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PostPosted: Sun Dec 02, 2018 3:23 pm 
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I recently purchased an AO820 microtome and I am now experimenting with it. I have been able to make a few sections with it but it turns out that the cell walls are shattered in all of them. Some of the sections are so bad that none of the cells are intact. I have no idea what I did wrong. Can someone help me?

I dehydrated the sections with the following series:
70% IPA for 2hrs
90% IPA for 2 hrs
100% IPA 2hrs,
100% IPA 2hrs,
100% IPA, overnight
toluene, 2hrs,
toluene, 2hrs,
molten paraffin wax 2hrs,
molten paraffin was 2hrs,
molten paraffin was 4 hrs

After sectioning I rehydrated using the following series:
2 changes of toluene for 30seconds
2 changes of 100% IPA for 30s
70%ipa for 30s
water for a few minutes

I also used a different way of rehydrating with xylene and ethanol with even worse results.


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PostPosted: Sun Dec 02, 2018 5:14 pm 
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Hi DeeJay,

I think you didn't mention what it was that you actually cut - the tips of your fingers, a shark tooth or a flower stem? :D

Bob


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PostPosted: Mon Dec 03, 2018 1:48 am 
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Hi, need more details to help. Have you any images, what are you sectioning, what type of blade are you using (e.g. disposable or steel re-sharpening), what thickness are your sections, how are you floating the sections before placing on slides, wax-type (melting/using temp for infiltration and embedding stages)....

As much detail as you have will help greatly.

John B.

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PostPosted: Mon Dec 03, 2018 9:28 pm 
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I am using a steel re-sharpening blade to cut plant material. I tried sectioning the stems of a hydrangea leaf. I also sectioned sellery. I tried both 20µm and 7µm cuts. Both had shattered cells. I included a picture of what it looks like. The first attempt I made with the 20µm cut was a bit better. It had much more stucture left. Unfortunately, I didn't take a picture of those.

I am floating the sections using a bowl of hot tap water since I don't have a tissue floating bath. I am using wax with a 56C melting point.
Attachment:
shattered cells_small.jpg
shattered cells_small.jpg [ 213.2 KiB | Viewed 1879 times ]


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PostPosted: Mon Dec 03, 2018 10:13 pm 
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Is this tissue on the slide and de-waxed?
Ideally an image of a section still in the wax section but dried onto a slide before de-waxing.

From this image I can't really tell you anything I'm afraid. Also are you able to take an image of your knife's edge under a stereo 'scope so that I can see it's condition, although a complete wax section would suffice for this also.

This is a complex question with a similarly complex answer and much more information is needed, including your stage of processing right from cutting the plant into pieces before you begin....

I can help you with this, but have nothing to go-on with this image I'm afraid.

Here's a good section, with good tissue integrity, as imaged still in the wax as mentioned above - no tears or nicks in either the wax or the tissue it'self,
Attachment:
ws_sonchus_asper_in_wax (1)_stitch.jpg
ws_sonchus_asper_in_wax (1)_stitch.jpg [ 106.69 KiB | Viewed 1874 times ]


This is a tragically poor section with scores in wax from a poor knife-edge, torn tissue from poor processing, etc,
Attachment:
ws_F1_in_wax_before_drying.jpg
ws_F1_in_wax_before_drying.jpg [ 331.97 KiB | Viewed 1874 times ]


Images such as these will be needed to give you useful advice, of the sections on slides and dried but not yet de-waxed.....

John B.

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PostPosted: Tue Dec 04, 2018 9:16 am 
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I'm absolutely no expert on this matter but I would suggest to make thicker cuts like 40µ or 60µ as a starting point. You knife edge has to be really sharp and honed for good cuts. You rotary microtome can't do a slicing cut so the knife really has to be perfect. The knife edge needs a free angle to the material so the steel does not rub on the block after the cut is done. The necessary free angle depends on the shape of the knife directly behind the cutting edge. How do you keep your knives sharp and honed? Some knowledgable people hone the knife directly before cutting to acheive the best results.

Bob


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PostPosted: Tue Dec 04, 2018 9:19 am 
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Here is a link to a good beginners document in german language about knife sharpening: http://www.mikroskopie-bonn.de/_downloads/Abenteuer_Klingen_Schaerfen.pdf


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PostPosted: Tue Dec 04, 2018 7:10 pm 
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I tried inspecting the section under the microscope directly after cutting but it appeared to be opaque. I could not get a clear image so I proceeded to rehydrate it. I didn't heat it after cutting. Maybe that would make it clear again.

I own a stereomicroscope so I can get an image of the knife. It is not a trinocular one so I'll have to improvise. I don't think I have time for that before the weekend. To be honest I didn't sharpen or hone the knife. I just bought them second-hand online. The add said that the knife was sharp and ready for use but now I am not so sure.

So far, I have been unable to make ribbons when cutting. The sections always curl up or crumple. I remedy that by using a thin brush to push the start of the section down. The sections also tend to stick to the block rather than to each oher. I will make some thicker cuts and take a picture before hydrating.

Thank you for your tips. I wouldn't know where to start myself :).


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PostPosted: Tue Dec 04, 2018 7:47 pm 
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DeeJay wrote:
I wouldn't know where to start myself :).

This is a very good place to start:
http://user.xmission.com/~psneeley/Personal/AO%20820%20Microtome.pdf

MichaelG.

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PostPosted: Tue Dec 04, 2018 9:25 pm 
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Oh, when you refer to rehydration, have you floated the sectionson water at about 42 deg C to allow them to expand and relax, before picking them up with a slide and drying then at an angle so that they stick to the slide, then when they are dry they must be immersed in a wax solvent to remove the wax. The, the rehydration may begin, if you intend to stain them with aqueous stain/s.
The entire process is very long and must be done correctly, an un-sharpened knife is of no use whatsoever I'm afraid.

You need to be certain you understand the complete process, of which sectioning is only part, before having a chance to produce sections and permanent slides.....

Have a look over some of my older posts where I have progressed through the entire process from a total beginner to making permanent slides - there's an awful lot of learning and work to be done old chap. I started to learn sectioning in the first part of 2015 I think, that's where my posts will begin.

John B.

Can you give a complete description of your entire process, before and after sectioning, right from the selection of a plant to use - a lot I know, but there's no shortcut with sectioning I'm afraid. The more you are able to tell us the more we can help.

p.s.with an unprepared knife you will not be able to section successfully - believe me, I know this to be true....

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PostPosted: Fri Dec 07, 2018 8:04 pm 
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Yes, I have floated them in water to allow them to relax but the water was closer to 50c. The complete process I used is as follows:
I cut 5mm pieces of the stems of the leaves of the hydrangea leaves. I put them in FAA for a couple of days. Then I dehydrated the sections as described in my original post. I don't believe I left anything out.

I took some pictures of the microtome blade with my stereo microscope. I added them to this post. Two of them are zoomed out and show most of the area that i was using. The others are at 40x magnification. Now that i have been able to view them on the screen, they look horrible.

So what is the best way to sharpen them? I have a whetstone that I use for my kitchen knifes. Can I use it for microtome blades as well?

I will try to take some pictures of the sectioned slide still covered in wax tomorrow. I have an adapter for my camera now so I should be able to take good pictures. In the mean time I will take a look at John B's posts.


Attachments:
File comment: zoomed out
right side.JPG
right side.JPG [ 276.66 KiB | Viewed 1728 times ]
File comment: zoomed out
left side.JPG
left side.JPG [ 264.4 KiB | Viewed 1728 times ]
some more damage.JPG
some more damage.JPG [ 176.45 KiB | Viewed 1728 times ]
rough edge.JPG
rough edge.JPG [ 179.6 KiB | Viewed 1728 times ]
more or less undamaged.JPG
more or less undamaged.JPG [ 210.08 KiB | Viewed 1728 times ]
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PostPosted: Fri Dec 07, 2018 8:23 pm 
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ARRRGGGHHHHH!! :o :shock: :!: :(
This brings back hideous memories..... :D

Hi, yes, the knife-edge is just the way mine went when I started with a 'rocking microtome' and such a steel resharpened blade.
I soon discovered that resharpening the blade is virtually impossible, to get both the finish and the angle needed for microtomy - this I learned very quickly!

My advice to you is to find then buy a disposable microtome-blade holder - of the 'low profile' type. Then of course you will be free of this tyranny and be able to use disposable blades, as do I with my Shandon rotary, very similar to your machine.

You'll see these details within my posts here as I started here from scratch and posted my long and hard progress. The acquisition of a reusable/disposable blade-holder and low-profile blades is absolutely essential in my experience.

John B.

Here's a holder that should fit your 'tome,
Here are the blades that go into it.

You can do it, but this is a vital requirement I'm afraid.....

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PostPosted: Fri Dec 07, 2018 8:37 pm 
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MicroBob wrote:
Here is a link to a good beginners document in german language about knife sharpening: http://www.mikroskopie-bonn.de/_downloads/Abenteuer_Klingen_Schaerfen.pdf


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PostPosted: Fri Dec 07, 2018 10:38 pm 
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MicroBob wrote:
Here is a link to a good beginners document in german language about knife sharpening: http://www.mikroskopie-bonn.de/_downloads/Abenteuer_Klingen_Schaerfen.pdf

DeepL will be working overtime ...

MichaelG.

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PostPosted: Fri Dec 07, 2018 11:30 pm 
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The Microscope and its Use, by Munoz and Charipper, has a large chapter on several methods for sharpening microtome knives. There are also instructions for examining the edge under the microscope and what to look for.

Unfortunately, I haven't been able to find a website where it can be downloaded for free. Archive.org has a method of "borrowing" an electronic copy, but I've never tried to use that.

https://archive.org/details/microscopeitsuse00mu

Abebooks has a copy for $6.00 plus $3.50 for shipping. I bought a copy from them several years ago, and the microtome knife use and sharpening information is very thorough. I haven't attempted to sharpen any of my microtome knives yet, that's on my to-do list for after New Year.

https://www.abebooks.com/book-search/ti ... noz-frank/

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A/O 10 Series Microstar
A/O 4 Series Microstar
A/O 4 Series Phasestar
A/O 4 Series Apostar
A/O Cycloptic Stereo
Several old monocular scopes in more or less decrepit but usable condition


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PostPosted: Sat Dec 08, 2018 12:31 am 
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Thanks, Rick

I have found the Muñoz & Charipper book at Hathi Trust:
https://babel.hathitrust.org/cgi/pt?id=mdp.39015006851300;skin=default;view=image

Members of 'partner institutions' can download the whole book, but even we mere mortals can view and save individual pages.
... Slightly laborious, but very worthwhile.

MichaelG.

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PostPosted: Sat Dec 08, 2018 6:40 am 
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MichaelG. wrote:
DeepL will be working overtime ...MichaelG.


Hi Michael,
I just mention these documents in german language when I think the fumbling with the foreign language might be worthwhile for the reader or when it is just the only text I know of on this topic. The linked document on knife sharpening has the advantage that it tackles the topic from the amatuer standpoint of today, no lab technician or company representative at hand, a knife in unknown condition... Some microscopy recipes are difficult to follow for the amateur today because they are based on the idea that you have unlimited supply of chemicals, buy matching equipment new, you use it all the time and cost is not important. Also there are sometimes old rules that need to be challenged.

Bob


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PostPosted: Sat Dec 08, 2018 11:04 am 
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MicroBob wrote:
MichaelG. wrote:
DeepL will be working overtime ...MichaelG.


Hi Michael,
I just mention these documents in german language when I think the fumbling with the foreign language might be worthwhile for the reader or when it is just the only text I know of on this topic. [ ... ]

Fully understood, Bob ... and greatly appreciated.

With tools like Google Translate and DeepL freely available, we can all make use of texts in many languages:
My comment was entirely positive ... I probably would never have found that document myself: You presented it, and I thank you !!

MichaelG.

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PostPosted: Sat Dec 08, 2018 11:44 am 
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Hi Michael,
I didn't see you comment negative, I just wanted to make my intentions clear for everybody. I myself found electronic translations quite effective for languages I have a little understanding of or that are written at least in our script. With Japanese or Russian it starts to become confusing, which is a pity as there probably is a lot to be found in these languages. I used to have a Pentax K-30 DSLR, a type that tends to give problems with the aperture solenoid. The russians had gotten quite far in identifying and explaining the problem. In an english speaking Pentax-forum it was possible to find a russian speaker who was able to do the necessary clarifications of the electronic translation and set a couple of repair guys going. :roll:

Bob


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PostPosted: Sun Dec 09, 2018 11:43 am 
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Quote:
Here's a holder that should fit your 'tome,
Here are the blades that go into it.

You can do it, but this is a vital requirement I'm afraid.....


Ouch! That's more than I paid for the microtome :(.

So everybody here thinks that the microtome blade is the culprit here. That leaves me with two options: either I find a way to sharpen the blade or I switch to disosables. I did some reading about sharpening and it seems to involve a lot of skills and patience... and I have neither of them. I think I will look into getting a holder for disposable blades.

Another thing I can try is to send the blade to a company that sharpens blades as a service. The only thing I am afraid of is that they are used to sharpen blades for lawnmowers and chisels. I think sharpening microtome blades is not comparable. What do you guys think?


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PostPosted: Sun Dec 09, 2018 12:44 pm 
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Hi again,

I've been through this exact-same scenario as I was learning how to section from the paraffin-method,as are you now. I also considered sending blades to be sharpened, and soon discovered that that was a no-no due to cost and the fact that the steel knives really last only about 2 days with a perfect edge, even less (perhaps only 2 sections...) when learning.....
I then bought new ones at about £25 each and found the exact-same thing.....

By this stage I had to have a new approach, and thankfully I found one, as detailed in one of my posts from 2015.

Basically to identify the blade as the very significantly major factor I, perhaps somewhat speculatively but even so, sticky-taped a single-sided razorblade to the ruined steel knife and tried this to section using my microtome of the time which was a rocking-microtome, before as now I progressed to the 'Mighty Shandon' rotary that I now use...

The result was indeed a definitive answer - the blade was the culprit - no question.

Here's a link to this thread with pictures that will show you how I progressed

The sooner you get a disposable blade-holder (make certain it'll fit into the 'jaws' of your 'tome of course, but I think the std low-profile holder should be fine) and a pack of blades (I just bought another pack of 50 feather S35 low profile blades for £25 on e-bay as always) the sooner you'll be able to crack-on.

Check-out the link and in the meantime I'll look for some more of my results for you....

John B.

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PostPosted: Sun Dec 09, 2018 12:49 pm 
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Hi again, here's a post of mine where I stuck a scraper-blade to the rockers steel blade and got perfect sections, the next stage that beckoned was the acquisition of a 'proper' microtome that took disposable blades - the rest, including the 'Mighty Shandon's arrival, is history!

Have a look at this thread and my short video therein...

I'll look for some more,

Aha - here's a good one, this thread includes a complete protocol......

Here's a video of the disposable blades in action, https://youtu.be/170HrJbGlo4?list=PL1NN ... qlxZgCyeTo
I haven't embedded it here as this isn't my thread, but just follow this address to my video of the Mighty Shandon and a disposable blade in action....
John B.

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PostPosted: Sun Dec 09, 2018 10:02 pm 
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The images in that thread look amazing, especially considering that they were made with dirt cheap razorblades. You have convinced me: I am going the disposable route.

For a metalworker it shouldn't be hard to create a disposable blade holder. It is basically just two strips of metal with a blade sandwiched between them. I would prefer using the standard razor blades because they are cheap and readily available. Also, you cannot use most of the blade anyway so I don't seen any benefit in using the official disposable microtome blades. Unless they are sharper than the razor blades.

I'd like to try your sticky tape approach first. Except I am going to replace the tape with magnets. They are pretty strong and should be able to hold the blade in place.


Attachments:
improvised disposable blade holder2.jpg
improvised disposable blade holder2.jpg [ 197.9 KiB | Viewed 1600 times ]
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PostPosted: Mon Dec 10, 2018 1:54 pm 
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Looks like good progress old chap!

The entire length of the disposables is used, in stages. The blades are able to be slid along the carrier 'from one end to the other' as it were.
I also use extra-long blades (because I bought a large number of them really cheap a while ago on e-bay.... :D ) of which I use essentially one end to 'rough and prepare' the wax block's face, and the other end to actually make the high-quality final cuts.... A sort-of two in one arrangement if you like.

You need to consider the requirement of a smooth and unobstructed 'run off' for the sections and ideally ribbon to travel across after cutting - your current arrangement seems a little difficult with regard to this aspect - I too found this to be a problem when I made my own clamp with screws, metal bar and wingnuts - then realised there was nowhere for the sections to go after the blade's edge! :oops:

Cheaper blade holders may be bought from India, and our own (U.K.) Brunel Microscopes sell such a holder.

Here's a screen-snip of the Brunel holder,
Attachment:
brunel blade holder.JPG
brunel blade holder.JPG [ 25.23 KiB | Viewed 1564 times ]


As seen, there's a lever on the LH end that clamps/releases the blade - simple, without screws too....

Hope this helps a bit,

John B. :D

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PostPosted: Wed Dec 12, 2018 8:56 pm 
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That holder looks interesting. I am also considering getting a cheaper holder from India. The Indian holder doesn't seem to have a quick release mechanism though.

But first, I'd like to know if I am able to make good 10µm cuts with a razor. I experiemented with the razorblade stuck to the microtome blade in the picture above. I made some pictures with the specimen still embedded in wax. What do you think? Is there any chance that they will turn out ok after hydrating?

Also, I added a picture of the mounted wax block (the cuts shown are not taken from this block) and the water batch. I noticed some white spots on the floating sections. They appear to be bubbles. Is that correct? How can they be avoided?


Attachments:
File comment: floating with bubbles
floating.jpg
floating.jpg [ 173.19 KiB | Viewed 1497 times ]
File comment: the mounted wax block
block mounted.jpg
block mounted.jpg [ 188.9 KiB | Viewed 1497 times ]
hydrangea.JPG
hydrangea.JPG [ 341.32 KiB | Viewed 1497 times ]
selery.JPG
selery.JPG [ 344.32 KiB | Viewed 1497 times ]
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PostPosted: Wed Dec 12, 2018 10:22 pm 
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Excellent my friend! You have just duplicated my own definitive test with such a blade temporarily-mounted. Your problems (EX-problems now, or at least greatly-reduced and now with a firm strategy in sight to eliminate!) have just been very clearly and repeatably revealed with respect to a good start with sectioning.
From this and the future (go-get one from Brunel - they're a fine outfit) acquisition of a holder you will rapidly begin to perfect your sectioning protocol/s and most importantly, understanding - the vital requirement before improvements are able to be formulated and tested of course.

I'd proceed from here as follows if I were you:

1) Order holder and blades (blades from internet are far cheaper and always genuine).
2) Continue with your test-rig for now.
3) Resist the temptation to switch your (main) attention towards processing etc (comments re your images sections following) and concentrate on the sectioning.

You have a major progression at your fingertips here, get the holder & blades and you'll master sectioning - for certain in my experience and my study of these images which are so much more informative.

Comments re images and what they tell me;

You need to attend to the shape of your wax-block's cutting face - the horizontal edges MUST be as parallel as you can get they, just by eye of course - don't over-think this bit.
All faces both horizontal and vertical must be straight and of a clean-looking (within reason) finish after trimming - yours are a little sub-optimal but again, no worries here for you at all.
Keep flotation water covered except when placing in or lifting out sections - dust will appear if you don't.

Sectioning (each block every session) is a two-part process, the 'roughing and 'polishing' of the block's face comprise 1 phase the final sections are the second.

Roughing/polishing - new blade which will not then be used for final-sectioning cuts (as you don't yet have disposables which are able to be used as one end for phase 1, the other end for phase 2, by sliding the blade across the holder as you move from phase to phase.) into the clamp as it is now.
Make first cuts at about 10-12µ smoothly and calmly until full-sections begin to appear - ribboning will be a bonus but at this stage don't worry - individual but well-cut sections are easy enough to handle.
When your happily cutting smoothly complete sections pause after about 20 or so and become attentive to any actual tissue to appear within the sections, then switch to thickness of about 5µ to 'polish' the face for about 3-6 sections if all goes (literally) smoothly in terms of no pieces missing from sections, wrinkles parallel to blade and compression (all sections cut will be smaller in the cut direction that the block-face due to this, all-sections - no worry, you're looking to exclude holes, tears etc.).
When you are ready, switch to 8-10 µ for your final cuts. This is an excellent and by no means over-thick section. It's often thought that thinner is better - it isn't! I section the vast majority of my slides at this range for many solid reasons, not because I can't go thinner - I have sectioned anthers and pollen at 1µ with the Mighty-Shandon)...

You sections and question re bubbles - an easy one!
The bubbles under the sections. A few different reasons exist....
a) water is a touch too warm (i.e. above about 45 deg) - I use between 42-44 deg myself (far too warm would be above say 50 deg in spite of what you'll have read)
b) bubbles are forming on bottom of your water container, not because your container's not clean enough, but because you need to 'stir gently' these peskies away before you begin flotation - they are less common in glass that in my full-on lab flotation black-line jobby - but simple brief stirring gets them away...
c) water not deep enough and slides disturbing these an any unseen bubbles during insertion of slide into water.

URK - just noticed all those bubbles on the surface :o of your water - nooooooooo get rid of these. :shock:

I've a short video of the section onto slide process,


Now, to the sections themselves;
In terms of cutting - very good, no scoring, tissue-tearing/loss or other integrity problems - your blade's edge and basic tissue-processing therefore looks to be off to a good start.

I think they may look a little 'sweaty' which can be a symptom of wax ant-medium still present in the wax, but this may be the view-point - don't worry at all about this - we're looking at the sectioning here.
The shrivelled-looking cells within the section can be, and very often are, quite normal - for the very delicate parenchyma tissue of plants - the walls of which contain in life a huge amount of water and suffer the harshness of processing far more than the more rigid walls of other cell-type for example lignified vessels. The stability of these other cells enable us to make sections that are easily close enough to life morphology for enjoyment and study....

If sectioning a succulent for example, stem or leaf, you'll find sectioning a greater challenge - your images have the look of an aloe-type succulent. They're easy to section but blighters to process - I always use them when I want to have a go at improving my technique for processing. Obversely soft but lignified dicotyledonous tissue is 'just right' - easy to process and easy to section - relatively.

Are they OK to take further - yes I think so - you're learning and these look good for this stage and will give you a lot of heuristic knowledge.

So, sections with bubbles under the actual section - no good at all - bubbles not under tissue regions - fine.
Float onto slides and briefly allow to drain (see video). Place nearly vertical to dry at RT onto slide under dust cover - for at least 24hrs.

De-wax with wax solvent such as Histoclear or Xylene (I only use the former) for 2 changes of about 10 minutes each. Remove solvent with OH (I use IPA) at 100% or nearest you have - 95% (nom) IPA is absolutely fine, for 2 changes of about 3 minutes each then onto OH/water rehydration stages for aqueous staining (I'd suggest Safranin - superb stain at 1% for about 5 minutes).

Water to remove excess stain the move through to 100% OH then to 100% solvent in preparation for resin mount....

Looking good - keep focus tight with your study here - many factors yes, but never of equal relevance at any one time.

Hope this helps - go get the holder and blades (if you have trouble finding cheap blades let me know)

Good luck - you're doing really well my friend! :D :D

John B.

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PostPosted: Wed Dec 12, 2018 10:34 pm 
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DeeJay wrote:
For a metalworker it shouldn't be hard to create a disposable blade holder.


In fact this is not problem at all but it is not as easy as expected by the layman. Most raw metals have tensions in them. When you remove material from them they deform. This is often no problem, but when you talk about µm it is a problem when the blade holder parts move 500 µm.

These knives are cheap and semm to be good: https://www.amazon.de/Tajima-Razar-Black-Klingen-TAJ-10312/dp/B010SRSMIQ/ref=sr_1_3?ie=UTF8&qid=1544653829&sr=8-3&keywords=tajima+razar+black

I have developed a holder for it to use them on a hand microtome and they cut well.

Razor blades are too thin. They would require perfect support to cut well. It is much easier to start with a more forgiving thicker blade.

Bob


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PostPosted: Thu Dec 13, 2018 9:19 pm 
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Joined: Sat Sep 15, 2018 9:42 pm
Posts: 12
Hello John. Thank you for your quick(and lengthy) reply. Without your help I probably would have given up.

You have given me a lot of useful tips. I was already planning to focus on sectioning rather than staining but it is always nice if someone can confirm that you're doing the right thing. The fact that I should cover the floatation bath and the drying sections makes sense but somehow hadn't occured to me :oops:. Also, thank you for your tips to get rid of the bubbles. Stirring shouldn't be hard since I am using a magnetic heater/stirrer to warm the floatation bath. :D I can simply add a magnet and stir until I have reached the correct temperature. Maintaining the correct temperature is more difficult since I don't own a contact thermometer.

Quote:
You need to attend to the shape of your wax-block's cutting face - the horizontal edges MUST be as parallel as you can get they, just by eye of course - don't over-think this bit.

The block on the photo wasn't trimmed yet. That is why it is so glossy. I made the block by wrapping aluminium foil around a 2x2x2 cm wooden block. I poured the wax in the created cup and put the plant material inside. After letting the block harden in the fridge, I stuck it to the wooden block by first trimming the back and then heating it and sticking it on a preheated wooden block. Is there anything I can improve in this process?

About the cheap blades, if you can tell me where to get cheap ones, that would be great. The ones I could find so far were all around $100 for 50 pieces.

I'll go ahead with the new sections I created. I'll hydrate then and see what they look like. If the cells are ok I might even try to stain them with safranin. I may have some. But I will focus on creating good sections first. I think I can improve if I apply the tips you gave me in the above comment.

@MicroBob, since I don't have any metalworker skills I think I'll just go for the disposable blade holder. If I had the skills though, I would have made something myself. It is very satisfying not to be dependent on off the shelf products.


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PostPosted: Thu Dec 13, 2018 9:58 pm 
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Joined: Tue Feb 03, 2015 9:42 pm
Posts: 3211
Location: Cumbria, UK
Hi, sounds like a good plan.

All you need to do with the bubbles is slightly disturb them just with a spoon or something - the stirring bar is not at all necessary my friend, just a complication for your workflow.

When I make just a few cuts for a bit of staining-practice for example I simply use a very-nearly full (of ordinary de-ionised water as bought in a supermarket for car batteries etc - very cheap and my std water for all lab work) 250ml glass lab beaker simply microwaved to about 50 deg, with a plastic petri-dish cover immediately placed over it out of the m/w and into the lab - it will stay in the optimal flotation range of about 40-46 deg for plenty of time before needing a simple re-heat...

To take all temperatures in the lab I use the superbly cheap, accurate (consistent certainly) infra-red 'point & shoot' thermometer with a little laser-like light to show you where your measurement comes from. This is a superbly simple method and may be used to instantly (e.g as you work) measure the surface temperature of water, wax, wax-block face, knife-edge, wax section as it's actually floating, flotation water obviously....
If I were you I'd order one of these beauties right now - you'll use it constantly throughout your workflow.

This is one example on e-bay, very similar to mine, even the same colour,
Attachment:
temp gun.JPG
temp gun.JPG [ 50.85 KiB | Viewed 1457 times ]


I also use this to measure molten wax temperature during embedding - even the wax in the moulds is simply measured.

Incidentally your method to cast your blocks is perfectly fine - but for use with your wooden-block system those rubbery ice-cube trays (silicone?) are perfect, as they are able to be scissor-cut into singles, pairs, 2x2s etc and work perfectly. They are very stable thermally, very easy to remove a newly cast block from and a nice cuboid-shape that's easy-peasy to trim. There's no need whatsoever to pre-heat your wooden blocks either - another chore gone!

Simple method I use when I use wooden blocks as I do from time to time for larger tissue:

Buy a cheap 'mortar-pointing trowel' - a tiny trowel used to replace loose mortar between brick-faces - perfect for this job.
Buy a cheap (about £10) chef's gas blowlamp (runs on cigarette lighter gas) as used to brown things.
Watch my short video about mounting a wax block onto a wooden block here,

The video 'features the trowel and the blowlamp.... It's so easy it's almost criminal!


The lamp's great for heating forceps, wax that's solidifying too fast etc.....

For the wax block embedding using the silicone trays and my home-made oven with programmable cheap thermostat see this early post of mine made when I was a beginner too and going through the learning process as you are now.

This is the post to see the wax-casting embedding in it's early stages....

Plenty more to come as you get going....

Viewer beware the post referred-to above contains images of my early work that some will find horrific and harrowing! :o (me for one)
I've made all the mistakes and learned something valuable from every single one! gulp... Some of the images of my early sectioning are almost frightening to see.... You've been warned good folk! :oops:

John B. :D
p.s. Where are you located DeeJay - I'm in the U.K. ?

_________________
John B


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PostPosted: Sun Dec 16, 2018 3:39 pm 
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Joined: Sat Sep 15, 2018 9:42 pm
Posts: 12
I did some more sectioning with a razorblade. I hydrated the hydrangea sections and the sections from the air root of an orchid. I think that the air root sections still contain some wax, even after soaking in xylene for 4 minutes. So I soaked the hydrangea sections for 12 minutes. They don't seem to contain any wax. I also stained the hydrangea section with safranine. I first added a drop of water and a coverslip to inspect the quality of the section. Later, I added a drop of safranine solution on the edge of the coverslip and let it penetrate. The water under the coverslip became colored but the section did not. Then, I lifted the coverslip and lowered it again. Hence the bubbles.

So, what do you think? Some of the cells still seem to be broken but the majority of them seem fine. I am a bit puzzled though. Some of the cells seem to be empty. Others are fine. Are they so big that at 10µm, I chopped off the top or the bottom?

I think I have a suitable ir thermometer. I have a small one that I use to check the temp of my radio controlled vehicles. I think they can be used for this as well.

I have been looking for sillicone ice cube molds to replace the aluminium foil procedure I was using. It turns out I had a square 2x2cm ice cube mold in my own freezer all along! They are not sillicone but maybe they work just as well. I am going to try that out tonight :D

I am going to try to stick the wax blocks to the wooden blocks using a trowel. I still have two of them from when I was working on the walls of my house. I already bought a butane powered bunsenburner which is easy to use so for now I am sticking with that.

So far, using the razor has seems to be the right choice. I am still looking for an affordable source of disposable microtome blades. I'd be gratefull if you can point me in the right dirction.

Also, I am based in the Netherlands.

Greetings and thanks again for your support.


Attachments:
File comment: Hydrangea, sectioned with a razor and hydrated. Stained with safranine
IMG_0037_klein.jpg
IMG_0037_klein.jpg [ 233.27 KiB | Viewed 1419 times ]
File comment: Hydrangea, sectioned with a razor and hydrated. Stained with safranine
IMG_0034_klein.jpg
IMG_0034_klein.jpg [ 276.86 KiB | Viewed 1419 times ]
File comment: Hydrangea, sectioned with a razor and hydrated.
IMG_0027_klein.jpg
IMG_0027_klein.jpg [ 223.61 KiB | Viewed 1419 times ]
File comment: Air root of an orchid. Some cells still seem to be broken
IMG_0018_klein.jpg
IMG_0018_klein.jpg [ 308.43 KiB | Viewed 1419 times ]
File comment: Air root of an orchid. This looks too dark. I suspect that there is still some wax present, even after soaking in xylene for 4 min
IMG_0017_klein.jpg
IMG_0017_klein.jpg [ 386.7 KiB | Viewed 1419 times ]
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