The wrong kind of green?


Back in 2016 I wrote a post entitled “The camera never lies?” which mused on the difficulties in interpreting photographs of stream beds taken with waterproof cameras.  My point was that quantities of algae can be highly variable and interpretation of conditions from a single photograph is fraught with difficulties.   Because waterproof cameras are now relatively cheap it is easy to take a photograph whilst out doing fieldwork and then include this in a report, making a point, perhaps, about the poor condition of a site.  However, the frequency of visits to a site is often such that the person who took the photograph has little idea of whether that growth is persistent or not.   The photograph at the top of the post (from the River Irt, Cumbria, just below Wastwater) shows just how variable the cover can be at a single site over the course of a year.

The question is still pertinent because, last year, an article appeared in FBA News suggesting that percent benthic algal cover, measured from bed photographs, was correlated with the level of phosphorus enrichment.   The example from the River Irt challenges this: the cover recorded here (average: 17%, maximum: 36%, based on direct estimates of the percent of the stream bed covered with algae, rather than via photographs) suggests quite a high level of enrichment, whereas the Irt is, in fact, just downstream of one of the most oligotrophic lakes in the country.   There are two plausible reasons for this: first, my experience of West Cumbrian rivers suggests that regulation can have a big effect on the quantity of filamentous algae present regardless of the amount of nutrients and, second, the article makes no differentiation between “good” algae and “bad” algae.   Not all algae are indicators of nutrient enrichment, so you really need to know the identity of the algae before leaping to a conclusion (see “The democratisation of stream ecology?” for one way of addressing this.

A further issue is illustrated by the second image – this time, showing the bed of the River Ehen, about 15 kilometres north of the River Irt and, once again, downstream from an oligotrophic lake.  Here, the total cover is even higher (average: 46%, maximum: 87%) but green algae represent a relatively small proportion of this (typically about a quarter).  The FBA News article makes little reference to the type of algae, just referring to “benthic algal growth”. Again, identity is important: the Ehen site has a lot of red algae and Cyanobacteria as well as green algae and, again, these are not necessarily indicators of enrichment.   Copious Lemanea fluviatilis in the Spring is, in fact, usually a sign of healthy conditions.


Seasonal variation in the appearance of the bed of the River Ehen, Cumbria.   Photographs taken at approximately the same spot at two monthly intervals.  The photograph at the top of the post shows variation in the River Irt, also in Cumbria.

One extra twist to the story comes when we compare visual estimates of cover with the biomass, as measured using a BenthoTorch (see “The complexities of measuring mass”).   Both show a trend of increased measured biomass linked to increased observed cover, but the relationships are very noisy.  Of particular interest is the row of points bouncing along the x axis, indicating visits where the eye saw lots of green algae but the BenthoTorch measured very little.  What I think is happening is that the green algae can form bright but relatively thin layers on the rocks on these occasions.  On other occasions, the measured biomass is much higher but if the green algae are covered with epiphytic diatoms, then they may not be so obvious to the naked eye, possibly leading to percent cover being under-estimated during surveys.   A final possibility, certainly plausible in the Irt, is that the green algal growths are associated with boulders whereas our biomass measurements are made on moveable cobbles.


Measuring biomass with a BenthoTorch in West Cumbrian streams.

There are, in other words, good reasons for the mismatch between observed and measured quantities of algae.  Neither is a perfect indicator of what is going on in a river.  If we are interested in the effect of algae on ecosystems, then measured biomass would seem to be the better indicator of the extent to which other ecological processes in a stream are likely to be affected by the algae that are present.  However, the general patchiness of algal growths, plus the relative ease with which visual estimates can be made probably outweighs their problems, at least from the point of view of basic assessments of ecosystem health.


The relationship between visual estimates of cover (x axis) and biomass measured using a BenthoTorch; a. all algae (Spearman’s rank correlation, r = 0.55); b. just green algae (r = 0.65).  Data from West Cumbrian lakes and streams, 2019-20.

There are, in other words, good reasons for the mismatch between observed and measured quantities of algae.  Neither is a perfect indicator of what is going on in a river.  If we are interested in the effect of algae on ecosystems, then measured biomass would seem to be the better indicator of the extent to which other ecological processes in a stream are likely to be affected by the algae that are present.  However, the general patchiness of algal growths, plus the relative ease with which visual estimates can be made probably outweighs their problems, at least from the point of view of basic assessments of ecosystem health.   Following on from this, I doubt that a photograph offers any  advantage over an inspection of the river bed for basic ecological health assessments.   You need to explore a reasonable length of river, including variations in velocity, substratum type and shade, to get some idea both of the quantity of algae, and its variability.

After a quarter of a century spent contemplating the relationship between algae and nutrients in rivers, however, I’m reluctant to endorse any system that implies that a “green” riverbed equates to poor ecosystem health based on a single visit.   These proliferations are, to use a human health analogy, the sniffles and colds of river ecosystems: most rivers suffer from them from time to time and most recover quite quickly.  The concern is when rivers are persistently green, as this suggests a long-term breakdown in the links between trophic levels.  The first response on seeing a green riverbed, in other words, should be to schedule a follow-up visit a month later to see if the situation persists rather than leaping to a rash conclusion.   The second response should be to work out which alga is responsible for the bloom as this, too, can help with subsequent diagnoses.

On a more positive note, anything that makes people notice and record algae in rivers has got to be a good thing.  Just don’t be seduced by any simplistic correlations that assume that algal growths are, by definition, a bad thing.  Algae are key components of the ecological engine of rivers and if that is not part of the thinking behind any river health assessment, then treat it with extreme caution.


Everall, N.C., Johnson, M.F., Clarke, A. & Gray, J. (2019).  The visual state of river beds and their associated invertebrate community biosignatures.  FBA News 77: 11-15.

Some other highlights from this week:

Wrote this whilst listening to:  early Fleetwood Mac, as I commemorated the death of Peter Green last weekend.

Cultural highlights:  Once Upon A Time In Iraq, on the BBC iPlayer, is a gruelling exposé of Western policy blunders and ideological naivity that brought the Middle East to its present dire predicament.

Currently reading:  Watching the English by Kate Fox.   Cultural anthropology on our own doorstep.

Culinary highlight:  Red Dragon (Aduki Bean) Pie – one of the best veggie alternatives to a shepherd’s pie that I have come across.  Recommended by Julie Gething.

Hilda Canter-Lund competition winners, 2020


The winner of this year’s Hilda Canter-Lund competition for the best algal-themed photograph is Davis Laundon of the Marine Biological Association in Plymouth for his photograph The Phycosphere.   Davis is a particularly appropriate winner of the competition as he, like Hilda Canter-Lund, has made a special study of chytrids, a group of simple fungi that were also the focus of Hilda Canter-Lund’s own research.  His current research at the Marine Biological Association looks at how thraustochytrids (fungus-like protists that, despite their name, are actually quite distant relations of true chytrids) interact with marine diatoms.

Davis’ picture, The Phycosphere shows a planktonic diatom Coscinodiscus photographed using confocal microscopy and fluorescence lighting.   A layperson looking at this image will see a pleasing circular semi-abstract composition against a dark backdrop and will probably be surprised if you told them that they were actually looking at living organisms.   Even if Davis had used straightforward brightfield lighting, then the yellow-brown colour of diatom chloroplasts might still have made it hard for a layperson to place as a distinct life-form.  Someone who knows about plankton, on the other hand, would have been able to recognise it as a centric diatom, and maybe name the genus.  The use of fluorescence lighting lifts the image in two ways: first, by making it more visually interesting, whether to a layperson or informed biologist, and second, because despite the artificial colours, it gives us more insights into the true nature of the organism than are possible with “ordinary” light.

Of particular interest, in this case, is the sprinkling of blue dots around the perimeter of the cell.  These are bacteria living in the thin mucus layer that surrounds every diatom cell.   Look back at the many images of live diatoms I’ve included in this blog and you will never see these diatoms but they are, undoubtably present.  They are, however, very small and difficult to discern with brightfield microscopy.  Only by adding a special stain that binds to DNA, then shining light of a particular wavelength through the microscope do we realise that it is there, as the stained DNA now fluoresces with a bright blue colour.


Michiel Vos:Carpodesmia tamariscifolia (Bushy Rainbow Wrack) framed by Himanthalia elongata (Thong  Weed) in a rockpool in Falmouth, Cornwall, U.K.   The picture frame encloses a depth of about a metre.  Davis Laundon’s The Phycosphere is at the top of the post.   The diatom is about a tenth of a millimetre in diameter (actually about 90 micrometres).

We have to go a little further west from Plymouth to meet our second prize winner, Michiel Vos.  Michiel (“Mick”) is originally from the Netherlands but is now a senior lecturer in microbiology at Exeter University’s Falmouth Campus.  His day-job doesn’t actually involve the study of algae, but he spends as much of his spare time as possible dabbling around the rock pools of the Cornish coast.  This particular image is a magical view of algae that could be the CGIed backdrop to a sequence in a fantasy movie.   The rainbow wrack is a perennial that forms a home to many animals, from sponges to tunicates.  The Bull Huss often attaches its egg cases to this alga and, in addition, many seaweed species grow epiphytically on it.   As for Davis’ image, we have an instantaneous hit of an intriguing image which then provides an entrée to a more sophisticated reading that reminds us of the interplay between organisms in the natural world.    Mick also writes a blog, An Bollenesor (“the rockpool hunter” in the Cornish language) is well worth a visit and contains much more of his stunning photography.

Our winners encapsulate the tensions encountered when photographing algae: Davis leans towards abstraction whilst Mick goes for a depiction of “other worlds” portrayal (see “How to win the Hilda Canter-Lund competition (2)”).   Davis offers us a very tech-heavy image whilst Mick leans towards the “decisive moment” (see “How to win the Hilda Canter-Lund competition”).   Both, however, fulfil the competition’s brief of celebrating the hidden world of this fascinating group of organisms.


Some other highlights from this week:

Wrote this whilst listening to:   The National and Bob Dylan’s new album Rough and Rowdy Ways

Cultural highlights:  Fanny Lye Deliver’d, a sometimes brutal portrayal of the religious turmoil in the aftermath of the English Civil War.   Ranters versus Puritans.

Currently reading:   Bill Bryson’s Down Under

Culinary highlight:  When I’m at a festival I seek out the stall selling Goan fish curry, and this week Felicity Cloake’s column in The Guardian described how to cook the perfect Goan fish curry.  To be honest, I haven’t eaten it yet, but the smell as it was cooking was quite something..

How not to win the Hilda Canter-Lund competition


The 2020 Hilda Canter-Lund competition for the best photograph of an alga is underway again, with a closing date of Friday 5 June.  Over the years I’ve written a few posts to encourage entries, by focussing on what makes a good entry for the competition (listed at the end of this post).  This time, however, I’m coming at the problem from a different angle because, each year, as we make our first review of entries in order to prepare a shortlist, the judges always reluctantly leave one or two images out due to fairly basic flaws that could have been corrected prior to submission.   At least two of our winners have used smartphones for their photos and even these now have basic editing capabilities, so there really is no excuse for a little cropping or tonal adjustment prior to submission, if that is what it takes.

A photograph is a record of a unique event.   It is objective, up to a point, but it reflects a decision, made by the photographer, about when to release the shutter.   The microscopist scans a slide, and picks out particularly well-presented organisms or cells, not overlain by other cells or detritus in the sample, and also for pleasing juxtapositions of cells or filaments.   The same applies to those who photograph larger algae.  Tiff Stephens, the 2016 winner, could have waited a few moments longer, taken a step along the deck to her left or right, or held the camera at a slightly different angle.   Each would have given her a slightly different image of essentially the same phenomenon.  Whether photographing landscapes or using a microscope, there is nothing sacrosanct about the image beyond it being a record of the photographer’s decision to press a button.  Indeed, I suspect that most of our shortlisted entries are not unique records of the phenomena they record, but one of a number of images, and that a second stage of decision-making is needed to select the image that will be used.


Tiffany Stephen’s Swell Life: winner of the 2016 Hilda Canter-Lund prize.   The images at the top of the post show the 2009 and 2010 winners of the Hilda Canter-Lund competition, by Mariano Sirioni and Ernesto Macayo respectively. 

Having challenged the idea that the image, itself, is sacrosanct, there is no particular reason why you should not apply a third stage of decision-making and edit the image to enhance the story that you want to tell.   The field of view that is recorded when you press the shutter release is somewhat arbitrary.  You may be able to modify this, in a generic sense, in your camera’s settings but we usually adjust these only rarely and it is easier to adjust the pictorial space post hoc, using crop and rotate commands in a photo editing package.  The microscopist is further limited because most microscope stages do not rotate so the orientation of an organism can only be adjusted after the image itself has been collected.  Similarly, those of us who are photographing larger algae have only the small screens on our cameras with which to check images in the field, possibly in the face of inclement weather.  There is no disgrace in some judicious imaging editing once we can examine the image on a large screen, and the rules allow for this, along with focus stacking and stitching, essential tools in the microscopist’s armoury.

What about adjustment of colour and tone?   Bear in mind that colour, in the macro world with which we are most familar is reflected and objects can only reflect those wavelengths that reach them.  That means that colour and tone, in underwater photography especially, is not really a fundamental property of the organism you are photographing.   Move the same alga from a deep location to a shallow one, and it will look different for no other reason than the amount and quality of light transmitted through the water will change.   The microscopist is less likely to deal with reflected light, as the camera will be recording light that has passed through a specimen but, here too, the light is far from natural.  It will depend on the type of bulb, the intensity of light that you are using and the set-up of the microscope itself.  Once again, the colour and tones recorded are not fundamental properties of the specimen.   Under such circumstances, there seems to be no particular reason not to use the “levels” and “curves” options in editing packages to produce an image that is visually pleasing.  The judges are looking for basic authenticity and honesty in the image, so as not to deceive or misrepresent the natural world to the viewer, but there is a wide tolerance around this criterion because, frankly, natural light is, itself, so changeable.

The pair of photographs below illustrate this point well.  I was walking through local woodlands as I was thinking about this post.  May in the UK is the time when woodland floors turn a spectacular violet-blue due to the flowers of the bluebell (Hyacinthoides non-scripta).   I took the upper photograph on my iPhone then walked a few steps into the woodland to remove the dead tree that runs diagonally across the foreground.  I went back to my original position and took another photograph.   No more than 30 seconds elapsed between the two pictures, but the colour balance is completely different.   It may be a product of the metering in the camera itself (I’ve cropped both to show the same scene but the upper image had more bluebells and less woodland than the lower one) and this introduces another source of variation: the oh-so-clever electronics inside even fairly basic cameras that are making decisions on your behalf.


Two images taken within 30 seconds of each other from the same spot in woodland near Shincliffe, Co. Durham, May 2020.  The images at the top of the post show the 2009 and 2010 winners of the Hilda Canter-Lund competition, by Mariano Sirioni and Ernesto Macayo respectively. 

Most scientists assume that photography offers a “truthful” account of the objects that they are recording.   That’s at odds with the approach of critical theorists in the arts and humanities who recognise how many interventions lie between any object and the final image that is presented to third party viewers.   Susan Sontag, for example, challenges the “presumption of veracity” – less of an issue, perhaps, for fine artists but almost everything we think of as “documentary photography” or “photojournalism” is loaded with presumptions by both photographer and viewer, and it is a small step from those disciplines to scientist’s efforts to use photographs as objective evidence in their research.

The Hilda Canter-Lund competition is, however, not about photography as a scientific tool, but as a means of communication.  Appreciating the artificial nature of photography should be a liberation not a constraint: you, as photographer, probably have as accurate a memory of the image you have captured as the jpeg or tiff file that represents the digital record of the moment you released the shutter.   So feel free to open up the file in an editing package and use your discretion to adjust all the factors that were either in-built constraints or impulsive spur-of-the-moment decisions.   And send the final image to us for consideration for the 2020 Hilda Canter-Lund prize.

You can find the rules of the competition at along with examples of recent shortlists to inspire you.


Sontag, Susan (1977).  On Photography.  Penguin Books, Hamondsworth.

Other posts on photographing algae

How to win the Hilda Canter-Lund prize

How to win the Hilda Canter-Lund prize (2)

How to win the Hilda Canter-Lund prize (3)(guest post by Chris Carter, twice winner of the competition)

How to win the Hilda Canter-Lund prize (4)


Some other highlights from this week:

Wrote this whilst listening to: still working through my resolution to listen to all Bob Dyla’s albums in sequence.   This week I listened to The Basement Tapes, Desire, Hard Rain (much underrated in my opinion) and Street Legal.  Also enjoyed Jagged Little Pill by Alanis Morissette.

Cultural highlights:  The Assistant is an excellent but gruelling film that references the predatory behaviour of Harvey Weinstein but manages to do this almost entirely by inference and implication.

Currently reading:  Tamed by Alice Roberts, about the domestication of plants and animals, is interesting but rather turgid so I’m alternating chapters with Slaves of New York, a 1986 short story collection by Tama Janowitz which I borrowed from my son’s bookshelf.

Culinary highlight:   Baked cod topped with a pesto made from garlic mustard (Alliaria petiolata) foraged from the garden and allotment.

The dark side of the leaf …


Having mentioned in my previous post that the epiphytes on the top and bottom surfaces of a Potamogeton polygonifolius leaf were different, I have produced a companion piece to the painting I showed in that post.   The new painting is of the lower surface, and shows a greater number of diatoms than are present on the upper surface.  In order to explain why this is the case, it is helpful to look at the structure of the Potamogeton leaves.  The first image, therefore, shows a section through a leaf. It is quite a thick section but we can see the upper epidermis, the palisade mesophyll cells below this, which have plenty of chloroplasts in order to capture the sunlight that the plant needs for photosynthesis.  Below this, we can see parenchymous tissue arranged to create some large internal air spaces which contribute to the leaves buoyancy. Finally, at the bottom, there is a single layer of epidermal cells.   All this is crammed into a thickness of about half a millimetre.


Part of a section of a leaf of Potamogeton polygonifolius.  The leaf vein is on the left, thinning to the leaf blade on the right.  The leaf blade is about half a millimetre thick.   The picture at the top of the post shows an artist’s impression of diatoms and Chamaesiphon cf. confervicolus on the lower surface of a Potamogeton polygonifolius leaf. 

 Viewed from the underside, these parenchymous tissues create polyhedronal chambers, ranging from about 100 to 200 micrometres (a tenth to a fifth of a millimetre) along the longest axis.  There are also a few stomata scattered across the leaf surfaces (see the right hand image below).

With this in mind, take a look at my impression of the epiphytes growing on the lower surface of a Potamotgen polygonifolius leaf.   There are a number of cells of Chamaesiphon cf confervicolius, as seen on the upper surface, but there are several cells of the diatom Achnanthidium minutissimum, growing on short stalks, plus a few long, thin cells of Ulnaria ulna, growing in small clusters on the leaf surface (there were a few other species present, but such low numbers that I have not included them here).    It might seem strange to think of two surfaces of a leaf having such different communities of epiphytes but that’s because we’re thinking like large land-dwelling organisms, not like algae.   The longest alga visible in the image of the leaf underside is Ulnaria ulna, at about a 10th of a millimetre in length.  Therefore, to get a realistic impression of the two images, we really need to put a distance of five of these between them, and then pack the gap with chloroplast-rich mesophyll cells inside the Potamogeton leaf.   Allowing for foreshortening, this distance is about five times the height of the image.


The structure of a Potamogeton polygonifolius leaf viewed from the underside.  The left hand image (100x magnification) shows a leaf vein running diagonally across the lower right hand side along with the polyhedron-shaped chambers; the right hand image (400x magnification) shows the outline of one of these chambers superimposed behind the epidermal cells with a stomata with two guard cells visible just above the centre.   Scale bar: 20 micrometres (= 1/50th of a millimetre). 

The epiphytes on the upper surface of the leaf get first dibs at the meagre Pennine sunlight, which then has to pass through the upper layers of the Potamogeton leaf, where the mesophyll cells will continue to feast on the tastiest wavelengths, leaving relatively meagre pickings for the epiphytes that hang around on the underside of the leaf.

Chlorophyll, the molecule that makes plants green, absorbs light over a relatively narrow range of wavelengths – predominately red and blue – and this means that there are plenty of other wavelengths awaiting an organism with different pigments.   Diatoms have chlorophyll, but they also have some carotenoids (principally fucoxanthin) that grabs energy from the green part of the visible light spectrum (which is reflected, rather than absorbed by chlorophyll) and passes it to the cell’s photosynthetic engine.  Having this capability means that they can survive in relatively low light, which is why we see more diatoms on the underside of the Potamogeton leaf than on the top.

And that, best beloved, is the story of how Potamogeton got its epiphytes …


Some other highlights from this week:

Wrote this whilst listening to: more Bob Dylan.   I’ve got to the mid-70s, which means the live version of Like a Rolling Stone on Before the Flood plus the great Blood on the Tracks.  Also, as I was reading Ian Rankin, I listened to John Martyn’s Solid Air.

Cultural highlights:  we’re watching the BBC adaptation of Sally Rooney’s Normal People

Currently reading:  Ian Rankin’s Rather be the Devil.

Culinary highlight:   A rather fine vegetarian chilli, from Felicity Cloake’s column in The Guardian last week.   Served with corn bread, using a recipe we got from a hand-me-down American housekeeping magazine during our time in Nigeria.


Whatever doesn’t kill you …


The previous post focussed mostly on the higher plants that I found in the short stream that connects White’s Level with Middlehope Burn.  I mentioned the mass growths of algae that I found growing immediately below the entrance to the adit, but I did not talk about them in any detail, instead spinning off on a tangent while I mused on why the water cress had a purplish tinge.

When I did find time to examine the algal floc, I found it to consist of a mix of three different algae, the most abundant of which was Tribonema viride, but there were also populations of a thin Microspora (not illustrated) and Klebsormidium subtile.   I talked about Tribonema in the drainage from the Hadjipavlou chromite mine in Cyprus last year (see “Survival of the fittest (1)”) and both Microspora and Klebsormidium are also genera that are known to frequent these habitats.  Indeed, there is evidence that the populations that grow in these extreme habitats have physiological adaptations that help them to cope with the conditions.  Brian Whitton, my PhD mentor, led several studies on these adaptations in the streams of the northern Pennines in the 1970s, and Patricia Foster did similar studies in Cornwall at about the same time.   There is probably a mixture of physiological strategies involved, including the production of low-molecular weight proteins, which bind the toxic metals, and the production of extracellular mucilage.  Most of the populations I find in such habitats have a distinctly slimy feel due to the production of extracellular polysaccharides, and it is possible that these play a role in trapping the metal ions before they can get into the cell and cause damage.


Filamentous algae from the drainage channel below White’s Level, upper Weardale, April 2020.  a., b. & c.: Tribonema cf. viride, showing the characteristic H-shaped cell ends.   d.  Klebsormidium cf. subtile.  Scale bar: 10 micrometres (= 100th of a millimetre).   The picture at the top of the post shows an artist’s impression of Chamaesiphon cf. confervicolus on the upper surface of a Potamogeton polygonifolius leaf. 

I also had a look at the algae growing on the submerged leaves of Potamogeton pergonifolius in the channel between the adit and Middlehope Burn.   One easy way of examining them is to add a small amount of stream water then shake the leaves vigorously in a plastic bag.  The result is a brownish suspension of algae that can be sucked up with a Pasteur pipette and placed on a microscope slide.  When I did this, I found a community that was dominated by a short cyanobacterium, closest in form to Chamaesiphon cf. confervicolus.  The other abundant alga in the sample was Achnanthidium minutissimum, which is often common in minewaters, along with smaller numbers of a few other species.  The total number of species in the sample was just 12, which is low by the standards of streams without metal pollution, but such suppression of all but the hardiest species is another characteristic effect of heavy metal pollution.

I’ve added a “cf” (from the Latin conferre, meaning “compare to”) to my identification of Chamaesiphon confervicolus because this is the closest name, based on a comparison with images in the Freshwater Algal Flora of Britain and Ireland.  However, it is not an exact match.  Whether this is because the metals have strange effects on Chamaesiphon (as we saw for diatoms in “A twist in the tale …”) or whether our knowledge of the species within this genus is imperfect is not clear.  But discretion is the better part of valour in this instance.  Chamaesiphon species fall into two groups: those that live on stone surfaces (see “Survival of the fittest (2)”) and those that live on algae and plants, such as the one we see today (another is illustrated in “More from the River Ehen”).   They consist of a single, elongate but gently tapering cell, attached at one end to the plant and enclosed in a sheath.   The upper end of the filament forms small spherical buds (technically “exospores”).  One reason that I am wary of calling this population C. confervicolus is that most illustrations of this species show a stack of exospores in the sheath, whereas the White’s Level population all had just a single exospore.


Chamaesiphon confervicolus, growing on Potamogeton polygonifolius in White’s Level outflow, April 2020.   Note the exospores at the end of the cell.  f. and g. show the sheath very clearly.  Scale bar: 10 micrometres (= 100th of a millimetre). 

The picture at the top of this post shows an artist’s impression of the Chamaesiphon cf confervicolus on the upper surface of the Potamogeton leaf.   I wanted to get some idea of the size, shape and arrangement of the epidermal and stomatal cells on the Potamogeton leaves and resorted to the tried and tested technique of painting a layer of clear nail varnish onto the leaf surface, then peeling this off when it had dried.  This had the added (and unexpected) benefit of also pulling of the epiphytes, giving some idea of their arrangement on the leaf surface at the same time.   One extra observation that this yielded was that upper surface was dominated by Chamaesiphon, growing in clusters, whilst the lower surface had greater representation of diatoms.   I’ve also tried to portray the chloroplasts in the stomata guard cells.  Plant epidermal cells generally do not contain chloroplasts, as their purpose is to protect the mesophyll cells that are the main centres of photosynthesis.  Guard cells of stomata, however, need energy to open and close the stomata so these are the exception to this rule.  I had not even been sure that I would see stomata on the upper surface of the cell, as these are mostly found on the underside of leaves; however, Potamogeton appears to have stomata on both surfaces.  As ever, there is a certain amount of evidence along with a dose of extrapolation.   Imagined, but not imaginary …

You can find a description of the terrestrial plant life of Slitt Mine and its environs in this post on Heather’s blog.


Foster, P.L. (1982).  Metal resistances of Chlorophyta from rivers polluted by heavy metals. Freshwater Biology 12: 41-61.

Harding, J.P.C. & Whitton, B.A. (1976).  Resistance to zinc of Stigeoclonium tenue in the field and the laboratory. British Phycological Journal 11: 417-426.

Robinson, N.J. (1989).  Algal metallothioneins: secondary metabolites and proteins.  Journal of Applied Phycology 1: 5-18.

Say, P.J., Diaz, B.M. & Whiton, B.A. (1977).  Influence of zinc on lotic plants. I. tolerance of Hormidium species to zinc.  Freshwater Biology 7: 357-376.

Sorentino, C. (1985).  Copper resistance in Hormidium fluitans (Gay) Heering (Ulotrichaceae, Chlorophyceae).  Phycologia 24: 366-368.

(Note that Hormidium is the old name for the genus Klebsormidium.  There is an orchid genus called Hormdium and, as this was described first, it takes priority.)


Some other highlights from this week:

Wrote this whilst listening to: Bob Dylan’s New Morning and Pat Garrett and Billy the Kid.   Also, Samuel Barber’s Prayers of Kirkegaard.

Cultural highlights:  The Netflix series Unorthodox, about a young woman fleeing a Hassidic community in New York.

Currently reading:  Agatha Christie’s A.B.C. Murders.

Culinary highlight:   Arroz con leche (Spanish rice pudding) served with peaches poached in madeira.

The littoral ecology of Lough Down


My recent overviews of the major groups of algae have been useful as a way of highlighting which families and orders I’ve neglected.  That’s mostly because my posts are largely reactions to the circumstances I find myself in, rather than as a comprehensive overview of the world of freshwater algae.  My travels, this week, have brought me to Lough Down, a little-known Irish lake to which many of us have journeyed over the past few weeks.   Today, I thought I would peer at the littoral zone in the hope that I might find an alga from an order I have not previously written about.

I seem to be in luck: there has not been much rain recently and there is recently-exposed mud.  When I look closely, I see tiny green spheres, each the size of a pin head, dotted across the mud surface.   These are vesicles of the alga Botrydium and, despite their bright green colour, they actually belong to the Xanthophyceae (see “When a green alga is not necessarily a Green Alga …”).   Below this pear-shaped vesicle there is a system of branched rhizoids which anchor the plant to the surface (see lowermost photograph).  Surprisingly, the whole plant is a single cell, a similar situation to the one we encountered in Vaucheria, another representative of the Xanthophyceae (see “The pros and cons of cell walls”).

The vesicle itself is green, and a close look reveals the presence of many chloroplasts and also (less easy to see without special stains) nuclei.   You can also see, on the image below, crystals of calcium carbonate which are deposited on the vesicle.   However, if the marginal mud where Botrydium thrives is flooded again, the cell contents divide into a large number of spores, each with a single nucleus and two flagellae, which are liberated.  On the other hand, if the pond continues to dry, then a different type of spore is produced, as the cell contents retracts into the rhizoids where it forms thick-walled spores, which can survive long periods of desiccation.   Once the mud is dampened again, these spores germinate into motile spores.


A vesicle of Botrydium granulatum spotted with crystals of calcium carbonate.  The photograph at the top of the post shows vesicles on the bed of a pond (both photos: Chris Carter).

Botrydium is a small genus, with just eleven species listed on AlgaeBase, of which just one, B. granulatum, is recorded from the UK and Ireland.  It fills in one glaring gap in my coverage of the Xanthophyceae, leaving just two Orders still to feature.  These are the ones containing the awkward little single cells and colonies that are difficult to identify and easy to confuse with similar-shaped forms in the Chlorophyta.   I’ll get around to writing about these one day.   Maybe I’ll find representatives from them on a future visit to Lough Down.  Who knows how much time I will have to become acquainted with this fascinating lake?


A complete Botrydium granulatum plant, showing the vesicle on the left with a series of rhizoids extending out below.  The lowermost rhizoids are obscured by soil particles (photo: Chris Carter).  All images in this post are from material collected from Pitsford Water, Northamptonshire.

 Some other highlights from this week:

Wrote this whilst listening to: J.S. Bach’s St Matthew’s Passion and Laura Marling’s new album, Song for Our Daughter.  My systematic review of Dylan’s back catalogue has reached the incomparable Blonde on Blonde.

Cultural highlights:  Portrait of a Lady on Fire.   French arthouse film.  I know, I know …

Currently reading:  Drawing to the close of The Mirror and The Light .

Culinary highlight:   Buying a bag of strong white flour after many abortive attempts.   And a Simnel Cake, in celebration of Easter.


Blessed are you that hunger …


At the time of writing, four of my five working days are given over to ecology whilst the fifth is spent volunteering for the local Foodbank, which is gearing itself for a huge run on the stocks built up from generous donations over the Christmas period.   It occurred to me last week that I spend four days extolling hunger in an ecological context whilst spending the fifth trying to alleviate it in a human one.

“Hunger” in an ecological context is a them to which I have returned several times over the years.   We set the threshold for “good ecological status” for attached algae at a point that we thought coincided with the algal community switching from species that were adapted to “stressed” (i.e. nutrient-poor) conditions to those adapted to compete when nutrients were not in short supply (see “What does it all mean?” and references therein).   I’ve also talked, in some of my posts, about the adaptations some algae have to scavenge scarce nutrients (“A day out in Weardale”).   We’ve then gone on to try to work out what that means in terms of nutrient concentrations in UK and European rivers (references at the end of the post).

So I was pleased to see a paper appear last week that confirms some of these hunches.   Broadly speaking, Eleanor Mackay and colleagues have shown, using in situ bioassays, that as the concentration of inorganic nutrients decreases so the algae make more use of phosphorus and nitrogen bound into organic complexes.  As the algae get more “hungry”, in other words, they become more adept at scavenging for the resources that they need.

The graph at the top of this post is the graphical abstract from the paper which summarises this, whilst the one below shows the response to organic sources of phosphorus as a function of the concentration of “soluble reactive phosphorus” (the standard measure of “inorganic” phosphorus).  I’ve added an arrow to the right-hand side of this which shows roughly the current UK threshold, based on the work mentioned above.   Ellie’s graph seems to be confirming that, once that limit is exceeded, the algae are no longer “hungry”, meaning that they no longer need the nutrients bound into organic complexes.  Because organic phosphorus utilisation depends upon production of phosphatase enzymes to break down the organic complexes to releasee the phosphorus, there must be a greater energetic cost to the organism than if there was a ready supply of inorganic phosphorus that they can access.  I have, I must admit, never seen any figures that quantify this cost.


Fig. 5c from Mackay et al. (2020):  The relationship between “soluble reactive phosphorus” and dissolved organic phosphorus use by algae in in situ bioassays.  The “response ratio” is the natural logarithm of the ratio between the chlorophyll concentration of the treatment and the chlorophyll concentration of the corresponding control.  The arrow on the right-hand side indicates the approximate position of the regulatory threshold for phosphorus (see note at end of post).  The figure at the top of the post is the graphical abstract from Mackay et al. (2020). 

Part of me, then, is reassured that the regulatory threshold for phosphorus is roughly in the right place.  The Environment Agency’s reliance on a single measure of inorganic phosphorus, measured infrequently, is often criticised by hydrochemists but we can take some comfort from knowing that other forms of phosphorus (more difficult to analyse and quantify) only become important at concentrations lower than the current UK targets.   There is still part of me, however, that sees room for improvement.  That there are relationships between algae and other plants and phosphorus is not in doubt, and I am sure that a shift in strategies for nutrient acquisition help to define this relationship, particularly at low concentrations.  However, the relationships are not very strong and predictions about the ecological benefits of lowering phosphorus concentrations are imprecise.

Adding another strand of evidence to the current decision-making process makes scientific sense, and looking at how organisms respond to nutrients, rather than just measuring chemistry and describing community structure, seems like a sensible way of doing this.   In situ bioassays clearly have potential, as this paper shows; however, they are time consuming.   An alternative would be to measure phosphatase activity directly.  The Environment Agency did, in fact, fund research on this in the late 1990s and David Harper used these assays in a DEFRA-funded project in the early 2000s, but they have never become routine.  That’s a shame because, particularly for catchment-level investigations, they could add a useful additional insight.

The downfall of all these methods is not science, but the “more-with-less” ethos that has prevailed in the UK public sector for the past decade.  Everyone recognises that diffuse nutrient pollution offers a challenge that current monitoring and decision-making processes struggle to address.  However, most of the serious research effectively concludes with “if you spend a lot more money, you’ll discover that the problem is more complicated than you initially thought”.   That’s a difficult message to pass up through managerial hierarchies trying to keep a cash-starved regulatory agency moving forward.


Mackay, E. B., Feuchtmayr, H., De Ville, M. M., Thackeray, S. J., Callaghan, N., Marshall, M., Rhodes, G., Yates, C.A., Johnes, P.J. & Maberly, S. C. (2020). Dissolved organic nutrient uptake by riverine phytoplankton varies along a gradient of nutrient enrichment. Science of the Total Environment 722: 137837.

Poikane, S., Kelly, M. G., Salas Herrero, F., Pitt, J. A., Jarvie, H. P., Claussen, U., Leujak, W., Solheim, A.S., Teixeira, H. & Phillips, G. (2019). Nutrient criteria for surface waters under the European Water Framework Directive: Current state-of-the-art, challenges and future outlook. Science of the Total Environment 695: 133888.


Note: regulatory threshold for inorganic phosphorus

The arrow indicating the approximate position of the regulatory threshold for phosphorus uses the current UK TAG phosphorus standard.   This is site specific, using altitude and alkalinity as predictor variables.  This means that a range of thresholds is possible and the position of the arrow reflects the average alkalinity (50 mg L-1 CaCO3) and altitude (75 m) in a database of river samples collected as part of DARES project. Note, too, that P standards are based on the Environment Agency’s standard measure, which is unfiltered molybdate reactive P.  This approximates to “soluble reactive P” or “orthophosphate-P” in most circumstances but the reagents will react with phosphorus attached to particles that would have been removed by membrane filtration.


Some other highlights from this week:

Wrote this whilst listening to: My lockdown project of listening to all Bob Dylan’s albums in sequence has brought me up to Bringing It All Back Home and Highway 61 Revisited.

Cultural highlights:  Bait, a low-key black and white British film from 2019.  Definitely sits in the “sub hero” genre that I much prefer to the crash, bang, wallop of most Hollywood blockbusters.

Currently reading:  About three-quarters of the way through Hilary Mantel’s The Mirror and The Light now.  Jane Seymour is gone; Anne of Cleeves coming up next.

Culinary highlight:   Grilled mackerel with sautéed potatoes, probably.  A close second was home-made tortellini filled with mushroom paté and served with garlic mustard (Alliaria petiola) butter.   Mrs K is forager-in-chief hereabouts.


Disagreeable distinctions …

When you look at an organism, how do you know what it is?   That’s a big question that hovers over many of the posts that I write.   I tell you the names of organisms and you believe me. Sometimes I do too.   The truth is that we take the way that our brains process the constant stream of signals that our eyes send us as we observe the natural world without a second thought.   The subject intrigues me, but I only manage to scratch the surface in the posts that I write (see “Abstracting from reality …” and “Do we see through a microscope?” for some of these speculations).

The plate below offers a case study in this process.  It shows a diatom we encountered in a recent ring test, and which most us agreed was either Fragilaria austriaca or something quite similar.   In binary terms, though, we have to be blunt: either it is Fragilaria austriaca or it is not which may have implications for subsequent recording and interpretation (see “All exact science is dominated by the idea of approximation”).   How come a group of experienced analysts can look at the same population of diatoms and reach different conclusions?   I’ve got two suggestions: the first is that we differ in how we process the images, and the second is that there are sources of systematic error which confound our attempts to seek the right answer.


Fragilaria austriaca” from Foreshield Burn, Cumbria, May 2019.

There are three basic strategies that we use to name an organism:

  • Probabilistic reasoning, through the use of keys which, in theory at least, have a logical structure that guides a user to the correct identity of an unknown specimen. In practice, this is not quite as straightforward as it sounds (see “Empathy with the ignorant …”) and, at some point, many of us will abandon the formal structure of a key and switch to …
  • Pattern recognition, which amounts to flicking through images until we find one that matches our specimen. We can then corroborate this preliminary match by checking the written description.  In practice, we will probably switch from probabilistic reasoning to pattern recognition and back again as we home in on the identity of an unknown specimen. Repeating this process several times will lodge a schemata of this species in our memories, leading to a third strategy:
  • Recall. In practice, most of us probably have seen many of the common and even less-common species so often that we can by-pass these first two steps completely because we recognise the species without recourse to any books.

Disagreements, then, arise partly because we use different books as part of our naming process, our prior experiences differ and because our discipline in checking measurements of our own specimens against descriptions is not always as good as it should be.   In many cases, especially with modern understanding of diatom species, boundaries between species are frequently being redrawn and descriptions of newer species can only be found in obscure journal articles, often behind paywalls, and knowledge of these often diffuses through the community of diatomists more slowly than it should.   However, our discussions about the identity of the mystery Fragilaria also revealed a further issue, which I’ve illustrated in the graph below.

When we switch from “pattern recognition” to “probabilistic reasoning” we often base decisions on categoric distinctions of continuous variables such as length and width.  In this case, the literature quotes a maximum width of four micrometres for F. austriaca, and this was an important factor contributing to decisions about the correct identity.  However, there were differences in our measurements which means that some decided that the population was too broad to qualify as F. austriaca whilst others decided that it fell within the correct range.   The likelihood, based on these graphs, is that at least some of us were making incorrect measurements but, at this stage, we don’t know who they are.


Measurements of width, stria density and length of the population of “Fragilaria austriaca”.  Six analysts were involved in total, using either the eyepiece (“E”), an image projected onto a screen (“S”) or a measuring program (“P”) to make measurements (some used more than one approach). The dashed lines show the upper and lower limits for each parameter.

But that, itself, brings me to another point: do we know the correct size range of Fragilaria austriaca?  In order to be sure, we would need measurements of both initial cells (the largest in a cell cycle) and cells at the point where they are about to undergo sexual reproduction (the smallest in the cell cycle), ideally from several populations.  As this is rarely the case, we actually have three problems: first, is the description reliable? Second, are your measurements accurate? Third, we are using a point on a continuous scale as a criterion for a categorical judgement which implies perfect knowledge of the size range of the target population.  Even if you are sure of your microscope’s calibration, the best you can say is that the largest valve that you saw in the sub-sub-sub-sub-sub-sub sample of the population that lived in the stream you sampled exceeds (or not) the largest valve that the original author measured in the sub-sub-sub-sub-sample that s/he examined.   Several of our measurements just tip over four micrometres, the maximum width quoted in the literature for Fragilaria austriaca but, given these other factors, is that enough to drive a decision?   Statisticians are more comfortable predicting means, modes and medians than predicting extreme values.   Taxonomists, by contrast, seem to have undue reverence for maxima and minima.

Molecular biologists are approaching similar questions with considerable vigour.   The arrival of metabarcoding and high throughput sequencing means that they have had to write complicated computer code (“bioinformatics pipelines”) to sort the millions of sequences that emerge from sequencers, matching as many as possible to sequences from organisms whose names we already know, in order to turn those sequences into data that biologists can use (see “When a picture is worth a thousand base pairs …”).   We are conscious that decisions about software and settings within packages contribute to variations in the final output for reasons that we cannot always answer to our satisfaction.  But, whilst engaged in these discussions about cutting-edge technology, I’m conscious that old-school biologists such as myself each perform our own private “bioinformatics” every time we try to name an organism and we don’t always agree on the outputs from these thought processes.   Molecular biology, in a roundabout way, holds up a mirror to the way that we’ve been used to operating and should make us ask hard questions.

Some other highlights from this week:

Wrote this whilst listening to: my elderly vinyl copy of Mike Oldfield’s Tubular Bells

Cultural highlights:  Milton Jones at Newcastle City Hall

Currently reading:  Hilary Mantel’s The Mirror and The Light

Culinary highlight: polenta served with a mushroom and cheese sauce.

Finally, breaking news: I’m going to be live at the Green Man festival this August.  More details of our event “Slime Time”, and all the other performers at Einstein’s Garden can be found here


When a green alga is not necessarily a Green Alga…


I will end this short series of posts on the organisation of the major groups of algae with a look at the Xanthophyceae, or yellow-green algae.   My old copy of West and Fritsch’s Treatise on British Freshwater Algae from 1927 includes this group of algae with the green algae, although we now know that, apart from a generally green appearance, these two groups of algae have very little in common.  The big differences lie, however, in the types of details that are beyond the purview of the casual natural historian, so you may well find yourself flicking back and forth between “green algae” and “yellow-green algae” as you try to put a name on a specimen.  The definitive test is to add some iodine to your sample, as the Xanthophyceae do not produce starch as a storage product, and so do not produce the characteristic blue-black colour in the cells.  However, iodine is messy stuff and most of us will struggle along without for as long as possible.

The five orders of Xanthophyceae are shown in the table below.   In contrast to the case for most algal groups where molecular studies have led to many revisions of traditional classifications, the Orders of the Xanthophyceae have proved to be quite robust when subjected to this type of scrutiny.   Two of the Orders have siphonous organisation, though the form that this takes is very different in each (see “The pros and cons of cell walls” for more about siphonous lifestyles).  Tribonematales is an Order of filamentous algae that can be difficult to differentiate from filamentous green algae, whilst the Mischococcales are easily confused with small Chlorophyceae.


The organisation of the Xanthophyceae into five orders.  Organisation follows Algaebase.   The image at the top of this post shows Tribonema smothering the surface of a pond in Norfolk (photo: Geoff Phillips).

That’s one of the mysteries of freshwater algae: to the lay observer, an organism such as Vaucheria looks very similar to Cladophora or another green alga.  Yet they are distant relatives, belonging to different Kingdoms (Chromista and Plantae respectively).  That means that they share the same genetic affinity to one another as they do to us, which is a staggering thought (see “Who do you think you are?”).   What we are seeing is two organisms supremely well adapted to living in similar habitats, which means that natural selection has, gradually, shaped two quite distinct gene pools in quite different ways to arrive at the same end-point.   Just as motor manufacturers have, in the hatchback, found a style of car that is well-adapted to urban living, so the rival algae manufacturing corporations (“Plantae Inc” and “Chromista plc”) have come up with two broadly similar models that are both well-adapted to life in lowland streams.  Just as, in the case of hatchbacks, you can lift up the bonnet and see differences in the engine (petrol, diesel, hybrid, electric) but within the same basic shape, so many of the big differences in algal groups concern their internal machinery not outward appearances.


Reproductive structures growing from a filament of Vaucheria frigida (photo: Chris Carter)


Maistro, S., Broady, P.A., Andreoli, C. & Negrisolo, S. (2009).  Phylogeny and taxonomy of Xanthophyceae (Stramenopiles, Chromalveolata).  Protist 160: 412-426.


Links to posts describing representatives of the major groups of Xanthophyceae found in freshwaters.  Only the most recent posts are included, but these should contain links to older posts (you can also use the WordPress search engine to find older posts).

Group Link
Botrydiales Botryidium: The littoral ecology of Lough Down
Mischococcales Watch this space …
Rhizochloridales Watch this space …
Tribonemetales Tribonema: Survival of the fittest (1)
Vaucheriales Vaucheria: When the going gets tough …

Some other highlights from this week:

Wrote this whilst listening to: Two Hands, by Big Thief

Cultural highlights:  Jon Hopkins at the Sage.  What Radio 3’s Ibiza night might sound like.

Currently reading: the last few pages of Bill Bryson’s The Body: A Guide for Occupants (454 pages) prior to starting Hilary Mantel’s The Mirror and The Light (904 pages)

Culinary highlight: fish pie.  Spécialitié de la maison.


Rhapsody in red


On an overcast winter day with just a sprinkling of snow on the fells the Lake District can appear very monochrome.  Look closely at the bed of some rivers, however, and you are confronted by a much more vibrant palette with browns, greens and reds vying for your attention.  Somehow, paradoxically, the stream algae are at their most prolific and vigorous when the rest of Cumbria’s biological diversity has hunkered down to wait for the onset of Spring.

One of the most conspicuous groups at this time of the year are the red algae.  The green algae, diatoms and cyanobacteria are there all year round, even if winter is the time when they are most abundant.  The red algae, however, are barely evident – and certainly not to the naked eye – during the summer months.   It is only when autumn is well underway that the first blushes of pinkish red appear on the stones lining the beds of rivers.   This is in contrast to the red seaweeds which can be found on our coasts all year round, and indeed, to the many red algae that inhabit warm tropical seas.  What is so different about red algae in streams that leads them to favour the colder periods of the year?   What is it about streams, too, as I rarely see red algae in lakes (Batrachospermum is the exception: see “More algae from Shetland lochs”)?

This post will not answer those questions, but will give a quick overview of the red algae we find in freshwaters, in the manner of an earlier post about green algae (see “The big pictures …”).   The table below shows the systematics of the red algae, following a molecular phylogeny study by Hwan Su Yoon and colleagues from 2006.   There are two sub-phyla, of which one, Cyanidophytina, has no representatives recorded from the UK or Ireland.   There are just eight species in this group of primitive red algae, associated mostly with extreme environments.

The other subphylum, by contrast, has over 7000 species, divided between six classes, but 94 per cent of these are marine.   There are just thirteen genera of red algae recorded from freshwaters in the UK and Ireland, but spread amongst five of these six classes.   This seems to suggest that an ability to thrive in freshwaters has evolved several times during the evolution of this group.


The organisation of the red algae (Rhodophyta) showing division into two subphyla and seven classes.  Pink fill indicates the classes that are represented in UK and Irish freshwaters.   Organisation follows Algaebase and Yoon et al. (2006).    The photo at the top of this post shows Audouinella hermainii in the River Ehen, Cumbria, in December 2019.

Of the five classes that do have freshwater representatives, well over half of the genera and species recorded from the UK and Ireland are found in the Floridiophyceae.   This class has over 6900 species (95% of all red algae) split between 34 orders, of which five contain genera found in UK and Irish freshwaters.   Of these, the Batrachospermales, one of the few red algal orders that is exclusively freshwater, contains five genera and eleven species, whilst the other four contain just one genus each.

The Batrachospermales contain two morphologically-distinct groups of genera: Batrachospermum, Sheathia and Sirodotia form one of these, whilst Lemanea and Paralemanea form the other (see links below for more details and images).   Whilst we have molecular evidence that suggests that the Batrachospermales are a natural group, it is hard to point to a single characteristic that helps someone more interested in identification than taxonomy.   In fact, it is the life-cycle that is most distinctive (“… diplohaplontic … heteromorphic and contains a reduced tetrasporophyte”) but few of us are as well-schooled in algal life-cycles now as our predecessors were (see “Reflections from the Trailing Edge of Science”).   A hundred years ago, we would have had to rely upon the same limited set of morphological characters for both identification and taxonomy; now the taxonomist’s toolkit has expanded considerably whilst identification is still mostly reliant on features we can see with the naked eye or a light microscope.  For the red algae, this is still mostly enough to answer questions about what species we have found but unravelling the logic behind a classification may need a broader perspective.


Organisation of the Florideophycae showing the orders that include genera found in UK and Irish freshwaters.  



Entwisle, T.J., Vis, M.L., Chiasson, W.B., Necchi, O. & Sherwood, A.R. (2009).  Systematics of the Batrachospermales (Rhodophyta) – a synthesis.   Journal of Phycology 45: 704-715.

Yoon, H.S., Müller, K.M., Sheath, R.G., Ott, F.D. & Bhattacharya, D. (2006).  Defining the major lineages of red algae (Rhodophyta).  Journal of Phycology 42: 482-492.

van den Hoek, C., Mann, D.G. & Jahns, H.M. (1995).  Algae: an Introduction to Phycology.  Cambridge University Press, Cambridge.


And some other cultural highlights from the week:

Wrote this whilst listening to: Dave’s Psychodrama,

Cultural highlights:  Dave’s performance of Black (from Psychodrama) at the Brits Award Show.  I would not normally have watched this but was stuck in a hotel room with no wifi reception and was totally blown away by the power of his performance.

Currently reading: Bill Bryson’s The Body

Culinary highlight: I’m trying to cook one meal each month using only UK-sourced ingredients, in order to help me focus on seasonal cycles.  My February effort was a beer and cheese fondue: very easy to cook, using beer from about 500 metres from my house (Durham Brewery’s Evensong) and a mixture of Cheddar and Lancashire cheeses from Durham Indoor Market.



Links to posts describing representatives of the major groups of red algae found in freshwaters.  Only the most recent posts are included, but these should contain links to older posts (you can also use the WordPress search engine to find older posts).

Group Link
Bangiophyceae Watch this space …
Bangiophyceae Watch this space …
Compsopogonophyceae Watch this space …
Achrochaetiales Something else we forgot to remember
Balbianiales The Hilda Canter-Lund prize
Batrachospermales Lemanea: The complicated life of simple plants

Batrachospermum: More algae from Shetland lochs

Hildenbrandiales More about red algae
Thoreales Watch this space
Porphyridiophyceae Watch this space …
Stylonematophyceae More pleasures in my own backyard