Paraplast from Amazon

[Olympusman]

Yesterday I ordered a kilo of Paraplast from Amazon - it arrived today. Pretty impressive. $20 for the Paraplast and $10 shipping. Search term Paraplast Amazon.

Mike

[discomorphella]

Mike,

You've just saved yourself countless hours of annoyance...your embedding and sectioning will be vastly better than trying beeswax and harder paraffin homebrew mixes.

David

[Olympusman]

I certainly hope so. I found out why my order arrived so rapidly. Electron Microscopy Sciences is in Hatfield, Pennsylvania - about 15 miles south of my front door.

Mike

[yvan_be]

Mike,

You've just saved yourself countless hours of annoyance...your embedding and sectioning will be vastly better than trying beeswax and harder paraffin homebrew mixes.

David

I've used a homebrew mixture of 95/5 W/W household paraffin/beeswax for years and I was very pleased with the results. Cutting artefacts were minor and I managed to cut really thin sections (1.5 - 2 µm) from small samples (onion root tips etc), using ordinary razor blades on a cheap rotary microtome (Euromex Mic 505/506). Ribboning was never a problem, within the usual section thickness range of 4 - 20 µm. Even the somewhat more "difficult" samples (mammal spleen, with trichloro-acetic acid decalcified mammal tissue, ...) sectioned well.

Later on I switched to Merck Histosec and still later to Carl Roth Rotiplast (I still have a 20kg bag somewhere in my garage...). I didn't found those to be superior, compared to my simple home brew. In some cases, such as plant samples with thick and difficult to penetrate cuticulae, on the contrary. Mind you: I never used paraffin mixtures genre Paraplast "Plus", Paraplast "Xtra", the stuff with DMSO or other "plastisizers" added.

I was lucky, I suppose, as the ordinary household paraffin I used (brand SPAAS, Belgium) was probably of exceptionaly high quality.

But I agree on the fact that microtomy has a rather steep learning curve and every problem eliminated in advance is a problem less to solve.

Yesterday I ordered a kilo of Paraplast from Amazon - it arrived today. Pretty impressive. $20 for the Paraplast and $10 shipping. Search term Paraplast Amazon.

Being somewhat stingy, I looked up some prices from my former suppliers. It seems that Paraplast is about the cheapest paraffin for histology availlable. That is: when bought in small packages. It costs here € 18.27 when bought in 1 kg packages. When bought in 8 kg packages, the price drops to € 15.37/kg.

Rotiplast (Carl Roth) is (only) slightly less expensive, costing € 18.15 when bought in 1 kg packages. However, when bougt in 20kg bags, the price drops significantly, to € 9.44/kg.

"Erbaplast", the paraffin for histology from the Italian brand Carlo Erba, costs € 28.12/kg and is only availlable in a 4 * 2.5 kg package.

The Spanisch brand Panreac has a paraffin "for clinical diagnosis", I suppose that means "for histology". Price on demand.
Prices mentioned are all for "regular" paraffins for histology, not containing DMSO.

[discomorphella]

You must have very good paraffin in Europe...I have used the modified paraplast mixtures (plus and extra) and found them to be superior to homebrew mixtures. I've also had lots of experience with home use of Steedman's wax (polyester wax) and paraffin/butyl methacrylate mixtures as well as glycol methacrylate. I prefer the polyester wax since its much easier to use (no xylene or other clearing agents required) and preserves tissue antigens better for immunofluorescence staining since its melts at 39 C. It cuts and ribbons well, but it can be tough to get sections to stay on the slides using regular albumin or gelatin adhesives. I've found that its necessary to coat with nitrocellulose to keep the polyester sections on. Even poly-lysine fails quite a bit. But polyester wax is 10X more expensive than paraplast plus. For home use its not prohibitive if you're not making many blocks, especially considering that 2-propanol is cheaply available at the supermarket at 99% for approximately 3 USD/liter. You don't need any other solvents so you save there. However its an expensive option if you're going to need kg/year.
Mike, the one thing you do need to watch out for is to thoroughly dehydrate and clear your samples. Even paraplast won't embed well if there's residual alcohol in your clearing agent, and if you put a poorly dehydrated sample into your clearing agent (e.g. xylene) if will shrink woefully and be more or less ruined. The old trick was to dehydrate some CuSO4-5H2O crystals (they go from deep blue to a pale blue when you drive off the water of crystallization, and get fragile and powdery) and put them into a small tube or beaker with a cotton or filter paper plug and keep them in the final alcohol jar. If you saw them rehydrating you knew it was time to change out the alcohol baths. I use molecular sieves now, but the CuSO4 works well.

David

[Olympusman]

Thanks Yvan and Discomorphella.
I just infiltrated a section of Eastern Milksnake in Paraplast that still had alcohol in it. I'll try sectioning that, but in the meantime start de-alcoholing another section for infiltration.

Mike

[yvan_be]

Contrary to what one reads in most textbooks, hot 2-propanol dissolves paraffin wax pretty wel.

It can be used as an intermedium between ethyl alcohol and wax. If memory serves me right, the method is described in an edition of the German classic "Romeis Mikroskopische Technik". I tried it a few times and it worked well. I'll see if I can find the book.

[discomorphella]

This is correct, 2-propanol will let you infiltrate with paraffin, but typically its used with a histological microwave oven. I've never tried it with a regular paraffin oven, and no histology lab I've ever worked in has used it outside of a microwave so I am not familiar with using it as a conventional paraffin solvent. Starting out its probably easier to take up more time with a clearing solvent following alcohol dehydration and get your sectioning technique working, then you can try more elaborate methods like hot 2-propanol or polyester wax (which does infiltrate directly from warm 2-propanol easily). Also, you don't need to use expensive (at least in the USA) ethanol. Reasonably dry USP grade 2-propanol (isopropyl alcohol) is readily available at supermarkets and drug stores at least here in the NW. The higher alcohols are also miscible with paraffin, such as t-butanol or 1-butanol; the more hydrocarbon chain(s) in the alcohol the more it resembles a regular hydrocarbon and the less the effect of the hydroxyl group (the OH is what makes the chemical an alcohol, isopropyl alcohol is also 2-hydroxy-propane) has on its properties. Insect and plant histologists use 1-butanol and t-butanol a lot since it hardens tissue less than ethanol. Isopropyl is less hardening than ethanol.
The really important thing is to find a protocol that works for you, i.e., doesn't poison you acutely or chronically, gives good sectioning results, doesn't break the budget etc. Isopropanol as a dehydrating agent and xylene as a clearing agent, WTIH GOOD VENTILAITON and PROPER GLOVES AND EYE PROTECTION is a reasonable starting place. Home depot sells xylene as a paint thinner, so do paint stores. Its not something to work with inside (take your jars outside or in the garage if you don't have a hood and wear good nitrile gloves approved for solvents, and of course use forceps to handle thing that have xylene on them).

[yvan_be]

This is from Romeis' Mikroskopische Technik, 16th edition, 1968:

[Olympusman]

Thanks, everyone, for sharing your experineces with parafin infiltration of specimens. I have learned a great deal from all of you and I hope this thread will continue with more microtomists sharing their techniques.

Mike

[yvan_be]

Can you prepare a really good Irish coffee? Meaning: a layer of whisky at the bottom, a layer of strong coffee on top of that, followed by a layer of freshly whipped cream?

The trick is to put in the whisky and to poor, slowly and gently, the coffee in, so that the two don't mix, but remain as separate layers.

If you can, that's a nice trick to put (delicate) samples trough. I used it to process hydra's, small Aurelia medusae and such. These are hard to manipulate: you can't use tweezers or soft brushes, as they're easily damaged. The only way to manipulate them is by using a wide mouth pipette.

You can use the "Irish coffee trick" for all kinds of samples.

Just take a jar and poor in some xylene. Poor in, slowly and gently, so that the layers don't mix, some 2-propanol, absolute ethyl alcohol, 2-etoxy ethyl alcohol ("cellosolve") or whatever alcohol you use for dehydration. Gently put in the sample, dehydrated in a first bath of absolute alcohol. It wil sinck trough the alcohol layer in the xylene. While the xylene enters the sample, the alcohol gradually leaves. Pippet off the alcohol en place the sample in a second bath of xylene.

For very delicate samples, who shrinck easilly when transferred from alcohol to xylene, a intermediary step can be used. The reagent of choice for that, in the past, was cedar oil. Cedar oil is difficult to get and very expensive these days, but it can be substituded by lavender oil, clove oil, or synthetic clove oil ("Eugenol").

On the importance of complete dehydration: It's interesting to notice that there are some conflicting opinions on the neccesity of *complete* dehydration. Some German authors claim, that at least some (botanical?) specimens benefit from partial dehydration only, prior to paraffin infiltration. In that scenario, the sample is dehydrated up to ethyl alcohol 95%, followed by xylene and wax... I never tried it. I'll look up the source in the next few days.