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


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.  

Burnhope Burn’s beautiful biofilms …

I have continued the series of studies that I started in “In search of the source of the Wear” with a three-dimensional diorama of the biofilm that I found at the mouth of Burnhope Burn, and can now compare it with the corresponding study from Wolsingham (see “The curious life of biofilms”).   The two big differences are the greater number of green filaments at Burnhope and the large numbers of cells of Navicula lanceolata at Wolsingham.   I suspect the two are linked: the Wolsingham biofilm was a mix of diatoms and organic particulate matter along with associated bacteria whilst the Burnhope biofilm was green algae and organic matter with diatoms in a subordinate role.  I speculated, in my earlier post, that Burnhope Burn’s location below a reservoir may have altered the hydrology of the stream such that green algae were favoured.   I wonder, too, if the presence of green algae then subtly shifts the composition of the biofilm matrix such that dense aggregations of Navicula lanceolata are not able to develop in the way that they could at Wolsingham.

There is something about the ecology of a few Navicula species that leads to the development of these aggregations (see “The ecology of cold days” for more about freshwaters, whilst “An excuse for a crab sandwich, really” and “A typical Geordie alga …” describes similar phenomena in brackish habitats).   Conversely, Nitzschia dissipata, which was the most abundant diatom at Burnhope Burn, never seems to form these dense monocultures.   Nitzschia dissipata was also much less common in the biofilm from Killhope Burn, just a few metres away from where I collected the Burnhope sample and where filamentous green algae are scarce.  wonder if this, too, is more than a coincidence and that N. dissipata is actually adapted to living within matrices formed by filamentous algae rather than on top of matrices dominated by diatoms and organic particulates?

I have seen a few other motile diatoms – Denticula tenuis is one – that seem to be more abundant in the presence of filamentous algae.   There may also be species that thrive when the matrix is composed largely of inorganic particles, as well as other species (Navicula angusta and N. notha are two that spring to mind) that may be naturally “understory” species that are never especially abundant in biofilms.   All this is pure speculation, but it is worth remembering that most of the insights into diatom ecology come from studies on cleaned valves which removes all traces of non-diatom algae, and also that the prevailing dogma of diatom sensitivity to their chemical environment is such that non-chemical factors are largely overlooked in academic studies.   No evidence, in this case, may just mean that no-one has asked the right questions.

In search of the source of the Wear …

Having investigated the microscopic world at Wolsingham (see “The River Wear in January” and “The curious life of biofilms), I decided that it would be interesting to head further upstream and see how much difference there was between the algae at the two locations.  I drove up to Wearhead on a cold Saturday morning to take a look but was immediately faced with a conundrum: the River Wear is formed from the confluence of two very different streams, both with extensive catchments on the moors of the northern Pennines.   One of these is Burnhope Burn, which is fed by Burnhope Reservoir, about a kilometre above Wearhead, and the other is Killhope Burn, which drains a large area of blanket bog, forestry and, importantly, abandoned metal mines.   Burnhope Burn is on the left of the photograph above whilst Killhope Burn comes in from the right.  I thought it might be rather interesting to take a sample from each and see how they compared.

The two streams look quite different to one another.   Burnhope Burn, its flow regulated by the reservoir, is the Cain of the pair whilst Killhope Burn is the unruly turbulent Abel.   This was apparent, too, when I was collecting the samples and, again, when I peered at them through my microscope.   Burnhope Burn’s biofilm was thicker and the most conspicuous algae that I could see were green filaments of Klebsormidium.  Killhope Burn’s was thinner and dominated by diatoms.   Many of the same diatoms were found in the two samples, but Burnhope Burn had more of the motile Nitzschia species that benefit from the tangled matrix of green algal filaments that thrived there.

Views of the biofilm from Burnhope Burn (a.) and Killhope Burn (b.) just above their confluence to form the River Wear, February 2018.

I’ve tried to capture the essence of the biofilm from Burnhope Burn in the schematic diagram below.  Compare this with the diagram of the biofilm from the Wear that I showed in my earlier post.   In both cases, we have a mix of organic and inorganic elements, with the organic matter further divided into living organisms and agglomerations of particulate matter.  A few of the species are common to both but there are also some notable differences.   The biofilm in the Wear, for example, had almost no green algae (though that may change over the coming months) whilst that from Burnhope Burn has many filaments of Klebsormidium.   There were motile diatoms at both locations but the species are different: Navicula lanceolata and N. gregaria at Wolsingham and Nitzschia dissipata at Burnhope Burn.  People usually describe differences in the ecology of diatoms in terms of their chemical environment but I sometimes wonder if, in the case of motile diatoms, the nature of the matrix within which they live also plays a role in determining which thrive.

The difference between Burnhope and Killhope Burns is a variation of the theme that I discussed in “Small details in the big picture …”.  Again, regulation of a river or stream plays a role in determining which species of algae can thrive.  However, whereas I found a lot of Platessa oblongella in the unregulated streams of the Ennerdale catchment, the more base-rich environment of the Pennines means that I am much less likely to find P. oblongella in these streams.  In fact, I don’t think I have ever seen it in north-east England (see distribution maps in “Why do you look for the living amongst the dead”).

That reminds me: I was going to write more about the ecology of Platessa oblongella before I was diverted by desmids and Wearhead.   Soon …

A schematic view of the vertical structure of a submerged biofilm from Burnhope Burn, Wearhead, February 2018.   a. Klebsormidium fluitans; b.  Phormidium; c. Nitzschia dissipata (valve view); d. N. dissipata (girdle view); e. Gomphonema cf. calcifugum (valve and girdle views); f. inorganic particles; g. fine particulate organic matter.  Scale bar: 20 micrometres (= 1/40th of a millimetre).


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.


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).


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.