A brief history of time-wasting …*

Having talked about diversity on a microscale in the previous post, I thought it would be interesting to place this in context by looking at the variations that I have observed in the River Wear at Wolsingham over the past decade or so.   The River Wear has seen some significant improvements in water quality over this period, but those have mainly affected sections of the river downstream from Wolsingham.  Most of the changes at Wolsingham are, therefore, giving us some insights into the range of natural variation that we should expect to see in a river.

I’ve got 31 samples from the River Wear at Wolsingham on my database, collected since 2005.  Over this period, nine different diatom species have dominated my counts: Achnanthidium minutissimum on 21 occasions, Nitzschia dissipata twice and Cocconeis euglypta, Encyonema silesiacum, Gomphonema calcifugum, Navicula lanceolata, Nitzshia archibaldii, N. paleacea and Reimeria sinuata once each.   I also have records for non-diatoms during 2009, during which time the green alga Ulothrix zonata, and two Cyanobacteria, Phormidium retzii and Homeothrix varians were the dominant alga on one occasion each.   In total, I have recorded 131 species of diatom from this one reach, although only I’ve only found 91 of them more than once, and only 59 have ever formed more than one percent of the total.   I’ve also got records of 22 species other than diatoms.

This – along with my comments in “The mystery of the alga that wasn’t there …” raises questions about just how effective a single sample is at capturing the diversity of algae present at a site.  .    In 2009 I collected a sample every month from Wolsingham and the graph below shows how the total number of species recorded increased over that period.   Typically, I find between 20 and 30 species in a single sample, and each subsequent month revealed a few that I had not seen in earlier samples.   Importantly, no single sample contained more than 40 per cent of the total diversity I observed over the course of the year.  Part of this high diversity is because of the greater effort invested but there is also a seasonal element, as I’ve already discussed.   The latter, in particular, means that we need to be very careful about making comments about alpha diversity of microalgae if we only have a single sample from a site.

Increase in the number of diatom taxa recorded in successive samples from the River Wear at Wolsingham.  In 2009 samples were collected monthly between January and December whilst in 2014 samples were collected quarterly. 

This seasonal pattern in the algal community also translates into variation in the Trophic Diatom Index, the measure we use to evaluate the condition of streams and rivers.  The trend is weak, for reasons that I have discussed in earlier posts, but it is there, nonetheless.   Not every river has such a seasonal trend and, in some cases, the community dynamics results in the opposite pattern: higher values in the summer and lower values in the winter.  It is, however, something that we have to keep in mind when evaluating ecological status.

Variation in the Trophic Diatom Index in the River Wear at Wolsingham between 2005 and 2015, with samples organised by month, from January (1) to December (12).   The blue line shows a LOESS regression and the grey band is the 95% confidence limits around this line.

All of these factors translate into uncertainty when evaluating ecological status.   In the case of the River Wear at Wolsingham, this is not particularly serious as most of the samples indicate “high status” and all are to the right of the key regulatory boundary of “good status”.  However, imagine if the histogram of EQRs was slid a little to the left, so that it straddled the good and moderate boundaries, and then put yourself in the position of the people who have to decide whether or not to make a water company invest a million pounds to improve the wastewater coming from one of their sewage treatment plants.

At this point, having a long-term perspective and knowing about the ecology of individual species may allow you to explain why an apparent dip into moderate status may not be a cause for concern.  Having a general sense of the ecology of the river – particularly those aspects not measured during formal status assessments – should help too.  It is quite common for the range of diatom results from a site to encompass an entire status class or more so the interpretative skills of the biologists play an important role in decision-making.   Unfortunately, if anything the trend is in the opposite direction: fewer samples being collected per site due to financial pressures, more automation in sample and data analysis leading to ecologists spending more time peering at spreadsheets than peering at stream beds.

I’ve never been in the invidious position of having to make hard decisions about how scarce public sector resources are used.  However, it does strike me that the time that ecologists used to spend in the field and laboratory, though deemed “inefficient” by middle managers trying to find cost savings, was the time that they learned to understand the rivers for which they were responsible.  The great irony is that, in a time when politicians trumpet the virtues of evidence-led policy, there is often barely enough ecological data being collected, and not enough time spent developing interpretative skills, for sensible decisions to be made.   Gathering ecological information takes time.   But if that leads to better decisions, then that is not time wasted …

Ecological Quality Ratio (EQR: observed TDI / expected TDI) of phytobenthos (diatoms) at the River Wear, Wolsingham) between 2005 and 2015.   Blue, green, orange and red lines show the positions of high, good, moderate and poor status class boundaries respectively.

* the title is borrowed from the late Janet Smith’s BBC Radio 4 comedy series

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My name is Legion …

I promised to write a little more about Gomphonema subclavatum, one of the diatoms we encountered in the previous post.   I picked this one out for more attention because it is one of many diatoms that have changed names in recent years and it is sometimes interesting to scratch around to understand why this has happened.

Had I seen this particular species fifteen years ago I would have called it Gomphonema clavatum without hesitation.  Although G. subclavatum was recognised as a distinct species back in the nineteenth century, for most of the twentieth century it was treated as a variety of G. longiceps, which Krammer and Lange-Bertalot then subsumed into G. clavatum.  If you look at their plate of G. clavatum, you will see a huge range of sizes and shapes so it is perhaps no surprise that people subsequently realised that there was more than one species lurking under this name.

Gomphonema subclavatum from Cregduff spring, Co. Mayo, Ireland, September 2017.  Photographs: Bryan Kennedy.  Scale bar: 10 micrometres ( = 100th of a millimetre).

When this happens, taxonomists ask which of the various contenders was the Gomphonema clavatum seen by the person who originally described the species.  This involves going back to the museum collection where that person deposited the material that they examined and taking another look.  This process of “typification” helps determine which of the forms is the rightful inheritor of the name.   Erwin Reichardt decided to have a go at this process for G. clavatum and went to examine the samples, now in the Museum für Naturkunde in Berlin, on which Christian Gottfreid Ehrenberg had based his original description.  However, he could find nothing that resembled G. clavatum, with the closest match being G. olivaceum.

I’m reading a biography at the moment that contains the warning that “history is always a matter of trying to think into the minds of people who think differently from ourselves”.  That serves as a useful reminder that Ehrenberg knew far less about the biology of diatoms than we do today, but was also limited by the technology available.  Not only were his microscopes far less sophisticated than ours but also capturing the essence of the organisms he saw in print was far from straightforward (see “Picture this?”).  The idea of Gomphonema clavatum that we had until Reichardt re-examined the type material was the result of a 180-year game of “Chinese whispers”: each generation matching their specimens to inadequate images and descriptions, then making their own images which, in turn, became the basis for their successor’s identifications.  By the time Krammer and Lange-Bertalot wrote their Flora, it was finally possible to reproduce high quality micrographs, rather than line drawings but over a century of taxonomic drift meant that their images are no longer connected to the right name.  Their plate actually shows two species: the larger forms with undulate margins belong to G. longiceps Ehrenberg 1854) whilst the smaller specimens are G. subclavatum.   That assumes, of course, that there are no further twists to come.  As I alluded in my previous post, morphology might not be telling us the whole story for this genus.

The unfortunate twist, also mentioned in my previous post, is that the taxonomic confusion in the past means that we don’t actually get any sharper ecological insights in the present as a result of unravelling these names.   Anyone looking at ecological data associated with “Gomphonema clavatum” from twenty years ago needs to know that this could represent either G. longiceps or G. subclavatum or one of a number of other species that have been split away in recent years.  There is always a hope that this better understanding of taxonomy will yield fruits as we go forward but I’m always suspicious that someone else might come along and rearrange things yet again…

Reference

Krammer, K. & Lange-Bertalot, H. (1986).  Süsswasserflora von Mitteleuropa. 2/1 Bacillariophyceae 1: Naviculaceae. Spektrum Akademischer Verlag, Heidelberg.

Reichardt, E. (2015). The identity of Gomphonema clavatum Ehrenberg (Bacillariophyceae) and typification of five species of the genus Gomphonema described by C.G. Ehrenberg.  Diatom Research 30: 141-149.

The biography to which I refer is Tom Wright’s new book on Paul (SPCK, 2018).

 

Baffling biodiversity …

Few of the participants in the UK / Ireland diatom ring-test that I described in my previous post felt any need to thank me for my choice of slide for our 50th test.  The slide came from a spring in County Mayo, Ireland, which is part of the Agricultural Catchments Programme, a large study into the effect of farming on water quality. The sample itself came from the stems and leaves of the submerged water cress (Nasturtium officinale*) plants which fill the entire channel.  It was a real stinker, with a mess of Gomphonema forms, several of which did not neatly fit any species description that we could find.   A conservative reckoning is that there were at least eight different Gomphonema “species” and that raises a further question about what it was about this habitat that led to so much diversity within a single genus within a single sample.

First, a quick tour around some of the Gomphonema forms that we found.   There was general agreement that the most common type was close to G. micropus Kützing 1844 but not a perfect match to published descriptions (the stria density, in particular, was too low).   The situation was further complicated because the status of G. micropus was questioned at times, with it being treated as a variety of G. parvulum and placed in the G. angustatum complex by different authorities during the 20th century.  Then there were a number of valves with more rounded ends and a higher striae density than G. micropus but which, if you look closely, are not symmetrical around the long axis.   We thought that these were close to G. cymbelliclinum Reichardt & Lange-Bertalot 1999.   Unfortunately, there were also quite a lot of valves that had intermediate properties, making it hard, in many cases, to say whether it was one species or the other.

Gomphonema cf micropus from Cregduff spring, Co. Mayo, Ireland, September 2017.  Photographs: Bryan Kennedy.  Scale bar: 10 micrometres ( = 100th of a millimetre).  The image at the top of the post shows Cregduff spring (photo by Lauren Williams)

Gomphonema cf cymbelliclinum from Cregduff spring, Co. Mayo, Ireland, September 2017.  Photographs: Bryan Kennedy.  Scale bar: 10 micrometres ( = 100th of a millimetre).

We also found some valves that were close to descriptions of Gomphonema utae Lange-Bertalot & Reichardt 1999 and some that were close to G. parallelistriatum Lange-Bertalot & Reichardt 1991.  We also found representatives of the G. parvulum complex, G. tergestinum and G. subclavatum (more about this one in the next post).

Gomphonema cf utae from Cregduff spring, Co. Mayo, Ireland, September 2017.  Photographs: Bryan Kennedy.  Scale bar: 10 micrometres ( = 100th of a millimetre).

Gomphonema cf parallelistriatum from Cregduff spring, Co. Mayo, Ireland, September 2017.  Photographs: Bryan Kennedy.  Scale bar: 10 micrometres ( = 100th of a millimetre).

So what is going on here?   There are, I suspect, two key elements to the story that we need to explain.  The first is the limits of species within Gomphonema.  I’ve touched on this before (see “Diatoms and dinosaurs”) and some recent studies that combine morphological and molecular biological evidence also cast doubt on our ability to differentiate within this genus using classical approaches.   Whilst I was struggling to disentangle the species in this sample, I had a conversation with an eminent taxonomist and she hinted darkly that Gomphonema was “over-described”.  There is a readiness to “split” established taxa and describe new species that, in her opinion, runs ahead of the evidence.

The limitations of taxonomy cannot explain all of the variation that we observed in this sample, so the second question to ask is what it is about the conditions here that allow so many representatives of one genus to thrive.   I’ve touched on this subject before (see “Baffled by the benthos (1)” and “Baffled by the benthos(2)”).  In these posts I introduced G. Evelyn Hutchinson’s “paradox of the plankton” in which he suggested that environments that look uniform, to mortals six orders of magnitude larger than algae are, in fact, considerably more heterogeneous  and, so offer more opportunities for “variations on a theme” to thrive.   In the second post I went on to suggest that this type of diversity imparts resilience to an ecosystem and so should be looked upon as a positive feature of the ecosystem when doing ecological status assessments.

There is, however, one final possibility that, to my knowledge, has not yet been explored.  The presence of transitional forms in the diatom assemblage at Cregduff may be an artefact of our inability to differentiate biological species based on a limited range of morphological criteria on offer. However, it is also possible that we are looking at a situation where the Linnaean species are not reproductively isolated from one another, allowing hybridisation.   The concept of a “hybrid swarm” is well known in some other groups (e.g. orchids) but has never been formally demonstrated in diatoms.  However, the wide morphological diversity within a single genus in one sample alongo with, in some cases, apparent continua of variation, does raise questions about speciation within thi genus.

The final twist to this story is that, from the point of view of current ecological status assessments, all this diversity has little effect.  Though everyone grumbled about the difficulties in naming the Gomphonema species, the results, as the box-and-whisker plot in the previous post show – were less variable than in many of our other ring tests.  What I suspect happened is that the underlying taxonomic confusion means that many of these taxa have “mid-range” scores for the TDI (and other indices), so the final calculation cancels out the identification issues.  Bear in mind that this may not always be the case!

* I understand that this is the correct name now, rather than Rorippa nasturtium-aquaticum.  See Al-Shehbaz, A. & Price, R.A. (1998).  Delimitation of the genus Nasturtium (Brassicaceae).  Novon 8: 124-126.

References

The two papers that deal with variation within Gomphonema to which I refer are:

Abarca, N., Jahn, R., Zimmermann, J. & Enke, N. (2014).  Does the cosmopolitan diatom Gomphonema parvulum (Kützing) Kützing have a biogeography? PLOS One 9: 1-18.

Kermarrec, L., Bouchez, A., Rimet, F. & Humbert, J.-F. (2013).  First evidence of the existence of semi-cryptic species and of a phylogeographic structure in the Gomphonema parvulum (Kützing) Kützing complex (Bacillariophyta).  Protist 164: 686-705.

Reaching a half century …

Last week saw a small career achievement as I sent out the result of the 50th diatom ring-test that I organise for the diatom analysts in the UK and Ireland.  “Ring-test” is the informal term for an inter-laboratory comparison, when two or more laboratories analyse the same sample and compare their results.  We started out doing regular ring-tests in 2007 for all the people who were analysing diatom samples for assessments associated with the Water Framework Directive, sending out five slides each year to staff in the UK and Irish environment agencies and contractors who worked with them. Now, a decade later, the scheme is still going strong, with participants from Germany, Sweden and Estonia joining the British and Irish contingents.

There are a number of similar schemes around Europe with the same basic model: the organiser sends out copies of a slide made from the same sample, all participants then analyse the slide and send in their results, which the organiser collates.   There is usually one or more designated “expert” against whose results everyone else is judged.  Most of the other schemes then organise a workshop at which participants gather to discuss the finer points of diatom taxonomy.   We have had workshops in the past, but these are not directly linked to the ring-tests.  Instead, we send out a report that summarises results and provides notes on the identification of difficult or unusual taxa.   The money we save on workshops means that we can circulate more slides.  I’m a great believer in “little and often” for this type of quality control.

A second feature of our scheme (which some of the other European schemes have also now adopted) is to use a panel of experienced analysts to provide the benchmark that other participants should achieve.   This means that we have an idea of both the average result and the scale of the variation associated with this.  We learned early on that some samples gave much less variable results than others, even when the analyses were performed by experienced analysts.  We use this knowledge to adjust the size of the “target” that participants must achieve.   The graphs below show the results for our most recent test.  The horizontal blue lines on the left hand graph show two standard deviations around the mean of the “expert” analyses (expressed as TDI).  This is the “warning limit”; if an analyst exceeds this then he or she should be looking at their results to see if they have made any mistakes.  The red line is the “action limit”, seven TDI units either side of the expert mean.   We know from other studies (see lower graph, left) that it is very unlikely that two replicate analyses have a greater difference than this, so analysts who exceed this should definitely be checking their results.

The results of the 50th UK / Ireland diatom ring test showing (left) difference in TDI and (right) number of taxa (N taxa) between experts and other participants.  Blue lines: mean TDI ± two standard deviations of expert panel’s mean; red lines: mean TDI ± 7.   Note that it is unusual for the between-analyst variability to be quite as narrow as it was for this slide.

The reason why we need flexible “warning limits” is illustrated in the right hand graph below.   This shows the similarity between two counts as a function of the diversity of the samples.   The relationship has a wedge-shape (illustrated by the blue line – the regression line through the 90th percentile of the data).   There are a number of reasons why two analysts are unlikely to get identical results, one of which is that they disagree on the identities of the taxa that they encounter (the reason why we are doing the audits in the first place).  But what a wedge-shaped relationship is also telling us is that there seems to be an upper limit to the similarity that can be achieved at any given diversity.   This is an inherent stochastic quality of the data and has nothing to do with the competence of the analysts.

Left: some of the data from which the “action limit” for the ring-tests was established.   These are the results of audits of 67 samples from Northern Ireland in which the original (“primary”) analysis was checked against the result of an independent (“audit”) analysis.   Right: The effect of diversity on the similarity between primary and audit analyses for the same dataset.

A further way in which our scheme differs from others is that no-one “passes” or “fails”.  That might seem counter-intuitive as this is supposed to be a test of competency.   A regular reader of this blog, however, should understand that there absolute truth is often elusive when it comes to identifying diatoms and other algae.  The hard objectivity needed for a real test of competency always has to be moderated by the recognition of the limitations of our craft.   Moreover, turning this exercise into a calibration exercise runs the risk of turning the analysts into machines.  Rather, we use the term “reflective learning”, encouraging participants to use the reports to judge their own performance relative to the experts, and to take their own corrective action.

Some of the organisations whose analysts participate use the ring-test as part of their own quality control systems, and will take corrective action if results stray across the action limit.  That seems to be a sensible compromise: quality control should be the responsibility of individual laboratories, rather than delegated out to third parties.   At the same time, organisations need to understand that the people who perform ecological analyses are professionals, not treated as if they are one more machine in a laboratory that needs to be calibrated.

 

If you are interested in joining the UK / Diatom ring test scheme, or just want to learn a little more about it, get in touch with me and I’ll do my best to answer your questions.

Reference

Kelly, M.G. (2013).  Building capacity for ecological assessment using diatoms in UK rivers.  Journal of Ecology and the Environment 36: 89-94.

 

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.

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.