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

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

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

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

Counting chamber seen from the side.

Grid layout of the Neubauer Improved hemocytometer.

Purpose of the hemocytometer

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

Preparing the sample

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

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

Counting the cells

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

Calculating the cell density

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

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

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

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

Things to watch out for

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

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

Objective-related factors

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

Lighting system

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

Specimen-related factors

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

Adjustments of the microscope

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

Maintenance-related factors

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

Stability of the photomicrographic system

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

The checlkist: how to obtain the best image quality

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

Types of cover glasses

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

 

Why cover glass thickness is important

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

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

Importance of the mounting medium

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

How to determine the thickness of a cover glass

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

What are the things that all types of microscopes have in common?
Microscopes can be very different (see Different types of microscopes. I therefore limit the answer to light microscopes. Things that optical microscopes have in common include:
Objectives, Oculars/eyepieces, stage (carries specimens), light source, focusing system.

Does glycerol mounting cure?
It will dry only up to a point, but will not (and should not) completely dry out. The glycerol will prevent the complete drying. This makes sure that a certain amount of water remains in the sample. A complete drying of the glycerol mounting medium could result in a shrinking and deforming of the specimen. Algae and other water organisms are especially sensitive to this. It is possible to protect the permanent slide by sealing the edges of the cover slip with nail polish.

Why do electron microscopes produce black and white images?
They produce B/W images because electrons do not have a color. Different wavelengths of light, in contrast, do possess colors that we can perceive. It is possible to artificially color electron microscopic images, however. But this does not reflect the “true” colors of the object.

Is pollen a microbe?
No, pollen are not considered microorganisms (microbes), because they are not capable of reproduction. Pollen do not divide to form more pollen. They form sperm cells for fertilizing the plant’s egg cell.

Why does the smell of hay infusions decrease over time?
As a hay infusion ages, different microorganisms start to grow (and others start to die out). Different microorganisms produce different substances which are responsible for the smell.

Is a bacterium too small to be seen under a compound microscope?
No, most bacterial can be seen with compound light microscopes from magnification of 400x up. If the resolution of the microscope optics is not very good, then it will be difficult to see them. You need phase contrast optics to be able to see bacterial well. They may be difficult to see using regular bright-field optics, because bacteria are transparent. Alternatively one may need to stain them. Beginners may have problems distinguishing bacteria from small specks of dirt and dust.

Which type of microscope would be best to use if you wanted a 3-Dimensional view of a virus?
Compound light microscope: It is not possible to see viruses with these microscopes. Resolution and magnification are not large enough.
Transmission electron microscope (TEM): It is possible to see viruses with TEMs, but they provide 2D views.
Scanning electron microscope (SEM): These are the ones that are able to visualize viruses in 3D

Why is it important to apply a cover slip at a 45 degree angle when making a wet mount?
Applying the cover slip at an angle (instead of dropping it down flat on the specimen) pushes the air to the side and therefore minimizes the risk of air bubbles.

Aperture: f/19.9, Exposure time: 10sec.

Aperture:f/22.6, Exposure time: 20sec.

In order to obtain good looking macro images one has to take several measures:

Technical set-up

• A solid, stable tripod: A heavy and stable tripod is absolutely necessary. Even the slightest vibrations (people walking, wind, shutter vibrations, etc.) will translate into a blurry image. I used a tripod designed for heavier video cameras.
• High f-stop values: Macro images generally have a low depth of field. In order to increase the depth of field it is necessary to increase the f-stop values. This again results in longer exposure times.
• Reducing shutter vibrations: Shutter vibrations can be minimized by using the mirror-lock up function of the camera. When the live-view feature of the camera is enabled, the mirror is in the “up” position. During release, the mirror does not have to swing up (because it is already up) and this significantly reduces vibrations. Not every SLR camera has a mirror lock-up feature, however. In this case, one can use a trick to completely eliminate shutter vibrations. Adjust the camera to have a long exposure time (let’s say 20 seconds). Cover the objective with a black cardboard, without touching the objective. Then release the shutter and wait 1 second for the system to stop vibrating. Then remove the black cardboard, this starts the exposure. One second before closing of the shutter cover the objective again. The opening and closing of the shutter (and mirror swing) will then take place while you have the black cardboard in front of the objective. Amateur astronomers use the same technique for taking vibration-free images of the night sky. They cover the telescope aperture with a dark cloth (without touching the telescope, of course).
• Long exposure times: This may come as a surprise for some. Long exposure times can result in more steady images because the duration of the vibrating camera/tripod system (after release) are much shorter than the total exposure time during which a steady image reaches the sensor of the camera.

Composition, artistic aspects etc.

• Sufficient ambient light: To prevent hard shadows, I decided to use the natural, indirect light of the room to take the picture of the rose. This results in longer exposure times, which makes a very stable tripod a necessity.
• Contrast: Make sure that the main object is set off from its background. This can be achieved either by blurring the background, or by making sure that the background has a distinctly different color.
• Exposure time: If you use a black background, then the camera may expose longer than necessary. The black background “fools” the camera into thinking that the image is too dark. This may result in the object being overexposed. Underexpose by 1-2 stops if the main object is too bright.
• Post processing: Crop the image and adjust the colors so that the background is completely black (if you use a black background). This will increase the impact of the picture.

Reproduction

Volvox reproduces both sexually and asexually. During asexual reproduction cells from the equator of the colony move to the inside and divide to form daughter colonies. The daughter colonies will grow and multiply. The mother colony will then rupture and release the offspring.

During sexual reproduction, Volvox colonies form sperm and egg cells (ova). The sperm cells will swarm out to find ova in other colonies. The fertilized egg cells will then form new colonies.

Growing and observing Volvox

Microscopists who are interested in observing Volvox should try to investigate water samples from ponds and puddles. It is also possible to grow Volvox at home. Volvox likes to grow in nutrient-rich water. Dilute some plant fertilizer in water and add some pond water containing Volvox (or other green algae that you want to grow). Place the container on the window sill for several days but prevent direct sunlight as this may cause overheating, and drives out the CO2 for photosynthesis from the water. Alternatively, you can also use a plankton net to catch the colonies.

For making permanent mounts, it’s probably best to use a water-based mounting medium such as glycerin gelatin. Solvent based media may dissolve the chlorophyll of the chloroplasts and may cause the cells to lose water and shrink.

Dandelion macro image.

Lens: Sigma 20-300mm APO DG MACRO (macro up to 1:2)
Camera: Canon EOS 450D
Exposure Time: 2 sec. (long time to minimize vibrations)
Aperture Value: 6.62 EV (f/9.9)
ISO Speed Rating: 100
Focal Length: 300.0 mm

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

Q: What is the principal advantage of an electron microscope over an optical microscope?
A: Electron microscopes have a far greater resolution compared to optical microscopes. Consequently, a much higher magnification is possible. Optical microscopes can magnify up to about 1000x, electron microscopes up to about 1 000 000x.

Q: How to increase resolution of image?
A: The resolution of am image can not simply be increased, once a picture has been taken through the microscope. Information which is not present in the first place can not simply be created. When taking pictures with the microscope, one should make sure that all the parameters are optimized to reach the maximum theoretical resolution. This includes a steady camera-microscope connection, the correct condenser diaphragm setting, the optimum mounting medium, etc.

Q: Parts of the microscope and their functions?
A: This question can not simply be answered in a line or two. I would recommend to watch the video, or read the post: Parts of a Compound Microscope

Q: What are some microbes that you can see under a microscope?
A: Ultimately you can see all types of microbes, provided you have the right type of microscope and use the appropriate technique. Viruses can be seen with electron microscopes, but not with light microscopes. Bacteria can best be seen with light microscopes that use phase contrast optics. Single celled eukaryotes (ciliates, algae etc.) as well as multicellular microorganisms can be seen with bight-field compound microscopes and also with stereo microscopes.

Q: How many different types of microscopes are there?
A: It depends on what system of classification you use and how many subdivisions you include. One common way to classify microscopes is into optical and non-optical microscopes. I already wrote a post on different types of microscopes:

Q: Which type of microscope would be best to use if you wanted a 3-dimensional view of a bacteria cell?
A: Here you have to be careful, the question can be misinterpreted. For true 3D, stereoscopic views two different images are needed.

There are two types of microscopes that provide 3-D (stereoscopic) views:

• Scanning electron microscopes: These devices scan the surface of the object. One single image is produced, which appears 3D (including “shadows” and surface texture). An example image can be found in this article:
• Confocal laser microscopes: These are highly specialized optical microscopes, in which a computer computes a final. In this case it is possible to compute two different pictures, one for the left and one for the right eye. The image is then truly stereoscopic

Q: Compare the kind of image obtained with scanning electron microscope with that obtained using transmission electron microscopy.
A: In short, scanning electron microscope (SEMs) produce images that have a 3D appearance, Transmission electron microscopes (TEMs) produce 2D images.

Q: Why is it desirable that microscope objectives be parfocal?
A: Parfocal objectives are not only desirable, but (in my humble view) a necessity for efficient microscopic work. Parfocal objectives manufactured in such a way that a change in objective will not result in a significant loss of focus. If the image is in focus using a 4x objective, then the image is also in focus when a 10x objective is used. Significant refocussing is not necessary with parfocal objectives.

Q: Part of the microscope that contains the ocular lens
A: One word answer: the eyepiece. Sometimes the terms “eyepiece” and “ocular lens” are used interchangeably, but the eye piece contains more than one lens element.

Q: Different types of microbes
A: The term “microbe” is a colloquial term which refers to organisms (living things) that are too small to be seen with the unaided eye. The term is somewhat unclear, because microscopic insects (and other multicellular organisms) generally are not included. Viruses are not alive and therefore do not quality as microorganisms. Without going into too much detail, microorganisms include prokaryotes (Bacteria, Archaea), microscopic fungi, single-celled algae and protozoa (ciliates and amoeba belong to this category, among others).

Q: Who invented the microscope?
A: Which microscope? There are many kinds. In 1931, Ernst Ruska and Max Knoll constructed the first prototype of an electron microscope. Optical microscopes as we know them today evolved over a longer time period. Many people contributed to the developments. Two notable figures are Antonie van Leeuwenhoek (1632 – 1723) and Robert Hook (1635 – 1703). Leeuwenhoek made single-lens microscopes with which he discovered bacteria. Hook constructed compound microscopes (composed of objective and ocular lenses) and coined the term “cell”.

Ranunculus repens pollen in dark field

Ranunculus repens pollen in bright field

Ranunculus repens

Now a few words concerning sample preparation. The pollen was collected by dusting the flower over a microscopic glass slide. The pollen was briefly dried in open air (about 1 hour) and then permanently mounted in Euparal mounting medium. The standard mounting medium for pollen is Glycerin gelatin, which is water based. I assume that the drying and the Euparal caused the pollen to shrink somewhat, but I yet have to make a comparison with fresh pollen.

Problems with the optics

• Dirty lens: This is due to immersion oil on the optical surfaces, which have collected dust and have hardened.
• Lens kit dissolving: Some lenses are glued together. Flower-like bubbles forming in the lens are an indication that the lens kit is coming loose.
• Fungi on the optical surfaces: This is a problem with microscopes which have been in use in areas of high humidity (such as the tropics). An anti-fungal coating of the lenses may prevent this.
• Scratches or cracks: These can occur if the objective is rotated into the specimen. You can see an extreme example of this in the following post: Dirty microscope objective: Its effect on image quality
• Loss of coating: Excessive rubbing or a wrong cleaning solution may remove the anti-reflective coating of the lenses.

Problems with the mechanics

• Stage drift: In this case, the stage slowly lowers due to its own weight. This can be fixed by tightening some screws.
  Focus difficult to turn: In this case the oil in the gears has solidified due to age and accumulated dust. Do not use force, it may increase the wear on the gears. It’s better to get the device cleaned.
• Mechanical stage difficult to move: Like with the focus knobs a solidified oil makes the stage difficult to move.
• Too much slack: Sometimes there is too much tolerance and turning the focus knobs. There may be too much slack in the gears, possibly due to too much wear.

Problems with electricity

• Old lamp: An old lamp will have a spectrum shifted towards the red. This is a disadvantage for digital photography. The sensors of the camera are very red-sensitive. Use a blue filter.
• Brittle insulation: Old power cables may become brittle and be a hazard.

The image above shows Lumbricus terrestris, the earth worm, in cross section. The red part in the center is the digestive system. You can zoom into the image. The only adjustment done to the image was a color correction. The image was not sharpened.

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

Field diaphragm is half open. The reflections are less.

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

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

Advantages of Köhler illumination for photography

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

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

• Advantages of Koehler Illumination
• Adjusting Koehler Illumination

Rule 1: Be weary about “department store” microscopes

Enthusiasts who want to pick up the hobby frequently encounter their first microscopes in department stores and toy shops. If you are serious about microscopy as a hobby, then I have to disadvise you from purchasing these devices. Microscopes are precision technical instruments and the low cost of toy microscopes simply does not allow them to keep up with the demands of the more serious enthusiast. The resolution of the optics is lower. Stability can also be an isue. It’s better to invest a bit more. You have to contact a retailer which is specialized for microscopes and who sells microscopes to hospitals, schools or research organizations.

Rule 2: Consider carefully if you want a stereo microscope or a compound microscope

Consider your areas of applications. Do you want to observe large or opaque specimens (stereo microscope) or are you more interested in observing small, transparent objects (compound microscope). If you want to do microscopy work with young children, then I would recommend stereo microscopes. See the other post for more info: Which Microscope for Children?. Compound microscopes allow you to observe much smaller specimens, but require you to engage in sample preparation (unless you purchase ready-made specimens).

Rule 3: The magnification is one of the least important criteria

Resolution, stability, extensibility, light intensity etc. also play a significant role. Get the big picture and look at the whole device. Do not get bogged down simply on magnification. Getting a high magnification is the easiest thing to achieve. Simply add a stronger eyepiece, or take a picture and enlarge it on the monitor. Magnification without resolution is meaningless. And a shaky plastic microscope will produce such an unsteady picture that you won’t be able to see much anyway.

Rule 4: Go for standards

Make sure that the microscope has exchangeable objective lenses manufactured according to the “160mm” standard. In this case you have a wide selection of different objectives available from different manufacturers. Infinity corrected optics are an alternative, but there is no universal standard. Some microscopes are not modular in design (“closed system”) and it is not possible to exchange parts later on. When choosing the microscope make sure that you also consider possible future interests and uses.

Rule 5: Consider your current interests

Microscopy does not have to be an entirely new hobby, it can also be a valuable extension of one of your existing pastimes. You may want to evaluate your current hobbies to see which type of microscope fits best.

• Choose a stereo microscope if you are collecting stamps, minerals, rocks, coins, trading cards, smaller antiquities, insects or other objects that are small enough to be placed directly on the stage. Also choose a stereo microscope if younger children should have access to the device.
• Choose a compound microscope of you are keeping a home aquarium, if you want to make specimen preparation (microtoming, staining, etc.) as part of your hobby.

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

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

• Oblique Illumination
• Increasing Contrast using Optical Methods

Making Patch stops for dark-field illumination.

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

Making patch stops of oblique illumunation

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

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

Making Rheinberg filters

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

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

Cocci in pairs and packets of four.

Short rods

Rods-slightly curved cells

The four pictures on the right show different bacterial species in phase contrast.

About phase contrast

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

How the bacteria were prepared

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

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

About the photographs

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

Encapsuled Trichinella spiralis larva in muscle. The larva is cut diagonally.

Longitudinal cross-section.

The larva (circular patterns) in cross-section.

Life Cycle of Trichinella spiralis

• T. spiralis larva are encapsuled in the muscle of the host animal. The pictures on the right show different cross-sections of this stage.
• A person who eats raw or undercooked meat will take these larva up into the body. The larvae are released by the stomach acid and pass into the intestine, where they mature and start to reproduce.
• The offspring larvae travel in the circulatory system throughout the body and settle in the muscles, where they encyst again.

Stacking

Microscopic images generally have a low depth of field. It is possible to take several images of different depth of fields and to combine them in such a way that the final image is sharp throughout. By carefully turning the fine-focus knob a specified amount, it is possible to section through a complete specimen. Care should be taken, however: If too many parts of a transparent specimen are in focus in the final image, then these parts may cover each other, thus reducing the information content of the final image. Two cell organelles which are located behind each other, both being in focus, will cover up each other, and it is not possible to say which part belongs to what organelle.

Stitching

In this method, different overlapping images are assembled together into a larger final picture. While stacking combines the images “vertically”, stitching produces a larger final image by “horizontally” combining them. By stitching, it is possible to overcome the limited field of view. Stitching can be accomplished by using a panorama software. When choosing the software, one should take care that it allows for the combining of images both horizontally and vertically (some only permit for horizontal combination). Microscopic images often do not offer much image complexity. For this reason, the software may have problems assembling the images automatically. It pays off to do a little planning beforehand.

• How large will the final image be? The processing requirements increase significantly with increased image size.
• What camera resolution should be used? The choice of camera resolution has a significant impact on the final image size and required processing power. One should first test, if a high camera resolution is indeed necessary of if it is not simply results in empty magnification. Read: Required camera resolution for photography through the microscope
• How much image overlap should be used? More overlap may make it easier for the software to automatically assemble the pictures, but at the same time more pictures are needed to cover the whole specimen (which again increases work time).

All of the images of the category Virtual Microscope were stitched together.

Background clean up

The optical surfaces (especially the lighting system and condenser optics) are rarely completely free of dust. These disturbances will be present in the image, whether or not a specimen slide is present. It is now possibly to mathematically subtract these disturbances from the image. A picture with and without a specimen has to be taken at the same magnification and using the same exposure time. The empty image (without specimen) is then subtracted from the image containing the specimen.

Alternatively, it is possible to clean the background by selecting the specimen without background and copying it to a new clean background. This system was employed when taking a photograph of the tick (See: Virtual microscope: The Tick). An automatic selection only works well if the specimen’s color or brightness is significantly different from the background.

Increasing contrast

Contrast enhancement is one of the methods which, when done correctly, does not result in any loss of image information content, provided that the image does not use the full brightness spectrum from white to black in the first place. Nearly all photo editing programs contain a “levels” or “histogram” function, with which one can adjust the contrast.

Sharpening

Sharpening the image may subjectively increase image quality, but it will not result in a higher information content. Excessive sharpening introduces artifacts, it may enhance image noise and may enhance irrelevant image components, such as dust and dirt. Before the image is sharpened, it is probably better to increase the contrast. This will sometimes also give an impression of a sharper and more pleasing image.

White balance adjustment

This is a critical adjustment if one wants to obtain reproducible results. Microscopic light will show a different color temperature, based on the intensity level. Turning up the light to a high intensity will also shift the color temperature towards the blue end of the spectrum. A lower intensity setting will increase the red components. The age of the light bulb also shifts the color temperature towards the red. Digital cameras can adjust the white balance automatically, but this may not be a reliable setting, as the camera uses a predefined standard. A specimen which contains many red components, for example, may fool the camera into thinking that the light source is too red. The camera will then shift the color balance toward the blue, which does not reflect the real nature of the specimen. This is a particular problem of colorful images of crystals and specimens which cover the full field of view, without a visible background from the lamp. Some cameras also have a custom white balance function. In this case an empty reference image without specimen is taken. The camera will then use this image as a basis for correcting the white balance of all subsequent images.

Photo editing software also permits users to automatically or manually adjust the white balance. An automatic setting will also take the specimen itself into consideration (just like in the automatic camera white balance setting described above), and the results may not be pleasing. I generally make white balance adjustments manually. In this case, one has to click on those parts of the image that should be considered white, usually the background.

Why bacteria are difficult to see

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

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

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

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

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

The general procedure of making a wet mount

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

Possible problems of making a wet mount

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

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

Mounting techniques

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

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

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

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

Reading materal

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

Comparison of resolution. There is practically no visible difference between 3MP and 12 MP. The limiting factor is therefore the microscope or specimen, and not the camera resolution.

The method of determining the optimum resolution is quite simple:

• Take a low-resolution and a high-resolution picture of the same specimen. I took two pictures each, one with a setting of about 3MP and one with the maximum resolution of 12MP. In both cases, the file compression was low (and JPG image quality high).
• Enlarge the low-resolution image to the size of the high-resolution image. In my case the low resolution (3MP) image had 2256 by 1504 pixels. It was enlarged to 4272 by 2848 pixels (12MP).
• Compare the two images. If the high-resolution image shows details that are not present in the low-resolution image, then this indicates a loss of information when taking a low-res photograph. Otherwise one can safely use the low-res setting of the camera. In this latter case the limiting factor for image quality is not the resolution of the camera, but rather the optics or the specimen.

The result? There was barely any discernible difference between the images. With the specimens that I used, it is perfectly OK to use the smallest camera resolution of 3MP. The limiting factor, so to say, is the optical system of the microscope and/or the specimen itself. For this reason, it is not necessary to choose a high resolution camera setting. Other specimens may deliver finer images, so the differences may become evident then, but from what I found, a small image resolution of 3MP is sufficient.

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

Water-insoluble mounting media that solidify

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

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

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

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

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

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

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

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

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

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

Water-insoluble mounting media that remain liquid

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

Water-soluble mounting media that solidify

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

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

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

Water-soluble mounting media that remain liquid

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

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

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

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

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

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

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

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

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

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

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

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

Optical Microscopes: These microscopes use visible light (or UV light in the case of fluorescence microscopy) to make an image. The light is refracted with optical lenses. The first microscopes that were invented belong to this category. The price of optical microscopes varies from very cheap to nearly unfordable (for the private person, at least). Optical microscopes can be further subdivided into several categories:

• Compound Microscope: These microscopes are composed of two lens systems, an objective and an ocular (eye piece). The maximum useful magnification of a compound microscope is about 1000x.
• Stereo Microscope (dissecting microscope): These microscopes magnify up to about maximum 100x and supply a 3-dimensional view of the specimen. They are useful for observing opaque objects.
• Confocal Laser scanning microscope: Unlike compound and stereo microscopes, these devices are reserved for research organizations. They are able to scan a sample also in depth. A computer is then able to assemble the data to make a 3D image.

X-ray Microscope: As the name suggests, these microscopes use a beam of x-rays to create an image. Due to the small wavelength, the image resolution is higher than in optical microscopes. The maximum useful magnification is therefore also higher and is between the optical microscopes and electron microscopes. One advantage of x-ray microscopes over electron microscopes is, that it is possible to observe living cells.

Scanning acoustic microscope (SAM): These devices use focused sound waves to generate an image. They are used in materials science to detect small cracks or tensions in materials. SAMs can also be used in biology where they help to uncover tensions, stress and elasticity inside biological structure.

Scanning Helium Ion Microscope (SHIM or HeIM): As the name suggests, these devices use a beam of Helium ions to generate an image. There are several advantages to electron microscopes, one being that the sample is left mostly intact (due to the low energy requirements) and that it provides a high resolution. It is a relatively new technology and the first commercial systems were released in 2007.

Neutron Microscope: These microscopes are still in an experimental stage. They have a high resolution and may offer better contrast than other forms of microscopy.

Electron Microscopes: Modern electron microscopes can magnify up to 2 million times. This is possible, because the wavelength of high energy electrons is very small. At the same time, the high energy electrons are pretty tough on the sample being observed. It may take a long time to completely dehydrate and prepare the specimen. Some biological specimens also need to be coated with a very thin layer of a metal before they can be observed.

• Transmission electron microscopy (TEM): In this case, the electron beam is passed through the sample. The result is a two dimensional image.
• Scanning electron microscopy (SEM): Here the electron beam is projected on the sample. The electrons do not go through the sample but bounce off. This way it is possible to visualize the surface structure of the specimen. The image appears 3 dimensional.

Scanning Probe Microscopes: It is possible to visualize individual atoms with these microscopes. The image of the atom is computer-generated, however. A small tip measures the surface structure of the sample by rastering over the surface. If an atom projects out of the surface, then a higher electrical current will flow through the tip. The amount of current is proportional to the height of the structure. A computer will then assemble the position data of the tip and the current to generate an image.

Conclusion: Microscopes can be classified based on the physical principle that is used to generate an image. Different microscopes visualize different physical characteristics of the sample (eg. elasticity can be visualized with acoustic microscopes). Image contrast, resolution (which determines magnification) and destructiveness of the sample are other relevant parameters.

There are several possibilities to reduce the vibrations:

Long exposure time: The microscope-camera system vibrates for the fraction of a second after shutter release. One should therefore use a long exposure time (2-5 sec.). The camera will therefore collect most of the light when the system is steady. Of course, this does not work for moving objects.

Very short exposure time: Alternatively the exposure time can be significantly reduced (about 1/250sec). This is so fast that the vibration will hardly be recorded. This requires much light, however.

Micro flashing: One can also use a flash system mounted above the light source. With experimentation, it is also possible to make a system like this oneself.

The cardboard technique: A similar technique is used in astronomy to make steady images. The camera is set to “bulb” (B) and the shutter is opened. The light intensity has already been adjusted, but the light source is covered by a dark piece of cardboard. The cardboard is then removed for exposure. This method is indeed free of vibrations but requires long exposure times to be practical.

Mirror lock up: Some digital SLR cameras have a mirror-lock-up function. The integrated mirror of the camera can be moved into an “up” position. This reduces the vibrations significantly because the mirror of the camera does not have to swing up during exposure.

Flexible camera-microscope connection: Here, the microscope does not carry the weight of the camera. Rather, the camera is mounted either on a tripod or on another separate system. A vibration of the camera is therefore not passed on to the microscope.

So what methods do I use? I use a combination of several measures: I use a mirror lock-up function, a self-timer (2 sec) and a “long” exposure time of about 2 sec. During the mirror lock-up, it is not possible to look through the viewfinder to focus. Focusing has to be done already beforehand, or one could use the live-view feature to focus and evaluate the picture on the LCD screen before exposure.

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

This the the parachute of a dandelion seed. The seed is not shown, it is attached to the long extension on the right. The leaves of the plant are toothed. The name “dandelion” comes from the French “dent-de-lion” meaning “lion’s tooth”. The microscopic observation reveals that the leaves are not the only part of the plant that have teeth. The fine hair of the parachute also show a tooth-like appearance.

Quite noticeable is the chromatic aberration, which can be seen as a blueish fringe around some of the hair.

This is a darkfield image of a tick. Ticks are blood-sucking arthropods. They possess 8 legs and are not insects, but rather are related to the spiders. Ticks are known to transmit various diseases, such as Lyme’s disease and encephalitis.

For more information on the tick, read the following post: The Tick (Ixodidae).

• 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)

This is a scan of maple leaf vascular tissue, done with a normal flat-bed scanner.

Method: Preparing the leaf was the difficult and time-consuming part. The leaf was boiled for several hours until the cells started to separate. I then carefully lifted the leaf out of the pot and placed it on a plate with water. The soft tissue was then removed with a stiff brush, trying not to damage the delicate veins. The veins were then rinsed in alcohol to remove the remaining chlorophyll, washed in water to remove the alcohol. The alcohol also shrinks the structures, but it will expand again when washed in water. The leaf skeleton was then, pressed and dried. Not all leaves work equally well! The leaves of some plant species are so stiff that the cells do not want to come off when boiled. Don’t waste your time on these leaves.

A confession: Because the stem of the leaves come off very easily, I had to scan it separately and then integrate it into the picture later using some photo editing. I could not scan the veins and the stem at the same time, because it then would not be flat on the scanner. You will also notice that some parts of the leaf are not in focus. This too is because the leaf was not completely flat on the scanner.

For teachers and parents: Boil some leaves with your students/children and let them prepare the leaf skeleton. Then observe the leaf skeleton under the stereo microscope.

For more information on the pine cone, have a look at the following post: Female Pine Cone (Pinus) The specimen size is approximately 20mm from left to right.

If you can not see anything, then you need to install a flash player. The image shows the cross section of the stem of the Aristolochia sipho plant. The image is an inverted (negative) image, and not a dark-field image. Why did I choose to invert the colors? The reason is surprisingly unscientific: it simply looks better… The diameter of the stem is about 6mm across. The annual rings are also visible.

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

Both systems compared.

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

Digitized slide showing vitamin C.

Digitized slide showing vitamin C.

There are several ways to digitize the slides:

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

About exposure time

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

About white balance

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

Macro image of the front lens of a dirty and cracked 40x objective.

A clean 40x objective provides a sharp and crisp image.

A dirty objective produces soft, low-contrasty images. The picture was taken with the above 40x objective.

Most devices were still in a reasonably good condition, with the biggest problems in the mechanics. A check of the optics revealed that most of them were still quite OK, but the 40x objective of one of the scopes was in a particular desolate condition. A macro image of the front lens can be seen on the right. I suspect highly that one of two things happened to the objective:

The objective could have been rotated into immersion oil and was subsequently not cleaned. Students sometimes want to use a lower magnification after they used the 100x oil immersion objective.

A second possibility is, that the “dirt” on the objective is in reality resin for making a permanent slide. Maybe some students attempted to make a permanent slide and used too much resin, and did not wait for the resin to dry out. The front part of the objective was then rotated into the resin.

The origin of the crack in the lens, remains a mystery. The lens is spring loaded , and retracts when crashed into the specimen.

The resulting image was not usable at all. I included two pictures of the same area, one with an intact 40x and one with the dirty and cracked objective from above. I think that the two images speak for themselves.

What do we learn from this? Proper microscope instruction saves money. And be really careful about using immersion oil and resin in the classroom. Don’t even get the students into the position of making such mistakes. In my view, a 100x oil immersion objective is not even necessary for most microscopic work (unless you deliberately want to teach the students different microscopic techniques). Remove the objectives from the microscopes and store them in a safe place.

Microbe hunting – a new term to an old pastime and hobby? The terms seems to be around now for over 70 years, but is still not used widely. Maybe it is time to establish this term a bit more. I have to admit that “Amateur Microscopy?” does sound a bit more “professional” (is this a paradox?), but the sentence “I’m a microbe hunter” flows much easier than “I’m an amateur microscopist”, so maybe this is enough justification to establish that term, even if amateur microscopists observe specimens other than microorganisms as well.

In any case, I herewith propose that the term “Microbe Hunting” be used interchangeably for amateur microscopy. Comments?

Bottom LED light of a stereo microscope. A single LED is sufficient. The glass stage plate has been removed.

Top LED light of a stereo microscope.

Rechargeable batteries deliver an operation time of approx. 50 hours with one charge.

• Long life span: LEDs have a life span of approximately 50 000-100 000 hours. A microscope which is in operation an unrealistic 12 hours per day would have a life span of 20 years+. Unlike tungsten or halogen light bulbs, LED lamps do not need to be replaced and are practically maintenance free.
• Low energy use: The electrical power is converted 80% into light, with nearly no heat production. The efficiency is therefore very high. This makes it possible to operate microscopes with batteries and without a power supply. LED microscopes can therefore also be used in classrooms in which the tables do not have an electrical power outlet. Incandescent light bulbs, in contrast, only convert 20% of the electrical energy to light, the rest is lost as heat.
• Not sensitive to movement: Hot tungsten or halogen light sources should not be moved during operation and when they are hot. Movement reduces the lifespan of the bulb becasue a hot filament breaks more easily when exposed to shock. LEDs are insensitive to movement. Bumping the microscope when the LEDs are on will have no detrimental effect on the LED.
• Can be switched on at full power: Incandescent light bulbs should not be switched on with the light intensity setting at maximum level. Rather, one should slowly increase the intensity after turning the light on. Sending maximum current through a cold bulb will reduce its lifespan. LEDs are insensitive in this respect. Sending a current through
• No color drift with age: LEDs do not significantly change their color or intensity with increasing age. Traditional incandescent light bulbs will shift towards the red end of the spectrum as they age. This is because over time a thin matallic layer will deposit on the inside of the glass bulb. A slight color shift can also be observed with LEDs, this is due to the aging of the resin of the LED.
• Nearly no heat produced: LEDs produce a cool light. LEDs pose no danger of overheating the specimen. This is important when observing water samples with live organisms. A heated specimen will drive out oxygen from the water sample which may possibly reduce movement of some (heterotrophic) organisms.

The following microscope features are probably not needed (or irrelevant) for introductory microscopic investigations in schools. These are the places where one can save money.

• Infinity optics: They are expensive and not yet very common for lower priced educational microscopes. Infinity optics are not interchangeable between manufacturers – to my knowledge there is no single world-wide standard. This can become problematic if individual objectives have to be exchanged, as one is bound to a particular manufacturer. The advantage of infinity optics becomes evident when working with filters or when doing photographic work. For this reason they are commonly employed in research-grade biomedical microscopes. Stay with the 160mm tube length standard objectives.
• 100x oil-immersion objective: This objective requires more experience. The danger is, that students confuse the oil-immersion objective with the regular air-objectives. There is the danger of contaminating these objectives with the immersion oil. Once the specimen slide is covered with oil, it is not possible to use lower magnification objectives anyway, without significant loss of quality. The oil may also stain the sticker of permanent slides. Using oil immersion objectives without oil is also not recommended. The image quality will be very low and students may rotate the objective into the specimen slide while searching for the best focus. If the oil is not properly cleaned, then dust will accumulate over time further reducing image quality. Choose a 100x oil objective if it is an educational goal to teach students to properly use this type of objective or if you need the high magnification (such as for observing chromosomes in cells).
• Phase contrast optics: Generally they are more expensive. These optics will distort the natural color of the specimen as it converts differences in refractive index into difference of brightness. They are useful for investigating unstained bacteria, and it is questionable if this type of investigation is suitable for introductory microscopic courses.
• Köhler illumination: This illumination system is useful when doing photographic work. For students it poses an additional diaphragm to worry about. The system has to be properly adjusted and aligned before use.
• Tension-free objectives: They are used for polarization work. They are expensive and probably they won’t sell them to you anyway if you tell them that you need the microscope for educational or amateur purposes. I’m just mentioning this for the sake of completion.
• Apochromatic objectives: These objectives are corrected for chromatic aberration and are primarily used for photographic work where high image quality is needed. Achromatic objectives are standard for educational work.
• Plan objectives: These objectives deliver an image which is in focus from center to edge. They are commonly used for photomicrographic work. They are expensive and deliver no significant advantages for educational work.

• 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.

Introducing the Microscope – Part 2

Introducing the Microscope – Part 3

A variety of different standards exist:

• Standard slide: 26 x 76 mm
• Geological slide: 75 x 50 mm
• Petrographic slide: 46 x 27 mm
• Thin sections slide: 48 x 28 mm

Microscope glass slides may be modified in a variety of ways:

• They may have a central indentation to carry several drops of liquid.
• They may have a frosted side to allow for easier writing with a marker.
• They may have polished corners to reduce the possibility of injury due to sharp corners.

Cover glasses (cover slips) exist in a wide range of different sizes, square, round, rectangular. Common sizes include:

• 18x18mm
• 20x20mm
• 22x22mm
• 24x24mm
• various rectangular sizes up to 24x60mm to cover nearly the whole slide.

Choose a cover glass that corresponds to the size of the specimen and the slide. The thickness of the cover glass is important, as it has a significant impact on the resolution of the image. The thickness should correspond to the thickness indicated on the objective lens. In many cases, the cover glass is 0.17mm thick, but there is often a small variation even in the same batch. For critical purposes, it may be necessary to measure the thickness of the individual cover glasses to find one close to the desired thickness (use a vernier caliper to determine the thickness).

• 1021 – Ibn al-Haytham (Alhazen) (965 -c.1039): describes the properties of magnifying glass in his Book of Optics.
• 1100s - Translation of Alhazen’s Book of Optics into Latin and spreading of the knowledge into Europe
• 1200s - Development of spectacles (Italy)
• 1590 – Hans Jansen and his son Sacharias Jansen: Invention of the compound microscope
• 1609 – Galileo Galilei (1564-1642): construction of a compound microscope with a convex and a concave lens.
• 1619 – Cornelius Drebbel (1572-1633): presents a compound microscope made of two convex lenses.
• 1625 – Giovanni Faber (1574-1629): coins the word microscope
• 1665 – Robert Hooke (1635-1703): publishes Micrographia, a collection of biological micrographs and the first basic publication dedicated to microscopy.
• 1673 – Anton van Leeuwenhoek (1632-1723): develops single-lense microscopes.
• 1678 – Cherubin d’Orleans: develops a binocular microscope out of two monocular systems
• 1690 – Christiaan Huygens (1629-1695): formulates the wave theory of light and constructs oculars made of two lenses and a diaphragm
• c. 1700 – John Marshall (1633-1725): Develops a microscope base with an illumination system
• 1712 – Christian Gottlieb Hertel: uses mirrors for illumination and constructs a micrometer eyepiece using horse hair for a grid
• 1744 – John Cuff (1708-1772): used a condenser lens to increase light intensity
• 1755 – Georg Adams (1704-1773): constructed microscopes with a revolving nose piece to change objectives.
• 1814 – Joseph Fraunhofer (1787-1826): besides constructing microscopes, his research contributed to the establishment of the wave-theory of light.
• 1830 – Joseph Jackson Lister (1786-1869): is able to correct both chomatic and spherical aberration
• 1834 – William Henry Fox Talbot (1800-1877): develops polarization microscopy and makes photomicrographs
• 1847 – Giovanni Battsta Amici (1786-1873): first person to use immersion objectives
• 1863 – Henry Clifton Sorby: development of a metallurgical microscope to observe meteorites.
• 1873 – Ernst Abbe (1840-1905): discovers the Abbe sine condition, the theory of microscopic imaging and resolution. This was a substantial discovery.
• 1893 – August Köhler (1866-1948): Köhler illumination invented
• 1935 – Frits Zernike (1888-1966): develops phase contrast microscopy, He recieves the Nobel Price in 1953.
• 1955 – George Nomarski (1919-1997): develops differential interference contrast microscopy.

Bright-field microscopy is useful for specimens, which possess a sufficiently high natural color contrast with the background, or for specimens that can easily be stained by dyes. Now, it is possible to increase the contrast by closing the condenser aperture diaphragm. This, however, results in a reduction of the resolution and introduces diffraction artifacts. The natural colors also become less visible, as the whole image darkens. To overcome these limitations of bright-field microscopy, different optical contrasting techniques were invented.

• Dark Field Microscopy: This is one of the easiest and cheapest contrast-enhancing techniques. The main light beam is not able to reach the objective (and therefore the eye), resulting in a black background image. Light is capable of striking the specimen, however. This light is then scattered into various directions, and is also picked up by the objective. The specimen will appear bright on a dark background. Dark-field illumination can be achieved in two ways. Either a specialized dark-field condenser is used, or a so-called patch-stop filter is inserted into the filter holder of the condenser. The patch-stop possesses a central black area which blocks the main light of the illumination system. The patch-stop may not result in a satisfactory image quality for all magnifications, it is advised to experiment with the size of the central black area. For more information: Darkfield Microscopy.
• Rheinberg Illumination: This contrast enhancing technique is closely related to the dark-field method. In this case the patch-stop filter is modified in such a way that the central black area is replaced with a strongly colored, transparent film. The color of the central area of the filter represents the background color of the microscopic image. The peripheral area of the filter possesses a different color. Specimens will then possess the color of the peripheral area. These filters can be easily made by printing the filter using a color printer on an overhead transparency.
• Phase contrast microscopy: This system was invented by Frits Zernike (who received the Nobel Prize for this invention in 1953). Transparent, colorless objects can differ from their surrounding medium (for example water, or the mounting medium) in that they possess a different refractive index. Using bright field microscopy alone, these objects would nearly be invisible. The phase contrast optics of a microscope is able to convert the differences in the refractive index into a difference in brightness. Depending on the system used, the specimens will either appear bright on a dark background, or dark on a bright background. Phase contrast microscopes need special phase contrast objectives and a dedicated phase contrast condenser. In many cases, the phase contrast objectives can also be used for regular bright-field work, with a slight decrease in image quality. Phase contrast microscopy is commonly used for the observation of bacteria, which are otherwise difficult to see.
• Nomarski Differential Interference Contrast (DIC): The theoretical background of this method is complex. The light of the microscope is split up into two beams by a specialized prism which is located beneath the condenser. One beam passes through the specimen, the other beam does not. The two beams therefore have to pass through different refractive indexes and are then allowed to interfere with each other. The result is an image which gives the impression of being three-dimensional. A cell, for example, will appear to be illuminated from the side, with one corner darker than the other. The individual cell organelles will appear to stand out (or be depressed). The 3-dimensional appearance is an illusion, formed by the shadows and highlights. The formed image is similar to oblique illumination.
• Polarization: This contrast enhancing method is commonly used when viewing bifringent speciems, such as starch grains, crystals and cellulose. The light from the illumination system passes through a polarizing filter and then through the bifringent specimen. These specimens are able to interact with the light in such a way, that the light is split into two components. This light continues, and passes through a second polarizing filter, where it is allowed to interfere. The specimens will appear as bright, colorful objects on a dark background. The colors can change when the filters are rotated. Dedicated polarizing microscopes possess a rotating stage and tension-free objective lenses. Possible tension in glass modifies the plane of the polarized light.
• Fluorescence: Certain specimens, such as chloroplasts or cell walls of plant cells, have the tendency to glow in a visible color when flooded with ultraviolet (UV) light. It is also possible to selectively stain the different parts of a cell with flurochomes (fluorescing stains) to visualize them. The UV light can either be passed through the specimen either from the bottom or from the top (“epi-illumination”). It is recommended to use fluorite objectives, otherwise the glass elements, the lenses, will start to glow as well.
• Oblique Illumination: In this method, the illumination system of the microscope is placed-off center. The light strikes the specimen from the side. The specimens appear darker on one side compared to the other side. It is also possible to use a patch stop filter which allows light to pass through only one side. The effect is, that the specimen seems to create a shadow and appears three-dimensional. See Oblique Illumination for sample images.
• Using Color Filters: Color filters absorb the complimentary color. A red filter will result in green chloroplasts to appear dark. A blue “daylight” filter is commonly used as well. It will absorb the red parts of the spectrum and will enhance the contrast of objects that possess a red color. The blue filter will also increase the resolution, as it allows only the passage of the shorter wavelengths.

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…..

Electron vs. Light Microscopes: Basic Differences

There are not many things that these two microscope types have in common. Both electron and light microscopes are technical devices which are used for visualizing structures that are too small to see with the unaided eye, and both types have relevant areas of applications in biology and the materials sciences. And this is pretty much it. The method of visualizing the structures is very different. Electron Microscopes use electrons and not photons (light rays) for visualization. The first electron microscope was constructed in 1931, compared to optical microscopes they are a very recent invention.

Electron microscopes have certain advantages over optical microscopes:

• The biggest advantage is that they have a higher resolution and are therefore also able of a higher magnification (up to 2 million times). Light microscopes can show a useful magnification only up to 1000-2000 times. This is a physical limit imposed by the wavelength of the light. Electron microscopes therefore allow for the visualization of structures that would normally be not visible by optical microscopy.
• Depending on the type of electron microscope, it is possible to view the three dimensional external shape of an object (Scanning Electron Microscope)
• In scanning electron microscopy (SEM), due to the nature of electrons, electron microscopes have a greater depth of field compared to light microscopes. The higher resolution may also give the human eye the subjective impression of a higher depth of field.

Electron microscopes have a range of disadvantages as well:

• They are extremely expensive.
• Sample preparation is often much more elaborate. It is often necessary to coat the specimen with a very thin layer of metal (such as gold). The metal is able to reflect the electrons.
• The sample must be completely dry. This makes it impossible to observe living specimens.
• It is not possible to observe moving specimens (they are dead).
• It is not possible to observe color. Electrons do not possess a color. The image is only black/white. Sometimes the image is colored artificially to give a better visual impression.
• They require more training and experience in identifying artifacts that may have been introduced during the sample preparation process.
• The energy of the electron beam is very high. The sample is therefore exposed to high radiation, and therefore not able to live.

When should one use optical (light) microscopes?

One big advantage of light microscopes is the ability to observe living cells. It is possible to observe a wide range of biological activity, such as the uptake of food, cell division and movement. Additionally, it is possible to use in-vivo staining techniques to observe the uptake of colored pigments by the cells. These processes can not be observed in real time using electron microscopes, as the specimen has to be fixed, and completely dehydrated (and is therefore dead). The low cost of optical microscopes makes them useful in a wide range of different areas, such as education, the medical sector or for hobbyists. Generally, optical and electron microscopes have different areas of application and they complement each other.

Different types of electron microscopes

There are two different types of electron microscopes, scanning electron microscopes (SEM) and transmission electron microscopes (TEM). In the TEM method, an electron beam is passed through an extremely thin section of the specimen. You will get a two-dimensional cross-section of the specimen. SEMs, in contrast, visualize the surface structure of the specimen, providing a 3-D impression. The image above was produced by a SEM.

Different types of light microscopes

The two most common types of microscopes are compound microscopes and stereo microscopes (dissecting microscopes). Stereo microscopes are frequently used to observe larger, opaque specimens. They generally do not magnify as much as compound microscopes (around 40x-70x maximum) but give a truly stereoscopic view. This is because the image delivered to each eye is slightly different. Stereo microscopes do not necessarily require elaborate sample preparation.

Compound microscopes magnify up to about 1000x. The specimen has to be sufficiently thin and bright for the microscope light to pass through. The specimen is mounted on a glass slide. Compound microscopes are not capable of producing a 3D (stereoscopic) view, even if they possess two eye pieces. This is because each one of the eyes receives the same image from the objective. The light beam is simply split in two.

• Tocopheryl acetate: this is a Vitamin E derivative which protects the skin from ultraviolet (UV) light. It is commonly found in ceams and other products that are applied to the skin.
• Allantoin: this substance has a moisturizing effect on the skin and increases its smoothness. It binds substances that irritate the skin and therefore protect the skin.
• Propylene Glycol: this one seems to be a pretty versatile compound. It is used as a moisturizing agent as well as for de-icing aircraft.
• Polyquaternium-7: An anti-static agent. It also forms a film around hair to protect it (it is also found in many shampoos).

]]> http://www.microbehunter.com/2009/01/22/shaving-foam/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 Kim http://microscopy.okim.info/?p=966

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

I also recommend the following post: Virtual microscope: female pine cone (Pinus)

 

 

 

 

Click on Observing a kiwi fruit to read the procedure.

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

Advantages of Drawing Microscopic Images over Photography

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

Drawing Techniques

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

Method:

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

Suitable objects for dry mounting:

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

A specimen for compound microscopy must fulfill several criteria:

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

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

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

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

Method:

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

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

Achromatic lenses: These lenses are designed to correct chromatic abberation for two colors (in contrast to apochromatic systems). Acrhromatic lenses are cheaper and popular in education.

Apochromatic lenses: These lenses are designed to correct chromatic aberration of three colors. They are more expensive and suitable for photographic work.

Arm: The arm connects the base of the microscope to the tube holding the eye piece (ocular).

Base: The bottom part of the microscope – it contains the lamp.

Binocular Head: The top part of a microscope designed to carry two eye pieces.In compound microscopes a binocular head does not give stereoscopic vision (3D).

C-mount: This is an adapter format to connect video cameras to the microscope.

Coarse Focus: Also referred to as rough focus, this knob raises and lowers the stage quickly. It should only be used in connection with the low magnification lenses.

Condenser Lens: This is a lens system which is mounted beneath the stage. It concentrates the light from the lamp so that the image of high power objectives is sufficiently bright. It is also necessary to increase resolution.

Cover Slip: This is a square piece of glass holding the specimen in place. The thickness of the cover slip influences the resolution of the image. Many objectives are manufactured for a cover slip thickness of 0.17mm.

Diaphragm: This is used to control the amount of light entering the objective. The diaphragm is a part of the condenser. By controlling the diameter, resolution and contrast (as well as the amount of light entering the objective) can be controlled. The effect of the diaphragm becomes more apparent at higher magnifications.

DIN Optics: This is a German standard of microscope optics. Objectives that are manufactured according to the DIN standard are interchangeable. The tube length is standardized to 160mm and the threads are standardized as well.

Diopter Adjustment: In binocular microscopes, the diopter adjustment is useful to compensate visual differences of the two eyes. This way it is possible to use the microscope without wearing glasses.

Eyepiece Lens: Also known as ocular lenses, they magnify the image of the objective. The eyepiece is the lens into which a person looks into when observing. The total magnification of a microscope is calculated by multiplying the magnification of the objective by the magnification of the eyepiece. Many eyepiece lenses have a magnification of 10x ot 15x.

Fine Focus: This focus knob moves the stage up and down in small steps. It is used to focus at different layers of the specimens.

Field of View: This is the diameter of the image that you see when looking into a microscope. The larger the field of view (FOV), the more of the specimen is visible. The FOV changes with magnification. The higher the magnification, the lower the FOV.

Focus: This refers to the changing of the distace between objective to specimen to obtain a crisp picture. In most cases the stage is raised or lowered with the coarse and the fine focus knob.

Head: This is the top part of the microscope. It carries the eyepiece(s) and other optical elements. There are several different types of heads: a monocular head is designed to carry only one eyepiece, a binocular head carries two (but does not give stereoscopic vision in compound microscopes) and a trinocular head is designed to carry a camera as well.

Illuminator: This is the light source of the microscope.

Immersion Oil: This is an oil which is used only with oil immersion objectives. A drop of oil is placed on the specimen and the objective is rotated directly into the oil. This way the resolution of the image is increased.

Interpupiliary Adjustment: It is possible to adjust the distance of the eyepieces of stereo or binocular microscopes. If a child only looks through a binocular microscope with one eye only, then this may be an indication that the distance is set to a too large distance.

Mechanical Stage: This type of stage is equipped with a slide holder and two knobs to turn. One knob moves the stage backwards and forwards, the other one moves the slide sideways.

Mirror: A mirror is an alternative to an electrical lamp. Mirrors are often used in field microscopes. It is important that the mirror is not directed towards the sun. This will result in overheating of the specimen and in eye damage.

Mechanical Stage: This type of stage is equipped with a slide holder and two knobs to turn. One knob moves the stage backwards and forwards, the other one moves the slide sideways.

Mirror: A mirror is an alternative to an electrical lamp. Mirrors are often used in field microscopes. It is important that the mirror is not directed towards the sun. This will result in overheating of the specimen and in eye damage.

Monocular Head: This is a microscope head is able to carry only one eyepeice (in comparison to a binocular head, which carries two).

Nosepiece (or revolving nosepiece, turret): This part carries the objectives. It can be rotated.

Numerical Aperture (N.A.): This number is imprinted on the objective lens. It is a measure of the resolving power of the objective (how fine a detail can be seen). The condenser aperture diaphragm should be adjusted to the same value of the N.A. of the objective, to obtain the best results.

Objective Lens: This is a highly magnifying lens system, it is located close to the specimen to be observed. The image of the objective is then magnified again by the ocular lens which is close to the eye.

Oil Immersion Lens / Objective: This is a specially designed objective lens (usually with a 100x magnification) which is used together with immersion oil. The objective is rotated into the immersion oil, with the consequence that the image is of a higher resolution and brightness. Oil immersion objectives have the word “OIL” written on it.

Parfocal: Parfocal objectives belong to one series. It is possible to change the magnification without requiring a refocusing. Parfocal systems are highly recommended for educational purposes.

Pointer: This is an arrow which can be seen when looking through some eye pieces.

Rack Stop: This is a safety measure which prevents you from turning the focus knob too far and from crashing the objective into the slide. You can adjust the rack stop if you need to get closer to the objective.

Resolution: This is the ability of the microscope to show two points as distinct objects. It is the distance at which two points can still be seen as separate points. The higher the resolution, the finer are the details which can be seen.

Reticle: This is a grid which can be seen through some eyepieces. It allows you to make size measurements.

Ring Light: This is a light source in the shape of a ring. The ring arrangement prevents shadows and gives an even illumination. They are used with stereo microscopes.

Slide: This is the glass plate on which the specimen is located. Some slides have one end frosted to allow for easier writing, others have a depression to hold some liquid, still others have smooth corners to remove sharp edges and prevent injuries.

Stage: This is the flat surface on which the slides are placed on. It can be moved up and down for focussing.

Stage Clips: These are clips that hold the slide.

Stereo: In optics, this refers to the ability to see depth (a 3-D image). Two separate eyepieces on a binocular head are not enough. You also need two separate objectives (as is the case with many stereo microscopes).

Student Proofed: Many microscopes used in schools are “student proofed”. This means that students can not remove parts of the microscope because special tools are required. Student proofed microscopes also have safety features like a rack stop and spring-loaded objectives.

T-mount: This is a standardiued adapter ring. It allows you to connect a photo camera to the microscope. Compare this to the C-mount, which is used to connect video cameras.

Tension Adjustment: A proper setting of the tension adjustment prevents the stage to automatically lower itself due to its own weight. If the tension adjustment is set too tight, then it is difficult to focus.

Trinocular Head: This microscope head has three exits, two for viewing (for binocular vision) and a third exit to connect a camera. Some microscopes also allow for taking photographs through a special adapter at the eyepiece, but a trinocular head offers more stability and is to be preferred for photographic work.

Widefield eyepiece lenses: These eyepieces cover a large field of view. More of the specimen can be seen when looking through them. This makes orientation easier.

X: The “X” stands for “times” as used in multiplication. It designates the magnification of the objective and the eyepiece. The total magnification is calculated by multiplying the magnification of the objective with the magnification of the eyepiece.

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.

The following list of terms can also be found in the glossary:

• Condenser: This is a system of different lens elements which is mounted beneath the stage of the microscope. It contains an iris diaphragm which controls the diameter of the light beam. The light beam should be adjusted to be larger or equal to the numerical aperture of the objective in use. Condensers can be moved up and down. The normal operating position is up.
• Base: This is the bottom part of the microscope, it contains the lamp.
• Coarse Focus: Also referred to as rough focus, this knob raises and lowers the microscope stage quickly. It should only be used in connection with the low magnification lenses.
• Eyepiece Lens: Also known as ocular lenses, they magnify the image of the objective. The eyepiece is the lens into which a person looks into when observing. The total magnification of a microscope is calculated by multiplying the magnification of the objective by the magnification of the eyepiece. Many eyepiece lenses have a magnification of 10x ot 15x.
• Fine Focus: This focus knob moves the stage up and down in small steps. It is used to focus at different layers of the specimens.
• Head: This is the top part of the microscope. It carries the eyepiece(s) and other optical elements. There are several different types of heads: a monocular head is designed to carry only one eyepiece, a binocular head carries two (but does not give stereoscopic vision in compound microscopes) and a trinocular head is designed to carry a camera as well.
• Mechanical Stage: This type of stage is equipped with a slide holder and two knobs to turn. One knob moves the stage backwards and forwards, the other one moves the slide sideways.
• Nosepiece (or revolving nosepiece, turret): This part carries the objectives. It can be rotated.
• Objective Lens: This is a highly magnifying lens system, it is located close to the specimen to be observed. The image of the objective is then magnified again by the ocular lens which is close to the eye.
• Stage: This is the flat surface on which the slides are placed on. It can be moved up and down for focusing.
• Stage Clips: These are clips that hold the slide.
• Trinocular Head: This microscope head has three exits, two for viewing (for binocular vision) and a third exit to connect a camera. Some microscopes also allow for taking photographs through a special adapter at the eyepiece, but a trinocular head offers more stability and is to be preferred for photographic work.

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

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

Materials: milk, water, slide, cover slip

Method:

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

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

Method 1:

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

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

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

About the ink:

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

Troubleshooting:

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

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

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

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

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

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

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

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

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

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

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

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

There are two possibilities to achieve a darkfield image:

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

Advantages of darkfield microscopy:

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

Some possible disadvantages of darkfield microscopy:

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

Light microscopes (optical microscopes) that are commonly used in schools come in two flavors – compound microscopes and stereo microscopes (also known as dissecting or binocular microscopes). In research or medicine the range of optical microscopes is naturally larger, some of which are variations or adaptations of the above two types. Strictly speaking, Laser Scanning Confocal Microscopes also belong to the category of light microscopes, but for educational purposes in schools, they are not relevant.

• Compound microscopes: When confronted with the term “microscope”, most people will have a picture of a compound microscope in their head. Compound microscopes are used to observe small, thin, translucent objects. In many cases it is necessary to prepare the specimen before it can be observed.
• Stereo microscopes: These microscopes are designed to view larger, opaque objects. Translucent objects can also be viewed. The magnification is lower than in compound microscopes, but with the advantage of a stereoscopic view.

Important: Compound microscopes with a binocular head should not be confused with stereo microscopes. Compound microscopes are not capable of delivering a stereoscopic (3D) image, even if they have a binocular head.

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

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

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

Method:

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

Troubleshooting:

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

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

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

Issues to consider:

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

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

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

Wikipedia explanation of stomata and guard cells.

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

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

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

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

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

I recommend the following steps:

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

I recommend the following accessories for each microscopic work place:

1. Tweezers: They are useful for placing the cover slip on the specimen and for picking up small specimens (insects, thin cuts, etc.).
2. Dropper: For placing a water drop between the slide and cover glass.
3. Scalpel: Useful for cutting away not needed plant tissue or algae. Do not include for young children.
4. Watch glass: For storing water for making temporary slides.
5. Slides: There are several types available. Some have a frosted side to allow for easier writing, others have rounded edges to decrease the possibility for injury.
6. Blue filter: Useful for compensating the red tint of old tungsten lamps.
7. Cover glasses: obtain those that correspond to the objectives. 0.17mm thickness is standard.
8. Needle: Useful for separating algae or to pick up very small samples of material to be observed.
9. Scissors: For cutting filter paper to remove excess water.
10. Small petri dish: For the storage of specimens that need to be kept in water (plant material, algae, pond water etc.). Cuts of plant material are stored in the dish before they are observed.
11. Plastic tray: For storing the above accessories.

The following accessories are also commonly used, but may not be recommended or necessary for each individual workplace. Safety is an issue as well!

• Razor blades: for making cuts through leaves and stems. Too dangerous to be stored in every workplace, and not always needed.
• Stains: Some stains are toxic, especially those that are used to stain the DNA inside the nucleus of cells (possibly carcinogenic!). Some may stain clothing irreversibly.
• Mounting media: These are used to make permanent mounts. They may contain organic solvents which are not healthy when inhaled. There is also the danger that students confuse them with the immersion oil…..
• Eldermarrow, styrodur or styrofoam: These are used to make thin cuts of plant material. The plant material is squeezed between two layers of this material and then cut. Eldermarrow is recommended. Styrodur and styrofoam also work but they are very tough on the razor blades and will make them dull extremely quickly.

Unplug the microscope. Open the bottom of the microscope to reach the lamp. Pull the lamp out and replace it with a new one. Never touch a new lamp with your fingers. Careful, the lamp may be hot!

A microscope lamp will last for many years. Tungsten lamps have the disadvantage that the color of the light will shift towards the red end of the spectrum and it may be necessary to exchange the lamp even before it burns out. Digital cameras are sensitive to the red end of the spectrum. This shift in color can easily be compensated by using a blue filter (“daylight filter”), but I have also seen older lamps which have a very pronounced red component.

When replacing a lamp, follow the instructions of the manufacturer. The following instructions are for microscopes that have a lamp compartment accessible from the bottom.

• Disconnect the microscope from the mains. It may also be a good idea to wait some time for the transformer to lose its magnetism (danger of electroshock otherwise? I don’t know, never tried it out… Energy is stored in the magnetic field of the transformer.)
• Turn the microscope to its side, taking care that the eyepieces do not fall out.
• Open the lamp compartment on the bottom of the microscope. Flip out the lamp.
• Remove the lamp and replace it with a new one. Do not touch the new lamp with your fingers, wear gloves or use a piece of cloth. Fat deposits will burn into the lamp and result in a darker image.
• Close the lamp compartment.

Lens paper, cotton swabs and an appropriate cleaning solution.

Darkfield microscopy is very sensitive to dust. Notice the bright spots (dust) on the dark background. This dust is not visible in bright field.

The following liquids can be used for cleaning:

• Water: Use water to remove dust from the body, stage, lamp (non-optical parts) of the microscope. Moisten a piece of lint-free cloth and remove the dust. Be careful when cleaning non-metal (plastic) parts, they may be scratched.
• Ether:alcohol (80:20 or 70:30, depending on manufactuer): Moisten a cotton swab or lens paper and clean the optical surfaces. The ether makes the solvent volatile, there is not much time for the cleaning solution to adversely affect the optical coatings and/or the lens kit.
• Cleaning fluid recommended by the manufacturer: Use as specified!

Do not use:

• Cleaning fluids made for computer screens: What do they contain? How aggressive (or not) are they? You do not want to risk the removal of the optical coating of the lenses.
• Cleaning fluids made for eye glasses: They too may contain substances that harm the coating of the lenses.
• Tissue paper or cloth made for eye glasses: they may contain additives that could scratch the optics. Use lens paper instead.
• Compressed air: Many compressed air cartridges for electronics contain additives that will form a milky layer on the optical surfaces. There is also the danger that you will frost the glass (expanding gasses have a cooling effect) and the rapid cooling may not be the best for the lenses. Don’t use them on optical surfaces.
• Other solvents: Acetone, xylol, etc. They may either attack the plastic parts and/or may dissolve the kit holding the lenses in place.
• Cotton swabs made for medical uses: they may contain disinfectants or other additives.
• Denatured or rubbing alcohol: Additives in the alcohol make the alcohol taste bitter. These additives also form a smear on the optical surfaces.

Cleaning the different parts:

• Cleaning the Eyepiece: First blow away the dust. Dust is able to scratch the surface. Then take a cotton swabs (real cotton, not artificial fibers). Moisten the swab with the appropriate cleaning fluid and make sure that there is no excessive liquid on the swab. Clean the eyepiece with the moistened swab. Then wipe the lens dry using lens paper. Do not apply too much pressure.
• Cleaning the Objectives: Immersion oil is first removed by carefully (no pressure!) wiping the objective with lens paper. You can then use the cleaning fluid recommended by the manufacturer. Do not dip the objective into the solvent (this should be obvious, but you never know what some folks are up to…). Apply the solvent to the lens paper.
• Cleaning the Microscope Stage: This can be done with a moist cloth dipped in an appropriate cleaning solution. Scratching of the stage should not be a problem (it’s made of metal), but do take care of the condenser lens!
• Cleaning the Microscope Body: The body can be cleaned either with a soft cloth and water, or dust can be removed by using pressured air (not those for electronics, which contain additives!)
• Cleaning the Condenser: The same things apply as for the objective and the eyepiece. First remove the dust, then use an appropriate cleaning solution with cotton swab and wipe dry with lens paper.

The effects of dust: Store the microscope in a dust-free environment and/or cover the microscope with a dust cover when not in use. Dust has several adverse effects:

• It disturbs the quality of the image. Especially dark-field illumination is very sensitive to dust. Even the smallest dust grains show up.
• Over the years the oil in the gearing (focus knobs, condenser knob, and of the mechanical stage, etc.) will collect dust and start to solidify, making the mechanics difficult to operate and increasing wear.
• Dust grains located between the condenser and the slide may scratch the condenser lens when moving the slide. The distance between the slide and the condenser lens is very small if the condenser is moved into the highest position.

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

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

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

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

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

The Koehler illumination must be adjusted before observation:

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

An opened field iris diaphragm increases the width of the light beam. This setting is used for low magnifications (large field of view)

An closed field diaphragm decreases the width of the light beam.

A closed field diaphragm.

• Uniform specimen illumination: Before the advent of Koehler illumination, a diffusing glass was placed over the light bulb. This had the disadvantage of reducing the light spectrum.
• Reduction in specimen heating: A heated specimen increases evaporation of the water beneath the cover slip and also reduces the dissolved oxygen, a potential problem when viewing live organisms.
• Reduction of light reflections in photographic work: Excessive light is eliminated reducing reflections inside the optical system. As a consequence the contrast of the photographic image increases.

The Koehler field diaphragm is designed to restrict the light beam only on this part of the specimen which is actually observed. Especially at high magnifications only a very small part of the specimen needs to be illuminated.
 
 

An opened condenser diaphragm increases the angle of the light beam.

A closed condenser diaphragm decreases the angle of the light beam. Notice that opening and closing does not change the width of the beam where it exits the condenser.

Many modern course microscopes are equipped with a condenser and an associated condenser diaphragm. The purpose of the condenser is to concentrate the light onto the specimen, its diaphragm regulates resolution, contrast and depth of field. There is a trade-off to consider:

• When the condenser diaphragm is closed, then the depth of field and contrast increase and
• the image will lose resolution and becomes darker.

It is up to the microscopist to find the optimum setting of the aperture diaphragm, but for optimum resolution the setting of the diaphragm should be more or equal to the numerical aperture of the objective (this value is printed on the objective).

Many beginning microscope users prefer to generally close the aperture diaphragm all the way. The image possesses more contrast and subjectively appears more crisp. The image looks less “washed-out” The increased depth of field also makes it easier to find the plane of focus.

There is, however, the danger of introducing optical artifacts:

• Dust grains on the cover slip or on the optical surfaces start to become more pronounced and may give the impression that they are part of the specimen.
• Structures become more pronounced than they actually are.
• The larger depth of field may result in some structures covering up other structures that are in front of, or behind them.
• The larger depth of field causes structures overlap more and it becomes more difficult in determining the layer in which they are located.
• Last but not least, the maximum possible resolution of the objective is not used.

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

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

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

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

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

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

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

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

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

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

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

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

Method – Plasmolysis.

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

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

What do the numbers and abbreviations on an objective mean? Especially when buying used microscopes from research laboratories or hospitals a basic knowledge of the text written on the optics can become handy. You don’t want to buy things that you don’t need.

• A or ACHRO (depending on brand): This signifies that the objective is an achromat. This means that chromatic abberration was corrected for 2 colors (in contrast to the expensive APOchromatic lenses). Achromatic lenses are those most commonly found in education, they are the cheapest.
• PLAN: These objectives produce an image which is in focus from edge to edge. They are used for photographic work and are more expensive.
• PLANAPO: This refers to a planapochromatic objective. It produces a flat image (in focus from edge to edge) and it is has a chromatic abberration correction for 4 colors. Expensive and not needed for educational work.
• PLANFL: A Planfluorite objective. A bit less expensive than the planapochromats but also not as fully corrected.
• 160: This represents the standard tube length of 160mm. Objectives with this standard are interchangeable between manufacturers.
• 0.17: This represents the thickness of the cover slip to be used in mm. Coverslips with a deviating thickness will result is an image of lower resolution.
• 4, 10, 20, 40, 100: This represents the magnification of the objective. The total magnification is calculated by multiplying the magnification of the objective with the magnification of the ocular (eye piece), which is usually 10x. The magnification is also indicated by the ring colors:
  • red: 4x or 5x
  • yellow: 10x
  • green: 20x
  • blue: 40x, 50x or 60x
  • white: 100x
• OIL: This designates an oil immersion objectives. Do not immerse non-oil objectives into immersion oil!
• WI: Water Immersion. Here water is used instead of oil.
• 0.65 (etc): This is the numerical aperture. This value indicates the angle to which an objective is able to receive light. This value also determines the resolution of the system. For maximum resolution, the iris diaphragm should be set to a value equal or larger than the numerical aperture of the objective in use.
• NCG or NC: These abbreviations stand for “No cover glass”. These objectives are designed to be used without a cover glass. They are useful in the medical area where blood smears etc. are observed.
• LWD or ULWD: These abbreviations stand for “long working distance” or “ultra-long working distance”. These objectives are able to work with a large specimen-objective distance and are used for specific applications.
• P, POL or SF: These objectives are designed to be used for polarization microscopy. The objectives are strain-free (SF) and will therefore not modify the polarization of the light. They are not necessary for simple polarization microscopy conducted in classrooms.
• PL or NH: These are designation of objectives used for phase contrast microscopy. A PL (positive low) objective produces an image of a specimen which is darker than the background, a NH (negative high) objective produces an image which is brighter than the background.
• NIC or DIC: Nomarski Interference Contrast or Differential Interference Contrast objectives produce an image of a specimen which appears to be slightly 3 dimensional. If you use a filter to achieve oblique illumination, then the result will look similar.

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

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

Before buying a second hand microscope, take care of the following points. The list is certainly not complete, but should give an overview of the things to look out for:

• Appropriate optics: Make sure that the microscope is equipped with the objectives that are needed for your task. Many microscopes from hospitals and research institutes are equipped with phase contrast or plan apochomatic objectives. These are expensive and not required for educational work.
• Are the focusing knobs easy to turn or has the lubrication oil already solidified? Do not force-turn the focus knobs, it will increase the wear on the gears.
• Does the condenser move freely up and down, or has the lubrication oil already solidified? Does the condenser stay up, or is it pulled downwards by its own weight?
• Does the iris diaphragm open and close without problems, or has the lubrication oil already solidified?
• Were the non-oil immersion objectives dipped into immersion oil? They are not designed for this.
• If the objectives are spring-loaded, does the front part of the objective retract properly when pushed in? Or was the objective covered completely with immersion oil, now solidified?
• Are there any fungi growing on the lens optics? This may be a problem of microscopes used in warm and humid areas.
• Does the stage stay where it is, or does it move down due to its own weight?
• Does the microscope generally make a good impression?

Additional points to consider for schools:

• If you want to equip a whole classroom with used (or new) microscopes, make sure that they are from the same series. Students are then confronted with the same device each time and do not have to re-learn the peculiarities of each instrument – more productive lab work. It also makes makes it easier for the teacher to explain the handling of the microscope. If you buy used instruments, then you may not be able to obtain a whole classroom set at once (unless another school, college or hospital replaces its equipment at once).
• Make sure that the used microscopes have the appropriate optics installed and not specialized objectives! You need achromatic bright-field objectives. Many research microscopes come with phase contrast objectives installed, however. This type of optics also can be made to work like bright-field objectives but they are more expensive and you don’t want to spend money on things that you don’t need.
• In recent years the so-called infinity optics became increasingly popular, especially in research. Be careful – infinity objectives are not compatible with the “traditional” DIN 160mm systems, and infinity optics from different manufacturers are also not compatible with each other. When purchasing infinity systems (if you can afford them…), be aware that the optics can possibly not be exchanged with other microscopes that you or your school owns.

Occasionally people ask me for advice about which type of microscope to buy as a present for their children. I once responded to a newsgroup question making a very strong point in favor for stereo microscopes for young children (approx. 5 years of age), and I would like to reiterate these points below. Read the article “Different Types of Light Microscopes” for a description of similarities and differences between the different microscope types. The following section reflects my own personal opinion on this issue.

In any case, I do not recommend the purchase of “toy” microscopes. If you invest a little more you are able to obtain a “real” instrument with substantially better image quality and flexibility, one which will retain the interest of the child (and parent!!) for a longer time. And especially for children a good image quality is necessary. An experienced microscopist may be able to interpret the “dark washed-out blob” as a cell, but children need crisper and clearer images to maintain their fascination – my personal opinion. There is the danger of disappointment if they do not see similar images as those printed on the box, and I am almost certain that many “toy microscopes” are not capable of keeping their promise. But this is my personal opinion, and the quality of these devices certainly varies as well.

Some of these microscopes are also sold with unrealistic magnifications up to over 1000x. Please understand that toy microscopes are useless at this magnification, for a range of reasons:

• Resolution is too low: The object that you want to see is magnified 1000x for sure, but you only see a washed-out blob with no detail.
• Stability is low:. There is a good reason why microscopes are made of metal and why they are heavy. Every vibration (walking) is magnified as well and transferred to the microscope.
• Image is dark: A high magnification requires a high light intensity. Many of these microscopes are not capable of delivering the required light intensity.
• And: bad depth of field, optics not corrected for lens errors, etc. etc.

I have to admit that even “toy” microscopes vary greatly in quality (and price). If you want to buy one of these, then I would recommend you not to give magnifications above 200x or 400x much weight and to read appropriate reviews beforehand. A cheap plastic scope with 1000x magnification is unrealistic. I have already seen some better quality “toy” microscopes but the price difference to a microscope manufactured according to the international DIN standard was not too big. The bottom line is that the child should enjoy working with the instrument.

Compound or Stereo Microscope?

Instead of simply listing the pros and cons of each type, I’ll make life easy by giving you two simple rules:

The younger the child the more you should tend towards stereo microscopes.

and:

If you intend to purchase a compound microscope, make sure that it works with the DIN standard. This allows for an exchange of objectives and guarantees a minimum quality.

There are many points that speak for stereo microscopes for young children, they are not only more “child friendly”, and more forgiving and easier to handle:

• The subjective visual impression of 3D samples (flies, hair, rocks etc.) can be quite fascinating. The view is, in contrast to compound microscopes, upright (!). A big advantage for orientation.
• Little to no sample preparation required for many objects. There is no need to cut and slice the specimens into the required thickness, even though this is possible as well. There is no need to prepare specimen slides with cover-slips. We use stereo microscopes in our school, and as a first introduction, we gave our students a post card and made them look at the colored dots that compose the image – fast, simple with an immediate result.
• Stereo microscopes have a low magnification, often not more than 40x. This means that there is less abstraction “from the real world”. A fly looks like a fly, only much bigger and more impressive. For a compound microscope you need to take the fly apart first and examine the individual parts, it’s too thick otherwise to be observed.
• Stereo microscopes also allow for an observation of non-transparent objects like rocks, fingernails (the dirt is pretty interesting…), skin, plant leaves etc. Stick a whole earth worm under the microscope and see how it looks like. Directly observe a dish of pond-water. If the child is already collecting rocks, insects, stamps, coins, etc. then a stereo microscope is the natural extension to observe these collected items.
• Some stereo microscopes also allow for a change in magnification, by zooming. This is not a necessity, though.
• Decent stereo microscopes can be cheaper than compound microscopes, because they are less complex.
• Stereo microscopes need less time for instruction. More instruction time needed for compound microscope. With compound microscopes, if you use a higher magnification and then turn the coarse-focus-adjustment knob into the wrong direction, you run the risk of ruining both sample and objective because you smash the objective into the specimen. Stereo microscopes have a large sample-objective distance.
• Stereo microscopes have a higher depth of field. It is therefore much easier to find what you are looking for.

Stereo microscopes also possess certain disadvantages:

• There may be some samples that you or your child is interested in but requires a higher magnification. For example, if you want to watch the nucleus of cells, then you are better off with a compound microscope. It is also not possible to observe bacteria, they are simply too small. As a comfort, a compound microscope with regular bright-field optics also does not allow you to view living bacteria, as they are transparent, you need expensive phase-contrast objectives (labs use them). Alternatively the bacteria have to be stained first, and then I doubt that novices will be able to recognize them as bacteria.
• Sample preparation may indeed be one of the activities that a child may be interested in, but stereo-microscopes do not require much preparation, while it is necessary in compound microscopes.
• Some younger children may have problems viewing through both eye-pieces (they can look through one of them if they want to).
• And possibly the biggest “problem”: There is the myth that microscopes have to magnify very much in order for the person to see much. Children may be disappointed if they hear that their stereo microscope only magnifies up to 40x, if their friend has a department store (“toy”) microscope which magnifies 1250x. This is where education comes into play – magnification is not everything, and a high magnification does not mean that one sees more, resolution also counts. I want to guarantee you that you won’t be able to see much at 1250x.

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

Method:

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

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

Method:

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

Troubleshooting:

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

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

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

Materials: Fresh pond water, wheat grains, glass beakers

Method 1:

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

Troubleshooting:

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