Biofilm blog - University of Hull
Posted: 23/05/2016 By: Samui Admin
Micro-organism play an important role in aquatic environments such as rivers, lakes and estuaries. These microbial assemblages ('biofilms') preferentially develop at water-air and water-sediment interfaces and are known to have, amongst others, a considerable influence on sediment stability and erodibility. This is mainly due to the secretion of extracellular polymeric substances (EPS) by the biofilm community. EPS ‘glues’ sediment particles together affecting sediment stability, flocculation and sediment transport processes. Despite the large impact of sediment dynamics on the development of coastal and fluvial environments, bio-stabilisation is still poorly understood due to the inherent complexity of biofilms and the large spatial and temporal (seasonality) variations involved.
Figure 1 - Image of a 3-week old biofilm grown under controlled conditions in the laboratory (Gerbersdorf and Wieprecht, 2015).
The reasons for studying biofilms within the Hydralab+ project are twofold. First, bio-stabilisation of sediments is an important effect of biofilms influencing sediment dynamics in coastal and fluvial environments. Changes in environmental conditions due to climate change may impact the sediment stabilising capacity of biofilms and hence the development of channels and bars in a range of aquatic environments. A better understanding of biofilm behaviour in natural systems is needed to predict robustly how environmental change will affect bio-stabilisation in the future. The second reason for studying biofilms is more practical and focusses on how to best represent biota in flume facilities. Traditionally, flume experiments have tried to incorporate biota by introducing small-scale plants (e.g. Braudrick et al., 2009; Tal and Paola, 2010) or artificial vegetation (Anderson and Smith, 2014) while research on biofilm stabilisation and sediment interactions has remained scarce so far (for an example see Thom et al., 2015).
Flume experiments are helpful to better understand natural biofilm behaviour and how this may change when the environmental conditions are altered. Flume experiments allow us to test how long it takes for a mature biofilm to develop and, in turn, how this affects the sediment stability. Also, do we find large differences between saline, brackish and freshwater environments in how biofilms develop and behave? Importantly, systematic tests can be done to examine the resilience and recovery of biofilms following climate-change related stresses such as droughts (simulating longer periods of low flow), floods (simulating increased storminess and flood risk) and increased salinity (simulating sea-level rise).
These experiments are important components of WP8 and WP9 of the Hydralab+ initiative in which we aim to represent climate change and complex sediment-biota interactions in flume experiments to provide informed solutions for climate-change adaptation. Below we outline the setup of our first explorative experiment aimed at growing and studying biofilm behaviour. And we have kept a blog to report on some of our observations and learnings.
The setup consists of transparent flume through which water is re-circulated by a garden pump. The flume is 13 cm wide, 52 cm long and 32 cm high. The flow rate in the flume is about 0.02 m/s with a water depth of about 0.1 m. A 2 cm thick layer of uniform sand-sized sediment is placed at the bottom of the flume for biofilms to attach to and develop on. A grow lamp creates a 16 hours light and 8 hours dark daily rhythm. In addition to visual observations, measurements involve temperature and a weekly qualitative assessment of the ATP levels, which may be a proxy for biological activity. Later on, measurement devices characterising sediment stability, EPS content and chlorophyll-a content may become available as well. A daily photo is made to capture the biofilm growth over time.
Figure 2. Experimental setup used to grow and study the growth of a natural biofilm developing in saline water. The grow lamp causes the purple lighting.
27 April 2016 Start of the experiment! We have prepared a working setup up, now it is up to the biofilms.
3 May 2016 A week into the experiment. No biofilm activity visible yet. Setup continues to work well with the water re-circulating and the lights set to a 16 to 8 hrs day-night rhythm.
4 May 2016 We have done an ATP measurement (see Okibe and Johnson, 2011) to obtain an indication of the level of biological activity in our flume. Three replicate measurements indicate an ATP level around 6000 RLU. This is quite high as for food purposes only ATP levels up to 30 RLU are allowed.
Figure 3 ATP measurement device and sticks. We are using the ATP measurements to estimate the level of biological activity in the flume (see also Okibe and Johnson, 2011).
11 May 2016 A bit over twee weeks into the experiment and still no visible biological activity in the flume. This is confirmed by another ATP measurement of only 400 RLU, which indicates that the biological activity in the flume has dropped dramatically over the last week. We don’t really understand why, but the recent warm temperatures (about 20 degrees outdoors, which resulted in an air temperature of 27 degrees in the lab) may have played a role and affected the establishment of a biological community.
18 May YES, finally some biological activity in our little re-circulating flume! Little green filaments can be seen in the upstream part of the flume. This is the area with the highest flow velocities, suggesting that higher flow velocities may be beneficial to the growth of biofilms. Also, the water appears more turbid and air bubbles have appeared at the water surface, mostly towards the down end of the flume. These are indirect indications of biological activity in the flume. These visual observations are confirmed by an ATP measurement of about 2800 RLU, which is much higher again than last week.
Figure 4 Formation of green filaments in the upstream part of the flume (left panel) and air bubbles at the water surface, mostly at the downstream end of the flume. These are the first visual indications of biological activity and biofilm formation after about 3 weeks of running the experiment.
More to come! We’ll keep you posted on any developments in the flume in the coming weeks.
Anderson and Smith, 2014, Wave attenuation by flexible, idealized salt marsh vegetation, Coastal Engineering, 83, 82-92, doi: 10.1016/j.coastaleng.2013.10.004.
Braudrick, Dietrich, Leverich and Sklar, 2009, Experimental evidence for the conditions necessary to sustain meandering in coarse-bedded rivers, PNAS, 106, 16936-16941, doi: 10.1073/pnas.0909417106
Gerbersdorf and Wieprecht, 2015, Biostabilization of cohesive sediments: revisiting the role of abiotic conditions, physiology and diversity of microbes, polymeric secretion, and biofilm architecture, Geobiology, 13, 68–97, doi:10.1111/gbi.12115
Okibe and Johnson, 2011, A rapid ATP-based method for determining active microbial populations in mineral leach liquors, Hydrometallurgy, 108, 195-198, doi: 10.1016/j.hydromet.2011.04.008
Tal, and Paola, 2010, Effects of vegetation on channel morphodynamics: results and insights from laboratory experiments, Earth Surf. Process. Landforms, 35, 1014–1028. doi: 10.1002/esp.1908
Thom, Schmidt, Gerbersdorf and Wieprecht, 2015, Seasonal biostabilization and erosion behavior of fluvial biofilms under different hydrodynamic and light conditions, International Journal of Sediment Research, 30, 273-284, doi: 10.1016/j.ijsrc.2015.03.015.