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.

 

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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.

 

 

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.