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Effects on biofilm growth due to the interactions between different levels of dissolved organic carbon… Chen, Jennifer Apr 20, 2015

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1           Effects on biofilm growth due to the interactions between different levels of dissolved organic carbon and nutrients    by Jennifer Chen    Faculty of Forestry University of British Columbia B.Sc. Forest Sciences    Primary Supervisor: John Richardson Secondary Supervisor: Scott Hinch    April 20, 2015              2  ABSTRACT   Biofilms consist of a complex community of microscopic organisms from various taxa that play an important role in any freshwater ecosystem. They perform various ecological functions, such as decomposition of organic matter, nutrient cycling, oxygen availability, water quality, and making up the base of the food web. From a management perspective, biofilms are also an indicator of ecosystem health and productivity since their overall growth can be determined by the amount of dissolved organic carbon (DOC) and nutrients available in the system. While many studies have examined the effects of varying nutrient conditions, there has been less research on the interactive effects between the two factors. This study, done through a fully crossed factorial experimental design, examines a gradient of increasing DOC vs. increasing nitrogen and phosphorus (N:P) concentrations. The addition of grazers Glossosoma penitum was used to determine the effect of grazers on biofilm accumulation. While the experiment itself did not show evidence of any statistically significant interactions between DOC and N:P (p-value 0.098 for biomass, and 0.366 for chlorophyll-α), the subsequent literature search yielded other studies that examine a significant coupling effect, as well as significant grazer influence on overall biomass and algal proportions.   Key words: biofilm growth, nutrient interaction, nitrogen, phosphorus, dissolved organic carbon                            3  TABLE OF CONTENTS  Abstract ...........................................................................................................................................2 Table of Contents ...........................................................................................................................3 List of Figures .................................................................................................................................0 List of Tables ..................................................................................................................................0 1. Introduction ................................................................................................................................5 1.1 Background ............................................................................................................................5 1.2 Biofilm Functions ...................................................................................................................5 1.3 Objectives ...............................................................................................................................8 2. Materials and Methods ..............................................................................................................9 2.1 Experimental Design ..............................................................................................................9 2.2 Biomass ................................................................................................................................11 2.3 Chlorophyll-α Extraction and Readings ...............................................................................12 2.4 Statistical Analyses ..............................................................................................................13 3. Results .......................................................................................................................................13 3.1 Grazer Influence ...................................................................................................................13 3.2 Overall Biomass ...................................................................................................................14 3.3 Chlorophyll-α .......................................................................................................................15 3.4 Biofilm Proportions ..............................................................................................................15 4. Discussion..................................................................................................................................16 4.1 Effect of Grazers ..................................................................................................................16 4.2 Total Biomass .......................................................................................................................18 4.3 Biofilm Proportions ..............................................................................................................20 4.4 Nutrient Interactions .............................................................................................................22 4.5 Future Research ....................................................................................................................23 5. Conclusion ................................................................................................................................25 Acknowledgements ......................................................................................................................25 References .....................................................................................................................................26 4  LIST OF FIGURES   Figure 1. Average biomass given DOC concentrations ................................................................29 Figure 2. Average biomass given nutrient concentrations ............................................................29 Figure 3. Average chlorophyll-α given DOC concentrations .......................................................30 Figure 4. Average chlorophyll-α given nutrient concentrations ...................................................30 Figure 5. Proportions of total to algal biomass .............................................................................31  LIST OF TABLES  Table 1. Statistical results for biomass ..........................................................................................31 Table 2. Statistical results for chlorophyll-α .................................................................................31                           5  I. INTRODUCTION   1.1 Background   Biofilms, in a broader sense, are defined as a community of bacteria or other microorganisms, such as algae, protists and fungi that form a layer on a solid surface. In the case of stream environments, they can be found on materials such as boulders, large wood, plastic materials, or on aquatic plants. As with any biotic community, they can be quite complex and the different components have varying ecological functions, both autotrophic and heterotrophic. Freshwater biofilms generally have two major portions, bacteria and algae, made up of taxa such as green algae, cyanobacteria, and diatoms (Sekar et al. 2004; Rickard et al. 2004). A study by Barranguet et al. (2005) examined the changing composition of biofilms over time and show that they follow a general “life-history” cycle. The biofilm initially starts off as heterotrophic bacteria colonizing a surface, followed by microalgae; this microorganism mixture forms extracellular polymeric substances (EPS) that hold the biofilm together, allowing for interactions between the two taxa and further colonization that increases biofilm thickness and some level of protection against grazers and moving waters. As the biofilm ages and thickens, overall photosynthesis decreases due to obstruction of light from continuous microorganism accumulation. Eventually, the biofilm ages to a point where shear stress from a moving current can easily displace the entire colony from the surface, allowing the cycle to begin again.    1.2 Biofilm Functions   In general, biofilms found in stream ecosystems are an important nutrient source since they provide the necessary energy and nutrients for higher trophic levels. As a major source of authochthonous energy, it is not surprising that biofilms also function as regulators of other 6  ecosystem components such as nutrient cycling, water quality, and being a part of the microbial food web. Much of the decomposition of organic matter in a stream is done by microscopic fungi and bacteria which contribute to the overall availability of those nutrients (Battin et al. 2003). The study by Lovatt et al. (2014) also examines how light and leaf litter leachates in varying quantities indirectly affect the overall composition of biofilm by changing the concentrations of phosphorus compounds and DOC in the stream. Since the leachates had a more significant impact under abundant light conditions, there are implications that autotrophic portions of biofilms benefit more from an increased input of DOC from leaf litter. This would indicate that the sources of DOC are also important in determining how biofilms partition their energy uptake.   The amount of ecosystem respiration performed by biofilm is also an indicator of overall productivity since the ecological process is related to biomass levels. The study by Haggerty et al. (2014) supports the fact that the increase in biofilm accumulation is highly correlated with the increase in ecosystem respiration. Understanding the rates of oxygen consumption is important from a management perspective since many aquatic organisms rely on a minimum concentration of oxygen. Various fish and invertebrate species will have different oxygen tolerance levels, but understanding how biofilms will respond to any changes in nutrient concentrations will determine how the stressors will affect larger organisms. Ultimately, changes in nutrient concentrations will be tied with oxygen availability because of how the biofilms use the increase or decrease of nutrients. Decomposition processes and ecosystem respiration are then linked to water quality which can be evaluated under different factors, such as oxygen consumption, turbidity, and nutrient spiraling (Kohler et al. 2010).      In addition to modifying water quality and nutrient availability, the microorganisms that make up biofilm are an important part of the microbial food web. Grazers such as the New 7  Zealand mud snail, Potamopyrgus antipodarum, can directly feed on the biofilms (Barranguet et al. 2005), followed by secondary consumers that then feed on the snails. Bacteria and fungi can feed on much finer organic matter that would generally be too small for other invertebrates to consume directly. This provides a pathway for fine particulate organic matter to travel through higher trophic levels and further contributes to the autochthonous energy input (Hargrave et al. 2010).  Understanding the complex food web interactions related to biofilm will also determine whether any trophic cascades are plausible given the situation where nutrient concentrations do change. Holomuzki et al. (2010) mention that herbivores have significant impacts on biofilm growth and that under high nutrient conditions, the increased biofilm biomass can support a  larger number of invertebrates which ties into other ecological processes, such as the increase in disease or parasite transfer in these grazers, or supporting a larger population of secondary consumers.    In addition to contributing to ecological processes, biofilms can also be used as an indicator in ecological risk assessments. McClellan et al. (2008) examine the retention of herbicides according to how biofilm organisms can take in the pollutants. The uptake of pollutants significantly altered the biofilm composition which can then be transferred to higher trophic levels. While the specific bioaccumulation effects were not examined in this study, understanding the uptake and retention of various pollutants through biofilm microorganisms is essential for identifying potential issues and thus restricting the use of industrial chemicals. Industrial applications for biofilms are also prevalent especially in the case of constructed wetlands where the filtration properties can be impeded or improved by the growth of biofilms. In the study by Zhao et al. (2009), the authors examined the clogging process of various biofilms in constructed wetland filtration systems and have concluded that there are cases where a 8  moderate level of biofilm accumulation can positively contribute to the filtration process since the microorganisms involved can perform the necessary decomposition of certain pollutants. Overall, biofilms are an important component of any natural system and can also be used in various industrial applications.     1.3 Objectives As with any other autochthonous source of energy, the overall productivity of biofilms is dependent on the amount of nutrients in the ecosystem. In most cases, nitrogen and phosphorous are the major limiting elements due to the fact that they make up an essential component in proteins which then function as enzymes or form overall tissue structure. While it is apparent that these nutrients are very important to an ecosystem as a whole, there have been fewer studies that examine some of the interactions between different nutrients. Crain et al. (2008) have found that nutrients and other compounds generally show a more interactive relationship where the effects can be either antagonistic or synergistic. An antagonistic relationship indicates that the coupling effect of the nutrients results in a net zero effect, while a synergistic nature indicates an increase in overall impact than if the two types of nutrients were examined separately. Knowing how their composition changes and what their optimum conditions for growth are can help with determining the overall productivity of a system and provide management implications. Ultimately, understanding biofilm composition and whether they are greatly affected by nutrient interactions can determine how the organisms will respond to certain environmental stressors (Franke et al. 2013).    In this experiment, the interactions between dissolved organic carbon (DOC) and other limiting nutrients (nitrogen and phosphorous) were examined under laboratory mesocosm 9  conditions. The objectives were to determine how total biomass and the proportions of bacteria to algae differ in biofilms that form under different nutrient conditions, followed by what the implications are for natural systems where concentrations of these nutrients can fluctuate greatly. The major predictions of this experiment were: (1) The presence of grazers will limit total biofilm biomass unless there are high levels of nutrients, (2) Nutrients are limiting and biofilm cannot take full advantage of increasing DOC levels, (3) Higher levels of DOC and low nutrients at a consistent ratio will yield a larger proportion of bacteria than algae, and (4) Higher nutrient levels compared to DOC will yield a larger proportion of algae to bacteria.   II. MATERIALS AND METHODS  2.1 Experimental Design The experimental setup is a fully crossed factorial design with three factors: concentration of dissolved organic carbon (DOC), concentration of nitrogen and phosphorous (N and P) at a consistent ratio, and larvae of the caddisfly Glossosoma penitum. The first factor for DOC consists of four added-concentration levels: 1 mg/L, 2 mg/L, 3 mg/L, and 4 mg/L. The second factor N:P consists of three levels of added-concentrations: 16:1 mg/L, 32:2 mg/L, and 48:3 mg/L, where the ratios are based on the amount of N and P atoms in the initial compounds and consistent with the Redfield 16:1 ratio found across a majority of natural aquatic systems (Letscher et al. 2015). There are a total of 24 treatments, each consisting of 3 replicates and resulting in a grand total of 72 experimental units. Both the DOC and N:P amounts were added to each of the experimental units in a fully crossed design so that it creates a gradient for each of these two factors.  10  In order to obtain biofilm for this experiment, rocks averaging 4 cm in diameter were retrieved from East Creek at the Malcolm Knapp Research Forest in British Columbia during early January to provide the initial colony of bacteria and algae. The stones were then brought back to the University of British Columbia Vancouver campus where the experiment took place. Seventy-two plastic containers (Richard’s Packaging, 2 kg Clear PET P/B Jar) were filled with tap water to set the environment for biofilm growth. Including stock solution volumes, the total volume in each of the containers was 1.5 L, and each container contained four, unglazed ceramic tiles (approximately 4 cm x 4 cm) and two stream rocks placed at the bottom of the jar. All containers were also fitted with air pumps and air stones to provide adequate circulation and dissolved oxygen.  Grazers retrieved from the same stream were also added to half of the experimental units to obtain the desired treatment type.    A stock solution of 6.63x10-3 M for the DOC concentrations was mixed using humic acid purchased from the UBC Chemistry Stores (Acros Organics humic acid, sodium salt 45-70%; CAS 68131-04-4). In the same way, a stock solution was also prepared for the N:P combination; 0.3871 M of NaNO3 (Caledon sodium nitrate; CAS 7631-99-4) and 0.0242 M of KH2PO4 (Fisher Scientific potassium phosphate monobasic; CAS 7778-77-0). Volumes of these stock solutions were then added to each of the containers using a micropipette in order to get the desired concentrations and volumes for each of the experimental units. After applying each of the treatments, the jars with starting biofilm colonies were left at room temperature (~19°C) under growing light conditions using LED Grow Light with Super Harvest Colors (310 x 310 x 38 mm dimensions). Following the four week growing phase, the ceramic tiles were then removed from the containers, wrapped with aluminum foil, and stored in a freezer to be used in the total biomass and chlorophyll-α analyses. 11   2.2 Biomass  From each of the individual containers, two ceramic tiles were used to measure total biomass of the accumulated biofilm. Samples were first scraped off the tiles into a separate container using a hard-bristled toothbrush and then rinsed with distilled water to form an algal slurry. Using a glass fiber filter with a vacuum below 20 kPa, algal slurry samples were dried on ash-free Whatman GF/F: 47 mm diameter (0.45 μm pore size) filters. Once all the water had been removed, filtered samples were allowed to air-dry for at least 24 hours before being weighed for total mass (organic + inorganic) and measurements are based on the assumption that all ash-free filters were roughly the same weight, averaging 0.128 g. A portion of the samples had also been placed in an oven at 60°C for at least 24 hours to determine the percent difference between air-dried and oven-dried mass for future calculations. After recording all air-dried and oven-dried weights, samples were then placed in a muffle furnace at 500°C for 1.5 hours to burn off any organic matter; the final weights being the remaining inorganic matter.  The resulting differences between the air-dried weights and ash-free dry mass were then calculated to determine the total biomass that grew on the two tiles per experimental unit. The air-dried weights were used instead of the oven-dried weights due to inconsistencies in the methodology since not all samples were placed in the oven. These differences between air-dried and oven-dried weights were quite small (average 0.4%) and were based on calculations with the                                          few samples that do have both air-dried and oven-dried weights and was not significant enough to affect further biomass calculations.   12  2.3 Chlorophyll-α extraction and readings  Similar to the process above, two of the ceramic tiles for each of the samples were scraped and used for further chlorophyll-α analysis. Once filtered through the same glass fiber filter apparatus and on ash-free Whatman 47 mm diameter (0.45 μm pores) filter paper, the filters were folded in half twice before placing in a 15 mL Falcon conical tube. The tubes were steeped in 10 mL of 90% acetone and placed in the refrigerator for at least 24 hours while covered in aluminum foil to prevent further chlorophyll degradation. All tubes were shaken at least once throughout the 24 hour steeping period.  Once the steeping period had been completed, samples were removed from the refrigerator and allowed to warm to room temperature (approx. 30 minutes) before being distributed into round cuvettes. Each cuvette was filled with 7 mL of the steeped solution (75-90% full), wiped with Kimtech Science precision wipes to remove any residual moisture, and then placed in a Turner Designs-700 laboratory fluorometer for the initial reading (Fb). Since calibration of the fluorometer was set to 0-200 μg/L, dilution of the more concentrated samples using 90% acetone was necessary; dilution levels varied between 1-25% of the original extracts. Following the initial reading, 0.1 mL of 0.1 (10%) hydrochloric acid (HCl) was added to the cuvettes and shaken to acidify the sample extract for the second reading (Fa) where living chlorophyll content has been destroyed. All cuvettes were rinsed with 90% acetone before continuing on with the next sample readings.   Calculations for the final chlorophyll-α concentrations followed the equation below:     (    )                    (     )     13  where Fb/Fa max is the acidification coefficient for a sample that only contains pure chlorophyll and has no presence of phaeopigments. The symbol   indicates the extraction volume (10 mL) and    refers to the sample volume placed in the cuvette (7 mL). Samples that have been diluted followed the same calculation process above and final values were simply multiplied by the inverse of the dilution ratio (e.g. multiplied by 4 if diluted to 25% of original concentration). Final values of chlorophyll-α (mg/m3) were converted to simple biomass (mg) and multiplied by the autotrophic index (AI) of 145.0 based on carbon:chl-α ratios listed by the United States Environmental Protection Agency (1985) and converted by Cole and Wells (2003); this allows for direct comparison with total biomass since prior to this conversion, the values only indicated pigment mass.   2.4 Statistical Analyses   Following the collection and measurements for all samples, a t-test assuming unequal variances using Microsoft Excel was first performed on the effects of the presence or absence of grazers to determine whether this factor could be eliminated from further analyses. The SAS (9.3) program was then used for further statistical analyses to determine whether there was a significant interaction between the different factors, mainly DOC and nutrients. The produced least square means and p-values were used for further interpretation of any trends in the data and the significance of each of the main factors.    III. RESULTS  3.1 Grazer Influence  14   The average total biofilm biomass in the presence of grazers is 4.6 mg; 5.2 mg for the absence of grazers. Chlorophyll-α levels also differed slightly between grazers and no grazers, averaged at 141.7 mg/m3 and 134.4 mg/m3, respectively. Since there were very few live caddisflies at the beginning of the experiment, the initial statistical test done using a simple t-test comparing average biomass and chlorophyll-α determined that there is no difference between the two means in both cases. The p-value comparing average biofilm biomass with grazers and no grazers was 0.459, while the p-value comparing average chlorophyll-α is 0.680. Both values are greater than the alpha level 0.05, indicating that grazer effects were negligible. As there were no significant differences between the two factor levels, this treatment was omitted from any further analyses (i.e. n = 6 per treatment, rather than n = 3).    3.2 Overall biomass  According to the SAS statistical analyses, there is no significant interaction between DOC and nutrients on total biomass, as indicated by the p-value 0.098 (> α = 0.05); the two factors were then examined separately. The average biomass for the four different DOC levels (1.0 mg/L, 2.0 mg/L, 3.0 mg/L, and 4.0 mg/L) was found to be 4.70 mg, 4.86 mg, 5.11 mg, and 4.92 mg, respectively. According to the statistical analyses however, this factor does not have a significant effect on biomass (p-value = 0.985; Table 1). While this also illustrates that the different levels of DOC do not differ, there is still a general trend where total biomass increases with an increase in DOC concentration (Figure 1). The level yielding highest average biomass was at 3.0 mg/L of DOC and the small drop at 4.0 mg/L DOC can be attributed to the statistical insignificance of each level, resulting in overlapping error bars.  15   Trends in average biomass based on the levels of nutrients also followed the same general pattern where an increase in nutrients insulted in greater biomass (Figure 2). The three levels 16:1 mg/L, 32:2 mg/L, and 48:3 mg/L of N:P yielded an average biomass of 3.87 mg, 5.00 mg, and 5.85 mg, respectively. Based on the p-value of 0.115, the nutrients also did not appear to have a statistically significant effect on overall biomass.   3.3 Chlorophyll-α  The SAS package reports a p-value of 0.366 (Table 2) on the interaction between DOC and nutrients and its effect on chlorophyll-α content, indicating that there is not a significant interaction. Examining just DOC levels, the average chlorophyll-α amounts are 1180.23 mg/m3, 2386.14 mg/m3, 2405.76 mg/m3, and 2163.46 mg/m3, respectively. The p-value for the DOC factor itself is 0.495, indicating that it did not have a significant impact on chlorophyll-α accumulation. Similar to the effects on biomass, there is a small trend where the increase of nutrients also increases the levels of chlorophyll (Figure 3); highest amounts occur at 3.0 mg/L of DOC and drops at 4.0 mg/L.   Chlorophyll levels according to amount of nutrients in a 16:1 ratio were 1023.72, 2132.92, and 2945.05 mg/m3 (Figure 4). Based on the p-value of 0.058, this factor was also insignificant in affecting the levels of chlorophyll-α.  3.4 Biofilm proportions  As there did not seem to be any trends within the different levels of DOC and within nutrient levels, the ratio of total biomass to algal biomass also remains consistent (Figure 5a, 5b). The average ratio is 2.6 for just DOC, and 2.7 for just nutrients; these two ratios are not 16  significantly different, indicating that the proportion of algae remains constant throughout all treatment types examined.    IV. DISCUSSION  4.1 Effect of Grazers From the start of this experiment, most of the caddisflies added into their respective containers were already dead or stressed from the change in environment, resulting in the negligible impact on the overall biofilm biomass and chl-α content. In most studies, however, there is a significant impact from grazers on the amount of biofilm accumulation under higher nutrient conditions since higher levels of biofilm can better support larger grazer populations (Anderson et al. 1999; Spanhoff et al. 2006). While nutrient availability can be limiting in some cases, other factors such as light may be more important; Martina et al. (2014) examined the effects of grazing on algal communities by comparing grassland and afforested stream sites with similar water chemistry and concluded that the grassland communities with more light exposure had greater overall productivity of biofilm, resulting in significant grazing effects by Helicopsyche sp. caddisflies. Alternatively, at the afforested sites where light was limiting, biofilm growth rates were much slower and had lower biomass over the same period of time, the lower number of grazers that could be supported did not significantly affect biofilm composition. Similarly, a study by Barranguet et al. (2005) compares light limitation and the effects of an aquatic snail, Potamopygrus antipodarum, on biofilm accumulation and reported an overall lower biomass for treatments that experience heavy grazing, even under adequate light conditions. Overall, the effects of grazers are limited by the amount of biofilm that can accumulate based on nutrient conditions, as opposed to the grazers being the limiting factor on 17  biofilm growth. The grazer species’ position on the trophic pyramid will also have an effect on how well they can benefit from high nutrient inputs (Davis et al. 2010).  Assuming general population dynamics, however, it is possible that under highly eutrophic conditions where biofilm communities are abundant, the effects of grazers become negligible and other density-dependent or independent factors become more limiting for biofilm.   While grazers generally do have a significant influence on biofilm growth under high nutrient conditions, the effectiveness of their grazing can also be dependent on the species of grazers and their feeding behaviour (Aguilera et al. 2013). The consumption rates and energy transfer of different taxa, such as molluscs (Barranguet et al. 2005), caddisflies (McNeely et al. 2006; Martina et al. 2014), chironomids (Maasri et al. 2010), protists (Chavez-Dozal et al. 2013), or amphipods (Moghadam and Zimmer 2014) can vary greatly. McNeely et al. (2010) examined the differences in carbon use between heptageniid mayfly nymphs and Glossosoma larvae and found that the caddisflies were more selective in their feeding and obtained most of their dietary requirements from the algal component of biofilm, even under low nutrient levels. Mayfly nymphs were non-selective and could feed on both the heterotrophic bacteria and algal components. In another case, Maasri et al. (2010) examined selective feeding behaviour for a freshwater fly, Eukiefferirlla claripennis, within the algal component. Since smaller grazers have a limited range in consumption, they are more selective in their feeding and these larvae showed a preference for the classes Chlorophyceae and Cyanophyceae, while avoiding Rhodophyceae, determined by their ability to break down these carbon sources. The differences in species feeding behaviour will have implications for energy transfer into higher trophic levels since some species are less likely to be preyed upon (e.g. having protective casings formed by Glossosma species or diurnal drifting).  18  Anti-predation behaviour from the bacterial component of biofilms is also an influential factor on its overall growth and accumulation. The marine protists, Vibrio fischeri, examined by Chavez-Dozal et al. (2013) are also selective feeders that mainly feed on the bacterial component of biofilms. This ultimately alters biofilm bacteria to algae proportions, but also induces the selective pressure for bacteria to resist grazing by secreting antiprotozoan enzymes. However, the type of protection provided by these enzymes is most common in marine environments and there has not been as much research thus far on freshwater anti-predation enzymes.   4.2 Total Biomass  Although the current experiment did not yield significant results between the different levels of DOC and N:P concentrations, many studies have concluded that these factors are particularly important in determining biofilm composition and total biomass. Ruegg et al. (2011) examined the nutrients most limiting to both autotrophic and heterotrophic biofilms by looking at changes in their community composition during an influx of nutrients from salmon carcasses. The authors state that autotrophic dominated biofilms were generally more N-limited and that heterotrophic biofilms were mainly P-limited; for both biofilms, the coupling of N and P are also an important factor. The addition of salmon did increase biofilm respiration by 3.2 times for dominantly heterotrophic biofilms while the overall growth for autotrophs was impeded by the increase in ammonia. This study also concludes that nutrients can limit the algal component of biofilm growth; increases in DOC composition will simply alter the composition rather than impede overall biofilm accumulation (i.e. biomass is relatively the same, but composition differs with DOC increase).  19              Other studies have shown that the light source may be more important than nutrient availability in determining biomass, especially under artificial light settings. This is reasonable due to the uneven distribution of light that could not be altered under the current laboratory circumstances; some experimental units received significantly more light than others. Mentioned previously, the study by Martina et al. (2014) found higher productivity and biomass accumulation at the light-abundant grassland sites compared to afforested areas; in absence of grazers, the average cell density per unit area was roughly 3.5 times greater than in areas with more shading. The increase in biomass is mainly attributed to an increase in the algal component, however, autotrophic productivity also translates to the productivity of their heterotrophic symbionts.           Stream flow rates are less important under mesocosm studies, but are quite important in determining persistence and overall biomass of biofilms in natural systems. Rickard et al. (2004) mainly examined the diversity of biofilms under different shear rates and found that with greater velocities, the bacterial diversity decreases. Bacteria that excrete autoaggregating enzymes tend to fare better under high flow rates, while under intermediate rates, coaggregating bacteria (i.e. bacteria that take advantage of other species’ aggregating properties) were more prevalent. Coundoul et al. (2015) examined friction velocities and shear stress and have also concluded that the higher flow rates are attributed to the slower rates of biofilm biomass accumulation.    Finally, another factor that can have an effect on the total biomass is the experimental time-frame and whether it mimics natural growing conditions (e.g. diurnal variations). While consecutive light duration is less applicable to natural systems, it does provide insight for the discrepancies between the data from this experiment and the conclusions of other experimental settings; the age of the biofilm is important since it takes time for a substantial amount of buildup. 20  Based on the findings from Barranguet et al. (2005), there are significant compositional changes in the biofilm that come with age. For example, the presence of extracellular polymeric substances (EPSs) is important for the integrity of the film, but is less important under slow or non-moving water scenarios. The age of the biofilm also determines the amount of grazing and by what species, which can also factor into the total biomass at different time periods (Chavez-Dozal 2013).   Understanding the factors most important in determining overall biomass can be beneficial for predicting the productivity of a system under different conditions, whether due to season or following disturbance events. Knowing the productivity of biofilm translates to predictions for the productivity of the overall community since they comprise the base of the food web and control ecosystem interactions through a bottom-up influence (Shurin et al. 2012).  4.3 Biofilm Proportions  Similar to the total biomass results, the amount of chlorophyll-α measured for each of the levels of DOC and nutrients were not significantly different from each other, showing that the ratio has remained the same throughout the different treatment levels. Most peer-reviewed literature that examines this concept support the hypothesis that a greater amount of DOC will result in a higher proportion of bacteria compared to algae. Oveido-Vargas et al. (2013) observed the coupling effects of organic carbon and phosphorus under different nitrogen levels and found that simply increasing DOC at low levels of nitrogen will result in a greater proportion of bacteria that outcompete the algae for nitrogen. Ruegg et al. (2011) also alluded to the fact that salmon carcasses also bring a source of organic carbon and that nitrogen and phosphorous influxes tend to yield greater chlorophyll content. Franke et al. (2013) have also found that an 21  increase in organic carbon does not necessarily affect biofilm respiration and that increasing nutrients can actually limit biofilm mineralization of DOC.  As mentioned previously, the light availability is also quite important in determining the algal proportion. While both the heterotrophic and autotrophic components rely on the other for survival, the ongoing production of sugars from the primary producers is also what allows these communities to persist.  In addition to nutrient levels, temperature also significantly affects the stoichiometry of carbon (indicator of total biomass) to chlorophyll. Ventura et al. (2008) found that the ratio of carbon to chlorophyll-α decreased with an increase in temperature, indicating a higher proportion of algae. Initial stoichiometric composition of these biofilm autotrophs and heterotrophs also differ due to their ability to store certain nutrients in different forms (McCall et al. 2014).  As with determining total biomass of biofilm, the age of these micro-communities are also an indicator of the proportional differences of bacteria to algae. In the same study by Barranguet et al. (2005), there are significant differences throughout the biofilm growth cycle. Younger biofilms tend to have a higher proportion of heterotrophic bacteria and are eventually colonized by algae. With age, biofilm thickening reduces light penetration and restricts growth of algae to the epilithic layer. The age of the biofilm also ties into the types of grazers that are present and selective feeders can also alter the biofilm composition.   Ultimately, knowing the biofilm composition can determine how a system will respond to environmental stressors. For example, heterotrophic biofilms are able to use organic forms of N and P if their inorganic counterparts are not as readily available (Franke et al. 2013); therefore, affecting cycling of these nutrients. Understanding the different ratios can greatly contribute to environmental assessment of various streams by indicating the levels of certain types of nutrients 22  and whether other abiotic factors, such as light availability and temperature, will also translate to higher tropic levels.   4.4 Nutrient Interactions  While this experiment did not find any significant interactions between DOC and nutrients, other studies do support the fact that the cycles between carbon, nitrogen, and phosphorus are interconnected in a complex manner. Most of these interactions are due to indirect effects on each other since one compound does not directly alter the chemistry of another; significant indirect impacts are due to alterations in other abiotic or biotic components of the system. Oveido-Vargas et al. (2013) have mentioned that low phosphorous levels can affect the ability of bacteria to take up DOC due to a significant coupling effect, mediated by added amounts of NH4+.  A similar case by Marcarelli et al. (2009) shows that nitrogen is often the limiting factor for autotrophic biofilms at highly disturbed sites, which then affects the ability for the heterotrophic component to take in organic sources of carbon. Ultimately, the study concludes that the two major components of biofilms will have different response rates to nutrient influx, thereby affecting the quantity of another nutrient or carbon source.  Rosemond et al. (2015) also examined an indirect effect of excess nutrients on the amount of organic carbon in a system. The authors found that the increase in algal productivity also increases mineralization of carbon available to higher trophic levels and decreases its residential time in the adjacent terrestrial environment. On the other hand, Rowe et al. (2014) modeled the effects of pH levels and nitrogen to find that the increase in plant productivity can actually increase the level of DOC and change the adjacent soil chemistry. They concluded that nitrogen pollution is interconnected with maintaining dissolved carbon pools and that the 23  persistence of higher DOC levels will remain even after soil conditions stabilize. Since Rowe et al. (2014) were examining stream acidity levels, the discrepancy with Rosemond et al. (2015) may not be directly related.  Either way, both of these findings indicate that there are various pathways in which carbon availability can be manipulated by nutrient levels. On the other hand, there have been fewer studies that examine the effects of DOC on nitrogen or phosphorus availability.  4.5 Future Research  Multiple aspects of this experiment could be altered in future studies in order to extract statistically significant information on the interactions between DOC and nutrients nitrogen and phosphorus. Alterations in experimental procedures could include a more consistent light source across all experimental units and increasing the increments for the different levels of DOC and N:P so that the differences are more drastic, but still within the levels of most natural ecosystems. Meticulous and timely transportation of grazers is also necessary to ensure the lowest level of stress on the organisms so that they can survive the duration of the experiment. A greater number of grazers added into each experimental unit will also likely show a more significant impact as it increases the chances that some individuals will survive to the end of the experiment and have higher rates of grazing. This would also emulate natural conditions where a higher population of grazers can be sustained with higher nutrient concentrations, thereby increasing the grazing impact.   Future studies on nutrient manipulation could also include the differences in nitrogen and phosphorus ratios since the Redfield ratio may not always be present, either due to geographic location or seasonal differences in nitrogen and phosphorus levels. Gusewell and Gessner (2009) 24  examined different N:P ratios on the decomposition rates of two main taxa, bacteria and fungi. The results showed that bacteria were more prevalent under low N:P situations where nitrogen was limiting, whereas fungi were more prevalent when N:P was high and phosphorus-limiting.  While the study did not examine the N:P effects on autotrophic algae, one can assume that N-limited sites will also show a similar trend where there is a greater proportion of bacteria and that algae will more easily take advantage of greater N:P ratios; this could be addressed in further studies that mainly focus on autotrophic biofilms. The nutrient stoichiometry of marine environments can also be further delved into since the biofilm communities are just as important at performing similar ecological functions. Letscher et al. (2015) uses a modeling approach to determine how variations in C:N:P ratios will affect how carbon is being cycled through the system.    In addition to nutrient cycling at various nutrient ratios, the impacts of various pollutants on the nutrient availability and carbon interactions can also provide management implications. Do pollutants decrease the availability of certain compounds through chemical reactions that immobilize nutrients? Then what is the response rate for how these aquatic communities are affected? One example that touches on this issue is a study by Courtney and Clements (2002) that examines how benthic communities react directly to heavy metal pollution, but it does not mention how the nutrients themselves can be affected.   The effects of temperature on the coupling properties of nutrients with DOC can also be addressed since the changes in global climate have been of increasing concern. It has been heavily studied that altered temperature regimes will have drastic effects on the community composition and since temperature is a master variable for most ecosystem components, understanding its effects can contribute to future management decisions. In the context of 25  biofilms, Ylla et al. (2014) show that an increase in temperature can increase biofilm decomposition rates and change the amount of available carbon for other organisms.     V. CONCLUSION  Overall, the experiment did not yield any significant interactions between DOC and the nutrients nitrogen and phosphorous. The grazer effects were negligible and can be attributed to issues with proper transportation methods. The most significant factors determining total biofilm biomass and algal proportions in this study are likely due to other factors that were not being measured for. According to past studies, however, there are significant interactions between DOC and nutrients and understanding their effects on biofilm growth and composition is necessary for determining the ecosystem response to various changes in environmental conditions.   ACKNOWLEDGEMENTS  First of all, I would like to thank my supervisor, John Richardson, for guiding me throughout the entire year, starting from the ideas and experimental design for the project, down to the final steps in data analysis and interpretation. I would also like to thank Alex Yeung, Liliana Lago, and Ana Chara for guiding me through various experimental procedures and the proper operation of specific lab equipment. Finally, I would like to thank Danielle Fontaine and Adrian Chan for assisting me throughout the experimental set-up and running of the experiment.       26  REFERENCES  Aguilera, M.A., Navarrete, S.A., and Broitman, B.R. 2013. Differential effects of grazer species  on periphyton of a temperate rocky shore. Marine Ecology Progress Series 484: 63-78.   Anderson, E.L., Welch, E.B., Jacoby, J.M., Schimek, G.M., and Horner, R.R. 1999. Periphyton  removal related to phosphorous and grazer biomass level. Freshwater Biology 41(3): 633- 651.   Barranguet, C., Veuger, B., Van Beusekom, S.A.M., Marvan, P., Sinke, J.J., and Admiraal, W.  2005. Divergent composition of algal-bacterial biofilms developing under various  external factors. European Journal of Phycology 40: 1-8.   Battin, T.J., Kaplan, L.A., Newbold, J.D., and Hansen, C.M.E. 2003. Contributions of microbial  biofilms to ecosystem processes in stream mesocosms. Nature 426: 439-442.   Chavez-Dozal, A., Gorman, C., Erken, M., Steinberg, P.D., McDougald, D., and Nishiguchi,  M.K. 2013. Predation response of Vibrio fischeri biofilms to bacterivorus protists.  Applied and Environmental Microbiology 79(2): 553-558.    Cole, T.M. and Wells, S.A. 2003. “CE-QUAL-W2: A two-dimensional, laterally averaged,  hydrodynamic and water quality model, version 3.1.,” Instruction Report EL-03-1, U.S.  Army Engineering and Research Development Center, Vicksburg, M.S.  Coundoul, F., Bonometti, T., Graba, M., Sauvage, S., Perez, and Moulin, F.Y. 2015. Role of  local flow conditions in river biofilm colonization and early growth. River Research and  Applications 31(3): 350-367.   Courtney, L.A. and Clements, W.H. 2002. Assessing the influence of water and substratum  quality on benthic macroinvertebrate communities in a metal-polluted stream: an  experimental approach. Freshwater Biology 47: 1766-1778.   Crain, C.M., Kroeker, K., and Halpern, B.S. 2008. Interactive and cumulative effects of multiple  human stressors in marine systems. Ecology Letters 11: 1304-1315.   Davis, J.M., Rosemond, A.D., Eggert, S.L., Cross, W.F., and Wallace, J.B. 2010. Nutrient  enrichment differentially affects body sizes of primary consumers and predators in a  detritus-based stream. Limnology and Oceanography 55(6): 2305-2316.   Franke, D., Bonnell, E.J., and Ziegler, S.E. 2013. Mineralisation of dissolved organic matter by  heterotrophic stream biofilm communities in a large boreal catchment. Freshwater  Biology 58: 2007-2026.   Gusewell, S. and Gessner, M.O. 2009. N:P ratios influence litter decomposition and colonization  by fungi and bacteria in microcosms. Functional Ecology 23: 211-219.   27  Haggerty, R., Ribot, M., Singer, G.A., Marti, E., Argerich, A., Agell, G., and Battin, T.J. 2014.  Ecosystem respiration increases with biofilm growth and bed forms: flume measurements  with resazurin. Journal of Geophysical Research: Biogeosciences 119: 2220-2230.   Holomuzki, J.R., Feminella, J.W., and Power, M.E. 2010. Benthic interactions in freshwater  benthic habitats. Journal of the North American Benthological Society 29(1): 220-244.   Kohler, J., Hachol, J., and Hilt, S. 2010. Regulation of submersed macrophyte biomass in a  temperate lowland river: interactions between shading by bank vegetation, epiphyton and  water turbidity. Aquatic Botany 92: 129-136.   Letscher, R.T., Moore, J.K., Teng, Y.C., and Primeau, F. 2015. Variable C:N:P stoichiometry of  dissolved organic matter cycling in the Community Earth System Model. Biogeosciences  12: 209-221.   Lovatt, C., Kominoski, J.S., Sakamaki, T., Macleod, B., and Richardson, J.S. 2014. Leaf-litter  leachate and light interactively enhance accrual of stream biofilms. Fundamental  Applications of Limnology 184(4): 297-306.   Marcarelli, A.M., Bechtold, H.A., Rugenski, A.T., Inouye, R.S. 2009. Nutrient limitation of  biofilm biomass and metabolism in the Upper Snake River basin, southeast Idaho, USA.  Hydrobiologia 620: 63-76.   Martina, L.C., Marquez, J., Principe, R., Gari, N., and Albarino, R. 2014. Does grazing change  algal communities from grassland and pine afforested streams?: A laboratory approach.  Limnologica 49: 26-32.   Maasri, A., Fayolle, S., and Franquet, E. 2010. Algal foraging by a rheophilic chironomid  (Eukiefferiella claripennis Lundbeck) extensively encountered in high nutrient enriched  streams. Fundamental Applications of Limnology 177(2): 151-159.   McCall, S.J., Bowes, M.J., Warnaars, T.A., Hale, M.S., Smith, J.T., Warwick, A., and Barett, C.  2014. Phosphorus enrichment of the oligotrophic River Rede (Northumberland, U.K.) has no effect on periphyton growth rate. Inland Waters 4(2): 121-132.   McNeely, C., Clinton, S.M., Erbe, J.M. 2006. Landscape variation in C sources of scraping  primary consumers in streams. Journal of the North American Benthological Society  25(4): 787-799.   Moghadam, F.S. and Zimmer, M. 2014. Effects of warming and nutrient enrichment on how  grazing pressure affects leaf litter-colonizing bacteria. Journal of Environmental Quality  43(3): 851-858.   Oveido-Vargas, D., Royer, T.V., and Johnson, L.T. 2013. Dissolved organic carbon  manipulation reveals coupled cycling of carbon, nitrogen, and phosphorus in a nitrogen- rich stream. Limnological Oceanography 58(4): 1196-1206.  28   Rickard, A.H., McBain, A.J., Stead, A.T., and Gilbert, P. 2004. Shear rate moderates community  diversity in freshwater biofilms. American Society for Microbiology 70(12): 7426-7435.   Rosemond, A.D., Benstead, J.P., Bumpers, P.M., Gulis, V., Kominoski, J.S., Manning, D.W.P.,  Suberkropp, K., and Wallace, B. 2015. Experimental nutrient additions accelerate terrestrial carbon loss from stream ecosystems. Freshwater Ecology 347(6226): 1142-1145.   Rowe, E.C., Tipping, E., Posch, M., Oulehle, F., Cooper, D.M., Jones, T.G., Burden, A., Hall, J.,  and Evans, C.D. 2014. Predicting nitrogen and acidity effects on long-term dynamics of  dissolved organic matter. Environmental Pollution 184: 271-282.   Ruegg, J., Tiegs, S.D., Chaloner, D.T., Levi, P.S., Tank, J.L., and Lamberti, G.A. 2011. Salmon  subsidies alleviate nutrient limitation of benthic biofilms in southeast Alaska streams.  Canadian Journal of Fisheries and Aquatic Sciences 68(2): 277-287.   Sekar, R., Venugopalan, V.P., Nandakumar, K., Nair, K.V.K., and Rao, V.N.R. 2004. Early  stages of biofilm succession in a lentic freshwater environment. Hydrobiologia 512(1-3):  97-108.   Shurin, J.B., Clasen, J.L., Greig, H.S., Kratina, P., and Thompson, P.L. 2012. Warming shifts  top-down and bottom-up control of pond food web structures and function. Philosophical  Transactions of the Royal Society Biological Sciences 367(1605): 3008-3017.   Spanhoff, B., Reuter, C., and Meyer, E. 2006. Epixylic biofilm and invertebrate colonization on  submerged pine branches in a regulated lowland stream. Archiv Fur Hydrobiologie  165(4): 515-536.   United States Environmental Protection Agency. (1985). Rates, constants, and kinetics  formulations in surface water quality modeling (Second Edition) (EPA Publication No.  68-03-3131 Athens, Georgia: U.S. Environmental Protection Agency.   Ylla, I., Canhoto, C., and Romani A.M. 2014. Effects of warming on stream biofilm organic  matter use capabilities. Microbial Ecology 68: 132-145.             29  FIGURES     Figure 1. Average total ash-free dry mass (mg) given the concentration of dissolved organic carbon (DOC). Individual standard error bars calculated according to n=18.    Figure 2. Average total ash-free dry mass (mg) given the concentration of nutrients nitrogen and phosphorous (mg/L) at a consistent 16:1 Redfield ratio (N:P). Individual standard error bars calculated according to n=24.    012345671.0 2.0 3.0 4.0Average Dry Mass (mg) Concentration of DOC (mg/L) 0123456716:1 32:2 48:3Average Dry Mass (mg) Concentration of Nutrients N:P (mg/L) 30   Figure 3. Average chlorophyll-α (mg/m3) given the concentration of dissolved organic carbon (DOC). Individual standard error bars calculated according to n=18.    Figure 4. Average chlorophyll-α (mg/m3) given the concentration of nutrients nitrogen and phosphorous (mg/L) at a consistent 16:1 Redfield ratio (N:P). Individual standard error bars calculated according to n=24.    0.00500.001000.001500.002000.002500.003000.003500.001.0 2.0 3.0 4.0Average Chlorophyll-a (mg/m3) Concentration of DOC (mg/L) 0.00500.001000.001500.002000.002500.003000.003500.004000.0016:1 32:2 48:3Average Chlorophyll-a (mg/m3) Concentration of Nutrients N:P (mg/L) 31    Figure 5. Proportions of biomass compared to chlorophyll-α derived algal biomass depending according to differing concentration levels for DOC and nutrients nitrogen and phosphorous (mg/L). Individual standard error bars calculated according to n=18 for DOC and n=24 for nutrients.    TABLES  Table 1. Type 3 test of fixed effects for total biomass (g) of biofilm, as calculated with the SAS program.P-values indicating the significance of each of the factors.   Effect DF F-value p-value DOC 3 0.05 0.985 Nutrients 2 2.24 0.115 DOC*Nutr 6 1.93 0.098   Table 2. Type 3 test of fixed effects for total chlorophyll-α of biofilm, as calculated with the SAS program. P-values indicating the significance of each of the factors.   Effect DF F-value p-value DOC 3 0.81 0.495 Nutrients 2 2.98 0.058 DOC*Nutr 6 1.11 0.366  00.050.10.150.20.251.0 2.0 3.0 4.0Biomass Per Area (mg/cm2) Concentration of DOC (mg/L) Total BiomassAlgal Biomass00.050.10.150.20.250.316:1 32:2 48:3Biomass Per Area (mg/cm2) Concentration of Nutrients N:P (mg/L) Total BiomassAlgal Biomass       

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