Doing Diatoms

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Charles
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Re: Doing Diatoms

#451 Post by Charles » Thu Mar 02, 2017 11:56 am

Good stuff Rod!

I use a squeeze bottle to direct a stream to forcefully rinse down the sides of the tubes as I add the water changes. I first direct the stream to the bottom to mix up the bottom concentrate and then down the sides. Here is an example of the squeeze bottle I use: http://www.ebay.com/itm/3-x-16oz-500ml- ... SwzaJYBMdY

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Re: Doing Diatoms

#452 Post by rnabholz » Thu Mar 02, 2017 2:14 pm

KurtM wrote:This is great stuff. Rod's methods diverge from mine, but it's a grand thing because it means there are two paths being explored, and I couldn't care less who the trailblazer is -- I'm intensely interested in finding the best route from A to B, however, and may the best method win!

I will add this for general interest: I find there's a big difference between centrifugation action in the electric machines versus the hand cranked variety. I would have thought that spinning is spinning, and the means of making the buckets rotate could have nothing to do with the results. But what do I know? The original Youtube video (link below) suggested 3 minutes in electric centrifuge or 1 minute in a hand cranked one, and my own experience is consistently that five minutes in my electric centrifuge barely accomplishes what 45 seconds does neatly in my hand cranker.

Ref: https://www.youtube.com/watch?v=l-uN2RPvDSM
More than one way to skin a cat as they say.

Can't explain why the hand crank seems to work better, that would seem to defy logic, but the proof is in the result.

I do think that not having to acquire a centrifuge at all may help encourage some to give diatoms a try, and that certainly should be marked in the plus column.

Hope everyone continues the collaboration, and others DO join in the fun.

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Re: Doing Diatoms

#453 Post by rnabholz » Thu Mar 02, 2017 2:27 pm

Charles wrote:Good stuff Rod!

I use a squeeze bottle to direct a stream to forcefully rinse down the sides of the tubes as I add the water changes. I first direct the stream to the bottom to mix up the bottom concentrate and then down the sides. Here is an example of the squeeze bottle I use: http://www.ebay.com/itm/3-x-16oz-500ml- ... SwzaJYBMdY
One of those bottles sits on my bench, and as you say, is very handy to have around.

I should have been a bit more precise in my description of the issue. The diatoms attach to the side of the cylinder during settling with the cylinder full of rinse water. Using the syringe to remove the water obviously lowers the water level in the tube, causing the Diatoms to come free of the wall and re-suspend.

The syringe tube must certainly gather some of these during the removal of the rinse water. Again, I don't think the loss is catastrophic or comprehensive for any given type, but is a consideration if your sample size is small, or you are targeting a form that you believe to be present but rare.

I read somewhere that diatom loss during cleaning and processing is unavoidable, but that doesn't mean we won't always wonder about the "one that got away", and that applies to whatever method or protocol you might choose to use.

Thanks,

Rod

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Re: Doing Diatoms

#454 Post by rnabholz » Sat Mar 04, 2017 4:16 pm

As I worked on the latest set of slides from the recent samples, I realized that I had never really talked about the process of getting mounted slides ready for ringing and labeling. So here are a few thoughts on that process.

After the process of mounting the slip to the slide, there are usually some things that that need to cleaned up before ringing or labeling the slide. Mountant can flow out from under the slip, or spatter during the curing process leaving spots or rings around the edge of the slip
Slide Prep Dirty.jpg
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While I know that some use solvent to clean these things up, I found that it tended to just spread things around. I found it much easier to just scrape away the problems.
Slide Prep Kit.jpg
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Using a scalpel it is very easy to just scrape the cured Pleurax away.
Slide Prep Scraped.jpg
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It will leave dust and chips on the slide. Use a dry paper towel to brush off the loose material, then a quick wipe with a paper towel with Windex will remove the last of the dust and clean up any fingerprints or other mess.

With that you are ready to ring. I won't spend much time on that process here as it has been discussed earlier in the thread.
Slide Prep Ringing.jpg
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Onward to Labeling - continued....
Last edited by rnabholz on Mon Mar 06, 2017 1:38 am, edited 1 time in total.

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Re: Doing Diatoms

#455 Post by rnabholz » Sat Mar 04, 2017 4:53 pm

Slide Prep Labels.jpg
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For labels I use Avery 5428 adhesive labels. They are 1" x 3/4" which work very well in the space available on both sides of the coverslip on a 1"x3" slide.

I hand write my labels. I use a 0.5 mechanical pencil, the fine lead allows me to get a pretty decent amount of information in that small space. I also believe that pencil is a better choice than many of the available ink alternatives from the standpoint of maintaining legibility over the years. Many inks will fade over time, and it would be a shame to have the information on the labels lost in that way.

On the left side label, I identify what is on the slide, the mountant type, cure process and add my name. So a typical slide would have "Diatom Strew", "Pleurax" "Inverted Cure" and my name.

The right side label carries, the name of the source, (lake, river, pond, etc) name, then usually a Geographic location, nearest city, town or other landmark. Sometimes a third line with more location information, and finally the date of collection.

With the labels written, next is placement on the slides. I use a forceps to take the label off of the sheet and place in on to the slide, it makes handling and precisely placing the labels much easier.
Slide Prep Label Placement.jpg
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One issue that you might encounter when labeling is what I call "cover slip-slip". Occasionally you will find that during curing, the cover slip will move slightly due to the bubbling and movement of the mountant under the cover. Moving it back while the mountant is still soft can sometimes create "waves" in the mountant that show up in the views. So I generally just let it be, the only effect being a wound to the pride of the mounter and of course it can crowd your label a bit at times as seen here.
Slide Prep Cover Movement.jpg
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As you can see, the label barely fit but the viewing "window" is unobstructed, so while I wouldn't enter it in the County Fair, it will still provide great views.

The next step in my process is to coat all of the labels with a clear nail polish. The paper in the labels can be stained by immersion oil, the nail polish prevents that from happening. It also has the effect of fixing the pencil lead on the label so it is less likely to be rubbed off with handling.
Slide Prep Label Protect.jpg
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The labels will show a mottled appearance when the polish is wet, but will dry clear and even with a pleasing finish that sheds any oil that might get on them.

The last steps are storage of the slides and samples. The slides go into storage boxes. I use 25 slot slide storage boxes. I store those boxes on edge, with the slides contained inside positioned with the cover slips facing up to avoid any movement of the slips or mounted material due mountant curing issues.

The remaining sample goes in a vial with its own label indicating "Cleaned Diatoms" source, location and date, and then into its own storage rack.
Slide Prep Storage.jpg
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That's it, next sample please..... ;^)
Last edited by rnabholz on Mon Mar 06, 2017 1:41 am, edited 1 time in total.

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Re: Doing Diatoms

#456 Post by Charles » Sat Mar 04, 2017 5:26 pm

Yes! Great technique. I too have made a mess trying to clean the splattered mountant by using the appropriate solvent, even after scrapping. But, like you said, it just makes a mess. There is still mountant around the edges of the coverslip even after scrapping and the solvent just pulls it out and makes a mess.

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Re: Doing Diatoms

#457 Post by zzffnn » Sat Mar 04, 2017 6:26 pm

Thank you for sharing, Rod!

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Re: Doing Diatoms

#458 Post by KurtM » Sat Mar 04, 2017 6:54 pm

Hey, your scraper scalpel looks just like mine! I ran into difficulties scraping the glass clean ... until it occurred to me to change the blade. Amazing how much it helps to use one that's nice and sharp.

Just thought I'd share that little gem of pure genius. :P
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Re: Doing Diatoms

#459 Post by rnabholz » Sat Mar 04, 2017 8:05 pm

Charles wrote:Yes! Great technique. I too have made a mess trying to clean the splattered mountant by using the appropriate solvent, even after scrapping. But, like you said, it just makes a mess. There is still mountant around the edges of the coverslip even after scrapping and the solvent just pulls it out and makes a mess.
Yes, it never seemed to end! I also worried about compromising the mount. So now I am an affirmed scraper.

Thanks

Rod

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Re: Doing Diatoms

#460 Post by rnabholz » Sat Mar 04, 2017 8:08 pm

Thanks zz.
KurtM wrote:Hey, your scraper scalpel looks just like mine! I ran into difficulties scraping the glass clean ... until it occurred to me to change the blade. Amazing how much it helps to use one that's nice and sharp.

Just thought I'd share that little gem of pure genius. :P
Ever heard the old saying about "Not the sharpest knife in the drawer"? Apply it here as you see fit........... ;^)

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Re: Doing Diatoms

#461 Post by Hobbyst46 » Mon Oct 09, 2017 1:52 pm

From my humble experience with diatoms on the one hand, and some acquaintence with diatom cleaning methods in research as described in the literature, as well as with chemicals, may I suggest avoiding H2SO4 (sulphuric acid) at ANY home laboratory.
H2SO4 is a strong oxidant. Even more so when hot. But only when it is concentrated. Concentrated H2SO4 is VERY VERy dangerous. Will burn paper as well as human skin, let alone eyes.
HCl is not an oxidant, hence will not oxidize the organic parts of the alga. It will dissolve calcium salts so it can be used to clean diatoms. But for this purpose, dilute HCl is fine. Commercial household muriatic acid is HCl 36%. Can be diluted 10-wise with water and still do its job at room temperature. However, HCl is still CORROSIVE and will rapidly damage the eyes, nose, throat upon contact. HCl vapors form upon heating and are harmful as mentioned.
35 % Hydrogen peroxide is not playstuff either. It reacts violently with some materials.

One way of diatom cleaning is with household bleach solution. Use a clear solution, not a cream or paste. 3-6% Hypochlorite is fine. It will destroy organic matter of the diatoms within minutes. No need to heat the mixture. Due to its basicisity, it may slightly dissolve the diatom skeleton, so if the treatment lasts more than 4-5 hours about 5 % of the diatoms may be lost. However, this disadvantage is far outweighed by its highest safety in comparison to strong acids.
If you add HCl to the bleach-containing mixture, chlorine gas is liberated. It is as effective as bleach for cleaning diatoms,
BUT IT IS HIGHLY CORROSIVE AND TOXIC. So you can start cleaning with bleach, then decant away the liquid, rinse 5-times with water, then if you wish, add dilute HCl. Weight an hour, decant the acid and rinse. I have cleaned pennate diatoms successfully by this method. Without hedrogen peroxide or dichromate ir permanganate.

May I recommend to anyone who cleans diatoms with chemicals to wear protective goggles, rubber gloves, a lab coat or other protective textile coat and shoes.
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Re: Doing Diatoms

#462 Post by Hobbyst46 » Mon Oct 09, 2017 1:55 pm

(Continued) - Sorry about that typo - Wait an hour, not weight an hour...
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Re: Doing Diatoms

#463 Post by wporter » Mon Oct 09, 2017 8:42 pm

Here's a manual for mounting diatoms, coccolithophores, etc., for some additional information:

https://www.google.com/url?sa=t&rct=j&q ... 7O1fJCN4wW


Apologies if it's been posted before. A search here didn't come up positive.

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Re: Doing Diatoms

#464 Post by MichaelBrock » Thu Oct 11, 2018 6:16 pm

I'm finally doing diatoms myself! I have cleaned that sample I collected from the St. John's river via the popular HP + Potassium Dichromate technique. Lots of small particles but seems clear of organic material. I'm in the process of making my first permanent slide using a small bit of Norland 61 I managed to get pending some Pleurax a generous UK member is sending me.

I cleaned the slide & coverslip pretty well w/ dish soap, alcohol, then rinsing in RO/DI water. However, the drop of diatom laden water beaded up in the middle of the cover slip and concentrated the diatoms in the middle. The "How to prepare diatom samples" from Sterrenburg suggests "powdered house hold cleaner". I have both "barkeeper's friend and "comet" but I haven't tried either yet. How do you guys clean your cover slips?

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Re: Doing Diatoms

#465 Post by MicroBob » Thu Oct 11, 2018 6:32 pm

Hi Michael,
your cleaning is probably fine, just add one step: Draw the cover slip quickly with the side you intend to use through the flame of a small alcohol burner. I don't know what happens, but the effect is there (especially if the cover slip shatters due to uneven expansion :lol:).

I only clean with alcohol or white spirit or brake cleaner and then the flame.

Bob

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Re: Doing Diatoms

#466 Post by MichaelBrock » Thu Oct 11, 2018 6:39 pm

Thanks Bob. I can't say that it would have occurred to me but I'll definitely give it a try. I have an alcohol lamp buried somewhere.
MicroBob wrote:Hi Michael,
your cleaning is probably fine, just add one step: Draw the cover slip quickly with the side you intend to use through the flame of a small alcohol burner. I don't know what happens, but the effect is there (especially if the cover slip shatters due to uneven expansion :lol:).

I only clean with alcohol or white spirit or brake cleaner and then the flame.

Bob

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Re: Doing Diatoms

#467 Post by Hobbyst46 » Thu Oct 11, 2018 10:18 pm

MichaelBrock wrote:I have an alcohol lamp buried somewhere.
Twice I purchased cheap 18x18mm coverslips (made in China) from local supply houses, and twice found them to be stained, as if it was dust, but it was permanent whitish spots. Got angry, and tried to clean them with water (failed) alcohol (failed) then liquid dish soap (Hepi or Fairy or Palmolive) - success. Then rinsed with DW, then a final wipe with ethanol-soaked filter paper. They are now transparent. If I place a drop of water on the coverslip, it does not spread as much as I wanted. MicroBob's method, of brief flaming of the coverslip, definitely works. I have done it in the past, mainly to sterilize coverslips. However, if the coverslip is exposed in the flame for too long - that is, a few seconds - it will either shatter or slightly bend and lose its flatness.

Congratulations on your progress! it is a lot of fun! I am on my way to work with NOA61 as well.
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Re: Doing Diatoms

#468 Post by MicroBob » Fri Oct 12, 2018 2:21 pm

These UV hardening adhesives are really interesting for microscopy purposes. There are no solvents and no shrinking.
I have found "LOCA TP-2500" to be a good mountant for radiolaria. It is used to stick new glass screens on mobile phones and is cheaply available. It has good penetration when supported with some vaccum cycles. For diatoms it will not have an high enough r.i..

Please report how you like Norland 61.

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Re: Doing Diatoms

#469 Post by MichaelBrock » Fri Oct 12, 2018 2:24 pm

Passing the cover slip through the flame definitely helped. Not perfect, but definitely better. I made two slides with the Norland 61 and had to wait until this morning to expose them to UV (although the small UV flashlight that I had was enough to start the cure and keep the cover slip in place. I'll need to pick up a UV bulb/lamp. "Overflow" (obviously still figuring out how much to us) scraped off easily before it was fully cured and the remainder cleaned up easily with a alcohol-dampened wipe.

I found quite a few mentions of using Norland 61 in slide preparation but very little by way of recommend process. This is the most detailed one I found:

http://tmitutorials.blogspot.com/2013/0 ... lides.html

and he mentions that bubbles can be an issue. It was in my slides. The first being worse than the second. His trick of heating the bottle upside down to get the bubbles to float to the "bottom" of the bottle might be worth pursuing.

I just did a quick scan through the strew and found lots to like. There isn't nearly as much non-diatom debris as I expected, almost none. Lots of broken frustules though, Including hints of huge centrics. This sample was a quick draw from the middle of the vial though and I'm sure I missed the big ones.

I found a few that really stood out and I'll revisit those and take better pictures. However, I have had no luck finding anything close to being similar in the "North American Diatoms" book from Vinyard or the Diatoms.org site. What other resources to you guys use to identify the diatoms?

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Re: Doing Diatoms

#470 Post by Hobbyst46 » Fri Oct 12, 2018 2:36 pm

MichaelBrock wrote:I found quite a few mentions of using Norland 61 in slide preparation but very little by way of recommend process. This is the most detailed one I found:
http://tmitutorials.blogspot.com/2013/0 ... lides.html
and he mentions that bubbles can be an issue. It was in my slides. The first being worse than the second. His trick of heating the bottle upside down to get the bubbles to float to the "bottom" of the bottle might be worth pursuing.
1. Air bubbles are a problem with all viscous mounting media, including warmed resins. I would hesitate to heat the NOA61 - I suspect that it will shorten its shelf life considerably. Last time I checked, keeping the bottle in the fridge was the way to have it last longer.
2. When using resins (Naphrax, Pleurax etc) the protocol includes a step of heating the slide to a high temperature, that drives off the bubbles.
3. The best weapon against bubbles is degassing. By creating vacuum around the slide. Putting the slide in a vacuum chamber. There were some posts about
DIY nice vacuum chambers for slides in this Forum. Never got to make it yet...
Good luck! waiting for more!
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Re: Doing Diatoms

#471 Post by MichaelBrock » Fri Oct 12, 2018 2:47 pm

Hobbyst46 wrote: 1. Air bubbles are a problem with all viscous mounting media, including warmed resins. I would hesitate to heat the NOA61 - I suspect that it will shorten its shelf life considerably. Last time I checked, keeping the bottle in the fridge was the way to have it last longer.
Good to know. I guess the key is selective photography :) I have the bit I have stored in the refrigerator now in a think envelope.
Hobbyst46 wrote: 3. The best weapon against bubbles is degassing. By creating vacuum around the slide. Putting the slide in a vacuum chamber. There were some posts about
DIY nice vacuum chambers for slides in this Forum. Never got to make it yet...
I'll have a look...turns out I have a few dessicant vacuum chambers and a vaccum pump laying about. Always helps to have another use to justify purchases :)

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Re: Doing Diatoms

#472 Post by MicroBob » Fri Oct 12, 2018 4:22 pm

Under vaccuum some mountants develop even more bubbles. To penetrate hollow particles it is said that repeated applications of a partial vaccuum for short periods is best. I do it that way but I haven't tried to proove this theory.

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Re: Doing Diatoms

#473 Post by Hobbyst46 » Fri Oct 12, 2018 4:34 pm

MicroBob wrote:Under vaccuum some mountants develop even more bubbles. To penetrate hollow particles it is said that repeated applications of a partial vaccuum for short periods is best. I do it that way but I haven't tried to proove this theory.
There are three possible sources of bubbles in mounting. One is the filling of dry frustules with liquid mountant that displaces air. The other is dissolved air in the mountant itself. The third is emission of vapors of a solvent, if the mountant contains such solvent (I believe that the NOA61 does not). The latter two types of bubbles occurs mainly upon heating or the application of vacuum.

Repeated applications, integrated with freeze-thaw cycles, were the method of choice in degassing liquids, AFAIK, but when dealing with viscous mountants, maybe it is not that important whether partial or deep vacuum - if I had a microscope slide vacuum chamber I could test it...


However, it all may depend on the viscosity of the mountant. I just finished the first test of NOA61. The ambient temperature here is 25C, and the adhesive is more like oil than like honey. I put the smallest drop on the 18x18mm coverslip, and it spread - it is slightly too much. To prevent bubbles I heated the slide at 80-85C (roughly) for a couple of minutes, but the result was that much liquid overflowed and crept out and coated the front side of the coverslip - will have to get rid of it... then 45min curing under the UV light (probably a much longer duration than necessary). The diatoms are OK, and there were no air bubbles. I will now repeat it, without heating this time.

Later edit: even without heating, mounting was smooth and easy, and there are no air bubbles!! Hurray to NOA61.
MichaelBrock wrote: I'll need to pick up a UV bulb/lamp
I am using a "black light" home UV tube. Pretty much like a fluorescent tube. Length 60cm, diameter 2.6cm. Connected directly to the house 220V (or 110V). I Installed it over a wall shelf so it only shines downward, and place the slides under it, a few cm below the glass tube. I never look at the tube itself. Guess it is a fairly inexpensive UV lamp.
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Re: Doing Diatoms

#474 Post by MichaelBrock » Fri Oct 12, 2018 7:39 pm

I am more than willing to believe that the bubbles are due to my technique (or lack thereof). I have also found that I do have debris in the slide, it's below the diatoms at slide level. I don't know if it was floating while the diatoms sank or if it was on the slide. We'll see if they are there next time.

Now that I'm "wasting" my work day (a disadvantage of working at home) staring at these slides I'm finding that 1) I really need to tune/clean my microscope 2) I need to figure out the camera controls to improve the image quality, and 3) diatoms are fascinating!

(the blue "thread" is something in the optical path below the condenser; I suspect that it is time open up the bottom again and replace the mirror)
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Re: Doing Diatoms

#475 Post by Hobbyst46 » Fri Oct 12, 2018 8:50 pm

I found somewhat similar blue marks that resulted from an unclean photo eyepiece.
Now I feel like hijacking the post, so will continue in a new one.
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Re: Doing Diatoms

#476 Post by MicroBob » Sat Oct 13, 2018 1:04 am

There are cheap UV lamps of the tube type to cure artificial finger nails. Very laboratory looking in pink! :shock: und

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Re: Doing Diatoms

#477 Post by rnabholz » Thu Oct 18, 2018 12:28 am

[quote="MichaelBrock"

Now that I'm "wasting" my work day (a disadvantage of working at home) staring at these slides I'm finding that 1) I really need to tune/clean my microscope 2) I need to figure out the camera controls to improve the image quality, and 3) diatoms are fascinating!

(the blue "thread" is something in the optical path below the condenser; I suspect that it is time open up the bottom again and replace the mirror)[/quote]

Your findings are consistent with mine, they are fascinating and demanding, but fun.

You have an interesting mix of forms in your image, lots of variety.

One thing you might consider is using the inverted cure process that was discussed earlier in the thread. The idea being that curing the slides with the coverslip down brings the frustules right to the bottom side of the slip. I makes a great difference in resolution and consistent focus across the slide.

I think you would have an advantage using an UV vs Heat curing mountant when it comes to facilitating this process. A 6" square pane of glass raised off the bench a few inches with your UV source underneath would be the perfect surface to place your slides slip down to cure. You might let the the inverted slides "settle" for a time before turning on the UV source to give the diatoms a chance to snug up to the slip.

Nice job so far, keep up the good work and please be sure to report your progress here.

Rod

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Re: Doing Diatoms

#478 Post by Hobbyst46 » Thu Oct 18, 2018 8:26 am

rnabholz wrote:...One thing you might consider is using the inverted cure process that was discussed earlier in the thread. The idea being that curing the slides with the coverslip down brings the frustules right to the bottom side of the slip. I makes a great difference in resolution and consistent focus across the slide.

I think you would have an advantage using an UV vs Heat curing mountant when it comes to facilitating this process. A 6" square pane of glass raised off the bench a few inches with your UV source underneath would be the perfect surface to place your slides slip down to cure. You might let the the inverted slides "settle" for a time before turning on the UV source to give the diatoms a chance to snug up to the slip.
Rod
Thanks for this important and helpful point. May I just add that, if the mountant is UV-curing adhesive, then, instead of the suggested glass pane, I would place the inverted slides (coverslip facing downward) on a steel wire frame that supports the slide by the two opposing ends only, so that the central portion of the slide, including the coverslip and its rims, will be totally exposed to the UV and not through a hard surface layer. This is because (1) the glass pane might block some UV light, (2) more importantly, excess mountant that spreads on the slide outside the coverslip edges may form a drop that contacts the glass pane; due to the UV, the drop might cure and in fact cement the slide to the glass pane.
At least that is my experience.
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Re: Doing Diatoms

#479 Post by rnabholz » Thu Oct 18, 2018 10:35 am

Having no experience with the UV mountant, I certainly defer to those who do.

If outflow around the edge of the slip could cause problems, the micro binder clips that I used for curing Pluerax on the hot plate could be used to hold the slides above whatever surface one might employ.

Whatever the details, I highly recommend the inverted cure process, it improves the finished product a great deal.
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Re: Doing Diatoms

#480 Post by Hobbyst46 » Thu Oct 18, 2018 12:36 pm

rnabholz wrote:Whatever the details, I highly recommend the inverted cure process, it improves the finished product a great deal.
Agreed. Will try to apply it.
Zeiss Standard GFL+Canon EOS-M10, Olympus VMZ stereo

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