Making a wet mount microscope slide

What is a wet mount?

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

Visit the Microscopy Shop!

>>> USA Shop | Germany Shop | UK Shop | Canada Shop <<<

As an Amazon Affiliate, I earn a commission but it does not cost you more.

Different types of wet mounts

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

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

Advantages and disadvantage of a wet mount

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

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

Disadvantages of wet mounts include:

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

Making the first wet mount: milk!

In your first wet mount we will be looking at a small amount of milk. Take a microscope slide and place a drop of water on the center of the slide. Then transfer a small droplet of milk into the water drop and mix carefully. You can use toothpicks or tweezers to do this. By mixing the milk with the water we make sure that the concentration of the fat droplets in the milk is not too high, otherwise it will be more difficult to see anything. Take a cover glass at its edge sand carefully lower it on the milk water mixture.

Carefully knock against the slide and look at the cover glass. does the cover glass move because it swims on top of the liquid? Does any liquid emerge beneath the cover glass? If this is the case, then you have used too much water or milk and should use some tissue paper to remove any excess water. too much visible water also runs the risk of making the objective wet. Then place the slide on the stage and start observing using the low power objective lens. only when the image is in focus should you use a larger magnification.

When making a wet mount, generally make sure that your sample is of the right size. If the specimen is too large, then prepare it first by cutting or tearing. Place a drop of water on the center of a slide Place the specimen into the water drop and make sure that it is completely covered by water. If necessary add an extra drop of water at the top. Carefully lower the cover glass at an angle on the water, this reduces air bubble formation. Remove excess water with tissue paper, or add more water with a dropper pipette at the side. Evaporating water can be replaced by adding more with a dropper pipette. The cover glass should not float, remove excess water with tissue paper.

Materials and Method

Let us now have a slightly more detailed look at making a wet mount. For making a wet mount you need these materials:

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

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

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

If the specimen is not in water:

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

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

How to prevent drying out

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

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

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 specimen 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.

Thicker specimens require more water between the cover glass and the slide. If too little water is used, then there is the real possibility that the specimen is sandwiched and squashed. Sometimes this is done deliberately in order to limit the movement of certain specimens. Water fleas, for example, can be immobilized this way. In other cases the pressure might be too high and the specimen is damaged.

If you need more space between the slide and the cover glass, then simply adding more water will not always work well. The cover slip then floats on top of the water and every small bump against the microscope or against the table will cause the cover glass to vibrate. A calm observation is much more difficult under these circumstances. What we need is a spacer which holds the cover glass at a defined distance over the slide. There are several ways of making such a spacer.

You can carefully attach small pieces of wax to the four corners of the cover glass. The wax should be first warmed by rolling it between your fingers into a small ball. Small pieces of soft wax are then carefully adhered to the four corners of the cover glass, which is then pressed against the slide, which does not yet contain the sample. If soft wax is not available, then one can also carefully dip the four corners of the cover glass into Vaseline. The wax or Vaseline will both adhere the cover glass to the slide and also increase the space for the sample. The liquid sample can then be applied from the side.

A second possibility is to use two additional cover glasses as a support. Place two cover glasses on a slide, slightly separated. Place a third one across these two, forming a bridge. You can then add your sample with a pipette. The water will be drawn in beneath the three cover glasses and hold them in place.

Last, it is possible to use slides which have a concave indentation. The amount of water that can be held by such a slide can be quite considerable and it may be difficult to observe if specimens float in and out of focus. Be aware that thick layers of water beneath the cover glass can significantly reduce the resolution of the image, especially for the higher magnification objectives.

How can air bubbles in wet munts be reduced?

Needless to say, the preferred method of reducing air bubbles depends on the characteristics of the specimen. Try out the following possibilities.

  • Cover slip placement: Lower the cover slip on the water droplet with an angle. This permits air to escape on one side.
  • Water placement: If the specimen is not fully submerged in the water droplet, add another droplet on top of the specimen before lowering the cover slip. Alternatively you can also place a drop of water on the cover slip itself, before it is lowered on the specimen.
  • Alternative mounting medium: Use a mounting medium other than water. Try immersion oil, nail polish or Euparal as a mounting medium. These mediums are hydrophobic and may therefore interact better with other hydrophobic specimens (such as bird feathers, fur, etc).
  • Break the surface tension: Add a small amount of detergent, such as soap. This will break the surface tension of the water. The water will therefore adhere better to some specimens, thus preventing bubbles. The soap may also harm some water organisms, however.
  • Apply a vacuum: You can remove air bubbles by placing the slide into a vacuum. THe bubbles will expand and move out beneath the cover glass.
  • Dehydrate the specimen: Place the specimen into alcohol. Some specimens will shrink and lose water and air. By placing the specimen into water again, the specimen will take up the water.
  • Remove oil and fat: Wash the specimen in alcohol. This will make the specimen less hydrophobic.
  • Add water: Air bubbles also form when the water starts to dry up. If the air bubble is large and reaches the side of the cover glass, you can add more water from the side of the cover glass.

Further Reading

15 thoughts on “Making a wet mount microscope slide”

  1. The types of microorganisms you want to see are quite large and easy to see, stains are not necessary and if you use stains, you have to use them in low concentrations, otherwise they might harm the cells. Stains are used commonly (but not exclusively) for bacteria, which are rather uninteresting to observe, however. Stains are chemicals that interfere with the biochemical processes and can actually harm the cells. For a science fair, I suppose that the visual observation is more important than the actual measurement and counting of different organisms present. To be on the safe side, so that folks have actually something to see, you might also consider getting some paramecia (aquarium shop) and some protoslo (TM) to slow down the movement of the cells to make them more easily visible (but not necessary). Oliver.

  2. We are working on a science fair projecttin which we want to measure the number of microorganisms we can find in different kinds of pond water. Do you think it would be easier for us to see them if we use a simple stain, or should we be able to see them without it.

  3. Do not take water itself, but rather some slimy stuff from rocks, wood or plants. If you can not see it with your unaided eye, then there is also not much to see under the microscope. 100x is too high a magnification and you will actually see less (because of a smaller area which is observed and because of a lower depth of field.).


  4. Must be something wrong with my pond. I can’t see anything even at 100x. Very, very frustrating. Seemed a lot easier in junior high 30 years ago.

  5. Great presentation! This was wonderfully helpful and made easy to understand. Currently, I am working with my daughter on a homework project with wet-mounts, but I am also a high school science teacher and would recommend and use this site again. Thank you for caring enough to create this site and thank you for giving of your time and energy.
    Well done.

  6. @Nicolas: Check my YouTube channel (icon at the very top of the page). I now have a video on cleaning eyepieces. Oliver

  7. Hello,
    Thanks for the feedback. I’ll make a video maybe during the Christmas break in a few weeks! Check the following post:
    As a general rule: do not clean the scope too much. Use lens paper to wipe off the oil and use synthetic oil (it does not turn hard). Do not use cleaning fluid for eye glasses. An 80:20 ether:alcohol mix has been suggested. It is very volatile and evaporates quickly.

  8. Thank you, great video and explanation. I love your site and it convinced me to buy a microscope for observation.
    So now I’m getting a nice second hand microscope soon, and I would like to know how to use oil, and especially how to clean the objectives and the occulars, and how to take care of the apparatus in general.

    Could you do something like this ?

    Tnak you very much.

  9. Thank you so much! This is just what we need to set up our slides. We’ve been working with our new microscopes with samples that were prepared for us. Now we can get busy with our own. Great explainations!

  10. very good presentation. It should be used in all biology courses as I had to experiment when I took the courses. Maybe the instructors did not know how to make slides.

  11. Well done, Oliver – an excellent appetizer for your new venture.

    I think your use of the video as a microscopy tool is really good.

    My passion is the Diatom. I have studied these little plants for years on Two Tree Island – a little island in the Thames Estuary. I have a number of AVI files of these little plants tearing around the screen. Some are quite slow and ponderous but others are very lively little critters, moving many times their body length in a minute. Would you like to assess them as a suitable contribution ? If so, do you have any means of electronic mailing – such as
    Best wishes Michael

  12. Hello,
    I’ll be adding a few more videos. I also use these videos for instructional purposes with my own students.
    Greetings, Oliver Kim

  13. Thank you for taking time to create these videos! We are using them as part of our home study program in Biology. We truly appreciate you!

    The Newman Family
    Canton, Georgia

Comments are closed.