BioImages: The Virtual Field-Guide (UK)

MAGNOLIOPSIDA (flowering plants)

Subtaxa (ie subgroups of this Class)

ACORALES (an order of flowering plants) Order 2 subtaxa  
ACORACEAE (sweet flags) Family 2 subtaxa  
Acorus calamus L. (Sweet-flag)
Plants at riverside
Species   1 ident refs
Acorus gramineus Aiton (Slender Sweet-flag) Species   1 ident refs
EU-DICOTS (dicotyledonous flowering plants)
Female flowers - close-up Flower Flower Flower - close-up Flower - female - side view Flower - side view Flower - side view Flower bud Flower bud - top view - close-up Flower head - close-up Flowers Flowers - in situ - top view Foliage Foliage Foliage - just coming into leaf and flower Inflorescence Leaf spray Leaf spray - with fruiting catkin Leaf spray - with fruiting catkin Leaf spray - with fruiting catkin Leaf spray - with fruiting catkin Partial umbel - underside showing bracteoles Plant Plant Plant Plant Plant Plant Plant - in cloud Plant - in situ Plant - in situ Plant - in situ Plant - in situ Plant - in situ Plant - in situ Plant - in situ Plant - in situ Plant - side view Shoot tip Snow-covered hedge Weather-damaged leaves - under surfaces
Subclass 1101 subtaxa 417 ident refs
LILIIDAE (monocotyledonous flowering plants)
Cut end of log showing thickness of Inflorescence - top view Plant Plant Plant Plant - in greenhouse display Plant - in situ
Subclass 337 subtaxa 236 ident refs
PRE-DICOTS (primitive angiosperms)
Flower and leaf - close-up Foliage and flower bud - close-up
Subclass 5 subtaxa 6 ident refs
(Climbing plants) (climbing plants)
Close-up Flower - face view Flower - female - side view Flower buds - just bursting Flowers Plant - in situ in hedge
Informal 15 subtaxa 2 ident refs
(Succulent plants) (succulent plants)
Plant - in situ
Informal 9 subtaxa 6 ident refs
(Tall Woody Herbs) (tall woody herbs)
Capitulum showing pair of bands of flowers moving apart as flowers open Cauline leaf Flower - side view Flower - two petals removed Flower buds - top view Inflorescence - top view - close-up Leaf base - top view Leaves - basal Partial umbel - underside showing bracteoles Plant Plant Plant Plant Plant - in cloud Plant - in situ Plant - part Plant - side view Plant -top view
Informal 156 subtaxa 90 ident refs

Suggested Literature

Identification Works

British Wild Flowers: British Wild Flowers
Irish Wildflowers: Irish Wildflowers
Wild Flowers of the British Isles: Wild Flowers of the British Isles
Crawley, M., 2005 The Flora of Berkshire
Floral Images: Floral Images
Wild Flower Finder: Wild Flower Finder
Fisher, J., 1991 (Rare species) A Colour Guide to Rare Wild Flowers
Garrard I. & Streeter D.T., 1998 The Wild Flowers of the British Isles
Find Wild Flowers: Find Wild Flowers
Hayward, J., 1995 A New Key to Wild Flowers
King, A., 1957 (Garden flowers) The Observer's Book of Garden Flowers
Wild Plants of the British Isles: Wild Plants of the British Isles
Poland, J. & Clement, E., 2009 The Vegetative Key to the British Flora
Rose, F. & O'Reilly, C., 2006 The Wild Flower Key - How to identify wild flowers, trees and shrubs in Britain and Ireland
Rose, F., 1981 The Wild Flower Key - British Isles - N.W. Europe
Stace, C., 2010 New Flora of the British Isles (Ed 3)
Stokoe, W.J. The Observer's Book of Wild Flowers
Leif & Anita Stridvall's Botanical Site: Leif & Anita Stridvall's Botanical Site
Cercle de Mycologie de Mons (Belgique): (Page perso de JJ. Wuilbaut): Cercle de Mycologie de Mons (Belgique): (Page perso de JJ. Wuilbaut)

Arable Plants

Francis, S.A., 2009 British Field Crops: a pocket guide to the identification, history and uses of arable crops in Great Britain
Wilson, P. & King, M., 2003 Arable Plants - a Field Guide


Fitch, W.H., Smith, W.G., et al, 1924 Illustrations of the British flora
Ross-Craig, S., 1948 Drawings of British Plants being Illustrations of the Species of Flowering Plants Growing naturally in the British Isles

Garden plants

Univerität Karlsruhe (TH) Botanischer Garten: Univerität Karlsruhe (TH) Botanischer Garten


Nelson, E.C., 2000 Sea Beans & Nickar Nuts
Trewella, S. & Hatcher, J., 2015 The Essential Guide to Beachcombing and the Strandline


Atlas Hymenoptera - pollens: Atlas Hymenoptera - pollens
Sawyer, R., 1981 Pollen Identification for Beekeepers


Bebbington, A. & J., 1996 Describing Flowers

BioInfo BioInfo ( has 428 general literature references to MAGNOLIOPSIDA (flowering plants)

MAGNOLIOPSIDA may also be covered by literature listed under:

(living things)
(vascular plants)

BioInfo BioInfo ( has 38064 feeding and other relationships of MAGNOLIOPSIDA (flowering plants)

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.


• 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.


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.


Thanks to David Rennison for helpful advice and discussions.


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.)

Brunel Microscopes,
Northern Bee Books,
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