THE RESILIENCE OF BIOFILM ON AN INTERTIDAL MUDFLAT
By CATHERINE BLAIS
B.Sc. Université du Québec à Trois-Rivières, 2009
A thesis submitted in partial fulfillment of the requirements for the degree of
MASTER OF SCIENCE in ENVIRONMENT AND MANAGEMENT
We accept this thesis as conforming to the required standard
...... Dr. Mary Lou Lauria, Thesis Supervisor WorleyParsons Canada
...... Dr. Matt Dodd, Thesis Coordinator School of Environment and Sustainability
...... Dr. Chris Ling, Director School of Environment and Sustainability
ROYAL ROADS UNIVERSITY September 2014
© Catherine Blais, 2014 THE RESILIENCE OF BIOFILM 2
Abstract
The key objective of this thesis was to determine the resilience of biofilm on an intertidal mudflat following a physical disturbance. Over the last decade the presence and importance of biofilm on intertidal mudflats has been a focus of scientific research as biofilm has been shown to play key physical and ecological roles in mudflat dynamics. An experiment took place in the summer of 2013 to assess the resilience of biofilm following a physical disturbance. Using chlorophyll a, total organic carbon, total carbohydrate as well as the taxonomic community as variables, the re-establishment of biofilm on an intertidal mudflat was monitored over a 45 day period. Biofilm was found to regenerate rapidly (between 5 to 9 days), and was linked to environmental variables such as the tide and taxonomic species present. The regeneration varied over time and baseline levels were reached after 28 days.
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Table of Contents
Title Page………………………………………………………………………………………….1 Abstract ...... 2 Table of Contents ...... 3 Acknowledgements ...... 7 Introduction ...... 8 Objectives ...... 10 Hypothesis ...... 12 Literature Review...... 12 Estuaries...... 12 Physical properties of estuaries...... 13 Threats to Intertidal Mudflats ...... 17 Biofilm ...... 17 Parameters influencing biofilm productivity...... 20 Biofilm succession and development...... 21 Ecological Importance of Biofilm ...... 22 Methodology ...... 24 Sampling Location ...... 24 Disturbed Sites ...... 28 Control Sites ...... 30 Monitoring ...... 31 Sampling Technique ...... 33 Use of marked rope...... 37 Sample collection and storage...... 38 Field duplicates...... 38 Chlorophyll a, Total Organic Carbon and Total Carbohydrate ...... 39 Chlorophyll a by fluorimetry...... 39 Total Organic Carbon (TOC)...... 40
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Total Carbohydrate...... 40 Quality Assurance and Quality Control (QA/QC)...... 41 Data analysis...... 41 Statistical analyses...... 42 Taxonomic Identification ...... 43 Identification and enumeration of microphytobenthos...... 43 Quality Assurance and Quality Control...... 46 Data analyses...... 47 Results ...... 50 Chlorophyll a, Total Organic Carbon and Total Carbohydrate ...... 50 Statistical results...... 53 Microphytobenthos Community Structure ...... 59 Microphytobenthos Community Diversity ...... 62 Relationship to the tidal cycle ...... 64 Discussion ...... 64 Chlorophyll a, Total Organic Carbon and Total Carbohydrate ...... 64 Microphytobenthos Community Structure ...... 66 Microphytobenthos Community Diversity ...... 67 Microphytobenthos diversity...... 68 Natural Variation ...... 69 Resilience of Biofilm to a Major Event ...... 72 Sea level rise...... 72 Climate change...... 73 Conclusion and Recommendations ...... 74 Works Cited ...... 76
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Figures
Figure 1. Roberts Bank Location ...... 11 Figure 2. Schematic representation of a tide cycle over a 24 hour period, based on data from Fisheries and Oceans Canada, for the Tsawwassen location (station ID: 7590) (Government of Canada, 2013)...... 15 Figure 3. Schematic representation of a tidal cycle over a 45 day period, based on data from Tsawwassen (station ID: 7590) and the lunar cycle...... 16 Figure 4. Schematic of Microbial Biofilm within Intertidal Sediments (updated from Decho 2000)...... 19 Figure 5. Sampling locations at Roberts Bank, outside of the Roberts Bank Wildlife Management Area...... 25 Figure 6. Schematic representation of the paired disturbed and control sampling plots. 26 Figure 7. Photographic representation of the steps involved in creating the disturbance plots...... 29 Figure 8. Control site marked with Surveyor Flags...... 30 Figure 9. Sampling technique showing core collection steps using a modified 2.75 cm diameter syringe tube...... 34 Figure 10. Sampling technique showing core collection steps using a scraper...... 35 Figure 11. Sampling style. (A) Horizontal. (B) Vertical. (C) Two samples (chemical and taxonomy) and one duplicate...... 36 Figure 12: Sediment sample collection using both the syringe core technique and scraper technique, with the marker rope on the ground...... 37
Figure 13. Relationship between Log10 (chlorophyll a), Log10 (total organic carbon), Log 10(total carbohydrate), and Day...... 52 Figure 14. Five microphytobenthic species found at Roberts Bank with the highest abundance, at each sampling site...... 61 Figure 15. Microphytobenthos Abundance, Shannon-Weiner Index (H’), Taxonomic Evenness (E) and Richness(S) for the duration of the experiment, for disturbed and controlled sampling sites...... 63
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Tables
Table 1 Sampling Area Locations ...... 27 Table 2 Tides and Dates during Sampling Days...... 32 Table 3 Classification of Plankton (modified from Wotton 1990) ...... 45 Table 4 Mean density prior and immediately following disturbance (mg/m2) ...... 51
Table 5 Results of ANOVA two factor Analysis with Log10 (chlorophyll a, TOC or total carbohydrate) as Dependent Factors and Treatment and Day as Independent Factors...... 54
Table 6 Results of Two-Sample t-test Analysis with Log10 (chlorophyll a, TOC and total carbohydrate) for each sampling day, grouped by treatment (Disturbed and Controlled)...... 56
Table 7 Results of Tukey Pairwise Comparison test for Log10 (chlorophyll a, TOC and total carbohydrate), controlled and disturbed sampling sites tested together, grouped per day...... 58 Table 8 Main taxonomic species found at Roberts Bank for all sampling sites. ...60
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Acknowledgements
I would like to thank my supervisor Dr. Mary Lou Lauria for all the support and advice throughout the duration of my master’s project, and her commitment to helping me create a quality thesis. I know her schedule was really hectic at the time and I appreciate her squeezing me in.
Obviously, I would not have been through the challenges of field sampling, data and statistical analysis without the help of Christopher Martin. I learned a lot from this experience and I hope to be able to bring some of it in my future works.
Thanks Karina Andrus for putting up with a stressed out, tired student, and for giving me the time needed to complete all the courses and write the thesis. I also would like to thanks my colleagues at WorleyParsons, who had to listen to my rambling about thesis and biofilm for months, and who were there to help me and support me in making it happen.
Merci à ma famille qui, malgré la distance, ma supporté tout au long de cette aventure.
Vous avez joué un grand rôle dans la réussite de ce projet.
Finally, thanks to all my friends who encouraged me and reminded me to go out once in a while.
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Introduction
Estuaries are characterized by a gradient of salinity ranging from freshwater to seawater, and are dominated by fine sedimentary material, which accumulate to form mudflats (McLusky
& Elliott, 2004). Estuaries are typically sheltered habitats where waves are dissipated by reefs, barrier islands, mud or sand bars, creating a habitat for thousands of species of birds, mammals, fish, and other wildlife that depend on the estuary to live, feed, and reproduce. Estuaries are biologically productive and provide areas for migratory birds to rest and re-fuel on migratory journeys, as well as spawning and nursing habitat for many fish species (United States
Environmental Protection Agency (US EPA), 2012).
At the boundary between the land and sea is a shallower section of the continental shelf: the intertidal zone. Kennish (1986) described the estuarine intertidal area as a broad, gently sloping region of muddy or sandy sediment covering up to hundreds of square kilometers along the coast. In contrast to subtidal habitats, which tend to be more physically predictable, intertidal habitats are influenced by a greater variability in physical factors (e.g., waves, tidal currents, erosion, slope, light, air exposure, temperature, salinity, and sediment stability) (Burd, Barnes,
Wright, & Thomson, 2008).
Coastal regions are predicted to experience dramatic change in future decades as a consequence of global climate change and local pressures from fishing, transport, habitat destruction and waste disposal (Johannessen & Macdonald, 2009). Historically, human populations have gathered around estuaries based on the characteristics of naturally navigable harbours and waterways that allow transportation of ships inland for commercial and industrial practices (Ross, 1998).
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Over the last 10 years, research has focused on specific components of the estuarine environment including biofilm, an important primary producer on intertidal mudflats (Beninger,
Elner, Moranais, & Decottignies, 2011; Chiu, Thiyagarajan, Tsoi, & Qian, 2006; Coulon et al.,
2012; Herlory, Guarini, Richard, & Blanchard, 2004; Kuwae et al., 2008). Biofilm is a three- dimensional matrix of organic and inorganic substances that grow on intertidal mudflats.
Biofilm colonization occurs through the settlement of phytoplankton, the proliferation of that microphytobenthos on and within the surface layers of the sediment and re-suspended microphytobenthos present in coastal systems. Similar to phytoplankton production in the water column, the establishment, growth and reproduction of biofilm communities are dependent on several physical, chemical and biological factors. Biofilm has been recognized as an important component of estuaries, because of its role in the mudflat food web and the stabilization it creates in the sediment.
To further understand the factors limiting biofilm growth, and the time necessary for biofilm to re-generate following a disturbance, an experiment was carried out in the summer of
2013 at Roberts Bank, in British Columbia (Figure 1). The Fraser River Estuary (FRE), comprised of Roberts Bank, Sturgeon Bank and Boundary Bay, is situated on the mainland coastline of the Strait of Georgia, South of Vancouver, BC (Figure 1). The FRE provides a suitable physical habitat for microphytobenthos establishment and large quantities of biofilm are known to exist (Catherine Berris Associates Inc., 2010; WorleyParsons, 2014).
Roberts Bank covers an area of 60 km2 and presents a range of habitats for foraging, resting, or roosting activities for staging waterfowl, shorebirds and birds. The species associated with Roberts Bank are supported by a diversity of habitats ranging from marshes to intertidal
THE RESILIENCE OF BIOFILM 10 mud and sand flats, eelgrass meadows, and biofilm. The significance of Roberts Bank for several million shorebirds and waterfowl as a stop-over on their annual migration resulted in a large portion of the area designated under section 4(2) of the Wildlife Act by The Ministry of
Forests, Lands and Natural Resources Operations, on September 7, 2011 (OIC 410/11, BC Reg.
155/2011), as a Wildlife Management Area (WMA) (Province of British Columbia, 2013).
Objectives
The purpose of this study is to examine the resilience of biofilm on an intertidal mudflat following a disturbance, such as sea level rise or a major storm event. The main objectives are:
. to measure biofilm regeneration following a disturbance ; . to assess the succession of the biofilm community following the disturbance by identifying genera; . to identify patterns of regeneration based on the local physical attributes at Roberts Bank; and . to evaluate biofilm recovery in the event of a major event.
To address these objectives, disturbance areas were created on the Roberts Bank’s mudflat and the regeneration progress was monitored through the quantification of chlorophyll a, total carbohydrates, total organic carbon (TOC) and taxonomic analysis.
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Figure 1. Roberts Bank Location
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Hypothesis
To date, research on biofilm re-establishment has been limited and many of the studies have been conducted under laboratory conditions. For example, Tolhurst, Consalvey, &
Paterson (2008) and Lundkvist, Gangelhof, Lunding, & Flindt (2007) determined that maximum biofilm biomass generally occurred approximately 10 to 13 days after introduction of sterilized sediments. The Tolhurst et al. (2008) study examined the micro-scale changes in the properties of sediment associated with the growth and development of biofilm over a period of seven weeks. The research team found that biofilm development was rapid, with changes in the sediment properties occurring after one day and a visible film forming after just three days.
There was an increase in microphytobenthos with time, and diatoms rapidly colonized the sediment and formed a biofilm (Tolhurst et al., 2008).
The hypothesis of this research study is that the resilience of biofilm will take the same length of time, or longer, for biofilm to re-establish in a natural setting than it did under the published laboratory experiment. The statistical hypotheses of this study were as follow:
. Null hypothesis (H0): the disturbed plots will not show a difference from controlled plots over a 45 day period following physical disturbance.
. Alternate hypothesis (H1): the disturbed plots will show a difference from controlled plots over a 45 day period following physical disturbance.
Literature Review
Estuaries
Estuaries are among the most productive environments on earth, creating more organic matter each year than comparably-sized areas of forest, grassland, or agricultural land (McLusky
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& Elliott, 2004). The high primary production found in estuaries is partly due to nutrients brought in by the tide and rivers, nitrogen-fixing organisms, and the decomposition of detritus,
(Castro & Huber, 2007), as well as the exchange of nutrients with deeper ocean water masses facilitated by the upwelling process; which is the entrainment of water from deeper layers to the surface. Along with tidal mixing, it constitutes the other major process for nutrient regeneration in the surface layer (Tomczak, 1996).
Physical properties of estuaries.
Estuarine waters are extremely variable in their salinities. While seawater is between 33 and 35 Practical Salinity Units (PSU), the salinity of freshwater is always less than 0.5 PSU
(McLusky & Elliott, 2004). The brackish water typically found in estuaries has salinities that range between 0.5 and 35 PSU. These fluctuations of salt and freshwater present many challenges to the bio-physiology of the living organisms in this habitat, and only a few euryhaline species (species tolerant of a wide range of salinities) are able to adapt (McLusky &
Elliott, 2004). This challenging habitat means that diversity in estuaries is generally low (Ross,
1998).
Water exchange and mixing contributes to a number of functions of estuaries: transportation of nutrients, distribution of fish larvae and invertebrates, flushing of waste, controlling salinity and the movement of sediments (Knox, 1986). The FRE has an open connection with the Strait of Georgia, which allows seawater to enter the estuary following a daily rhythm of the tides. Tides in the FRE are semi-diurnal (two high waters and two low waters each day) and are comprised of eight stages, presented in Figure 2.
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During a flood tide, when the saline water is drawn from the Strait of Georgia into the estuary, the Fraser River flow is slowed or reversed as a wedge of salt water is pushed up the river channel (Bendell-Young et al., 2004). During the ebb tide, when the tide is going out, the salt water retreats, releasing the freshwater into the estuary (Bendell-Young et al., 2004). At the interface between the salt and freshwater, entrainment (mixing) occurs and saltwater is mixed into the outflowing freshwater (McLusky & Elliott, 2004).
The freshwater flow coming from the Fraser River floats on top of the saline water coming from the open sea, and gradually mixes vertically from the bottom to the top. This gradual vertical mixing leads to an outgoing stream of fresher surface water (McLusky & Elliott, 2004).
In addition wind can enhance surface currents and upwelling, resulting in increased nutrient entrainment (Dransfeld, 2000; K. Yin, 1994).
Productivity and exposure time to light is also dependent of the spring and neap tides on a fortnightly cycle. Spring tides occur when the moon is full or new (Figure 3). At these times, the high tides are very high and the low tides are very low. Neap tides happen when the moon is at first or third quarter and the tidal range is at its minimum (Figure 3). There is about a seven- day interval between springs and neaps. The full cycle from spring to neap tides occurs each lunar month (28 days).
At Roberts Bank, in the FRE, the nutrients brought daily by the tidal exchange of marine and freshwater, as well as the day long solar radiation in spring and summer on the wide and shallow mudflat create a suitable environment for biofilm growth.
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(2) (4) (3) (6) (8) (7) (5) (1)
Figure 2. Schematic representation of a tide cycle over a 24 hour period, based on data from Fisheries and Oceans Canada, for the Tsawwassen location (station ID: 7590) (Government of Canada, 2013).
(1) Slack High Water-1: The water level has reached its highest level within the lunar day. (2) Ebb Tide-1: The water falls over several hours, exposing the intertidal zone. This portion of the tide cycle has the greatest tidally-driven water velocities. (3) Slack Low Water-1: The water stops falling, reaching the daily low tide, a period of typically two (2) hours. (4) Flood Tide-1: The water rises for several hours, covering the intertidal zone. (5) Slack High Water-2: The water rises to its second highest level, lower than the first Slack High Water. (6) Ebb Tide-2: The water falls for a few hours, but with a lower range compared to the first ebb tide, resulting in lower water velocities. (7) Flood Tide-2: The water rises for a few hours, with a lower range compared to the first flood tide. (8) Slack Low Water-2: The water stops falling, reaching the second daily low tide, for a period usually longer than the first Slack Low Water.
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Figure 3. Schematic representation of a tidal cycle over a 45 day period, based on data from Tsawwassen (station ID: 7590) and the lunar cycle. Spring tides are represented by the gray boxes, while the white boxes represent the neap tide.
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Threats to Intertidal Mudflats
Tidal flats face a number of natural and anthropogenic threats including predicted sea level rise, natural disaster, erosion, invasive species, loss of habitat, coastal development and pollution. Storms, wind-induced waves, hurricanes, prop scarring, etc., can contribute to the erosion of tidal flat, as well as tidal current velocities.
Recent estimates put predicted levels of sea rise at 50 cm in the next one hundred years
(Bornhold, 2008; Lemmen, Warren, Lacroix, & Bush, 2008; Intergovernmental Panel on Climate
Change (IPCC), 2013). Past changes in sea level have greatly affected estuarine coastlines and could have rapid and significant present-day effects (Holligan & Reiners, 1992; Lemmen et al.,
2008). Rising sea levels could render intertidal flats into subtidal habitat (Little, 2000); a zone that occurs below low water and is rarely exposed to the atmosphere. Erosion and higher sea level rise could lead to decreases in biofilm availability as a food source. Some of the predictions related to climate change are increases in flood risk, threats to coastal defenses (such as dykes), as well as the migration of the intertidal area inshore (Fujii, 2012).
Biofilm
In intertidal systems, microbial processes play critical roles in the remineralisation of nutrients and primary production (Decho, 2000). These processes are localized within a matrix called biofilm. Biofilm is composed of a mixture of the following elements:
. Microphytobenthos; . A mucilaginous matrix of Extracellular Polymeric Substances (EPS); . Microbes; . Organic detritus; and
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. Sediment.
Microphytobenthos are unicellular, eukaryotic algae and cyanobacteria that grow within the upper several millimeters of illuminated sediments, typically appearing as a subtle brownish or greenish shading (MacIntyre, Geider, & Miller, 1996). The microphytobenthic communities, which are typically dominated by diatoms, drive the production of biofilm and related carbohydrates (Admiraal, 1984; Underwood & Kromkamp, 1999).
Comprised primarily of carbohydrates (polysaccharides), the EPS matrix provides a protective microenvironment from the rapidly changing physical and chemical conditions experienced at intertidal mudflats (Decho, 2000) as well as a method of attachment to sediment particles (Wang, 2003). The physical state of the EPS ranges from a gelatinous continuum to a dissolved solution (Decho, 2000) with semi-solid forms observed as a light green or brownish layer. Figure 4 represents an updated schematic of microbial biofilm, based on Decho (2000).
As microphytobenthos are photosynthetic, they are constrained by the depth of maximum light penetration; usually the top 2 mm of sediments (De Brouwer & Stal, 2001; Herlory et al.,
2004). These organisms are known to exhibit vertical migrations within sediments spurred by changing conditions in the physical environment, such as light levels and water immersion/emersion (Guarini, Blanchard, Gros, & Harrison, 1997; Smith & Underwood, 1998).
This vertical migration is synchronized with daily emersion periods (Admiraal, 1984; Consalvey et al., 2004), reducing the risk of microphytobenthos being dislodged by the ebbing tide as well as reducing predation from grazers (Herlory et al., 2004). These migrations are facilitated by the secretion of EPS (Decho, 2000).
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Figure 4. Schematic of Microbial Biofilm within Intertidal Sediments (updated from Decho 2000).
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Parameters influencing biofilm productivity.
Biofilm establishment, growth, and productivity are limited by several physical and chemical parameters such as tide, light, salinity, temperature, and nutrient availability. These parameters fluctuate on spatial and temporal scales, resulting in fluctuations to the total community primary productivity (Dransfeld, 2000), that can be defined as the amount and rate of production which occur in a given ecosystem over a given time. Differences in productivity are common, particularly on daily and seasonal scales (Dransfeld, 2000; Ross, 1998). Other seasonal factors associated with spring freshet will also impact phytoplankton and microphytobenthos growth. For example, the silt-laden discharge of the Fraser River increases the turbidity levels in the FRE, which is known to result in reduced concentrations of chlorophyll a near the river mouth (Parsons, Stronach, Borstad, Louttit, & Perry, 1981).
Biofilm located in intertidal habitats is influenced by tidal cycles. Mudflats experience dramatic changes in the physical and chemical environment over a tidal cycle. These changes induce vertical migration behaviour of microphytobenthos in order to optimize productivity (i.e., light levels) while minimizing removal (i.e., ebb flows). These daily movements of microphytobenthos within the biofilm can influence the spatial and temporal distribution of biofilm biomass and productivity (Consalvey et al., 2004; Easley, Hymel, & Plante, 2005).
A limited number of studies looked at both sediment type and tidal height as factors explaining microphytobenthos variability on intertidal mudflat, and tidal height was found to be the second most significant variable influencing microphytobenthos variability, after sediment type (Brotas et al, 1995; Jesus et al., 2009). Tidal height was also found to be more important in the distribution of the total amount of microphytobenthic biomass due to desiccation effects,
THE RESILIENCE OF BIOFILM 21 available irradiation, temperature, re-suspension of algae and sediment particles, grazing, and emersion/immersion rhythm of vertical migrations, while sediment type was more important in taxonomic composition and photo-regulation strategies (Brotas et al., 1995; Jesus et al., 2009).
Biofilm succession and development.
Bare substrates are generally colonized rapidly by bacteria, followed by heavy diatom settlement within four weeks (Gould & Gallagher, 1990). Diatom settlement is largely from the water column, where suspended phytoplankton settles. Diatoms present in biofilm may re- suspend into the surface waters through physical processes such as waves, tides and currents, and therefore influence the primary productivity of the water column (De Jonge & van Beusekom,
1992), and the pelagic trophic food web (Consalvey et al., 2004). Before re-settling in the sediment, benthic diatoms survive in the water column. Once attached, their growth and community structure is further dependent upon the physico-chemical and biological nature of both the ambient water and the surface. Light and Beardal (1998) also stated that buried chlorophyll a may represent an important source of primary production during sediment resuspension events, based on the capacity of chlorophyll a, occurring below the sediment photic zone, to exhibit photosynthesis when subjected to light (Steele & Baird, 1968).
The rates and success of the colonization of biofilm species are heavily influenced by temperature (Hillebrand & Sommer, 1997) and immersion period (Hudson & Bourget, 1981).
Biofilm development in temperate climates generally reaches a maximum during the summer
(June and July) and decreases significantly during the winter months to a minimum in February as a result of reduced temperature, lower light intensity and shorter day-length (Chan, Chan, &
Walker, 2003).
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Ecological Importance of Biofilm
Pomeroy (1959) first pointed out the importance of microphytobenthos to primary productivity in coastal ecosystems, and it is estimated that the contribution is up to 50% of estuarine primary productivity (Underwood & Kromkamp, 1999). Microphytobenthic photosynthetic organisms can regulate oxygen concentrations at the sediment surface, which can mediate nutrient transformations and fluxes between the sediment and overlying water column
(Rysgaard, Christensen, & Nielsen, 1995). These processes can be of critical importance to the cycling of nutrients in many marine systems (Light & Beardall, 1998).
Biofilm is recognized as an important food source (Middleburg & Barranguet, 2000).
Muddy sediments usually include digestible and nutritive food sources, such as benthic microalgae or bacterial communities (Coelho et al., 2011). At low tide, diatoms and associated bacteria are concentrated in the first two centimeters of the sediment (Saint-Béat et al., 2013) and meiofauna and deposit feeders, comprising herbivorous and bacterivorous species, feed on the biofilm. The biofilm communities provide a substantial source of energy for grazers, deposit feeders, and filter feeders (through re-suspension) (Cahoon, 1999). Furthermore, some microphytobenthos species are preferred prey of invertebrate and fish species (Sullivan &
Currin, 2000). Such predation pressure can act as a top-down control as a proportional relationship between microphytobenthos production and macrobenthos biomass has been observed (Herman, Middelburg, & Heip, 2001; Hillebrand, Worm, & Lotze, 2000). Ross (1998) observed that the community with the highest microphytobenthos productivity also had the lowest total biomass; a hypothesized effect of increased predation stimulated by increases in primary production.
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More recently, biofilm has been shown to be an important food source for higher level consumers, such as the migratory western sandpiper (Calidris mauri) and dunlin (Calidris alpina) (Kuwae et al., 2008; Mathot, Lund, & Elner, 2010). Biofilm grazing as a foraging mode for western sandpiper and dunlin was first suggested by Elner et al. (2005). Mathot et al. (2010) found a large proportion of sediment in the stomach content of both species, suggesting sediments were ingested as a medium to consume biofilm. Their results showed that stomach contents for both species were dominated by volumes of sediment that exceeded invertebrate remains by an average approaching 9:1 for western sandpipers and 2:1 for dunlin (Mathot et al.,
2010).
As well as being ecologically important, biofilm existing on intertidal sediments is also known to increase sediment stability through the production of extracellular polymeric substances (EPS) which acts to bind sediment grains and form complex matrices in the surficial layers (Lucas, Widdows, & Wall, 2003). The biofilm add to the stabilization of fine sediments and increases its critical erosion threshold (Tolhurst et al., 1999; Trites, Kaczmarska, Ehrman,
Hicklin, & Ollerhead, 2005). Reductions in sediment stability can lead to a dynamic system where regular disturbances can occur, resulting in a reduction of productivity. These reductions are primarily from physical damage caused by the movement of larger sediments (i.e., sand particles in wave action) (Delgado, Jonge, & Peletier, 1991).
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Methodology
Sampling Location
To test the resilience of biofilm on an intertidal mudflat, a field experiment took place in summer 2013. The approach chosen was to identify sample and control sites, clear an area in a known biofilm and monitor re-growth over time. The experiment ran from July 17, 2013 to
August 31, 2013.
Due to the WMA, and the time restriction imposed by the tidal cycle, the sampling locations were located between the furthest South-East boundary of the WMA and the Roberts
Bank causeway, where sites could be safely reached in the same sampling period. This sampling area is known to have high densities of biofilm (WorleyParsons, 2014).
As the goal of the experiment was to evaluate biofilm resilience, the general area of the sample locations was pre-determined using ArcGIS software based on previous site data
(WorleyParsons, 2014) and not on a random design. These general coordinates were uploaded into a handheld GPS and used by field personnel to locate the area of sampling. Once the area was determined, locations were selected based on site-level factors including the avoidance of standing water and distance from drainage channels (Figure 5). For each location, two sampling plots were created: one control and one disturbed (Figure 6). The paired sampling plots were located approximately 50 metres from each other in what visually appeared to be similar physical characteristics. Locations of the established plots are provided in Figure 5 and in Table 1.
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Figure 5. Sampling locations at Roberts Bank, outside of the Roberts Bank Wildlife Management Area.
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Figure 6. Schematic representation of the paired disturbed and control sampling plots.
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Table 1
Sampling Area Locations
Site ID Treatment Northing Easting
REG01 Disturbed 490213 5433430 REG02 Controlled 490173 5433458 REG03 Disturbed 490517 5433269 REG04 Controlled 490466 5433303 REG05 Disturbed 490657 5433032 REG06 Controlled 490647 5432987 REG07 Disturbed 490449 5432768 REG08 Controlled 490408 5432730 REG09 Disturbed 490088 5433028 REG10 Controlled 490033 5433099
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Disturbed Sites
On July 17, 2013, after locating a suitable sampling location, a 5 m2 buffer zone was established using wooden stakes and rope. At the centre of the 5 m2 plot, a 1 m by 2 m sampling area was established using surveyor flags (Figure 7). Two sediment samples (Day-1) were collected before disturbance (1 sample for chlorophyll a, carbohydrate and total organic carbon
(TOC), and 1 sample for taxonomy). The technique used to collect these samples is described in the following section.
The 5 m2 plot was cleared of biofilm using a commercial squeegee. The squeegeed material was collected in a bucket and removed from the sampling location (Figure 7). The sampling area was a rectangular shape of 1 m x 2 m in the centre of the buffer zone. The sediment within the sampling area was disturbed using a rake and a shovel to mix and turn the sediments within the top 50 cm, mimicking a physical disturbance (Figure 7). This process was assumed to homogenize the sediments and their photopigment densities. Once the sampling area was completely disturbed, the sediment was smoothed using the floor squeegee (Figure 7). The disturbed plot and its paired control plots were sampled just after the disturbance (Day 0) and systematically afterwards for a period of 45 days.
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(1)
(2)
(3)
(4)
Figure 7. Photographic representation of the steps involved in creating the disturbance plots: (1) buffer area delimited by ropes and flag; sampling area marked with surveyor flags. (2) biofilm was removed from the buffer area in a bucket. (3) sampling area was disturbed using a rake and shovel. (4) levelling out with the surrounding sediment.
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Control Sites
Once each disturbance site was created, field crews located an appropriate control site, approximately 50 m away. The location was chosen based on physical features, sediment type and visual appearance. Control sites were identified with two surveyor flags (Figure 8). The technique used to collect samples was identical to those used at the disturbance site.
Figure 8. Control site marked with Surveyor Flags.
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Monitoring The sites were sampled once prior to disturbance, immediately following disturbance, and then every three to five days for a 45 day period, during a peak summer growing season. All samples were collected under Fisheries and Oceans Canada (DFO) Scientific Collection Licence
Number XR 160 2013. Each visit, 2 sediment samples (1 sample for chlorophyll a, total carbohydrate and total organic carbon (TOC), and 1 sample for taxonomy) were taken from each sample plot (disturbed and control), for a total of 20 samples per day, plus 2 duplicates. The sample acquisition method was identical for both chemical and taxonomy samples. The samples were taken during the low tide in order to provide sufficient access to all sites. The dates, tides, and time of each sampling event are presented in Table 2. A photo record was taken each sampling day to provide a visual record of each sampling location throughout the experiment.
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Table 2
Tides and Dates during Sampling Days
Date Day Tide (m) Low tide at Sampling period 17-Jul-13 D-1, DO 1.4 7:21 5:30 - 10:45 18-Jul-13 D1 1.1 8:16 6:40 - 9:50 22-Jul-13 D5 0.4 11:41 9:35 - 12:10 26-Jul-13 D9 1.4 14:39 11:25 - 13:45 30-Jul-13 D13 1.6 6:46 5:55 - 9:15 03-Aug-13 D17 1.2 10:08 8:20 - 10:45 07-Aug-13 D21 1.2 12:29 10:55 - 13:50 10-Aug-13 D24 1.7 14:08 10:35 - 13:20 14-Aug-13 D28 1.5 5:44 5:45 - 8:00 18-Aug-13 D32 0.8 9:45 8:20 - 11:15 22-Aug-13 D36 1.2 12:53 11:45 - 15:45 27-Aug-13 D41 1.6 4:57 6:20 - 8:30 31-Aug-13 D45 1.6 8:43 7:20 - 9:42
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Sampling Technique
The intended sampling technique was to use a modified 2.75 cm diameter syringe tube.
Samples were successfully collected on Day -1 and Day 0 using this technique. However, after immersion, the texture and consistency of the disturbed substrate increased in liquidity. This resulted in cores being difficult to extract such that samples could not be reliably collected. With the sampling crew already on site, and the desire to optimize the sampling day, equipment already available on site was used to carry on the experiment. Therefore, a modified technique was used on Day 1 through Day 45. Both techniques are described in the following section:
Syringe tube (Figure 9): Cores were collected at a depth of approximately 5 cm then removed and slowly ejected through the top end of the syringe using the plunger. The plunger was pushed until only the top 2 mm of the sediment core extruded the syringe, the top 2 mm was then removed from the core by using a spatula across the top edge. All 10 cores, each with a surface area of 5.94 cm2, were placed within one black centrifuge tube to create a composite sample. A new syringe was used at each sample location.
Scrape sampling technique (Figure 10): To measure the correct area to sample, a pencil
(183 mm long) was used as a length guide. Samples were collected using a 41 mm wide plastic scraper, and delicately scraping the top 2 mm of biofilm off the surface. The depth of the scrape was controlled with the angled edge of the scraper, which was exactly 2 mm. The scraper was rinsed with de-ionized water in between each sample. Based on the space available at each sample location, due to standing water or cyanobacteria, the chemical and taxonomy samples were aligned either horizontally or vertically (Figure 11).
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Figure 9. Sampling technique showing core collection steps using a modified 2.75 cm diameter syringe tube: (1). The core was collected at a depth of approximately 5 cm (2) then removed and slowly ejected through the top end of the syringe using the plunger. (3) The plunger was pushed until only the top 2 mm of the sediment core extruded the syringe. (4) The top 2 mm was then removed from the core by using a spatula across the top edge of the syringe.
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Figure 10. Sampling technique showing core collection steps using a scraper. A pencil was used as a length guide, while the depth was controlled by the angled edge of the scraper, which was 2 mm in width.
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Figure 11. Sampling style. (A) Horizontal. (B) Vertical. (C) Two samples (chemical and taxonomy) and one duplicate.
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Use of marked rope.
A marked rope was used with both techniques. Upon reaching the sampling location, a
2 m rope was laid on the ground, along the bottom line of the sampling area marked with flags.
The rope was marked into 80 mm units (Figure 12). The units on the rope used for each sample
(chemical and taxonomy) plus the duplicate were noted every time, so no two samples were taken from the same location, confirming that disturbed sediments were never re-sampled.
Figure 12: Sediment sample collection using both the syringe core technique and scraper technique, with the marker rope on the ground.
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Sample collection and storage.
Each sample taken was placed in a 60 mL black centrifuge tube. The tubes were labelled and sealed in an individual amber bag. Samples intended for chemical analysis were stored in a handheld cooler with dry ice until transferred to a -80ºC freezer at the end of the day. The samples were held in the -80ºC freezer until transferred to the laboratory for analysis. All samples were shipped on dry ice and remained below -30ºC during transport. The 143 samples
(131 samples plus 12 duplicates) were transported to Aquatech Enviroscience Laboratories
(AEL), Victoria, BC. Samples were placed in Styrofoam coolers with dry ice and transited for less than 24 hours.
The centrifuge tube with the sample intended for taxonomic analysis was filled with 5%
Lugol’s solution (acetic acid and iodine as a preservative) in filtered seawater, and gently shaken to mix the sediment and the preservative. The samples were stored in a backpack and transported to a storage box at the end of each day. The samples were held in a dark cooler until they were transferred to the laboratory for analysis. The 143 samples (131 samples plus 12 duplicates) collected in the field were shipped to EcoAnalysts Inc., Moscow, Idaho, for taxonomic identification of the microphytobenthos.
Field duplicates.
One field duplicate was collected on each day, randomly at one of the sampling sites.
Field duplicates were collected to measure the precision of sampling effectiveness. The field duplicates were collected from the same sampling area, though different core and scraper were used to collect the sample. A comparison of the field duplicate to the actual sample was conducted to establish the percent variability in data due to sample collection protocols. This
THE RESILIENCE OF BIOFILM 39 comparison could not take into account the natural variability in the sediment. However, the field duplicates were taken in the same 1 m x 2 m sampling area as the actual sample, and natural variability was deemed to be minimal.
Chlorophyll a, Total Organic Carbon and Total Carbohydrate
Once at AEL, each wet sample was weighed, freeze-dried at -60°C, and then re-weighed to determine dry weight. Once dried, the samples were homogenized using a pestle and mortar, grinding the sample until all obvious clumps had been broken and the sample was a fine powder.
A subsample of approximately 100 mg was taken for each of the analyses (chlorophyll a, TOC and total carbohydrate).
Chlorophyll a by fluorimetry.
The assessment of key photopigments, primarily chlorophyll a, is a common method of estimating biomass and the potential productivity of photosynthetic organisms in the marine environment. The strong positive relationship between photosynthetically active cells and biofilm biomass has been employed as a cost effective method of community assessment
(Pinckney, Papa, & Zingmark, 1994). Chlorophyll a is the primary photosynthetic pigment of all oxygen-producing photosynthetic organisms and is present in all algae and cyanobacteria. For this study, the concentration of chlorophyll a in each sample was measured by fluorimetry.
Fluorimetry measures fluorescence, which is the molecular absorption of light energy at one wavelength and its nearly instantaneous re-emission at another, usually longer, wavelength.
Certain substances produce fluorescence of a characteristic wavelength, allowing identification and quantification of several significant compounds in biological specimens.
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Total Organic Carbon (TOC).
In sediments, there are three basic forms of carbon: Elemental, Inorganic and Organic.
Elemental carbon includes charcoal, soot, graphite, and coal. Inorganic carbon forms are derived from geologic or soil parent material sources and are present in soils and sediments typically as carbonates, while naturally-occurring organic carbon forms are derived from the decomposition of plants and animals. In addition to the naturally-occurring organic carbon sources are sources that are derived as a result of contamination through anthropogenic activities (Schumacher,
2002). The organic matter in the sediment can be quantified by the total organic carbon (TOC), and was determined by using a thermal combustion elemental analyzer (EA) coupled via a flow reducing interface with continuous flow isotope ratio mass spectrometer (IRMS).
Total Carbohydrate.
Carbohydrates are an important component of extracellular polymeric substance (EPS) secreted by epipelic diatoms found in estuarine biofilm. Epipelic diatoms live at the water/sediment interface, and grow on the surface of sediment. Carbohydrates are composed mainly of three basic elements: carbon, hydrogen and oxygen, and are the product of photosynthesis that store the energy absorbed in the form of glucose (Castro & Huber, 2007). In the photosynthesis process, the energy absorbed by chlorophyll transforms carbon dioxide and water into carbohydrates (glucose) and oxygen (Castro & Huber, 2007). Total Carbohydrate
(TCH) levels were determined based on the amount of Glucose, and were determined using a phenol-sulfuric acid assay measured at 485 nm using a UV-Vis spectrometer.
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Quality Assurance and Quality Control (QA/QC).
The QA/QC program applied to the collection, storage and analysis of samples and was designed to confirm a systematic and reproducible program. The QA/QC program included:
. A total of 12 field duplicate samples, or approximately 9% of the samples;
. Sample inventory lists derived from field records and double checked prior to transfer to AEL;
. Sample transfers conducted in coolers packed with dry ice, at a measured temperature of at least -30ºC. At all other times, the samples were held in a -80ºC freezer;
. Review of sample temperatures recorded by the laboratory upon receipt of the samples;
. Laboratory QA/QC programs designed to confirm the precision (closeness of repeated measurements to one another) and accuracy (closeness of a measurement to the actual value) of the instruments; these were specific to each type of analysis; and . Each batch of samples included, depending on the analysis, Quality Assurance and Quality Control (QA/QC) samples: a Reagent Blank (RB), Calibration Standard (CAL) STDs, a Sample Spike (SPK), Check Standard (CHS), and a Method Replicate.
Data analysis.
Data were delivered by AEL as parts per billion (ppb), or ng/g. Glucose was used as a calibration standard for the Total Carbohydrate analysis and as such, data were presented in an
Amount of Glucose. As the technique used to collect the baseline data (D-1 and D0) was different than the technique used to collect samples on the following days, the surface area of biofilm collection was different. This difference in sampling was normalized by converting values to density estimations (mg/m2). This was conducted using Equation 1, where the Sample
Weight is the total weight of sample (g) measured before sediment subsampling.
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ng cm2 Batch results ( )*Sample Weight (g) 10,000 ( ) mg g m2 Density of Parameter ( ) = * ng (1) m2 Total Sample Area (cm2) 1,000,000 ( ) mg
The Total Sample Area of 59.39 cm2 for the Core technique was calculated by multiplying the Syringe Area (1.375 cm2 * π) by the number of cores (10). The Total Sample Area of 75.03 cm2 for the Scrape technique was calculated by multiplying the pencil length (18.3 cm) by the
Scraper width (4.1 cm).
A key assumption to this analysis is that only the biofilm within the top 2 mm were collected in the field. This conversion also makes the assumption that the measured
Chlorophyll a pigments are attributable to the microphytobenthos, which are understood to be contained within the top 2 mm of the intertidal sediments (De Brouwer & Stal, 2001; Herlory et al., 2004).
Statistical analyses.
The statistical analyses were completed using SYSTAT 13.1 (Chicago, USA). Prior to analysis, each parameter (chlorophyll a, TOC, total carbohydrate) was tested for normality using the Shapiro-Wilks test. None of the parameters were found to possess a normal distribution, therefore the data were normalized using a Log10 transformation. To analyse the data, a two- factor Analysis of Variance (ANOVA) was used to determine if the measured parameters differed between disturbed and controlled sites (Treatment), over the course of the experiment
(Day), and an interaction of these (Treatment * Day).
The results obtained with a two-factor ANOVA serve only to indicate whether the means of the dependent variable differ significantly when considering the independent variable. The
THE RESILIENCE OF BIOFILM 43 treatment effects (disturbed vs controlled) were further explored using a paired T-test. In order to determine specific differences among individual days, a Tukey’s post-hoc test was used.
The significance level of the results obtained is reported as a p-value with a number between 0 and 1. The p value or calculated probability is the estimated probability of rejecting the null hypothesis (H0) when that hypothesis is true. A small p-value (< .05) indicates strong evidence against the null hypothesis, so the hypothesis is rejected. A large p-value (> .05) indicates weak evidence against the null hypothesis, so the hypothesis is not rejected.
Taxonomic Identification
Identification and enumeration of microphytobenthos.
EcoAnalysts used the Palmer Maloney counting chamber method for counting and identifying microphytobenthos to genus level. To homogenize and evenly disperse the cells, samples were gently inverted a minimum of 30 times, then 0.1 mL of the sample was randomly collected and placed in a Palmer Maloney counting chamber using a micropipette. The counting chamber contained the sample within a defined area on a glass microscope slide.
A Leica DMIL Light Emitting Diode (LED) compound light microscope with 20X, 40X, and 63X magnification was used to visually identify microphytobenthos. This microscope was integrated with a Leica DFC450 digital camera (5 megapixel resolution). The combination of microscope and camera allowed single cells greater than approximately 4-5 µm to be identified.
Identification was focused on nanoplankton species (Table 3). Once in the counting chamber, the samples could be enumerated to determine cell concentration.
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Optimal efficiency and accuracy of identification/enumeration occurs at approximately
15-20 units per field of view. If the sample concentration was too high or low, the subsample was adjusted by adding or removing small volumes of sample and filtered seawater. If the volume was changed, the new volume and concentration were recorded on the datasheet.
Natural units were identified using the latest taxonomic references (John, Whitton, &
Brook, 2011; Pfeil, 2010; Siver, Hamilton, Stachura-Suchoples, & Kociolek, 2005; Tomas, 1997;
Wehr & Sheath, 2003). The term natural units is used to distinguish between single cell and colonial algae. Colonial algae can form aggregations of multiple cells. In this instance, the aggregation is defined as one natural unit. Taxa were identified at the genus level. Some species were able to be identified due to unique and readily identifiable traits. Microphytobenthos were identified and counted to the lowest practical taxon until at least 300 natural units were encountered. Counts were conducted using the transect method, where taxonomists follow a straight line across the entire counting chamber, enumerating and identifying all organisms within the field of view.
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Table 3
Classification of Plankton (modified from Wotton 1990)
Plankton Group Size Range (µm) Examples Megaplankton >20,000 Jellyfish, Salps, Tunicates Macroplankton 20,000-2,000 Krill Mesoplankton 200-2,000 Copepods Microplankton 20-200 Protozoa, Rotifera Nanoplankton 2-20 Diatoms Picoplankton 0.2-2 Bacteria Femtoplankton <0.2 Marine Viruses
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Quality Assurance and Quality Control.
Microphytobenthos samples were stored in optimal conditions (i.e., cool and dark) at
EcoAnalysts. Taxonomic identification included two laboratory QA/QC steps:
(1) High quality digital images were taken of each taxon encountered to permanently archive
specimens. These images included taxon name, photographer/taxonomist name, date,
representative scale bar, and project ID number.
(2) A minimum of 10% of all samples were analyzed by two phycologists to confirm taxonomic
accuracy and reproducibility of the processing and analysis methods. In the event of
discrepancies, taxonomists re-examined the digital images and/or the sample. Any
outstanding errors were adjusted in the final data, according to the recommendation of both
taxonomists. The following criteria applied to the QA/QC:
. The common algae identified by both taxonomists should match.
. Taxa accounting for more than 10% relative abundance should be identified similarly by both taxonomists.
. The percent community similarity index calculated from the two counts will meet established criteria. If any of these criteria were not met, samples were re-analyzed.
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Data analyses.
Calculating cell density.
To estimate microphytobenthos cell density in each sample, the length, width, and depth of each counting chamber were recorded and multiplied to calculate the volume. If applicable, the counted volume was multiplied by the dilution or concentration factor. This density was then multiplied by the total sample of volume (mL) to provide a total number of cells per sample. To be consistent with the chlorophyll a, TOC and total carbohydrate results, the number of cells per sample was extrapolated to the entire 1 m2 quadrat. The total number of cells per m2, for each sampling site, was calculated by using Equation 2.
N VT V CR cm 2 x SC 10,000 (2) 2 2 Surface Area cm m
In Equation 4, x is the total algal cell density, N is the total number of cells counted, Vsc is the volume of sample counted, CR is the concentration or dilution factor (if applicable, or CR=1) and VT is the total sample volume.
The Total Sample Area of 59.39 cm2 for the Core technique was calculated by multiplying the Syringe Area (1.375 cm2 * π) by the number of cores (10). The Total Sample
Area of 75.03 cm2 for the Scrape technique was calculated by multiplying the pencil length (18.3 cm) by the Scraper width (4.1 cm).
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Community diversity.
Diversity encompasses two different concepts of variety: richness and evenness.
Richness refers to the number of units per unit area, and is the simplest way to measure species diversity by counting the number of species present in a designated area. Diversity depends on the distribution of species within an area, or evenness, and refers to their abundance, dominance, or spatial distribution.
Taxonomic Diversity was calculated using the Shannon-Weiner Index (H’). The
Shannon-Weiner Index is one of the most commonly used diversity indices, and is a measure of overall biodiversity. The Shannon-Weiner index of diversity assumed that individuals were randomly sampled from an independently large population, and that all the species were represented in the sample. The index is unitless but provides a relative comparison of diversity among communities. Values range from zero (0) to infinity with low values indicating a low amount of uncertainty, and hence low diversity. The index was calculated by using Equation 5, where s equals the number of taxa; and pi equal the proportion of the ith taxa (the probability that any given individual belongs to the taxa).
s H' pi ln(pi ) (3) i1
The Evenness (E) component of H’ was calculated by using Equation 4, where H’ is equal to the Shannon-Weiner Diversity Index; and H’max is the maximum possible value of H’.
H’max = LnS, where S equal the number of identified taxa.
H' H' E (4) H'max LnS
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Evenness index numbers are presented between 0 and 1. An Evenness index close to 1 indicates an even distribution of abundance amongst species, while a number close to 0 indicates an uneven distribution.
In ecology, ‘richness’ is often used interchangeably with ‘diversity’. For this study, richness is used as the number of different microphytobenthos species found within the sample plots, per day. Taxonomic Richness (R) was calculated for each sampling location. Richness was calculated as the number of taxa identified within each sampling plot, per day.
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Results
Chlorophyll a, Total Organic Carbon and Total Carbohydrate
Following disturbance, 41% of the chlorophyll a density remained on the sampling site, while 64% of the total carbohydrate density remained (Table 4). However, the density of the total organic carbon level showed an increase of 15% after disturbance (Table 4).
For chlorophyll a, the pre-disturbance mean density value (98.70 ± 7.81 mg/m2) for the disturbed sites was reached between Day 32 (74.81 ± 6.986 mg/m2) and Day 36 (101.34 ±
16.871 mg/m2). The same situation happened with TOC, which reached its pre-disturbance mean density value (25,719.87 ± 1,728.46 mg/m2) between Day 32 (18,121.12 ± 665.76 mg/m2) and Day 36 (26,834.70 ± 3,223.82 mg/m2). Total carbohydrate mean density value never reached the pre-disturbance level (12,915.08 ± 1,364.53 mg/m2), but was the closest on Day 41
(12,072.28 ± 1,877.18 mg/m2).
When normalized using a Log10 transformation, the data showed a temporal trend for all three variables (chlorophyll a, TOC and total carbohydrate) (Figure 13). A decrease was observed for both disturbed and controlled sites on Day 1. The variable slowly decreased until
Day 9, where the value remained quasi-constant or slightly increased until Day 28, where they started to increase (Figure 13).
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Table 4
Mean density prior and immediately following disturbance (mg/m2)
Day -1 Day 0 % Difference
Chlorophyll a 98.70 ± 7.81 41.39 ± 0.94 -59%
TOC 25,719.87 ±1,728.46 29,742.16 ± 2,522.92 +15%
Total carbohydrate 12,915.08 ± 1,364.53 8,267.48 ± 526.28 -36%
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Figure 13. Relationship between Log10 (chlorophyll a), Log10 (total organic carbon), Log 10(total carbohydrate), and Day.
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Statistical results.
The two-factor ANOVA analysis considered the effect on two separate defined factors, and their interaction, over an independent variable.
For chlorophyll a, the small p-value (˂ .05) indicates that the H0 is rejected. A significant difference was found in the density between disturbed and controlled (Treatments), among Days, and with the interaction between Treatment and Day (Table 5).
For TOC, the p-value was significant (p ˂ .05) for the difference between days only.
There was no difference between the disturbed and controlled sampling location based on the
TOC density, as well as no significance difference between the treatment (disturbed or controlled) and the sampling day (Table 5).
A significant difference (p ˂ .05) in the mean of total carbohydrate levels was found between control and disturbance (Treatments), among Days, but no interaction between
Treatment and Day (Table 5).
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Table 5 Results of ANOVA two factor Analysis with Log10 (chlorophyll a, TOC or total carbohydrate) as Dependent Factors and Treatment and Day as Independent Factors.
Sources df Mean F-ratio p Squares
Chlorophyll a Treatment 1 0.673 39.764 .000 Day 13 0.143 8.447 .000 Treatment * Day 13 0.056 3.329 .000 Total organic carbon Treatment 1 0.000 0.007 .934 Day 13 0.190 14.096 .000 Treatment * Day 13 0.009 0.678 .781 Total carbohydrate Treatment 1 0.316 7.058 .009 Day 13 0.142 3.175 .000 Treatment * Day 13 0.046 1.019 .439 Note: df = degrees of freedom; F = test statistic; p = p-value. Numbers in bold indicate significance of the p-value.
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The significant treatment (disturbed or controlled) effects were further explored using a paired T-test. Comparisons of controlled and disturbed treatments were completed on each individual day. Within the T-test, an adjustment method was used, the Bonferroni correction, in which the p-values are multiplied by the number of comparisons.
For chlorophyll a, based on the p-value, no significant difference was found between treatments prior to disturbance (Day -1), concluding that the two groups were statistically the same prior to disturbance. However, significant differences (p ˂ .05) between Treatments were observed on Day 0, Day 1 and Day 5 (Table 6) (Figure 13).
For TOC, no significant differences were found between Treatments on any days using the Bonferroni Adjusted p-value; however, using the more liberal uncorrected p-values, differences were found on Day 17 (Table 6). The same results were found for total carbohydrate, where no significant differences were found between Treatments on any days using the
Bonferroni Adjusted p-value; but using the p-values, differences were found on Day 0 and Day 1
(Table 6) (Figure 13).
The results indicate that after Day 9 for chlorophyll a and Day 5 for total carbohydrate, there was no difference between the disturbed and controlled sampling location (H0 was rejected). Day 17 was the only day where there was a significant difference for TOC (Table 5)
(Figure 13).
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Table 6 Results of Two-Sample t-test Analysis with Log10 (chlorophyll a, TOC and total carbohydrate) for each sampling day, grouped by treatment (Disturbed and Controlled).
Log10 (chlorophyll a) Log10 (TOC) Log10 (total carbohydrate) t p-value B. A. T p- B. A. t p-value B. A. p-value value p-value p-value
D-1 1.136 .289 .867 1.039 .329 .988 0.922 .384 .000 D0 6.473 .000 .001 0.278 .788 .000 2.862 .021 .063 D1 3.675 .006 .019 0.324 .754 .000 2.305 .050 .150 D5 3.114 .014 .043 0.980 .356 .000 1.913 .092 .276 D9 0.863 .413 .000 -0.740 .480 .000 0.229 .824 .000 D13 1.404 .198 .594 0.357 .730 .000 0.520 .617 .000 D17 0.213 .837 .000 -2.386 .044 .132 -0.022 .983 .000 D21 1.248 .247 .741 -0.074 .942 .000 -0.401 .699 .000 D24 0.265 .798 .000 -0.750 .475 .000 -0.209 .840 .000 D28 0.720 .492 .000 -0.766 .466 .000 0.278 .788 .000 D32 1.542 .162 .485 0.545 .601 .000 0.918 .385 .000 D36 0.661 .527 .000 -0.064 .951 .000 0.878 .406 .000 D41 1.896 .107 .320 1.361 .222 .667 0.668 .529 .000 D45 0.257 .806 .000 -0.534 .612 .000 -0.354 .736 .000 Note: B. A. p-value = Bonferroni Adjusted p-value. Degree of freedom (df) was 8 for D-1 to D36, and 6 for D41 and D45. Numbers in bold indicate significance of the p-value.
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The significant day effects were explored using a post-hoc Tukey’s test where controlled and disturbed sampling sites were grouped per day for each variable. The Tukey's test compared the means of each individual day to the mean of every other day. It was applied simultaneously to the set of all pairwise comparisons and identified any difference between two means that was greater than the expected standard error.
For the chlorophyll a variable, the post hoc Tukey test result showed a significant difference (p ˂ .05) between baseline conditions (Day -1) and Day 1, Day 5, Day 9, Day 1,
Day 17, Day 21 and Day 28. No significant differences with pre-impact samples were observed past Day 28 (Table 7).
The post-hoc Tukey test on total organic carbon showed a significant difference (p ˂
0.05) between baseline conditions (Day -1) and Day 5, Day 9, Day 13, Day 17, Day 21, Day 24 and Day 28; no significant differences were observed past Day 28 (Table 7).
For total carbohydrate, the post hoc Tukey test showed a significant difference (p ˂ .05) between baseline conditions (Day -1) and Day 5, Day 9 and Day 13. No significant difference was noted between Day -1 and Day 0 or Day 1. No significant differences were observed past
Day 13 (Table 7).
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Table 7 Results of Tukey Pairwise Comparison test for Log10 (chlorophyll a, TOC and total carbohydrate), controlled and disturbed sampling sites tested together, grouped per day.
Chlorophyll a TOC Total carbohydrate
Day (i) Day Difference p-Value Difference p-Value Difference p-Value (j) between between between the means the means the means D-1 D0 0.186 .093 -0.030 .000 0.093 .999 D-1 D1 0.252 .003 0.044 .000 0.222 .529 D-1 D5 0.315 .000 0.195 .018 0.334 .037 D-1 D9 0.395 .000 0.393 .000 0.362 .015 D-1 D13 0.284 .000 0.305 .000 0.325 .050 D-1 D17 0.205 .039 0.249 .000 0.238 .409 D-1 D21 0.223 .015 0.301 .000 0.256 .292 D-1 D24 0.194 .065 0.286 .000 0.244 .364 D-1 D28 0.247 .003 0.297 .000 0.283 .160 D-1 D32 0.106 .864 0.172 .069 0.107 .997 D-1 D36 0.026 .000 0.034 .000 0.036 .000 D-1 D41 0.017 .000 0.048 .000 0.037 .000 D-1 D45 0.041 .000 0.052 .000 0.133 .987 Difference of the means = difference of the means of each individual day to the mean of every other day. Numbers in bold indicate significance of the p-value.
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Microphytobenthos Community Structure
When extrapolated into the total number of individuals per square meter for the duration of the experiment, a total of 590 million microphytobenthic individuals were found at the five combined disturbed sites, while 775 million individuals were found at the five combined controlled sites.
In total, 67 different taxonomic groups were identified. The five main taxonomic species found at Roberts Bank, for the duration of the experiment, and all the sampling sites combined, are presented in Table 8. Nitzschia sp. numerically dominated the microphytobenthic community, followed by the sub-dominant Leptolyngbya sp. Gyrosigma/Pleurosigma, Navicula and Amphora sp. These were followed by 62 other taxa which comprised the remaining 30.17% of the community.
Figure 14 presents the five taxonomic species found with the highest abundance at each sampling site during the experiment. Nitzschia sp. dominated the sampling sites with the exception of REG02, which was dominated by Leptolyngbya sp. Leptolyngbya sp. was the second most abundance species at almost every sampling site. Overall, the same species were present on each sampling day, but in different relative proportions.
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Table 8
Main taxonomic species found at Roberts Bank for all sampling sites.
Main species % of the mean Type
Nitzschia sp. 30.89 Diatoms
Leptolyngbya sp. 24.46 Cyanobacteria
Gyrosigma/Pleurosigma 5.43 Diatoms
Navicula sp. 5.26 Diatoms
Amphora sp. 3.79 Diatoms
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Figure 14. Five microphytobenthic species found at Roberts Bank with the highest abundance, at each sampling site. REG 01, 03, 05, 07 and 09 are disturbed sampling sites, REG 02, 04, 06, 08 and 10 are the control sites.
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Microphytobenthos Community Diversity
Following the disturbance, the microphytobenthos abundance experienced a sharp decrease
(Figure 15). Taxonomic richness also experienced a significant decline after D-1 (Figure 15).
However, as both the disturbed and controlled sites experienced the decline, the manual disturbance cannot be assumed to be the cause and therefore could be due to an external environmental variable not taken into account in this study. Conversely, taxonomic evenness experienced an increase following disturbance, for both disturbed and controlled sampling sites
(Figure 15). Taxonomic diversity, based on the Shannon-Weiner Index, was variable throughout the entire experiment across the sampling sites and between the paired sampling sites (Figure
15).
Microphytobenthos abundance, diversity, evenness and richness did not return to baseline conditions until Day 32, a date that coincided with the beginning of the spring tide.
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Figure 15. Microphytobenthos Abundance, Shannon-Weiner Index (H’), Taxonomic Evenness (E) and Richness(S) for the duration of the experiment, for disturbed and controlled sampling sites.
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Relationship to the tidal cycle
When comparing the values for chlorophyll a, TOC and total carbohydrate, as well as the community structure indices, a clear relationship appears between the variation in concentrations and the tidal pattern. For each variable, the values started to decline toward the end of the spring tide, and increased around Day 28, which is also the start of the following spring tide.
Discussion
The variables (chlorophyll a, TOC, total carbohydrate, taxonomic communities) examined during this study provides an indication of how a biofilm community would regenerate in the event of disturbance. More specifically, links were observed between recovery and the tidal and hydrologic conditions.
Chlorophyll a, Total Organic Carbon and Total Carbohydrate
An immediate decrease in chlorophyll a and total carbohydrate was observed at the disturbed sites compared to the control sites. This decrease was observed across all the disturbed sites. Chlorophyll a concentrations returned to undisturbed levels after nine days, while carbohydrate levels returned to an undisturbed condition five days after disturbance. However, some variability did occur with the individual plots, indicating the actual recovery period may have been as long as nine days for carbohydrate.
Several studies have described the link between chlorophyll a and extracellular carbohydrates in intertidal mudflats. Chlorophyll a production is linked to the amount of light available for absorption as well as available nutrients. Total carbohydrate, a product of photosynthesis, therefore has level closely related to chlorophyll a (Sutherland, Grant, & Amos,
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1998; Underwood, Paterson, & Parkes, 1995; Underwood & Smith, 1998). The secretion of EPS
(comprised of carbohydrates) is believed to be a mechanism by microphytobenthos to avoid re- suspension during immersion (Easley et al., 2005). These same microphytobenthic taxa are photosynthetic and, with increased carbohydrate secretion, are able to remain associated with the sediments and grow. Due to this dependence, a linear relationship between carbohydrates and chlorophyll a occurs within biofilm.
TOC levels were similar between disturbance and control sites throughout the study period, and in some cases, increases at the disturbance sites were observed. This increase might be due to the action of churning the top 50 cm of sediment during disturbance. The maximum value of mean TOC at the disturbance sites was observed immediately following disturbance.
The organic carbon content of marine and estuarine sediments is determined by multiple factors, including sediment grain size, organic matter source, sedimentation rate, and preservation rate (Pelletier et al., 2011). Organic carbon is a product derived from decaying vegetation, bacterial growth, and metabolic activities of living organisms or chemicals. In the marine environment, organic carbon tends to accumulate in low energy environments which are comprised of fine sediments such as silt and clay (Bergamaschi et al., 1997; Mayer et al., 1985;
Sutherland, Petersen, Levings, & Martin, 2007). Given the location of the study site, organic carbon inputs are likely introduced from the Fraser River as well as marine drift (i.e., macroalgae and eelgrass); these are in addition to carbon provided by the microphytobenthos. Sutherland,
Elner, & O’Neill (2013) noted that the depositional conditions in the landward northern corner of the Roberts Bank causeway, where the regeneration plots were established, have led to an increase in organic matter content, namely in association with the development of porous, silt-
THE RESILIENCE OF BIOFILM 66 dominated sediments. This indicates the location of the regeneration plots were in an area of high organic carbon deposition. This continual deposition would likely result in the accumulation of carbon, leading to high carbon content with the upper sediments.
Microphytobenthos Community Structure
At Roberts Bank, the freshwater inputs of the Fraser River expose the microphytobenthos community to a full range of marine, brackish and freshwater species throughout the year. The greater range of environmental conditions can lead to a scenario where frequent, and low intensity disturbances prevent dominance by a single taxa (Connell, 1978). At Roberts Bank,
Nitzschia sp., a strictly marine species (Spaulding & Edlund, 2008b), was dominating throughout the experiment, although its proportion varied between sampling days and location. The change in dominant species was observed for both the disturbed and controlled sampling sites. This indicates that unmeasured environmental variables likely impacted the species present on each sampling day.
Several authors have reported that spatial distribution patterns of epipelic diatom species are related to salinity and nutrient gradients or to tidal elevation (Oppenheim, 1991; Underwood,
Phillips, & Saunders, 1998). In estuaries, diatom species have been shown to tolerate a wide range of environmental conditions (Admiraal, 1984) as well as having defined broad ecological niches (Underwood et al., 1998; Underwood & Provot, 2000). Two of the species found
(Achnanthidium sp. and Melosira sp.) (Figure 14) are freshwater diatom species (Potapova,
Spaulding, & Edlund, 2008; Spaulding & Edlund, 2008a). The presence of these two species could be linked to the tidal cycle; with a lower density, the freshwater sits on top of the marine water and can be pushed further across the mudflat on a flood tide. Amphora sp.,
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Gyrosigma/Pleurosigma and Navicula sp. (Figure 14) are found in marine and freshwater habitats (Spaulding & Edlund, 2009; Spaulding, 2011; The University of British Columbia,
2012). While Navicula sp. was never the dominant species on any sampling location, it is an important contributor to biomass and productivity (The University of British Columbia, 2012).
Leibleinia sp and Leptolyngbya sp., the two remaining species with the highest abundance
(Figure 14), are cyanobacterial species. Cyanobacteria are a group of photosynthetic bacteria that are widely distributed in the marine and freshwater environment and many species can tolerate a wide range of salinity and temperature (Komarek, 1992). Cyanobacteria are the most abundant members of the picoplankton and account for at least half the ocean’s total primary production (Castro & Huber, 2007).
Microphytobenthos Community Diversity
A sharp decrease in taxonomic abundance was observed on Day 1, for both control and disturbance sampling sites. This indicates that this loss of abundance was not due to the disturbance process, but to an uncontrolled environmental or physical factor. The taxonomic abundance did not come back to pre-disturbance level until Day 32.
On Day 1, the taxonomic abundance at the disturbed sites was lower than at the controlled sites (Figure 15). This decrease is attributed to the removal of the established biofilm community. The initial removal of biofilm was assumed to have removed the microphytobenthos from the sediment surface. Following this, colonization would have been due to settlement of water column phytoplankton or re-suspended microphytobenthos, or by the growth of species that were not removed from the sediments. This colonization would have been
THE RESILIENCE OF BIOFILM 68 dependent upon the densities in the water column and the surrounding sediments. Additionally, with the removal of biofilm, any settled microphytobenthos would be susceptible to removal without the development of the biofilm EPS. As Tolhurst et al. (2008) reported the development of the EPS layer can take up to five days, it is likely that the protective layer had not been fully developed one day following disturbance. Without a fully developed EPS layer, the rate of microphytobenthos establishment and density can be expected to be reduced.
Microphytobenthos diversity.
After an initial increase on Day 1, the diversity index across the sampling sites at Roberts
Bank remained constant between 1.3 and 1.8 until Day 28, where it decreased below the average number, especially at the disturbance sites. The sharper decrease was experienced between Day
28 and Day 32 for the disturbance sites. At the same time, taxonomic richness decreased and evenness increased. The increase in evenness combined with decrease in richness allows for the conclusion that not only were the most dominant taxa reduced in density, but other species were entirely removed from the community. As this was noted in both the disturbed and controlled treatments, it does not appear that there is a pronounced succession or competition of species.
As such, the microphytobenthic community is shown to maintain consistent levels of taxonomic diversity during periods of natural temporal variation.
After reducing following disturbance, the taxonomic richness remained lower until D32, when the levels started to increase toward their original levels. This period coincides with the beginning of the spring tide, which create more mixing in the water. This mixing and input of more water could have brought in new species, as well as providing enough nutrients for less dominating species to settle. Two factors also promoted biofilm growth during that period: the
THE RESILIENCE OF BIOFILM 69 longer emersion period associated with Spring tide, and the low water events occurring during day time.
Oldeland, Dreber & Wesuls (2010) stated that the overall biological activity of an environment can be evaluated by the evenness of species distribution. A low evenness means a high biological activity, due to at least one dominant species, while a decreased evenness can be due to the reduction of resources, favouring the dominance of an increased sub species (Oldeland et al., 2010). Immediately after disturbance (D0), the taxonomic evenness of both the controlled and disturbed sampling location increased, indicating a natural environmental effect not linked to the physical disturbance. The evenness remained stable until Day 28, when a sharp decrease was observed. As with taxonomic richness, this can be linked to the beginning of the spring tide; an increase of species richness across the mudflat led to a reduction of the overall species evenness.
Natural Variation
The results of this study allowed a clear relationship to be seen between biofilm density and composition and the tidal cycle. Several authors have shown the tidal influence on microphytobenthos communities (Jesus et al., 2009; Patil & Anil, 2005; Ribeiro, Brotas, Rincé,
& Jesus, 2013). These authors have identified the spring-neap tidal cycle as an important factor for influencing phytoplankton growth in estuaries.
The timing of this study was designed to commence with a manual disturbance on July 17,
2013, at the onset of a large spring tide (Figure 3). This time was selected to maximize the duration of site emersion for sampling purposes. As an effect of this timing, sampling began during a spring tide, before maximum tidal ranges were achieved. Over the course of the
THE RESILIENCE OF BIOFILM 70 remaining spring tide, a continuous decrease in biofilm biomass was observed. The minimum values of chlorophyll a, TOC and total carbohydrate were observed on Day 9, half-way through the neap tide. It is possible that observed decreases in biofilm biomass compared to baseline conditions (D-1) were a result of the dynamic shift in the intertidal environment marked by a direct link to the spring-neap tidal cycle.
During the following neap tide, and subsequent smaller spring tide, biofilm productivity remained low. This period occurred between Day 9 and Day 28. On Day 28, the next spring tide commenced and was at its near peak tidal range by Day 32. This coincided with a return to baseline (D-1) biofilm conditions. The spring tide that began on Day 28 possessed a lower maximum tidal range compared to the initial spring tide experienced at the commencement of the study. This pattern shows that the biofilm biomass and microphytobenthos community experienced higher density and productivity in the lead up to and during spring tides; however, during the peak tidal range spring tides, microphytobenthos were removed from the biofilm.
Dransfeld (2000) reported that planktonic and benthic diatoms are photosynthesizing during tidal emersion and, depending on light penetration, during inundation. Diatoms and cyanobacteria that are not attached to the substrate are suspended in the water column during tidal mixing, where the photosynthetic activity continues. Due to this cycle of suspension and deposition, diatoms and cyanobacteria are continuously redistributed, a process which is directly influenced by water currents and velocity. During periods of standing water, phytoplanktons settle out of the water column and establish on top of intertidal substrates where production of
EPS permits motility within the sediment.
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On a daily tidal cycle, phytoplankton settlement is most likely to be greatest during the
Slack High Water. The semi-diurnal nature of the tides at Roberts Bank and the asymmetric nature of the high waters suggest that the highest Slack Water will provide the greatest potential for phytoplankton settlement due to the lower initial velocities, resulting in a longer period of calm water. However, during the first Ebb Tide, shear stress at the biofilm-water interface will likely be greatest due to the large tidal range; this will have the highest potential for removal/resuspension of microphytobenthos.
Dominance of microphytobenthic diatoms in the water column is often reflected in the biofilm microphytobenthic community and productivity, suggesting that hydrodynamic conditions might play an important role in the biofilm diatom community structure (Patil & Anil,
2005). Patil and Anil (2005) found that maximum biofilm diatom density was found during calm periods such as the slack period rather than during highly disturbed period such as ebb tides, allowing the water column phytoplankton to settle in the intertidal sediment. Furthermore,
Dransfeld (2000) reported reductions in biofilm densities during spring tidal periods which were attributed to the re-suspension of microphytobenthos by stronger tidal currents. However, the species composition and their percentage contribution to the biofilm diatom community varied with time and the exposure period (Patil & Anil, 2005). This change in species composition might explain the enhanced biodiversity found by Levings and Thom (1994) in intertidal areas with fast tidal current.
The increased tidal currents often observed on the ebb flows, and more pronounced during a spring tide, also increase the bed shear along the sediment-water interface, and thus increase phytoplankton re-suspension in the water column (Lauria, 1998).
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Resilience of Biofilm to a Major Event
Sea level rise.
Currently, Roberts Bank experiences a tidal cycle which is semi-diurnal, having two low and two high tides, with different heights, over a day. Different areas of the mudflat have different exposure periods, based on elevation and location. With predicted sea level rise, areas that were previously exposed at low tide might be exposed only at the lowest low tides, or only during the spring tide.
Hill et al. (2013) reported that for the Roberts Bank area, a 60 cm rise in relative sea level may be experienced by 2035 and a 1 m rise by 2060. A simple scenario of the mudflat response to rising sea level was created by retaining the present elevation range with respect to the higher sea level. The simulation suggests that the mudflat surface would be substantially reduced to
56% for a 0.62 m sea level rise, and to 40% for a 1.02 m sea level rise (Hill et al., 2013). Should this level of sea rise occur the area available for optimal biofilm growth would be greatly reduced, exacerbated by the process of coastal squeeze by which the area of intertidal habitat is reduced, thus preventing biofilm from migrating inshore due to the physical presence of hard coastal defences (Fujii, 2012). Tidal wetlands and estuarine areas that would normally migrate inland as sea level rises would be unable to do so in many areas of the Fraser River delta because of the dyke barriers (Taylor, 2004).
Based on the need for microphytobenthos to settle in muddy, nutrient-rich substrates, and the incapacity for the shoreline to recede, the resilience of biofilm at Roberts Bank would be reduced as the sea level rises.
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Climate change.
The Fraser River has an average annual flow of 3630 m3/s (Cameron, 1996). The Fraser
River, which is fed by rain and snowmelt, has an increase in March, a large peak freshet in June
(up to 10 000 m3/s), gradually decreases in July, August and September, and then remains near the minimum levels (as low as 700 m3/s) throughout the rest of the year (Yin et al., 1997;
Johannessen, Macdonald, & Paton 2003; Masson & Cummins 2004). Climate change could result in changes in the intensity and timing of the Fraser River’s peak flow in the spring (Taylor,
2004). Within a climate warming scenario, much of the precipitation over the delta and the surrounding coastal mountains would fall as rain rather than snow, which could result in winter precipitation increases by 200 mm and summer precipitations decreases by 40 mm (Taylor,
2004).
An increase in the intensity of the Fraser River’s freshet also has the potential to increase the turbidity in the estuary. As turbidity plays an important role in light availability to the microphytobenthos, it could be expected that the photosynthetic potential would be impacted.
Anthony et al. (2004) noted that suspended sediments can generate up to 80% of the variation on light availability. Diatoms are known to generally be adapted to low light levels (Lionard,
Muylaert, Gansbeke, & Vyverman, 2005) as well as having developed photo-adaptive mechanisms such as up-regulation of photopigment (Jesus et al., 2009) and vertical migration
(Underwood et al., 2005). These mechanisms allow diatoms to survive in a turbid estuary. A study by Pratt et al. (2014) demonstrated the consequences of increased turbidity on the productivity of microphytobenthos in estuarine intertidal sandflat systems. A three-fold reduction in rates of net primary production was observed, as well as greater effects on photosynthetic efficiency.
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Light also plays a major role in nutrient exchange as microphytobenthos regulate the flux of nutrients remineralised in sediments to the water column through the photosynthesis process
(Pratt et al., 2014). Therefore, an increase in turbidity due to greater riverine run-off is likely to have a significant impact on the productivity and nutrient exchange in the estuary, impacting the biofilm biomass.
Conclusion and Recommendations
The purpose of this study was to examine the resilience of biofilm on an intertidal mudflat following habitat loss or disturbance. Using chlorophyll a, total organic carbon, total carbohydrate as well as the taxonomic community as variables, the re-establishment of biofilm on an intertidal mudflat was monitored over a 45 day period. This study allowed a number of conclusions:
(1) After nine days, there is no significant difference between the disturbed and control condition;
(2) There is a strong relationship between natural variability in biofilm biomass and the lunar cycle;
(3) Pre-disturbance levels are reached after 29 to 32 days, and is likely linked to the tidal cycle;
(4) Natural variations in biofilm biomass and community composition occur at Roberts Bank; and
(5) The presence of a dyke along the shore will prevent the natural movement of the mudflat inland as sea level rises in the future.
Based on previous published studies discussed in the literature review and the results of this experiment, biofilm has proven to be resilient to disturbance. However, as this experiment
THE RESILIENCE OF BIOFILM 75 was performed during the peak biofilm growing season, different results and timings could be present at different times of the year and should be evaluated in the future.
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