Doing Diatoms - part 2

Here you can discuss sample and specimen preparation issues.
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Sure Squintsalot
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Re: Doing Diatoms

#1 Post by Sure Squintsalot » Tue Mar 14, 2023 9:35 pm

In the 9 months since I've started "Doing Diatoms" I've developed a sophisticated technique for collecting, preserving, and observing diatoms based entirely on things I misread, misinterpreted, and entirely misunderstood. This has cost me, but on occasion I get lucky. Like the centric frustule posted above, collected in a tea basket dragged through seawater from a panga.

I think these things are pretty cool, here photographed under (HA)DIC illumination. No way I'm getting sub-micron resolution with my set-up:
Screenshot 2023-03-14 152242.jpg
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Someone else thought these things were cool enough to stydy in detail:
Screenshot 2023-03-14 152722.jpg
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These guys report that "C. centralis valves trap light and modify its spectrum to match the optical absorption of the specific chlorophyll a and c photoreceptors...."

Read it here: Wavelength and orientation dependent capture of light by diatom frustule nanostructures https://www.nature.com/articles/srep17403

Their prep process is pretty interesting, though:

Diatoms were filtered out of the culture medium, washed 3 times in Milli-Q water, centrifuged at 4500 rpm for 10 min between each washing and dried overnight at 60 °C. Dried diatom valves (2 mg) were placed in hydrogen peroxide (H2O2, 10 mL) solution (30%) and stirred at 90 °C for 24 h. After addition of HCl solution (37%, 1 mL) the sample was centrifuged at 4500 rpm for 10 min. Finally, the cleaned frustules were rinsed with Milli-Q water, centrifuged (4500 rpm, 10 min) 3 times and stored in ethanol (96%).

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Re: Doing Diatoms

#2 Post by Sure Squintsalot » Wed Mar 15, 2023 12:24 am

That's the thing about plankton collection, fixing, and preservation. There are many different ways to skin this cat and sometimes they contradict one another. In my case, my entire process had to be as air-travel friendly as possible: no toxic materials, nothing flammable, and nothing too weird. Ideally it would have been built from things found in any trash bin near a beach, and I came pretty close too, once using a torn fabric shopping bag and some scrap fishing line found in a parking lot, which got me a pretty awesome, impromptu plankton sample.

My ocean harvest travel kit, however, looks like this:
Screenshot 2023-03-14 171753.jpg
Screenshot 2023-03-14 171753.jpg (83.31 KiB) Viewed 38896 times
The baskets are a gold 150 micron coffee screen and a 450 micron stainless tea basket that nests neatly within. I "found" these in my cupboard but you can buy these at any home goods store (like I ended up doing after my wife found hers in my plankton kit). Some string, fishing swivels, and a take-up reel will round out the collection eqipment.
The 150 micron screen and 450 micron screen compared. The larger screen does a pretty good job of filtering out larger zooplankton like copepods and larvae. This is actually kind of useful when preparing a slide.
The 150 micron screen and 450 micron screen compared. The larger screen does a pretty good job of filtering out larger zooplankton like copepods and larvae. This is actually kind of useful when preparing a slide.
Screenshot 2023-03-14 180510.jpg (202.48 KiB) Viewed 38896 times
After a 5-15 minute drag, the sampler is brought on deck and individually rinsed out in the half-bottle using %90 IPA (available anywhere), the top half of a 2 liter plastic bottle, and a syringe for rinsing off the goods. I get a coarse plankton sample as well as a fine sample, per the two screens. The plankton/IPA soup is then funneled into a couple of sample vials for processing on shore. The syringe may also be used to draw off more clean IPA to further concentrate the sample bottle with plankton:
Screenshot 2023-03-14 174108.jpg
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This technique is not without its drawbacks:
  • Plankton nets are notorious for either mangling or destroying softer ciliates and any delicate life form not made out of glass or stone. The best way to collect plankton, apparently, is also the most hands-off: Collecting a large volume of water and letting everything settle to the bottom over a few hours to be carefully drawn off. That's my next experiment; using gravity and a 3 liter water bladder suspended from a tree with a beer in hand, waiting for my plankton to "settle down".
  • Left unattended, salt is precipitated out of solution after a time and may contaminate or destroy delicate internal structures. The IPA absorbs the water within the sample leaving the dissolved salts with no place to go and so, they crystallize, usually within plankton walls or on appendages. Don't leave your samples on the shelf for a few months without a few rinses in distilled water, THEN preservation in IPA.
  • I'm not interested in using nasty, unconventional chemicals for any part of this process, though I'll draw the line at Lugol's Iodine or Formalin as I've already ruined a few samples to bacteria eating everything in overlooked sample vials that hadn't been kept in IPA. For example, bleach is a pain in the ass to fully rinse from a sample and if you're impatient, you'll be tempted into using the centrifuge to speed the process along, further destroying what you've worked so hard to obtain. Mercury-based preservation compounds are also a major PITA to dispose of properly.
  • I do not know what the lifetime is of plankton prepped, fixed, and preserved in IPA; the nearest I've come to seeing anything in the literature is the use of 90% ethanol following fixing in formalin. I'm figuring on 90% IPA being the poor man's 90% ethanol. We'll see.
Did I hear someone ask about slide prep?

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Re: Doing Diatoms

#3 Post by imkap » Wed Mar 15, 2023 8:39 am

Thanks for the detailed info :)

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Re: Doing Diatoms

#4 Post by Hobbyst46 » Wed Mar 15, 2023 1:06 pm

Citation: "Diatoms were filtered out of the culture medium, washed 3 times in Milli-Q water, centrifuged at 4500 rpm....."
1. That protocol is fairly standard. Actually, it is quite minimal, for diatom cleaning. The reason is that their diatom stock is a culture, in which typically there are very few contaminants. By comparison, diatoms directly collected from nature (river, lake, beach) are often a minority within a nasty mixture of silt, micro gravel, organic debris etc.

2. Isopropanol is OK as preservent, if you can stand the smell. It is less volatile than ethanol, so keeps better on the long run. Storage in dark-colored flasks or vials is also advantegeous.

3. IMO, for diatom collection, centrifuges are optional, not a must. Diatoms settle from water within a few minutes even in a simple static test tube. Moreover, when the sample contains silt, centrifugation will likely sediment the diatoms and silt together, that is, contaminate the sample even more.

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Re: Doing Diatoms

#5 Post by Sure Squintsalot » Thu Mar 16, 2023 5:12 am

Hobbyst46 wrote:
Wed Mar 15, 2023 1:06 pm
Citation: "Diatoms were filtered out of the culture medium, washed 3 times in Milli-Q water, centrifuged at 4500 rpm....."
1. That protocol is fairly standard. Actually, it is quite minimal, for diatom cleaning. The reason is that their diatom stock is a culture, in which typically there are very few contaminants. By comparison, diatoms directly collected from nature (river, lake, beach) are often a minority within a nasty mixture of silt, micro gravel, organic debris etc.

2. Isopropanol is OK as preservent, if you can stand the smell. It is less volatile than ethanol, so keeps better on the long run. Storage in dark-colored flasks or vials is also advantegeous.

3. IMO, for diatom collection, centrifuges are optional, not a must. Diatoms settle from water within a few minutes even in a simple static test tube. Moreover, when the sample contains silt, centrifugation will likely sediment the diatoms and silt together, that is, contaminate the sample even more.
I agree, so far as prepping for SEM goes. In addition to slowing down the pumping-down of the sample chamber, at 20,000+ volts, electron beams easily vaporize any organic contaminants which are then re-deposited within the beam guide, screwing up aiming, focus, and stability of the beam; I can speak from direct experience. The poor shmo manning the instrument then has to take the entire beam guide apart and spend 4 hours cleaning and reassembling the thing. I thought it was interesting because having only ever looked at inorganic, non-biologic materials in an SEM, I'd never realized how big a PITA it was to remove volatiles from a sample. You're right I shouldn't be surprised that this fairly standard.

But I can only imagine such cleaning procedures working with compact, centric diatoms exclusively. There is no way pennate diatoms could survive these procedures intact. In fact I'm amazed by how few images and studies are done on pennate diatoms. Is this an example of diatom investigations self-selecting to centric varieties because only they survive a stiff cleaning? Even on this site, how many plankton images have we seen of these?:
Pennate phytoplankton collected from Puerto Galera using collecting tools and process detailed above.
Pennate phytoplankton collected from Puerto Galera using collecting tools and process detailed above.
Screenshot 2023-03-15 225247.jpg (65.19 KiB) Viewed 38835 times
I just wish ethanol weren't so expensive and hard to come by: $80/gal + hazmat fee + shipping! And yet we use cubic miles of that stuff every year in our gas tanks.

I've pored through dozens of plankton collection procedures from the US EPA, National Marine Authorities, The UN, and tons of academic procedures and about 25% will mandate the use of a centrifuge; likely because these are ocean-going collection procedures free of dealing with silt and other fine suspended crap. And they they probably could care less about getting the perfect photo of the prettiest plankton. I agree that there is no value in centrifuging a dirty plankton solution. I've had pretty good luck simply lightly stirring a silty solution and drawing off the cloudy rinse-water repeatedly until it was clear- yielding diatoms from filthy diatomaceous earth.

Here are a few more (Ceratium (?)) pennates, and a radiolara (?) that I don't often see here or in academic papers aside from the rare characterization:
Ceratium
Ceratium
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Ceratium colony! Never seen one of these before!
Ceratium colony! Never seen one of these before!
Screenshot 2023-03-15 233232.jpg (107.99 KiB) Viewed 38827 times
Radiolara (?)
Radiolara (?)
Screenshot 2023-03-15 232241.jpg (117.54 KiB) Viewed 38827 times

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Re: Doing Diatoms

#6 Post by Phill Brown » Thu Mar 16, 2023 10:48 pm

Thanks for taking the time to share your efforts.
Definitely some worthy examples to make permanent mounts.
I'd be interested to see DF Vs DIC, mostly because I'm unlikely to ever get DIC, it's just too spendy to justify whereas DF is within reach of the most modest budget.
Keep up the good work.

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Re: Doing Diatoms

#7 Post by Sure Squintsalot » Fri Mar 17, 2023 12:11 am

I'm happy to show these off, Phill; glad you and others here can appreciate them!

As for DF, I'd love to do that, but I don't know how. I bought the eBay 3D printed insert set for my condenser, but I can't seem to make anything work with my assortment of objectives (mainly DIC and metallurgical). If I need to have DF-specific objectives, then, I guess, I'm screwed.

As I mentioned in an earlier post, I'm using "HA-DIC".... a highly technical variation on standard DIC in which I half-assed the condenser prism with a plexiglass Sanderson prism. At less than $30, and half the price of that 3D printed insert set, it seems to work well enough for my purposes!
Screenshot 2023-03-16 180323.jpg
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I do, however, use a full DIC-capable turret. My HA-DIC solution, as useful as it is, still can't compare to a proper, fully dedicated DIC set-up.

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Re: Doing Diatoms

#8 Post by Phill Brown » Fri Mar 17, 2023 8:35 am

It's for another thread but I'm confident there would be a flood of options for how to achieve DF with what you already have.

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Re: Doing Diatoms

#9 Post by JWW » Fri Mar 17, 2023 9:12 pm

Personally I'd love to see a new thread instead of 17 pages to sift through.

-JW:

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Re: Doing Diatoms

#10 Post by Hobbyst46 » Sat Mar 18, 2023 11:03 am

JWW wrote:
Fri Mar 17, 2023 9:12 pm
Personally I'd love to see a new thread instead of 17 pages to sift through.

-JW:
Likewise. Especially in view of these very nice photos of the fragile diatoms. And the beautifully machined Sanderson prism holder.
Would very much appreciate more technical details. I see a Nikon Optiphot with trans-illumination, but:
1. Thickness of the prism ?
2. Objective ?
3. Any other optical component ?
Thanks in advance.

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Re: Doing Diatoms

#11 Post by Sure Squintsalot » Tue Mar 28, 2023 10:30 pm

Oliver was kind enough to make some of your dreams come true and create a new thread, Doing Diatoms- Part 2.

Onward!
Phill Brown wrote:
Thu Mar 16, 2023 10:48 pm
I'd be interested to see DF Vs DIC, mostly because I'm unlikely to ever get DIC, it's just too spendy to justify whereas DF is within reach of the most modest budget.
Thanks for the prod, Phill. As a result, I've been going a little nuts with the dark field imaging. None of my stops will work with my fancy pants 20x objective so I had to buy a crappier E-Plan! My condenser isn't the best for this sort of thing, either. Nevertheless, here are a few DF images:
DIC at left, DF at right
DIC at left, DF at right
Screenshot 2023-03-28 160902.jpg (100.9 KiB) Viewed 38436 times
Space Invader!
Space Invader!
Screenshot 2023-03-28 161423.jpg (76.04 KiB) Viewed 38436 times
Blue Fluorescence? or refraction into blue spectrum?
Blue Fluorescence? or refraction into blue spectrum?
Screenshot 2023-03-28 162138.jpg (41.43 KiB) Viewed 38436 times
Screenshot 2023-03-28 162511.jpg
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The oddities never cease!
The oddities never cease!
Screenshot 2023-03-28 162651.jpg (99.93 KiB) Viewed 38436 times
I've got to admit, the DF is a lot of fun. I feel like I've gotten a whole new set of slides to look through!

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Re: Doing Diatoms - part 2

#12 Post by apochronaut » Wed Mar 29, 2023 1:40 am

Your careful preparation which preserves such exquisitely delicate structures, certainly deserves examination in as many illumination techniques as is practical. DF, for objects that are essentially made of glass is pretty ripe for exploitation. These are quite remarkable. It is amazing that some of them survive the rigours of their ennvironment but many I guess are built to just break and then continue dividing.
For about 75 years, 20X apochromats were .65 N.A., sometimes a little less. I have numerous examples of them made right up to the mid. 80's and they all yield fantastic images. Was .65 the N.A. because manufacturers could go no higher? No, it was because those objectives could be used directly for D.F. with a dry D.F. condenser. Once you go to .75, you can't, you need an oil D.F. condenser. I wrote the following for the recent thread " Illumination techniques for beginners". It clarifies as succinctly as I could, why these higher N.A. .75 objectives don't work for dry D.F.
Even N.A. .65 is very tricky with patch stops due to centering difficulties and achieving success is almost impossible. Definitely a dedicated condenser is a major plus.
I'm not sure what Nikon has to offer for the Optiphot 1(?) in terms of condensers that are economical for DF but there are other condensers out there of very good specification at ridiculously bargain prices. AO oil DF condensers were the standard in hospitals and doctor's offices over about 50 years for the detection of syphilis. They are circular mirror condensers or cardioid mirror condensers, with a sealed silver mirror section, a threaded nose extendable by about 10mm and an attached AO dovetail that can be removed by loosening 2 screws in about 10 sec. This reveals a circulsr neck that can receive any other dovetail, printed or machined to fit your microscope if the condenser is tall enough using that dovetail location. If not, a custom dovetail can be mounted on the bottom with three existing threaded holes. It is the easiest high quality DF condenser that can adapt to almost any other microscope avaiable . There still seem to be hundreds around, I have only ever seen 1 failed one, and if you pay more than 100.00 for one, you payed too much. It will work at N.A. .80 or lower and the later versions made from about 1948 to 1985 ( the 214F) cover a field of about 900 microns but I have not actually meaured that. I have used a 25X .65 plsnapo with one at a 20mm f.n. but with a 20X , it doesn't quite fill a 20mm field. Still useable, though.They are mostly plated brass as far as I can tell, with some aluminum. This is by far the best under the radar oil DF condenser for those not wanting to pay the huge sums being asked for big 4 versions.

I wrote the following for the recent thread " Illumination techniques for beginners". It clarifies as succinctly as I could, why these higher N.A. .75 objectives don't work for dry D.F but it could just as easily refer to .65 objectives which still suffer from some of the technical ills. One overlooked problem with DF is that it more than any other technique seems to amplify ca. Condensers such as an abbe with a patch stop will contribute to ca in the DF image. The better the alignment, the less the ca but there is a big difference between an abbe with a patch stop working at .65 N.A. and an oil DF condenser working at .65 N.A. Many oil DF condensers are completely ca free, some are prone to very little and virtually none when apochromats are used.

The principle of DF is very simple. It is a modified form of circular oblique lighting ( COL) that results in total internal reflection in the slide itself, (TIR). There are 2 conditions for successful DF illumination .
1) Illumination by a source that is at least .20 N.A. above the N.A. of the objective in order to eliminate flare and scatter.
2) Completely uniform symmetrical illumination, therefore precise centering. DF condensers do most of that for you, whereas with patch stops at higher N.A's you might as well take up plate balancing and go on ( fill in your country of choice)'s Got Talent.

DF condensers have a minimum and maximum N.A., which creates a ring of illumination, like a thin doughnut. It's not really a cone, it is more like the walls of a funnel, projecting it's illuminated cross section on the slide. If the objective can't see it, then theoretically you can have DF.
This N.A. range of that doughnut is often stamped on the condenser but if not, a slide thickness specification may be, which is a related specification. The maximum N.A. is usually about .20 higher than the minimum, so since any dry DF condenser cannot have a maximum N.A. higher than 1, therefore it's minimum will be approximately .80 to .85 , maybe .90 is possible but that would be a really precise tightly engineered condenser and would require high illumination. Never seen one like that. This is where it gets a bit confusing because technically if the objective cannot see the projected ring and therefore the minimum condenser N.A. is higher than the objective, there should be DF. That gets stated a lot in print and by people who have never actually done DF, just read about it. In practice, when the objective N.A. is too close to the lower condenser N.A., a whole group of elements from glass impurities to sample refraction and extraneous reflection cause illumination scatter, turning DF into grayfield, which is kind of like dawn. In order to get really good DF with a good dark background you need to have the condenser's minimum N.A. .20 above the objective at least to be safe, which means an oil DF condenser, for an objective of .75.
This is evidenced by the Reichert Univar microscope and maybe the Polyvar too. They both had the advantage of a really good DF condenser that had a quite wide illumination funnel or ring between 1.40 and 1.20 N.A., so very high. Since all of the objectives for those microscopes were entirely universal and used for all contrast methods, they all had to be D.F. ready. They made every objective with an N.A. above .75 with a built in iris diaphragm. 3- 40X, 2-63X and 4- 100X. So, .75 seems to be an accepted cuttoff point, since they made a 25X .65 planapo which works stellar with their oil condenser and did not come fitted with an iris. For the 10 or so objectives they made with N.A.s below .50, they also fitted s dry DF condenser into the substage.

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Re: Doing Diatoms - part 2

#13 Post by Phill Brown » Wed Mar 29, 2023 9:32 am

Some beautiful images of great subjects.
I'll dig out my Labophot 2 and try some Nikon options.
Will add some images of what is used.
Fairly critical with DF is to get the condenser top lens as close to, if not touching the slide.
That dictates where the patch is best placed, not always where a filter holder is.
Always happy to be corrected.

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Re: Doing Diatoms - part 2

#14 Post by apochronaut » Wed Mar 29, 2023 11:01 am

Because DF relies on total internal reflection in the slide, it is the slide thickness that determines all other vertical spacial relationships in the system.

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Re: Doing Diatoms - part 2

#15 Post by Phill Brown » Wed Mar 29, 2023 11:45 am

apochronaut wrote:
Wed Mar 29, 2023 11:01 am
Because DF relies on total internal reflection in the slide, it is the slide thickness that determines all other vertical spacial relationships in the system.
A Gap between the condenser and the slide? Good luck with that if it works.

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Re: Doing Diatoms - part 2

#16 Post by apochronaut » Wed Mar 29, 2023 12:45 pm

You misinterpreted. I never mentioned the word gap. I said "the slide thickness determines all other vertical spatial relationships in the system" . The layer of oil or in the case of a dry condenser air is necessarily variable, depending on the slide thickness. It's simple geometry based on the angle of incidence that the minimal N.A. of the condenser achieves at the slide and the thickness of the slide.
The fact that you find that the condenser needs to " be as close to if not touching the slide" is only due to the conditions that you have : the condenser, type of oil and slide thickness. That is not always necessarily the case, though. There is no, one size fits all. The slide thickness rules all the rest and often the condenser focuses best when backed away from the slide some.
I have used, let me count them, 10 oil DF condensers on 12 or more different microscopes, no 11 condensers and a couple or 3 dry ones plus various slide thicknesses and oils, and the condenser focus is not a constant.
Last edited by apochronaut on Wed Mar 29, 2023 3:24 pm, edited 1 time in total.

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Re: Doing Diatoms - part 2

#17 Post by Phill Brown » Wed Mar 29, 2023 2:39 pm

IMG-20230329-WA0004.jpg
IMG-20230329-WA0004.jpg (58.83 KiB) Viewed 38327 times
F.O.V is 0.32mm with 40x .65 .
Taken with Lumix G3 micro 4/3 NDPL2X with micro 4/3 adapter.
Slide thickness is 1.3mm,if that matters.
Euromex iscope which is my least favourite but current model.
I don't do stacking, maybe this year I'll get there.
It's only to show contrast of DF with .65.
I have a 100x .85 LWD/Dry which works perfectly well without oiling the condenser.
I don't have any AO anything so I'll stay away from that.
Image has no photo shop or corrections. Just as is.
For DF I'll leave focusing the condenser to anyone who wants to try that.

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Re: Doing Diatoms - part 2

#18 Post by Sure Squintsalot » Wed Mar 29, 2023 4:12 pm

Wow! That's pretty damn good for being right out of the box! What kind of diatom is that, anyway?

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Re: Doing Diatoms - part 2

#19 Post by apochronaut » Wed Mar 29, 2023 5:16 pm

Very typical image quality for a .65 achromat with a dry DF condenser, which I assume is a DF condenser, not a modified BF condenser? If it were an abbe condenser with a patch stop, that would be exceptionally good, just to get the thing centered.
Your 1.3mm slide just illustrates the slide thickness point. Most DF condensers require the slide to be thicker than what is currently the standard in order to provide the N.A.along with field coverage. Dry condensers less picky than oil condensers. As the slide thickness changes, so by necessity does the condenser distance from the bottom of the slide, in order to maintain the condenser f.o.v. The standard slide thickness today is around 1.0 mm. DF condensers either have the slide specification stamped on them or it is provided in the catalogue.

I would be much more interested in an image using your 100X .85 objective with that condenser. .85 is typically what an oil DF condenser can support.

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Re: Doing Diatoms - part 2

#20 Post by Phill Brown » Wed Mar 29, 2023 6:10 pm

IMG-20230329-WA0005.jpg
IMG-20230329-WA0005.jpg (57.59 KiB) Viewed 38279 times
F.O.V 0.12 ish. Same diatom.
I would say it's not it's best performance, I'd also say it's not very often great,it would be the first I'd let go if this was the best it can do.
WD 3.2mm. metallurgical. 100x .85 Pl.
Lives on my inverted scope.
Again it's contrast in DF, sometimes it's ok.
Edit. Thinking about the 100x living on my inverted and not being used for some time.
Took it out to check, it's very dusty.
Epic fail #2
After not labelling the slide.
Last edited by Phill Brown on Wed Mar 29, 2023 6:31 pm, edited 1 time in total.

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Re: Doing Diatoms - part 2

#21 Post by zzffnn » Wed Mar 29, 2023 6:23 pm

Phil Brown, NA 0.85 with a dry (not immersed in oil) condenser is theoretically impossible and against laws of physics. Are you sure you did not make a typographical error there? Or was the numerical aperture of your “NA 0.85” restricted without you knowing it?

Apochronaut was being polite and subtle there, by not calling “impossible” like I did.

If you don’t believe us, I suggest you open a new thread with the thread title “is it possible to achieve NA 0.85 darkfield with a dry condenser”, then just ask members to answer yes or no (or ask them to provide technical/ scientific references).

I have done quite some darkfield imaging, with DIY patches + oiled Abbe condenser or oiled dedicated darkfield condensers. My avatar image here (the phacus image) was shot right around NA 0.7-0.75 or so.

With a dry (not immersed, so NA 0.7-0.9) darkfield condenser, even NA 0.65 is very very hard to achieve. With immersion, it is easy, even with DIY patches and Abbe.

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Re: Doing Diatoms - part 2

#22 Post by Phill Brown » Wed Mar 29, 2023 6:45 pm

I stand corrected.
IMG-20230329-WA0006.jpg
IMG-20230329-WA0006.jpg (90.68 KiB) Viewed 38253 times
.80.
As for .65 dry being difficult I can only say I took it for granted it isn't.
Condenser is dry.
It's a thread including diatoms in DF.
If anything it's a good example of why not to add a 100x dry for rare occasions.
Lots of things are impossible, getting DF after focusing the condenser is likely one of them.
I don't do it but I'm happy for anyone to keep trying.

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Re: Doing Diatoms - part 2

#23 Post by zzffnn » Wed Mar 29, 2023 6:59 pm

Phil Brown, I would be very interested to see what your condenser is and how you set up condenser + sample + scope in one photo.

Since yours is an infinity scope, is it possible that you restricted aperture after the objective (before the tube lens / somewhere inside your scope), without knowing it?

How you mount the camera or what camera it is does not matter. So you don’t have to include those details.

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Re: Doing Diatoms - part 2

#24 Post by apochronaut » Wed Mar 29, 2023 8:55 pm

Phill Brown wrote:
Wed Mar 29, 2023 6:45 pm
I stand corrected.IMG-20230329-WA0006.jpg
.80.
As for .65 dry being difficult I can only say I took it for granted it isn't.
Condenser is dry.
It's a thread including diatoms in DF.
If anything it's a good example of why not to add a 100x dry for rare occasions.
Lots of things are impossible, getting DF after focusing the condenser is likely one of them.
I don't do it but I'm happy for anyone to keep trying.
All microscope condensers must be focused. You are focusing your condenser by moving it on it's track whether you think you are or not and quite logically, a thick slide requires the condenser to have a different point of rear focus than a thin slide does, since the object the condenser is illuminating is in a different location.
Your slide is quite thick, so obviously the condenser needs to be very high in order to focus properly, hence the tight proximity to the slide bottom. You are still focusing it, though.
zzffnn's thought that your optical pathway may have a restriction has a lot of merit. I would start with the rear diaphragm of the objective. Many of that type have a threaded in plastic rear baffle/diaphragm. If it has one like that and you remove that, do you still get DF with that .80 objective?

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Re: Doing Diatoms - part 2

#25 Post by Phill Brown » Wed Mar 29, 2023 9:42 pm

I'm here to contribute to diatoms in DF, I pulled the objective marked as 100x .80 from the inverted scope and put it on an unmodified freely available scope. The 100x is also available,if it's not what it says that's just the way it is.
If my objectives marked .65 aren't what they say that's just the way things are.
I use stuff that says Canada balsam on it.

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Re: Doing Diatoms - part 2

#26 Post by apochronaut » Wed Mar 29, 2023 11:38 pm

o.k. as it is. DF is a bit tricky for the uninitiated so I feel it necessary to nail down the specifications and dispel assumptions. Specs. are quite specific, assumptions not. New microscopists can end up in a sandwich between the the two. I've been doing DF since 1972. A little while.

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Re: Doing Diatoms - part 2

#27 Post by zzffnn » Thu Mar 30, 2023 4:52 am

apochronaut wrote:
Wed Mar 29, 2023 11:38 pm
o.k. as it is. DF is a bit tricky for the uninitiated so I feel it necessary to nail down the specifications and dispel assumptions. Specs. are quite specific, assumptions not. New microscopists can end up in a sandwich between the the two. I've been doing DF since 1972. A little while.
When I commented above, I also wanted to let other newbie microscopists know that the general rule is that dry condenser will not support DF for NA 0.8 at objective.

Of course, when one modifies the specific settings, (such as mounting a wide tube NA 0.8 infinity objective on a narrow tube 160mm tube scope), actual parameters would change (for example actual NA may fall below 0.65), so the general rule would not apply anymore.

There is always specific equipment designs and settings that may break a general rule, but such occurrence is not common. It is much safer for beginners to follow the general rules (instead of counting on the exceptions to occur).

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Re: Doing Diatoms - part 2

#28 Post by Phill Brown » Thu Mar 30, 2023 5:57 am

My preferred scope is a Watson Hilux 160mm.
Someone offered the Euromex for sale, I paid £270, I offered the seller to back out as they had bought new only a few months earlier and not given it a fair go.
I have a Labophot 2 which I regard as recent and more relevant to the OP who has dedicated many hours to diatoms.
My intention is only to contribute what is realistically achievable,ideally on a restricted budget and where possible doesn't require a lathe.
Whenever I have mentioned this 100x dry objective I've tried to make it clear I would not recommend it.

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Re: Doing Diatoms - part 2

#29 Post by Phill Brown » Fri Mar 31, 2023 6:14 pm

IMG-20230331-WA0007.jpg
IMG-20230331-WA0007.jpg (74.88 KiB) Viewed 38120 times
This is with a x20 .5 abbe condenser and DIY patch.
IMG-20230331-WA0009.jpg
IMG-20230331-WA0009.jpg (76.85 KiB) Viewed 38120 times
Same abbe condenser with DIY patch x40 .65
Watson microsystem 70.
As I am still not winning with stacking they are unprocessed.

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Re: Doing Diatoms - part 2

#30 Post by Sure Squintsalot » Wed Apr 26, 2023 4:54 am

"Doing" diatoms involves way more than simply catching them. I'd argue that catching them is the easy part. The preparation, mounting, viewing, photography, and post-processing are the hard parts of "doing" diatoms. Never mind the literature research that goes into understanding what these things are! Sure it can be fun, but man! There are a lot of frustrations along the way. A few fruits from some labors:


Part of a radiolarian. I'd guess.
Screenshot 2023-04-25 223208.jpg
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Not at all sure what this is, but I've seen a fair number of them in my Pacific Ocean plankton drags.
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Screenshot 2023-04-25 223253.jpg (89.76 KiB) Viewed 37722 times
I've no idea what these are. I don't even ever see parts of these, just entire units like this one here. Whatever glue is holding the legs together at the center must be pretty strong!
Screenshot 2023-04-25 223412.jpg
Screenshot 2023-04-25 223412.jpg (70.6 KiB) Viewed 37722 times
In plan view, their star-shape is obvious. What ARE these things?
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Screenshot 2023-04-25 223441.jpg (122.95 KiB) Viewed 37722 times
Things worth posting in this thread
  • Sewing up a respectable plankton net out of $5 in materials
  • Substituting photoshop "Spot Removal Tool" for clean optics
  • Air travel with your plankton collection/preservation kit
  • Interesting scientific articles in plankton science
  • Plankton photo vs. drawing- what's better?
Anyone else want to pitch in?

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