Acrylate Histology



SOP Acrylate-Histology



Version 02







1      Aim and Purpose


Preparation of samples for histological analysis. Especially well suited for bone tissue samples.


2      Place of Application


IOBM histological laboratories, Lottestraße 59.



3        Method


3.1       Cutting

Material:           dissecting set, cutting board, container with 80% ethanol, fixed and undecalcified samples, pencil for marking cassettes and embedding cassettes.


Method:            Prepare formalin-fixed tissue parts of interest and place them in marked cassettes.  To avoid drying of the samples, store them in 80% ethanol until dehydration.


3.2       Dehydration

Material:           Samples in marked cassettes, increasing alcohol concentrations in the autotechnicon, acrylate-inflitration solutions.


Methode:          Place cassettes in the metal basket. Open the autotechnicon by pressing the up/down button. Mount the basket in the holder above the container 1 (70% ethanol) If this is not possible, press the turn button until the holder is positioned above the first container. Close the device by pressing the up/down button and set the timer to 24. Start the process by pressing automatic/manual. The indicator for the selected program lights up.




Transfer the samples to the first infiltration solution after completing the autotechnicon run. The samples need to remain in the first infiltration solution for at least 24h before being transferred to the second infiltration solution. The incubation time of the second infiltration step is at least 24h as well. Both infiltration steps need to occur at 4°C. The infiltrated samples can be used for embedding in.


The duration of the steps during dehydration is dependent on the size of the samples. The following protocol is optimized for small samples (approx. 10x10x5 mm):


·        70% Ethanol           1-5h

·        70% Ethanol           1h

·        80% Ethanol           1h

·        80% Ethanol           1h

·        96% Ethanol           1h

·        96% Ethanol           1h

·        96% Ethanol           1h

·        96% Ethanol           1h

·        100% Ethanol         1h

·        100% Ethanol         1h

·        100% Ethanol         1h

·        100% Ethanol         1h


3.3       Infiltration


Material:           Dehydrated tissue in embedding cassettes, glass containers with infiltration solution, refrigerator


Remove cassettes from the autotechnicon.


Infiltration I (24h- at 4°C in the refrigerator)

Methyl methacrylate (MMA) by Merck 8.00590 1000ml destabilized

3,3g Benzoyl peroxide (dried) (BPO) (Merck 801641)

100ml Nonylphenol


Infiltration II (24h- at 4°C in the refrigerator)

MMA 1000ml

3,3g BPO

100ml Nonylphenol polyglycol ether acetate



3.4 Embedding


Material:           bottles with snap-on caps, embedding solution, infiltrated tissue, water filled tray, refrigerator

Embedding solution

MMA 1000ml

6,6g BPO

100ml Nonylphenol


Add starter shortly before embedding:

N,N Dimethyl-p-toluidin Merck 8.00590 250ml

(smaller units of this chemical can be obtained from Sigma-Aldrich)

500µl / 100ml embedding solution


Starting embedding process imediately after adding the starter! Pour 10-15 ml in each bottle wih snap-on cap. Adjust the sample and place a small, pencil-marked paper tag with the sample ID in the bottle. Close the bottle air-tight wih the snap-on cap. Place the bottle in the water filled tray. The water level should be at approximately the same height as the embedding solution within the bottles. Let the samples cure at 4°C over-night (or longer).


MMA used here is principally destabilized via aluminium oxide!


3.5 Sectioning

Driving out the blocks


Wrap the bottles with the cured blocks inside in a towel and break the glass by striking them with a hammer. Carefully remove the towel and dispose of the glass shards.


The blocks are now ready for sanding.


Preparation of microscope slides

Chrome alum solution

4g Chromium (III) potassium sulfate

100ml Aqua dest.

Filter after complete dissolution and store at 4°C



0,5g Gelatine

50ml Aqua dest. heat (not over 60°C) until completely dissolved

add 2ml chrome alum solution. Mix well and add 3 small thymol crystals. Store at 4°C. Use a Pasteur pipette to place a drop on each slide. Spread the gelatine evenly with a finger.





Roughly sand the block down to a diamond shape around the sample in order to remove excessive acrylate:

The face of the block can then be carefully breached by fine sanding until the actual sample is reached. Make sure that the surface is smooth.

The steepness of the diamond edges determines the cut face. The larger the area, the more of the blade will be worn down.




Use a rotational microctome with a blade suited for cutting acrylate blocks.

The blade can be marked to ensure that the blade is worn down evenly. The first cuts until the right layer is reached can always be done with the same position on the blade. Move the blade to an unused (or less used) position for the actual sections. For these also sectioning fluid should be used. Apply the fluid with a brush and use it for pulling the section onto the blade as well.


Thickness of sections:

Toluidin blue stain: 4µm

van Gieson/ von Kossa stain: 4µm

Calcein label: 12µm





Example for commonly used blade markings: A= initial cutting area; X= worn down position; between the two lines on the left= position currently in use.




Always cut the sample from a pointed, narrow side. Wider sample edges or cortical bone should be positioned in the upper part of the block and sectioned diagonally.

                   Right                               Wrong                              Right                              Wrong



Carefully pull the sections on the gelatine coated slides and apply a few drops of stretching solution.

Stretch the sections with two brushes to avoid folds and wrinkles. Cover the slide with a piece of poly ethylene foil and place it in a dry block. From now on apply a little pressure at all times, e.g. by placing a 1 cm³ piece of steel or lead on the dry block.


When all sections of one session are completed, stack them with some empty slides on top and bottom and place them in a press. Moderately tighten the press and let the sections dry over-night at 60°C.


Take the slides out of the press and remove the PE-foil before placing them in a staining rack.

Use only plastic racks for von Kossa staining!

Calcein labeled sections can be directly covered with mounting solution and a glass coverslip.

Sections for toluidine and von Kossa staining need to be deplasticized. Incubate for at least 15 min in 2-Methoxyethyl acetate (Merck 806061). Subsequently the sections need to be hydrated by transferring them through a decreasing ethanol concentration series. 1 min each: 2x 100%, 2x 96%, 80%, 70%, 50% Ethanol and finally once aqua dest.

Following this the sections are ready for staining.



Sectioning fluid

1l aqua dest. + 1ml Tween 20


Stretching solution

80% Isopropanol with a drop of monoglycol-butylether