Dear Stefan,

interesting question adn aspect of staining undecalcified grounded samples of bone tissue. I would like to ask honsetyl for your input regarding:

1) the resin you use, and naturally

2) the stain /staining method you use.

From what you have stated in your question it might be that H2O2 and /or acidic treatment might be the culprit, but this one can confirm only after knowing more in detail about your technique(s). Regards and best wishes meanwhile, from the "West" to the "East", Wolfgang

Dear Stefan! Based on embedding human bone or teeth in Technovit(R) 7200 or T9100 resin, I use  for Toluidin blue staining first 5% Cirtic acid and than 15% H2O2 with a final clearing in running water to modify the staining intensity. In my experience it seams to be difficult/impossible to get a constant color intensity directly  by staining.

Best regards

Vitus Stachniss

Dear Matthew Ibbs,

In our technique we are grinding down a block with a total height of about 3mm in increments of 40 µm steps, than stain again and photograph it with a scanning microscope. Its a sort of serial grinding not cutting. The surface stained layer, which is usually 5 to 10 mm thick is thereby completely removed. What you see in this kind of histology is always the upper stained part of the block. I cannot explain why the staining intensity declines with the repeated use of acetic acid and hydrogen peroxide. If anything, I would have expected that the staining intensity increases because the tissue is more intensly demikneralized and the plastic is getting more penetrable due to H2O2.

Dear Vitus Stachniss,

that is a lovely stained specimen. My congratulations.

To give an impression of the differences in staining intensity please have a look at the attached serial ground surfaces of a dental implant. Our embedding medium is Technovit 7200 VLC by the way.

Could you please be so kind to give me some further informations about your technique?

In your experience, does it make any any difference if you use citric acid before or after the H202? We use it after, but have never tried it the other way arround.

How long do you usually clear the specimens in running water?

Thanks and best regards

Dear Stefan - thanks for Your feed back, here my RE.

1.  "some further informations": I wrote a hand book, issued in 2005 about my sawing and grinding technique and subsequent digital photo reproduction technic. I send you a link for a dropbox download. From a DFG (Dtsch. Forsch. Gemeinsch.) Research Follower Symposium in 2006 in Ulm/Donau my PPT elaborate is available. Please return your email address to [email protected]

2. "citric acid bevor or after H2O2": I only know concerning tooth material; it is essential to remove the "smear layer" of dentin and enamel before 15%-H2O2 application. The structures will be more clearly stained.

3. "how long clear the stains": approx. between 5 sec and 2 minutes usually.

4. possible reason for reduced staining in the second and further layer: T7200 is highly sensitive against ethanol, which might be part of an staining receipt.

Best regards

Vitus

Dear Stefan,

is this a vanGieson stain, with picrofuchsin as dye? I ask, because in histotechnic the vG needs to be rinsed after staining in 96% ethanol. And it is always a matter of "not too short" and "not too long", because the acid-fuchsin is soluble in diluted ethanol. For paraffin-slides it is recommended to go directly into absolute ethanol (3x), not to loose the color. Picric acid is soluble in water. So the longer the water-rinse, the less intense the yellow.

I find it very difficult to produce exactly the same staining intensity on paraffin-slides. So for my un-experienced and non-digital eye your stainings look very similiar from step to step.

Another point could be the interaction of plastic and dye; and the eveness of infiltration.

Rinsing in acetic acid before staining may improve dye-binding because vG needs a low pH for optimal working. This leads to the wanted ionization of the staining-substrate.

Dear Gudrun,

it is a Giemsa variant we use that is called Levai-Lazcko.

Unfortunatly Technovit 7200 is rather sensitive to ethanol and I fear that it will produce cracks in the block and blur the image.

The stainings look comparatively similar because we increased the staining times in the later steps. It works, but it is rather imprecise and poorly reproducible. We need the  standardization because we want to perform automated histomorphometry and difference in coloring lead to problems there.

Thank you for your suggestions.

Thanks for clearifying. It's rather astonishing, that Methylenblue/Azur II  and Pararosanilin lead to yellow results. ; )

Since I know for a long time the original paper from Laczko & Levai 1975 *) (see below) and have used this sequence also in my work I only would like to add here - for better understanding of novices in the field -  that use of stain (after) < Levai-Lazcko > may be misleading and has been used in this thread (as this happened unfortunately with other papers/references) trivially and perhaps erroneously for several but unknown reasons ("American spelling errors", you can see that googling < Levai-Lazcko >). When a technique used becoming more popular, it is usual & quite normal to be referencing such paper according to the original published paper and the order of authors therein. 

Mrs. Lang, just for your kind information, this method has no Pararosanilin dye stuff added separately to the staining solutions(s). The solution(s) consist(s) only of (basic cationic dyes) = Azure II, Methylene blue and – as counterstaining – one uses either basic fuchsin or Safranin O if adhering to the original reference of Laczko & Levai 1975 . So the stain does not implicate to be designated as a TRUE GIEMSA sequence (despite having some staining effects like…   ).

The quested “yellow” [= resin area] in the images in my opinion is no colour of staining but perhaps due to the treshold of automatic imaging (no correct – “underexposure” for visualizing better the faint blue and red tones visible in the embedded tissue slices.

An earlier published technique using these dye components for staining epoxy resin section (i. e. in LM-resin sections prior to EM) has been: Humphrey CD and Pittman FE (1974), A simple Methylene Blue-Azure II-basic Fuchsin stain for epoxy-embedded Tissue sections; Stain Technol 49 No1, 9-14, which is far superior, reliable and time-efficient as compared to most of the multiple AM-bF (sometimes also ) modifications I have seen and replicated in our EM-Lab.

*) Laczko & Levai1975; Mikroskopie [WIEN] 31(1) pp.1-4,1975

 Laczkó, Jenö; Lévai Géza,:  A simple differential staining method for semi-thin sections of ossifying cartilage and bone tissues embedded in epoxy resin. [Dt. Untertitel: "Eine einfache Färbemethode zur Differenzierung des verkalkenden Knorpel- und Knochengewebes nach Einbettung in Epoxyharz"]:

“….were stained Azure II –Methylene Blue in alkaline solution and counterstained with basic fuchsin or Safranin O in water. Differential staining effects were obtained …..” (which IMHO is a better variant of the so called RICHARDSON -stain for semithin resin sections.

Stain solutions:

1. 0.25% Azure II and 0.25% Methylene Blue in 0.5% Na2CO3  (freshly prepared mixing STOCK solutions:

1% Azure II, 1% Methylene blue, 1% Na2CO3 in water in proportions 1 : 1 : 2)

2. 0.5% basic Fuchsin or Safranin O in water.>

Dear Dr. Muss, thank you. I appreciate your detailed explanation. I looked after the misunderstanding of pararosanlin vs. basic fuchsin. Basic fuchsin with CI 42500 is also called pararosanilin. ( http://stainsfile.info/StainsFile/dyes/42500.htm )

Chapeau!, Mrs.Lang (forgot to consult Conn's 10th Ed. on "Biological Stains, 2002.

In my field (LM/TEM satining resin sections) I haven't seen the term "pararosaniline", just only "basic Fuchsin". Thank you for pointing to this my mistake!

Best regards, Wolfgang

The yellow stain is a result of our special technique. We are only photographing the stained surface of a 3mm high block. What appears as yellow are the deeper lying unstained tissue layers. It is not an effect of the dyes themselves but is caused by the unusually thick tissue block.

Dear Professor Tangl,

I am familiar with the cutting-grinding technique according Donath as we performed it here in the lab at Malmö University.

I am really interested to the method of staining the same plastic-embedded sample at different layers by grinding it down progressively and staining it. I think the picture of the bone-implant block you posted in this thread was of very high quality. 

I have one question: at what thickness do you start the process (I presume thicket than the usually 30-40 µm, but how thin to still have beautiful pictures as those you showed) and how much do you grind away between two layers, to remove the staining? 

Thank you in advance if you find the time to read my question and reply. 

Kindest regards

Silvia

Dear Dr. Galli,

We have meanwhile developed the step-wise grinding into a standard technique in our laboratory. Usually we first grind the specimen to 400 µm, stain it and produce a virtual histology of the plane and then do the same at 250 and 100µm. These three steps give you images that are far enough from each other that the information is not to redundant. Of course the quality at 400 µm is not as good as that at 100 µm, but it is still interesting enough to justify the additional work. We usually don't grind to 40 µm anymore because the risk to lose the specimen is rather high but the quality is not so different too a 100 µm plane (although my teacher Prof. Donath would probably disagree). If you really need too, you can still grind down from the 100 µm step to 40µm.

If you want many layers very close to each other I recommend 20 µm steps. In this way you have removed all the color from the previous step. I should mention that the work and the time that is necessary explodes when you choose very thin layers. The pictures I posted here on Oct 24, 2014 were done in 50µm steps. It can be helpful to glue several (small) specimens onto one Plexiglas slide and process them together.

To give you an impression what the 400/250/100 µm steps look like you can watch the virtual histology using the following links to webimages on our server:

http://histomorphometrie.meduniwien.ac.at/virtual_histology/P244/5826A_1_02_103micron_P244_20x/index.html

http://histomorphometrie.meduniwien.ac.at/virtual_histology/P244/5826A_1_01_252micron_P244_20x/index.html

http://histomorphometrie.meduniwien.ac.at/virtual_histology/P244/5826A_1_00_412micron_P244_20x/index.html

Maybe you will have to copy the addresses and paste them into your browser. You can then zoom in and out with the scroll wheel.

It is a rat femur with a healed defect. The screws are part of the external fixation. The thickness of the step is included in the names of the links.

I hope this helps. If anything is unclear please write.

Kind regards

Stefan