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=The Application of Trichrome Staining Methods yo Embryological Technique=
=The Application of Trichrome Staining Methods to Embryological Technique=
By J. S. Baxter  
By J. S. Baxter  

Latest revision as of 05:31, 11 February 2020

Baxter JS. The application of trichrome staining methods to embryological technique. (1940) J Anat. 75(1): 137-140. PMID 17104841

The Application of Trichrome Staining Methods to Embryological Technique

By J. S. Baxter

Department of Anatomy, University of Bristol

Thispaper summarizes the results of seven years’ experience with trichrome staining methods in the preparation of several mammalian embryonic forms (including the human) for teaching and research. I have adapted the original methods of Masson (1929) and Heidenhain (1915) for these purposes and the modifications are described below.

Many embryologists seem to be deterred from the use of trichrome stains because they consider them complicated and time-consuming; also, these disadvantages seem to be accentuated by the fact that in their work large series of sections must be stained. My experience, however, is that these objections are more apparent than real; there are only a very few stages in these staining methods where individual treatment of the slides is essential, and the beauty and clarity of the finished sections more than compensate for the slight additional time required for their preparation.

Preliminary Treatment of Embryos

Several points in the preliminary treatment of the material are of special importance. A number of fixatives have been tested and, of these, I have found Bouin’s fluid to be excellent as a preliminary to the Masson staining methods. Formol-bichromate and Zenker-formol are also fairly good. With the Heidenhain method, on the other hand, the best staining results have been found with fixation in a fluid containing mercuric chloride; but a greater or lesser degree of shrinkage takes place. Fixation in 10% formalin has not been found to be satisfactory for this type of work. I have had no experience with the Presbyterian Hospital technique (Lillie, 1988), in which formalin-fixed material is mordanted upon the slide with Bouin’s fluid. In view of the results reported, this method might be worthy of trial with formalin-fixed embryos.

Most of the embryos used in this study required to be decalcified, and, after a number of trials, I now use as a routine measure, when necessary, 2% nitric acid in 70 % alcohol. The action is quite slow (a 35 mm. rabbit embryo requires 7-8 days for thorough treatment, the fluid being changed every two days). The optimum time may be exceeded without untoward results if later the embryo be washed very thoroughly in 70% alcohol to which a small quantity (0-25 %) of potassium alum has been added. This counteracts the slight swelling effects of the nitric acid.

In the further treatment of these embryos my aim has been to combat shrinkage as far as possible, and at the same time to preserve the mutual relations of the embryonic structures. Double embedding in celloidin and paraffin has proved undoubtedly the best method for the preparation of serial sections of embryos of which I have had experience. Sections are cut upon a sliding microtome (using the water-on-the-knife method of Huber, as elaborated by Heuser (McClung, 1929)). The procedure admittedly takes more time than simple paraffin embedding, but the shrinkage is much less and the tissue-relationships are infinitely better preserved.

The dioxan-paraffin method as used by Mossman (19387) has been found of service. Dioxan has been substituted for the higher concentrations of alcohol in dehydration and has been used also as a vehicle in the transition of embryos from alcohol to paraffin. Shrinkage is rather more marked than when double embedding is used, noteWith ordinary paraffin embedding methods shrinkage has been found difficult to combat and the distortion due to unequal expansion of the paraffin sections upon the slide during spreading has later caused complications in reconstruction. I have also tried the methyl benzoate-celloidin-paraffin method (Romeis, 1932) but I have not found the results satisfactory.

Modified Trichrome Staining Methods of Masson

Masson (1929) described a number of trichrome staining methods. Of these, I have found the iron-haematoxylin : ponceau-acid fuschin : light green technique most suitable for embryos.

In order to economize in time and labour during the staining of slides, glass staining troughs and racks are very helpful. I mount my embryological sections usually on 3 x 2 in. slides; so troughs and racks of the following dimensions are most useful: troughs 10x 8x7em., racks 8-5x6:5x3cm. Each rack is grooved to accommodate 10 slides. (20 slides arranged back to back cun be accommodated in one staining rack, but this is not reeommended.)

Steps of the method

(1) Xylol.

(2) Absolute alcohol. If the embryo has been doubly embedded in celloidin and paraffin, pass the slides through two baths of equal parts of absolute alcohol and ether. The object of this is to remove as completely as possible the original celloidin mass used for embedding. I have found that if the original celloidin used for embedding be not fully removed a very undesirable precipitation of stain upon the slide occurs later.

(3) 0-5 % celloidin dissolved in equal parts of absolute alcohol and ether. This fresh celloidin coating is necessary if hot staining solutions are subsequently used.

(4) Remove slides, drain off all superfluous fluid, allow partly to dry in the air and then immerse in 80% alcohol.

(5) 60% alcohol.

(6) 30% alcohol.

(7) Distilled water.

(8) Mordant sections in 5% iron alum heated to 45° C. for 5 min.

(9) Wash with tap water for 2-3 min.

(10) Stain in Regaud’s haematoxylin heated to 45° C. for 5 min.

(11) Place the staining rack and slides in distilled water.

(12) Decolorization is carried out in picric acid-aleohol (as recommended by Masson (1929)). Masson states that this fluid has a rapid action. My experience with embryos has been quite different. Decolorization is usually slow and delicate, which is a virtue. In many instances I have found that 20 or 30 min. may elapse before sections are properly affected. My custom is to take one slide from each stained batch and study the progress of decolorization under the microscope. This then gives a definite idea of the time necessary for treatment of that batch.

The remainder are treated together and the process is stopped just before the terminal stage by immersion in tap water. Final differentiation of each in picric acid-aleohol is then checked by direct observation under the microscope, which takes mercly a few minutes.

Very rarely one finds an embryo where the excess of haematoxylin is rapidly removed by picric acid-alcohol. When the test slide of a series shows this, the remainder should be treated with a weaker decolorization solution (such as 1 part of the picric acid-alcohol to 2, 3, or more parts of 95% alcohol). This is the only step in this method where individual treatment of the slides is necessary.

(18) Wash the slides thoroughly in running tap water (30 min. at least). This removes all traces of picric acid from the tissues, which is very important, and this removal may be accelerated by passing the slides into 70% alcohol for 10 min. or so and then back to water.

After differentiation, all nuclei should be clear blue against an almost colourless background. Incomplete differentiation affects the further steps in the method.

(14) Distilled water for 5 min. :

(15) Diluted ponceau-acid fuchsin stain—1 hr. Details of the preparation of this stain are given by Masson (1929). I have found the proportion of acid fuchsin in his original mixture too strong for embryos and my best cytoplasmic staining results have been obtained by immersion for 1 hr. in 4 parts of the original ponceau de xylidene stain added to 1 part of the acid fuchsin stain, the resultant mixture being then diluted ten times with 1 % acetic acid.

(16) Rinse in distilled water for 1 min.

(17) Immerse in 1 % aq. phosphomolybdic acid for 5 min.

(18) Wash in distilled water for 1-2 min.

(19) Stain now in a 2% solution of light green? in 1 % acetic acid for 3 min.

(20) Pass directly into a bath of 1 % acetic acid for 2 min.

(21) 95% alcohol for 2 min.

(22) Absolute alcohol for 2 min.

(23) Absolute alcohol-ether (equal parts) until the celloidin pellicle is dissolved.

(24) Xylol.

(25) Mount in salicylic acid balsam.

Some further notes may be helpful to other workers. Slides stained by this method seven years ago seem as brilliant to-day as they were when first made. No particular care has been taken to avoid exposure of these slides to excess of light.

Fast Green F.C.F.2 has been tried as a substitute for light green in this method. It is much more powerful than light green; 0-5-0-75 % solution in 1% acetic acid is the correct strength in which to employ it. Collagenous tissue after this stain is blue-green in colour and is especially noticeable after formalin fixation.

Modified Azan Stain of Heidenhain

I have mentioned above that the optimum results with this stain are obtained after fixation in a fluid containing mercuric chloride. Susa’s fluid and mercuric | chloride-acetic acid are the best of these. Again in this method I use staining troughs and racks as described in the other.

Steps of the method

The sections are brought to water in the same manner as in the previous method, but since fixation has usually been in a fluid containing mercuric chloride they are treated with a weak solution of iodine in 70% alcohol for 5 min. to remove any sublimate crystals in the tissues. The excess of iodine is then removed by immersion for a short time in a bath of 50% alcohol, containing 0-25 % sodium thiosulphate. Then the procedure is as follows:

(1) Stain the sections in azocarmine for 45 min. at 55°C. Allow to cool for 10-15 min. Details of the preparation of the azocarmine stain are given in the text- book of Romeis (1982, p. 413). I have used azocarmine G in 0-1 % solution.

(2) Rinse in distilled water.

(3) Differentiate in anilin-alcohol (anilin oil 1 c.c. in 1000 c.c. 90 % alcohol). This differentiation is important and must be controlled under the miscroscope. The object is to extract the stain from the tissues until the nuclei are sharp and of a

1 The light green stain which I have hitherto used is described as “ Lichtgriin F.S. Yellowish”’, and was obtained from Dr G. Grubler and Co., Leipzig. The light green stain manufactured by many American firms, and certified by the Stain Commission, would serve equally well.

2 Fast Green F.C.F. (dye content 94%), certification number N.G.f. 2.

deep red colour. It is often helpful to dilute the anilin-alcohol solution with distilled water. .

When the sections have to be examined under the microscope, interrupt the process of differentiation by dipping the slide several times in acetic-alcohol (prepared as in stage 4). If it has not reached the desired stage, continue by transferring the slide again to anilin-alcohol, re-examining until the extraction of stain is satisfactory.

(4) Wash for a short time in acetic-alcohol (glac. acet. acid 1 c.c. in 100 c.c. 95 % alcohol) to stop the differentiation.

(5) Immerse in 5 % aqueous solution of phosphotungstic acid for 1 hr.

(6) Wash quickly in distilled water.

(7) Stain in the following mixture for 30 min.: orange G. 2-0 g., anilin blue (water soluble) 0-5 g., glac. acet. 8 c.c., distilled water 100 ¢.c. Before use, dilute this with three times its volume of distilled water.

Fast Green F.C.F. has been substituted in this staining mixture for anilin blue. The results have been very satisfactory but 0-35 g. Fast Green must be used instead of 0-5 g. anilin blue.

(8) Wash for a very short time in distilled water.

(9) Differentiate in 95 % alcohol. The slides are examined individually under the microscope at this stage. One must bear in mind that further slight extraction of the anilin blue-orange G stain will occur in the next stage.

(10) Absolute alcohol.

(11) Xylol.

(12) Mount in balsam. :

This azan staining method gives very beautiful results; but it must be stressed that fixation is important, a fixative containing mercuric chloride being recommended. Also, the differentiation of the azocarmine stain with anilin-alcohol has to be carefully controlled.


I have described two trichrome staining methods which have proved in my hands very satisfactory for the study of mammalian embryos. It must be remembered that these embryos were those in which epithelial and connective tissue differentiation had commenced. Young embryos, on the other hand, are best studied after the use of an iron-haematoxylin nuclear stain followed by a simple cytoplasmic counterstain (such as orange G in 1% solution).

Trichrome stains definitely require more care in their application and consume more time than simple embryological stains. I am strongly of the opinion, however, that more use should be made of these methods by embryologists who work upon the later stages of development.


Hetpennatn, M. (1915). Z. wiss. Mikr. 32, 361-72.

Lituik, R. D. (1938). J. tech. Meth. 18, 75-81.

McCuune, C. E. (1929). Handbook of Microscopical Technique, 1st ed. New York: Hoeber.

Masson, P. (1929). J. tech. Meth. 12, 75-90.

Mossman, H. W. (1937). Stain Tech. 12, 147.

Romets, B. (1932). Taschenbuch der mikroskopischen Technik, 13th ed. Munich.

Cite this page: Hill, M.A. (2020, April 6) Embryology Paper - The application of trichrome staining methods to embryological technique (1940). Retrieved from https://embryology.med.unsw.edu.au/embryology/index.php/Paper_-_The_application_of_trichrome_staining_methods_to_embryological_technique_(1940)

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