MicrobeHunter.com Microscopy Forum

Anyone who has tried this is familiar with the medium shrinking excessively as it dries.
About 6 months ago I mounted a small fly in a deep cell (a plastic ring 1.5 mm deep, 1cm diameter); no cover slip.
Let this dry for a week in a closed container (to keep out dust). Then stored it upside down for 6 months.
Today, I added fresh Euparal and a cover slip.
The clear arcs on the blue ring appear to be air bubbles beneath the cover glass, fortunately outside of the ring.

The bottom of the ring seems to have deformed from its original circular shape (?) but the inner circle stayed intact.
The method has promise.

Spacers for insect mounting on slides

Self-adhesive plastic 'paper sheet hole protectors' work OK for making a well-slide.
The ones I have are 15 mm diam with a 7 mm hole; 0.1mm thick.
Here is an ant head mount in Euparal made several years ago (then still learing how to make acceptable 'ringing').
I am sure that one could stack the rings to make a deeper well. Point I am making here is that Euparal has no effect
on the plastic, and vice versa, even after, probably, 5 years.

Just out of curiosity, you just dropped an entire dead fly into euparal and didn't get any air bubbles? Did you use new or thinned euparal? Did you dewater the fly first? Where did you get your euparal, anyway? And how did you manage to get NO air between the cover slip and the well?

Dead fly soaked in 5% KOH for a couple of hours until it looked transparent - time depends on size of insect and how
thick is the exoskeleton; springtails maybe 1 hr, bluebottle fly likely overnight.
Thoroughly rinsed in water. Dehydrate in alcohol. Clear, using something miscible with Euparal (e.g. cedarwood oil);
for Canada Balsam mounting use Xylene.
Place insect on slide in the smallest amount of clearing agent to keep it completely wet.
Add a drop of mountant, can let this thicken for up to 6 months. Normally, after about 1 hour add a final drop.
Carefully add a coverslip.
If coverslip is scrupulously clean and you add it carefully, there should be no trapped bubbles - I rarely have a problem.
I got my Euparal from a UK company; it's no longer listed on their site.

Is the choice of potassium hydroxide over sodium hydroxide just a matter of waiting time?

I make 500 mL of 5% KOH and it lasts for >1 year.
I read somewhere, a long time ago, that NaOH solutions deteriorate over time; I forget how and forget why.
Regardless, KOH works OK for me.
10% KOH seems more like the standard, I prefer the weaker 5% with longer time.

TonyT wrote: ↑

Wed Feb 15, 2023 2:53 pm

Dead fly soaked in 5% KOH for a couple of hours until it looked transparent - time depends on size of insect and how
thick is the exoskeleton; springtails maybe 1 hr, bluebottle fly likely overnight.
Thoroughly rinsed in water. Dehydrate in alcohol. Clear, using something miscible with Euparal (e.g. cedarwood oil);
for Canada Balsam mounting use Xylene.
Place insect on slide in the smallest amount of clearing agent to keep it completely wet.
Add a drop of mountant, can let this thicken for up to 6 months. Normally, after about 1 hour add a final drop.
Carefully add a coverslip.
If coverslip is scrupulously clean and you add it carefully, there should be no trapped bubbles - I rarely have a problem.
I got my Euparal from a UK company; it's no longer listed on their site.

Thanks TonyT....
Your process sounds pretty involved. I wish I had your patience and attention span.
I have an insect specimen bank waiting for me to get done with my load of plankton strews and copepods. Currently, I use Permount for everything and use a vacuum pump and chamber to get air bubbles out; it takes about 20 minutes. Afterwards, Samples get baked at 100deg. C. for a few hours under a light load to flatten things down uniformly. When cooled, the globs of hardened Permount around the cover slip fracture off like glass indicating cured mountant. My oldest heat-cured slides are about 6 months old and I'm not seeing any degradation.

By contrast, some of those slides (that weren't heat-cured) will loosen up if held in hand too long. I'm not entirely sure this heat-curing process would work with the large volume of mountant in a well, but I aim to try.

A few questions, though:
1) What's the value in rendering an insect body transparent?
2) How do you rinse out the KOH without destroying microstructures?
3) What are you going to use when you run out of Euparal?

My microscope, an Olympus BH2/BHS, is designed for tranmitted illumination, thus the need for transparent subjects.
If you use reflected illumination there in no need for transparancy. Specimens examined by reflected light are
probaly best mounted dry.
Removing KOH is done simply by placing the specimen in water for about 20 seconds, and changing the water about
five times.
Use Canada Balsam

KOH and NaOH should be fairly equi-potent for this purpose.
They "deteriorate" over time if stored in an open flask, since they absorb carbon dioxide from air and are converted to carbonates.
Carbonates are less effective for destroying organic matter (insect guts).
Be very careful in handling those chemicals. They can cause extreme burns, especially to the eyes.
Wear rubber gloves and eye protection.
Preferably store them in properly sealed durable plastic containers (PP falcon tubes, for example, are fine), not glass containers.
If only a glass flask is available, it should not be stoppered with a glass stopper, because the chemical corrodes glass and can "glue" the stopper to the flask.