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.

Advertisements

The curious life of biofilms …

My explorations of the microscopic world of the River Wear have now gone one step further with the transformation of the schematic representation that I presented in The River Wear in January into a three-dimensional diorama.   This shows the “biofilm” on the top of submerged stones, with a layer of Navicula lanceolata at the top (the chocolate brown layer in the photograph from the earlier post) intermingled with small Gomphonema cells on long stalks and some cyanobacterial filaments.   A large part of the biofilm, however, is inorganic particles and aggregations of organic matter.

I’m curious about why this biofilm is thickest in the winter, not just in the River Wear but in many other rivers too.   Part of the reason is that the organisms that form this film can outpace the bugs that want to eat them at this time of year but this is not the whole story.    As the image shows, the biofilm is about far more than just algae, so we need to know a little more about all that organic matter that takes up so much of the space in the picture.   Where does it come from and why does it accumulate on stone surfaces?

The story starts with the polysaccharides that algae and other microorganisms (fungi and bacteria) secrete as they grow.   These polysaccharides play several roles – they provide the stalks for diatoms such as Gomphonema, they help motile diatoms such as Navicula move and they also ensure that any enzymes that the organisms secrete stay in the proximity of the cell while they perform their functions.  However, as well as servicing the organisms that produce them, they also alter the chemical and physical environment on the stone surface.   Organic and inorganic particles, for example, can be trapped amongst the stalks of diatoms such as Gomphonema, but there are also chemical interactions.  River water contains dissolved organic matter, the end-result of the breakdown of organic matter such as leaves further upstream.   This can flocculate to form small particles which can be physically trapped, or it may be adsorbed onto the various polysaccharides in the biofilm.

If you think of a snowball rolling down a hill and growing in size as more and more snow gets stuck on the outside, you have a very rough idea of how a biofilm grows.   Simply being a biofilm is enough to help it become a bigger biofilm, as the wide range of biological, chemical and physical interactions that take place will increase the quantity of living and dead organic material, along with inorganic particles.  The supply of organic material varies through the year, and is greatest in autumn, following leaf fall (see “A very dilute compost heap …”).  The biofilm, unlike the snowball, is largely static; it is the water around it which is moving, bearing with it the raw materials to help it grow.  However, the biofilm also bears the seeds of its own destruction: all that organic matter – whether produced by algae in situ or imported from upstream – makes it a nutritious food source for the small invertebrates that inhabit the stream bed.  I often see midge larvae eating their way through both living and dead matter when I am examining samples under my microscope.   They are there throughout the year, but are busier in the warmer months when, as a consequence, the biofilms are thinner.

Curiously, despite having collected this sample from a stretch of the Wear where I could feel the strength of the current pushing against my legs, flow has relatively little effect on biofilms.   There is a thin layer just above the bed of the river where there is almost no current, due to frictional drag and the biofilms exist in this zone.   Only when the discharge becomes so strong that the stones themselves are overturned do we see major losses to the biofilm itself.   I have seen a medium-sized summer spate in the Wear lead to the opposite effect: a rapid increase in biofilm thickness, presumably because the invertebrates were more vulnerable than the smaller algae.

I will return to the same location on the River Wear in March to see how things have changed.

References

Blenkinsopp, S.A., & Lock, M.A. (1994).  The impact of storm-flow on river biofilm architecture.   Journal of Phycology 30: 807-818.

Liu, W., Xu, X., McGoff, N.M., Eaton J.M., Leahy, P., Foley, N. & Kiely, G.  (2014).  Spatial and seasonal variation of dissolved organic carbon (DOC) concentrations in Irish streams: importance of soil and topography characteristics.  Environmental Management 53: 959-967.

Lock, M.A., Wallace, R.R., Costerton, J.W., Ventullo, R.M. & Charlton, S.E. (1984).  River epilithon: toward a structural-functional model.  Oikos 42: 10-22.

Stevenson, R.J. (1990).  Benthic algal community dynamics in a stream during and after a spate. Journal of the North American Benthological Society 9: 277-288.

 

Empathy with the ignorant …

One of my ongoing projects is to produce a “beginner’s guide” to freshwater diatoms.  I wrote one about twenty years ago but the taxonomy is now seriously out of date.   I also illustrated it with line drawings rather than photographs.  Strange to think now but this was the last years of film photography and the quality of photomicrograph that we now take for granted was a lot harder to achieve.   Nonetheless, I like to think that it played a useful role, if not as the definitive guide to the diatoms of the UK then as a “phrase book” that helps them get understand the foreign language that hardcore diatomists talk.

Most of us refer to identification guides, informally, as “keys” yet, paradoxically, experienced biologists often skip the keys and flick through the pictures, using pattern recognition in preference to probabilistic reasoning to name specimens.  There is a reason for this: most keys are not very good.  I’ll go further: by the time a biologist has enough experience of a group of organisms to write a key, s/he has forgotten what it is like to be a beginner.   What is missing from most keys is empathy with the ignorant.

Let’s take as an example, an early couplet in the key in Freshwater Diatoms of Central Europe which asks if the cells you are looking at have internal septa.   Septa are thin, silica plates which project into the cell space (see figure a. below) and are useful diagnostic characters for some araphid diatoms, in particular.   Most keys to freshwater diatoms would have a couplet such as this at an early stage.   However, using septa as a diagnostic characteristic carries some disadvantages because they are part of the structure of girdle bands not the valve.   In the case of Tabellaria, the most common septa-bearing genus in freshwater (see image below), the cells often disintegrate during slide preparation so that our struggling beginner is faced with a few valves (b.) and a larger number of girdle bands (c. and d.).   Most of the features that are useful for identification are on the valve itself but here, just to spice up the beginner’s experience, the septa are on the girdle bands.   So the beginner has to detect a thin plate of silica (virtually a flat piece of glass) that is mounted between a slide and a coverslip (two more flat pieces of glass) in order to progress to the next step which asks about characteristics on an entirely separate structure.

Tabellaria flocculosa from the River Broom in north-west Scotland with septa indicated by arrows.   a. whole frustule in girdle view; b. valve; c. and d. girdle bands.  Scale bar: 10 micrometres (= 100th of a millimetre).

Just to make matters more confusing, some diatoms (Rhoicosphenia and some Gomphonema, for example) have “pseudosepta”, which are similar to septa but are part of the valve itself rather than the girdle band.  This should not be a problem when using keys because questions about septa usually come after questions about whether a raphe is present or not.  That should have steered our beginners away from Gomphonema and Rhoicosphenia except that one valve of Rhoicosphenia has a very short raphe that a beginner might well have missed.

Rhoicosphenia abbreviata from the River Derwent, north-east England with pseudosepta indicated by arrows.   a. and c. inner (concave) valve; b. outer (convex) valve (pseudosepta present but not in focus); d. whole frustule in girdle view.  Scale bar: 10 micrometres (= 100th of a millimetre).   Photos: Ingrid Jüttner.

Whilst, in theory, the logical structure of a key should take the user infallibly to the right taxon, in practice, users tend to use the key only until the point where they encounter a couplet that cannot be easily tackled.  At this point, they switch from probabilistic reasoning to pattern recognition – they flick through the images, in other words, until they find one that matches the specimen that they are trying to identify.   Then they use the descriptions to confirm (or not) their hunch.   The key may fail because the writer assumed too much knowledge on the part of the user, because the specimen is not “typical” (that’s for another post!) or the user’s equipment is not as good as the writer expected.   I suspect that the first of these is most often the case, because the experts who write the keys have, quite simply, forgotten what it is like to be a beginner.

I do use keys a lot when teaching because I think that the repetition of a series of (more-or-less) logical steps drives home the elements of diatom morphology that beginners need in order to put names on different genera and species.  Once they have got these basics in their heads, then I am happy for them to switch to pattern recognition rather than probabilistic reasoning.   What I suspect happens is that schemata of most of the genera get lodged in their memory, and they can then use this information to find the right set of images from which to match their unknown specimen.   The key is an important part of this process so effort put into writing a really good key should be worthwhile.

The problem lies in understanding what we mean by “really good”.   There is a risk that we define the quality of identification guides in terms of taxonomy whereas didactics plays an equally important role and we cannot assume that someone who is an expert on the former is as knowledgeable about the latter.   The Field Studies Council AIDGAP guides set a fine example by insisting on end-user testing to ensure the usability of keys, but these principles have not filtered through to the wider academic community.   Remembering what it was like to be a complete beginner is a good start.

That’s laid out the theory. Next job is to turn this into practice ….

References

More about the development of keys to identify diatoms in “The decline and fall of a CD-ROM

I also contributed to a European standard which set out the principles behind writing good keys for applied ecology:

EN 16164:2013.  Water quality – Guidance standard for designing and selecting taxonomic keys.   Comité Europeen  de Normalisation (CEN), Geneva,  12pp.

The River Wear in January

The series of events that eventually gave birth to this blog started with a visit to the River Wear at Wolsingham on the first day of 2009.  I had visited on a whim, intending to blow away the cobwebs after lunch on New Year’s Day, but with no real plan.  But I thought it would be interesting to pull on my waders and have a look at the river bed and, while I was there, I may as well collect a sample too.   Those observations and that sample must have triggered something in my mind, because I returned every month after that and, on each occasion, the samples and observations generated sketches which, in turn, made me curious about the factors that drove the algal communities in our rivers.

I thought it would be interesting to repeat that exercise during 2018 as my thinking has moved on over the past nine years.  I’m essentially visiting the same site and making the same observations but, this time, filtering them through deeper beds of experience.   The River Wear at this point is about 30 metres wide, a broad, shallow, riffled stretch, skirting the small town of Wolsingham roughly at the point where Weardale broadens out from a narrow Pennine valley to the gentler landscape of the Durham coalfield.  There are a couple of small towns upstream but the ecological condition of the river is still good.  Although there are still concerns about concentrations of heavy metals arising from the mines that are scattered around the upper parts of the valleys, I can see no serious effects of toxic pollution when I look at the plants and animals that live at Wolsingham.

If you follow this blog you will not be surprised to hear that, even in the depths of winter, algal communities in the River Wear are thriving Most of the larger stone surfaces are covered with a discernible brown film, up to a couple of millimetres thick.   The very top layer is dark brown in colour, with a lighter brown layer beneath this.   When I put a sample of this under my microscope, I saw that it was dominated by gliding cells of Navicula lanceolata, though other diatoms were also present (described in more detail in “The ecology of cold days”) and there were also a few thin filaments of a blue-green alga.

A submerged cobble photographed in situ in the River Wear at Wolsingham, January 2018, covered with a thick diatom-dominated biofilm.

I’ve included a picture of the view down my microscope because one of the questions that I’ve been trying to answer over the past few years is how we construct an understanding of the microscopic world using microscopy (see “The central dilemma of microscopy” and “Do we see through a microscope?”).   Of course, a single view field of view does not convey all the information I require, so my understanding is actually built up from observations of a large number of separate fields.  The boat-shaped cells of Navicula lanceolata were almost ubiquitous in these, as were patches of amorphous organic matter (“fine particulate organic matter” – see “A very dilute compost heap …”).  In total, I found 15 different species of algae in my preliminary analysis, of which Navicula lanceolata comprised about half of the total, with thin filaments of the cyanobacterium Phormidium and the diatom Achnanthidium minutissimum each constituting about 15 per cent.

A view of the biofilm from the River Wear, Wolsingham in January 2018.

However, my earlier comment about the biofilms having distinct layers means that simply observing what organisms are present will not tell us the whole story about how those organisms are organised within the biofilm (see “The multiple dimensions of submerged biofilms …”) so the next step is to hypothesise how these organisms might be arranged in the biofilm before I disrupted their microhabitat with my sampling.   The schematic diagram below attempts to capture this, but with a few provisos.  First, I said that the biofilm was a couple of millimetres thick but my portrayal only shows about a tenth of a millimetre; second, there is considerable spatial and temporal variation in biofilms and my depiction amalgamates my direct observations in January 2018 with information gleaned from a number of other visits.   Gomphonema olivaceum (probably a complex of two or three species in this particular case), for example, is often more prominent than it was last week, and I have also omitted Achnanthidium minutissimum altogether.   I suspect that this is less abundant in the mature biofilms but that the cobble surface is a patchwork of different thicknesses, reflecting different types of disturbance.   That raises another issue: the scale at which we generally collect samples is greater than the scales at which the forces which shape biofilms operate.   The whole image below, for context, occupies about the same width as a single bristle on the toothbrush that I used to collect the sample.

It is difficult to convert what we “see” back to the original condition when working under such constraints and, inevitably, decisions are guided by what others before us have written.  That brings a different set of problems: Isaac Newton may have seen further by “standing on the shoulders of giants” but Leonardo da Vinci’s usually rigorous objectivity lapsed on at least one occasion when his eye was led by assumptions he had inherited from earlier generations (see “I am only trying to teach you to see …”).   What my picture is actually showing, in other words, is a mixture of what I saw and what I think I should have seen.   This two-way process in art extends from the very earliest drawings we make through to the most sophisticated Old Masters so I am in good company.  In truth, I am not trying to depict a particular point in space or time so much as to encapsulate the idea of a biofilm from that river that is more than a random aggregation of cells.

A schematic view of the vertical structure of a submerged biofilm from the River Wear, Wolsingham, January 2018.   a., Navicula lanceolata (valve view); b., N. lanceolata (girdle view); c. Navicula gregaria (valve view); d. N. gregaria (girdle view); e. Gomphonema olivaceum (valve view); f. G. olivaceum (girdle view); g. Phormidium; h. inorganic particles; i. fine particulate organic matter.  Scale bar: 20 micrometres (= 1/40th of a millimetre).

References

You can find out more about the condition of the River Wear (or any other river or lake) using the Environment Agency’s excellent Catchment Planning webpages

Three good books that discuss the relationship between pictorial representation and the mind are:

Cox, Maureen (1992).  Children’s Drawings.   Penguin, Harmondsworth.

Gombrich, E.H. (1977) Art and Illusion: a study in the psychology of pictorial representation.   5th Edition.  Phaidon, London.

Hamilton, James (2017).  Gainsborough: a Portrait.   Weidenfield & Nicholson, London.

 

 

Algae behaving selfishly …

My most recent trip to Ennerdale Water was on a wonderful windless winter day, offering perfect reflections of the snow-dusted peaks beyond the lake. It was a cold day but I was well wrapped-up and could enjoy both the long-distance views and the close-ups of nature around the lake’s margins.   One of the small streams that I crossed as I skirted the perimeter of the lake had patches of green algae growing on its submerged stones and even a quick examination showed it to be coarser than the green algae that covered most of the larger stones on the lake bed itself, as well on those in the River Ehen, just below the outfall.   When I managed to get specimens under my microscope I saw that the algae on the lake bed was Spirogyra (which I have seen here before; see “A lake of two halves”) whilst that in the inflow stream was Oedogonium.

I’ve written about Oedogonium before, and lamented the problems we face when we try to identify the species within this large genus (see “The perplexing case of the celibate alga”).   Ironically, a couple of weeks after I wrote this, I encountered a population of Oedogonium in another Cumbrian stream that did have sexual organs (see “Love and sex in a tufa-forming stream”).  However, this was the exception that proves the rule, as I have not seen a sexually-mature population of Oedogonium since.  The population I found beside Ennerdale was not sexually mature either but it did show a different, but equally effective, means of going forth and multiplying.

In the left hand diagram below we see a vegetative cell from an Oedogonium filament that has split open, allowing a vesicle to be extruded within which a single zoospore has formed.   This has a ring of flagella at one end, resembling a monk’s tonsure (you can just see these flagella in the photograph).   The other two photographs show the monk’s bald pate, though the fringe of flagella is not very clear.    The transparent vesicle swells and eventually ruptures, releasing the zoospore, which swim around for an hour or so, before settling on a new substratum and growing into new filaments.

Zoospores of Oedogonium from a stream flowing into Ennerdale Water, January 2018.   Scale bar: 25 micrometres (= 1/40th of a millimetre). 

In my material, the new filaments were mostly attached to mature Oedogonium filaments; however, this is probably partly an artefact and, in the field, they would almost certainly also settle on rocks and other surfaces too.   You can see, in the diagram below, how the “bald” end of the zoospore has started to differentiate into a holdfast that will secure the cell to the substrate whilst, over time, the other end will start to divide to produce the first cells of the new filament.  The whole process is described in a series of papers by Jeremy Pickett-Heaps (see reference list below).

Why did I see zoospore formation in this particular sample?   I don’t know for sure but it may be because I let a longer than usual time elapse between collecting and examining the sample.   This one had sat around in a cool box and fridge for four days, whereas I usually manage to check them within 24 hours.   Neglect can be a useful tool in the phycologist’s arsenal, as many freshwater algae see no need to indulge in anything more taxing than routine cell division for as long as the habitat keeps them replenished with whatever light, nutrients and other resources that they need.   Only when this is no longer the case do the algae start to channel resources into survival strategies.

Oedogonium zoospores germinating into new filaments, both epiphytic on mature filaments.   From a stream flowing into Ennerdale Water, January 2018. .   Scale bar: 25 micrometres (= 1/40th of a millimetre). 

Although I used the phrase “go forth and multiply” in an earlier paragraph, these Oedogonium cells are actually “going forth” rather than “multiplying” as the process we are watching only produces a single new cell.  However, were this zoospore to be released in a stream rather than a sample bottle, then there is a good chance that it would have been washed downstream and that a few of the many zoospores might have settled on a suitable habitat away from the constraints of their former home.   Asexual reproduction is a dispersal mechanism that results in the spread of genetically-identical copies of the parent cell.  For a sessile organism, this strategy allows a single genotype to move on from less-favourable locations and to exploit the potential of nearby locations.

The word “reproduction” is misleading as the mixing of genetic material that we associate with sex doesn’t take place.  The end product is a clone of a successful Oedogonium filament growing somewhere else.   However, taking the “sex” out of “asexual” removes a huge potential for innuendo, and readers who have battled this far through a post on nondescript green filaments deserve a reward.  So let’s finish with Woody Allen’s definition of masturbation as “sex with someone you love” and suggesting that the cytological huffing and puffing involved in zoospore production may not have the romance of sex but nor does it lead to any of the complications which result from sex either.   The alga gets offspring that are 100% identical to itself, just slightly further downstream and there is no risk of mixing with inferior genotypes.   That’s about as “selfish” as the “selfish gene” can get.

References

Pickett-Heaps, J. (1971).   Reproduction by zoospores in Oedogonium. I. Zoosporogenesis.   Protoplasma 72: 275-314.

Pickett-Heaps, J. (1971).   Reproduction by zoospores in Oedogonium. II. Emergence of the zoospore and the motile phase. Protoplasma 74: 149-167.

Pickett-Heaps, J. (1972).   Reproduction by zoospores in Oedogonium. III. Differentiation of the germling.  Protoplasma 74: 169-173.

Pickett-Heaps, J. (1972).   Reproduction by zoospores in Oedogonium. IV. Cell division in the germling and the possible evolution of the wall rings.   Protoplasma 74: 195-212.

See also “The River Ehen in March” for some further perspectives on asexual reproduction in algae.

View from near our sampling site on Croasdale Beck, looking towards Ennerdale Bridge, January 2018.

 

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.

Diatom hunting in the Pirin mountains

I started 2018 peering down my microscope at a sample that I collected whilst in Bulgaria back in the summer.   I have written about my trip to the Pirin mountains before (see “Desmids from the Pirin mountains”) but the diatom sample that I collected from Lake Popovo had remained unexamined since I got back.

I had waded into the littoral zone of this steep-sided corrie lake and picked up a few of the smaller stones, which I had then scrubbed with the toothbrush stowed in my rucksack to remove the thin film of diatoms.  These, like most of the algae that I collect on my travels, get treated to a bath in local spirits to ease the journey back to the UK.  This is not an ideal preservative for soft-bodied algae but is not a problem when your primary interest is diatoms with their tough silica cell walls.  Once I got back, I had them prepared and mounted ready for inspection, but then got distracted by other things and have only just got around to having a proper look.

The two most abundant taxa were the Achnanthidium minutissimum complex (probably at least three species) and Cymbella excisiformis.  Together, these constituted over eighty per cent of all the diatoms in the sample.  Ten years ago, I would have called the organism I was calling Cymbella excisiformis by a different name, Cymbella affinis, but opinions have shifted more than once.  The original Diatomeen im Süsswasser-Benthos von Mitteleuropa has images of C. affinis that are actually C. tumidula, and also describes C. excisa as a separate species.   However, the most recent view is that C. affinis and C. excisa are two names for the same species, with C. affinis taking precedence.   To confuse matters yet further, the population illustrated below shows a gradation of features from “C. affinis” to “C. excisiformis”, suggesting that the use of length:width as a discriminating factor is over simplistic.  Krammer tried to explain his rationale for distinguishing between these species in his 2002 monograph but he uses the name “C. excisa” for the organism called “C. affinis” in our 2017 English edition.  Confused?  You will be ….

Cymbella excisiformis” from Lake Popovo, Pirin Mountains, Bulgaria, August 2017.   Based on Lange-Bertalot et al. (2017)’s criteria of length:breadth 4.2-5.3 in C. excisiformis compared to 3.1 – 3.8 in C. affinis, images a., b. and c. are C. excisiformis whilst d., e., f. and g. are C. affinis.   Scale bar: 10 micrometres (= 1/100th of a millimetre).

This is another good example of points that I have made several times before: that we should always try to identify populations rather than single cells, and that we should treat dimensions stated in the literature as indicative rather than definitive (see “More about Gomphonema vibrio”).    Length:width, in particular, can change a lot during the life-cycle of the diatom.

Species of Gomphonema were also present in the sample.  Though not numerically abundant (none constituted more than one per cent of the total count), they included some large cells which, in addition, have extensive mucilaginous stalks, so their contribution to total biomass is greater than their low abundance suggests.   I’ll write more about the ecology of these species in the next post.   Finally, I also found other Cymbella species, as well as some Encyonema and Encyonopsis, and a few valves of Eucocconeis flexella, a relative of Achnanthidium and Cocconeis which has a distinctive diagonal raphe.

Gomphonema spp. from Lake Popovo, Pirin mountains, Bulgaria, August 2017.   h., i.: G. acuminatum; j.: G. truncatum; k.: unidentified girdle view; l.: G. pumilum.  Scale bar: 10 micrometres (= 1/100th of a millimetre).

There is, at this point in time, no official Bulgarian method for assessing the ecological status of lakes using diatoms so I have evaluated Lake Popovo as if it were a low alkalinity lake in the UK instead.  Using the method we developed, this one sample has an Ecological Quality Ratio of 0.92, which puts it on the border between high and good status.   Looking around the lake, I see no reason why it should not be firmly in high status but, at the same time, I am using an evaluation method that was designed for lakes 2000 kilometres away, so maybe we should not expect perfect results.    However, I have performed similar exercises at other lakes far from the UK and also got similar results (see “Lago di Maggiore under the microscope”) which points to a basic robustness in this approach.

The outflow of Lake Popovo leads into a cascade that ends in the first of a series of lakes, the “Fish Popovski” lakes.   I wrote about the desmids in this lake back in September (see “”Desmids from the Pirin mountains”) and will return to this sample in order to describe the diatoms in another post.   But, meanwhile, the assemblage at Popovo reminded me of the littoral algae in another lake that I really should tell you about …

Miscellaneous diatoms from Lake Popovo, Pirin mountains, Bulgaria, August 2017.   m.: Cymbella sp.; n.: Encyonema neogracile; o. and p.: Eucocconeis flexella (raphe valve and girdle view respectively).  Scale bar: 10 micrometres (= 1/100th of a millimetre).

Lake Popovo, photographed from close to the location from which my sample was collected.  The brass plate on the rock at the right hand side gives the altitude as 2234 metres above sea level.  The photograph at the top of the post shows Lake Popovo against a backdrop of the Pirin mountains.

References

Hofmann, G., Werum, M. & Lange-Bertalot, H. (2011).   Diatomeen im Süßwasser-Benthos von Mitteleuropa. A.R.G. Gantner Verlag K.G., Rugell.

Krammer, K. (2002).  Diatoms of Europe volume 3: Cymbella.   A.R.G. Gantner Verlag K.G., Ruggell, Germany.

Lange-Bertalot, H., Hofmann, G., Werum, M. & Cantonati, M. (2017).   Freshwater Benthic Diatoms of Central Europe: Over 800 Common Species Used In Ecological Assessment (edited by M. Cantonati, M.G. Kelly & H. Lange-Bertalot).   Koeltz Botanical Books, Schmitten-Oberreifenberg.

The UK lake diatom assessment method is described in:

Bennion, H., Kelly, M.G., Juggins, S., Yallop, M.L., Burgess, A., Jamieson, J. & Krokowski, J. (2014).  Assessment of ecological status in UK lakes using benthic diatoms.  Freshwater Science 33: 639-654.

Details of the calculation can be found in the UK TAG method statement.