Frithjof A.S. Sterrenburg (The Netherlands)


(A downloadable version of this article in cross-platform 'rtf' format and more suitable for printing is here.)


Diatoms have been, and still are, favourite objects for microscopists. Although they should always be studied first in their natural, living, state, the fine structure of their frustules on which identification depends only becomes visible after the cell contents (especially the chloroplast) have been removed by “cleaning” ― or rather, destruction by oxidation. Various methods have been described in the professional literature, but since this is expensive, a survey of suitable procedures may be helpful to the general microscopist. The methods described have been selected for their simplicity and have been used in several decades of practice. Comments and suggestions are welcomed at the above e-mail address.




Fossil "rocky" material (diatomites) may require specialised treatment and will not be considered here. Assume that the sample consists of a gathering like:

- sediment from a pond or a coastal marine mudflat

- scrapings from stones or piles

- leaves of aquatic plants

- harvest from a plankton net

The sample will contain ― apart from (hopefully) diatoms ― mineral debris (sand, mud, silt) and organic matter (from plant debris to small animals). The aim is to remove both as well as possible while losing as few diatoms as possible. Mineral debris is removed by sedimentation, organic matter (including the diatom cell contents) is removed by oxidation. Sedimentation and oxidation will be described further on.

Samples must be fixed, e.g. with formalin, immediately after collection. Add about a tenth of the sample volume of 40% formalin and swirl. Investigation of the cell contents requires special fixatives and falls outside the scope of this primer.


General note:


Throughout these notes, the phrase "discard the supernatant" will occur. Do not pour off the supernatant, as this may disturb the sediment and you may lose material. The best way for samples of a reasonable volume is siphoning off with small diameter soft plastic tubing. The speed of draining can be sensitively controlled by pinching the tube. A good method for minute samples is to use a plastic disposable eye-dropper.


Separate the coarse organic debris first. Objects like plant leaves or stems take ages (and gallons of oxidant) to decompose. On the other hand, they cannot be discarded because they may bear rich populations of diatoms (epiphytes!). The same is true for small stones or shells (epilithic species) and even for sand grains (epipsammic species).

Sample pre-treatment aims at detaching the diatoms from such substrata. Procedure:

1) Remove excess water.

If the sample contains a large volume of water, pour the entire gathering ― leaves, stems, sand, algae, shells etc. ― through a household sieve (plastic, mesh about 1 mm), collecting the "fluid" fraction (which also contains the sand/silt). What remains on the sieve (the solid fraction) goes into a generously sized glass beaker. Let the fluid fraction settle completely (check in direct sunlight, the supernatant should not be "milky"), discard supernatant, resuspend the sediment with just enough water and add to the beaker containing the solid fraction. These steps simply remove excess water and may not always be necessary: if the sample does not contain large plant fragments etc., just let the material settle and discard the excess water.

2) Detach diatoms.

Pour some hydrochloric acid (household quality will do) onto the material. Calcareous matter (limestone, shells) will dissolve with production of foam. Stir and leave until foaming subsides, add some more hydrochloric acid until no more foam develops.

Then add enough water to cover the sample by a layer of a couple of cms. Heat gently and let simmer for about half an hour. Beware of fumes! This detaches the diatoms by dissolving mucus. The process can be assisted by scrubbing the leaves, stems or stones with a plastic toothbrush.

Pour through a plastic sieve and collect the fluid, pour some more water over the residue in the sieve to wash out remaining diatoms. Collect this water too, discard contents of sieve.

3) Remove acid.

Let the fluid settle completely, discard supernatant. Add water, mix thoroughly with the sediment, let settle and discard supernatant. Repeat at least three times, the last time with distilled water. This removes the acid and the calcium chloride into which the calcareous matter has been converted.


This pre-treatment procedure ensures that you will not try to oxidize more stuff than is necessary, whilst avoiding major loss of diatoms. You'll always lose a few! At this stage the volume of the sample has become much more manageable and the "raw" material can be stored for further processing if you add formalin. Incineration and mounting (see further on) allow quick examination under the microscope.

Not all materials require such pretreatment. The most unfavourable situation has been assumed (leaves, stems present). Samples like scrapings from stones, plankton catches or rich harvests of periphyton may not require anything but a rinse in hydrochloric acid just to be sure no calcareous matter is left.





Materials from museum collections require special care. If the material has been oxidised already and is stored in distilled water, slides can be made without further processing. There are two special cases, however:

1) Tiny fragments of mica with specks of sample.

2) Samples in glass tubes that have dried out and form a hard cake that sticks to the tube.

Museum material is precious and there may be very little of it. Yet, it may be possible to collect specimens from tubes that are on record as being "empty" ― and indeed look so! Some concessions may have to be made to cleanliness, the purpose is to at least recover whatever diatoms may be left...

1) Mica fragments.

Place the fragment in a small test tube (#1). Add a few drops of concentrated (30%) hydrogen peroxide. Let stand in sunlight for two days or so, or gently heat to about 60° C. for an hour or so (water bath). Swirl gently every now and then. This will detach most of the diatoms from the mica. Take out the mica with pincers and transfer it to another test tube (#2) with some distilled water.

Let the fluid in tube #1 settle completely and the supernatant can then be very gently removed with an eye-dropper, followed by two rinses of distilled water. Check critically (preferably in direct sunlight) that no material sticks to the wall of the tube! If it does, swirl briefly and let settle again.

If the sample is minute, it's easy to lose everything. In that case a slide can be made at this stage, without further rinsing. Some peroxide remains, but this evaporates when you make the slide. To remove any residual organic matter, use incineration (see further on). Note that if the material was of marine origin, rinsing may be unavoidable, to remove the salt.

ESSENTIAL: take out the mica fragment from tube #2, let dry and make a slide of it too! Place the mica on a small blob of mountant on a slide, the surface that carried the material uppermost. Put another small blob of mountant on a cover-slip, turn over the slip, drop it onto the mica and heat gently for an evenly spread and bubble-free layer of mountant. Apply some pressure (place a small weight on the cover before heating). The microscope image will not be perfect, but I've had a case where the only type specimen present in the entire sample had remained stuck to the mica!

The water that remains in tube #2 is added to tube #1 ― it still may contain the single diatom you're looking for ....

2) Dried-out samples.

The natural inclination is to scrape a bit from the surface. NEVER (!) do this, probably the uppermost layer will merely contain the finest silt or the smallest diatoms in the sample and damage will also ensue.

Procedure: add a few drops of concentrated hydrogen peroxide. This softens the cake and after some time you can re-suspend the sample in distilled water. Collect your subsample. For both subsample and the original sample replace the peroxide with distilled water (let settle and rinse, repeat 2x), add some formalin for storage.

CAUTION: if the dried-out sample is of "raw" (unoxidized) material, peroxide may lead to foaming, see "Beware of peroxide" below.





******** BEWARE OF PEROXIDE! *********

A lively reaction will occur when an oxidant is added to hydrogen peroxide (see under "oxidation"). But even when used alone, peroxide has nasty surprises in store.

Adding hydrogen peroxide to unoxidized material may result in quite severe foaming and even "brewing up" of the lot. I have been surprised by truly explosive reactions after adding hydrogen peroxide to 40-year-old dried-out mud cakes! The phenomenon appears to be limited to muddy samples (including dirty periphyton) ― do some muds contain catalytic minerals?

The most treacherous aspect of peroxide is that this "brewing up" may take quite some time (several minutes) to start, when nothing much appears to happen. Then it may chain-react in almost no time to catastrophic intensity.

Constantly keep an eye on a peroxide brew and if foaming seems to get out of hand, pour everything QUICKLY into a MUCH larger wide beaker or better yet: a flat dish. NEVER (!) put a stopper on the tube or you may have to collect the remnants of the sample from the ceiling.










1 For very small samples, see under "Dry (herbarium) materials".

2 Just two methods of oxidation have been selected here. Both yield good results and require the minimum of widely available chemicals. Of course, many more have been described in the literature, but all seem to yield comparable results. Patience is more important than the chemical brew ― do not hurry.

3 Always treat only a small portion of the sample, say a layer of a few mm in a 50 ml beaker, not large quantities. Use wide beakers with a flat bottom and straight (not: conical) sides, do not use small-diameter tubes.

4 If hydrogen peroxide is used, refer to "Beware of peroxide" first. Always use wide beakers or dishes here. Glassware must be heatproof (Pyrex etc.) as it becomes boiling hot very quickly.

5 All oxidants are corrosive and fumes are toxic.

6 Home-cooking of diatom samples knows no immutable laws but requires flexibility, adapting your procedure to the material in question. Samples differ greatly in "difficulty" !



Peroxide method:

1) When the sample has settled completely, discard supernatant

2) Add a small (!) quantity of concentrated (30%) hydrogen peroxide

3) Let stand for several minutes. If alarming foaming already occurs, let this subside and only then add a little more peroxide. Repeat until foaming becomes less violent.

4) If no serious foaming has occurred several minutes after the first small amount of peroxide has been added, add more peroxide until the volume is about 5x that of the original sample.

5) Heat gently (water-bath) for 30 minutes or so (depending on amount of organic dirt). Constantly watch, foaming can still get out of hand.

6) Take the beaker out of the water-bath and place it on the bench, preferably in a wide dish or on a plate. (If the reaction gets out of hand, you can then save the sample if it boils over).

7) Add a VERY SMALL pinch of finely powdered potassium bichromate, just a speck. A violent reaction will occur, swirl and let subside. Only then add a little bit more of the bichromate. Continue this until the reaction has stopped, the contents of the beaker must now be orange in colour.

8) Let settle completely, discard supernatant, resuspend with ample water and repeat this at least twice.

For plankton catches and other samples with very little organic dirt, steps 1-3 may be sufficient.






Sulphuric acid method:

This has the advantage of not causing violent foaming. Check that all calcareous matter has been removed first, otherwise the sample will become totally useless because gypsum crystals will form.

1) When the sample has settled completely, discard supernatant

2) Add concentrated sulphuric acid (battery acid from garages will do) until the volume is twice that of the original sample.

3) Add potassium bichromate. In contrast to the peroxide method, no special care is necessary here, as no violent reaction occurs. Just add enough bichromate to make for a strong solution.

4) Let stand for 24 hours or more, or speed up the reaction in a water-bath (60 degrees or so). Even so, it may take several hours before the sample is clean. The sediment should look greyish and no plant fragments etc. should remain.

5) Let settle completely, discard supernatant and rinse several times as described above.

The sulphuric acid method seems to remove resistant "dirt" somewhat better than the peroxide method, mainly because the oxidation reaction is not as abrupt as with peroxide. But again, the principal point is patience, not the chemistry involved.








1 After oxidation, sedimentation aims at removing as much of the mineral "dirt" (sand, silt, clay) as possible while losing as few diatoms as possible. Especially with very fine silt/clay, this may be difficult and some concessions may be necessary: the cleaner you want the sample to be, the greater the chance of losses. Attempts at getting "nice, clean" samples may be incompatible with quantitative investigations!

2 Especially in sedimentation, you'll have to adapt your procedure to the nature of the sample. There are no standard time-schedules: some stuff settles in a few minutes, other samples may take hours. The only good method is individual checking, see further.

3 It is assumed that the sample has been pre-treated (see under "Pretreatment"), calcareous matter has been dissolved in hydrochloric acid and organic matter has been destroyed by oxidation, and that both the heaviest (very small gravel) and the floating (plant fragments) muck have been removed.

4 When "suspending" or "resuspending" samples, do NOT violently shake the material as this may damage fragile diatoms. Instead, swirl the fluid around for as long as it takes to suspend the sediment.

5 To discard the supernatant, use siphoning as described earlier.

6 It may take many rinses to clean the sample sufficiently. To economise on distilled water consumption, the first rinses can be carried out with tap water. End with at least two rinses of distilled water, see further.

7 Always check (preferably in direct sunlight) that no material sticks to the wall of the beaker glass. If it does, "twist" the beaker quickly by half a turn while it stands on the bench and let settle.




Sedimentation will be time-consuming. Although centrifugation speeds up the work considerably, it has a disadvantage: during sedimentation, the chemicals used for oxidation get a chance to slowly diffuse out. These chemicals have penetrated the diatom valves and are "trapped" in the minute cavities. Centrifugation may leave insufficient time for them to leach out so that oxidant residues may contaminate the sample. "Natural" sedimentation is slow enough for the chemicals to leach out.


Sedimentation procedure:

The literature contains recipes giving standard times for the sedimentation process, but the situation will differ for a tiny, clean epiphytic sample in a 5 ml tube and for a clay sample in a 50 ml beaker! The recipe given here is suitable for any volume, but it should be noted that processing of small samples is always preferable. The aim is to remove the heaviest fraction (called "sand" here) and the lightest fraction (called "clay" here), with the middle fraction (let's hope "diatoms") being retained. Before the "dirt" fractions are discarded, they are examined under the microscope to verify the ABSENCE of diatoms. For this check, darkfield illumination and low power (100x) are ideal.

1) Separate "sand":

1 Suspend sample in beaker #1, let settle briefly (e.g. 20 seconds). Decant supernatant into another beaker ("#2") for further processing.

2 Again add water to "sand" in beaker #1, swirl, let settle briefly, again decant supernatant into beaker #2. Keep "sand" in beaker #1.

3 Resuspend contents of beaker #2, let settle for 20 seconds, decant into beaker #3.

4 Add water to "sand" residue in beaker #2, swirl, let settle briefly, decant into beaker #3.

5 Pour contents of beaker #2 into beaker #1 ― this is the "sand". Beaker #3 will become the sample.


CHECK: add some water to the "sand" in beaker #1, swirl and check a drop of the suspended "sand" for diatoms. If none are present, discard the "sand". If diatoms are still present, repeat entire procedure for the "sand", shortening the settling-time.

When you read this, it may seem like juggling with too many balls (beakers), but it's self-explanatory when you do it...


2) Swirling trick:

Heavy diatoms like Trachyneis, some Centrics or Diploneis spp. may sink almost as quickly as the "sand". What may help is the "swirling trick".

Put a small quantity of the "sand" into a so-called watch glass. Add a few drops of distilled water. Place the watch glass on the bench and GENTLY swirl the fluid by shoving the watch glass with a circular motion over the bench . The sand will collect in the middle. Quickly collect the fluid with a pipette.

Repeat and check for absence of diatoms in the "sand". Try faster or slower swirling speeds. If the diatoms cannot be separated in this manner, do not discard the "sand" fraction but make separate slides of it. Label these correspondingly, e.g. "heaviest fraction".


3) Separate "clay":

Suspend sample and let settle. When the supernatant about 1 cm above the sediment still contains diatoms (collect with pipette and check under the microscope), let settle for some more time. When the water about 1 cm above the sediment no longer contains diatoms, discard the supernatant. Resuspend the sediment and repeat until the discarded supernatant is no longer cloudy.

"Clay" may be impossible to get rid of and you may have to settle for a "dirty" sample in the end.



A phenomenon seen with marine littoral mudflat samples is the following. When tap water is used for sedimentation, the supernatant will no longer be milky after a couple of rinses. Subsequent rinses with distilled water will nevertheless again release large quantities of "clay". Sometimes 6 rinses were necessary and the volume of the sample was eventually reduced to less than a fifth! Apparently some physico-chemical mechanism is involved.


4) In extreme cases: sieving.

After removal of "sand" and "clay", the middle fraction should contain the diatoms with a minimum of dirt. In some cases (notably marine littoral mudflats or saltmarshes) it may be impossible to separate the diatoms from tiny mineral grains in this manner. In that case, sieving through fine-mesh (20 microns) plankton gauze may be the last resort. This will, of course, also remove the smallest diatoms!



To prevent fouling, add some drops of 40% formalin, but to quote the curator of a renowned collection "we have finally succeeded in breeding a formalin-loving fungus". According to some reports, formalin may cause some erosion (only visible in SEM), addition of some hexamethylene-tetramine (buffer) prevents this. Alcohol (70%) is OK but it evaporates quickly unless sealing is really hermetic.

Label with full data (exact location, type of sample, date, collector etc.), use waterproof ink (if you use a printer check that the ink does not come off when wet!!) and varnish the label. Self-adhesive labels and transparent adhesive tape have been found to deteriorate quickly ― sometimes in a decade ― and if long-term documentation is desired it's better to stick to old-fashioned stuff like gummed paper. Secure the cap of the bottle by wrapping with paper-tape. Dipping into molten paraffin is another good (old-fashioned...) method.






Two key issues in making diatom preparations:

1) The slide and the cover-slip must be CLEAN. To test: a drop of water must spread evenly. Simplest: a household cleaner in powder form. Apply some powder to a small wad of moist toilet paper and rub both sides of slide and cover-slip. Rinse under the tap, dry with toilet paper.

NOTE: some luxury types of toilet paper apparently contain something like lanolin or whatever and are totally useless because the glass gets a nice coating of grease... Tissues may be equally useless. Buy cheap ...!

2) NEVER, repeat NEVER, make a preparation on the slide, always on the cover-slip. Preparation on the slide results in serious deterioration of image quality.


To make a slide:

1 clean slide and cover, dry

2 apply a drop (or two) of material to the COVER SLIP, spread evenly (breathing on the slip helps)

3 dry WITHOUT disturbing the slip (diatoms will clot together otherwise), preferably without heating. When the sample is dry, dry by gentle heating for a few minutes to avoid droplets of condensation in the finished slide (moisture is "trapped" in the tiny cavities of the diatom valve).

4 apply a SMALL blob of mountant to the SLIDE

5 turn over the cover slip, place it on the blob of mountant

6 gently heat until all bubbles have escaped and let cool.




For highest contrast use a mountant with high refractive index like Naphrax or Zrax (RI= circa 1.7). Canada Balsam has an RI of circa 1.5 and is not suitable for the more delicate diatoms because it yields very low contrast. For very robust diatoms like heavily silicified Centrics or large Pinnularias, however, Canada Balsam may actually yield better results than high RI media because excessive contrast is avoided.

Varnishing of the edges of the cover-slip prevents "cracking" of the mountant (after decades) but is not absolutely necessary. Canada Balsam slides keep for at least 150 years. Naphrax has been in use for several decades and no deterioration has been reported. Some exotic 19th-century mountants that have been used for slides in museum collections were unstable (crystallization).

NOTE: if a mountant with high RI is unobtainable, do NOT experiment with varnishes etc. but use Canada Balsam if you wish to assure survival of your slides for future reference. Personal examination has shown that well-made 19th-century Canada Balsam slides can still yield satisfactory images if contrast enhancement methods like DIC or phase-contrast are used ― and at least, slides made with Canada Balsam do not deteriorate.





A handy method to quickly examine samples. This is also the only procedure that leaves diatom frustules and aggregates intact, permitting verification of heterovalvarity (e.g. Achnanthes), growth in chains or sessile life-style. Procedure:

1 fix fresh material in formalin (a couple of hours)

2 let sample settle, decant supernatant

3 pour on distilled water, let settle, decant

4 repeat twice

5 apply one (or two) drops to cleaned cover-slip, spread, dry

6 heat cover-slip over (not: "in") a spirit or gas flame, the material will turn black, continue to heat until it becomes grey or white. Take care that the cover does not start to warp or melt!

7 mount as usual

Organic matter (chloroplasts, blue-greens etc.) will be burnt nicely but mineral dirt will remain. You can partially remove this by first applying fractionated sedimentation to the fresh sample ― see under Sedimentation, but beware of loss of material.



   All comments to the author Frithjof Sterrenburg are welcomed.


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