Summertime blues …

My reflections on the effects of the heatwave on freshwater algae continued with the latest of my regular visits to the River Wear at Wolsingham.  A comparison of the picture above with that at the head of “Spring comes slowly up this way …” says it all: the sun was shining and the gravel berms that I usually use to enter the river were occupied by families with barbeques whilst their children splashed around in the water.   At times such as this, a grown man picking up stones and then vigorously brushing their tops with a toothbrush would have invited too many questions, so I slunk off 100 metres or so downstream and found a quieter spot to explore.

The biofilm in the main channel of the River Wear at Wolsingham, July 2018. 

The first thing I noticed was that the biofilm coating the submerged stones at the bottom of the river had a greenish tinge, rather than its usual chocolate brown appearance.  It also was more crusty and less slimy to the touch than I usually see in this river.  When I got a specimen under the microscope, I could see that the composition was completely different to that which I had observed in previous months.   Most samples from this location that I’ve looked at in the past have been dominated by diatoms, with occasional spring flourishes of filamentous green algae.  Today, however, the sample was dominated by small green algae – a group that I am not very confident at identifying.   My rough estimate is that these formed about three quarters of all the algae that I could see, with diatoms and cyanobacteria each accounting for about half of the remainder.   The most abundant greens were a tiny single-celled alga that I tentatively identified as Keratococcus bicaudatus, along with a species of Scenedesmus and Desmococcus communis.   There were also a number of cells of Monoraphidium arcuatum and some of Ankistrodesmus sp.

Two views of biofilms from the River Wear, Wolsingham in January 2018.   Left: from the main channel; right: from pools at the edge of the channel.

Green algae from the River Wear at Wolsingham, July 2018: a. Desmococcus communis; b. Monoraphidium arcuatum; c. Scenedesmus sp.; d. unidentified, possibly Keratococcus bicaudatus.  Scale bar: 10 micrometres (= 1/100th of a millimetre).

However, there were also pools at the side of the channel, away from the main current but not so cut off that they were isolated from the river itself.   These were dominated by dense, brown filamentous growths, very similar in appearance to the Melosira varians flocs I described in “Some like it hot …”.  The filaments, however, felt coarser to the touch and, in close-up, could be seen to be branched, even without recourse to a microscope.   Once I got these under the microscope, I could see that they were filaments of Cladophora glomerata, another green alga, but so smothered with epiphytic diatoms (mostly Cocconeis pediculus) that they appeared brown in colour.

This combination of Cladophora glomerata and Cocconeis pediculus in the backwaters were as much of a surprise as the green-algae-dominated biofilms in the main channel.   These are species usually associated with enriched rivers (see “Cladophora and friends”) and, whilst I have seen Cladophora in the upper Wear before, it is an unusual occurrence.   Just as for the prolific growths of Melosira varians described in “Some like it hot …” it is tempting to leap to the conclusion that this must be a sign that the river is nutrient-rich.  However, the same conditions will apply here as there: “nutrients” are not the only resource that can limit plant growth and a steady trickle of phosphorus combined with warm, sunny conditions is just as likely to lead to prolific growths as a more conventionally “polluted” river.

Cladophora filaments smothered by the diatom Cocconeis pediculus in a pool beside the River Wear at Wolsingham, July 2017.   The frame width of the upper image is about 1 cm; the scale bar on the lower images is 20 micrometres (= 1/50th of a millimetre).

Another way to think of these situations is that, just as even healthy people are occasionally ill, so healthy streams can go through short periods when, based on a quick examination of plants and animals present, they exhibit symptoms associated with polluted conditions or simply (as for the first sample I described) different to what we usually expect to find.   A pulse of pollution might have passed downstream or, as seems to be happening at the moment, an unusual set of conditions lad to different organisms thriving.   Just as the ability to fight off infection forms part of a doctor’s understanding of “health”, so I expect that the River Wear will, in a few weeks time, be back to its usual state.   Healthy ecosystems, just like healthy humans, show “resilience”.   The irony is that, in this case, the “symptoms” are most obvious at a time when we are enjoying a summer better than any we’ve had in recent years.

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A question of scale …

It has taken some time to convert the observations from my last visit to the River Wear (see “Spring comes slowly up this way …”) into a picture.  Then, if you remember, the river was balanced between its “spring” and “summer” guises, the cool, wet weather that we experienced in March seems to have held the plants and animals that I usually see at this time of year back.   The result was a patchiness that was easy to see with the naked eye, but harder to visualise at the microscopic level.

First there were quite a few diatoms, Achnanthidium minutissimum in particular, – that suggested a thin biofilm subject to grazing by invertebrates (and I could see some chironomid larvae moving amongst the biofilm as I was sampling).   However, there were also diatoms such as Ulnaria ulna and Gomphonema olivaceum that suggested a thicker biofilm.    And finally there were filaments of the green alga Stigeoclonium tenue, mostly in discrete patches.   I never see healthy filaments of Stigeoclonium tenue smothered in epiphytes, which I have always assumed to be due to the copious mucilage that surrounds the plants.  However, I wondered if, nonetheless, Stigeoclonium contributes to overall habitat patchiness for the diatoms, as they subtly alter the way that water flows across the stone, reducing drag and shear stress in a way that favours Gomphonema and Ulnaria.   This is just speculation, of course…

That brings me back to a familiar theme: the problems of understanding the structure of the microscopic world (see “The River Wear in January” and “Baffled by the benthos (1)”) and, tangentially, to a paper on organisms’ responses to climate that was quoted at a scientific meeting I attended recently.   In this, Kristen Potter and colleagues demonstrated that there was typically a 1000 to 10,000 fold difference between the scale at which the distribution of organisms is studied and the size of those organisms.   That might be enough to draw out some coarse-scale patterns in distribution of species, but organisms actually live in microclimates, which may be patchy and which can often be quite different to the prevailing macroclimate (the difference between being exposed to full sun in open grassland and in the shade of a forest being a good example).   They suggested that the ideal spatial resolution is between one and ten times the organism’s length/height.

I see no reason why the same challenge should not also apply to the pressures faced by organisms in rivers where, again, we can get a certain amount of useful information from a coarse analysis of distribution in relation to (let’s say) average nutrient concentrations in a reach, but cannot really understand the reasons behind the spatial and temporal variation that we see in our data.  This mismatch between the scale at which organisms respond and the scale at which we study them is, I suspect, an even bigger problem for those of us who study the microscopic world.

A second illustration came at the same meeting in a talk by Honor Prentice from the University of Lund in Sweden.  She was dabbling in molecular biology years before this became a fashionable pastime for ecologists and has, over her career, developed some fascinating insights into how the structure of both plant communities and populations of individuals vary over short distances.  Her work has focussed on the island of Öland in Sweden which has the largest extent of alvar (limestone pavement) in Europe.   The system of grikes (the slabs) and clints (the fissures which separate the grikes) create quite different microclimates – the cool, moist conditions in the latter can create bog-like conditions with much lower pH than the limestone clints.   These differences influence not just the composition of the community but the genetic structure of species within these communities too.  I left thinking that if she could detect such differences at a scale barely more than one order of magnitude greater than the organisms, then how much more variation am I missing, with perhaps a five order of magnitude difference between organism size and sampling scale?

Based on these two studies, we would need to sample biofilms at a scale of about 1 mm2 in order to get a meaningful understanding of habitat patchiness in stream benthic algae.  That might just be possible with Next Generation Sequencing technologies, though I am not sure how one would go about collecting environmental data at that scale needed to explain what is going on.  Meanwhile, I am left with the coarse approach to sampling that is inevitable when you are five orders of magnitude bigger than the organism that you want to collect, and my imagination.

References

Potter, K.A., Woods, H.A. & Pincebourde, S. (2013).  Microclimatic changes in global change biology.   Global Change Biology 19: 2932-2939.

Prentice, H.C., Lonn, M., Lefkovitch, L.P. & Runyeon, H. (1995).   Associations between allele frequencies in Festuca ovina and habitat variation in the alvar grasslands on the Baltic island of OlandJournal of Ecology 83: 391-402.

 

Spring comes slowly up this way …*

I took a few minutes out on my trip to Upper Teesdale to stop at Wolsingham and collect one of my regular samples from the River Wear.  Back in March, I commented on the absence of Ulothrix zonata, which is a common feature of the upper reaches of rivers such as the Wear in early Spring (see “The mystery of the alga that wasn’t there …”).   I put this down to the unusually wet and cold weather that we had been experiencing and this was, to some extent, confirmed by finding prolific growths of Ulothrix zonata in late April in Croasdale Beck (see “That’s funny …”).   Everything seems to be happening a little later than usual this year.   So I should not have been that surprised to find lush growths of green algae growing on the bed of the river when I waded out to find some stones from which to sample.

These growths, however, turned out to be Stigeoclonium tenue, not Ulothrix zonata (see “A day out in Weardale”): it is often hard to be absolutely sure about the identity of an alga in the field and, in this case, both can form conspicuous bright green growths that are slimy to the touch.   Did I miss the Ulothrix zonata bloom in the River Wear this year?   Maybe.   Looking back at my records from May 2009 I see that I recorded quite a lot of narrow Phormidium filaments then but none were apparent in this sample.   That taxon thrived throughout the summer, so perhaps, again, its absence is also a consequence of the unusual weather.

Growths of Stigeoclonium tenue on a cobble in the River Wear at Wolsingham, May 2018.  

The photograph illustrates some of the problems that ecologists face: the distribution of algae such as Ulothrix zonata and Stigeoclonium zonata is often very patchy: there is rarely a homogeneous cover and, often, these growths are most prolific on the larger, more stable stones.   I talked about this in Our Patchwork Heritage; the difference now is that the patchiness is exhibited by different groups of algae, rather than variation within a single group.   Ironically, the patchiness is easier to record with the naked eye than by our usual method of sampling attached algae using toothbrushes.   That’s partly because we tend to sample from smaller substrata (the ones that we can pick up and move!) but also because of the complications involved in getting a representative sample.   We have experimented with stratified sampling approaches – including some stones with green algae, for example, in proportion to their representation on the stream bed – but that still means that we have to make an initial survey to estimate the proportions of different types of growth.

Under the microscope, therefore, the algal community looks very different.   There are fewer green cells and more yellow-brown diatom cells, these dominated by Achnanthidium minutissimum, elegant curved cells of Hannaea arcus and some Navicula lanceolata, still hanging on from its winter peak.   The patterns I described in The mystery of the alga that wasn’t there … are still apparent although the timings are all slightly adrift.

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

The schematic view below tries to capture this spatial heterogeneity.  On the left hand side I have depicted the edge of one of the patches of Stigeoclonium.   Healthy populations of Stigeoclonium do no support large populations of epiphytes, probably as a result of the mucilage that this alga produces.  My diagram also speculates that the populations of Gomphonema olivaceum-type cells and Ulnaria ulna may be living in the shadow of these larger algal growths, as neither is well adapted to the fast current speeds on more exposed rock surfaces.  Finally, on the right of the image, there are cells of Achnanthidium minutissimum, small fast-growing cells that can cope with both fast currents and grazing.   I have not included all of the taxa I could see under the microscope, partly because of the space available.  There is no Hannaea arcus or Navicula lanceolata and I have also left out the chain of Diatoma cells that you can see on the right hand side of the view down the microscope.

The speckled background in the image of the view down the microscope is, by the way, a mass of tiny bacteria, all jigging around due to Brownian motion.  The sample had sat around in the warm boot of the car for a few hours after collection so I cannot be sure that these were quite as abundant at the time of collection as they were when I came to examine it.  However, some people have commented on the absence of bacteria – known to be very abundant in stream biofilms – from my pictures, so these serve as a salutary reminder of an extra dimension that really needs to be incorporated into my next images.

Schematic view of the biofilm from the River Wear at Wolsingham, May 2018.  a. Stigeoclonium tenue; b. Gomphonema olivaceum complex; c. Ulnaria ulna; d. Meridion circulare; e. Achnanthidium minutissimum.   Scale bar: 10 micrometres (= 1/100th of a millimetre).

* Samuel Taylor Coleridge, Christabel (1816)

 

Our patchwork heritage* …

The problem with the case I set out for a “switch” from a winter / early spring biofilm community to a summer / autumn assemblage is the sample that I was writing about contained elements of both.   This, I think, is another aspect of an issue that I touched upon in “The River Wear in January”: that the scale that we work at is much greater than the scales at which the forces which shape biofilms operate.   There is no intrinsic driver for this switch beyond the physical forces in the river but each stone will have a slightly different history.  A smaller cobble will be more likely to be rolled than a boulder, as will one that is not sheltered from the main current, or not well bedded into the substratum.  The sample I collect is a composite from the upper surface of five separate cobbles so will blend these different histories.   The more stable stone might have more Navicula lanceolata and Gomphonema olivaceum whilst the recently rolled might be dominated by early colonisers such as Achnanthidium minutissimum.

The same processes can even work on a single stone.   Arlette Cazaubon, a French diatomist, now retired, wrote several papers on this topic (see references at the end of this post).  She highlighted how the diatom assemblages differed across the surface of a boulder, depending on the exposure to the current.  However, that is only part of the story.  The picture at the top of the post was taken in January, when I was collecting my first samples of the year.  You can see the streak where I ran my finger through the biofilm and some other marks, perhaps where the heel of my wader had scuffed the stone (I’m trying to keep my balance in the middle of a northern English river in January whilst holding a waterproof camera underwater, remember).   But such damage could have arisen just as easily from twigs or stones that were being washed downstream.   Taken together with Arlette’s work, it shows how a mature Navicula lanceolata / Gomphonema olivaceum assemblage can live alongside a pioneer Achnanthidium minutissimum assemblage.

A schematic view of the biofilm in the River Wear at Wolsingham, March 2018.   a. Navicula lanceolata; b. Gomphonema olivaceum complex; c. Fragilaria gracilis; d. Achnanthidium minutissimum.   Scale bar: 10 micrometres (= 1/100th of a millimetre).

I’ve tried to depict that in the schematic diagram above.   On the left-hand side there is a mature biofilm, with long-stalked Gomphonema species creating a matrix within which motile diatoms such as Navicula lanceolata live whilst, on the right, there is a pioneer community dominated by Achnanthidium minutissimum.   However, whilst this patchiness is a natural phenomenon, it can contribute to the variability we see in ecological data and, indirectly, to an impression that ecological data are not precise.   If I were to divide the diagram above into two halves, the left-hand side would return a higher TDI than the right.  This is because the diatoms on that side have broader ecological tolerances than those on the other (the sample size, by the way, is far too small to do this seriously but I just want to make a point).   In practice, however, the entire diagram represents little more than the width of a single bristle of the toothbrush that I use to collect samples so a sample is, inevitably, an amalgam of many different microhabitats on a stone.  Our assessment of the condition of the river represents the average of all the patches across the five stones that form a typical sample on that day.

The importance of patchiness in determining the structure and composition of stream communities has been known for some time (see review by Alan Hildrew and Paul Giller in the reference list).   What we have to remember when trying to understand phytobenthos is that patchiness is, to some extent, embedded in the samples we collect, rather than being something that our present sampling strategies might reveal.

* “… for we know our patchwork heritage is a strength not a weakness ..” Barack Obama: inaugural address, 2009

Reference

A useful review on patchiness in stream ecosytems (several other papers in this volume also discuss patchiness in freshwater and marine environments):

Hildrew, A.G. & Giller, P.S. (1994).  Patchiness, species interactions and disturbance in the stream benthos.  pp. 21-62.  In: Aquatic Ecology: Scale, Pattern and Process (edited by P.S. Giller, A.G. Hildrew & D.G. Rafaelli).   Blackwell Scientific Publications, Oxford.

Some of Arlette Cazaubon’s papers on variability in diatom assemblages across the surfaces of single stones:

Rolland, T., Fayolle, S., Cazaubon, A. & Pagnetti, S. (1997). Methodological approach to distribution of epilithic and drifting algae communities in a French subalpine river: inferences on water quality assessment. Aquatic Science 59: 57-73.

Cazaubon, A. & Loudiki, M. (1986). Microrépartition des algues épilithiques sur les cailloux d’un torrent Corse, le Rizzanese. Annals de Limnologie 22: 3-16.

Cazaubon, A. (1986). Role du courant sur la microdistribution des diatomées epilithiques dans une Riviere Méditerranéenne, L’Argens (Var, Provence). pp. 93-107.   Proceedings of the 9th Diatom Symposium.   Bristol.

Cazaubon, A. (1988). The significance of a sample in a natural lotic ecosystem: microdistribution of diatoms in the karstic Argens Spring, south-east France.  pp. 513-519.   In: Proceedings of the 10th Diatom Symposium, Joensuu, Finland.

The mystery of the alga that wasn’t there…

I was back at the River Wear at Wolsingham a few days ago for my second visit of the year (see “The River Wear in January” and “The curious life of biofilms” for accounts of the first visit).   I had wanted to go out earlier in the month but we’ve had a month of terrible weather that has translated into high river flows.  Even this trip was touch and go: the river was about 30 cm higher than usual and the gravel berm that usually stretches out under the bridge on the left bank was largely submerged.

Compare the image of the substratum with the one I took in January: that one had a thick film with a chocolate-brown surface whilst the March substratum had a much thinner film lacking any differentiation into two layers.  When I put a small sample of the biofilm under my microscope, I could see that it was dominated by diatoms with only a few strands of green algae.   Many of the diatoms that I saw in January were still here in March but Navicula lanceolata, which comprised over half the algal cells I saw in January was now just 15 per cent of the total whilst Achnanthidium minutissimum was up from about 15 per cent to about 40%.    However, as A. minutissimum is a much smaller cell, N. lanceolata still formed more of the total biovolume.   One other difference that I noticed as I peered down my microscope was that there was much less amorphous organic matter in the March sample compared with the one from January.

The substratum at the River Wear, Wolsingham on 24 March 2018.   The photograph at the top shows the view from the road bridge looking downstream.

When I looked back at notes I had taken after my visit in March 2009, I saw that the riverbed then had been covered with lush growths of the green alga Ulothrix zonata (you can see a photograph of this in “BollihopeBurn in close-up”).   I did not see this on my visit last week.  That might be because the high water level means that I could not explore as much of the river as I wanted, but it was more likely a consequence of the preceding conditions.   The graph below shows at least three separate high flow events during March, the first of which associated with the melting of the snow that fell during the “Beast from the East”.   I suspect that these high flow events would have both moved the smaller substrata (the ones I usually pick up to sample!) scouring away the biofilms in the process.

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

River levels at Stanhope, 20 km upstream from Wolsingham across March 2018 showing three separate high flow events.  A screenshot from www.gaugemap.co.uk.

The final graph shows the trend in the three algae that I’ve been talking about over the course of 2009, which is similar to what I am seeing in 2018 except that that the timing of the decline in Navicula lanceolata and Ulothrix zonata along with the increase in Achnanthidium minutissimum is slightly different.   In very broad terms N. lanceolata is typical of winter / early spring conditions, favoured by thick biofilms partly created by the matrix of stalks that Gomphonema olivaceum and relatives creates.   Achnanthidium minutissimum, on the other hand, is the most abundant alga through the summer and early autumn.  It is a species that thrives in disturbed conditions, such as we would expect after the weather we’ve experienced this March.   However, we must not forget that the grazing invertebrates that thrive

during the summer months also represent a type of disturbance.  Ulothrix zonata thrives in the late winter / early spring window (see “The intricate ecology of green slime”).   I would have expected it to have persisted beyond March but, as I said earlier in the post, I may have missed some as it was difficult to get a good impression of the whole reach due to high flows.

This moveable switch between a “winter” and “summer” state creates a problem when we are sampling for ecological status assessments.   The Environment Agency has, for as long as I have worked with them, had a “spring” sampling window that starts on 1 March and runs to the end of May.  As you can see, this straddles the period when there is a considerable shift in the composition of the flora.   I’ve always suggested that they wait as long as possible within this window to collect diatom samples to increase the chance of being past the switch.  However, with a huge network to cover in a short period, along with other logistical considerations, this was always easier said than done.   I’ve worked closely with the Environment Agency to manage as much of the variation in their diatom analyses as is possible (see “Reaching a half century …”); one of the mild ironies is that simply being a huge Behemoth of an organisation can, itself, be the source of some of the variation that we are trying to manage.

Trends in approximate biovolume of three common taxa discussed in this post in the River Wear at Wolsingham during 2009.  

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.

 

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.