Diatom selection experience

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Hobbyst46
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Diatom selection experience

#1 Post by Hobbyst46 » Sat Jun 20, 2020 2:32 pm

Hello all,

To make some reasonable slides from chemically cleaned diatom samples, that are plagued with silt and sand (of the same size distribution as the frustules), I pick single diatoms and transfer them to a cover slip. My only target is a slide of clean frustules only, without silt, but not necessarily defined patterns. Rather, a "randomly arranged slide".

I am quite familiar with the literature on mounting strew and arranged slides. Have prepared strew slides in the past.

The tip (pun): For a picking bristle, since animal eyelashes or hairs and pulled glass needles are inaccessible to me, I have tried fibers from extra-soft toothbrushes. Of several toothbrush brands, most fibers were found to be uniformly thick (and too thick). However, some brands have nicely tapered and pointed to a tip of a diameter 10-20 um. They are translucent, so the frustule is visible, and being made of nylon, pick the diatoms fairly well. The (expected) difficulty is to release the diatom from the tip onto the coverslip. Especially since I did not apply any adhesive to the coverslip.

My question to experienced diatomists : supposing that I just place the frustules on the bare clean coverslip, without any adhesive, then proceed to mount in Pleurax. Will the frustules stay attached more or less where they were placed, or anyway, somewhere on the coverslip ? or will they be swept by the bubbles when the resin boils, and disappear ?
Last edited by Hobbyst46 on Thu Aug 13, 2020 12:55 pm, edited 1 time in total.
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Re: A tip and a question about a slide of clean diatoms without silt

#2 Post by Zuul » Sat Jun 20, 2020 3:09 pm

Hobbyst46 wrote:
Sat Jun 20, 2020 2:32 pm
since animal eyelashes or hairs and pulled glass needles are inaccessible to me
You don’t have eyelashes? (Sincere apologies if you suffer alopecia.)

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Re: A tip and a question about a slide of clean diatoms without silt

#3 Post by Hobbyst46 » Sat Jun 20, 2020 3:16 pm

Zuul wrote:
Sat Jun 20, 2020 3:09 pm
Hobbyst46 wrote:
Sat Jun 20, 2020 2:32 pm
since animal eyelashes or hairs and pulled glass needles are inaccessible to me
You don’t have eyelashes? (Sincere apologies if you suffer alopecia.)
Literature says PIG'S eyelash.
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Re: A tip and a question about a slide of clean diatoms without silt

#4 Post by Wes » Sat Jun 20, 2020 4:03 pm

These are two problems I'm currently dealing with.

To get rid of the silt and other fine particles two very experienced diatomists told me to get a sieve/mesh (15-20 µm pore size). If the particles are sticking to the diatoms after cleaning or if you get clumps of diatoms as often happens after boiling de-calcified diatomite in sulfuric acid you can "shock" them by 30 seconds boiling in 0.5% w/v NaOH followed by immediate neutralization (runs the risk of dissolving the frustules if left for too long).

If anyone knows of a good source for a the relevant pore size sieve/mesh please share. I looked on ebay but the ones there are not good (or too expensive).

Regarding the diatoms sticking to the coverslip I was told to flip the slide upside down (with some sort of support as not to let the coverslip make contact with the hot plate) after reaching the temperature of 180˚ C. You leave it like that for a couple of hours and then you should have a nice strew slide. If you can get a hot plate with a temperature regulator that would help you eliminate the boiling of the mounting media by slowly raising the temperature to allow solvent evaporation.

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Re: A tip and a question about a slide of clean diatoms without silt

#5 Post by Charles » Sat Jun 20, 2020 4:56 pm

You will have frustules migrating to the ends of your coverslip without a mountant.
I haven't tried this, but, it should work. Try dissolving a very small bit of pleurax (dip a end of a needle) in alcohol and place a drop on a coverslip and let dry. After placing your frustules on the coverslip, add another drop of the alcohol with trace pleurax and let dry by gentle heating. Then add your pleurax and mount the coverslip.

For cleaning silt and other particles, it's best to treat it with HCl and Sulfuric acid, which will get rid a lot of your detritus. If your diatoms are the same size as other particles you are trying to get rid of, put a sample in a beaker or test tube with distilled water, swirl or agitate with a pipet, let settle for about a minute and pour off the top sediment with the diatoms. The heavier particles will settle first. After you have done that for a few times, this time after swirling and agitating, let settle for about 30 minutes and siphon off the water and repeat a few times. I would keep the sediment and siphoned water until you check to make sure most of the diatoms are not still in those portions.

To get the frustules clean, I add a flake or two of pure soap, agitate and mix well, let settle for 30-45 min and siphon off and rinse, repeating until there is no more soap.

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Re: A tip and a question about a slide of clean diatoms without silt

#6 Post by MicroBob » Sat Jun 20, 2020 5:43 pm

Hi together,
I use this stainless mesh in different mesh sizes: https://www.ebay.de/itm/Edelstahlsieb-E ... SwRfxcKWKH
I have used the 25µ mesh for cleaning steps. Some small diatoms will be lost but there is a price to pay. With even smaller mesh sizes it will be increasingly difficult to get the water through the mesh.

To remove silt it should work to first use sieves to separate the material for size. in a given size the silt will settle quicker than the diatoms. Whithout sieving before the problem will be that big diatoms settle as fast as small silt particles. I have used this method before but haven't monitored it precisely enough to describe where the limitations are.

@Charles: That method with the thinned Pleurax sounds very good!

Bob

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Re: A tip and a question about a slide of clean diatoms without silt

#7 Post by Hobbyst46 » Sat Jun 20, 2020 7:08 pm

Thanks a lot Wes, Charles and Bob, for the comments.

A few words about how I cleaned the diatoms.

First, the raw sample - from floating plants and sunken stones in a small river; I thought that the sample would be mainly diatoms, but found it to mainly consist of silt and sand - as fine mud.
Cleaning consisted of several steps, including preliminary sieving through a nylon 25um mesh plastic cloth. Sieving was only partially efficient.
Several cleaning steps followed: EDTA+SDS, boiling in persulfate (sodium PS/ammonium PS), bleach, many rinsings with water after each step, checking the pH etc.
I do not have the safety facilities to work with strong acids or 30% peroxide, so I stick with gentle procedures. Nevertheless, these steps might have changed the dispersion of the silt and sand particles (big lumps were disintegrated, maybe). Any way, the product is a mixture that cannot be fractioned by sieving.
Also, the final amount of diatoms is fairly small, so sedimentations are liable to "eliminate" them from site.

Thus I started picking them one by one:
1) Inspired by Charles, adopted the stereo microscope for the task.
Edit: sideways illumination, that creates quasi-darkefield, is better than transmitted illumination, since some diatoms are practically invisible in brightfield.

2) Fabricated a "mechanical finger" to lift them, and out of an old mechanical stage, made an X-Y slide holder. Installed these gadgets on the stereo base, around the base glass window.

3) Found a tapered pointed fiber, from an extra-soft toothbrush, for the finger (as outlined above). This fiber is flexible and, at least for a cetrales species and a long needle-like species (Synedra maybe) works fine !

So - Bob, I ordered the steel meshes, 25um and 42um, for future work. Will construct sturdy sieves.*
Charles - I like the suggestion about using Pleurax as cement. Worth trying.
Wes - I will try to slow down the heating.

* Edit: one disadvantage of most inexpensive sieves (be they nylon or steel) is that they are woven. Non-woven would be better, but apparently expensive.
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Re: A tip and a question about a slide of clean diatoms without silt

#8 Post by Hobbyst46 » Wed Jul 01, 2020 8:44 pm

Update:
The "bristle" made of a toothbrush hair works, but is somewhat too thick.
I sought a thinner bristle. And found a glass fiber. Not one that was pulled from a tube. Just a fiber. Glass fibers, in the form of glass wool, is a know heat isolator. So I just pulled a tiny bit out and managed to cut a single fiber, length of ~ 2mm I think, diameter 10um. Not pointed or tapered - just a uniformly 10um diameter fiber. I glued it to the mechanical finger with nail polish.
And started the practice of playing golf with diatoms. At least with this centrales it works well.
Below is the setup. The stereo microscope is a 10X-40X zoom, with a 2X Barlow lens and 10X23 eyepieces. On the front of the stage - a slide holder, made from an old spare mechanical stage, and a replacement spring-loded left jaw that catches the slide. The tip of the mechanical finger is shown (not the glass fiber of course...). In the back - a quasi-dark field illuminator, made from a LED strip (12 LEDs), and shaded with a blackened metal tilted "roof" to protect me from the bright LED light.
The slide in this case is a "five well" slide. They are not wells actually, only circles bordered by a whallow white layer of something, to provide five separate spaces for samples. Their depth is probably of the order of 10 um. I show here a catch of a diatom and its placement in a new clean space, in isolation (in fact, social distancing).

The camera is a cheap 5MP USB 2 eyepiece camera, inserted into one of the two eyepiece tubes.
The photos compare bright field view to dark field view. I finally realized that room light through the other eyepiece was affecting the dark field. So for the last photo I covered it with black cloth.
Attachments
Stereo microscope stage, ready for the isolation.jpg
Stereo microscope stage, ready for the isolation.jpg (87.16 KiB) Viewed 3656 times
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Re: A tip and a question about a slide of clean diatoms without silt

#9 Post by Hobbyst46 » Wed Jul 01, 2020 8:46 pm

(continued) the focus is not the best, sorry. When the fiber is raised, either it or the diatoms on the slide could be in focus - not both.
In photos 2 and 4, the round diatom is carried at the tip of the fiber.

I must say, for me, it was easier to find this fiber than to pull an elegant tapered tip glass pipette from a tube, like some experts do. The challenge, of course, was to attach the 10 um diameter fiber to a #32 hypodermic needle - which by comparison is huge !
Attachments
1) strew source. 40X mag. brightfield. fibre above, at ~1 oclock.jpg
1) strew source. 40X mag. brightfield. fibre above, at ~1 oclock.jpg (55.3 KiB) Viewed 3655 times
2) fiber above strew source. 40X mag. brightfield.jpg
2) fiber above strew source. 40X mag. brightfield.jpg (59.97 KiB) Viewed 3655 times
3) strew source. 40X mag. dark field. fibre above, at ~1 oclock.jpg
3) strew source. 40X mag. dark field. fibre above, at ~1 oclock.jpg (46.4 KiB) Viewed 3655 times
4) fiber above strew source. 40X mag. dark field.jpg
4) fiber above strew source. 40X mag. dark field.jpg (43.47 KiB) Viewed 3655 times
5) isolated frustules. fiber end touches the slide. 75X mag. second eyepiece covered. dark field.jpg
5) isolated frustules. fiber end touches the slide. 75X mag. second eyepiece covered. dark field.jpg (26.45 KiB) Viewed 3655 times
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Re: A tip and a question about a slide of clean diatoms without silt

#10 Post by Hobbyst46 » Wed Jul 01, 2020 8:53 pm

Charles: I tried to attach the glass fiber with sugar water (fructose syrup actually) but was too impatient to wait for drying, so I switched to nail polish.
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Re: A tip and a question about a slide of clean diatoms without silt

#11 Post by Charles » Wed Jul 01, 2020 10:30 pm

Excellent work around Doron!

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Re: A tip and a question about a slide of clean diatoms without silt

#12 Post by Hobbyst46 » Thu Jul 02, 2020 8:02 am

Thanks Charles.
I noticed that your diatoms, the ones that you pick under the stereo microscope view, are clearly visible - the contrast against the background is very good.
Is it "ordinary" transmitted illumination ? the original lamp of the microscope ?
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Re: A tip and a question about a slide of clean diatoms without silt

#13 Post by MicroBob » Thu Jul 02, 2020 8:10 am

Hi Doron,
congratulations to this nice setup! For some samples it probably is much more worthwhile to pick good frustules out than to try to clean the sample to a high grade.
Do you use a sticky substance on the tip of your fibre? Is the fibre attatched to the objective and moves with the focussing action?

Bob

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Re: A tip and a question about a slide of clean diatoms without silt

#14 Post by Charles » Thu Jul 02, 2020 11:38 am

Hobbyst46 wrote:
Thu Jul 02, 2020 8:02 am
I noticed that your diatoms, the ones that you pick under the stereo microscope view, are clearly visible - the contrast against the background is very good.
Is it "ordinary" transmitted illumination ? the original lamp of the microscope ?
I use both transmitted light, original lamp from the base as well as two top lighting from each side provided by two IKEA LED lamps.

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Re: A tip and a question about a slide of clean diatoms without silt

#15 Post by Hobbyst46 » Thu Jul 02, 2020 1:20 pm

MicroBob wrote:
Thu Jul 02, 2020 8:10 am
Hi Doron,
congratulations to this nice setup! For some samples it probably is much more worthwhile to pick good frustules out than to try to clean the sample to a high grade.
Do you use a sticky substance on the tip of your fibre? Is the fibre attatched to the objective and moves with the focussing action?

Bob
Thanks, Bob
There is no sticky substance on the tip. Maybe I will try it in the future. At the moment it is all experimental (as Q of Q-branch would say). When the frustule in the strew slide is strongly attached to the slide surface, the fiber is unable to lift it - sumtimes it scrapes and destroys the poor frustule. Sometimes it operates like a chisel or lever - the tip pushes under the frustule, and being flexible and springy, the diatom will jump and land who knows where... :lol:
But often it works well as is. I will post some images of the transfered diatoms, valve view and maybe even girdle view...

Below is a photo of the setup from a the right side of the microscope. The mechanical finger is a 3mm OD stainless steel tube (2 in the photo), inserted into a 4mm OD tube (1 in the photo) that serves as sleeve. They slide coaxially like a telescope. The back end of the inner tube is stuck into a plastic tube (3) that serves as retainer as well as handle for rotation around the axis. The front end is a series of sections of shafts hypodermic needles, of decreasing diameters (4); the tip receives the glass fiber (not visible in the photo). Sleeve (1) is epoxy-glued (6) to another short horizontal section of the same tubing as 2. This section, in turn, serves as sleeve for two opposing needles (#15 gage; 5) that together form an axis for the finger. The needles are glued to a rack made of Lego pieces (7). The finger rocks up and down on its horizontal axis, by means of pressure of the tip of a Moore&Wright 0.01mm micrometer (8). Also shown are the slide holder, including jaws and fine X-movement mechanism (9), and the tilted roof of the LED-strip illuminator (10).
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diatom isolation setup.jpg
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Re: A tip and a question about a slide of clean diatoms without silt

#16 Post by MicroBob » Thu Jul 02, 2020 4:48 pm

Hi Doron,
thank you for the explanation! Building this ingenious setup must have taken quite some time. And there are probably few other designs that include a micrometer as well as Lego bricks! :lol:

For my attempts at diatom arranging I used a simpler setup but was not that happy with it. It made diatom arranging feasible for ordinary mortal but the productivity is low. Möller would have died from old age after the first 200 of his 20000 species type plates. :lol:
I have some harshly cleaned diatom materials that contain lots of broken bits and don't give nice strew slides but contain beautiful diatoms. For these I would like to try Möllers free hand method. Generally I have a steady hand but this will for sure be a challenge!

Bob

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Re: A tip and a question about a slide of clean diatoms without silt

#17 Post by Hobbyst46 » Thu Jul 02, 2020 6:01 pm

MicroBob wrote:
Thu Jul 02, 2020 4:48 pm
Hi Doron,
thank you for the explanation! Building this ingenious setup must have taken quite some time. And there are probably few other designs that include a micrometer as well as Lego bricks! :lol:

For my attempts at diatom arranging I used a simpler setup but was not that happy with it. It made diatom arranging feasible for ordinary mortal but the productivity is low. Möller would have died from old age after the first 200 of his 20000 species type plates. :lol:
I have some harshly cleaned diatom materials that contain lots of broken bits and don't give nice strew slides but contain beautiful diatoms. For these I would like to try Möllers free hand method. Generally I have a steady hand but this will for sure be a challenge!

Bob
Thanks. I had considered the Klaus Kemp setups, engineering-level manipulators, the setup you posted, the one shown by Charles, the MacLaughlin book, Steve Beats and others. I learnt that if the mechanical finger can lift a diatom, a sophisticated micromanipulator is not a must.
My hands are unstable, so I will be very happy to isolate clean diatoms, and get rid of the huge amount of silt and debris - not even dream of arranging them.
Actually, the challenge is twofold: many diatoms are so strongly attached to the slide that the finger cannot lift them; and, many of those that do cling to the finger, refuse to let go and settle in the new place. As mentioned in the past by other microscopists...

BTW: I had the micrometer from some old project stuff; but I believe that a very fine pitch long screw would also do the job.
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Re: A tip and a question about a slide of clean diatoms without silt

#18 Post by MicroBob » Thu Jul 02, 2020 8:02 pm

I think the importance of arranged slides has recedet: One can assemble the arrangement from individual photos and gets a result that can be shared via internet. But separating and mounting individual diatoms is still nearly as useful as ever.

I used skin fat on target surface and seal hair as I think Möller did. But the picking and placing still was difficult.

It also is a question of how clean exactly the material is. Recently I worked on harshly chemical cleaned material that has no inclination to stick somewhere so clean it is. For strew slides it was enough to shovel a bit of material on the cover slip, it willingly spread itself in the mounting process. My self cleaned material would tend to stick together more and when applied as a suspension would adhere to the donor slide more.

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Re: A tip and a question about a slide of clean diatoms without silt

#19 Post by Hobbyst46 » Tue Jul 14, 2020 1:29 pm

MicroBob wrote:
Thu Jul 02, 2020 8:02 pm
...Recently I worked on harshly chemical cleaned material that has no inclination to stick somewhere so clean it is. For strew slides it was enough to shovel a bit of material on the cover slip, it willingly spread itself in the mounting process. My self cleaned material would tend to stick together more and when applied as a suspension would adhere to the donor slide more.
My cleaned diatom slurries, regardless of the exact cleaning process (gently cleaned with various chemicals except strong acids and 30% H2O2), seem to strongly adhere to the donor slide. Only the centric large (~50um) Thalassiosira agrees to be picked up with the mechanical finger.
So I re-read stuff and found that Stuart R. Stidolph from NZ suggested a ground ("frosted") glass slide, made by honing an ordinary slide with carborundum powder, as donor. Instead, I simply placed the drops of raw diatom slurries onto the frosted surface of frosted slides ("Super-frost" or similar). Indeed, the diatoms appear to hold less to the slide and easier to pick-up with the thin glass fiber. The illumination is top-light, epi-oblique, anyway, with a horizontal LED strip.
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Re: A tip and a question about a slide of clean diatoms without silt

#20 Post by MicroBob » Tue Jul 14, 2020 2:33 pm

The use of frosted slides is a good idea I have never heard of before. Once I have sent diatom material I had cleaned to a degree suitable for the light microscope to someone who photographs diatoms with a SEM. For him the material was not usable as the SEM showed a lot of remaining dirt. Some diatoms also still stuck together a lot.
Attachments
30_Klebtest UV-I  Bob 304 klein.jpg
30_Klebtest UV-I Bob 304 klein.jpg (93.25 KiB) Viewed 3044 times
30_Klebtest UV-I  Bob 101 klein.jpg
30_Klebtest UV-I Bob 101 klein.jpg (122.31 KiB) Viewed 3044 times

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Re: A tip and a question about a slide of clean diatoms without silt

#21 Post by MichaelG. » Tue Jul 14, 2020 4:33 pm

Not sure how I missed this this, Doron ... but I must add my congratulations

Incidentally ... The slides with multiple cavities are similar in concept to these:

http://www.hendley-essex.com/microscope ... 4586826589

[ but I note that yours have individual rings]

MichaelG.
Too many 'projects'

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Re: A tip and a question about a slide of clean diatoms without silt

#22 Post by Hobbyst46 » Tue Jul 14, 2020 7:54 pm

MichaelG. wrote:
Tue Jul 14, 2020 4:33 pm
Incidentally ... The slides with multiple cavities are similar in concept to these:
http://www.hendley-essex.com/microscope ... 4586826589
[ but I note that yours have individual rings]
Thanks, Michael
The slides I used were inherited from a very old stock, expiration date years before 1990... the Hendley ones are modern and probably better - but I have switched to frosted slides anyway.

I pick the diatoms from the frosted surface and place them (actually, drop them) on the transparent surface of the same slide.

Here are some of my isolated diatoms on the "stock" slide: in air, no mountant or coverslip, just lying on the glass, and imaged with the 40X0.75 Neofluar Ph2 with the Ph3 setting of the condenser. Not the "proper" setting for the objective, but convey the mood. Pictures are resized and cropped.

I believe that the circular arc thing is a girdle band from a disrupted Thalassiosira. Lying on its side. When I pick them up with the glass fiber, they look like that.
One Thalassiosira is lying on its side, so to speak, girdle view; maybe I when I mount them in Pleurax, some will stay in girdle view.

There are many dust particles around - they accumulated during the isolation work, and provide a certain astronomical sight, to my eyes...
The elongated diatom is Pleurosigma or Gyrosigma - must work on the identification. "Thalassiosira" is also a guess. Might be Biddulphia.
Attachments
diatom dry, bare slide, 40X0.75.JPG
diatom dry, bare slide, 40X0.75.JPG (37.54 KiB) Viewed 3014 times
Thalassiosira and girdle (tentative) dry, bare slide, 40X0.75 stack of 2.jpg
Thalassiosira and girdle (tentative) dry, bare slide, 40X0.75 stack of 2.jpg (11.99 KiB) Viewed 3014 times
Thalassiosira (tentative) girdle view, dry, bare slide, 40X0.75 stack of 3.jpg
Thalassiosira (tentative) girdle view, dry, bare slide, 40X0.75 stack of 3.jpg (9.26 KiB) Viewed 3014 times
Last edited by Hobbyst46 on Tue Jul 14, 2020 9:27 pm, edited 1 time in total.
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Re: A tip and a question about a slide of clean diatoms without silt

#23 Post by Hobbyst46 » Tue Jul 14, 2020 8:01 pm

MicroBob wrote:
Tue Jul 14, 2020 2:33 pm
The use of frosted slides is a good idea I have never heard of before. Once I have sent diatom material I had cleaned to a degree suitable for the light microscope to someone who photographs diatoms with a SEM. For him the material was not usable as the SEM showed a lot of remaining dirt. Some diatoms also still stuck together a lot.
SEM has its virtues but lacks the colorful light effects !
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Re: A tip and a question about a slide of clean diatoms without silt

#24 Post by Hobbyst46 » Thu Jul 30, 2020 9:41 pm

Update:

Been busy training to "herd" (as someone coined it) diatoms to isolate them from debris- and silt-ridden strew slide (coined "donor slide").

Some findings:
1) The glass fiber from glass-wool (see above) is a very good tool on the mechanical finger. Mine is perhaps too long - 3-4 mm. It is the long rod in the images below, running left-low to right-high.
2) The stereoscope - 10X-40X and a 1.5X Barlow lens, yielding a 60X maximum magnification - is fine for the larger frustules. 20X eyepieces were ordered to try and catch the smaller frustules.
3) A strew slide on a frosted glass donor slide is batter than on a plain glass slide.
4) A strew slide on an optically black self-adhesive tape, on top of a flat surface (just any slide), is a tough competitor for the frosted, and often the winner.
5) The top-side-light from a LEd strip illuminator (see above) works well with the above opaque surfaces.
6) A black background under the microscope glass stage improves the view. See images below. The fiber has been pressed onto the surface to bring it in focus with the frustules. 60X magnification.
7) As expected, high relative humidity in the air weakens the attraction of frustules to the "fishing-pole", namely the glass fiber. Rubbing the end of the glass fiber on a piece of nylon pantyhose helps a little.
8) A fine (pun) accessory: short acupuncture needles. They have a "handle" and are much thinner than sewing pins/needles.
Attachments
6) Strew slide on the frosted surface of a frosted slide.jpg
6) Strew slide on the frosted surface of a frosted slide.jpg (98.94 KiB) Viewed 2387 times
7) Strew slide on an optical black tape on top of a slide.jpg
7) Strew slide on an optical black tape on top of a slide.jpg (58.5 KiB) Viewed 2387 times
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Re: Diatom selection experience

#25 Post by Hobbyst46 » Thu Aug 13, 2020 1:30 pm

Hello all,

I modified the name of this post thread, to refer to various aspects of Diatom selection and isolation. Picking them from a strew or storage slide onto an "arranged" (target) slide. Without any artistic ambition, just isolation from the silt and debris (that filtration/sedimentation did not remove).

Updates:
1. Again, the use of miniature (1/2") paper binder clips to hold the slide on top and a few mm above the hot surface (as suggested by rnabholz) is successful.

2. There are several (or many) recipes of pre-mounting adhesives (=fixatives) to stabilize the selected diatoms in their place on the target coverslip. Judging them on the basis of the physical properties of their components is difficult, at least when the mountant is Pleurax. The adhesive I am now trying is gelatine in water/IPA/acetic acid. Although the local weather is highly humid, diatoms do not stick to the adhesive on the target coverslip, regardless of adhesive layer thickness, unless moistened. I moisten not by breathing, which is ineffective and blows away the diatom, but by storage of the coverslip near to a sheet of wet filter paper in a closed box for 12 hours.

3. Because my diatoms were gently cleaned, many of them are not completely separated into valves and girdles - so they break during mounting. It seems that they break by the mere pressure of the slide on the coverslip right after the Pleurax drop is placed. I mount by placing the slide on top of the coverslip, to keep the diatoms near the coveslip. Apparently, this configuration crashes them. Definitely not the boiling or expansion of the resin.

4. So, trying spacers. Placed coverslip fragments on the adhesive layer imediatley after the diatom selection. Problems - the required resin layer was very thick; the fragments did not hold well and some were lost in the process. Will try fragments of 90um thick aluminum foil instead (as suggested by Charles) but these will be placed under the adhesive.

5. Another possibility, instead of as in 4) above, is resting the slide on pillars that widen the gap between the slide and coverslip - so after they are united with a Pleurax drop in between, the resin layer will be thick (without the need for spacers) and will safely hold the diatoms without crashing them.

6. 18mm diameter coverslip are conveniently stored in 12-well Coastar plates, in which 4-pole Lego pieces are placed as stand-offs.

Work is in progress.
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Hobbyst46
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Re: Diatom selection experience

#26 Post by Hobbyst46 » Thu Aug 20, 2020 8:45 pm

Update.
Still looking for an appropriate cement for diatoms to be mounted in Pleurax.
Literature data about Gum Tragacanth are somewhat variable; in the meantime I try gelatine. Heating to >90C seems to improve the retention of the diatoms on the gelatine, not for a true arrangement yet.

Here are some images of my favorite centric isolated diatom in Pleurax. They are slightly modified in post processing, resized and cropped (not by the same factor). valve view. Could not achieve girdle view because the boiling Pleurax apparently pushes them flat onto the surface...
Attachments
Neofluar 40X0.75 Ph2, phase contrast.JPG
Neofluar 40X0.75 Ph2, phase contrast.JPG (84.33 KiB) Viewed 1936 times
Planapo 63X1.4 Ph3, phase contrast.JPG
Planapo 63X1.4 Ph3, phase contrast.JPG (43.26 KiB) Viewed 1936 times
Planapo 63X1.4 Ph3, oblique enhanced, stack.jpg
Planapo 63X1.4 Ph3, oblique enhanced, stack.jpg (49.5 KiB) Viewed 1936 times
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Hobbyst46
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Re: Diatom selection experience

#27 Post by Hobbyst46 » Wed Sep 09, 2020 3:46 pm

Update:

So after further experiments, I arrived at a practical (probably not ideal) method of isolation of cleaned diatom frustules from silt/debris/powder and mounting them in Pleurax.
It works for isolation - not for arrangements.

1. Fish the cleaned diatoms out from the dry raw mixture (prepared by letting a drop of suspension of diatoms dry out on a black surface on top of a slide) with a glass rod, diameter 6-10um, attached to a mechanical finger that moves up and down (like a seesaw).

2. Place them on either a cement-coated coverslip (a) or into drops of water on coverslip (b), as follows :

(a) Lay a small drop of the liquid gelatine cement (prepared as above) on the coverslip and let spread to the rims and dry out on the shelf (10-12h).
Then place diatoms one by one. When isolation is complete, heat the coverslip in an oven at 110C, for one hour.


(b) Lay a tini-tiny drop of distilled water from a syringe (disposable 1-2ml syringe, very thin hypodermic needle, say #27) onto the center of the clean dry coverslip. The drop remains
in the form of a tiny hemisphere. Place diatoms one by one on top of the drop. Some of them sink, others flow and tend to attach to each other. With the rod, push and nudge
to separate. When isolation is complete, let dry out.


3. Mount in Pleurax as usual. I do it over an alcohol burner. Three slides are placed, inverted and supported on paper clips, on top of an aluminum plate (dimensions ~ 150 x 70 x 10 mm) on a tripod over a burner. Heat gradually to 180C, then keep at about the same temperature for 15 min. Heating to 190-200 or higher causes intensive boiling and detaches the coverslip from the slide, so not recommended...

Note 1: method b above (water drops) is less elegant than method a (cement method), yet it does keep the diatoms on the slide, not far away from their initial positions. I chose it after several futile attempts to improve the gelatine cement method. My thought was: since strew slides are prepared by letting a drop of diatom suspension dry out on the glass, then mounting, it appears that for some reason, wet diatoms attach better (to glass) than dry diatoms. Perhaps, this holds for gently cleaned frustules, that still contain trace amounts of protein or other organic residue that may function as adhesive. But it worked for me with chemically cleaned fossil diatoms as well.
So far, method b works better than method a.
One more advantage of the water drops method is that dusty frustules are being washed with the water prior to sinking onto the glass surface.

Note 2: Gelatine is chemically similar to polyamide, and Diatoms.nl state that polyamide cement is compatible with Pleurax. This is logical, since polyamide, like protein, does not dissolve in alcohol. There are different gelatine-based recipes, which of them is best for Pleurax I do not know.

Note 3: still do not know whether gum-tragacanth cement is compatible with Pleurax. Hanna (inventor of Pleurax) advocated against it, though it was long ago.

Note 4: The above experiments were performed in very warm and humid days, which made both the gelatine cement and Pleurax quite free-running. Slight modifications are expected in colder and dryer weather.

Hope to show some photos of isolated diatoms in the future.
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MicroBob
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Re: Diatom selection experience

#28 Post by MicroBob » Wed Sep 09, 2020 7:26 pm

Hi Doron,
nice to hear of you progress!
According to your note 2: Michel Haak suggests polyacrylamide, not polyamide. I'm no chemist and don't know how far this is apart. http://www.diatoms.eu/de/node/77
The high curing temperature of Pleurax really narrows down the number of possible candidates for adhesives.

Bob

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75RR
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Re: Diatom selection experience

#29 Post by 75RR » Wed Sep 09, 2020 7:43 pm

Hobbyst46 wrote:
Thu Aug 20, 2020 8:45 pm
Here are some images of my favorite centric isolated diatom in Pleurax.
Very nice images and cool diatom!
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Hobbyst46
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Re: Diatom selection experience

#30 Post by Hobbyst46 » Wed Sep 09, 2020 7:48 pm

MicroBob wrote:
Wed Sep 09, 2020 7:26 pm
Hi Doron,
nice to hear of you progress!
According to your note 2: Michel Haak suggests polyacrylamide, not polyamide. I'm no chemist and don't know how far this is apart. http://www.diatoms.eu/de/node/77
The high curing temperature of Pleurax really narrows down the number of possible candidates for adhesives.

Bob
Thanks Bob, I stand corrected ! it is far apart... yet both substances are soluble in water and not in alcohol (or at least, much less in alcohol). The amide chemical groups makes them so.
Zeiss Standard GFL+Canon EOS-M10, Olympus VMZ stereo

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