.



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

This is a simple but somewhat time-consuming preparatory technique. It is possible to isolate the vascular bundles of certain leaves and prepare them for microscopic observation. The prepared leaf veins make an ideal specimen for stereo microscopy. The microscope allows the students to perform a quality-check of their preparation. 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.

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