Fixatives, Stains and Staining Schedules

In this article we will discuss about the fixatives, stains and staining schedules used for anatomical studies of plant.

Fixatives:

A large number of chemicals such as ethyl alcohol, formalin, acetic acid, chloroform, mercuric chloride, chromic acid, picric acid, osmic acid, etc. are used singly or in combinations as fixatives for anatomical studies (For principle of fixation — see Cytology section). Amongst these, formalin and formalin-aceto-alcohol (FAA) are most commonly used in Anatomy.

Formalin:

A 5% to 10% aqueous solution of commercial formalin (35% to 40% concentration) is used for fixation. For making a 5% solution, 5 ml of comm. formalin is taken and dist. water is added to make the volume 100 ml (Plant Micro-technique by Johansen).

Formalin, as a rule, penetrates slowly and is one of the best hardening agents. It is a powerful reducing agent. It does not precipitate proteins or render them insoluble in water. Formalin neither destroys not preserves fats but, more or less, preserves phospholipids.

Formalin-Aceto-Alcohol:

This fixative, commonly called FAA, is the standard fixative for plant tissues meant for anatomical studies.

A number of variations have been proposed, but the standard proportions are:

70% (or 50%) ethyl alcohol — 90 ml

Glacial acetic acid — 5 ml

Formalin (40% formaldehyde i.e. comm. formalin) — 5 ml

or,

70% ethyl alcohol — 85 ml

Glacial acetic acid — 5 ml

Formalin — 10ml

The lower percentage of alcohol is more suitable for delicate materials, such as bryophytes. For hard and woody materials, it is better to decrease the amount of acetic acid and to increase the amount of formalin, i.e. use the second mixture, since formalin penetrates more slowly. The material may be left in FAA almost indefinitely without any appreciable damage.

The minimum time of fixation is 18 hours. Materials to be sectioned are removed with forceps and washed in running water 1/2 to 1 hour and then handled freely. This is a corrosive liquid, and if it comes in contact with the skin it should be washed-off immediately.

Stains:

A large number of natural and synthetic dyes are used as anatomical stains, such as hematoxylin, brazilin, hematein, Bismarck brown, eosin, fast green, gentian violet, light green, safranin, Sudan IV, etc. (for principle of staining, see Cytology section).

Hematoxylin:

Hematoxylin is a chromogen obtained from the heartwood of the plant Hematoxylin campechianum. The dye solution itself has no affinity for tissues, unless iron or aluminium is present as a mordant. The colour effect of hematoxylin depends upon the pH of the solvent and after-treatment of the tissue.

In acidic medium the colour is red and in alkaline medium it is blue. Hematoxylin stain can be prepared according to various schedules, such as Heidenhain’s iron hematoxylin, Delafied’s hematoxylin, Ehrlieh ‘s hematoxylin, etc.

Heidenhain’s Iron Hematoxylin:

0.5% solution of hematoxylin in distilled water is used. Practically, a 10% solution in absolute ethyl alcohol is prepared and then diluted with distilled water to 0.5% when required. The solution is allowed to stand for a few days to ripen (into hematein). The solution may also be ripened more quickly following the method described under Ehrlieh’s hematoxylin. On ripening, the solution attains a rich wine-red colour.

A more stable solution can be made by adding 5 ml of 10% absolute ethyl alcohol solution to 100 ml methyl cello solve, 50 ml distilled water and 50 ml tap-water that contains calcium compounds in solution.

If, on shaking, the solution does not acquire a rich wine-red colour, add a pinch of sodium bicarbonate and shake vigorously. Ripening occurs almost immediately, and the solution can be used at once. This solution lasts longer than simple aqueous solutions and does not spoil at night temperature.

Hematoxylin staining requires mordanting, which can be done by ferric ammonium sulphate (iron alum) or ferric chloride; 2 to 5% solutions are used for mordanting and 1 to 3% solutions for differentia­tion.

Ehrlieh’s Hematoxylin:

This is extensive used with safranin on woody tissues.

It is prepared in the following way:

Distilled water — 50 ml Glacial, acetic acid — 5 ml

Absolute ethyl alcohol — 50 ml Hematoxylin crystals — 1 g

Glycerine C.P. — 50 ml Aluminium potassium sulphate — to excess.

Keep the solution in a dark place until it becomes deep red in colour. It may also be ripened or matured in 3 to 4 hours by exposure to a quartz mercury lamp. Place the solution in a wide and shallow evaporating dish at a distance of about 2 feet (61 cm) from the mercury lamp, stir the solution frequently during exposure.

Delafield’s Hematoxylin:

This is used in mixture with safranin for staining woody plant tissues. To 400 ml of a saturated aqueous solution of ammonium aluminium sulphate add drop by drop a solution of 4 gm. of hematoxylin crystals in 25 ml of 95% ethyl alcohol. Expose this to light and air for 4 days.

Then add 10 ml G.P. glycerin and 100 ml methyl alcohol. Allow this solution to stand for 2 months, exposed to air, till the colour turns sufficiently dark. Alternatively, expose the solution to a quartz mercury lamp for 2 hours as described above.

Bismarck Brown Y:

This is a synthetic dye. It is obtained from coal-tar and belongs to the azo group. Bismarck brown is a basic dye. Its solubility is 1.36% in water and 1.08% in alcohol. A 1% solution in 70% ethyl alcohol is used. Cellulose and mucin walls are stained bright brown. The stain is permanent and it rarely over-stains. It is generally used for relatively soft plant tissues without lignin, such as bryophytes.

Eosin:

Eosin is available both in bluish and yellowish form. Yellowish eosin is more frequently used. It is a coal-tar dye, acidic and is a fluorine derivative. Solubility in water is 44.2% and in alcohol 2.18%. It is a valuable cytoplasmic stain, although not much used by botanists. Usually a 1% aqueous solution is used.

Light Green:

It is a coal-tar dye, acidic and belongs to diamino-triphenyl methane group. Solubility is 20.35% in water and 0.82% in alcohol. It is a very good stain for cytoplasm and cellulose walls. Staining occurs very rapidly. Irrespective of the solvent, a 0.2% to 0.5% solution is used.

Usually the dye is dissolved in 90% or 95% alcohol. Sometimes it is dissolved in absolute alcohol and diluted with clove oil. By combining light green with alcoholic Sudan IV, cutinized and suberized tissues can be differentiated from lignified tissues.

Safranin O:

It is a coal-tar dye, basic in reaction and belongs to the azin group. Its solubility is 5.45% in water and 2.41% in alcohol. It is a very important stain for botanists. It stains lignified, suberized, cutinized and chitinized structures as also nucleoli, chromosomes and centrosomes.

Various formulae are available for making safranin solution; but generally a 1% solution in 50% alcohol is good enough for most plant materials. A 1% aqueous solution can be used for temporary staining and for this special water solution safranin is available; because although it is soluble in both water and alcohol, it dissolves better in alcohol than in water. Other stains will be mentioned as and when required.

Precautions for Staining:

1. While staining free-hand section, take a few drops of the required reagent in a watch glass staining cube) and, after transferring the sections, always keep it covered by another watch glass.

2. Transfer the section from one reagent to another with a section lifting spatula or a scalpel. Use of brush for transferring sections is not advisable because it causes transference of the reagent too. However, a fine brush may be used, blotting it each time after use. Needles should not be used as they may prick the sections and damage them.

3. Dehydration is influenced by atmospheric humidity. Hence it often becomes difficult to com­pletely dehydrate sections during the rainy season. Fans should be switched-off during staining in order to reduce evaporation of alcohol during staining.

4. The watch glass should be placed on a piece of white paper while taking out sections from stain. They can be easily located against a white background.

5. If sections turn whitish on being transferred to xylol or balsam from absolute alcohol, it indicates incomplete dehydration. If staining is inadequate (either excess or less), it can be re-stained. In all such cases the sections should be passed down through the alcohol grade in the opposite order as it has come up.

6. While mounting the sections in Canada balsam or any other mounting medium, the cover glass should be dipped in xylol and then placed over the section gradually with the help of a needle.

If the balsam appears to be too thick, it should be diluted first with a little xylol. Ordinarily, balsam is neutral, and has a light straw colour. If the balsam is reddish, it indicates acidity. Sections should not be mounted in acidic balsam, because it fades the stain quickly.

7. After mounting, the sections should be kept on a hot plate (37° to 45°C) for at least 24 hours. It helps to drive away air bubbles and also dries the slides.

8. Clearing:

In many plant materials, various cell inclusions often interfere with staining and makes the stained preparation difficult to study. Such sections can be cleared by transferring them from 50% alcohol to a dish of water with a fine forceps or a brush.

Then, using a needle or forceps they are placed in a watch glass containing some sodium hypo-chlorate, household bleach or para-zone and kept for not more than 5 minutes. The whole section may dissolve if left long enough. Wash the sections thor­oughly in water after bleaching, don’t immerse the brush in the bleach, the bristles will dissolve.

Staining Schedules:

Two main types of stains are in use:

1. Temporary Stains – whose colour fades or which gradually damages the sections, and

2. Permanent Stains – whose colour lasts for many years.

1. Temporary Stains:

a. Methylene blue:

1% aqueous solution is used. The stain is usually mixed with glycerine, 10 ml of 1% aqueous stain is added to 90 ml of 50% glycerine. Sections are directly mounted into this medium. Macerated tissues can also be stained with this mixture.

A drop of washed macerate in water is mixed with a drop of the mixture on a slide and the cover glass is put on. All cell walls turn blue, except cutinized walls which remain unstained. The intensity of the blue colour depends upon the chemical and physical nature of the cell wall. Various wall layers stain differ­ently.

b. Chlor-zinc-iodine solution (Schult’s solution):

This solution consists of — Zinc chloride 30 g, potassium iodide 5 g, iodine crystals 1 g and distilled water 140 ml. Sections are placed on the slide and 1 or 2 drops of this solution are added. This is blotted off after 2-4 minutes and a drop of 50% glycerine is added. Alternatively, 50% glycerine is added directly and sections are mounted in the mixture.

Cellulose wall turns blue, starch turns blue-black, lignin and suberin turn yellow and moderately lignified wall turns greenish-blue. This stain swells the cell walls and eventually dissolves them. So, observations must be made immediately.

c. Chlorazol black:

A saturated solution of the dye in 70% alcohol is used. Cell walls turn black or grey. It is particularly good for showing pitting.

d. Phloroglucin and conc. HCl:

Add a drop of saturated aqueous solution of phloroglucin to the section taken on a slide and then the HCl. Some prefer to use 20% HCl. Lignin turns red.

e. Sudan IV:

Stain in a saturated alcoholic solution for about 10 minutes and wash rapidly in alcohol. Mount in 50% glycerine. As fat is soluble in alcohol, keep in alcohol no more than necessary. Sudan IV is a specific stain for fat which turns orange. Lecithin, latex, wax, cutin and resins also stain. Chloroplasts are stained a dull red.

f. Iodine with potassium iodide:

An aqueous solution of 1 % iodine and 1 % potassium iodide is used. The sections are stained for a few minutes and observed directly or after mounting in 50% glycerine. An aqueous 1% iodine solution may also be used. Starch turns blue, cellulose, inulin deposits, proteins and alkaloids turn brown and pectin, cutin, callose and cork turn yellow.

g. Eosin:

An 1% aqueous solution of eosin is sometimes used. It gives general yellowish colour to the tissue.

2. Permanent Stains:

Permanent staining is always accompanied by complete dehydration and clearing followed by mounting in Canada balsam or euparal etc. (For details see Cytology section).

Depending upon the number of stains used, it is called:

(a) Single staining,

(b) Double staining, etc.

(a) Single Staining:

Single staining is done generally in case of thallophytes and bryophytes where no lignified tissue is present and the cell wall material is mostly cellulose:

I. Heidenhain’s Iron Hematoxylin:

1. Transfer the sections from water to a mordant (a mordant is any substance that combines with and fines an dye-stuff in material that cannot be dyed direct) containing iron, aluminium or copper, because hematoxylin cannot stain tissues without mordanting with one of these three metals.

Ordinarily, ferric ammonium sulphate (iron alum) is used. The strength of the solution is usually 3% but varies from 2% to 4%; weaker solutions being more suitable for softer tissues such as algae. The solution should be freshly prepared before use.

A permanent mordanting mixture can be prepared by mixing 500 ml. distilled water, 5 ml. glacial acetic acid, 0.6 ml. C.P. sulphuric acid and 15 g of clear violet coloured alum crystals. The time required for mordanting should not exceed 2 hours and, with thin sections, 1 hour is sufficient.

2. Wash thoroughly in running water for 5 minutes and then rinse in distilled water to remove salts present in tap-water, which interfere with staining.

3. Stain in 0.5% aqueous Heidenhain’s hematoxylin solution. As a rule, the sections are left in the stain at least as long as they were left in the mordant; but, in most cases, 24 hours is the optimum time.

4. Wash off excess stain with water.

5. Destain in 2% ferric ammonium sulphate (or, ferric chloride). The time required for de-staining varies with the nature of the material, the thickness of sections, etc. Differentiate under a microscope. When the sections appear grayish-black, de-staining is complete.

6. Wash in running water for 30 minutes to 1 hour to remove all traces of the de-staining solution, which, if left, will gradually fade the stain.

7. Dehydrate in 50%, 70% and 95% alcohol, keeping at least 5 minutes in each. Give two changes in absolute alcohol at 5 minutes’ interval.

8. Keep in absolute alcohol-xylem mixture (1: 1) for 5 minutes.

9. Give two changes in xylol at 5 minutes’ interval.

10. Mount in Canada balsam and dry on a hot plate.

N. B.:

Although hematoxylin staining is rather cumbersome, the result is well worth it as different cellular components, including chromatin matter, take up various shades of stain.

II. Bismarck Brown Staining:

This is a suitable stain for soft and non-lignified plant tissues such as bryophytes and thallophytes.

The schedule is:

1. Take the sections from water and pass through 30% and 50% alcohol keeping 5 minutes in each. 30% alcohol may as well be omitted.

2. Stain the sections in Bismarck brown (1% solution in 70% ethanol) for 5 to 15 minutes. The duration of staining depends upon the concentration of the stain and the thickness of sections. If the stain appears to be relatively more concentrated, it may be diluted with a few drops of 70% alcohol. If the sections are rather thick, stain for a shorter period so that the stain remains light.

3. Pass through 80% (or 90%) and 95% alcohol keeping 5 minutes in each. Give two changes in absolute ethyl alcohol keeping at least 5 minutes in each.

4. Clear in xylol, preferably giving 2 changes of at least 5 minutes’ duration.

5. Mount in Canada balsam, Euparol or DPX mountant. All cell walls turn brown.

III. Light Green Staining:

Single staining can also be done using light green instead of Bismarck brown. Staining is done after dehydration in 90% alcohol as light green is dissolved in 95% alcohol (1% solution in 95% alcohol). The rest of the staining schedule is the same as in Bismarck brown staining.

IV. Ehrlieh’s Hematoxylin Staining:

1. Keep the sections in 30% alcohol for 5 minutes.

2. Stain in matured Ehrlieh’s hematoxylin solution for 5 to 30 minutes.

3. Wash out excess stain with 50% alcohol.

4. Dehydrate, clear and mount in Canada balsam.

This is an excellent stain for algae, fungi and small bryophytes.

(b) Double Staining:

Double staining is resorted to in case of all sections having both lignified and non-lignified tissues, i.e., in case of pteridophytes, gymnosperms and angiosperms. One of the two stains is specific for lignified tissues and the other stains the non-lignified tissues, i.e., mainly cellulose. The sections with two different stains show more contrast. There are numerous double staining schedules.

Some of these, which are usually followed in the class, are discussed:

I. Safranin and Delafield’s Hematoxylin Staining:

1. Freshly mix safranin (1% in 50% alcohol) and matured Delafield’s hematoxylin in the proportion of 1: 4. Filter the mixture. This stock mixture can be used up to a week, but should be filtered each time before use.

2. Stain the sections in this mixture for 2 to 6 hours. Some sections need less time. If it is more convenient to stain overnight, use safranin-hematoxylin mixture in the proportion of 94: 6.

3. Transfer the sections to 50% alcohol containing 2 to 3 drops of conc. HCl. This solution removes the stain, acting first on safranin. The duration of de-staining has to be learnt by experience. Generally, sections should be removed when they still appear slightly dark or over-stained under microscope.

4. Transfer the sections to 95% alcohol and keep for 5 minutes. This is followed by 5 minutes in absolute alcohol.

5. Keep the sections in alcohol-xylol mixture (1: 1) for 5 minutes.

6. Transfer to xylol and keep for 10 minutes.

7. Mount in Canada balsam and dry on a hot plate.

Lignin turns red, cellulose turns dark blue and cellulose walls with some lignin become purple.

II. Safranin and Fast Green Staining:

1. Keep the sections in 30% alcohol for 5 minutes.

2. Stain in a 1% solution of safranin in methyl cello solve 50% alcohol for 2 to 24 hours, or even 48 hours (Prepare safranin solution by dissolving 4 g of safranin in 100 ml each of 95% alcohol and distilled water followed by 4 g of sodium acetate and 8 ml of formalin. The acetate intensifies the stain and formalin acts as a mordant.).

3. Wash off the excess stain with running water for a few moments.

4. Differentiate and dehydrate with 95% alcohol to which 0.5% picric acid has been added. Ordi­narily, about 10 seconds of treatment is sufficient for differentiation.

5. Stop action of the acid by immersing the slide in 95% alcohol to which 4 to 5 drops of ammonia per 100 ml of alcohol has been added. This treatment should not be continued for more than 2 minutes, as alcohol extracts stain.

6. Dehydrate in absolute alcohol for 10 seconds.

7. Counterstain in fast green for not more than 15 seconds (Prepare a nearly saturated solution of fast green in equal parts of methyl cello solve and absolute alcohol and 75 parts clove oil.). This stain can be used repeatedly.

8. Pour the fast green stain back into dropping bottle and rinse off the excess stain with clove oil. Used clove oil may be used.

9. Clear in a mixture of 50 parts clove oil, 25 parts absolute alcohol and 25 parts xylol — for a few seconds (Actually this reagent mixture, after use, is collected in a bottle and used in step 8.).

10. Remove the clearing mixture by treating for a few seconds in xylol (Add 3 to 4 drops of absolute alcohol to this xylol to take care of any moisture that may be inadvertently brought over.).

11. Give two changes in pure xylol for at least 10 minutes’ interval and then mount in Canada balsam.

The safranin appears a brilliant red in chromosomes, nuclei and ii} lignified and cutinized cell walls; while the fast green gives a bright green colour to cellulose cell walls and cytoplasm. Both the colours are permanent and they persist for many years.

III. Safranin and Light Green Staining:

1. Keep the sections in 30% alcohol for 5 minutes.

2. Stain the sections in 1% solution of safranin in 50% alcohol for 30 minutes. The exact duration of staining depends upon the intensity of the dye solution. Some anatomists (Johansen etc.) prefer to make a 1% solution of the dye in 95% alcohol. A part of this stock solution is diluted with an equal volume of distilled water when needed for use. If this proves to be a strong solution, it may be further diluted with 50% alcohol.

3. Dehydrate in 70%, 80% and 90% alcohol, keeping 5 minutes in each.

4. Counterstain in 1% light green in 95% alcohol, for 15 seconds to 1 minute. The exact duration of staining is to be determined by experience. It depends on the intensity of the dye solution. If required, the staining solution may be further diluted by adding more 95% alcohol.

5. Dehydrate in absolute alcohol for 5 minutes.

6. Pass through alcohol xylol (1: 1) mixture keeping the sections in it for 5 minutes.

7. Clear in xylol for at least 10 minutes.

8. Mount in Canada balsam and dry on a hot plate.

Lignified walls turn red and cellulose walls green. Often the entire section turns green indicating over-staining in light green and under-staining in safranin. In such cases, the duration of safranin staining should be increased, while reducing the duration of light green staining.

The light green solution may also be further diluted. Sometimes, both the stains appear to be rather fade. To counteract this, the duration of treatment in different alcohol grades should be reduced. Although both the stains are permanent, light green appears to fade rather quickly.