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.

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.

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).

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.

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.

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.

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.

• 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.

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

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

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

Method:

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

Alternative method:

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

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

Other Ideas:

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

Troubleshooting:

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