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

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

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

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

Counting chamber seen from the side.

Grid layout of the Neubauer Improved hemocytometer.

Purpose of the hemocytometer

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

Preparing the sample

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

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

Counting the cells

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

Calculating the cell density

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

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

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

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

Things to watch out for

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

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

Objective-related factors

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

Lighting system

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

Specimen-related factors

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

Adjustments of the microscope

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

Maintenance-related factors

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

Stability of the photomicrographic system

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

The checlkist: how to obtain the best image quality

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

Types of cover glasses

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

 

Why cover glass thickness is important

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

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

Importance of the mounting medium

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

How to determine the thickness of a cover glass

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

Ranunculus pollen mounted in air with cover glass.

Ranunculus pollen mounted in water.

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

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

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

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

All observations were made using a 20x achromatic objective.

Results

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

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

What about Glycerin Gelatin (glycerol gelatin, jelly)?

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

Lessons learned

What can we learn from these observations?

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

Some philosophy

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

External Links, References

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

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

Field diaphragm is half open. The reflections are less.

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

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

Advantages of Köhler illumination for photography

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

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

• Advantages of Koehler Illumination
• Adjusting Koehler Illumination

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

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

• Oblique Illumination
• Increasing Contrast using Optical Methods

Making Patch stops for dark-field illumination.

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

Making patch stops of oblique illumunation

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

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

Making Rheinberg filters

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

Cocci in 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.

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.

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.

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.

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.

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

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

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

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

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

I recommend the following steps:

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

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

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

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

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

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

The Koehler illumination must be adjusted before observation:

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

When the polarizing filters are turned into a 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.

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

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

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