A) Botany

B) Parasitology

                a. Plant parasites
                b. Animal parasites
                c. Filterable infectious agents



c) Zoologie

 

Fixation

Any material taken from a living organism begins to change very quickly. Therefore, various methods are used to preserve and fix the original state. For blood smears, tissue fluid, and bacterial cultures, alcohol or heat fixation is most commonly used. The best method for each is indicated in these instructions. Fixative solutions are used to preserve the structure of tissue sections, of which the following are the most frequently used. A single fixative solution that preserves all cell and tissue components equally well does not exist. Therefore, the following instructions specify which fixative is most suitable for each method.

A. Smear Preparations:
A small amount of specimen material is spread in the thinnest and most even layer possible onto the coverslip or slide using a annealed platinum needle or scalpel. Alternatively, the following method, particularly recommended for blood preparations, can be used: Place a very small drop of blood, etc., onto a slide and hold a second slide at an acute angle to the drop. The drop will then spread along the edge of the second slide. Now, slowly slide this slide along the first slide, which serves as a base, thus carrying and spreading the drop. Pure cultures or material that is difficult to spread are best thinned beforehand with a drop of water. Then, allow the smear to air dry completely. Only then should the coverslip or slide, held with Cornet forceps and with the smear side facing up, be passed through a flame three times. Air drying is accelerated by repeatedly agitating the preparations. Air-dried preparations can be fixed not only by heat but also by immersion in absolute alcohol or a mixture of equal parts absolute alcohol and ether for approximately ten minutes. Staining is best achieved by applying the staining solution dropwise from a pipette. The staining time depends on the type of preparation and varies depending on the staining solution. Generally, 15 seconds to 1 minute is sufficient. Gentle warming accelerates the staining process. Where numerical values ​​can be given, they are listed in the following instructions. After staining is complete, the smears are carefully rinsed with water, patted dry with blotting paper, and dried, which can again be done with gentle warming. The preparation is then covered with a drop of cedarwood oil and examined without a coverslip. If a permanent slide is to be prepared, a coverslip is cemented in place with Canada balsam or Malinol. If special fixation methods are required, the necessary information can be found in the following instructions.

B. Tissue Sections:
The duration of fixation depends on the fixative and the type and size of the tissue sample. Where information is available, it has been provided for the fixative mixtures listed below.

 

Fixation solutions

1. Bouin

 

Pikrinsäure 1% ig wässrig	15 ccm
Formalin 40%	5 ccm
Eisessig	1 ccm

Alkohol absolut	60 ccm
Chloroform	30 ccm
Eisessig	10 ccm

Starkes Gemisch =
Chromsäure 1 %ig	15 ccm
Osmiumsäure 2 %ig	4 ccm
Eisessig	1 ccm

Formalin 40 %ig	10 ccm
Wasser dest.	100 ccm

Formalin 40 %ig	10 ccm
Alkohol 80 %ig	20 ccm

Sublimat	4,5 g
Natriumchlorid	0,5 g
Wasser	80 ccm
Formalin 40 %ig	20 ccm

Sublimat	5,0 g
Kaliumbichromat	2,5 g
Natriumsulfat krist.	1,0 g
Wasser dest	100 ccm
Formalin 40 %ig	5,0 ccm

   

Kaliumbichromat	2,5 g
Natriumsulfat krist	1,0 g
Wasser dest	100 ccm

                         

Kaliumbichromatlösung 3 %ig	80 ccm
Formalin 40 %ig, neutral	20 ccm

    

Sublimatlösung gesättigt wässrig	76 ccm
Formalin 40 %	20 ccm
Eisessig	4 ccm

Sublimat	4,5 g
Kochsalz	0,5 g
Trichloressigsäure	2,0 g
Eisessig	4,0 ccm
Wasser dest	80,0 ccm
Formalin 40 %ig	20,0 ccm

Sublimat	5,0 g
Kaliumbichromat	2,5 g
Natriumsulfat krist	1,0 g
Wasser dest	100,0 ccm
Kurz vor dem Gebrauch zufügen: Eisessig	5,0 ccm

Paraffin embedding:    


 

A. Botany


Astra Blue - Auramine - Safranin
Differentiation of cellulose and lignified cell walls

Required solutions:

A.	Astrablau FM	0,5 g
	dest. Wasser	100 ml
	Weinsäure	2,0 g
B.	Auramin O saturated aqueous solution
C.	1 g Safranin O in 100 ml distilled water. Materials used: Plant sections, embedded in paraffin, synthetic wax, or unembedded. Fixation: Formaldehyde or ethanol.

Staining procedure:

1. Transfer dewaxed sections from water to solution A for 1–5 minutes.
2. Rinse with distilled water.
3. Stain in solution B for 1–5 minutes; the sections will now have a greenish tint.
4. Rinse with distilled water.
5. Stain in solution C for 1–5 minutes.
6. Rinse with distilled water.
7. Dehydrate in acetone (changed three times).
8. Embed over a 1:3 mixture of phenol and petroleum ether in balsam or malinol.

Staining result:
Cellulose cell walls and only pure cellulose are intensely blue. Lignified cell walls range from sulfur yellow to brick red, depending on the degree of lignification. The stronger the lignification, the more intense the yellow hue.

Note: The use of a blue filter enhances the contrast of auramine and safranin.
Lit.: Maácz and Vágás, “Microscopy” Vol. 16 (1961), pp. 40 - 43
Required solutions:
   A. Strasburger's picric aniline blue solution
   B. 0.5 g picric acid in 100 ml 96% alcohol. Material used: Stems of Zea mays.

Staining procedure:

1. Stain sections for 1–2 minutes in solution A.
2. Rinse in distilled water.
3. Dehydrate in solution B.
4. Convert to xylene balsam or malinol.

Staining result:
Lignified elements yellow. Non-lignified elements blue.
Required solutions:

A. Dissolve 4 g of safranin in 200 ml methylcellosolve, add 100 ml 95% alcohol and 100 ml distilled water. Water, then add 4 g sodium acetate and 8 ml 40% formalin.
B. Saturated solution of picric acid in alcohol.
C. Saturated solution of aniline blue, water-soluble in alcohol. Before use, mix 78 ml of B with 22 ml of C Material used: Any lignified tissue.

Staining procedure:

1. Stain the sections in A for at least 2 hours.
2. Wash thoroughly with tap water.
3. Differentiate in B.
4. Stain for 2 hours in the mixture of B and C.
5. Wash in absolute alcohol for 10 seconds.
6. Lighten in clove oil.
7. Transfer in xylene.
8. Preserve in balsam or malinol.

Staining result:
Lignified or cutinated walls are stained bright red,cellulose walls are stained blue.

Lit.: Johansen, 1940, Plant Microtechnique, 1st edition.
Required Solutions:
   A. Prepare a saturated aqueous solution of Auramine O.
   B. Dissolve 1 g of Crystal Violet in 50 ml of distilled water and 50 ml of isopropyl alcohol. Mix equal parts of A and B before use. Materials Used:
       Sections of fresh, preserved, or bleach-treated material.

Staining Procedure:

1. Stain the sections for 5–10 minutes in the mixture of A and B.
2. Rinse briefly in distilled water. Differentiate in isopropyl alcohol (changed once) and dry with filter paper.
3. Transfer to terpineol (changed twice).
4. Encase in Malinol.

Staining Result:
Lignified membranes turn yellow. Cellulose turns violet.

Lit.: Beck, Mikrokosmos 48 (1959): 94 - 95, Franckh'sche Verlagshandlung, Stuttgart.

 
Required Solutions:

  A. 0.2% aqueous solution of Pure Blue
  B. 0.2% aqueous solution of Brilliantcrocein Immediately before use, mix equal parts of A and B. Materials used: As above.

Staining procedure:

1. Stain sections for 5–15 minutes in the mixture of A and B.
2. Rinse briefly in isopropyl alcohol (changed once).
3. Dry with filter paper.
4. Transfer to terpineol (changed twice).
5. Contain in Malinol.

Staining result:
Wood membranes stained red, cellulose stained blue. Literature: As above.
 
Necessary Solutions:

A. Malachitgrün	0,5 g
Acid fuchsin	0,5 g
Alcohol	75 ccm
distilled water	75 ccm

Staining procedure:

1. Add sections to the substrate.
2. Stain in solution A for 10 minutes.
3. Wash in 96% alcohol.
4. Lighten in carbolic-xylene, then embed in balsam or malinol over xylene.

Staining result:
   Membran cellulose: red
   Lyngeal parts:         green
Ref.: Nečesaný, V. (1955), Prirodovedecky Spornik Ostravskeho kraje, 16: 184–202
 
Necessary Solutions:

A.	Ethyl alcohol	90 ccm
	Propionic acid	5,0 ccm
	Formaldehyde 40%	5,0 ccm
B.	Sodium hypochlorite 1%
C.	Potassium hydroxide 10% in 95% alcohol
D.	Sudan Blue	0,5 g
	Butyl alcohol (tert.)	10,0 ccm
	Dissolve at room temperature, add: ethyl alcohol shake well, filter	190,0 ccm
E.	Safranine O	2,0 g
	Cellosolve)	100,0 ccm
	Ethyl alcohol 95% 0g	50,0 ccm
	Sodium acetate 4% aqueous	50,0 ccm
	Formaldehyde 40%	4,0 ccm
D.	Congorot	0,05 - 0,1 g
	Solution D	25,0 ccm
	Solution E	5,0 ccm

Staining procedure:

1. Fix in solution A, 24–48 hours.
2. Cut to 50–100 µm thickness, immerse in 50% alcohol.
3. Extract in acetone, 5 minutes.
4. Bleach in solution B, 5 minutes.
5. Saponify in solution C, 15 minutes.
6. Rinse with 50% alcohol, 3 times.
7. Stain in solution F, 30 minutes at 55°C.
8. Rinse in 50% alcohol, 2 times.
9. Embed in Malinol.

Staining result:
    Cellulose and cytoplasm: light red
    Wood tissue and cork:     orange-red
    Rubber:                            blue
Ref.: Addicott, F.T. (1944), Stain Tech., 19: 99–102.
Johansen, D. A. (1940), Plant Microtechnique, pp. 41–42, 62 (McGraw-Hill Book Co., New York & London).
Necessary Solutions:

A. Safranin	1 g
Alkohol 50 %ig	100 ccm
B. Lichtgrün FS	1 g
Nelkenöl	100 ccm

Staining Procedure:

1. Stain sections in solution A for 5–10 minutes.
2. Wash in water.
3. Dehydrate in an ascending alcohol series.
4. Counterstain with solution B for 2–5 minutes (microscopic inspection).
5. Lighten in clove oil.
6. Wash in xylene, embed in balsam or malinol.

Staining Result:
  Wood fibers: red
  Cellulose:     green
Ref.: Chamberlain, C. J. (1932), Methods in Plant Histology, 5th ed. (Univ. of Chicago Press)
Required solutions:

   A. Dissolve 0.1 g of thionine in 100 ml of 5% phenol solution with warming.
   B. Saturate solution of Orange G in absolute alcohol.
       Materials used: Pathological plant sections, fixative of choice,
       Paraffin embedding.

Staining procedure:

1. Deparaffin as usual.
2. Stain in solution A for approximately one hour.
3. Dehydrate in the ascending alcohol series.
4. Differentiate in solution B.
5. Rinse in absolute alcohol.
6. Embed over xylene in balsam or malinol.

Staining result:
   Parasites purple-violet, cellulose walls yellow-green, xylem blue, chromosomes blue, spindles purple.
Ref.: Conn, 1953, Biological Stains, 6th edition, Williams & Wilkins Company, Baltimore.
 

Required solution:
   Dissolve 5 mg of resorcinol blue and 5 mg of Martius yellow in 10–15 ml of distilled water. Add a few drops of highly diluted ammonia until the solution         has an olive tint.
   Material used: Pollen tubes in styles, unfixed and unembedded.

Staining procedure:

1. Lightly moisten thin styles or ovaries with water and crush between two microscope slides. Alternatively, make longitudinal sections of thicker styles or ovaries by hand and then crush them.
2. Cover the material spread on a microscope slide with the above solution and allow it to act for 2–5 minutes.
3. Carefully aspirate excess staining solution or rinse with a little water.
4. Place a coverslip on top, press down, and view under a microscope with strong illumination.
5. If permanent slides are desired, rim the coverslip with coverslip putty or coverslip varnish.

Staining result:
    Pollen tubes blue, background light yellowish-green.
Ref.: Nebel, 1931, Stain Technology, 6, 27-29, Publisher: Williams & Wilkins Company, Baltimore.
Required Solution:
Bacteriological ink
Materials Used: Pollen dust from hazel, alder, pine,
spruce, coltsfoot, dandelion, pumpkin, and others.

Staining Procedure:

1. Thoroughly mix a small amount of ink with an equal amount of distilled water.
2. Place a drop of the diluted ink onto a microscope slide, pour some pollen dust next to it, and rub them together well.
3. Spread the drop as thinly as possible using a second slide or a coverslip and allow it to air dry.
4. View under a microscope with strong illumination.

Staining Result:
   The pollen grains stand out clearly in their various shapes against the dark background.
Note: Once the smear has dried completely flat, a permanent slide can be prepared by cementing a coverslip onto it.

 


Required solutions:

   A. Fixing solution consisting of 20 ml glacial acetic acid and 60 ml 100% alcohol
   B. Orcein stock solution: 1 g of orcein is boiled in 50 ml 45% acetic acid for 2 hours using a reflux condenser. After cooling, the solution is filtered. Material used: Meristem cells from root tips.

Staining procedure:

1. Fix the material in solution A.
2. Transfer to a mixture of equal parts alcohol and concentrated hydrochloric acid for 5 minutes, then wash thoroughly in pure alcohol.
3. Place on a slide coated with protein glycerin.
4. Stain for 3–4 minutes with a mixture of equal parts by volume of solution B and 45% acetic acid.
5. Press the material firmly onto a second slide and filter the liquid using filter paper.
6. Wrap the two slides coated with the material tightly in blotting paper soaked in 95% alcohol, place them in a Petri dish, and cover.
7. After 5 hours, remove the blotting paper and soak the slides in alcohol until they separate easily. The material will now adhere firmly to the coated protein glycerin.
8. Allow to drain and then enclose in Euparal.

Staining result:
   Chromosomes appear purple to brownish-red; cytoplasm and nuclei are barely stained.
Ref.: B. P. Kaufmann, cited in Conn, Darrow, and Emmel, Staining Procedures, 2nd ed.

 
Required solutions:
   A. 4% iron alum solution, freshly prepared
   B. 0.5% aqueous haematoxylin solution
   C. 1.2% aqueous picric acid solution
Materials used: Root tips, sections.

Staining procedure:

1. Fix in Flemming's chromosmic acid, embed in paraffin. Transfer the deparaffinized sections from water to A for 30 minutes.
2. Wash in running water for 10 minutes.
3. Stain for 30–60 minutes in B.
4. Wash for 10 minutes in running tap water.
5. Differentiate in C, check under a microscope.
6. Wash for 20 minutes in running tap water.
7. Dehydrate in the ascending alcohol series, adding one drop of ammonia to either the 70% or 80% alcohol solution.
8. Pass through xylene and encapsulate in balsam or malinol. Counterstaining can be performed.

Staining result:
   Chromosomes are stained bluish-black and are well differentiated.
Ref.: Hutner, Stain Technology, 9, 57-59


Required solutions:
   A. 1% aqueous solution of safranin
   B. 1% aqueous solution of gentian violet
   C. 0.5% aqueous solution of orange G
Materials used: Root tips, e.g., of Vicia faba, delicate longitudinal sections.
Fixation is best performed in Flemming's chromosmium acetic acid.

Staining procedure:

1. Deparaffin the sections in the usual manner.
2. Stain for 1–3 hours in solution A.
3. Rinse in distilled water.
4. Transfer to solution B for approximately 2–5 minutes.
5. Rinse with distilled water.
6. Add dropwise solution C.
7. After approximately 1–5 minutes, rinse with water.
8. Dehydrate in 96% alcohol.
9. When the last traces of staining dissipate, quickly cover the slide with clove oil or terpineol and check under a microscope whether the staining was successful.
10. Pass through xylene and encapsulate in balsam or malinol.

Staining result:
   If the staining is successful, the chromosomes in dividing cells will be bright red, the nucleolus red with a usually bluish tinge. The spindle fibers will             appear blue, and the cytoplasm brownish-yellow.
Ref.: Flemming, W., 1891, Arch. mikr. Anat, 37, 249–298
Required solution:
   Carmine-acetic acid according to Schneider.
  Material used: Root sections.

Staining procedure:

1. Cut 0.5 cm long root sections between the pith into thin longitudinal sections.
2. Cover the sections on the slide with carmine-acetic acid.
3. Place a large coverslip on top and heat the preparations over a low flame for about 10 seconds until the acetic acid boils.
4. After cooling, gently squeeze the sections if necessary and examine them under the microscope.
5. Not suitable for permanent slides.

Staining result:
   The nuclei and nuclear divisions appear clearly stained red.
Required solution:
   Methyl green-acetic acid
   Material used: T       hin longitudinal sections of broad bean (Vicia faba).

Staining procedure:

1.   Pour methyl green acetic acid over sections on the slide and allow to act briefly.
2.    Rinse quickly with water to which a trace of acetic acid has been added.
3.    Absorb excess liquid with filter paper, place a coverslip on top, and examine under a microscope. Not suitable for permanent slides.

Staining result:
The dormant and dividing cell nuclei appear fixed and distinctly green within the cells.
Pianese stain
   

Necessary solutions:
A. Pianese-Farbgemisch		0,6 g
Alkohol		50 ccm
Wasser dest.                                                                                      erwärmen, abkühlen lassen, filtrieren B. Salzsäure-Alkohol 3 %		150 ccm

Staining procedure:

1. Place sections in 50% alcohol.
2. Stain with solution A for 30–60 minutes.
3. Wash with alcohol.
4. Differentiate with solution B for 30 seconds.
5. Wash with alcohol.
6. Lighten in clove oil, embed in balsam or Malinol.

Staining result:
   Fungal hyphae: pink
   Host tissue:      green
Ref.: Pianese, G. (1896), Beitr. Path. Anat. u. Allg. Path. Ergb. I, 193

 

 

B. Parasitology

 
                 a. Plant parasites
                 b. Animal parasites
                 c. Filterable infectious agents
 

a) Plant parasites
Bacterial staining according to Gram

 

Smear Preparations

Required solutions:

A. Carbolgentiana violet solution
B. Iodine-potassium iodide solution 1:2:300
C. Carbol fuchsin 1 part
     Distilled water 10 parts

Staining procedure:

1. After fixing, stain with solution A under gentle warming for 1-3 minutes.

2. Discard the staining solution (do not rinse).

3. Transfer to solution B for 1-3 minutes.

4. Discard the staining solution.

5. Decolorize with 96% alcohol until no more color scum is released, approximately 15 seconds.

6. Re-stain with solution C for 5-30 seconds.

7. Rinse briefly with water, dry.

8. Embrace in Canada balsam or Malinol.

Staining result:
   Gram-positive bacteria:   blackish-blue
   Gram-negative bacteria: red

 
Required Solutions:
   A. Carbolgentian Violet Solution
   B. Iodine-Potassium Iodine Solution 1:2:300
   C. Bismarck Brown Solution

Staining Procedure:

1. Adhere the sections and de-paraffin.
2. Stain in Solution A for 2–5 minutes.
3. Pour off the staining solution (do not rinse).
4. Transfer to Solution B for 1–2 minutes.
5. Decolorize with 90% alcohol until no more color runs.
6. Rinse with water.
7. Re-stain with Solution C.
8. Rinse with 60% alcohol.
9. Dehydrate with absolute alcohol.
10. Lighten in xylene and embed in balsam or malinol.

Coloring result:
Gram-positive bacteria: black-blue Gram-negative bacteria: brown
Gram positive are: 
            Bacillus diphtheriac                Bacillus anthracis 
            Bacillus tetani                         Mycobacterium leprae 
            Bacillus tuberculosis               Micrococcus tetragenus

Pneumococcus Streptococcus           Staphylococcus Actinomyces
Bacillus rhusiopathiae suis                 Bacillus of mouse septicemia
Bacterium rhinoscleromatis                Bacillus smegmatis
Yeast cells
 

Gram-negative words are:
Bacterium tussis convulsivae	Bacterium typhi
Bacterium influencae	Bacterium paratyphi
Bacterium pneumoniae Friedländer	Bacterium pestis
Bacterium pyocyaneum	Bacterium mallei
Bacillus oedematis maligni	Bacterium coli
Bacterium aegyptiacum	Bacterium cholerae
Bacterium abortus infectiosi	Bacterium necrophorum
Micrococcus gonorrhoeae	Micrococcus melitensis
Micrococcus intracellularis Spirochaeta pallida	Spirochaeta Obermeier

 
Bonhoff staining (sections)

Required solutions:

A. Concentrated alcoholic methylene blue solution                      4 parts
     Carbol fuchsin                                                                        15 parts
     Distilled water                                                                         20 parts
     The mixture must always be freshly prepared.
B. 1% aqueous acetic acid
C. Aniline-xylene

Staining procedure:

1. Transfer sections from absolute alcohol to microscope slides, rinse, and fix.
2. Stain in solution A for 2 minutes (heat once over a flame until steaming occurs).
3. Discharge the staining solution and rinse with water.
4. Differentiate with solution B.
5. Rinse in water.
6. Dehydrate with solution C (changing several times) for 3–5 minutes.
7. Transfer to xylene and embed in balsam or malinol.

Staining result:
   Bacteria: light blue
   Tissue and nuclei: red
Required solutions:
   A. Bunge mordant
   B. Carbol-gentian violet

Staining procedure:

1. Mix a small amount of pure culture with an equal amount of water and spread onto a completely grease-free slide.
2. Dry by passing the slide three times over a flame.
3. Drip the mordant through a paper filter.
4. Apply the mordant for 1 minute while gently warming.
5. Rinse with water.
6. Re-stain with solution B for 3-5 minutes while gently warming.
7. Rinse with water.
8. Dry, then encapsulate in balsam or Malinol.

Staining result:
   Flags: blackish-brown
   Bacteria: violet Smear Preparations


Required Solution:
A. Carbolfuchsin                                              1 part 
     Distilled Water                                          10 parts

Staining Procedure:

1. Fix smears.
2. Stain in solution A for 5–10 minutes.
3. Rinse in water.
4. Dry, then encase in balsam or Malinol.

Staining result:
   Bacteria: red

Required solutions:
   A. Löffler's methylene blue solution
   B. 0.25–0.5% aqueous acetic acid

Staining procedure:

1. Stain in solution A for 5–30 minutes.
2. Rinse in water.
3. Differentiate in solution B.
4. Transfer to 90% alcohol for 2–5 minutes.
5. Lighten in xylene, embed in balsam or malinol.

Staining result:
   Bacteria: blue


Frosch stain (smear preparations)
 

Required solutions: A. Fuchsin 1 %ig wässrig, schwach alkalisch
B. Patentblau konz. wässrig	3 Tropfen
Essigsäure konz. (Eisessig)	1 Tropfen
Wasser dest.	20 ccm
C. Wasser dest.	30 ccm
Essigsäure konz. (Eisessig)	1 Tropfen

 
Staining procedure:
Fix in absolute alcohol.

Stain in solution A for 10–30 seconds.

Counterstain and differentiate in solution B until the preparation (in the thinner parts of uneven smears) has taken on a greenish-blue hue.

Rinse in solution C.
Dry with blotting paper and encase in balsam or Malinol.

Staining result:
Germ bacilli: bright red
Erythrocytes: green
Note: Staining fails with old, decomposed material and very old smears.
Hansen stain (smear preparations)

Required solutions:
   A. Löffler's methylene blue
   B. Safranin 3% aqueous

Staining procedure:

1. Heat fixation.
2. Stain in solution A, 1 minute.
3. Rinse in water.
4. Re-stain in solution B, 15–20 seconds.
5. Rinse in water.
6. Dry, encase in balsam or malinol.

Staining result:
   Bang bacteria:                                      blue
   Accompanying bacteria and substrate: red

Section preparations
   Required solution: A. Carbolthionine

Staining procedure:

1. Stain the sections (paraffin sections) in solution A, 2–5 minutes.
2. Rinse in water.
3. Dehydrate in absolute alcohol.
4. Lighten in xylene, embed in balsam or malinol.

Staining result:
   Bacteria: blue-violet
   Nutrients: light blue
Claudius stain (smears)

Required solutions:

1. Methyl violet, concentrated aqueous
2. Picric acid, 0.5% aqueous
3. Clove oil

 

1. Safranin in aniline water (analogous to aniline water gentian violet)
2. Concentrated safranin Alcoholic
3. Fluorescein saturated in clove oil
4. Methyl green saturated aqueous
5. Staining procedure:
6. Fix in Müller's solution
7. Post-cure in alcohol for a few minutes
8. Stain in solution A for 5 minutes
9. Dehydrate in solution B
10. Transfer to solution C
11. Embrace in clove oil, then in alcohol
12. Re-stain with solution D
13. Dehydrate with alcohol, lighten in xylene, embed in balsam or malinol

  Oelt's stain (smears)

Required solutions:

1. Safranin 2% aqueous
2. Acetic acid 2%
3. Staining procedure:
4. Stain the air-dried smears in solution A while boiling.
5. Rinse with water.
6. Differentiate with solution B, 10 seconds.
7. Rinse with water.
8. Dry, encase in balsam or Malinol.

Staining result:
    Bacillus body: red,
    capsule: yellow

Azure eosin stain (smears)

Required solution:
    Azure eosin solution


Staining procedure:

1. Add approximately 2 drops of solution A to air-dried, fiber-fixed smears.
2. After 30 seconds, allow the staining solution to run off.
3. Add 20 drops of distilled water and allow to act for 1–7 minutes.
4. Dry between sheets of blotting paper.
5. Embed in balsam or Malinol.

Staining result:
     Bacilli: blue
     Capsules: reddish-violet to pink

Note: This staining method also makes disintegrated anthrax bacilli and empty capsules, such as those found in anthrax shadows in decomposing material, visible. The staining times must be strictly adhered to, as excessively long staining will result in an overly intense blue color, making the capsules difficult to see.

Kühne-Weigert staining (sections)

Required solutions:
   A. Lithium carmine solution
   B. Hydrochloric acid-alcohol solution
   C. Crystal violet, concentrated aqueous solution     100 ml
        Hydrochloric acid                                                   10 drops
   D. Lugol's solution


Staining procedure:

1. Stain sections in solution A for 3 minutes.
2. Briefly rinse in solution B.
3. Rinse in water.
4. Stain in solution C for 5–10 minutes.
5. Transfer to solution D for 1–2 minutes.
6. Carefully dab off the staining solution.
7. Decolorize in aniline oil until no more violet flecks appear and the section shows carmine color again.
8. Lighten in xylene, embed in balsam or malinol.

Staining result:
      Bacilli: deep blue-violet
      Tissue: red

Capsule staining according to Klett (depiction of anthrax bacillus capsules)

Necessary solutions:

1. Aqueous-alcoholic methylene blue solution.
2. Aqueous-alcoholic fuchsin solution.

Staining procedure:

1. Pass air-dried smears through a flame.
2. Stain in solution A, heating until boiling.
3. Rinse thoroughly with water.
4. Re-stain with solution B, 5 seconds.
5. Rinse with water.
6. Dry, encase in balsam or Malinol.

Staining result:
     Inner parts of the bacteria: blue
     Cover: light rose red
     Contours: dark red

Capsule staining according to Räbiger (representation of anthrax bacillus capsules)

Necessary solution:

1. Formalin conc. Pure 100 cc
2. Gentian Violet 15 g

Mix well and filter after 12 hours.


Staining procedure:

1. Stain unfixed preparations in solution A for 20 seconds.
2. Rinse with water.
3. Dry, then encase in balsam or Malinol.

Staining result:
      Capsules: reddish violet
      Bacilli: deep blue


Neisser Staining (Smears)

Required Solutions:

1. A. Neisser I and II
2. (Mix 1:1 before use.)
3. B. Neisser III

Staining Procedure:

1. Stain heat-fixed smears in solution A for 10–20 seconds.
2. Rinse briefly in water.
3. Counterstain with solution B for 10 seconds.
4. Rinse with water and dry with blotting paper.
5. Embal in balsam or Malinol.

Staining Result:
     Bacillus body: brown
     Polar granules: dark blue-black
Note: Pseudodiphtheria bacilli react negatively to this stain. A clearer visualization of the polar granules is obtained by treating the smears with Lugol's solution containing 1% lactic acid for 3–5 seconds before the first rinse.

Stoltenberg Staining (Smear Preparations)

Required Solution:
      A. Stoltenberg Stain

Staining Procedure:

1. Stain the fixed smears in solution A for 1 minute.
2. Rinse with water, dry.
3. Embal in balsam or Malinol.
4. Staining Result:
5. Bacillus body: green
6. Polar granules: red
7. Ljubinsky Staining

Required Solutions:
A.  Methyl Violet 2B                 0.25 g 
     Distilled water                      95 cc
     Glaceous acetic acid             5 cc
B. Bismarck brown                    0.1 g
    Distilled water                     100 cc

Staining procedure:

1. Fix with gentle heat.
2. Stain in solution A, 30–20 seconds.
3. Wash in water.
4. Stain with solution B, 30 seconds.
5. Wash in water.
6. Dry (on blotting paper).

Staining result:
     Polar particles: bluish-black
     Bacteria: yellowish-brown

Ref.: Blumenthal, J. M. & Lipskerow, M. (1905), Centbl. Bact. I Abtl., Orig. 38, 359–366

Gram staining (sections)

Required solutions:
A. Carbolgentiana violet, Gram
B. Lugol's solution
C. Carbol fuchsin
(Dilute 1:10 with water before use.)

Staining procedure:

1. Fix in Müller's solution.
2. Stain paraffin sections in solution A for 5–20 minutes.
3. Rinse in water.
4. Mordant in solution B for 1–2 minutes.
5. Differentiate in 90% alcohol until the section appears nearly colorless.
6. Rinse briefly with water.
7. Re-stain with solution C.
8. Transfer in 96% alcohol and absolute.
9. Lighten in xylene, embed in balsam or malinol.

Staining result:
Diphtheria bacilli: bluish-black
Background: light red

Küster fluorescence method (smear preparations)

Required solution:
A. Auramine O               0.1 g
    Distilled water           100 cc
    Liquid carbolic acid       5 cc

Staining procedure:

1. Stain the fixed smears with solution A for 10 seconds.
2. Rinse vigorously in very soft tap water or a dilute ammonia solution for approximately 10 seconds.
3. Air dry or dry between lint-free blotting paper.

Staining result:
       Diphtheria bacilli: blue-green
       Granules: golden yellow

If pseudodiphtheria bacilli are present, any granules will not appear bright golden yellow, but at most a faint greenish-yellow.
Feedback geben

 
Ziehl-Neelsen stain (smears)

Required solutions:
A.   Carbol fuchsin, Ziehl-Neelsen
B.   Hydrochloric acid-alcohol
C.   Löffler's methylene blue

Staining procedure:

1. Fix the sputum smears with heat.
2. Stain with solution A for 3 minutes, warming until bubbles form.
3. Wash in water.
4. Decolorize with solution B until only a pink hue remains.
5. Wash in water.
6. Counterstain with solution C.
7. Wash in water.
8. Dry, then encase in balsam or Malinol.

Staining result:
       Acid-fast bacteria: red.
      Other organisms:   blue.
      Background:          faint blue or unstained
Lit.: Ziehl, F. (1882), Deut. Med. Woch., 8: 451.
Neelsen, F. (1883), Centbl. Med. Wis., 21: 497–501

Mühlpfordt staining (smears)

Required solutions:

A. Victoria Blue 4 R 3% aqueous solution
B. Hydrochloric acid-alcohol solution
C. Pyronin G 1% aqueous solution

Staining procedure:

1. Drop solution A onto the air-dried specimen and heat once until bubbles form.
2. Rinse with water.
3. Differentiate with solution B for 5 minutes.
4. Rinse with water and dry with blotting paper.
5. Counterstain with solution C for 10–20 seconds.
6. Dry and encase in balsam or Malinol.

Staining result:
     Tubercle bacilli:                    deep blue
     Substance:                           crimson red
     All accompanying bacteria:  red

Schaedel stain for red-blue colorblind individuals (smear preparations)

Required solutions:

   A. Methyl Violet BB                 1 g
       Absolute alcohol                10 ml
      Carbol water 2%                 90 cc
B. Hydrochloric acid-alcohol
C. Bismarck brown, conc. aqueous

Staining procedure:

1. Stain fixed smears with solution A, boiling three times.
2. Rinse thoroughly with water.
3. Decolorize with solution B until the preparation appears gray.
4. Rinse with water.
5. Re-stain with solution C for 1–2 minutes.
6. Rinse with water.
7. Dry, then encase in balsam or Malinol.

Staining result:
     Bacilli:        blackish-violet
     Substrate:  light brown

 
Koch-Ehrlich staining (sections)

Required solutions:
A. Carbol fuchsin
B. Hydrochloric acid-alcohol
C. Concentrated aqueous methylene blue solution
D. Acetic acid 0.25%
Staining Procedure:

1. Fix in Müller's solution or an ascending alcohol series.
2. Stain with solution A at 37°C for 30 minutes or 12 hours at room temperature.
3. Decolorize with solution B for 10–15 seconds.
4. Wash with 70% alcohol until no more color runs and the preparation appears colorless.
5. Re-stain with solution C for 5 minutes.
6. Rinse in solution D, then in alcohol.
7. Lighten in clove oil, embed in balsam or Malinol.

Staining Result:
        Tubercle bacilli:                   red
        Tissue and other bacteria: blue

Haitinger and Schwertner Fluorescence Method

(Smear Preparations)

Required Solutions:
A.  Acridine Yellow 0.2% Aqueous
B.  Hydrochloric Acid-Alcohol

Staining Procedure:

1. Stain heat-fixed smears in solution A for 30 seconds.
2. Rinse with water.
3. Differentiate with solution B for approximately 10–15 seconds until the preparation appears colorless in daylight.
4. Rinse with water.
5. Air dry or dry with lint-free blotting paper.

Staining result:
     Tubercle bacilli:     golden yellow
     Substance:            dark blue-black
In very thick areas of the smear, the cells of the substance appear faintly greenish.

Fluorescence method according to Hagemann-Herrmann

(Smear preparations)

Required solutions:
 A.   Auramine O                       0.1 g
       Distilled water                   100 cc
       Liquid carbolic acid              5 cc
B. Hydrochloric acid-alcohol
C. Potassium pyrrmanganate solution 1:1000
D. Löffler's methylene blue

Staining procedure:

1. Pour solution A over heat-fixed slides.
2. Heat until bubbles burst, then allow to act for 5 minutes.
3. Rinse in running water.
4. Differentiate with solution B until the slide appears colorless, approximately 15–20 seconds.
5. Rinse in running water.
6. Immerse in solution C for exactly 5 seconds.
7. Rinse in running water.
8. Immerse in solution D for 1 second.
9. Rinse with water and dry between lint-free blotting paper.

Staining result:
     Tubercle bacilli:                                            bright yellow
     Non-acid-resistant material, cells, mucus:   dark brown shadow.
Floodstaining method according to Bachmann and Finke (sections)
 

Necessary solutions:
A. Auramine O		0,1 g
Wasser dest.		100 ccm
Liquid carbolic acid		5 ccm
B. Denatured alcohol (ethyl alcohol)		100 ccm
hydrochloric acid		0,4 ccm
Sodium chloride C. Löffler's Methylene Blue		0,4 g

Staining procedure:

1. Rinse dewaxed sections thoroughly with water.
2. Stain with solution A for 15 minutes.
3. Rinse with water.
4. Differentiate with solution B for 3 minutes. (Renew the solution after 90 seconds.)
5. Rinse with water.
6. Stain with solution C for 30 seconds.
7. Rinse with water.
8. Dry carefully with lint-free blotting paper.
9. For permanent slides: Embed in fluorescent-free cedarwood oil and store in the dark.

Staining result:
     Tubercle bacilli: bright yellow
Fluorescence method according to Schallock (sections)

Necessary solutions:
A. Auramine O	0,1 g
distilled water                                                                  B. Hydrochloric acid alcohol	100 ccm
C. Thiazole yellow	0,1 g
distilled water	100 ccm

 
Staining procedure:

1. Transfer dewaxed sections from water to solution A at 37°C for 15 minutes.
2. Differentiate with solution B at 37°C.
3. Rinse in water.
4. Counterstain with solution C for 2 seconds.
5. Rinse in 50% alcohol.
6. Rinse in water.
7. Cover with glycerin, rim the coverslip with wax.

Staining result:
       Tubercle bacilli: bright yellow
       Tissue:              pale blue
Note: Material fixed in formalin for more than 4-5 days is no longer suitable for staining.
 
Required Solutions:
A. Aniline Water-Gentian Violet
B. Nitric Acid-Alcohol 10%
C. Fuchsin, Conc. Aqueous
D. Hydrochloric Acid-Alcohol 3%
E. Malachite Green                                1 g
    Distilled Water                               100 ml

Staining Procedure:

1. Fix with heat.
2. Stain with solution A for 5 minutes.
3. Decolorize with solution B.
4. Rinse with water.
5. Add solution C.
6. Heat until bubbles rupture.
7. Rinse with water. Decolorize with solution D.
8. Rinse with water.
9. Counterstain with solution E, 10–15 seconds.
10. Rinse with water, dry, and encase in neutral balsam or Malinol.

Staining result:
         Leprosy bacilli:                            deep violet
         Tubercle bacilli:                           ruby ​​red
          Cell fragments and other bacilli: green
Lit.: Cardelus-Dalfó, C. (1959), Laboratorio (Granada) 27, 531–533.
Ref.: Z. wiss. Mikrosk., Vol. 64, No. 6: 380 (1960).
Rüdel stain (smear preparations)

Necessary solutions:
A. Carbolfuchsin                    1 part
    Distilled water                    1 part
B. Sulfuric acid 2.5%
C. Löffler's methylene blue    1 part
    Distilled water                 2-3 parts

Staining procedure:

1. Stain the fixed smears with solution A while heating until bubbles burst. Rinse several times with solution B.
2. Rinse with water.
3. Counterstain with solution C for a few seconds to a maximum of 1 minute.
4. Rinse with water.
5. Dry, encase in balsam or Malinol.

Staining result:
     Leprosy bacilli: red
     Background: blue
     Tubercle bacilli: unstained
     Unna stain (sections)

Required solutions:
A. Thymene Victoria blue
B. 25% saline solution
C. 1% aqueous pyronin G

Staining procedure:

1. Stain celloidin sections in solution A for 12 hours.
2. Rinse in water.
3. Transfer to solution B for 2 minutes.
4. Rinse in water.
5. Immerse in absolute alcohol until decolorized.
6. Rinse in water. Counterstain with solution C, 5 minutes.
7. Rinse in water.
8. Dehydrate in alcohol.
9. Lighten in bergamot oil, embed in balsam or malinol.

Staining result:
    Bacilli:                       dark blue
    Protoplasm, especially granoplasm and nuclei, keratohyalin, and
    stratum corneum:      dark red
 
Schlirf stain (smears)

Required solutions:
A. Schlirf solution A
B. Lugol's solution
C. Schlirf solution B

Staining procedure:

1. Stain the fixed smears in solution A, 1 minute.
2. Rinse with water, dry between blotting paper.
3. Mordant with solution B, 1 minute.
4. Differentiate in alcohol by first rinsing solution B from the specimen and then adding more alcohol dropwise while swirling until no more color bleeds out, approximately 30 seconds.
5. Re-stain with solution C, 2 minutes.
6. Rinse in water.
7. Dry, encase in balsam or Malinol.

Staining result:
     Gonococci:                   deep pink
     Gram-negative flora:    faintly stained

Unna-Pappenheim staining (smears)

Required solution:
A. Methyl green pyronine, Unna-Pappenheim

Staining procedure:

1. Fix air-dried smears in methyl alcohol.
2. Stain in solution A for 3–5 minutes.
3. Rinse with water.
4. Dry, then encase in balsam or Malinol.

Staining result:
Gonococci:        red
Nutrients:          blue-green
Staining according to Jensen (smear preparations)

Required solutions:
A. Methyl Violet 6 B 0.5% aqueous
B. Lugol's iodine potassium iodide solution
C. Neutral Red                               0.5 g
     Distilled water                          500 ml
     1% Acetic Acid                             1 ml

Staining Procedure:

1. Pass air-dried specimens once through a flame (smear facing upwards), allow to cool.
2. Stain with solution A, 15–30 seconds.
3. Rinse with solution B.
4. Rinse with alcohol.
5. Add fresh absolute alcohol, pouring from the edge of the slide outside the smear area, and repeat until decolorization is complete.
6. Re-stain with solution C, 15–30 seconds.
7. Rinse with water.
8. Dry, then encase in balsam or Malinol.

Staining result:
     Gonococci:                       bright red
     Other cocci and bacteria: dark violet
Staining of choice (smears)

Required solution:
A. Wahl's solution

Staining procedure:

1. Stain the fixed smears in solution A for 5–15 seconds
2. Rinse briefly in water
3. Dry, encase in balsam or Malinol

Staining result:
Gonococci:                          reddish violet to black
Leukocyte nuclei:                pale bluish-green to light green
Plasma:                               light yellow or colorless, light green in thickened areas
Epithelia:                             yellowish-green


Unna-Pappenheim staining (sections)

Required solution:
A. Carbol-methyl green-pyronine solution

Staining procedure:

1. Stain the unmounted sections with solution A for 5–10 minutes in test tubes placed in water at 30–40°C.
2. Cool rapidly by placing in cold water.
3. Wash in water until the initially greenish water turns bluish-red.
4. Mount the sections.
5. Dry lightly with blotting paper.
6. Immerse in alcohol until no more red color is released.
7. Lighten in xylene for 30 seconds, then embed in balsam or malinol.

Staining result:
     Gonococci:            bright red
     Nuclei:                   pale green
     Protoplasm:           pale pink

Fluorescence method according to Haitinger and Schwertner (smear preparations)

Required solution:
A. Acridine Yellow 0.2% aqueous

Staining procedure:

1. Stain heat-fixed preparations in solution A for 5 minutes.
2. Rinse with water, dry.

Staining result:
    Nuclei of the pus bodies:     greenish-yellow
    Plasma of the pus bodies:   dark green
    Gonococci:                           intense golden yellow
    Background:                         dark blue
 
Required solutions:

1. Aniline water gentian violet
2. Safranin in aniline water, analogous to aniline water gentian violet
3. Potassium iodide solution, Grams
4. Staining procedure:
5. Stain the fixed smears in solution A for 5 minutes.
6. Transfer to solution B for 10 hours
7. Incubate in solution C for 1 minute.
8. Differentiate in absolute alcohol.
9. Dry and encase in balsam or Malinol.

Staining result:
      Threads:             blue-violet
      Knocking ends:   yellow-red

Staining according to Weigert (sections)

Necessary solutions:
A. Bismarck brown solution
B. Aniline aqueous gentian violet
C. Iodine-iodine solution, Gram
D. Aniline
E. Acid fuchsin 0.2% aqueous

Staining procedure:

1. Pre-stain the paraffin sections in solution A.
2. Rinse briefly in water.
3. Stain in solution B for 5 minutes.
4. Rinse in water and dry with blotting paper.
5. Transfer to solution C for 1–3 minutes.
6. Dry with blotting paper.
7. Decolorize with solution D until no more color is released.
8. Immerse in alcohol for 2–5 minutes. Counterstain with solution E, 3 minutes.
9. Rinse in water, 2 minutes.
10. Dehydrate in alcohol.
11. Lighten in xylene, embed in balsam or malinol.

Staining result:
    Mycelium: dark blue
    Flask: fuchsin red
    Nuclei: brown
    Wirtz, Spore Staining

Necessary solutions:
A.  Malachite Green          5 g
     Distilled water             95 cc
B. Safranin O                  0.5 g
    Distilled water            100 cc

Staining procedure:

1. Fix the smears with heat.
2. Stain with solution A, 30–60 seconds.
3. Heat until bubbles form.
4. Wash off excess stain under running water, 30 seconds.
5. Stain in solution B, 30 seconds.
6. Wash under running water.
7. Dry (on blotting paper).

Staining result:
    Spores: green 
    Cells:    red
Lit.: Wirtz, R. (1908), Centbl. Bakt., I. Abt., Orig., 46, 727-728  

Necessary solutions:
A. Malachite Green	5,0  g
distilled water	95 ccm
B. Eosin, water-soluble, yellowish	2,5  g
distilled water	100 ccm

Dyeing instructions:

1. Heat fixation.
2. Cover with solution A, boil for 20 seconds, then allow to stand for another 30 seconds.
3. Rinse in water.
4. Counter-stain with solution B for 1 minute.
5. Rinse with water, dry, and encase in neutral balsam or Malinol.

Staining result:
    Spores:              green
    Bacterial body:  pinkish-red

Spore staining according to Möller (demonstrating tetanus bacillus spores)

Required solutions:
A. 5% chromic acid solution
B. Carbol fuchsin
C. 5% sulfuric acid
D. Aqueous methylene blue solution

Staining procedure:

1. Fixed smears are first immersed in chloroform for 2 minutes to remove any fat droplets that could mimic spores after staining.
2. Rinse with water.
3. Immerse in solution A for 2 minutes.
4. Rinse with water.
5. Stain with solution B for 1 minute.
6. Differentiate in solution C.
7. Rinse with water.
8. Counterstain with solution D for 30 seconds.
9. Rinse with distilled water.
10. Dry, then encapsulate in balsam or Malinol.

Staining result:
     Spores:   red
     Bacilli:     blue
 
Keil staining (smears)

Required solution:
A. Astraviolet 2% aqueous

Staining procedure:

1. Cover air-dried smears on carefully cleaned slides with solution A for 3–5 minutes.
2. Gently warm.
3. Rinse with water, dry, and encase in balsam or Malinol.

Staining result:
      Pallida spirochetes: reddish-violet
Note: By avoiding any irritation, such as fixation, the characteristic shape of Spirochaeta pallida is preserved as much as possible.

Becker staining (smears)
 

Necessary solutions:
A. Ruge's solution: Formalin	20 ccm
Glacial acetic acid	1 ccm
Water	100 ccm
B. Tannin	10 g
Carbolic acid	1 g
Water                                                                                 C. Carbolfuchsin, Ziehl-Neelsen	100 ccm

 
Staining procedure:

1. Fix thin, air-dried smears of irritant serum in solution A, changing the solution 2-3 times for 1 minute each time.
2. Rinse in water for 10 seconds.
3. Mordant in solution B while warming (not exceeding 50°C) for 30 seconds.
4. Rinse in water for 30 seconds.
5. Stain in solution C while gently warming for 30-45 seconds.
6. Rinse with water.
7. Dry and encapsulate in neutral balsam or Malinol.

Staining result:
     Spirochaeta pallida and recurrent spirochetes:  intense red.
Note: The steep whorls of the pallida spirochetes are well preserved.


Burri Negative Stain

Required Solution:
    A. Burri Ink

Staining Procedure:

1. On a microscope slide, mix one drop of irritant serum or one drop of fluid scraped from the cut surface of an organ with one drop of distilled water and one drop of Burri ink.
2. Spread the mixture as evenly as possible in a thin layer using the edge of a microscope slide or coverslip.
3. Allow to air dry for approximately 30–60 seconds.
4. Examine by oil immersion.

Staining Result:
     The spirochetes stand out unstained against the uniformly brown-stained background.

 
Required Solution:
A. Cyanochine Eisenberg

Staining Procedure:

1. Rub one drop of specimen with one drop of solution A as evenly as possible on a microscope slide and spread in a thin layer.
2. Allow to air dry.
3. Examine by oil immersion.

Staining Result:
     The spirochetes stand out as sharply defined, colorless or at most faintly pink-stained spirals against the blue-violet background.


 
Manson stain (smears)

Required solution:
A. Borax-Methylene Blue, Manson
(Dilute 1:40 with water before use.)

Staining procedure:

1. Fix the air-dried, preferably thin, smears with methyl alcohol for 5 minutes.
2. Stain with solution A for 10–15 seconds.
3. Rinse with water until the preparation has a greenish-blue hue.
4. Dry with blotting paper.
5. Examine immediately by oil immersion or encapsulate in neutral balsam or Malinol.

Staining result:
     Erythrocytes:                     greenish
     Leukocytes and parasites: intense blue

Staining with azure eosin (smears)

Required solution:
A. Giemsa azure eosin solution

Staining procedure:

1. Fix air-dried, very thin smears in methyl alcohol for 2–3 minutes or absolute alcohol for 15–20 minutes.
2. Blot with blotting paper.
3. Stain with solution A (10 drops to 10 cc of absolutely neutral water) for 30–40 minutes.
4. Rinse thoroughly with water.
5. Dry between blotting papers.
6. Encapsulate in neutral balsam or Malinol.

Staining result:
    Chromatin of the parasites and Schüffner's stippling: bright red
    Protoplasm of the parasites:                                        blue
    Leukocyte nuclei:                                                         violet-reddish-violet
   Erythrocytes:                                                                 pinkish-brownish-red
   Note: With older preparations, this method only works if the preparations have been stored absolutely dry.

To create the effects of masonry staining and Schüffner's stippling, it is advantageous to add a drop of 0.1% potassium carbonate solution to the water used to dilute the paint solution.


Potassium carbonate solution
... Staining of old dry smears according to Giemsa (smear preparations)

Necessary solutions:
A. Azure-eosin solution
B. Weise buffer solution
C. 1% aqueous sodium carbonate solution
D. 0.1% aqueous sodium phosphate (NaH₂PO₄ • 2 H₂O)
E. 0.1% aqueous eosin

Staining procedure:

1. Since the preparations are already sufficiently fixed by the prolonged exposure to calcium chloride, immediately
2. stain in diluted azure-eosin solution (10 drops of solution A to 10 cc of solution B and 1 drop of solution C) for 45 minutes.
3. Rinse vigorously with water.
4. Differentiate in solution D, to which 2–3 drops of solution E are added to every 10 cc before use. The exposure time is 1–4 minutes.
5. Differentiation is complete when the blue hue of the smear has changed to reddish-brown. A pure eosin color indicates overdifferentiation and should be avoided.
6. Rinse with water.
7. Dry, then encase in neutral balsam or Malinol.

Staining result:

1. Parasite chromatin: bright red
2. Parasite protoplasm: blue
3. Leukocyte nuclei: violet-reddish-violet
4. Erythrocytes: pinkish-brownish-red
5. Schilling stain (smear preparations)
6. Thick blood drop method for detecting sparse parasites.

Required solution:
A. Azure eosin solution

Staining procedure:

1. On a thoroughly cleaned slide, allow a thick drop of blood to air dry for 1–2 hours.
2. Pour solution A (diluted 10 drops to 10 cc of distilled water) over the unfixed slide.
3. In a horizontal position, yellow haemoglobin clouds will appear after 2-3 minutes.
4. Pour fresh, diluted solution A from one side, rinsing off the old solution. Stain for 30-60 minutes.
5. Rinse with water and dry by holding the slide upright (do not use blotting paper).
6. Examine by oil immersion.

Staining result:
The parasites are somewhat distorted due to the slow drying process, but the staining otherwise corresponds completely to the familiar appearance of azure-eosin staining.
Note that, due to their basophilic ground substance and azurophilic, pale reddish granules, platelets can mimic parasites to the untrained eye.


Azure-Eosin Staining (Sections)

Required Solutions:
A. Giemsa Azure-Eosin Solution
B. 0.5% Acetic Acid

Staining Procedure:

1. Fix in absolute alcohol, embed in paraffin.
2. Prepare very thin sections.
3. Stain with solution A (dilute 10 drops in 10 cc of distilled water) in a covered dish for 20–24 hours.
4. Rinse with water.
5. Differentiate with solution B until the section appears reddish.
6. Wash in water, dry between sheets of blotting paper.
7. Differentiate in absolute alcohol until the section appears pale blue.
8. Dry, lighten in xylene, encase in neutral balsam or Malinol.

Staining result:
      Malaria plasmodia:                       pale blue
      Chromatin of malaria plasmodia: bright red
      Erythrocytes:                                pink
Jäger staining (smear preparations)

Required solutions:
A. 1% sublimate solution                 100 ml
     Absolute alcohol                           50 ml
     Glaceous acetic acid                      3 drops
B. Potassium iodide solution
C. Acidic hemelaune
D. 0.1% aqueous eosin

Staining procedure:

1. Fix moist smears in solution A for 12 minutes.
2. Immerse in 70% alcohol to which solution B has been added until a pale yellow color is achieved, for 10–15 minutes.
3. Rinse with water.
4. Stain in solution C for 10 minutes.
5. Rinse with tap water until a blue hue appears.
6. Re-stain with solution D for 1–2 minutes.
7. Dehydrate in alcohol, transfer to xylene.
8. Embark in neutral balsam or Malinol.

Staining result:
Nuclei of amoebae:  red
Other cells:               blue

Staining according to Jäger (sections)

Required solutions:
A. Sublimation solution 1% 100 ml
     Alcohol absolute 50 ml
     Glaceous acetic acid 3 drops
B. Potassium iodide solution
C. Acidic hemelaune
D. 0.1% aqueous safranin

Staining procedure:

1. Fix the material in solution A as soon as possible after death for 1 hour or longer.
2. Rinse with water.
3. Treat with 70% alcohol to which sufficient solution B is added until a pale yellow tint appears.
4. Embed in paraffin, mount, and deparaffinize.
5. Stain with solution C for 10–15 minutes.
6. Rinse with water until a blue hue appears.
7. Re-stain with solution D for 2–3 minutes.
8. Dehydrate in alcohol, lighten in xylene.
9. Embed in neutral balsam or Malinol.

Staining result:
     Nuclei of the amoebae: saffron red
     Other cells:                    blue

 



Required solutions:
 A. Physiological saline solution
 B. Bender-Hettche brilliant cresyl blue solution

Staining procedure:

1. Trim one part trichomonad pus with three parts of solution A.
2. Place a loop of suspended pus with a diameter of 4–6 mm onto a coverslip and rub one drop of diluted solution B into it.
3. Examine the specimen with a medium-strength drying system.

Staining result:
      Leukocytes, epithelia, lymphocytes: blue
      Nuclei:                                               deep blue
      Trichomonads:                                   unstained
Gerlach stain (smear preparations)

Required solution:

A. Carbolfuchsin, Ziehl-Neelsen          3 cc
Löffler methylene blue                         6 cc
Distilled water                                    50 ml

Staining procedure:

1. Fix the air-dried smears in methyl alcohol.
2. Rinse with water.
3. Swipe off any remaining water (do not dry).
4. Add solution A dropwise and heat until steaming occurs.
5. Agitate the specimen in the warm staining solution.
6. Pour off the staining solution and repeat the staining process twice more with fresh staining solution.
7. Rinse with water.
8. Dry and cover with a coverslip.

Staining result:
    Ganglion cells:       blue
    Negric corpuscles: red with a blue internal structure
    Erythrocytes:          unstained or pale red


Mann-Bohne staining (sections)

Required solutions:
Mann eosin-methyl blue
Absolute alcohol: 30 ml
1% alcoholic sodium hydroxide solution: 5 drops
Distilled water 30 cc
Acetic acid 1 drop

Staining procedure:

1. Fix thin sections of ammonite in alcohol at 37°C for 30–40 minutes.
2. Embed in paraffin (rapid embedding).
3. Cut, mount, and remove the paraffin.
4. Stain in solution A for 30–240 seconds.
5. Briefly rinse in water.
6. Rinse in absolute alcohol.
7. Immerse in solution B for 15–20 seconds.
8. Briefly rinse in alcohol. Rinse in water, 1 minute. Immerse in solution C, 1–2 minutes.
9. Dehydrate in alcohol.
10. Lighten in xylene, embed in balsam or malinol.

Staining result:
       Negro bodies: red
       Tissue: blue
       
 
The causative agent itself is unknown. If a smallpox pustule is used to inoculate the cornea of ​​a rabbit with a fine incision, a whitish opacity becomes noticeable after two days. Microscopic examination reveals round structures in the corneal epithelium, known as Guarni corpuscles. These same structures are also found after inoculation of the cornea with vaccine lymph, but not after inoculation with the contents of the varicella vesicles. Guarni corpuscles are specific to variola. Paschen was able to demonstrate the presence of extremely small, round corpuscles, called Paschen elementary bodies, in the smallpox pustule.


Guarni corpuscles Paschen Staining (Smear Preparations)

Required Solutions:
 A. Löffler's Flagella Mash
 B. Carbol Fuchsin

Staining Procedure:

1. Scrape the variola pustule with a coverslip, collect the emerging tissue fluid with the edge of a coverslip under slight pressure, and spread it onto a microscope slide in the manner of a blood smear.
2. Allow to dry completely in air for approximately 12–24 hours.
3. Fix in absolute alcohol for 1–10 hours or in methyl alcohol for 5–10 minutes.
4. Mordant in solution A for 3 minutes while warming until vapors just rise.
5. Rinse in water.
6. Stain in solution B while warming for 2–4 minutes.
7. Rinse in water, dry, and encase in balsam or Malinol.

Staining Result:
      Elementary particles stained dark red on a light red background.
Note: The Paschen method is suitable for children's lymph, variola pustule contents, and smear preparations of infected rabbit corneas.

Staining according to Sorgenfrei (section preparations)

Required solutions:
  A. Flemming's chromosmium acetic acid.
  B. Ehrlich's hematoxylin solution, of which 1 part is diluted with 19 parts of 2% potassium alum solution for use.
  C. A mixture of equal parts safranin aniline solution and 1 g of safranin O dissolved in 100 ml of absolute alcohol and 50 ml of            water.
  D. Tannin-picric acid = 25 g tannin, 0.1 g picric acid, 100 ml distilled water.

Scrap a small piece of thickened corneal epithelium from the cornea desensitized with 5% cocaine solution.
Quickly tear apart the fragments in a drop of 1% acetic acid under a dissecting microscope or loupe until the smallest, thinnest epithelial clumps, composed of only 2-3 cell layers, are formed.

      3. Carefully absorb the acetic acid with blotting paper.
      4. Fix in solution A for 15 minutes.
      5. Wash in running water for 10 minutes.
      6. Stain in solution B for 15 minutes.
      7. Wash in water for 10-15 minutes.
      8. Re-stain in solution C for 30 seconds.
      9. Mordant for 10-15 minutes and differentiate in solution D.
    10. Rinse in alcohol until a change to violet occurs.
    11. Lighten in xylene, encapsulate in balsam or malinol.

Staining result:
    The Guarni corpuscles are stained blue-violet and stand out clearly against the saffron-red surroundings.

Heidenhain-van Gieson staining (sections)

Required solutions:
  A. Haematoxylin, Heidenhain I (= iron salt)
  B. Haematoxylin, Heidenhain II (= haematoxylin)
  C. van Gieson's solution

Staining procedure:

1. Fix corneal scrapings, prepared as above, in Flemming's chromosmium acetic acid (30202) for 15 minutes.
2. Rinse in water for 10–15 minutes.
3. Stain in solution B, diluted with an equal amount of distilled water, for 24 hours.
4. Rinse in water.
5. Differentiate in solution A under microscopic control until cellular substances appear bright.
6. Rinse in water.
7. Stain in solution C for 3–5 minutes.
8. Rinse briefly in water, approximately 30 seconds.
9. Dehydrate in alcohol, lighten in xylene.
10. Encase in balsam or Malinol.

Staining result:
Guarni corpuscles bluish-black, nuclei light brown, centrosomes reddish-brown, protoplasm yellow, connective tissue red, leukocytes dark brown.



 
The minute, bacteria-like structures found in the gut of body lice by Riketts and von Prowazeki are only found in body lice infected with typhus blood. Whether they are the causative agents of typhus is still undecided, but their detection is diagnostically significant.

Castaneda staining (smears)

Required solutions:
  A. Dissolve 23.86 g of disodium phosphate in 1 liter of distilled water
  B. Dissolve 11.34 g of monopotassium phosphate in 1 liter of distilled water.
  C. Buffer solution = 88 ml of A mixed with 12 ml of B.
  D. Löffler's methylene blue solution.
  E. 0.5% solution of safranin O in distilled water.

Fix air-dried smears from suspected body lice in methyl alcohol for 5 minutes.
Staining in a mixture of: 20 ml buffer solution C
1 ml 40% formalin
3 drops of D for 2–3 minutes.

  3. Rinse in water for 30 seconds.
  4. Counterstain in E for 1–2 minutes.
  5. Rinse briefly in distilled water.
  6. Dry and encapsulate in balsam or Malinol.

Staining result:
     Rikettsia blue, cells pale pink, nuclei stained red.

Giemsa staining (sections)

In sections, rickettsia are found particularly in the stomach of lice. Giemsa's azure-eosin solution is used for staining, following the procedure described for sections of malaria plasmodia. The rickettsia stain pale ruby ​​red.
 
The causative agent of psittacosis is not yet known. Detection in human samples from patients is usually very difficult. A common method is intraperitoneal inoculation of mice with an emulsion of infectious material. The mice are euthanized after approximately three weeks if they do not die spontaneously, as asymptomatic infections also occur. Abundant, stringy exudate is found between the intestinal loops in the abdomen and on the parietal and visceral peritoneum of infected animals; this exudate is used for examination. The so-called Levintha bodies, characteristic of psittacosis, are detected. They are always found within large, vacuolated peritoneal epithelial cells, sometimes enclosed in dense clumps and streaks within leukocytes, and sometimes freely outside the cells.

Castaneda Staining (Smears)

Required Solutions:
Use solutions A, B, C, D, and E as specified under Typhus.

Staining Procedure:

1. Fix air-dried exudate smears in methyl alcohol for 5 minutes.
2. Stain in a mixture of: 95 ml buffer solution C
    5 ml formalin 40%
   10 ml Löffler's methylene blue for 3–4 minutes.
3. Rinse briefly in distilled water for approximately 30 seconds.
4. Counterstain in safranin solution E for 2 seconds.
5. Rinse in distilled water.
6. Dry and encase in balsam or Malinol.

Staining result:
    Levinthal bodies appear as fine, pale blue, coccoid structures; the cells are stained pink.
Note: Due to the very high risk of infection, examinations for psittacosis may only be carried out with the utmost precautions and only at specially equipped institutes.
Ref.: Harald Mohs, Zbl. Bakter. I Orig., Vol. 136, No. 1/2 (1936).
 
Herzberg Stain (Smear Preparations)

Required Solution:
A. Herzberg Victoria Blue Solution
(Always filter fresh before use)

Staining Procedure:

1. For infectious disease investigations, pass virus smears through a flame three times; otherwise,
2. allow to air dry for 24 hours. Smear preparations should not be older than 10–15 days.
3. Stain in solution A for 3–10 minutes.
4. Rinse in distilled water with gentle agitation for 30 seconds.
5. Rinse again with distilled water.
6. Dry, then examine by oil immersion.

Staining Result:
    Elementary particles, depending on the species: light, dark, or blackish blue
Note: The staining power can be increased by adding 0.25 ml of saturated citric acid solution to 5 ml of freshly filtered staining solution.


Hagemann Fluorescence Method (Smear Preparations)

Required Solution:
A. Primulin                      0.1 g
    Distilled Water           100 cc
    Liquid carbolic acid       2 cc

Staining procedure:

1. Stain air-dried, preferably thin smears with solution A for 15 seconds.
2. Rinse with water and dry.

Staining result:
   Virus bodies: white to bluish-white fluorescent
   Substrate:     dark violet

Note: Highly infectious material for humans can be killed by fixation before staining without affecting the staining result. For this purpose, three passes of the air-dried smears through a flame are suitable, followed by treatment with 96% ethyl alcohol or 4% formalin for 5–10 minutes. However, the most beautiful stains are obtained when staining untreated specimens. Iron Haematoxylin-Duropicrofuchsin

Tissue Staining

Necessary Solutions:
A. Iron Haematoxylin Weigert A and B
     (mix 1:1 before use)
B. Duropicrofuchsin                               1.25 g
    Distilled Water                                    100 cc

Staining procedure:

1. Fix in Bouin or Helly, embed in paraffin.
2. Add sections to a tray, de-paraffin, and immerse in water using a descending series of alcohols.
3. Stain in solution A for 2 minutes.
4. Wash with running water for 10 minutes.
5. Stain with solution B for 3–5 minutes.
6. Rinse with water.
7. Dehydrate, lighten in xylene, and embed in balsam or Malinol.

Staining result:
    Nutrients:               dark brown
    Connective tissue: bright red
    Muscle tissue:       yellow
Ref.: Weigert, K. (1904), Ztschr. f. wiss. Mikr., 21 : 1


 

Necessary solutions:
A. Eisenhaematoxylin Weigert A and
    (mix 1:1 before use)
B. Acetic acid 1% aqueous
C. Acid fuchsin cloth fast yellow                                                        1.5 g
    distilled water90cc
    Glacial acetic acid                                                                           12 drops

Staining instructions:

1. Fix in formalin, Zenkcr, Helly or Bouin, embed, dewax, introduce into water by descending alcohol series.
2. Transfer and stain in solution A, 3 - 5 minutes.
3. Rinse in water.
4. Differentiate with solution B.
5. Wash in running water, 10 minutes.
6. Stain in solution C, 5 minutes.
7. Rinse with water.
8. Differentiate with solution B.
9. Wipe the bottom and free surface of the slide with filter paper.
10. Transferred into absolute alcohol, lightened into xylene, embedded into balsam or malinol.

Staining result: 
   Kernels:                    dark brown 
   Muscle tissue:           red 
   Connective tissue:     yellow
Lit.: Wallart, J. & Houette, Ch. (1934), Bull. Hist.appl., 12 B. 254 - 256.55


 


Necessary solutions:

    A. Aldehyde fuchsin working solution: = Aldehyde fuchsin stock solution

(0.75 g aldehyde fuchsin in 70% alcohol)	25 ccm
Alcohol 70%	75 ccm
Glacial acetic acid	1 ccm
B. Pontacylbiauschwarz SX	0,6 g
Potassium dichromate	0,4 g
distilled water	80 ccm
C. True Yellow	2,0 g
Alcohol 95 %ig	100 ccm

Staining procedure:

1. Fix in Bouin, Zenker, or formalin.
2. Dewax and soak in 70% alcohol.
3. Stain in solution A for 30 minutes.
4. Wash in 70% alcohol and transfer to water.
5. Stain in solution B for 15 minutes.
6. Wash in water.
7. Differentiate in 70% alcohol (until no further color separation occurs).
8. Stain in solution C for 5 minutes.
9. Wash in 95% alcohol.
10. Dehydrate, lighten in xylene, and embed in balsam or malinol.

Staining result:

Elastic fibers:	dark red
Collagen fibers:	bright yellow
Mucin:	reddish purple
Cores:	blue-green
Cytoplasm:	yellow to green

Lit.:   Green, J. A.; Wood, M. L. (1959), Stain Tech., 6 : 313

 
Necessary Solutions:
A. Kernechtrot Aluminum Sulfate 5.2 g
     Distilled Water 95 ml
B. Phosphorus Tungstenic Acid 5.0 g
    Distilled Water 95 cc
C. Aniline Blue-Orange Glacial Vinegar

Staining Procedure:

1. Fix in Carnoy, Bouin, or Held.
2. Stain the dewaxed sections in solution A for 30 minutes.
3. Rinse with water.
4. Mordant with solution B for 10–15 minutes.
5. Rinse with water.
6. Stain with solution C for 5–10 minutes.
7. Rinse with water.
8. Differentiate and dehydrate with isopropyl alcohol (microscopic inspection).
9. Lighten in xylene, embed in balsam or Malinol.

Staining result:
   Nuclei:                                        red
   Collagenous connective tissue: blue
   Erythrocytes:                              orange
   Glial fibers:                                 red
   Protoplasm, muscle:                  reddish-orange
Ref.: Geidies, H. (1954), Mikrokosmos 10: 239
This newly developed dye is comparable to the aforementioned nuclear red in its ease of application and lightfastness, but it stains the nuclei a more intense red, approximately the shade of the previously used azo carmine.

Required solutions:
A. True-to-nucleus ruby: Dissolve 0.1 g in 100 ml of distilled water without heating and add 1 ml of glacial acetic acid. The solution is stable             
     indefinitely.
B. Aniline blue-orange according to Halmi-Konecny ​​=
    0.1 g aniline blue
    0.3 g orange G
    0.5 g phosphotungstic acid
    1 ml glacial acetic acid
    100 ml distilled water, boil, cool, filter
    Formol-fixed material

Staining procedure:

1. Stain in solution A for 5 minutes.
2. Rinse in distilled water.
3. Immerse for 5 minutes in 5% phosphotungstic acid.
4. Rinse in distilled water.
5. Transfer to B for 8 minutes.
6. Rinse in distilled water.
7. Briefly differentiate in 96% alcohol, lighten in xylene, and embed in balsam or Malinol.

Staining result:
   Nuclei:                                        intensely red
   collagenous connective tissue:  blue
   erythrocytes:                               orange
   glial fibers:                                   red
   protoplasm and muscle stained: reddish-orange.
Note: Heidenhain Azan solution can be used instead of B with the same results.
Required Solutions:
A. Hemalaun, acidic, Mayer
B. Hydrochloric acid-alcohol
C. Erythrosine                                            1 g
    Distilled water                                    100 cc
    Formalin 40%                             a few drops
D. Safran du Gatinais                                2 g
     Water                                                100 cc
     Boil for 1 hour, allow to cool, add:
     Tannin 5% aqueous solution                1 cc
     Formaldehyde 40%                              1 cc

Staining instructions:

1. Fix, embed. Mount the sections, deparaffinize, and introduce them into water using a descending alcohol series.
2. Transfer to solution A and stain for 4–6 minutes. (Nuclei must be well stained; check under a microscope. Collagenous tissue should remain unstained; differentiate with solution B if necessary.)
3. Wash with water for 10 minutes. 5. Stain with solution C for 5 minutes.
4. Rinse with water.
5. Differentiate with 70% alcohol for a few seconds.
6. Wash with water.
7. Stain with solution D for 5 minutes.
8. Rinse quickly with water and dry (with blotting paper).
9. Dewater quickly with absolute alcohol.
10. Lighten in xylene, embed in balsam or malinol

Staining result:
    Nuclei:                                                  dark blue
    Cytoplasm:                                           reddish
    Muscle, nerve, elastic fibers:                red
    Collagenous tissue, cartilage, bone:    yellow
Ref.: Romeis, Mikr. Techn. (1948) § 711

Necessary solution:
A. Muchaematein, Mayer

Staining procedure:

1. Adhere the sections, deparaffinize, rinse with a descending series of alcohols, and immerse in water.
2. Stain with solution A for 5–10 minutes.
3. Absorb excess stain with filter paper.
4. Rinse in water for 3–5 minutes.
5. Wash thoroughly with 95% alcohol.
6. Dehydrate, lighten in xylene, and embed in balsam or malinol.

Staining result:
     Mucine:                            deep violet
     Nuclei:                              light blue
     Connective tissue:            light gray or unstained
Ref.: Laskey, A. M. (1950), Stain Tech., 25: 33
  

Necessary solutions:
A. Alcian Blue 8 GS	1 g
distilled water	100  ccm
Glacial acetic acid	10 Drops
chloroform                                                                                     B. Haemalaun, Mayer	a few drops
C. Eosine, yellow	0,1 g
distilled water	100 ccm

Staining procedure:
      1. Fix in picric acid solution
      2. Immerse the paraffin sections in water.
     3. Stain with solution A for 10–40 seconds.

Note: A longer staining time must be strictly avoided, as otherwise strong overstaining will occur.
      4. Wash in water.
      5. Stain in solution B for 5–10 minutes.
      6. Counterstain with solution C, if desired.
      7. Dehydrate, lighten in xylene, and embed in balsam or malinol.

Staining result:
     Mucin: blue-green
Ref.: Steedman, H. F. (1950), Quart. J. Microsc. Sc. 91:477
Required solution:
A. Mucicarmin conc., Mayer

Staining procedure:

1. Fix in alcohol and embed in paraffin or Celloidin. Mount sections and deparaffinize.
2. Stain in descending order of alcohol concentration in solution A for 10–15 minutes.
3. Wash with water.
4. Dehydrate with absolute alcohol.
5. Lighten in xylene, embed in balsam or malinol.
6. Note: If sections embedded in celloidin were used, lighten with terpino oil or oregano oil.

Staining result:
    Mucin: red
Ref.: Mayer, P. (1896), Mitt. Zool. Stat. Naples, 12, 303–330
 
Required Solutions:
A. Iron Haematoxylin, Weigert A and B (equal parts mixed immediately before use)
B. Metanil Yellow 0.25 g
     Acetic acid 0.25% 100 ml
C. Mucicarmine Solution

Staining Procedure:

1. Fix in formalin 1:10, embed in paraffin.
2. Adhere the sections, remove the paraffin.
3. Wash in a descending series of alcohols, then in water.
4. Stain in solution A for 1 minute. Wash in water. Stain in solution B, 30 seconds.
5. Wash in water.
6. Stain in solution C, 45 minutes.
7. Wash in water, dehydrate, embed in balsam or Malinol.

Staining result:
   Nutrients:               black
   Connective tissue: yellow
   Mucine:                  red
Ref.: Mayer, P. (1896), Mitt. Zool. Stat. Naples, 12, 303–330;
Masson, P. (1923), Traité de Path. Medicale (Maloine et Fils, Paris)
 


Required Solutions:
A. Iron Alum                                                                5 g
    Distilled Water                                                     100 cc
    Dissolve in the cold, add:
   Celestin Blue                                                          0.5 g
   Boil for 3 minutes, allow to cool, filter, pour into:
   Glycerin                                                                    7 cc
Note: Solution is stable for several months.
B. Haemalaun, acid, Majer
C. Picroorange 8.2% in 80% alcohol
D. Acid fuchsin                                                            1 g
    Trichloroacetic acid                                                 3 g
    Distilled water                                                      100 ml
E. Phosphorustungstic acid 1% aqueous
F. Aniline blue, water-soluble                                      2 g
   Acetic acid 2% aqueous                                       100 ml

Staining procedure:

1. Fix in formaldehyde sublimate, embed in paraffin.
2. Mount sections, immerse in 90% alcohol.
3. Remove the mercury precipitate.
4. Wash thoroughly in water. Stain in solution A, 3–7 minutes.
5. Wash in water.
6. Stain in solution B, 3–7 minutes.
7. Wash in water, at least 2 minutes.
8. Rinse in 95% alcohol.
9. Stain in solution C, 2 minutes.
10. Stain in solution D, 5 minutes.
11. Rinse with water.
12. Briefly immerse in a mixture of equal parts solution D and 80% alcohol.
13. Treat with solution E, 5–10 minutes.
14. Rinse with water.
15. Stain with solution F, 2–10 minutes.
16. Rinse with water.
17. Dehydrate with absolute alcohol.
18. Lighten in xylene, embed in balsam or malinol.

Staining result:
    Nuclei:                                bluish-black
    Cytoplasm, muscle tissue: red
    Erythrocytes:                      red-orange
    Connective tissue:             blue
    Fibrin:                                vivid red
Ref.: Lendrum, A. C. (1949), J. Path. & Bact., LXI 443-448
 


   

Necessary solutions:
A. Water blue	1	g
Orcein	0,75	g
Glycerin	20	ccm
Ethyl alcohol	50	ccm
Acetic acid 5%                                                             B. Eosin 1.25% alcoholic C. Hydroquinone 1% aqueous D. Safranine O 1% aqueous E. Potassium dichromate solution 0.5%	100	ccm

Staining procedure:
 1. Fix in 10% formaldehyde, embed in paraffin or Celloidin.
 2. Immerse in water, stain in a mixture of:
    10 cc of solution A
    3 cc of solution B
    3 cc of solution C for 10 minutes.
 3. Wash thoroughly in water.
 4. Stain with solution D for 10 minutes. 
 5. Wash with water.
 6. Immerse in solution E for 10–30 minutes.
 7. Wash in water, dehydrate with absolute alcohol, lighten in bergamot oil.
 8. Check under a microscope, differentiate with alcohol and bergamot oil if necessary, until overstaining by safranin is reduced.
 9.  Embed in balsam or Malinol.

Staining result:
   Epithelial fibers:         red
   Nuclei:                       pale violet
   Plasmosomes:           red
   Cytoplasm:                 blue to violet
   Neutrophil granules:   sky blue
   Elastic fibers:              red
   Collagen fibers:          blue
Ref.: Carleton, H. M. & Leach, E. H. (1947), Histological Technique, page 285,
(Oxford University Press)
Unna, P. G. (1910), Histotechnique of Leprous Skin, (Voss, Leipzig)
Necessary solutions:
A. Haematoxylin - Weigert A
B. Haematoxylin - Weigert B
C. Eosin, yellowish, 1% aqueous
D. Ferric chloride (anhydrous), 10% aqueous
E. Naphthol Green B, 1% aqueous
F. Acetone-Xylene 1:1

Staining procedure:

1. Adhere the paraffin section and immerse it in water.
2. Stain in a freshly prepared mixture of equal parts of solutions A and B for 6 minutes.
3. Wash in water and stain in solution C for 3 minutes. Wash in water, immerse in solution D, 5 minutes.
4. Rinse thoroughly with water, stain with solution E, 5 minutes.
5. Differentiate in 1% acetic acid, 2–3 minutes.
6. Dry thoroughly, dehydrate with acetone, lighten in solution F, embed in balsam or Malinol.

Staining result:
   Connective tissue:            green
   Muscles and cytoplasm:   pink
Ref.: Lillie, R. D. (1954), J. Techn. Meth., 25:32
Required solutions:

1. Congo red 1% aqueous
2. Lithium carbonate, saturated aqueous
3. Delafield hematoxylin solution
4. Material fixed in formalin or alcohol can be embedded in Celloidin or paraffin or used as a frozen section.

Staining procedure:

1. Adhere the section and immerse it in water.
2. Stain with solution A for 10–30 minutes.
3. Immerse in solution B for 15 seconds.
4. Decolorize with 80% alcohol until no more color scum is released.
5. Wash with running water for 15 minutes.
6. Stain with solution C for 5–10 minutes.
7. Wash with water, dehydrate, lighten with xylene, and embed in balsam or malinol.

Note: If sections embedded in celloidin were used, dehydrate with isopropyl alcohol instead of ethyl alcohol.

Staining result:
    Amyloid:    red
    Nuclei:      blue
Ref.: Bennhold, H. (1922), Münch. Med. Woch., page 1537;
Cowdry, E. V. (1952), Laboratory Technique in Biology & Medicine;
Page 11; (Balliere, Tindell & Cox, London);
Hess, G. (1942), Arch. Path., 34:92-105;
Taren, A. G. (1936-37), J. Lab. Clin. Med., 22:975-977
Necessary Solutions:
A. Carbol-Fuchsin, Ziehl-Neelsen 5 cc
     Distilled Water 45 cc
B. Methyl green 1% in carbolic acid 5%

Staining procedure:
Fix and embed in paraffin.
Add the section to a mount and immerse in water.
Stain with solution A for 15–45 minutes.
Wash with water and dry carefully (with blotting paper).
Dehydrate with alcohol.
Differentiate and counterstain with solution B for 2–3 minutes.
Wash with alcohol.
Lighten in xylene and embed in balsam or malinol.

Staining result:
    Hyaline substance: red
    Nuclei:                     light green
Ref.: Carleton, H. M. & Leach, E. H. (1947), Histological Technique (Oxford University Press) Pages 307-308



  Necessary solutions:
A. Potassium permanganate 2.5%                           1 cc
    Sulfuric acid 5%                                                    1 cc
    Distilled water                                                        6 cc
B. Sodium bisulfite 2% aqueous
C. Ferrous trioxyhaematein, Hansen
D. Aldehyde fuchsin working solution:
    Aldehyde fuchsin stock solution                           25 cc
    Alcohol 70%                                                         75 cc
   Glaceous acetic acid                                              1 cc
Note: This solution can be stored at room temperature for several months. (Preparation of the
         Aldehyde fuchsin stock solution: Dissolve 0.75 g of aldehyde fuchsin in 100 cc of 70% alcohol)
E. Picroindig carmine                                                1 g
    Distilled water                                                    100 cc

Staining procedure:

1. Fix according to Bouin, embed in paraffin or Celloidin.
2. Mount sections, dewax, and immerse in water using a descending alcohol series.
3. Oxidize in solution A for 20–30 seconds.
4. Rinse briefly with water.
5. Wash in solution B.
6. Rinse in running water for 30 seconds.
7. Stain with solution C for 1 minute.
8. Rinse in running water.
9. Stain with solution D for 2 minutes.
10. Rinse briefly in water.
11. Counterstain with solution E for 20 seconds.
12. Dehydrate, lighten with toluene, and embed in balsam or malinol.

Staining result:
    Elastic fibers, cartilage substance, mucin: intense red
    Nuclear chromatin:                                     black or brown
    Basophilic cytoplasm:                                 gray
    Acidophilic cytoplasm, muscle fibers:         green
    Collagen fibers:                                          blue
Ref.: Bull. Micro, appl., 2nd Series, Volume 3, Nos. 11 and 12 Nov. - Dec. 1953 A. Resorcinol fuchsin, Weigert
B. Nuclear red aluminum sulfate 5.1 g
    Distilled water 100 cc

Staining procedure:

1. Fix in Susa, Helly, or Stieve, embed in paraffin, deparaffin.
2. Transfer sections from 80% alcohol into solution A, stain for 10–30 minutes.
3. Stain in solution B for 5–10 minutes.
4. Wash with water.
5. Differentiate with 96% alcohol.
6. Dehydrate, lighten in xylene, embed in balsam or Malinol.

Staining result:
   Nuclei:              red
   Elastic fibers:   blackish-blue
Ref.: Weigert, C. (1898), Zbl. Path., 9:289-292.
   

Necessary solutions: A. Aluminum chloride 2% aqueous
B. Congorot	2 g
Sodium citrate	2,5 g
Glycerin	1 ccm
distilled water	97 ccm
C.Aniline blue	1,5 g
Orange G	2,25 g
Resorcin	3 g
Phosphoromolybdic acid	1  g
distilled water	100 ccm

Staining Procedure:

1. The material can be fixed in 10% formaldehyde or used as a frozen section.
2. Wash the section in water, immerse in solution A for 10 minutes.
3. Wash with water, dehydrate, stain with solution B for 10 minutes.
4. Wash with water, repeatedly and thoroughly by immersion.
5. Stain with solution C for 5–10 minutes.
6. Rinse carefully, dry with blotting paper.
7. Dehydrate with alcohol, lighten in oregano oil, wash in xylene, embed in balsam or malinol.

Staining Result:
   Elastic fibers: light red
   Fibrin:            dark blue
   Erythrocytes: yellowish-orange
Ref.: Krajian, A. A. (1941), Canadian J. Med., 3:207.

 

Necessary solutions:
A. Orcein	1 g
Alcohol 70%	100 ccm
Hydrochloric acid, concentrated B. Ferrous hematoxylin, Weigert A     Weigert B     Mix both in equal parts before use. C. Hydrochloric acid-alcohol	1 ccm
D. Azocarmine G	0,1 g
Boil distilled water,     let it cool, filter it, and add:	100 ccm
Glacial acetic acid	1 ccm
E. Phosphoric tungstic acid	5 g
distilled water	95 ccm
F. Naphthalene Green B	1 g
distilled water	100 ccm
Glacial acetic acid	1 ccm

Staining procedure:

1. Fix in Zenker, Helly, Bouin, or Carnoy.
2. Transfer sections from 70% alcohol into solution A; stain for 12 hours.
3. Wash in water until excess stain is completely removed.
4. Stain in solution B for 12–30 minutes.
5. Wash in water.
6. Briefly differentiate in solution C.
7. Wash under running water for 6 minutes.
8. Stain in solution D (microscopic inspection is necessary, as overstaining is common).
9. Wash in water.
10. Decolorize collagen fibers with solution E, changing the bath 2–3 times; total treatment time is 3–6 hours.
11. Wash in water.
12. Stain with solution F for 15–30 minutes.
13. Transfer to 95% alcohol and shake for 30 seconds.
14. Dehydrate, lighten, and embed in balsam or Malinol.

 Staining Result:

Elastic fibers:	black
Collagen fibers:	green
Skeletal muscles:	bright red
Smooth muscle:	pink
Epithelial cells:	red
Nuclear chromatin:	black

Lit.: Mollier, G. (1938), Zts. Wis. Mikr., 55:472
Required Solutions:
A. Silver nitrate 0.1% aqueous solution                                                         3 ml
    Distilled water                                                                                         100 ml
    Mix well, add:
   
    Sodium chloride 0.1% aqueous solution                                                    1 ml
    Mix well, if necessary adjust the pH to 7.8 by adding a few drops of highly diluted ammonia solution.
Note: This solution is not stable and must be stored away from light.
B. Hydroquinone                                                                                             1 g
    Sodium sulfite                                                                                            10 g
    Distilled water                                                                                           100 cc
C. Gold chloride 0.5% aqueous
D. Oxalic acid 2% aqueous
F. Sodium sulfite 5% aqueous

Staining procedure:

1. Fix in formalin or Bouin.
2. Transfer to a 2% solution of ammonia (specific gravity 0.89) in 70% alcohol for several hours.
3. Wash thoroughly in water, dehydrate, and embed in paraffin.
4. Mount sections, remove paraffin, and immerse in water using a descending series of alcohols.
5. Stain in solution A under cover of light at 58°C for 16 hours.
6. Immediately transfer to solution B at 18–20°C for 5 minutes.
7. Wash thoroughly in water.
8. Tone in solution C for 10 minutes.
9. Wash twice in water for 15 seconds each time.
10. Reduce in solution D until the nerve fibers are clearly visible, approximately 3 minutes.
11. Wash in running water.
12. Fix in solution E for 3–5 minutes.
13. Wash thoroughly.
14. Dehydrate, lighten, and embed in balsam or Malinol.

Staining result:
    Nerve fibers:        purple to black
    Nuclei:                  red
    Neurofibrils:          purple
    Keratin:                 yellow
    Bone cells:            black
Ref.: Romanes, G. J. (1950), J. of Anatomy, 84, II, 104-116
Required solutions:
  A. 4 g sodium hydroxide per analysis are dissolved in exactly 100 ml distilled water = stock solution.
  B. 10% silver nitrate solution 5 ml, pure pyridine 25 drops, distilled water 20 ml, absolute alcohol. 15 ml. This solution is stable for 1-2 months.
  C. Hydroquinone 1 g, 250 ml distilled water, formalin 70 ml, acetone 40 ml.

Staining procedure:

1. Fix in formalin 1:9 water for at least 3 days, but not longer than 3 months.
2. Prepare frozen sections of 25-30 µm, wash thoroughly in distilled water.
3. Detoxify for 5-8 minutes at 60°C in diluted A. Use 10 ml of A per 50 ml of distilled water for each of the following: for nerve fibers of the brain, spinal cord, ganglia, and eye; 6 ml for nerve cells; 5 ml for sympathetic ganglion cells; 10 ml for axial cylinders; 5 ml for sheaths; 0.5 ml for neurokeratin.
4. Wash in distilled water, changing the solution several times. Add water until the wash water, upon the addition of a few drops of 1% aqueous phenolphthalein, no longer turns red.
5. Immerse the sections in solution B, warmed to 60°C, until they are chestnut brown.
6. Reduce for a few minutes in solution C.
7. Wash thoroughly.
8. Dehydrate in the usual manner and transfer to balsam or Malinol.

Staining result:
    Aximal cylinders and neurofibrils stained black; ganglion cells stained brownish.

Note: For gilding, place the sections as described in step 7 in a 0.2% gold chloride solution for 5 minutes, then wash in distilled water, transfer to a 5% aqueous sodium thiosulfate solution for one minute, and then proceed as described in step 8. For neuroglia, fix in 110 ml formalin, 16 g ammonium bromide, and distilled water instead of formalin 1:9. Water 500 ml, specifically microglia 2–3 days, protoplasmic glia 15–30 days, fibrous glia 1.5–2 months. Material thus pretreated can then be stored in 10% formalin.
Lit.: Lobo 1937. Sobre una nova técnica de impregnacao do sistemo nervosa. Patologia general. Vol. 22, pp. 4–7
 


Required solutions:
A. Chromium alum mordant = 2.5 g of chrome alum are heated to a boil in 100 ml of distilled water. When the solution is boiling vigorously, the flame  is extinguished.
     Then, while stirring, 5 ml of 30% acetic acid are added, and finally, while continuously stirring with a glass rod, 5 g of finely powdered neutral          copper acetate are added.
B. Haematoxylin solution according to Böhmer or Delafield.
C. 2 g of borax and 2.5 g of potassium ferricyanide are dissolved in 100 ml of distilled water while heating.

Staining procedure:

1. Fix in formalin 1:10 and prepare frozen sections.
2. Transfer the sections from water to solution A without alcohol treatment for several hours.
3. Wash thoroughly in distilled water.
4. Immerse in solution B for 24 hours.
5. Wash thoroughly in tap water and differentiate in C until the gray matter is clearly distinguishable from the white matter.
6. Wash thoroughly in tap water and stain via the ascending alcohol series, encapsulating xylene in balsam or malinol.

Staining result:
   The myelin sheaths are stained dark blue, degenerations red.
Ref.: C. Benda, Neurol. CentralbL, Vol. XXII, 1903, No. 3, pp. 139–140.
Required solutions:
A. 8–9% solution of potassium permanganate in distilled water.
B. Potassium sulfite-oxalic acid: Immediately before use, mix equal amounts of 1% potassium sulfite solution and 1% oxalic acid solution.
    The mixture has a short shelf life. In contrast, the separate solutions have an unlimited shelf life.
C. Immediately before use, mix 2 ml of a 1% gold chloride solution with 2 ml of a 5% sublimate solution and 10 ml of distilled water. This amount is sufficient for a maximum of 4 sections. The separate gold chloride and sublimate solutions are stable.
D. 2% aqueous solution of sodium thiosulfate.

Staining procedure:

1. Fix in formalin 1:0, frozen sections 25–30 µm thick.
2. Collect the sections in water onto strips of filter paper and transfer them to A. Transfer the sections back into water after 10 minutes in the same manner. The sections become so brittle in the potassium permanganate solution that the usual transfer with glass rods is not possible. In pure water, they regain their normal firmness.
3. Rinse very briefly with distilled water.
4. Immerse the sections in B until they are decolorized.
5. Briefly rinse in two trays of distilled water.
6. Transfer to C and leave in the dark for 20 hours.
7. Rinse thoroughly in distilled water (changed several times) and fix for 1 minute in D to prevent darkening.
8. Rinse thoroughly in distilled water (changed several times), then proceed through the ascending alcohol series, encapsulating xylene in balsam or malinol.

Staining result:
   The connective tissue fibers are very sharply impregnated in deep black on a violet background.
Reference: Cited in Romeis, Microscopic Technique, 15th ed., 1948, § 1882 Required Solutions:

A. Mordant I = Dissolve 5 g of potassium dichromate and 2.5 g of fluorochrome in 100 ml of water by boiling. Filter after cooling.
B. Mordant II = Dissolve 2.5 g of fluorochrome in 100 ml of distilled water by boiling. When the solution boils, extinguish the flame and
    add 5 ml of glacial acetic acid and then, while stirring continuously with a glass rod, add 5 g of finely powdered neutral cupric acetic acid.
C. Dissolve 1 g of haematoxylin in 100 ml of 96% ethyl alcohol. The solution must be well-matured, i.e., have a brown, not yellow, hue.
D. Mix 4 ml of ferric chloride solution with 96 ml of distilled water. Immediately before use, mix equal parts of C and D.
E. Dissolve 2 g of borax and 2.5 g of potassium ferricyanide in 100 ml of distilled water with warming.

Staining procedure:

1. Fix in 10% formalin.
2. Partrate the pieces, which should be no more than 2 cm thick, in A for 4–14 days; the white substance should eventually turn brown, not yellow.
3. Wash in the dark in 70% alcohol until it no longer stains.
4. Embed in celloid and prepare sections.
5. Pickling of sections: 24–48 hours at 37°C in B.
6. Rinse in 70% alcohol and stain in mixture C-D for 24 hours.
7. Rinse in tap water and differentiate in E until the white substance (myelin sheaths) appears black, while the gray substance appears light yellow to brownish, on average 15–30 minutes.
8. Wash in tap water for 24 hours, possibly with the addition of a few drops of aqueous lithium carbonate solution.
9. Dehydrate, xylene, and encapsulate in balsam or malinol.

Staining result:
   The myelin sheaths are bluish-black, all other tissue is yellowish-brown.
Lit.: Mayer, Paula (1909). On the technique of myelin sheath staining. Neurolog. Zbl., Vol. 28, pp. 353–354.

 


Required solutions:
  A. Silver nitrate solution 3% aqueous, freshly filtered before use.
  B. Ammoniacal silver nitrate solution: 5 drops of the purest 40% sodium hydroxide solution are added to 10 ml of a 10% silver nitrate solution while          shaking. A brown precipitate forms. Now strong ammonia is added drop by drop while shaking constantly until most of the precipitate is dissolved        again. There should not be an excess of ammonia. Therefore, some grains of the brown precipitate are left undissolved. These are
      filtered off and the filtrate is mixed with distilled water. Water topped up to 20 ml
 C. Gold chloride solution: 3 - 5 drops of a 1% solution of yellow gold chloride are dissolved in 10 ml of distilled water. Water dripped. D. 5% aqueous        solution of sodium thiosulfate.

Staining procedure:
Use fresh material if possible, frozen sections 5 - 10 µ thick. Do not use metal instruments.

1. Fix in 40% formalin 1 part and 9 parts tap water for 24 - 48 hours.
2. Wash in running water for 2 - 3 hours.
3. Cutting on the freezing microtome and collecting the sections in distilled water. Water, section thickness 5 - 10 µ.
4. Another 1 - 2 hours in distilled water, changed 3 - 4 times. Water.
5. Place in A for 24 hours.
6. Quickly steep through distilled water. water for about 2 - 3 seconds. If the cut remains in the dest for too long. water, perfect impregnation will not occur.
7. Place in B for 10 - 20 minutes until the cut has taken on a yellowish tint.
8. Quickly strain through 2 - 3 bowls of distilled water. Water.
9. Reduce in acid-free Formol (1 part formula: 4 parts tap water) for 10 minutes; the sections will turn slate gray to blackish.
10. Wash in tap water for 15 minutes.
11. Place in C until the brown tone of the preparation has changed to gray or gray-violet.
12. Fix in D for 1 - 2 minutes.
13. Wash carefully in tap water for 1 - 2 hours.
14. Carry out through the ascending alcohol series, carbolxylol, xylene (no longer than absolutely necessary) and enclose in balsam or malinol.

Staining result:
    Neurofibrils colored black on a light background, as well as the pericellular network structures of the ganglion cells. Even the thinnest
    axis cylinders stand out clearly.
Lit.: Bielschowsky, Journal for Psychology and Neurology, Volume 3, pages 169 -189 and
Volume 4, pages 227 – 236
 

Required solutions:
  A. Fixation solution = 5 ml formalin, 5 ml glacial acetic acid, 90 ml 80% alcohol. The result of Bodian's method largely depends on the type of         
      fixation. Bodian therefore specified around 10 different fixing mixtures for the various elements. In most cases you can get by with the above                mixture.
  B. Weigh out 1 g of silver protein and dust it directly from the weighing paper onto the surface of 100 ml of water in a very fine distribution. Leave to        stand until the powder has sunk to the bottom. Only then is the powder completely dissolved by shaking. The solution is not heated.
  C. Hydroquinone 1 g, sodium sulfite anhydrous (Na2SO3) 5 g are dissolved in 100 ml of water.
  D. Gold chloride solution 1% aqueous, to which 3 drops of glacial acetic acid are added to 100 ml before use.
  E. Oxalic acid solution 2% aqueous
  F. Sodium thiosulfate solution 5% aqueous.

Staining procedure:

1. The fixed and deparaffinized sections come from water for 12 - 48 hours at 37 ° C in B, to which 4 - 5 g of metallic copper (copper wire or copper shavings, carefully cleaned) per 100 ml were added immediately before the sections were placed.
2. Rinse in distilled water. Water.
3. Reduce in C for about 10 minutes.
4. Wash in distilled water that has been changed at least three times. Water.
5. Transmitted in D for a duration of 5 - 10 minutes.
6. Wash in distilled water. Water.
7. Adjust in E until the cuts are slightly reddish or bluish.
8. Wash in distilled water. Water.
9. Fix in F for 5 - 10 minutes.
10. Wash in distilled water. Water, dehydrate, enclose in balm or malinol.

Strain result:
   Axial cylinders and neurofibrils are colored black or purplish-black
Note: The silver protein solution can only be used once. Celloidin sections must be freed from celloidin before impregnation. Cuts of old formol material are placed in 5% acetic acid overnight before impregnation. This prevents the collagen fibers from coloring. Counterstaining with acridine red, azan solution or light green-orange according to Foley is possible and can be recommended.
Lit.: Bodian, 1937, Anat. Rec. 69, pp. 153 - 162

  When using eosin-methylene blue solutions, the following must be observed: To consistently achieve usable results, only clean glass vessels that have not come into contact with acid may be used. The slides or coverslips must be cleaned only with ether-alcohol, never with acid or hydrochloric acid-alcohol. Even repeated rinsing in pure water does not remove the last traces of acid, so proper staining cannot be obtained. The water used to dilute the staining solutions must be absolutely neutral. Its suitability must be tested as follows: Add 3 drops of bromocresol purple solution to 100 ml of distilled water and then, while continuously shaking, add 1% sodium carbonate solution dropwise until the yellow color of the water changes to violet. This violet hue must persist for at least one minute. If the solution turns yellow or colorless again during this time, the amount of sodium carbonate added was insufficient, and another drop should be added. The number of drops determined in this way is then added to every 100 ml of the water to be used for dilution. This water test must be repeated each time, as water stored in the examination room can change its acidity daily due to atmospheric influence.

The water should be at least at room temperature; otherwise, it should be warmed. However, temperatures above 40°C should be avoided. Dye precipitates can be removed by pouring absolute alcohol over the preparations after staining, rinsing them with water, and then drying them. Preparations that are 2–24 hours old stain best. Older preparations stain more slowly and usually show a reddish-brown erythrocyte stain. In overly thick smears or overstained preparations, the erythrocytes often have a narrow blue border or appear blue altogether.

Correctly stained specimens show a red to violet-red hue in bright light. Specimens that are stained for too short a time appear gray to, at most, a pale pink. Overstained specimens have a pronounced violet to blue tint. In both cases, improvements can be achieved by first removing the cedarwood oil with xylene, then drying the specimens, and then re-immersing them in the staining solution for a few minutes. In the case of overstaining, rinse them vigorously with water for an extended period, or—if this is unsuccessful—immerse the specimens in methyl alcohol for a few seconds.

Jenner staining (fresh smears)

Required solution:
A. Jenner's Eosin-Methylene Blue

Staining procedure:

1. Immerse the thinnest possible air-dried, unfixed smears in solution A for 3 minutes.
2. Add the same amount of distilled water as was used in the staining solution, mix well, and stain for 10–15 minutes.
3. Rinse briefly in distilled water.
4. Dry with blotting paper.
5. Permanent slides: Enclose in Canada balsam neutral or Malinol.

Staining result:
   Nutrients:                     blue
   Eosinophilic granules: deep red
   Neutrophilic granules: brownish-violet
   Erythrocytes:               light red
   Platelets:                     pale blue

May-Grünwald staining (fresh smears)

Required solution:
A. May-Grünwald Eosin-Methylene Blue

Staining procedure:

1. Overlay well-air-dried, unfixed smears with solution A for 3 minutes.
2. Add an equal amount of distilled water, mix thoroughly by swirling, and stain for 5–10 minutes.
3. Rinse thoroughly with distilled water.
4. Blot carefully with blotting paper and allow to air dry completely.
5. Permanent slides: Enclose in Canada Balsam Neutral or Malinol.

Staining result:

Cores:	blue
Eosinophilic granules:	bright red
Basophilic granules:	dark blue
Neutrophil granules:	light red to purplish-red granules
Erythrocytes:	bright red
Platelets:	light blue

May-Grünwald staining (for older or difficult-to-stain specimens)

Required solution:
A. May-Grünwald Eosin Methylene Blue

Staining procedure:

1. Fix the smears in absolute alcohol for 10–20 minutes.
2. Stain for 10–15 minutes in a mixture of one part solution A and two parts distilled water.
3. Rinse with water and dry between blotting paper.
4. Permanent slides: Encase in Canada Balsam Neutral or Malinol.

Staining result:
The result is the same as described for fresh smears.

Leishman staining

Required solution:
A. Leishman Eosin Methylene Blue (10680 and 20220)

Staining procedure:

1. Cover air-dried smears as thinly as possible with solution A for 2 minutes.
2. Add twice the amount of distilled water. Mix with water by swirling and stain for 10 minutes.
3. Rinse thoroughly with distilled water.
4. Dry between sheets of tissue paper and in the open air.
5. Permanent preparations: Encase in Canada Balsam Neutral or Malinol.

Staining result:
   Erythrocytes: light pink to brownish-red
   Lymphocyte nuclei: deep dark blue to blue-violet
   Lymphocyte protoplasm: light blue
   Neutrophilic, polymorphonuclear leukocyte nuclei: deep blue to blue-violet
   Neutrophil polymorphonuclear leukocyte granules: red
   Eosinophil leukocyte nuclei: blue-violet
   Eosinophil leukocyte granules: deep dark red
   Basophil leukocyte nuclei: blue-violet
Required solution:
A. Wright Eosin Methylene Blue

Staining procedure:

1. Cover very thin, air-dried smears with solution A for 2 minutes.
2. Add the same amount of distilled water, mix, and stain for 5 minutes.
3. Rinse thoroughly with distilled water.
4. Dry with blotting paper and air dry.
5. Permanent preparations: Encase in Canada Balsam Neutral or Malinol.

Staining result:
  The result is the same as described by Leishman.


Giemsa staining

Required solution:
A. Azure-Eosin, Romanowsky-Giemsa

Staining procedure:

1. Fix air-dried, thin smears in methyl alcohol for 5 minutes.
2. Stain in solution A (diluted 10 drops to 10 cc of water) for 20–30 minutes.
3. Rinse with distilled water.
4. Dry between blotting paper and in the open air.
5. 5. Permanent slides: Encase in Canada Balsam Neutral or Malinol.

Note: Buffered water can be used instead of distilled water.


Staining result:
     Nuclei:                             blue-violet
     Eosinophilic granules:     red
     Neutrophilic granules:     red-violet
     Basophilic granules:        blue to blue-violet
     Lymphocyte protoplasm: light blue
    Erythrocytes:                    reddish
    Protozoan nuclei:             Bright red
    Platelets:                          blue with a violet core
   

Giemsa stain (in a thick drop)

Required solution:
A. Azure-Eosin, Romanowsky-Giemsa

Staining procedure:

1. Preparation of the slide: Spread a large drop of blood onto a slide thoroughly cleaned with ether-alcohol until it is approximately 1 cm in diameter. The blood layer should not be too thick, otherwise it will easily flake off and be difficult to stain. Allow the blood to dry at room temperature (beware of flies) or in an incubator at 37°C. Higher temperatures should be avoided, as the blood layer will easily flake off.
2. Fix and stain simultaneously with solution A (dilute 10 drops in 10 cc of distilled water) for 3 minutes.
3. Any greenish-yellow haemoglobin clouds that now appear are poured off and, after tilting the slide, rinsed with fresh, also diluted, staining solution.
4. Re-pour diluted staining solution and stain for 45 minutes or longer.
5. Rinse by pouring distilled water laterally.
6. Dry by holding the slide upright (do not use blotting paper).

Staining result:
   Well-stained preparations appear reddish-violet, poorly stained ones blue. Eosinophilic leukocytes (red dots) and basic erythrocytes (blue dots) are     easily identifiable.
Note: To avoid the difficulties that can arise in preparing absolutely neutral water, the use of buffered water with a buffer mixture according to Weise is recommended.

Staining according to Giemsa-Pappenheim


Required solutions:
  A. May-Grünwald eosin-methylene blue
  B. Romanowsky-Giemsa azure eosin

Staining procedure:

1. Air-dried, thin smears (layer side down) undercoat with solution A, 3 minutes.
2. Add an equal amount of distilled water and stain, 1 minute.
3. Pour off the staining solution (do not rinse).
4. Pour over solution B (dilute 15 drops to 10 cc of distilled water) and stain, 10–20 minutes.
5. Rinse thoroughly with distilled water.
6. Dry between blotting paper and in the open air.
7. Permanent slides: encase in Canada Balsam Neutral or Malinol.

Staining result:
   Nuclei:                                                    reddish violet
   Plasma of lymphoid cells:                       light blue
   Lymphoid azure granules:                      bright purplish-red
   Myeloid azure granules:                         violet to violet-brown
   Neutral granules:                                    brownish to bluish-pink
   Eosinophilic granules:                            brownish-orange to brick red
   Mast granules:                                       ultramarine with a violet tinge
   Erythrocytes:                                          pink to bluish
   Basophilic punctation of erythrocytes:   intense cobalt blue

MacNeal stain (smear preparations)

Required solution:
A. Tetrachrome Stain, MacNeal

Staining procedure:

1. Cover thin, air-dried smears with solution A for 2 minutes.
2. Add an equal volume of distilled water, mix, and stain for 5 minutes.
3. Rinse thoroughly with water. Dry (with blotting paper), then embed in balsam or Malinol.
4. Dry on blotting paper.

Staining results:
   Erythrocytes:                                      yellowish-red
   Polymorphonuclear neutrophils:         nuclei dark blue, granules reddish-purple
   Cytoplasm:                                          pale pink
   Eosinophils:                                         nuclei blue, granules orange to red, cytoplasm blue
   Basophils:                                            nuclei purple or dark blue, granules deep purple
   Lymphocytes:                                      nuclei dark purple, cytoplasm sky blue
   Platelets:                                             granules violet to purple
Ref.: MacNeal, W. J., (1922), J. Amer. Med. Assoc., 78, 1112

Vital (Supravital) Staining

Necessary Solutions:
  A. Alcoholic Brilliant Cresyl Blue Solution
  B. Azure Eosin Solution

Staining Procedure:

1. Place one drop of solution A onto a carefully cleaned slide and spread it in the manner of blood smears.
2. Spread the blood onto the emulsion side of the slide as usual, but not too thinly.
3. Place in a humid chamber, stain-side up, and cover the chamber.
4. After 5–10 minutes, remove the slides and allow them to air dry.
5. Fix the air-dried smears in methyl alcohol for 3–5 minutes.
6. Stain in solution B (diluted 10 drops to 10 cc of neutral water) for 20–30 minutes.
7. Rinse with water and dry between sheets of blotting paper.
8. Enclose in Canada balsam neutral or Malinol.

Staining result:
   This method is primarily used to visualize the vital staining substances of erythrocytes (granulofilamentous substance, basophilic granulation,         
   polychromasia). It produces very delicate blue images of the network structure while preserving all other components.
 

1. Fix air-dried smears in methyl alcohol for 3 minutes.
2. Do not rinse, but immediately transfer to the diluted Löffler's methylene blue solution for another 3 minutes.
3. Rinse with water, dry between sheets of blotting paper.
4. Examine by oil immersion under full illumination.

  A. Boric acid 2 g, pure methylene blue 1 g, distilled water 100 ml = Manson-Black I
  B. Pure sodium hydroxide 0.28 g dissolved in 100 ml distilled water Water = Manson-Black II

Staining Procedure:

1. Fix air-dried blood smears for 3–5 minutes in methyl alcohol.
2. Rinse with water and allow to air dry.
3. Stain for 15–20 seconds in a mixture of 6 drops of A, 8 drops of B, and 10 ml of distilled water.
4. Rinse carefully with distilled water and dry between blotting paper.
5. Examine by oil immersion.

Necessary solutions:
A. distilled water	95	ccm
iron alum	5	g
Celestine blue     Boil for 3 minutes, let cool, filter, add:	0,5	g
Sulfuric acid (concentrated)	2	ccm
Glycerin                                                                             B. Haemalaun, Mayer C. Lithium carbonate solution, saturated aqueous	14	ccm
D. Alcohol 70% ig	99	ccm
Hydrochloric acid, concentrated.	1	ccm
E. Barlow's Trichrom sicc.	1,55	g
distilled water	60	ccm
Nitric acid (concentrated) Boil vigorously for 5-10 minutes, let cool, add:	0,25	ccm
Glycerin                                                                                 Top up with distilled water to 150 cc Note: Solution has a limited shelf life	50	ccm
F. Acetic acid 1% aqueous	100	ccm
Lissamine True Yellow	0,1	g

  1. Fix in Bouin's solution or alcohol.
  2. Stain in solution A for 2 minutes.
  3. Rinse under running water.
  4. Stain in solution B for 2 minutes.
  5. Rinse and blue with solution C.
  6. Differentiate with solution D.

Check under the microscope until the cytoplasm lightens and the nuclei become visible.
  7. Stain in solution E for 5–15 minutes.
  8. Wash under running water.
  9. Counterstain with solution F or G.
 10. Rinse in water.
 11. Dehydrate with alcohol.
 12. Lighten in xylene, embed in balsam or malinol.

Mast cell granules: cartilage, Keratin,	orange to red
Bacteria:	pink to red
Mucin:	zart rosa
Cores:	blue
other tissue components:	grün oder hellgelb (je nach Gegenfärbung)

1. Fix in 10% formaldehyde or 80% alcohol, embed in paraffin.
2. Mount the sections, deparaffinize with xylene.
3. Immerse in water, through a descending alcohol series.
4. Stain with solution A, 6 minutes.
5. Rinse under running water, 2–3 minutes.
6. Wash in an ascending alcohol series.
7. Stain with solution B, 5 minutes.
8. Rinse thoroughly in 95% alcohol (change containers).
9. Stain with solution C, 2.5 minutes, shaking occasionally.
10. Rinse thoroughly in 95% alcohol (change containers). Dehydrate completely with alcohol.
11. Brighten with xylene, embed in balsam or malinol.

1. Fix in Helly's solution for 6–24 hours, then embed in paraffin.
2. Mount sections, remove paraffin, and introduce them into water using a descending alcohol series.
3. Stain in solution A for 1 hour.
4. Wash under running water.
5. While still moist, check under a microscope. If beta cells do not appear bluish-violet, stain again for one hour with solution A and check again.
6. Dehydrate by an ascending series of alcohols.
7. Clear in toluene, embed in Canada balsam or malinol.

1. Fix in equal parts ether and alcohol for 5–15 minutes.
2. Wash thoroughly in a descending alcohol series.
3. Wash in water.
4. Stain in solution A for 20 minutes.
5. Wash in water.
6. Differentiate in 0.5% hydrochloric acid.
7. Wash in running water for several hours.
8. Bluent in saturated aqueous lithium carbonate solution.
9. Wash well in water.
10. Pickle in solution B for 15 minutes.
11. Wash with water.
12. Mordant in solution C for 5 minutes.
13. Wash well in water. Stain in solution D for 20 minutes.
14. Wash thoroughly with water.
15. Air dry, then embed in balsam or Malinol.

 

Necessary solutions: A. Aluminium acetate 2% aqueous
B. Alizarin Red S	1 g
Distilled water C. Polychrome Methylene Blue, Unna	99 ccm

1. Pickling of the alcohol-fixed and paraffin-embedded sections in solution A for 24 hours.
2. Wash in water.
3. Staining in solution B for 24 hours.
4. Wash in water.
5. Wash in 95% alcohol at 60°C.
6. Counterstaining in solution C.
7. Wash, dehydrate, lighten in xylene, and embed in balsam or malinol.

1. Fix in Zenker or Bouin, embed in paraffin.
2. Adhere sections, deparaffinize.
3. Mordant in solution A for 30 minutes to 3 hours using a descending alcohol series.
4. Wash in water.
5. Stain with solution B for 1–3 hours.
6. Wash in water.
7. Differentiate with solution A (check under the microscope).
8. Wash in water for 5–10 minutes.
9. Opponent stain with solution C for 2–5 minutes.
10. Dehydrate, lighten in xylene, embed in balsam or Malinol.

  A. Sudan solution, Daddi
  B. Emeralum, acidic, Mayer

Staining Procedure:

1. Use frozen sections.
2. Fix in formalin or Helly.
3. Transfer sections to 50% alcohol.
4. Stain with solution A for 30 minutes.
5. Immerse over 50% alcohol in water.
6. Counterstain with solution B for 4–6 minutes.
7. Rinse with running water.
8. Embed in glycerin gelatin Kaiser or gelatinol.

Necessary solutions:
A. Sudanschwarz	0,1 g
Alcohol 70 %ig	100 ccm
B. Kernechtrot aluminum sulfate	5,2 g
distilled water	95 ccm

1. Fix the still-moist smears with equal parts ether and alcohol for 1-2 minutes.
2. Stain with solution A for 1 minute.
3. Dehydrate in 70% and absolute alcohol by repeated immersion.
4. Lighten in xylene, embed in balsam or malinol.

1. Fix in Regaud, embed in paraffin, de-paraffin, immerse in 95% alcohol over absolute alcohol.
2. Immediately immerse in solution A, stain for 2–4 hours.
3. Wash off excess stain with 95% alcohol and water.
4. Differentiate with solution B, 3–5 seconds.
5. Wash with 70% alcohol and water.
6. Immerse in solution C, 5 minutes.
7. Transfer to solution D, 20 minutes.
8. Wash several times in water.
9. Immerse in solution E, changing the bath three times, 1–2 minutes each time.
10. Wash in water, dehydrate, lighten in xylene, embed in balsam or Malinol.

1. Fix and embed.
2. Mount the sections, deparaffinize, and introduce them into water using a descending alcohol series.
3. Stain in solution A (heated to 62°C) for 6 minutes.
4. Rinse in water.
5. Introduce in solution B for 10 minutes.
6. Rinse in water.
7. Mordant in solution C for 1 minute.
8. Wash in water.
9. Counterstain with solution D for 3–6 minutes.
10. Dehydrate by an ascending alcohol series (microscopic control, as safranin is alcohol-soluble), lighten in xylene, and embed in balsam or malinol.

E. Sodium metabisulfite                                                                              0,5 g
Distilled waterWasser dest.                                                                                   99 ccm
Hydrochloric acid, concentrated.                                                                               1 ccm

1. Fix in 4% formaldehyde solution in 90% alcohol, embed in paraffin.
2. Deparaffin, add to water via a descending alcohol series.
3. Stain in solution A, 10 minutes.
4. Wash in water.
5. Immerse in solution B, 6 minutes.
6. Rinse thoroughly in water.
7. Adjust in solution C, 10 minutes.
8. Rinse well with water, 5 minutes.
9. Immerse in solution D, 15 minutes.
10. Rinse with solution E, changing the bath three times for 2 minutes each time.
11. Wash in water, 2–5 minutes.
12. Clarify in xylene via an ascending alcohol series, embed in balsam or malinol.

1. Fix in Bouin or formalin, embed in paraffin, de-perfume.
2. After treatment with absolute and 95% alcohol, immediately stain in solution A at 55°C for 2–4 hours.
3. Remove excess dye with 95% alcohol and water.
4. Differentiate by rapid immersion in solution B for 3–5 seconds. Then perform four washes with 70% alcohol. The final alcohol bath must be free of dye.
5. Wash in water (differentiation check under a microscope).
6. Opponent colors with solution C for 5–10 seconds.
7. Wash in water, dehydrate, lighten in xylene, and embed in balsam or malinol.

1. Fix in formalin 1-2 weeks before staining.
2. Wash in water for at least 6 hours.
3. Dehydrate twice in 95% alcohol for 1 hour each time.
4. Stain at 45-55°C in solution A for 16-18 hours.
5. Remove excess dye with 95% pure alcohol.
6. Rinse in water.
7. Differentiate with solution B, changing the bath several times.
8. Rinse several times in 70% alcohol while shaking.
9. Wash in water.
10. Embed in gelatinol.