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Preparing an anther squash

Class practical

This practical involves fine handling of plant material, and a complex multi-stage process of staining to prepare microscope slides. It also involves sophisticated interpretation of those slides. For students who have grasped the stages of meiosis, it would provide a valuable test of understanding. For students who are struggling to follow the main stages of the process, it may be confusing.

Lesson organisation

This will depend on the amount and type of anther material you have available, and the access you have to microscopes – with or without digital feeds. Preparing the slides is time-consuming, as is searching the slides to find cells at different stages.

Apparatus and Chemicals

For each group of students:

Access to stereomicroscopes (when working on the anthers)

Access to high power microscopes (when studying the slides – Note 1)

Suitable plant material as a source of anthers (Notes 2 and 3)

Microscope slides


Ethanoic alcohol to fix anther tissues (if using ethano-orcein stain – see Note 4)

Hydrochloric acid (Note 5)

Glycerol (if using toluidine blue stain – Note 6)

Stains (Note 7)

Health & Safety and Technical notes

1 Using the microscope: Lighting the slide is crucial, and students will need to use the condenser correctly, in order to achieve good contrast. Use of fine focus (and an oil immersion lens if available) will allow better interpretation.

2 In this practical, you are looking for cells that are currently manufacturing pollen. It is important to obtain very young anthers to study, in which the pollen mother cells on the inner wall of the anther are still active. If the anthers are yellow and obviously filled with pollen, they will be too mature for cell division to be taking place. Look for white, translucent anthers.

3 Some anthers do not take up the stain very readily, so it is wise to use a number of different sources of material. Generally, the smaller the number and the larger the size of chromosomes, the better the preparation and the easier it is to interpret the results. Possible sources of material are corms or bulbs (in late summer or early autumn – before sprouting) of Crocus, Narcissus, hyacinth, tulip and Lilium, or young buds of Pelargonium sp., Tradescantia sp., and Impatiens sp., or winter buds of Aesculus (horse chestnut) and Fraxinus (ash).

4 Ethanoic alcohol (“Farmer’s fluid”) is 3 parts absolute ethanol (highly flammable) to 1 part glacial ethanoic acid (Hazcard 38 A describes this as flammable and corrosive). Mix just before use, adding the acid to the alcohol. You can use IDA or 95% ethanol as a fix instead (Hazcard 40 A – highly flammable and harmful due to the presence of methanol), but chromosomes may not be as clearly defined. The minimum fixation time for developing anthers is 10 minutes.

5 Hydrochloric acid: Hazcard 47A describes this as corrosive or irritant at higher concentrations than 1.0 M, but it would be low hazard at this concentration, although eye protection is still recommended.

6 Glycerol: Hazcard 37 describes this as low hazard.

7 Stains:

Preparing ethano-orcein stain: Grind 1.5 g of solid orcein (described on Hazcard 32 as low hazard) with a pestle and mortar. In a fume cupboard, mix 90 cm3 of glacial ethanoic acid (Hazcard 38A describes this as flammable and corrosive) with 110 cm3 of distilled water and bring to the boil. Pour the boiling mixture over the orcein and stir very thoroughly (still in the fume cupboard). Leave overnight, then filter and store in a tightly-stoppered dark bottle. If it overstains, dilute with 45% ethanoic acid and try again.
Using ethano-orcein stain:

  • Put the fixed tissue into a watch-glass or small beaker and cover with a mixture of 9 parts ethano-orcein stain to 1 part of 1.0 M hydrochloric acid.
  • Warm to 60 °C gently using a thermostatically-controlled hotplate, or a spirit lamp (keeping the watch-glass just warm enough to be painful to touch on the back of the hand). Keep at 50 °C to 60 °C for ten minutes.
  • Add more drops of stain if evaporation dries out the preparation.
  • Transfer the stained tissue to a clean slide, using a mounted needle. Cut off and discard any tissue you do not need and wipe away any stain.
  • Add one drop of fresh orcein stain (with no hydrochloric acid).
  • Break up the tissue into very small pieces with a needle.
  • Cover with a coverslip and squash using Method A or Method B (see Investigation).

Preparing toluidine blue stain: Hazcard 32 describes this as low hazard. Dissolve 0.5 g of solid toluidine blue in 100 cm3 of water.

Using toluidine blue stain:

  • Place a small piece (2-3 mm) of the tissue to be stained on a clean slide.
  • Place three drops of 1.0 M hydrochloric acid on the tissue. Warm the slide very gently in a small Bunsen burner flame. Do not let the liquid boil or dry up. After about 1 minute, the pieces should appear fuzzy, showing that the cells are separating from each other.
  • Blot off the acid with filter paper or a paper tissue.
  • Add three drops of toluidine blue solution. Warm the slide as before for one minute.
  • Blot off the stain. Add one drop of fresh stain.
  • Mount the specimen by putting the stained tissue in the centre of the slide, and covering with a suitable-sized drop of mountant liquid (propane-1,2,3-triol (glycerol)), water or stain. Make sure the specimen is wetted and there are no visible air bubbles. Lower a coverslip slowly using a needle or forceps to leave a thin film of liquid over the specimen and no air bubbles.
  • Squash using Method A or Method B (see Investigation).


SAFETY: Take care with the acids and stains involved in preparing the slide. Avoid skin contact and wear eye protection as appropriate.


Location of anthers in a bulb

a Cut vertically down through the bulb just to one side of the apex.

b Discard the portion of the bulb that does not contain the bud; the bud is usually yellow-green and is in the centre of the bulb.

c Remove the white, fleshy leaf bases from around the bud and discard them.

d With a mounted needle, ‘unwrap’ the flower bud by peeling away the yellow leaves that tightly enclose it. Now pick away the petals of the flower until you find the anthers; they may be attached to tiny petals or standing free in the centre of the flower, around the style.

e Proceed to ‘Preparation of anthers’.

Location of anthers in an inflorescence

f Use a stereo microscope if the anthers are hard to see within the inflorescence.

g Select the very smallest single bud on the inflorescence, which will also be the youngest.

h Carefully hold the base or stalk of the bud with forceps and pick the green sepals off with a mounted needle.

i Remove the small white petals that are around the outside of the flower and expose the ring of anthers in the middle of the flower. In some flowers, the anthers may themselves be attached to very small petals; do not discard these by mistake.

Preparation of anthers for further investigation

j Anthers should appear white and translucent. If in doubt, try a smaller bud (Note 2). Leave them attached to the flower throughout the staining process as they are very easily lost.

k If using ethano-orcein stain, fix in ethanoic alcohol (Note 4).

l Stain using one of the techniques in Note 7.

m Detach two or three anthers and transfer to a clean slide. Use a small amount of material to ensure cells are well-separated by the squashing process.

n Break up with a mounted needle and discard the filament and any other relatively unstained portions.

o Cover with a coverslip and squash using method A or method B below.

Method A  Place the slide and coverslip on a double layer of paper towel and fold the paper over the coverslip. Make certain that the slide is on a flat surface and squash down on the coverslip with a strong vertical pressure, using your thumb. Do not twist or roll the thumb from side to side.

Method B  Tap the coverslip about 20 times by dropping a wooden mounted needle or a pencil, blunt end down, from a height of about 5 cm onto the middle of the coverslip.

p Search the slide and, at first, try to get a general idea of the apparently different types of cell that you can see. There may well be mature or maturing pollen grains and cells showing almost homogeneous nuclei, as well as cells clearly undergoing division.

q In the cells that are actively dividing, the nuclei may show obvious thread-like structures (chromosomes). Damage to the nucleus or incomplete staining may have occurred. Compare several cells that seem to be at different stages of cell division. Sometimes cells break up and you may see free chromosomes in the field of view.

r Draw three or four different cells each showing some nuclear structure, but at different stages. If your microscope has a digital image feed, you could store some digital images instead. Focus carefully up and down with the fine focus as you draw; try to see the whole length of the chromosomes.

Teaching notes

Pollen is manufactured in the anthers within the flowers of most angiosperms. The anthers form even while the flower is still a very young bud; they produce pollen grains in the locules of the anther by the time a clearly recognisable flower bud or flower has grown. Students will need a basic understanding of flower structure to complete this practical work.

The manufacture of pollen by an angiosperm requires meiosis to occur, producing haploid nuclei. If an anther contains pollen, it is usually possible to squeeze the loose grains out of the loculi during the staining process, so that a squash of the anther wall produces gametogenic cells.

The act of drawing a nucleus, rather than just looking, often encourages more careful interpretation of the stage it has reached.

Mature pollen grains have a more dense cytoplasm than meiotic cells. The volume of cytoplasm in the maturing pollen grain relative to the volume of nucleus is greater than a meiotic cell. The cell wall of the maturing pollen grain is somewhat thicker than that of meiotic cells.

Patterns of chromosome alignment or distribution within the nuclei are repeated in many cells. The existence of recognizable patterns suggests that an ordered process of chromosome distribution is taking place. This could be confirmed by observation of living cells using phase contrast and other illumination techniques.

The use of a light microscope places limits on the resolution of images seen. It may be difficult for inexperienced students to find nuclei and confirm with certainty that meiosis is taking place. Configurations of first prophase, showing chiasmata, may be identified and are diagnostic – these will never be found in (for example) root tip cells dividing by mitosis. An occasional sighting of a tetrad of cells with nuclei clearly at second telophase would also be diagnostic. In these preparations, there are usually a large number of stages of the division where chromosomes are clearly ‘doubled’; this contrasts nicely with mitotic preparations where is it usually hard to see double strands at any stage.

Health & Safety checked, September 2009

Related experiments

Investigating mitosis in a root tip squash


Page last updated on 24 November 2011