Thoughts about diatom cleaning

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JackyMac
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Re: Thoughts about diatom cleaning

#31 Post by JackyMac » Mon Jun 11, 2018 9:39 pm

Using a blowtorch to prepare diatom slides.

I tried it, having read about the method and spoken to people at PMS meetings.
I was very unsuccessful. My little 'blow torch' based on a butane lighter was too hot. The diatoms carbonised nicely but the coverslip buckled and cracked, Actually in one of the attempts it exploded into smithereens!
I used an Aluminium plate,.
I was told this was too conductive.

I haven't had another go yet, but I will and control the heating much more carefully. I will try with different thicknesses and qualities of cover glasses too. I think mine - cheapo cheapo ones - were too flimsy.

If you get it right, apparently, you wash the ashy mix off the buckled coverglass and remount.
We'll see.

Charles
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Re: Thoughts about diatom cleaning

#32 Post by Charles » Tue Jun 12, 2018 12:24 am

I've never tried incineration but probably will, just to see how it looks after.

Hobbyst46
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Re: Thoughts about diatom cleaning

#33 Post by Hobbyst46 » Tue Jun 12, 2018 9:29 am

JackyMac wrote:Using a blowtorch to prepare diatom slides.

I tried it, having read about the method and spoken to people at PMS meetings.
I was very unsuccessful. My little 'blow torch' based on a butane lighter was too hot. The diatoms carbonised nicely but the coverslip buckled and cracked, Actually in one of the attempts it exploded into smithereens!
I used an Aluminium plate,.
I was told this was too conductive.

I haven't had another go yet, but I will and control the heating much more carefully. I will try with different thicknesses and qualities of cover glasses too. I think mine - cheapo cheapo ones - were too flimsy.
I am working on diatom cleaning right now, using GENTLE chemicals, NO sulfuric/nitric/conc HCl, NO conc H2O2, NO centrifuge, and hope to report soon.
In between I attempted incineration, in my small apartment and without a hot plate.

The temperature of the plate and coverslips depends on many factors but here are the results of my measurements, that should provide a useful starting point:
I am using a flat smooth-surface stainless steel plate, size 92x33x2mm, which is the remains of an IKEA shelf supporter. Probably a 303 or 304 SS, not SS316. I won't use aluminum, at least for safety reasons.
To heat with an alcohol burner, I place a tripod, height 140mm, over the burner. On the tripod I place a 150x150mm SS wire mesh (hole size 7mm), which prevents soot somewhat and helps to spread the heat. Please see the photo below.
To heat over an ordinary kitchen propane-butane gas stove, I place the same wire mesh and plate on the stove. The flame diameter is 60-70mm.
I measure the temperature of the plate with a digital thermometer equipped with a type-K thermocouple. I touch the sensor tip to various points on the top surface of the plate. A roughly constant temperature is obtained within 10min.

Results: with the alcohol burner, the temperature of the plate is 200-250C. On the gas flame, the temperature is 300-350. This temperature should suffice for incineration. Whther or not it will soften the frustules and strongly attach them to the glass I do not know.

I ran some incinerations of cheapest Chinese 18mm, 0.17mm coverslips on top of the plate. Dried diatom suspensions on the slips. Incineration was nice and easy, there were no deformations, no cracks, buckling, no sputtering of the coverslips, and the only thing to watch was that the plate must be placed horizontally without any slopes, otherwise the coverslips can slide away. On the gas flame, incineration was complete. With the alcohol burner some black dots remained.

Whether or not the heat caused any disintegration of the diatoms themselves I cannot report, since some frustules were broken after the preliminary chemical cleaning step, prior to the incineration. Many diatoms were undamaged, though.
Please note, that the hot plate remains hot a long time after you put out the flame. I gently push the coverslips (that are on the plate) aside with the tip of a spatula, then handle them using a tweezers.
Hopefully this information will help others. The above data should hold in other "home labs", provided the same composition of cooking gas, the same oxygen content in air, a similar mesh, and especially a SS plate of a similar size. A somewhat thicker plate would take more time to reach constant temperature. A wider/longer plate will need a stronger flame (say, in proportion to the squared flame diameter), to reach a similar temperature.

BTW, a common commercial kitchen metal plate from a kitchenware store, made of thin perforated steel, provides a lower temperature than than 200C, and is not good enough for the incineration.


Error correction (edited June 23, 2018) - the steel plate thickness is 2mm, not 3mm.
Attachments
Alcohol burner+tripod+stainless steel mesh+stainless steel plate.jpg
Alcohol burner+tripod+stainless steel mesh+stainless steel plate.jpg (83.05 KiB) Viewed 8236 times
Last edited by Hobbyst46 on Sat Jun 23, 2018 11:31 am, edited 3 times in total.

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Re: Thoughts about diatom cleaning

#34 Post by Hobbyst46 » Tue Jun 12, 2018 10:46 am

@JackyMac
Hello and welcome. Just discovered your web site/blog. So you are trying sonics/ultrasonics to separate diatoms from mineral sediments or debris? are the diatoms strongly attached to the other particles?

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Re: Thoughts about diatom cleaning

#35 Post by JackyMac » Tue Jun 12, 2018 9:46 pm

Hobbyist46
My diatoms, unlike a lot in the forum, are long dead. Just the siliceous element remaining. And they come embedded in mud, mineral and plant material .
My source is a lump of Oamaru diatomite and I have been following various methods to remove them from the brash,
The ones I have located to date poor things, have, after a wait of 30 million plus years, been washed with plain soap, boiled in Patio Cleaner (16% HCl), rinsed, boiled in H2SO4 - not sure of the strength but it is strong, rinsed and then boiled in HNO3. All of this accompanied by copious washing in distilled water and gravitational settling. With me peeking at the material en-route and taking pictures of the wonders within.

Ideally the only thing left should be the siliceous material, but I have obviously not actually managed to eliminate everything else, yet.

So to answer your question is maybe. Some of them seem reluctant to come out from behind the grot, others float quite nice and free.

If you look at my blog there are pictures from all stages in the - well it must be three years now- experiments.

With the 'final' result from all that boiling I have tried the louspeaker technique - but what I did just seemed to smash them up. They are actually somewhat fragile - at least the interesting ones are. Some Coscinodiscus sp. seems to survive everything, Hemiaulus just fractures leaving only a stump. The triangular ones are, as someone else commented, almost impossible to get whole, although I have seen their valve view when they were floating in acid. Bear in mind I am trying to separate small siliceous structured lumps from among other small siliceous not particularly structured lumps. And it seemed to me that the sonic treatment induces vertical waves in the liquor, not radial ones. I would have thought it desirable to create a radial movement to sort out the heavy and light elements. (simlar to the separation you get in gel electrophoresis)

I had much greater success with a set of filters (250mu down to 5mu) and gravity washing. That got rid of some of the brash.

Right now I am experimenting with picking them, initially from the suspensions. And today with a very bent, but perfectly usable glass needle dry picking Biddulphia from a dried up dish of the residue.
Now I need to make some adhesive to put on the coverslip. Currently I'm looking here because I saw a recipe earlier.

Aside: I do have a suitable chemistry background and a ventilated cabinet in which the most dangerous of these methods were performed. In this case my laboratory was definitely without walls.


Thanks for the welcome, it's always nice to meet like-minded people.

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Re: Thoughts about diatom cleaning

#36 Post by Charles » Wed Jun 13, 2018 11:53 am

JackyMac wrote: Now I need to make some adhesive to put on the coverslip. Currently I'm looking here because I saw a recipe earlier.
I don't know if this is the one you was looking for but I got this recipe from Klaus Kemp:

This is best done with a heater with a magnetic stirrer and your mixing beaker should be in a larger beaker or container of water to prevent scorching.

-Boil 20ml of distilled water (the amount can be 10ml or whatever you want as long as you use equal amounts of Glycerin and IPA. Believe me, 50-60ml of this will last you a lifetime) in a beaker with magnetic stirrer rod in place.
-Activate the stirrer and slowly add the Gum tragacanth until the mixture is the consistency of heavy syrup.
-Turn off the heat.
-Add 20ml of Glycerin (the stirrer should still be going)
-Add 20ml of IPA (90% and the stirrer should be still mixing)
-Add 3-4 drops of Glacial Acetic acid.
-Let all ingredients mix well.
-Filter the mixture into a clean storage container.

The way to get the adhesive onto the coverslip:
Wash you hands really well, especially your index finger. Get a clean coverslip and using a toothpick, pick up some adhesive on the tip and touch it to the coverslip. It should be a very small amount and not a glob. Dip your clean index finger in IPA and let dry. Then rub the adhesive on the coverslip between thumb and index finger until the adhesive becomes tacky. A final straight firm stroke with the index finger straight across the surface of the coverslip and it should be ready for mounting your forms. The single stroke 'smooths' the adhesive out and gives you 'reference' lines to arrange your diatoms.

Transfer the diatoms to the adhesive on the coverslip and after you are finished, breathe on the adhesive and diatoms to set them in the adhesive. Then heat the coverslip under very low heat until you no longer see the adhesive. Then use your mountant to mount the coverslip to a slide.
This adhesive will remain useful on the coverslip for weeks, so if you are making a large arrangement or need to stop and come back to it later, you can. Just be sure to cover it so dust does not get on it. Just by breathing on the adhesive on the coverslip again when done with the mounting and it makes it 'liquid' again.

Hobbyst46
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Re: Thoughts about diatom cleaning

#37 Post by Hobbyst46 » Wed Jun 13, 2018 12:15 pm

JackyMac wrote:Hobbyist46My diatoms, unlike a lot in the forum, are long dead. Just the siliceous element remaining...
Thanks for the detailed information. Solid, mixed, millions of years of age sediments sound like a difficult challenge. I have noticed some sophisticated separation techniques in the literature. However they are all too complex for home use, and include a cleaning step and a sieving step. I add to my future mission list to get some 2 million year sediment rocks, dating from the era when the region was a covered by the archaic sea, and process it with citric acid, EDTA etc.

MicroBob
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Re: Thoughts about diatom cleaning

#38 Post by MicroBob » Wed Jun 13, 2018 3:51 pm

I wrote something abou diatom arrangment in german here : http://www.mikrohamburg.de/Programm/Pro ... 170218.pdf

Your glue base should fit to the mountant you are goint to use..

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Re: Thoughts about diatom cleaning

#39 Post by JackyMac » Sat Jun 16, 2018 12:10 am

I'm currently waiting a delivery of Gum Tragacanth to make up Klaus Kemp's adhesive, so I can hold the picked diatoms and get them in permanent mounts.
Just for 'fun' this is one of the triangular forms I got out today. https://jackymac07.wordpress.com/2018/0 ... tar-prize/
it's a Trigonium articum and even though it's damaged I'm really pleased with it. However i shall probably lose it before I fix in in a permanent mount.

I've amended the link
Last edited by JackyMac on Sat Jun 16, 2018 10:27 pm, edited 1 time in total.

Hobbyst46
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Re: Thoughts about diatom cleaning

#40 Post by Hobbyst46 » Sat Jun 16, 2018 7:30 am

JackyMac wrote:I'm currently waiting a delivery of Gum Tragacanth to make up Klaus Kemp's adhesive, so I can hold the picked diatoms and get them in permanent mounts.
Just for 'fun' this is one of the triangular forms I got out today. https://wordpress.com/post/jackymac07.w ... s.com/2475
it's a Trigonium articum and even though it's damaged I'm really pleased with it. However i shall probably lose it before I fix in in a permanent mount.
Sorry, I could not open the link.

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Re: Thoughts about diatom cleaning

#41 Post by Hobbyst46 » Sat Jun 16, 2018 8:16 pm

Hello everyone,

I attempted a mild diatom cleaning process, for home microscopists who cannot rely on strong mineral acids and oxidizers, nor a centrifuge. I started with enzymatic digestion, using a commercial food additive - capsules of a mixture of amylase, protease, lipase and other constituents. Only slight cleaning was achieved. I did not use Pancreatin, since I could not find it at acceptable price (shipping rates count too).

Recently, Hildebrand and coworkers from California cleaned live diatoms with a detergent (SDS, AKA SLS) + metal complexant (EDTA). I tried it on live diatoms from two sources: (1) MARINE - sand from the bottom of small natural beach ponds. It contained algae detritus and smelled somewhat foul. (2) FRESHWATER - brownish-yellow submersed algae and slime from an ornamental freshwater pond. In both cases I wrung the stuff into water, swirled the muddy water to remove sand, filtered the mass through a 0.5mm mesh cloth, and filtered the filtrate through a 25 micrometer mesh cloth. The retained dark green-brown solids on the cloth were back-rinsed into centrifuge test tubes and rinsed with distilled water several times. All operations were based on gravitational settling of the solids.

The cleaning solution is the mixture: 3.5% Na2EDTA and 2% SDS in water. Add the Na2EDTA to water, then slowly and dropwise add caustic soda (say, a 10% solution) with patient stirring/shaking in between, until dissolution of the Na2EDTA is almost complete. Filter through paper. The final pH should be 6-7. Add SDS and dissolve it by stirring. All is done at room temperature. The resulting cleaning mixture is a colorless, foamy liquid.

The above chemicals are commercial, solid white powders or pellets. Caustic soda must be handled under appropriate safety measures - goggles and rubber gloves. Na2EDTA and SDS are much less hazardous, although eye protection is always recommended with any chemical.

I added the cleaning solution to the dark-colored diatom+detritus suspension at a ratio of about 4:1. Within about 6 h after mixing, the solids mostly precipitated and the supernatant liquid was colored dark brown-orange. Such coloration is indicative of separation of at least the plant pigments from the frustules. Several rinsing cycles after 16 h left a grey precipitate in a colorless, foam-less liquid. Microscope examination showed substantial though incomplete cleaning. Further cleaning was obtained with 3% (pharmaceutical grade) H2O2 (added 2-3:1 to the diatoms) overnight or, alternatively, 3.5% clear home bleach solution for 1h, followed by 6-7 rinses with DW. The final product is a grey precipitate that consists mostly of diatoms (photo below). In contrast to my experience with conc acids and/or H2O2 boilings and centrifugations, many of the diatoms were preserved. Large (200-400micrometer long) pennates still broke into sections.

Attached are some example photos. Diatoms were mounted in either air or PS-CBO. The latter medium is temporary (highly viscous, non-solidifying, sealable with gel nail polish or paraffin), useful up to about 6months. An 40x1.0 iris oil objective was used throughout, with oild immersion, by stopping down the iris diaphragm. The condenser was oild to the slide as well.

The photos are resized to <1024 pixels and cropped. In the brightfield photos, background was whitened in software. Single frames.

In my opinion, these relatively mild cleaning methods can produce clean diatoms at home with relative safety. Occasionally I added short incineration at about 300C. However, no significant improvement was seen. Incineration will be dealt with in the future.

Thanks to helpful comments & suggestions by MicroBob, Charles, zzffnn, desertrat & others along this thread.
Attachments
Pond diatoms cleaned by EDTA+SDS then 3% H2O2.jpg
Pond diatoms cleaned by EDTA+SDS then 3% H2O2.jpg (117.41 KiB) Viewed 8102 times
marine diatoms 40x~0.7 oil air brightfield_1.jpg
marine diatoms 40x~0.7 oil air brightfield_1.jpg (189.54 KiB) Viewed 8102 times
marine diatoms 40x~0.7 oil air brightfield.jpg
marine diatoms 40x~0.7 oil air brightfield.jpg (232.74 KiB) Viewed 8102 times
marine diatoms 40x~0.7 oil air 4jpg.jpg
marine diatoms 40x~0.7 oil air 4jpg.jpg (169.35 KiB) Viewed 8102 times
marine diatoms 40x~0.75 oil air darkfield.jpg
marine diatoms 40x~0.75 oil air darkfield.jpg (80.91 KiB) Viewed 8102 times
Last edited by Hobbyst46 on Mon Jun 18, 2018 9:35 pm, edited 3 times in total.

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Re: Thoughts about diatom cleaning

#42 Post by Hobbyst46 » Sat Jun 16, 2018 8:21 pm

continued - example photos. the small squares or rhomboids with shiny rims are perhaps salt crystals, due to salt traces in spite of the rinses with DW.
Attachments
freshwater diatom 36micron, 40x~0.7 oil darkefield PS-CBO.jpg
freshwater diatom 36micron, 40x~0.7 oil darkefield PS-CBO.jpg (89.57 KiB) Viewed 8101 times
freshwater diatom 160micron, 40x~0.7 oil darkefield PS-CBO.jpg
freshwater diatom 160micron, 40x~0.7 oil darkefield PS-CBO.jpg (61.79 KiB) Viewed 8101 times
freshwater diatom 130micron, 40x~0.7 oil darkefield PS-CBO.jpg
freshwater diatom 130micron, 40x~0.7 oil darkefield PS-CBO.jpg (78.79 KiB) Viewed 8101 times
freshwater diatom 72micron, 40x~0.7 oil darkefield PS-CBO.jpg
freshwater diatom 72micron, 40x~0.7 oil darkefield PS-CBO.jpg (147.17 KiB) Viewed 8101 times

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Re: Thoughts about diatom cleaning

#43 Post by zzffnn » Wed Jul 04, 2018 4:48 am

Inspired by Doron's nice incineration results, I would try the following cleaning protocol:

1) incineration on kitchen stove fire first, to reduce bulk, (see Doron's thread for exact how-to's viewtopic.php?f=10&t=6148), but a large high temperature cooking container may be required for large sample volume;

2) rinse with distilled water twice, to remove metal ions;

3) soak in cold 31% hydrochloric acid (from HomeDepot) for 1 or 2 days, as I am very comfortable using it (now that I have a backyard), this step should also remove organic materials left over from incineration;

change this step to soak with EDTA + SDS (Ethylenediaminetetraacetic acid + sodium dodecyl sulfate), if not comfortable with acids, again, refer Doron's thread viewtopic.php?f=10&t=6148 and this current thread for details;

4) rinse with distilled water three times;

5) soak in cold 30% hydrogen peroxide for 1 or 2 days, as this is the most expensive agent, or soak in 3% hydrogen peroxide instead, if I want to clean diatoms gently or cheaply. Boiling may work better here, per the research paper, but I am too lazy to do long boiling and may forget to turn off stove. This step should further remove organic materials left over from incineration.

6) rinse 3 times with distilled water.

This protocol is for those who is comfortable with strong evaporating acid (30% hydrochloric acid) and is lazy and/or have tendency to forget turning off reactions, but prefer not to handle carcinogen (dichromate). All you have to remember and look after is the incineration step #1; all other steps can be left for much longer without ill effects.

Dichromate protocol can be found in Rob's original thread:
viewtopic.php?f=10&t=3036&p=42559&hilit ... ate#p42559

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Re: Thoughts about diatom cleaning

#44 Post by Hobbyst46 » Wed Jul 04, 2018 5:03 pm

zzffnn wrote:I would try the following cleaning protocol:

1) incineration on kitchen stove fire first, to reduce bulk, (see Doron's thread for exact how-to's viewtopic.php?f=10&t=6148), but a large high temperature cooking container may be required for large sample volume;
Burning a mass of organic stuff at home can be unpleasant and smelly - start with a small sample, please. Also, I suggest that the initial sample be on a large coverslip to facilitate handling into a rinsing tank/tube.
3) soak in cold 31% hydrochloric acid (from HomeDepot) for 1 or 2 days, as I am very comfortable using it (now that I have a backyard), this step should also remove organic materials left over from incineration;

change this step to soak with EDTA + SDS (Ethylenediaminetetraacetic acid + sodium dodecyl sulfate), if not comfortable with acids, again, refer Doron's thread viewtopic.php?f=10&t=6148 and this current thread for details;
If, for any reason, you want to run both options in series, first HCl then EDTA+SDS, rinse in between many times with DW and check with a pH paper that all acid has been removed (pH=7) before adding EDTA.

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Re: Thoughts about diatom cleaning

#45 Post by zzffnn » Wed Jul 04, 2018 5:15 pm

Hobbyst46 wrote:......Burning a mass of organic stuff at home can be unpleasant and smelly - start with a small sample, please. Also, I suggest that the initial sample be on a large coverslip to facilitate handling into a rinsing tank/tube.
...........
I agree that starting small is better, if goal is initial trial and not large scale harvest. In case of very small starting sample, it probably does not matter if incineration should go first or last.

Looking at SEM image of HCL + H2O2 cleaned diatoms from that paper, SDS + EDTA may not be necessary (as an extra following step, assuming one is willing to use the acid). Yes, HCL kills the detergent property of SDS (most acids will destroy detergent property), so it has to be used separately from SDS.

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Re: Thoughts about diatom cleaning

#46 Post by Hobbyst46 » Wed Jul 04, 2018 6:34 pm

zzffnn wrote:
Hobbyst46 wrote:......Burning a mass of organic stuff at home can be unpleasant and smelly - start with a small sample, please. Also, I suggest that the initial sample be on a large coverslip to facilitate handling into a rinsing tank/tube.
...........
Yes, HCL kills the detergent property of SDS (most acids will destroy detergent property), so it has to be used separately from SDS.
Yes! but this point I forgot! I was thinking of another issue: EDTA solubility. Pure EDTA (acid) is a weak acid itself. It will not dissolve in water but in base. Even the disodium salt of EDTA is hardly soluble, you need to add some base to dissolve it. So, since HCl is a strong acid (pH=0) it must be removed completely, otherwise when adding the EDTA+SDS solution, EDTA will precipitate out.
I will try to find a better diversity of epiphytic diatoms to work on. My previous freshwater sample so far was dominated by needle like diatoms which I now think are Synedra.

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Re: Thoughts about diatom cleaning

#47 Post by Hobbyst46 » Thu Aug 02, 2018 1:12 pm

The diatom cleaning adventure, including incineration, is yet alive.
I collected a bunch of freshwater algae, wrung it into water, tried to avoid sand and mud and silt, was rewarded with a very dark brown precipitate, and subjected it to four alternative cleaning methods.
At this initial point, and as a temporary and not entirely reliable indicator, I judge the product quality only by the color of the final precipitate in the test-tube, after treatment. Intuitively, if the final solid mass is light-colored, diatoms are clean. But is the reverse true? not necessarily, because in principle, the removal of organic residues from frustules can differ from the destruction of other organic stuff (and certainly from the destruction of inorganic stuff).

preliminary results:

1. Incineration->conc. H2O2->conc. HCl. Pros: Requires less chemicals than method 2. Cons: darker-color product than in method 2; hazardous chemical
2. conc. H2O2->conc. HCl. Pros: Lightest color product; fast. Cons: Expensive/hazardous chemicals.
3. EDTA+SDS->dil. H2O2 Pros: Harmless(relatively) chemicals. Cons: repetitive treatments, slow; darker-colored product than in methods 1-2.
4. Boiling->dil. H2O2 Pros: Almost no chemicals. Cons: darker-colored product than in methods 1-2.
5. No. 4 above->Home stain remover ("vanish")
Pros: Harmless chemicals. Cons: darker-colored product than in methods 1-2.
6. No. 3 above->conc. HCl Pros: None. Cons: Excessive agglomeration, non-separable combinations of diatoms and detritus diatom Clean product; hazardous chemical
7. No. 5 above->conc. HCl Pros: Clean product Cons: hazardous chemical

Incineration was done by drying the original wet precipitate in a Pyrex petri dish placed on a steel mesh over the stove flame. The result was a brown-grey powder, of volume about 1/4 the volume of the original wet precipitate, hence the subsequent saving on chemicals.
And one more amazing, though trivial, fact: A dark-color mass is not a reliable indicator of non-clean diatoms.

As the total of the above experiments becomes messy, I hope to demonstrate the more important of them in separate posts.

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