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.

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It’s all about the algae

Just a short post to point you all towards an article I wrote for Royal Society of Biology’s magazine The Biologist.  It is a broad overview of the reasons why we use algae to assess the condition of our lakes and rivers in Europe and is illustrated with three of Chris Carter’s beautiful images, and the print edition will have even more of these.  Take the figure legends with a pinch of salt (we didn’t write these!): neither Tolypella nor Chaetophora are particularly common in the UK.   Navicula, on the other hand, is common but the legend makes no mention of this.

Whilst I have your attention, I will also point you towards a short article that I wrote for the most recent Phycological Bulletin, the newsletter of the Phycological Society of America.  This offers a few more hints to anyone thinking about entering the Hilda Canter-Lund competition next year.

Certainly uncertain …

Back in May I set out some thoughts on what the diatom-based metrics that we use for ecological assessment are actually telling us (see “What does it all mean?”).  I suggested that diatoms (and, for that matter, other freshwater benthic algae) showed four basic responses to nutrients and that the apparent continua of optima obtained from statistical models was the result of interactions with other variables such as alkalinity.   However, this is still only a partial explanation for what we see in samples, which often contain species with a range of different responses to the nutrient gradient.  At a purely computational level, this is not a major problem, as assessments are based on the average response of the assemblage. This assumes that the variation is stochastic, with no biological significance.  In practice, standard methods for sampling phytobenthos destroy the structure and patchiness of the community at the location, and our understanding is further confounded by the microscopic scale of the habitats we are trying to interpret (see “Baffled by the benthos (1)”).  But what if the variability that we observe in our samples is actually telling us something about the structure and function of the ecosystem?

One limitation of the transfer functions that I talked about in that earlier post is that they amalgamate information about individual species but do not use any higher level information about community structure.  Understanding more about community structure may help us to understand some of the variation that we see.   In the graph below I have tried to visualise the response of the four categories of response along the nutrient/organic gradient in a way that tries to explain the overlap in occurrence of different types of response.   I have put a vertical line on this graph in order that we can focus on the community at one point along the pollution gradient, noting, in particular, that three different strategies can co-exist at the same level of pollution.  Received wisdom amongst the diatom faithful is that the apparent variation we see in ecological preferences amongst the species in a single sample reflects inadequacies in our taxonomic understanding.  My suggestion is that this is partly because we have not appreciated how species are arranged within a biofilm.  I’ve tried to illustrate this with a diagram of a biofilm that might lead to this type of assemblage.

Schematic diagram showing the response of benthic algae along a nutrient/organic gradient.  a.: taxa thriving in low nutrient / high oxygen habitats; b.: taxa thriving in high nutrient / high oxygen habitats; c.: taxa thriving in high nutrient / low oxygen habitats; d.: taxa thriving in high nutrients / very low oxygen habitats.   H, G., M, P and B refer to high, good, moderate, poor and bad ecological status.

The dominant alga in many of the enriched rivers in my part of the world is the tough, branched filamentous green alga Cladophora glomerata.   This, in turn, creates micro-habitats for a range of algae.  Some algae, such as Rhoicosphenia abbreviata, Cocconeis pediculus and Chamaesiphon incrustans, thrive as epiphytes on Cladophora whilst others, such as C. euglypta are often, but not exclusively, found in this microhabitat.  Living on Cladophora filaments gives them better access to light but also means that their supply of oxygen is constantly replenished by the water (few rivers in the UK are, these days, so bereft of oxygen to make this an issue).   All of these species fit neatly into category b. in my earlier post.

Underneath the Cladophora filaments, however, there is a very different environment.  The filaments trap organic and inorganic particulate matter which are energy sources for a variety of protozoans, bacteria and fungi.   These use up the limited oxygen in the water, possibly faster than it can be replenished, so any algae that live in this part of the biofilm need to be able to cope with the shading from the Cladophora plus the low levels of oxygen.   Many of the species that we find in highly polluted conditions are motile (e.g. Nitzschia palea), and so are able to constantly adjust their positions, in order to access more light and other resources.   They will also need to be able to cope with lower oxygen concentrations and, possibly, with consequences such as highly reducing conditions.  These species will fit into categories c. and d. in the first diagram.

A stylised (and simplified) cross-section through a biofilm in a polluted river, showing how different algae may co-exist.   The biofilm is dominated by Cladophora glomerata (i.) with epiphytic Rhoicosphenia abbreviata (ii.), Cocconeis euglypta (iii.) and Chamaesiphon incrustans (iv.) whilst, lower down in the biofilm, we see motile Nitzschia palea (v.) and Fistulifera and Mayamaea species (vi.) growing in mucilaginous masses.

However, as the cross-section above represents substantially less than a millimetre of a real biofilm, it is almost impossible to keep apart when sampling, and we end up trying to make sense of a mess of different species.   The ecologists default position is, inevitably, name and count, then feed the outputs into a statistical program and hope for the best.

A final complication is that river beds are rarely uniform.  The stones that make up the substrate vary in size and stability, so some are rolled by the current more frequently than others.  There may be patches of faster and slower flow associated with the inside and outsides of meanders, plus areas with more or less shade.   As a result, the patches of Cladophora will vary in thickness (some less stable stones will lack them altogether) and, along with this, the proportions of species exhibiting each of the strategies.  The final twist, therefore, is that the vertical line that I drew on the first illustration to illustrate a point on a gradient is, itself, simplistic.  As the proportions vary, so the position of that line will also shift.  Any one sample (itself the amalgamation of at least five microhabitats) could appear at a number of different points on the gradient.  Broadly speaking, uncertainty is embedded into the assessment of ecological status using phytobenthos as deeply as it is in quantum mechanics.  We can manage uncertainty to some extent by taking care with those aspects that are within our control.   However, in the final analysis, a sampling procedure that involves an organism 25,000 times larger than most diatoms blundering around a stream wielding a toothbrush is invariably going to have limitations.

The same schematic diagram as that at the start of this article, but with the vertical line indicating the position of a hypothetical sample replaced by a rectangle representing the range of possibilities for samples at any one site. 

Notes from Berlin

Whenever I pass though Liverpool Street Station on the eastern side of London, I always take a moment to pause beside “Kindertransport – the arrival” sculptures by Frank Meisler.   They show Jewish refugee children arriving in Britain with their small amounts of luggage, fleeing from Nazi persecution.   In this age of fear and distrust of migration, they remind me that these fluxes are nothing new, and remind me that we need to show generosity towards the helpless and dispossessed.  As I walked past Friedrichstrasse station during a visit to Berlin last week I saw a companion piece to the Liverpool Street sculptures.  However, the statue at Friedrichstrasse (“Züge in das leben, Züge in den tod, 1938 – 1945” – literally: “trains to life, trains to death”, also by Frank Meisler) differs in one important respect: the German children at the start of their journey are facing in two directions: some are awaiting the Kindertransports to the west but two are heading towards the death camps.

Züge in das leben, Züge in den tod, 1938 – 1945.  Bronze sculpture by Frank Meisler at Freidrichstrasse station, Berlin (photo taken in low light – apologies!).  The photograph at the top shows “Kindertransport – the arrival, also by Frank Meisler, at Liverpool Street Station, London.

Berlin does not flinch at confronting its recent past.   I spent the hour or so between the end of my meeting and my trip out to the airport at “Topographie des Terrors” – a museum on the site of the former Gestapo and SS Headquarters.  It was a reminder of what the children depicted in Meisler’s sculptures were fleeing from.  After a gruelling hour, I was ready for a change of atmosphere and turned towards the Martin Gropius Bau – one of Berlin’s best galleries, just a short distance away.  By coincidence, one of the exhibitions was a series of etchings by Lucien Freud, himself a Jewish refugee (though not part of the Kindertransport).   Freud’s portraits have an intensity that can be unsettling to a viewer; however, seen immediately after my immersion in Nazi atrocities, they had the opposite effect.  I was left wondering how my reactions would have differed had I seen the exhibition before, rather than after, Topographie des Terrors.  The irony of a Jewish refugee’s art being exhibited so close to the former headquarters of the powers that forced him to flee in the first place did not escape me either.

A year ago, I was here on the night that Donald Trump was elected president of the USA (see “Remembrance in Berlin”).   This time, my visit started just a day after the AfD (“Alternative for Germany”) won 74 seats in the Bundestag at the federal elections, becoming the third largest party.   Anger at Germany’s own tolerant policy to refugees was one of the reasons that they did so well, particularly in the former eastern states.   As I write, I am not sure how this meshes with my statement about Berlin’s ability to confront the less savoury aspects of its past.  There is a willingness to do this within the Federal government, and the middle-class, well-educated Germans I meet share a desire to look back objectively.  The rise of AfD is worrying partly because it has happened in a country that has tried so hard to learn from its past.   Let’s hope that Germany does not forget to remember ….

The Martin Gropius Bau rising above a section of the Berlin wall.  Topographies des Terrors is behind the wall.  The right hand image shows the row of cobblestones marking the path of the wall in places where it is no longer standing.

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).

Lessons from School Knott Tarn …

As not everyone could join us on our excursion on Friday afternoon, we repeated the exercise on Saturday morning, heading to a small tarn just a short walk from Windermere and Bowness.   Despite its proximity to two of the busiest towns in the Lake District, there were very few other people around to disturb our peace whilst we collected samples.   As at Kelly Hall and Long Moss Tarns, Dave had his plankton net out, but we also explored a boggy region at one end, finding more patches of Sphagnum but also extensive growths of Utricularia minor (Lesser Bladderwort), one of a small number of aquatic carnivorous plants.   Dave was particularly pleased by this find as he associates this particular plant with rich hauls of desmids.

It was tempting to linger in the sunshine beside School Knott Tarn but the green tinge of the water that dripped out of the Sphagnum squeezings in particular was enough to lure us towards the Freshwater Biological Association’s laboratories in order to start examining our samples.

Utricularia minor (Lesser Bladderwort) from School Knott Tarn, near Windermere, September 2017.   Several of the spherical bladders which trap small invertebrates are visible on the plant.

My selection of photographs below shows just a part of the diversity that we encountered during our microscopic examinations.  I was using a borrowed set-up and the images are all from photographs of the desmids displayed on computer monitor, which is far from ideal.   Some of the larger desmids – one large Closterium species in particular – were too large to fit onto the screen and have had to be omitted from this account.  There were also a number of cells of Eremosphaera (see “More from Loughrigg Fell”) and some Cyanobacteria (Merismopedia was quite common) so this is a very partial description of our microscopical adventures in School Knott Tarn.

The first two desmids, Spirotaenia condensata and Cylindrocystis gracilis, belong to a group of desmids called “saccoderm desmids”.  These are more closely related to filamentous green algae of the Zygnemetaceae that are old friends of this blog (see “Concentrating on Carbon, for example) and, in fact, we could think of these genera as being unicellular analogues of their filamentous cousins.   Spirotaenia, with its helical chloroplast, for example, recalls Spirogyra whilst Cylindrocystis’ two star-shaped chloroplasts is reminiscent of Zygnema.  Mesotaenium, which we did not see in this sample, has a plate-like chloroplast similar to that in Mougeotia.

The next two illustrations both show species of Micrasterias.  Of these, M. compereana generated a vigorous discussion amongst our experts. This would have been described as M. fimbriata using the latest British floras but a paper has been published recently which uses molecular data to demonstrated the need to split the species. Finally, we have representatives of Euastrum and Haplotaenium, two genera that we also met at Dock Tarn (see “Damp days in search of desmids …”) although the species are different.   Haplotaenium differs from Pleurotaenium in the number and form of the chloroplasts and also because it lacks a terminal vacuole.

Desmids from Sphagnum squeezings from School Knott Tarn, September 2017: a. Spirotaenia condensata; b. Cylindrocystis gracilis; c. Micrasterias compereana; d. Micrasterias crux-meltensis; e. Euastrum oblongum; f. Haplotaenium rectum.  Scale bar: 25 micrometres (= 1/40th of a millimetre).

Four more desmids are illustrated on the lower plate.   Of these, we have seen Netrium digitus in Dock Tarn and the illustration there is better than this one, showing the undulating nature of the chloroplast margins quite clearly.   The desmid below this, Closterium closterioides caused some confusion at first.   We usually associate Closterium with lunate (moon-shaped) cells (see “More from Loughrigg Fell”) but this species is straight, sending me towards the section on Netrium in my Flora.  However, Netrium lacks terminal vacuoles whereas this specimen has prominent vacuoles at both ends.   We also found a variety, C. closterioides var. intermedium, in the same sample.

The final desmid that I have illustrated is a filamentous form: Desmidium schwartzii.  In contrast to Hyalotheca dissilens (see “Desmids from the Pirin mountains”) there is no obvious mucilaginous sheath around this specimen, but this may be an anomaly of this population or an artefact of the microscopy set-up.   We are looking at the side view of a chain of cells but if we were to look at the end view of one cell it would be triangular in this particular species.  The chloroplast fills most of the cell and has projections running into the corners of the cells.  However, as the filaments of the cells are slightly twisted, these projections appear to shift in position from cell to cell, giving a helical appearance.  I’ve tried to illustrate this with a schematic diagram.

More desmids from Sphagnum squeezings from School Knott Tarn, September 2017: g. Netrium digitus; h. Closterium closterioides var. closterioides; i. C. closterioides var. intermedium; j. Desmidium schwartzii Scale bar: 25 micrometres (= 1/40th of a millimetre).

This short post gives some idea of the diversity in a single sample from a single Tarn.   Dave handed all the samples we collected over to David Williamson on his way back south and we’ll get a fuller list of their diversity in due course.  This one sample occupied me for the latter part of Saturday morning and all of the afternoon.   On Sunday, I moved on to look at another sample and I’ll write about that in another post very soon.

A schematic view of a chain of Desmidium cells, showing the arrangement of the chloroplast seen in apical view (k.) and the implications of slight twisting of the filament on appearance (l.).  Diagram adapted from John et al. (2011).

Reference

John, D.M., Whitton, B.A. & Brock, A.J. (2011). The Freshwater Algal Flora of the British Isles. 2nd Edition. Cambridge University Press, Cambridge.

Neustupa, J., Šťastný, J. & Škaloud, P. (2014). Splitting of Micrasterias fimbriata (Desmidiales, Viridiplantae) into two monophyletic species and description of Micrasterias compereana sp. nov.  Plant Ecology and Evolution 147: 405-411.