Hello,
Beginners question: how long does it take for Euparal in Isopropanol to cure/settle/dry?
6 weeks ago I received a beginners kit from Klaus H. And some example slides from Wolfgang (thanks again)
Started with botanical stem slices (50u thickness approximately) with WAsim III (one combined solution variant) de-hydration with Isopropyl alcohol (10 seconds-30 seconds-2 minutes). Embedded in Euparal in Isopropanol.
Slides are on a warming plate 40 Degrees Celsius for 6 weeks now: and still the Euparal has not cured/dryed completely: when cleaning the slides for observation it is very easy to move the coverglass and distroy the slide.
I received some example slides from Wolfgang dated September 2019 sing Euparal: the are dry/cured completely.
What am I doing wrong?
Guidance appreciated on how to improve my way of working.
Best,
Maarten
Hi Maarten,
ZitatSlides are on a warming plate 40 Degrees Celsius for 6 weeks now: and still the Euparal has not cured/dryed completely:
I put my slides over night into the lab-oven at 50° and then they are done! I can only think, that the thickness is too high. Is it possible, that your cuts are 100 µ or more?
Could you show a picture?
Slide thickness: yes this may be the issue as cutting with the table microtome and the Leitz blades is still a struggle.
Will try and get a picture up tomorrow.
Thanks for feed back.
Hi Maarten,
Euparal takes about 6 months to cure completely. My slides sit on a hot plate at about 45 to 50 degree for one week, befor I do the labeling. That's enough to put them into a slide box in upright possision but one should be carefull when cleaning the cover glass. So Klaus seems to be somewhat hasty ... ;D
If You have a hot plate too You might want to weight the slides with a 8 nut on the cover glass to keep the specimen level.
Good luck for the picture! Camera adaption is not always easy ...
Best
Jörg
First try on a photo. (well trying to figure out how to upload (the upload instruction appear to be in the 2009 edition of the forum and I have no access there).
Feed back on cut (thickness OK?), staining and photo quality appreciated.
(https://www.mikroskopie-forum.de/pictures010/261479_48180537.jpg)
Hi Maarten,
the picture upload has been changed since 2009. I attatch a picture that shows how to do this now. There are size limits to be considered. Irfan View is a nice program to reduce images in resolution and also to reduce to a given file size. It is ideal to use the forums image host as most external image hosts have a questionable future.
The section looks nicely made. There is a bit of residual red stain in the left side that could have been washed out a bit better. On the right side some torn cell walls seem to be drawn over the section. When have you started to make botanic sections? Not bad!
Bob
Bob,
Thank you.
This is my 2nd try on sections and staining and first photo with my newly acquired Trino head.
Maarten
Good Morning Maarten,
the photo looks fine so far. When the whole section ist the same it's not to thick and the artefacts Bob mentioned will vanish when You geht used to slicing. Are You using a microtom knife or single use blades? Even if these are expensive, they should be used only once.
After staining and rinsing with Aqua dest. You can try slow differenciation by leaving the stained sections in Aqua dest. for up to 24 h. This will remove the red from the cells where it does not belong.
Best
Jörg
Thank you all for guidance: new MK2 version: feed back appreciated
- Klaus: practicing on getting thin cuts
- Jorg: New cut with Table microtome, SHK and fresh knife (Yes I was reusing an old knife :-[): so less torn cell walls now
- Bob: Check: I think I need to improve on the staining and washing: we have a lot of green residual stain
Protocol:
* AFE 2 days => 70 % alcohol
* Table microtome, SHK and fresh knife, Schneidehilfe
* A drop of 70% alcohol on knife and sample each cut
* Use a small brush to move the cut
* 70 % Alcohol => 30 %, toilet paper to remove the fluids
* 5 min staining on the slide itself: 2 drops WAsimIII combined solution from Klaus
and a bit of moving/shaking the slide to distribute the stain very 30 seconds
* 30 sec Alcohol 30 %=> 30 sec 70 % => 30 Sec 90% (on the slide): I loose a lot of red stain here
* Isopropanol 30 seconds 3 times
* Euparal
Noot: still practicing on taking pictures as well
Hi Maarten,
then let us take the next step now. :)
- Fixation is fine, You may also try to cut fresh samples and do a slice fixation instead (min. 20 min. AFE, then rinse with Ethanol 70%)
- Important: bring the slices into Aqua dest. prior to staining. This should happen in steps (descending Ethanol row of 70% Ethanol, 50%, 30%, drop some water in, more water when the sections stops dancing ..., water).
- Do the staining for 15 minutes with some heating but not boiling (all steps best done with the slices sitting in a watch glass ....), shake now and then during the staining time
- rinse with Aqua dest.
- leave section in Aqua dest. for 1 to 24 hours to differentiate, change Aqua dest. now and then, You can stop the Process, if the water is clear. There is no danger to overdifferentiation.
- Get the slices in Isopropanol quick (no ascending Ethanol row)
(3 * fresh Isopropanol 100% for some seconds, 2 * fresh Isopropanol for 1 minute, 2* fresh Isopropanol for 5 minutes each).
- Euparal
Best
Jörg
Retry No 3: updated staining protocol as per Jorg guidance:
Protocol v 3 (UPDATES IN CAPITAL)
* AFE 2 days => 70 % alcohol
* Table microtome, SHK and fresh knife, Schneidehilfe
* A drop of 70% alcohol on knife and sample each cut
* Use a small brush to move the cut
* DESCENDING ALCOHOL 70 => 50 => 35 => 0 %
* STAINING NOW IN WATCH GLASES
- (no longer on the slide itself) diluted (2 drops + a little remaining water in watch glass) WAsimIII combined solution from Klaus
-15 minutes on the warming plate approx 50 Degrees Celcius and a bit of moving/shaking the slide to distribute the stain
* DIFFERENTIATION 1.5 hour in aqua dest (mainly loosing some red)
* (skipped the ascending Ethanol steps)
* Isopropanol : twice fast change, twice 1 minute, twice 5 minutes
* Euparal
Observations:
- I think the cut is still to thick?
- Stain remains in some cells: how to prevent / correct this?
note: working on Photo quality and parfocality: updated version
Hi Maarten,
looks fine so far. I think that even the red colour in some bark parenchyma cells would still be removed if they were left in distilled water for a longer period of time. On the other hand: you won't get rid of all artefacts even with a verry, very long time of differentiation.
best
Jörg
Hallo Maarten,
da du ja Deutsch verstehst schreibe ich in Deutsch. Die wichtigsten Informationen hat Jörg dir ja schon gegeben.
Die dunklen Flecken sind gefärbte Zellwände, das hat man oft wenn der Schnitt etwas dicker ist als eine Zelle. Die Zellen sind ja 3dimensionale "Behälter" komplett von einer Zellwand umhüllt. In einem dünnen Schnitt sind die oben und unten liegenden Wände abgeschnitten, so bleibt nur das "Häkeldeckchen-Muster" übrig.
Diese störenden Reste kann man mit einer Behandlung der Schnitte mit Natrium-Hypochlorid entfernen. Handelsname in Deutschland ist Clorix. Es wird nicht konzentriert sonder verdünnt eingesetzt. Über Dauer der Einwirkung und Konzentrationen können dir die Spezialisten sicher Auskunft geben. Nach der Behandlung muss sehr gut ausgewaschen werden eventuell sogar noch neutralisiert mit Essigsäure. Sonst wird die anschließende Färbung durch Oxidation zerstört.
Hi Maarten,
yes, Klaus is right. Sodium dichlorite or potassium hypochlorite bleache the cell walls and dissolve remains of cell content. If used to long, both also dissolve the middle lamella of the tissue and you will end up with single cells at last.
So if You want to use Klorix (the blue bottle, not the green one) You should use 1 part Klorix and 4 parts Aqua dest. You also can use Eau de Javel, which is sold by DM (at least in Germany). To geht used to ist You might also dilute with 2 parts of Aqua dest.
I usually use times around 60 to 90 seconds with lens controll.
Important: rinse until You don't smell chlorine anymore.
Best
Jörg
Jorg und Klaus: danke.
I will practice my cutting to get thinner slides (both fresh and AFE stored) and try the NaClO bleach.
Best
Maarten
Morgen,
New cut and stain (WA-sim III) Ginkgo biloba, leave and leave-stem, collected and stored for 2 months in 70 % of Alcohol (no AFE available at that moment in time).
No klorix used yet.
Still struggling in getting thin cuts (Table Microtome, SHK, Schneidehilfe, lots of Alcohol, new knife for each series of cuts).
The leaves cut relatively easy and relatively thin but still having some torn cell walls.
The stem cuts are thicker and "snap" and jump away when cutting: so probably are to "hard" caused by 2 months in Alcohol?
(I will try to soak the sample for 1-2 days in a mix of Alcohol and Glycerol and see whether this improves the cutting).
Feed Back and tips on moving forward appreciated.
and the stem cut
Guten Morgen Maarten,
Blätter sind schwierig zu schneiden, man muss sie gut einklemmen, damit sie sich nicht bewegen können beim schneiden. Der Schnitt durch den Blattstiel ist doch sehr gut gelungen. So wie es aussieht noch etwas keilförmig. Das liegt sicher auch an ungenügender Klemmung. Ich nehme dafür immer Karotte oder Styrodur.
Zitatso probably are to "hard" caused by 2 months in Alcohol?
Das ist sicher richtig. Alkohol härtet das Gewebe. Deshalb ist das AFE besser, weil die Essigsäure den Härtevorgang ausgleicht. Aber das AFE hast du ja jetzt.
Klaus,
Danke.
When I sampled the Ginkgo I was on holiday: only 70 % of Alcohol available.
Somehow my idea was to start with AFE (2 days-1 week) to stop al the chemical processes in the sample and afterwards move the sample to 70 % Ethanol.
Question: so is it better to STORE the samples in AFE ?
Best,
Maarten
ZitatQuestion: so is it better to STORE the samples in AFE ?
In Ethanol werden sie halt stärker gehärtet. Es ist immer abhängig vom Material, nicht alle Gewebe verhalten sich gleich.
Hello,
As per Klaus H. guidance I now tried to support the Ginkgo leave-stem with Styropor (the green pellets they put in mail-boxes the prevent damage) while cutting.
They are way less damaging to the knife sharpness compared to the Styrodur from the DIY shop.
Still no clean cut and in my impression the thickness is still an issue?
Feed Back appreciated.
Best,Maarten
Hi Maarten,
that is a too thick section for sure. One useful point to remeber is that a section is always the result of not one but two cuts. So if in a cut the knife just scratches over the object instead of cutting it the material is still there and will add to the thickness of the next cut. You might install a magnifying glass on a flexible arm at your cutting place to monitor the cutting process more closely. I try to move the blade along the object with little pressure until I see that it actually enters. Then I finish the cut with a slicing movement. This works quite well for me.
Bob
Bob,
Thank you: I have been practicing cuts for approx. 4 weeks now, Table microtome, SHK, new Leitz knives every time, Schneidehilfe , yes/no carrot or styropor support, lots of alcohol and not getting a cut that is OK.
I really enjoy the exercise including the staining but to be honest having no succes is not vey encouraging.
I took a 1 afternoon course on staining and cutting they were using a "Rasierklingenmikrotom": and the cuts were a bit more thin but damaged: so this does not look like a useful alternative.
I bumped into a rotary microtome (clone from India) but need a to find a knife holder for that one.
Not sure it will be much help as wax embedding the sample is one more step complicating the process.
Next steps, continue practicing with the table microtome:
- Will try your approach (and find a loupe add on for my spectacles)
- As per Jorg's guidance will try to cut some fresh material and apply the AFE etc AFTER the cut
Best,
Maarten
Hi Maarten,
at first: staining seems to be fine now!
Yes, the last section is not thin enough but Ginkgo is not easy to slice.
To get used to Your microtome equipment start with easy things like the petiole of ivy leaves (Hedera helix). They are quite easy to slice and 8 out of ten should be fine. If so, You can be sure to have mastered Your tools and if sections of other specimens are not so good I think You will be able to figure out what to change.
I usually use carrot to embed my samples for sectioning. If You use any plastic it should be Styrodur.
Best
Jörg
p.s.
A rotational microtom or sliding microtome will not help from learning the section technique needed for each of them and they do differ. If You get one, You'll have to start from scratch.
Especially a rotanional microtom is best used for zoomolgical samples embeded with paraffin. On the other hand: in the time I'll get my sliding microtom ready I'll have the sections ready in AFE using my cylinder microtome ... so I normaly don't use it. ;)
There are exceptions: blossoms for example or cones, that would disintegrate without a paraffin embeding ...
Jorg,
Thanks again for feedback.
Next steps, continue practicing with the table microtome:
- Will try Bobs' approach (and find a loupe add on for my spectacles)
- As per your guidance will try to cut some fresh Hedera helix material and apply the AFE etc AFTER the cut.
Will keep you updated,
Best
Maarten
Maarten,
good luck! At the end You won't need a loupe any longer if Your eyes are still strong: if You get no slice or a very thin one, You simply drop the next section and go on.
For example: when doing a new specimen I usually try to get 6 to 8 fine slides. Therefor I need about 12 to 16 sections to be on the good side. When präparitions are ready (Mircotom on the taple, watch glas with AFE at hand, knife holder with a fresh blade, Ethanol and a fine brush ready and specimen fixated in the cylinder) it will take about 5 minutes to get the 16 slices needed in good quality.
So it's usually 10 minutes to build the setup, 10 minutes to clean everything after sectioning and 5 minutes sectioning. ;)
Best
Jörg
Don't get frustraded: when specimens are hard to slice I often get only two out of ten. And I think I know my tools ;)
Hi Maarten,
the cylinder microtome / blade holder method really works but there is some room for errors left. So a bit of experience is needed to get the process straight. I'm sure you will succeed soon.
Some further ideas:
- the object has to be held firmly: The thickness of the section varies a lot when the cylinder moves within the microtome or the object is pushed to the side. I use carrot as it is firm and friendly to the knives. On Jörgs MKB-website there is a description how to prepare the carrot - very useful.
- Does your cylinder microtome have slack between outer and inner cylinder - many newer ones have.
- The blade must never touch anything hard like the table surface. A good check of the blade's sharpness is cutting a firmly held paper tissue.
The rotary microtome is only for paraffine sections. Since the cylinder microtome / blade holder method works so well for botanic objects that hold together well I would suggest to concentrate on this now. Some people nevertheless don't get along well with this method - here an intermediate alternative would be cutting fresh, carrot embedded material with a sliding microtome with angled blade. With the rotary this work work.
Bob
Hallo Bob,
Your observation on my microtome may be spot on, it is 30 years old.
In the market section a tischmikrotom from Swift is offered: would this be a better option?
Best
Maarten
Hi Maarten,
the 30 years won't have done any harm - but was it a good fit from new? Can you wiggle the inner cylinder back and forth? I can't comment on the Swift microtome.
A sloppy fit might be curable: Since this is not a crank pin of a race engine two longitudinal stips might be enough to lead to a closer fit. Typical material for this could be beer can sheet metal. You would look for a three line fit, on one side cylinder angainst cylinder, 120° to both sides a strip that closes the gap.
In my old Sartorius hand microtome the cylinder fits very well - until the screw that closes the jaw is tightened that is. 8)
Bob
Well,
Total failure again in cutting slices of Hedra helix: slices are way to thick and uneven.
Tried the carrot: no succes:
- no there is hardly any play in the inner and outer tube of the microtome
- the underside of the SHK holder bumps into the top of the carrot halfway the cut
(see picture of the underside knife with blue circle)
( the part the knife rests on! So I have the knife sticking out 5 mm extra to remedy this)
- slices are so thick I do not even consider staining them and / or view them under the scope
I'm a bit out of options now.
Hi Maarten,
I can recommend to cut the carrot as described on the MKB website (inside out) it really clamps better this way. Remove the peel completely as it can contain sand!
I don't have an SHK holder but I'm sure that it is not intended that there is a gap between blade and holder. The Leica blades are fairly thin and need all the support they can get. The blade should rest against the back rest. This ensures that the edge is at a constant height over it's length. The cuts are made in a slicing action and this doesn't work well when the blade is not parallel to the base. So there are still a few options! ;)
Bob
Bob,
Thanks again, also for the heads up 8), next steps:
- The SHK has adjustment screws: wil try to get the blade flush to the support
- cut WITHOUT the carrot support only the "schneidehilfe"
I have not yet been able to locate the recommended blades for your printed knife-holder: as I have 2 of those and 2 initial delivered blades will try and switch to using that one instead and see whether the cuts improve.
Best,
Maarten
Hello,
Hedera helix cut: still no success:
* fresh cut => AFE 20 min => Ethanol
* SHK: now with Leitz 818 flush to the bottom (so no gap at te back side between knife and holder: as consequence the cutting angle has changed)
* Table microtome WITH "Schneidehilfe" plus "back-2-back"carrot support as per Bob's/Jorg's guidance: the slices are way too thick
* Table Microtome "Schniedehilfe" only: slices are somewhat thinner now but:
- I think the cuts are still way to thick?
- Also the staining is to intense (15 min WA sim III => wash => 24 Hrs of differentiation in aqua dest => 99.9 Propanol => Euparal).
Your feed back and guidance on next steps is appreciated.
I will start using the Bob's "Printed" knife holder for next cuts to see whether this helps.
Best,
Maarten
Maarten,
yes, You'r right, the section shown on the photo is way to thick.
But you startet with a piece of Ivy sprout, not petiolus. This is much harder to section, especially when embedded with carrot.
If You choose a petiolus that is about 3 mm in diameter You won't need the carrot, the "Scheidehilfe" will do. Sprouts I usually cut without embedding (= freistehend).
Try to move the SHK holder with only a slight pressure: I managed to deformate mine (about half a mm) because I pressed it far to much down on the glas plate of the microtome. No other person was able to get a good cut with it and it had to be mendet at Darmstadt (thanks Detlef!).
Best
Jörg
p.s.
Perhaps You can manage to visit Würzburg in spring 2020, as I will show the cutting technique there.
Ha Jorg,
Thanks for feedback.
This time I switched from SHK to Bob's printed knifeholder with Olfa LBB blades (baumarkt) and Schneidehilfe only (so no carrot support) plus an easier sample to cut: the leafstalk (petiole) of a Pelargonium.
Ik think we are getting somewhere now: as always please feed back on this result.
Best,
Maarten
Hallo Maarten,
und jetzt wäre schon der exakte Vergleich mit einem Schnitt mit dem SHK durch das Pelargonium interessant. Ich habe mit Efeu auch keine guten Erfahrungen gemacht.
Hi Maarten,
this looks great now! And you have selected a beautiful object for you first success - is this Pelargonium or Pelarfonium?
You should be able to get the same result with the SHK holder, I think there is still something wrong with the assembly of it. Can you post a picture of it, especially from the side? One image assembled as used for the last cut and one with the knurled nuts and the top removed and the blade positioned?
My blade holder is cheaper on consumables but won't cut significantly better.
Bob
Hi Maarten,
nothing to complain about. Sectioning and staining fine.
But You should be able to come to the same good results with the SHK assembly.
Best
Jörg
Thnx for all guidance and feed back, next step: the SHK again
Bob: Pelargonium graveolens (corrected: tnx).
Best,
Maarten
Hello,
As per Klaus request I retried the Pelargonium cut with SHK, new Knife, plenty of Ethanol.
(Note: I'm asked questions at the pharmacy on my drinking habits: again 4 new 100 ml bottles of Ethanol).
Wel: no succes: fragmented cuts (zerissen): I must be doing something wrong using the SHK.
Note: I tried an overview picture using a different Photo setup: not very successful as you can see uneven lighting.
Additional SHK cut: twice the thickness
As per Bobs guidance: the SHK knife setup.
Anything here that might cause the unsuccessful cuts?
Hi Maarten,
I can't see any error with the SHK setup. The petiolus You've cut with it seems to be smaller than the one You prepared befor with the printed knife holder, which might be a reason for the different results.
On the other Hand I think we are at a point now, where I must actually see, how You do the sectioning to get some new clue or another. Or You might take your self the time and do some 100 sections of the same specimen with both methods to get a hint what might go wrong ...
Best
Jörg
Jorg: tnx for feed back.
Yes it was a fresh petiolus part, so size may differ a bit.
As for the last 2 weeks I did approx over 750 cuts and nearly 80 slides: so I will give it a few days before starting a 100 cut comparison run. :-\.
Next ambitious project: try and do a CLA (Clean, Lubricate, Adjust) of the BH2 Trinocular 8).
Best,
Maarten
Hi Maarten,
the last sections with the SHK holder look quite good, I wouldn't see the torn cell walls as a safe indicator for a problem with blade or holder, they sometimes simply tear.
In my view the assembly of blade and holder should be ok. The maximum clamping force is needed right behind the edge, the blade should have no play there. The clamping force comes from the knurled nuts in the middle of the holder. The other support should be in the back of the flat area. In the first image of your last post it looks as if there were a gap - true? If there is a gap you might close this by inserting a strip of cardboard. The blade should idealy be securely clamped before the knurled nuts are fully tightenend so the cap of the holder exerts some spring action in pressing on the blade.
750 sections x 0,05mm = 37,5mm! ;D
Bob
Adjusted Photo set-up: stitched 9 pano slides into 1 overview.
(Note: pano stitch as workaround to build an overview picture as I'm using a 4* objective plus micro 4/3 camera plus 2.5* NFK projection eyepiece)
SHK and WAsimIII: slightly thicker cut.
Next step: 50 cuts with SHK and 50 with Bob's printed holder (to practice cutting and compare).
Bob, Jorg, Klaus: thanks for tips and guidance: you guy's are great!
Best,
Maarten
Hallo Maarten,
"not too bad!" ;) ;D ;)
Unser großer Lehrer Dieter Krauter hat vor vielen Jahren einmal gesagt: "Üben sie, Üben sie wenn sie mal 1000 Schnitte gemacht haben werden sie sehen: es geht schon!"
Schöner Schnitt und gelungene Färbung! :)
Guten Morgen Maarten,
da kann ich mich Klaus nur anschließen und danke für die Blumen. :)
Herzliche Grüße
Jörg
Next step: feed back appreciated!
As per Klaus and Jorg's challenge 50 cuts with Bobs printed knife holder + OLFA blades and 50 with SHK + Leitz blades both using the "Scheidehilfe".
Hedera helix leave stem (petioles) this time.
Observations:
- Both series of cuts are slightly to thick
- New knives used after 25 cuts
- Need to change the camera setting for exposure and white balance to full manual next time
- I expect stacking will improve the picture
- All cuts with Bob's knifeholder went smooth (only 1 failure)
- Something must be wrong with the way I handle the SHK knife holder as 30 % of the cuts failed,
and all of the cuts with the second knife required 100 % increase in thickness (picture is from the first 25 thin cuts)
Best Maarten
A new run of cuts to see whether we can get thinner slides:
- 25 cuts: using Bobs printed OLFA knife holder, fresh knife, schneidehilfe, lots of alcohol during he cutting
- most cuts have a damaged centre
- Wasim III using the Klaus instructions (no heating)
- reduced the clearing: 5 minute in aqua dest: to see whether we can hold more of the red colour: no succes
Next try:
- "in between" cuts on thickness
- heating during the last minute of the staining phase
- try and prevent the "dirt" getting into the slides
(think the dirt is caused by lint's of paper-fibre, as I used paper towels to clean the Euparal dropper in between sessions: the Euparal may be contaminated already,
a the Euparal is nearly gone I will have to ask Klaus for a fresh bottle anyway)
Best,
Maarten
Hi Maarten,
that look very well now - nice to see that you continue with the effort! To find the perfect section thickness for object and sectioning process you might change to a 5 sections - review under stereo microscope - next 5 sections routine.
Did you find chrystals in your stem sections? I once have wrecked a blade in an instant by probably cuttling into one.
Concerning the dirt: to me your images look very clean - is that after a lot of cleaning work in image editing? There will be a point of diminishing return: You spend a lot of effort and only save a tiny amount of editing time.
Bob
Bob,
Tnx for feedback: appreciated as I have little comparison material and still have to learn what to look for ...
Have not encountered any crystals yet: good to know this can happen.
Still lacking a stereo scope .....
Actually no cleaning in photo-post-processing yet .
There is some dust on the slides as they are fresh, Euparal only partly dried: so I cannot clean them yet :(
In this case I wouldn't worry about the Euparal and your process, to me your images are very clean.
"Clean" can mean a lot of different things when buying slides - since you look for perfection I would clean them myself before use. In this case here you don't need any special adherence properties so wiping with "Wundbenzin" will do nicely.
Slides out of older boxes (decades) often have damaged surfaces with a milky look that doesn't clean up any more.
Bob,
Tnx again.
I'm using relative older slides (got several 1000 for free), put a full box of 50 of them in a plastic slidebox with holes cut into the box and then the dishwasher .....convenient and helps to remove the white "haze".
Net result is perfect pre-cleaned object-slides.
Using Zeiss coverglasses 1.5 with strict thickness tolerance: clean/un dust them with an old cotton towel.
Thanks for the "wundbenzine" cleaning recommendation.
My issue is the lint and small dirt partikels introduced When building up the final slide (object-slide + sample + Euparal + cover glass). I eliminated one source of lint: fibers from the paper towel used to clean the Euparal-dropper.
Plus the slides collect some dust during their 48 hours stay on the hot-plate and When I take picture after 24 hours (no patience to wait any longer) I cannot un-dust them without disturbing the -only partially cured slide...
Best