Desmid diversity …

Back in September, I wrote about a joint British Phycological Society and Quekett Microscopical Club field weekend looking at desmids in the Lake District (see “Desmid Masterclass”, “Lessons from School Knott Tarn” and “Different tarn, different desmids …”).  Dave John sent some of the samples that we collected to David Williamson, the UK’s leading expert on desmids but, at 92, too frail to join us, and he has now sent back some fine drawings illustrating the range of desmids that he encountered.

Two of the tarns (Long Moss Tarn, Kelly Hall Tarns) are already recognised as Internationally Important Plant Areas (IPAs) for desmids because of their desmid diversity and containing internationally very rare desmids (based largely on David Williamson’s records) so their diversity is not a complete surprise to us.  Nonetheless, David found a total of 129 desmid taxa in the three tarns, whilst another desmid specialist, Marien van Westen, identified almost 160 desmids in another set of samples from the same tarns.

The drawings are arranged in three plates, one for each tarn.   Desmids identified by David Williamson from the three tarns are illustrated.  The desmids have been numbered and the captions prepared by David John who is analysing the findings and comparing them with surveys dating back to the 1970s.   David Williamson has drawn the taxa at different scales to roughly balance the arrangement on the collage, and adjusted the sizes so important details are visible.   No details of the chloroplasts are given since all samples had been preserved in formalin.  A few of the desmids, particularly those that are very long, have not been included in the plates.

Desmids from Long Moss Tarn (SD 292 936), September 2017.   Long Moss Tarn is shown in the photograph at the top of this post.

Desmids from Kelly Hall Tarn (SD 289 933), September 2017.

Desmids from School Knott Tarn (SD 427 973), September 2017.

Key

1-Actinotaenium diplosporum; 2-Actinotaenium turgidum;  3-Bambusina borreri;  4-Closterium acerosum var. borgei; 5-Closterium angustatum;  6-Closterium archerianum var. pseudocynthia;  7-Closterium archerianum; 8-Closterium attenuatum;  9-Closterium baillyanum var. alpinum; 10-Closterium baillyanum; 11-Closterium closterioides; 12-Closterium costatum; 13-Closterium dianae var. arcuatum; 14-Closterium dianae var. minus;  15-Closterium didymotocum; 16-Closterium incurvum; 17-Closterium intermedium; 18-Closterium kuetzingii;  19-Closterium lunula; 20-Closterium navicula;  21- Closterium setaceum; 22-Closterium striolatum; 23-Cosmarium amoenum; 24-Cosmarium anceps; 25-Cosmarium binum; 26-Cosmarium brebissonii; 27-Cosmarium contractum;  28-Cosmarium davidsonii; 29-Cosmarium debaryi;  30-Cosmarium depressum; 31-Cosmarium formosulum; 32-Cosmarium hostensiense; 33-Cosmarium incrassatum var. schmidlei; 34-Cosmarium margaritatum; 35-Cosmarium margaritiferum; 36-Cosmarium monomazum var. polymazum;  37-Cosmarium obtusatum;  38-Cosmarium ornatum; 39-Cosmarium ovale;  40-Cosmarium pachydermum; 41-Cosmarium pachydermum var. aethiopicum; 42-Cosmarium perforatum var. skujae; 43-Cosmarium portianum; 44-Cosmarium punctulatum;  45-Cosmarium quadratum; 46-Cosmarium quadrum; 47-Cosmarium subochthodes var. majus; 48-Cosmarium subtumidum var. groenbladii;  49-Cosmarium subundulatum; 50-Cosmarium tetragonum var. ornatum ; 51-Cosmarium tetraophthalmum; 52-Cosmarium variolatum;  53-Cosmocladium tuberculatum; 54-Desmidium aptogonum; 55-Desmidium swartzii; 56-Docidium baculum; 57-Euastrum ampullaceum; 58-Euastrum ansatum;  59-Euastrum bidentatum var. speciosum; 60-Euastrum gemmatum; 61-Euastrum luetkemulleri; 62-Euastrum oblongum; 63-Euastrum pectinatum; 64-Euastrum pulchellum; 65-Euastrum verrucosum; 66-Gonatozygon aculeatum; 67-Gonatozygon brebissonii; 68-Groenbladia undulata; 69-Haplotaenium minutum;  70-Hyalotheca dissiliens;  71- Micrasterias americana var. boldtii; 72-Micrasterias compereana; 73-Micrasterias crux-melitensis; 74-Micrasterias denticulata; 75-Micrasterias furcata; 76-Micrasterias pinnatifida;  77-Micrasterias radiosa; 78-Micrasterias rotata; 79-Micrasterias thomasiana; 80-Micrasterias truncata; 81-Netrium digitus; 82-Netrium digitus var. latum; 83-Netrium interruptum;  84-Penium exiguum; 85-Penium margaritaceum; 86-Pleurotaenium coronatum var. robustum;  87-Pleurotaenium ehrenbergii; 88-Pleurotaenium truncatum; 89-Sphaerozosma filiforme; 90-Staurastrum arachne;  91-Staurastrum arctiscon; 92-Staurastrum bieneanum; 93-Staurastrum boreale var. robustum; 94-Staurastrum cristatum; 95-Staurastrum dilatatum; 96-Staurastrum inconspicuum; 97-Staurastrum kouwetsii; 98-Staurastrum lapponicum; 99-Staurastrum maamense; 100-Staurastrum polytrichum; 101-Staurastrum productum; 102-Staurastrum quadrangulare; 103-Staurastrum striolatum; 104-Staurastrum teliferum; 105-Staurastrum tetracerum; 106-Staurodesmus convergens; 107-Staurodesmus convergens var. wollei; 108-Staurodesmus cuspidatus var. curvatus; 109-Staurodesmus megacanthus; 110- Xanthidium antilopaeum; 111-Xanthidium antilopaeum var. laeve; 112-Xanthidium antilopaeum var. polymazum; 113-Xanthidium cristatum.

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Small details in the big picture …

I’ve written about Platessa oblongella, a small diatom common in low alkalinity environments, before (see “A tale of two diatoms …” and links therein) but my travels around west Cumbria are gradually revealing more and more about the ecology of this organism, so bear with me as I explain my latest findings.

My first graph shows how the distribution of this diatom varies in different types of water body in the Ennerdale catchment.   I have analysed 223 samples from this small area over the past few years and, within this dataset, there is a very clear distinction between situations where Platessa oblongella is abundant and situations where it is very rare.   I have very few records from Ennerdale Water itself (present in just two out of 27 samples, and never comprising more than 2.7% of all diatoms in the sample) nor from the River Ehen, which flows out of the lake (present in just 16 out of 164 samples, and always £ 1% of all diatoms).  By contrast, in Croasdale Beck and in streams that flow into the north-west corner of the lake, it is present in 28 out of 32 samples, with a maximum relative abundance of 69%.   In ten samples it forms more than 10% of all diatoms present.   Several of my samples from the small streams were collected from just a few metres above the point where they joined the lake, which makes the distinction between these streams and the lake that much more intriguing.

My theory – based on data I showed in A  tale of two diatoms  is that Platessa oblongella is a species of disturbed habitats and that the littoral zone of a lake, whilst subject to some turbulence, is less disturbed than the rough world of an unregulated stream.  The contrast between the River Ehen immediately below the dam at the outfall of the lake and the various small tributary streams also supports this idea.

Differences in percentage of Platessa oblongella (including P. saxonica) in epilithic samples from Ennerdale Water and associated streams.  Data collected between 2012 and 2018 (along with one sample from River Ehen collected in 1997).   The photograph at the top of the post shows Ennerdale Water, photographed in January 2018.

Some of the populations I looked at seemed to consist of two distinct forms, one broader than the other.   This variability is quite common in Platessa oblongella and Carlos Wetzel and colleagues recently published a paper which suggests that these are, in fact, two distinct species.   When I first started looking at diatoms, John Carter, my mentor, used the name Achnanthes saxonica, but Krammer and Lange-Bertalot, in the revised Süsswassserflora, regarded this as a synonym of Achnanthes oblongella, a species first found in Thailand.   Wetzel’s study shows, as well as the difference in valve width, differences in the fine details of the striae between the two species.   They also decided that both species belonged in the genus Platessa, rather than Achnanthes.

Platessa oblongella (top) and P. saxonica (bottom) from Croasdale Beck, October 2017.  Scale bar: 10 micrometres (= 1/100th of a millimetre).

Valve width is, however, a very useful criterion, as the histograms below show.   The left hand graph shows a distinctly bimodal distribution of widths in specimens from Croasdale Beck, whilst the right hand graph shows a much tighter, and clearly unimodal, range.   This comes from another tributary stream flowing into the Ehen about 500 metres below the lake itself.  Quite why two species can co-exist in one stream but only one is present in another is not clear.

The modes of these populations are very close to the median widths for P. saxonica (narrow, ± 4/5 – 5 mm) and P. oblongella (broader, ± 6.5 mm) respectively but, as the left hand histogram shows, there is some overlap.    You might have trouble, for example, deciding whether a valve that was 5.5 mm wide was a “fat” P. saxonica or a “thin” P. oblongella.   My standard advice in situations such as this is that we should identify populations not individuals although, in the case of Croasdale Beck, this will still leave a grey area between the “fat” and “thin” valves where a judgement call is necessary.   In this case I think I could have done it because the P. saxonica valves in this stream tended to have a greater length:breadth ratio than those of P. oblongella, though I have not actually quantified this.

Width of valves in populations of “Achnanthes oblongella” from a) Croasdale Beck, and b) an unnamed tributary stream of the River Ehen, October 2017. 

There is more to say about the ecology of these species, but I have probably written enough for now.  I will leave you, for now, to bask in the rare sensation that occurs when diatom taxonomists make a situation clearer rather than more opaque, and return to this subject in a future post.

References

Carter, J.R. (1970).   Observations of some British forms of Achnanthes saxonica Krasske.  Microscopy: Journal of the Quekett Microscopical Club 31: 313-316.

Wetzel, C.E., Lange-Bertalot, H. & Ector, L. (2017).  Type analysis of Achnanthes oblongella Østup and resurrection of Achnanthes saxonica Krasske (Bacillariophyta).  Nova Hedwigia, Beiheft 146: 209-227.

Change is the only constant …

The diatoms I saw in my sample from the littoral of Lake Popovo (described in the previous post) reminded me of an assemblage that I had seen in another lake which, apart from its location, has much in common with Popovo. This lake is Wastwater, in the western part of the English Lake District (see “The Power of Rock …”).  Like Popovo, it is situated in a remote a region of hard volcanic rocks and, as such, has very soft water and is subject to few of the pressures to which most of our freshwaters are subject.  The photograph above shows me sampling Wastwater in about 2006 (more about this photograph, by the way, in “A cautionary tale …”).

I wrote about Wastwater when I was writing my book Of Microscopes and Monsters, the precursor of this blog.   I focussed, in particular, on an experiment that my friend Lydia King had performed as part of the research towards her PhD.  Her previous work had established that there were relationships between the types of algae that she found in lakes in the Lake District and the amount of nutrients that they contained.  She also saw that the types of algae she found depended upon how acid or alkaline the water was.  But the water chemistry only explained a part of the variation in the algae and now she wanted to find out about the variation that was not explained by this.   In particular, she wanted to know how much of the variation was due to the way that the algae interacted with each other.

Lydia’s experiment involved putting clay pots into the shallows at the edge of Wastwater and then watched how the algal communities changed over the course of six weeks.  She also examined small parts of the pots at extremely high magnifications using a scanning electron microscope.   These micrographs, and subsequent conversations with her, had inspired some of my early paintings and I returned to this subject several times, finally producing a series of three pictures that showed changes in the algae over time.

The microbial world of the littoral zone of Wastwater after two weeks of colonisation showing unidentified small unicellular blue-green alga,  unidentified small unicellular green alga; thin filaments of Phormidium,  Achnanthidium minutissimum and Gomphonema parvulum.

The first of these shows the surface of the plant pot after being submerged in Wastwater for two weeks.   You could think of this as a patch of waste ground that was, at the start of the experiment, bare of vegetation.   If we watched this patch over a number of weeks, we would notice some plants appearing: scattered stalks of grass, perhaps some rosebay willow herb, dock or plantains. A gardener might dismiss these as “weeds”, although this term has no ecological meaning but ecologists prefer to think of these as “pioneers”: plants adapted to colonising new habitats, growing quickly (which might mean producing lots of seeds in a short space of time or producing rhizomes or runners) and covering the ground.  This same process has taken place on Lydia’s plant pot in Wastwater: the “weeds” in this case are scattered thin filaments of the blue-green alga Phormidium, the diatoms Achnanthidium minutissimum and Gomphonema parvulum plus a number of spherical green and blue-green cells that she couldn’t identify.   Such is the scale that we are working at that this open landscape still contains about 92000 cells per square centimetre.

The microbial world of the littoral zone of Wastwater after three weeks of colonisation.   The composition is similar to that in the previous figure but the density of cells is greater.

When she came back a week later, much of the empty space had been infilled; there were now about 300,000 cells per square centimetre.  These mostly belonged to the same species that she had found the week before.  The difference is that they are now rubbing up against each other and this has some important consequences.  All plants need light and nutrients to grow and algae are no exceptions.   Sunlight provides the energy for photosynthesis but now, at week three, the density of algae is such that there is a chance that some of the light will be intercepted by a neighbouring cell.   The total amount of sunlight that filters through the water to the pot surface is already much lower than that available at the lake surface; now it has to be shared out between many more cells.   At this point, properties such as fast growth rates that helped our pioneers to colonise the plant pot become less relevant, and it is algae that are better adapted to capturing the limited light that will survive.

So when Lydia came back to Wastwater after six weeks, she saw a very different community of algae on her pots.   There was still a lot of Achnanthidium minutissimum, but rising above these was the elegant art deco shape of Gomphonema acuminatum (also found in Lake Popovo) which, importantly for our story, grows on a long stalk.  There are also cells of “Cymbella affinis” (the correct name at the time that Lydia was working but see comments in the previous post about the nomenclatural history of this species).   This, too, grows on a long-stalk, the better to grow above the Achnanthidium and other pioneers.   If we continue to use the analogy of a patch of wasteland, then it has now reached the point where it has been invaded by shrubs such as hawthorn and blackthorn.   However, in a terrestrial habitat this would happen two or three years after the first pioneers had arrived, not six weeks as Lydia had observed for the algae.   She also found the diatom called Tabellaria flocculosa which forms filaments.  These often start out loosely-attached to the substratum but more often break free and become entangled around the other algae.   In our “wasteland” analogy, these would be the brambles.

The microbial world of the littoral zone of Wastwater after five weeks of colonisation.  Gomphonema acuminatum, “Cymbella affinis” and Tabellaria flocculosa have now joined the assemblage seen in the two earlier dioramas.

The experiment finished shortly after this, terminated when the apparatus was overturned.  Whether by a wave or by vandalism, Lydia will never know but this event is, itself, a metaphor for the harsh world in which benthic algae have to survive.  In real life, the many cobbles in the littoral zone will be rolled by wave action or, as we have seen in other posts, invertebrate grazers could have removed much of the “shrubbery”, leaving a “pasture” composed of the tough, fast-growing species such as Achnanthidium minutissimum to dominate samples.   The “successions” we see in the microscopic world not only take place much more quickly than those in the macro world, but they also rarely have a stable “climax”: just a brief pause before the next onslaught from the physical, chemical and biological processes that shape their existence.

References

King, L., Barker, P. & Jones, R.I. (2000). Epilithic algal communities and their relationship to environmental variables in lakes of the English Lake District. Freshwater Biology 45: 425-442.

King, L., Jones, R.I. & Barker, P. (2002). Seasonal variation in the epilithic algal communities from four lakes of different trophic state. Archiv für Hydrobiologie 154: 177-198.

What a difference a storm makes …

I was back at Croasdale Beck last week and noticed a rather dramatic change to the meander just upstream from our regular sampling spot.   If you look at the photograph that heads the post “A tale of two diatoms …”, you’ll see the stream flowing around this meander.  Now, however, it has cut a new, shorter channel that bypasses the meander altogether.   We visited the stream just a few days after Storm Ophelia had passed through although, judging by the grass growing on the gravel of the abandoned meander, it was not necessarily this particular event that reshaped the stream.

Croasdale Beck is an unruly tributary of the River Ehen, rising on the fells above Ennerdale Water and tumbling down across rough grazing land and some semi-improved pasture (as in the picture above) before joining the Ehen in Ennerdale Bridge.   This is not the first time that we have seen conspicuous changes in the channel after a storm.  The magnitude of the flood is illustrated by the hydrograph below, which went off-scale for a period, as the discharge exceeded 3000 mega litres per day (300 MLD is the approximate limit for safe wading, in my experience).   I noticed that there was much less green algae present than we usually record at this time of year, although the diatom film was still quite thick.   Some of the stones that I picked up to sample had the slimy biofilm on the underside, suggesting that they had been recently rolled by the flooded river.   Croasdale Beck has no lake to buffer the rise and fall of the floodwaters and a huge amount of energy is carried down in a short period of time as the water surges downstream.

By the time we had arrived, the floodwaters had subsided and the sheep were contentedly grazing the surrounding land.  The stream itself was almost back to base flow (in contrast to the River Ehen which was still only just wadable).  Only the meander looked different …

The hydrograph for the River Ehen, as the aftereffects of Storm Ophelia make their way downstream.

Buffers for duffers …

In Ecology in the hard rock café I wrote about the challenges of living in an aquatic world where carbon – one of the raw materials for photosynthesis – was in short supply.   What I did not write about in that post is that this carbon also gives freshwater some useful additional properties.   In brief, rainwater is not pure water, but absorbs carbon dioxide from the atmosphere.  This, in turn, makes rainwater slightly acidic and, when it falls onto rocks, this weak acid dissolves the minerals from which the rock is made.  This adds two other forms of carbon to the water – bicarbonate and carbonate (the latter, particularly, from limestone).

Each of these three types of carbon in freshwater can convert to either of the other two types, with the speed of the reaction depending on the balance between the forms (the “law of mass actions”).  In essence, the reactions proceed until equilibrium is obtained, and this equilibrium, in turn, depends upon the pH of the solution.  These processes are summarised in the diagram below.

Relationship between pH and the proportion of inorganic carbon as free carbon dioxide (or carbonic acid, H2CO3 – orange line), bicarbonate (HCO3 – green line) and carbonate (CO32- – blue line).

The chemistry behind this is not easy to explain but a consequence is that any attempt to shift the pH (e.g. by adding acid) causes an automatic adjustment in the balance between the different forms of carbon.  Some of the hydrogen ions that could make the water acid are, instead , bound up as bicarbonate, and the pH, as a result, does not change.  The greater the quantity of inorganic carbon in the sample, in other words, the greater the capacity of the water to resist changes in pH.   The carbonate, bicarbonate and free carbon dioxide together act as a “buffer”, a chemical shock absorber.   Think of it as equivalent to the responsible use of a credit card or savings account to defer the cost of an unexpected bill (a car repair, for example) so that your current account does not go overdrawn.

Because life largely evolved in well-buffered marine systems, the enzymes that run our cells generally work best within a narrow range of pH (approximately 6-9).   Cells – unicellular life forms in particular – get stressed if pH strays outside this range, so the greater the buffering capacity, the easier it is for cells (life at high pH can bring additional complications, but we don’t have time to go into those here).  “Alkalinity”, as I mentioned in the earlier post, is the measure that ecologists use to assess the strength of the buffer system in a lake or river.  The principle of the measurement is straightforward: we add a dilute acid very slowly and watch what happens to the pH.   At first, nothing happens but, as soon as the water’s natural buffering capacity has been exceeded, pH drops rapidly.

I have a small portable alkalinity titration kit which involves adding drops of bromophenol blue indicator to a sample of stream or lake water.  This gives the water a blue colour when the pH is greater than 4.6.  As the pH falls, the solution becomes colourless and, eventually, turns yellow.   If you look at the graph above you will see that, at pH 4.6 most of the bicarbonate (HCO3) has been converted to carbon dioxide so the buffering capacity is pretty much non-existent.  This means that I can use the quantity of acid that is needed to make the bromophenol blue change colour as a measure of the buffering capacity of the water.

Alkalinity titrations beside Ennerdale Water (see top photograph) using a Hanna HI 3811 alkalinity test kit.  The right hand image shows acid being added to the water sample with a 1 ml pipette.  The blue colour shows that pH has not yet dropped below 4.6.

All this talk of chemical equilibria seems to be a long way from the natural history that is the core business of this blog.  Yet, at the same time, these reactions describe natural phenomena every bit as real as the plants and animals that attract the interest of naturalists.   Geology and chemistry ultimately create the context within which biology flourishes, but it is rare to meet a chemist who can talk with a naturalist’s passion.  I think that this is partly because chemistry tends not to describe tangible features of the landscape but, instead, quickly gets lost in abstract equations.  However, it is also a matter of culture: chemists need clinical separation from the mud and filth to maximise precision, whilst ecologists feel the lure of the field.  There is, nonetheless, a very basic and necessary link between the chemistry and ecology of aquatic systems.   Geology may shape a landscape but chemistry is one of the key mediators that determines the types of plants that cloak the hills and vales.  We ignore it at our peril.

Taking desmids to the next dimension …

Participants at the British Phycological Society / Quekett Microscopical Club field weekend at the Freshwater Biological Association, September 2017.  Scale bar: one metre (= 1,000,000 micrometres).

A theme that has run through this blog over the years has been that what you see down a microscope is often a highly distorted view of reality and at the end of our weekend of desmid hunting, Chris Carter gave a talk that also made this point, using desmids as a case study.  In essence, we had spent much of Saturday and Sunday morning peering down microscopes at three-dimensional objects that appeared, as a result of the very shallow depth of field that is characteristic of high magnification images, two dimensional.   We were then matching these to two-dimensional representations in the Floras and identification guides that we had to hand.  Dave explained a few tricks that experts use, such as applying gentle pressure to a coverslip with a fine needle, to turn desmids in order to see them from other angles but, mostly, we were restricted to very flattened views of desmids.

Chris has tackled this problem from several directions over the years, including experiments with anaglyphs (see “Phworrrrhhh …. algal sex in 3D!“) as well as the very careful manipulation of a long, cylindrical Pleurotaenium that won the Hilda Canter-Lund prize earlier this year.   He has also produced a number of plates with desmids laid out almost as if on an engineer’s drawing board, with front, top and side views.   Several of these are on Algaebase, but one example is reproduced below.  Microscopists learn to use the fine-focus control to appreciate the depth of the objects that they are examining and Chris also shows how it can reveal the nature of surface ornamentation on different parts of the cell.  The temptation, given a series of photos such as these (excluding the side view) would be to use “stacking” software to produce a single crisp image.  This is appropriate in some situations but you are, in truth, just producing a crisp two-dimensional image rather than offering any insights into the true shape of the cell.

Staurastrum furcatum from Botswana, photographed by Chris Carter.

Another technique that can be used to generate three-dimensional images is, of course, scanning electron microscopy.  However, this is beyond the budget for anyone outside a major institution.  This has helped greatly get a better understanding of the morphology of diatoms, in particular, but the third dimension comes at a price.   Scanning electron micrographs take us to an opaque, monochrome world, purged of the vivid colours that the microscopic world usually offers us.

Chris’ pièce de résistance, however, was a three-dimensional model of a Staurastrum, produced by the 3D printing company Shapeways and loosely-based on various pictures of S. furcatum and presented to him as a 70th birthday present by his son.  The main point is to demonstrate the symmetry and gross features of a typical Staurastrum rather than to be a taxonomic blueprint. The designers were very helpful but it does hint at what is possible with modern technology.

Chris Carter’s three-dimensional model of Staurastrum.  It is about six centimetres across.   You can buy your own copy from Shapeways by following this link …

The missing ingredient in this recipe is imagination.  Or, to be more precise, the viewer’s imagination as Chris has clearly demonstrated that he is not lacking in that department.  Once you have a sense of the three-dimensional form of a Staurastrum, you be able to use that knowledge every time you look at a two-dimensional image of a desmid through a microscope.   Seeing, as Ernest Gombrich reminds us in his great book Art and Illusion, is as much about using prior experiences to interpret the raw data collected by our optic nerves as it is about the patterns of light that stimulate our retinas.   Just as a child can look at a two-dimensional image of a cat in a book and match this to the real creatures that he or she encounters, so knowing about Staurastrum’s third dimension helps us to interpret the flat shapes that we see.

At a more basic level, all identification is a matter of matching the objects we see either to schemata stored in our memory or to patterns in books.  This, in turn, helps us to understand why the microscopic world seems so strange and mysterious to those who do not study it.  It all comes down to having (or not having) the prior experiences that generate recognition.   At one level, there are gasps of astonishment as people with none of these schemata in their memories encounter the beauty of desmids for the first time.  And then there is Frans Kouwets, another speaker at the meeting , who is busy cataloguing 750 different species of one genus, Cosmarium.   And in between there are the rest of us …

Frans Kouwets explains his fascination with Cosmarium to the British Phycological Society / Quekett Microscopical Club field meeting at the Freshwater Biological Association in September 2017.

Different tarn, different desmids …

Geoff and Chris, two of our band of desmid hunters, chose to stay in the FBA’s brand new holiday apartments and, rather than cross the lake to join us on Saturday morning they headed out to Moss Eccles Tarn, in the area between Esthwaite Water and Windermere.   One of Dave’s first dips into one of their samples yielded an almost pure monoculture of another filamentous desmid, Spherozosma vertebratum which presented some beguiling abstract patterns on my computer monitor.

Spherozosma vertebratum from Moss Eccles Tarn, September 2017.   Scale bar: 25 micrometres (= 1/40th of a millimetre).

Curiously, after our first encounter with Spherozosma vertebratum we did not see it in any of our other dips into the Moss Eccles samples although there were plenty of other desmids on display.   The most abundant of these was Staurastrum productum and, usefully, there were examples showing both apical and side views.   The three arms are distinctive (and distinguish it from relatives such as S. arachne which have five) and you can also see the knobbly “verrucae” on the spines as well as a broad mucilaginous envelope around the cells.

Staurastrum productum in side (left) and apical (right) views.  Images photographed from a computer monitor so apologies for their poor quality.  Scale bar: 25 micrometres (= 1/40th of a millimetre).

Another desmid with spines and mucilage was quite common.  This was Staurodesmus bulnheimii.  Spines slow the rate of sinking so are associated with several genera of predominately planktonic desmids.   The star-shaped arrangement of colonies of the diatom Asterionella formosa play a similar role (see “Little bugs have littler bugs upon their backs to bite ‘em”).   There were also several cells  of a small Cosmarium species, including some that had recently divided and the image shows how one cell has split down the central isthmus and a new semicell is growing back on each of the two daughter cells.   Finally, I have included an illustration of Micrasterias radiosa.  To the uninitiated this may look little different to M. compereana, illustrated in the previous post, but if you look closely you will see that the incisions between the lobes are much deeper in M. radiosa.

One sample from Moss Eccles Tarn kept me busy for half the morning and this account describes only part of the diversity.   Note how the differences between this and the School Knott Tarn sample are not just in the genera and species present but also in the life-forms I found.  The School Knott sample was from a Sphagnum squeezing whilst the Moss Eccles sample was from a plankton net.  That explains why I saw more spine-bearing desmids in the latter.  If I had looked at a plankton sample from School Knott and a Sphagnum squeezing from Moss Eccles, I might have found a different balance of life-forms between the two tarns.   But time was running out and I had to move on …

More desmids from Moss Eccles Tarn, September 2017: a. Staurodesmus bulnheimii; b. Cosmarium quadrifarium var. hexastichum; c. Euastrum cf. gemmatum.   Scale bar: 25 micrometres (= 1/40th of a millimetre).

Micrasterias radiosa from Moss Eccles Tarn, September 2017.   Scale bar: 25 micrometres (= 1/40th of a millimetre).