Botanical Microtechnique Part 2. Staining Botanical Sections.

by Jim Battersby, UK

  

 Click here for part 1 of the series.

Botanical Microtechnique Part 2

Staining Botanical Sections

We will assume that sections have been cut from the embedded blocks on a microtome, and are now ready for attaching to slides and staining. As stated in part one, there are lots of directions and hints on using microtomes in the literature (i.e. NBS booklet 15), and the purpose of these little monographs is to bring together some of the techniques which have been updated.

There are a huge number of staining techniques for botanical sections, but I will only look at two more general and widely used methods, Safranin & Alcian Blue and Safranin & Fast Green.

Adhering sections to slides

Once the sections have been cut they should be adhered to the slide using a section adhesive. Traditionally, many of the adhesive formulas use dilute formaldehyde solution to float and flatten the section, these work very well but need to be used in a well ventilated area - preferably near an extractor fan. The first formula - a variation on Haupt's adhesive was sent to me by Colin J Kirk; the second one from Chamberlain (1932) does not use formalin and the sections are floated out in water.

  1. Haupt's gelatine fixative

5g Glycerine Jelly

45mls Water

Warm and stir until the jelly has dissolved, then allow to cool.

Having smeared a thin film adhesive on the slide and float sections on 2% formalin (2.5ml commercial formaldehyde in 97.5ml water). Warm gently until section flattens (do not overheat or this will melt the wax and ruin the section), then blot carefully with damp filter paper. Dry at 37C as long as possible - 30 mins minimum.

2. Land's gum fixative

1g Gum Arabic

1g Potassium Dichromate

98ml Water

Mix the gum with a little of the (cold) water into a paste; dissolve the pot. dichromate in the remaining (warm) water; then mix together warm and stir until dissolved, then allow to cool.

Smear a thin film of the adhesive on the slide and follow the above method but using water instead of formalin.

Alternatively, in this latter method, the sections can floated out on warm water (about 45C) until they are flattened out, and are then 'picked up' directly onto on a slide which has been thinly smeared with the section adhesive. Place the slide into the water under the section and lift out horizontally, centre the section on the slide on the black centring guide, carefully dab away excess water with filter paper, and leave to dry at about 35C to 40C overnight. An ideal 'floating out' bath is a non-stick baking tin. This can be kept at approx 45C on a home made hotplate, or photographic developer warming plate. Blotting & drying section as above.

N.B. It is important to keep the sections dust free whilst they are drying.

 

Dewaxing

The slides first have to dewaxed. Traditionally, this is done by immersing in two changes of xylene (not less than 10 minutes in each), however Hystoclear is a new (and safer) alternative to xylene and is used in the same way.

Safranain / Alcian Blue

This method was first developed by John H. Nicholls who I had the pleasure of meeting over 20 years ago, and was published in the Balsam Post (October 2001 etc.) by Charles Bailey. The following updated schedule was written by Colin J. Kirk who was especially patient in helping me to clarifying some of the stages.

1% Alcian Blue 1g Alcian blue (Aq)

2.5ml Acetic acid (glacial)

0.2ml Formalin 40%

to 100 ml Water (deionised)

Premix the acetic acid - formaldehyde solution in the water. Add the Alcian Blue with stirring. Stir until homogenous.

1% Safranin O 1g Safranin O

100ml 50% isopropyl alcohol

Staining Schedule

Bring sections to water. (After dewaxing, place slides in two changes of absolute alcohol (5mins each), followed by 5 minutes each in 75%, 50%, 25% alcohol, and finally 1 minute in water.)

Stain for 30 minutes in the Alcian blue

Wash in water to remove excess stain 3 or 4 changes of water in a coplin jar OR 10 - 15 secs in gentle running water.

Stain for 10 mins in 1% Safranin solution

Wash in water as above.

Differentiate in 50% isopropyl alcohol until the section shows a clear blue in parts

Dehydrate 100% alcohol I (dip and reverse)

Dehydrate 100% alcohol II Dehydrate 100% alcohol I (dip and reverse)

Clear in Xylene I Dehydrate 100% alcohol I (dip and reverse)

Clear in Xylene II Dehydrate 100% alcohol I (dip and reverse)

Mount in Practamount or Canada Balsam

 

 

Safranin O / Fast Green FCF

The following schedule is based on Eric Marson's (NBS Booklet 17), amended and updated:

1% Safranin in Cellosolve alcohol 1g Safranin O, alcoholic

50ml Cellosolve

25ml 95% alcohol

25ml Purified water

1g Sodium Acetate

2ml Formalin

Premix the cellosolve and alcohol and stir in the Safranin O until dissolved.

Add the Formalin to the water and dissolve in the sodium acetate then mix the two solutions.

Fast Green FCF in Cellosolve alcohol 0.2g Safranin O, alcoholic

50ml Cellosolve

50ml 95% alcohol

Other regents required: 1% Picric acid solution in water

1% Lithium carbonate solution in water

OR

100mls 50% alcohol with a few drops of ammonia

Staining Schedule

Bring sections to 75% alcohol (After dewaxing, place slides in two changes of 100% alcohol (5mins each), followed by 5 minutes in 75% alcohol).

Stain for 1 hour to 24 hours Safranin O

Rinse in water to remove excess stain

Rinse in 1% Picric acid solution (dip and reverse)

Rinse in 1% Lithium carbonate solution (dip and reverse) OR 50% alcohol to which a couple of drops of ammonia have been added (dip and reverse)

Rinse in 75% alcohol (dip and reverse)

Rinse in 100% alcohol (dip and reverse)

Stain in Fast Green FCF (dip and reverse)

Dehydrate 100% alcohol I (dip and reverse)

Dehydrate 100% alcohol II Dehydrate 100% alcohol I (dip and reverse)

Clear in Xylene I Dehydrate 100% alcohol I (dip and reverse)

Clear in Xylene II Dehydrate 100% alcohol I (dip and reverse)

Mount in Practamount or Canada Balsam

HINTS & TIPS

175mm wide mouth bottles are ideal for storing stains and reagents, and for the 'dip and reverse' stages above, although staining jars should be used where the slides are kept in the stain for any length of time. Use tweezers to move slides from one bottle to the next.

Wear disposable gloves during staining, except for using xylene when Nitrile gloves should be worn.

Wife off excess xylene with paper towels (do not touch the specimen) before mounting. The paper should be well whetted and sealed in a plastic bag before disposal. Remember xylene fumes are toxic and highly flammable - take sensible precautions.

 

Bibliography and further reading

Chamberlain, C.J. Methods in Plant Histology University of Chicago Press (4thEd. 1932)

Out of print, usually available from http://www.abebooks.com/

Gurr, E. The Rational Use of Dyes in Biology London: Leonard Hill, 1965

An excellent reference book covering a huge range of staining techniques. Out of print, sometimes available at: http://www.abebooks.com/

Johansen, D.A. Plant Microtechnique New York: McGraw Hill, 1940

Superb book, the plant histologist's bible! Again, out of print, often available at: http://www.abebooks.com/ Highly recommended.

Marson, J.E. Practical Microscopy Ipswich: N.B.S., 1983

Another excellent reference book covering a huge range of techniques for the practical microscopist, including making and using hotplates for wax imbedding, using hand and mechanical microtomes for sectioning wax blocks etc. 5 from:

http://www.savonabooks.free-online.co.uk/ Highly recommended.

Ruzin, S Plant Microtechnique and Microscopy  Oxford University Press Inc, USA 1999

A superb modern reference book, full of practical information, well written and designed, but of limited use to the amateur microscopist. 42.50 from: http://www.amazon.co.uk

With special thanks to:

Charles Bailey, Colin J.Kirk , Eric Marsden, John Nicholls, Stephen Ruzin and Alan Taylor,

who were all both patient and helpful with my questions.

Further information:

Health & Safety data on individual chemicals & compounds can be found at http://ptcl.chem.ox.ac.uk/MSDS/

Alternative chemical names:

Formaldehyde, Formalin (app.37% formaldehyde solution).

IMS, Industrial Methylated Spirit

Propionic acid, Propanoic acid.

Iso-Propyl Alcohol, IPA, Propan-2-ol, 2-propanol,

Cellosolve 2-Ethoxyethanol

See also http://www.marigoldindustrial.com/charts/names/names.html

If any readers have difficulty if finding supplies of any of the above or Practamount mountant, or have any questions of comments please contact the writer.

All comments to the author Jim Battersby are welcomed.

 

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