BioImages: The Virtual Field-Guide (UK)

PINOPSIDA (conifers)

Subtaxa (ie subgroups of this Class)

Suggested Literature

Identification Works

Auders, A.G., 2013 RHS Encyclopedia of Conifers
Mitchell, A.F., 1972 Conifers in the British Isles: a descriptive handbook
Conifers Workshop (Hampshire Flora Group): http://hantsplants.org.uk/articles.php Conifers Workshop (Hampshire Flora Group)
Sell, P.D., 2012 Conifers

BioInfo BioInfo (www.bioinfo.org.uk) has 31 general literature references to PINOPSIDA (conifers)

PINOPSIDA may also be covered by literature listed under:

BIOTA
(living things)
Eukaryota
(eukaryotes)
PLANTAE
(plants)
TRACHEOPHYTA
(vascular plants)
Trees Trees and shrubs Trees, shrubs, woody and climbing plants (via Pinopsida)

BioInfo BioInfo (www.bioinfo.org.uk) has 4036 feeding and other relationships of PINOPSIDA (conifers)

Further Information

Lab. techniques Examining and Identifying Pollen

The following account describes a simple technique for collecting pollen from flowers and preparing it for microscopic examination. It is based on Appendix D of White, 1999.

Materials:

• Glycerine Jelly
• Safranin in cellosolve [poisonous - do not swallow]
• Alcohol: preferably isopropanol (=isopropyl alcohol)
• Water

Only small bottles of the above chemicals are required.

The alcohol is used to de-wax the pollen. Isopropanol is preferred to ethanol (ethyl alcohol) as the latter is said to cause permanent contraction of the cytoplasm. Ether (diethyl ether) is even better but highly flammable and evaporates so readily that it's difficult to use and store; it's also much harder to obtain.

• Laundry marker for labelling slides and pipettes
• Clean glass microscope slides (say 10)
• Coverslips
• A slide box to hold the slides vertically, so that the surfaces do not touch.
• A few disposable plastic pipettes.

Obviously you'll need a compound ("slide") microscope, ideally with a x100 oil-immersion objective. A dissection microscope is also useful to check the progress of staining but has insufficient magnification to identify pollen.

Disposable pipettes cost a few pence each. If you plan to reuse them, label with the reagent used - a laundry marker is ideal. If used carefully to avoid contamination they last for years.

Preparation:

The first thing is to prepare the slides for collecting pollen. We'll prepare glycerine jelly smears which are used to capture a thin layer of pollen.

Glycerine jelly smears with pollen on all look the same, so label the slides before you start! Either stick on a blank slide label or write a number in the top left corner with the laundry pen. The other reason for labelling is that it is quite difficult to see which side of the slide the smear is on, and only too easy to smear the pollen on the wrong side - it is very annoying to watch all your pollen wash away as soon as you start the prep.! Put the labelled slides in the box.

Warm the glycerine jelly by placing the bottle in a bath of hot water. (An old single portion beans can with the lid cut off is ideal.) When the jelly has melted, dip in a clean fingertip and make a large smear on a clean slide. Give it a few minutes to set, then wash and dry your finger and transfer a smear from this smear to the other slides (a smear from a smear). I do two fingerprint-sized smears on each slide, to give me two chances.

The master slide is reusable for another batch of smears so put that in the box too.

The smears will remain tacky for two to three weeks, depending on temperature, but they will last a whole summer if the box of slides is stored in a re-sealable polythene bag in the fridge.

Collection of pollen from flowers:

Collection of pollen is as simple as touching the anthers a few times against the smear. For many flowers this can be done in the field, although it may be necessary to remove a few petals. A pair of fine forceps is useful for this and for very small flowers which can be plucked and touched against the smear. Don’t worry about getting too much pollen; unless the smear is very thick, only a monolayer will stick.

Sampling of pollen from insects:

You could use the same technique to sample pollen load — even from the living insect, although the sample would be biased towards the pollen most recently collected. Or you could manually transfer the pollen onto the slide. A mounted needle dipped in glycerine jelly would even enable you to sample the pollen from specific hairs (taking care not to contaminate your glycerine jelly).

Pollen may be extracted from honey by diluting in warm water, then filtering. A similar technique would give an unbiased sample from pollen load. It may help to first wet it with a little alcohol.

Examining the pollen:

It’s useful to have some 2" squares of paper tissue to hand (paper tissue cut into squares is fine, paper serviettes are better, loo paper is too dusty) to soak up fluids, wipe spills etc. You'll also need a receptacle for waste tissue and to catch the washings; an old margarine carton is fine. Lab coat or old clothes are also recommended and you may want to protect the carpet.

With a fine pipette, drip a few drops of isopropyl alcohol onto the smear and let them run off. Make sure the whole smear is wetted. This only takes a few moments and dewaxes the pollen. (This is when you find out if you put the pollen on the wrong side of the slide!) Soak up the excess with paper tissue, taking care not to touch the smear itself.

Using a glass or smooth metal rod or fine pipette, add a drop of Safranin in cellosolve/alcohol. It will initially form a discrete drop, but this will soon creep over the smear. Tilt the slide to help it run in the right direction. Leave for a few seconds then gently wash off with a few drips of water. Again, soak up the excess with paper tissue. Check the pollen under a dissection microscope: it need only be slightly coloured - enough to see it. Add another droplet of stain if necessary and repat. Don't overstain as this obscures detail. Some pollens (eg umbellifers, pink family) stain more easily than others (eg borage family).

When sufficiently stained, gently lower a coverslip onto your prep. and examine immediately. Don’t let the pollen dry out. The x10 objective is usually appropriate for gross structure, with x40 for surface detail; only the very smallest pollens (forget-me-nots!) require oil-immersion (x100). A green filter often makes the red-stained details stand out better, especially for photography. Try to avoid sliding the coverslip or crashing into it with the objective as this may cause the grains to clump or burst.

"Glycerine Jelly for Pollen" contains basic fuchsin instead of safranin but gives similar results.

White, 1999, describes how to prepare permanent mounts using aqueous mountant or glycerine jelly.

Sawyer, 1981, and the related CD enable identification of pollens from most common flowers.

A word of warning: safranin, like most microscopical stains, will stain most things it comes into contact with including fingers, worksurfaces and sinks. Glazed or stainless steel should be OK, but modern polymer or geological materials could be permanently marked.

Finally, pollen is the most efficient contaminant known to science! Work clean. Keep slide boxes and chemical bottles closed when not in use. Wash slides and coverslips thoroughly before re-use.

Acknowledgments:

Thanks to David Rennison for helpful advice and discussions.

References:

Sawyer, R., 1981, Pollen Identification for Beekeepers, Cardiff Academic Press, ISBN 0 906449 29 4
(There is also an illustrated CD related to this. Both are available from Northern Bee Books)

White, J., 1999, Pollen, its Collection and Preparation for the Microscope, NBS
(available from Brunel Microscopes.)

Suppliers:
Brunel Microscopes, http://www.brunelmicroscopes.co.uk/
Northern Bee Books, http://www.beedata.com/beebooks.htm
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