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Factors Affecting Cyanobacterial Ecology in a Restoration Wetland on Poplar Island, Chesapeake Bay Austin Boardman, REU Fellow Maryland Sea Grant Dr. Judy O’Neil, Research Assistant Professor Horn Point Laboratory, University of Maryland Center for Environmental Science Dr. Kevin Sellner, Research Associate Smithsonian Environmental Research Center Abstract A wetland habitat at the Poplar Island ecosystem restoration project (Cell 6) has recently become home to a variety of birds, which use the pond for its nearby food and protection from predators. Additionally, a variety of cyanobacteria inhabit the site, including Anabaena, Oscillatoria, and Microcystis. Toxins produced by some of these cyanobacteria resulted in bird fatalities after a Microcystis bloom in summer 2012. Here, we examine the role that temperature, nutrients, and species composition plays in predicting how blooms might occur, whether toxins are produced, and how nitrogen fixation is performed. We found that cultures of cyanobacteria (primarily Anabaena) grow well in temperatures as high as 30°C, and that Anabaena from Cell 6 produces low levels of microcystin. Nitrogen fixation levels in cultures tended to spike periodically, possibly in response to fluctuations in limiting nutrients. Finally, nutrient concentrations played an important role in cyanobacterial growth and nitrogen fixation. Keywords: Microcystis, Anabaena, microcystin, nitrogen fixation, eutrophication, cyanobacteria, Poplar Island Introduction Poplar Island Poplar Island, near Talbot County, MD, has long served as a shelter for birds, turtles, and a host of other organisms. Unfortunately, sea level rise in the 1900s caused much of the island to erode, reducing its area from 440ha to just 2ha (Erwin et al. 2007). Recently, a solution to this problem was found with the need to dispose of sediment removed from shipping channels in the Chesapeake Bay. Beginning in 1998, the U.S. Army Corps of Engineers began transporting non-contaminated sediment from harbors near Baltimore to the former site of Poplar Island and used the dredge material to form wetland and upland habitats tailored specifically to certain species of wildlife (Burton (n.d.)). One major wetland habitat on the island, Cell 6 (Figure 1), is currently the subject of an investigation led by the Maryland Environmental Service (MES) and the Smithsonian Environmental Research Center (SERC) after a bloom of the cyanobacterium Microcystis in the summer of 2012. Toxins (microcystins) produced by this cyanobacterium were associated with bird mortalities (Maryland Environmental Service 2013)—a major roadblock in the central

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mission of the restoration project. Further, Cell 6 is also highly nutrient-rich, especially in nitrogen and phosphorus (MES, unpublished data) and varies greatly in salinity, with areas ranging from 9.1-14.1. These conditions make it favorable for a variety of species of cyanobacteria, many of which can produce harmful toxins (Johnston and Jacoby 2003; Smith 1983). Microcystis Microcystis aeruginosa is one cyanobacterium that is known to produce harmful toxins, namely microcystins. These hepatotoxins are known to have detrimental effects on mammals, but also have been associated with deaths of birds, turtles, and other vertebrates that live in or near aquatic habitats (Chen et al. 2009). As such, researchers have been closely monitoring aquatic habitats at Poplar Island to control blooms that may arise. M. aeruginosa tends to form harmful blooms in the late summer and thrives at higher temperatures, typically above 25°C (O'Neil et al. 2012; Robarts and Zohary 1987; Thomas and Walsby 1986). For the remainder of the year, when conditions are not favorable, Microcystis will fall to the sediment and remain dormant until conditions improve (Welch and Barbiero 1992). In some cases, it will outcompete other cyanobacteria in brackish water (O'Neil et al. 2012). Previous research has estimated the salt tolerance of Microcystis to be between 7-14, with the ability to withstand salt-shocks of up to 17 (Kotut and Krienitz 2011; Tonk 2007). Habitat nutrient composition also plays a major role in determining the onset of a Microcystis bloom. Blooms are often associated with high levels of nitrogen (Jacoby et al. 2000), as well as high total phosphorus and nitrogen to phosphorus ratio (Johnston and Jacoby 2003). Highly eutrophic habitats are also associated with increased toxin production (O'Neil et al. 2012). Considering all of these conditions, Cell 6 seems to be an optimal habitat for a toxic Microcystis bloom in the late summer when the water reaches a suitable temperature. Cyanobacterial Ecology / Nitrogen Fixation A variety of factors influence the algal composition of a body of water, including temperature, salinity, and nutrient availability. Macronutrients such as nitrogen and phosphorus are of particular interest to those studying algal ecology. Specifically, nitrogen fixing cyanobacteria (diazotrophs) are likely to outcompete non-nitrogen fixers when nitrogen levels are low (Horne and Commins 1987), phosphorus is high (Trimbee and Prepas 1987), and the ratio of nitrogen to phosphorus is low (Smith 1983). This is because diazotrophs are often poor competitors for phosphorus, meaning that if other cyanobacteria have an ample supply of dissolved inorganic nitrogen (DIN), they will outlive the nitrogen fixers that cannot survive in low-phosphorus environments. Likewise, nitrogen-fixing cyanobacteria will be successful in low-DIN environments because of their ability to convert atmospheric N2 into useable NH3 (Moisander et al. 2012; Whitton and Potts 2000) Nitrogen fixation has been observed in many genera of cyanobacteria, including Anabaena (Figure 2) and Oscillatoria (Figure 3)—two groups that have recently been found in Cell 6 (Morgan State University Estuarine Research Center, unpublished data). The presence of these diazotrophs in Cell 6 is curious, considering the high levels of DIN and relatively high salinity. Our research seeks to answer the many questions surrounding the cyanobacteria populations at Poplar Island. For example, the conditions (temperature, salinity, etc.) that may lead to additional Microcystis blooms in Cell 6 are unclear. It is also unknown how these factors can

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influence toxin production. Additionally, it is unknown what role Anabaena and Oscillatoria may be playing, especially with regard to toxin production and nitrogen fixation. This project seeks to investigate the reasons these diazotrophs are so abundant in a eutrophic environment and identify the factors that may affect rates of nitrogen fixation, namely availability of nutrients. For this project, we hypothesize that:

• Microcystis will form blooms in late July/early August 2013. • Microcystis will grow best at temperatures 28-30°C. • Nitrogen fixation in Anabaena will increase as DIN decreases. • Anabaena, in addition to Microcystis, will produce toxins.

Materials and Methods Phase I: Bloom Monitoring & Temperature Assay Seven water samples were collected from the NWC, SEC, SW16, and SWP sites (aka Sites 1, 3, 4, and 5) at Poplar Island’s Cell 6 (Figure 4) on May 29, 2013. At each site, measurements for temperature, salinity, conductivity, DO, and percent saturation were taken using an appropriately equipped YSI Pro 2030. pH was taken using a General Tools PH-501 pH meter. Water was also tested for phycocyanin and chlorophyll a (chl a) fluorescence using a Turner Designs Aquafluor fluorometer. Samples were then processed as follows:

• Samples 1 and 2 from each site were filtered through 64µm mesh to remove zooplankton and through a GF/F filter to remove phytoplankton. 100ml of the filtered water from each site was placed in autoclaved flasks. BG-11, a media commonly used for culturing cyanobacteria (Anderson 2005), was prepared at the salinities measured in the field and 100mL was added to each flask. These served as controls and were placed in the environmental chamber.

• Samples 4, 5, and 6 from each site were filtered through 64µm mesh and 100mL distributed to flasks containing 100mL BG-11, prepared as above, and stored in the environmental chamber.

• 50mL of Sample 7 from each site was decanted to a 50mL disposable capped tube, fixed with Lugol’s iodine solution, stored on ice, and transferred to Morgan State University for species ID and enumeration.

• Sample 8 from each site was collected in 250mL amber bottles and stored at -20°C for microcystin assay.

Three surficial sediment samples were also collected at each site using 60mL polypropylene syringes. Syringes were placed directly into sediment to a depth of at least 5cm, and capped with a rubber stopper, then stored on ice. Upon returning to the lab, sediment samples deeper than 5cm were extruded from syringes and discarded. The remaining sediment was deposited into 250mL flasks. 100mL of GF/F filtered water and 100mL of BG-11 media at the proper salinity was added to each flask, and the contents swirled. Samples were placed in the environmental chamber. Light in the chamber was set at a level appropriate for cyanobacterial growth (~100µE m-2 s-1), on a 14:10 L:D cycle. Initial incubation temperature was 22°C, and was increased by 2°C every 3 days, to a final temperature of 30°C, where they were held for over 10 days. Chl a and phycocyanin fluorescence in each sample was measured every three days to monitor for blooming cyanobacteria (presumably Microcystis). Previous research suggests that a chl

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a/phycocyanin ratio of <10 can be used as an indicator of Microcystis dominance in field samples. Phase II: Toxin Analysis Materials for the phosphatase inhibition assay were obtained from New England Biolabs, Inc. (Ipswich, MA). For this assay, 80µL NEB-BSA buffer was added to all wells in a 96-well plate. Samples for a standard curve were generated using a 5ng/mL microcystin standard, diluted in NEB-BSA buffer to levels of 2.5, 1.25, 0.63, 0.31, and 0.16 ng/mL. 40µL of each standard was added to the plate, except for a 0ng/mL standard and a blank, to which 40µL and 80µL of additional substrate were added, respectively. Standards were made in triplicate. Approximately 50mL from each culture was subsampled and placed in glass vials at the beginning and end of the temperature assay for microcystin analysis. Samples were stored at -20°C, and later thawed to release toxins. Samples were then centrifuged for 1 minute at 5000rpm to separate any cell material from the supernatant. 40µL of each sample was added to the plate. Samples at 25% dilution were also made and added to the plate to increase the sensitivity of the test in case microcystin levels were exceptionally high. All samples were made in triplicate. Using a multichannel pipette, 40µL PP1 enzyme was added to each well, followed immediately by 40uL pNPP substrate. Samples were mixed by gently pipetting up and down several times. Any bubbles that formed were popped using a toothpick. The plate was left to rest for 60 minutes, and then placed in a spectrophotometer and absorbance was read at 405 nm. A standard curve was produced in Excel and sample values were compared against standards. Phase III: Nitrogen Fixation Tests Cyanobacterial cultures from the temperature assay were periodically subsampled and tested for nitrogenase activity through Acetylene Reduction Assay (ARA) (Hardy et al. 1973). For this process, 4mL from each culture was transferred to 10mL gas chromatography vials and sealed with rubber caps. 0.6µL acetylene gas (generated from calcium carbide) was then injected into each vial. Nitrogenase activity was measured in the headspace of each vial by detecting the ethylene content of a 100µL gas sample in a gas chromatograph using ARA (Capone 1993). This process takes advantage of nitrogenase’s ability to preferentially fix acetylene—rather than atmospheric nitrogen—and convert it to ethylene. Readings were taken at time intervals of approximately 4 hours, for up to 24 hours. GC peaks for samples were compared against a 100ppm standard and converted to nmol ethylene using the following equation:

!"#$%&  !"#$!"#$%#&%  !"#$

×(!"#$%&#&'(  !"##$!%&"')×(!"#$%#&%  !"#!$#%&'%("#)×(!"#  !ℎ!"#  !"#$%&)

In order to standardize ethylene volumes per cell, all samples identified and enumerated with light microscopy using a Sedgwick-Rafter chamber at 200x. The number of filaments, cells, and heterocysts was counted for each species and squares were counted until either 50 units of the most common taxa were observed or 100 squares had been examined. In some cases, highly concentrated sample were diluted at a 1:10 ratio with distilled water. The number of cells per mL was calculated as follows:

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!"##$  !"#$%&' ×!"#$%  !"#$%&  (!"  !"#$%&!)!"!#!$%  !"#$%&  (!"  !"#$%&!)

 ×!"!#$  !"#$%&  !"  !"  !ℎ!"#$%  (1000)!"#$%&  !"  !"#$%&!  !"#$%&$'

Once samples were counted, the nmol ethylene produced was divided by the number of cells or by the number of heterocysts to find the average nitrogenase activity per cell. The instantaneous rate of nitrogen fixation was then calculated by dividing this value by the number of hours from the previous reading. Phase IV: Nutrient Bioassays To examine what effect nutrients such as nitrogen and phosphorus have on cyanobacterial growth, Oscillatoria was grown in three different nutrient treatments. First, 8.3mL Oscillatoria culture from the temperature assay was placed in flasks containing 41.6mL BG-11 media to achieve a 1:5 dilution. Three flasks were set-aside as controls. The remaining flasks had nutrients added as follows:

• Flasks 4, 5, and 6: +1mM NH4 • Flasks 7, 8, and 9: +50µM PO4 • Flasks 10, 11, and 12: +1mM NH4 and +50µM PO4

These high concentrations of nutrients reflected the levels measured at Cell 6 in May 2013. (Though nutrient data for June and July were collected weekly by the Smithsonian Environmental Research Center, data were not available at the time of this report’s publication). Samples were incubated at a temperature of 27°C for 7 days, under the same light conditions described for the temperature assay. Readings for pigment fluorescence were taken at 0, 4, and 7 days. A second nutrient bioassay was also performed using water samples collected from the Sassafras River on August 1, 2013, when Anabaena spiroides was known to be present (Maryland DNR 2013) 100mL of river water was dispensed into flasks and nutrients were added as follows:

• Flask 1: Control. No nutrient addition. • Flask 2: +20µM NH4 • Flask 3: +2µM NH4 • Flask 4: +2µM PO4 • Flask 5: +1µM PO4 • Flask 6: +20µM NH4 and +2µM PO4 • Flask 7: +20µM NH4 and +1µM PO4

These nutrient levels reflected the historical range of nutrients that may be found in the Sassafras River. Cultures were incubated in the same conditions as above for 4 days. Pigment fluorescence and nitrogenase activity were measured for each. Results Phase I: Bloom Monitoring & Temperature Assay As shown in Table 1, the measurements taken May 29, 2013 for temperature, salinity, and dissolved oxygen varied greatly between sites, despite the fact that all were within the same body of water. Pigment fluorescence also varied greatly, with the lowest phycocyanin at Site 1. Pigment fluorescence seemed to loosely correlate with the dissolved oxygen levels. Taxonomy

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results compliment the fluorescence data, showing Sites 3, 4, and 5 to be much higher in cyanobacteria than Site 1. In the temperature assay, cyanobacterial growth seemed to increase as temperature increased. As shown in Figures 5-6, chl a fluorescence tended to decrease in higher temperatures, while phycocyanin fluorescence increased. Thus, the chl a:phycocyanin ratio decreased over time, falling below the threshold of 10 in some cases. Since a low chl a:phycocyanin ratio is often indicative of a cyanobacterial bloom, it can be concluded that generally cyanobacteria exhibited stronger growth at higher temperatures. Phycocyanin fluorescence of sediment samples was consistently low throughout the temperature assay. Though a scum (possibly Oscillatoria) did develop in many of the cultures, no additional experiments were performed on these. Monitoring done by MES officials found that Microcystis was not identified at Cell 6 during the course of this study. Phase II: Toxin Analysis Microcystin was found at comparable concentrations in the four cultures that were tested, with levels ranging from 0.366 to 0.375 ng/L (Figure 7). At sites 3, 4, and 5, microcystin concentration was less at the end of the temperature assay than at the beginning, while the opposite was true at Site 1. Phase III: Nitrogen Fixation Tests Nitrogen fixation was measured in all cultures on 6/17/13 and 6/24/13 and normalized to the number of Anabaena heterocysts counted for each vial. (Though some other diazotrophic cyanobacteria were present in the samples, their concentrations were negligible compared to Anabaena, as seen in Figure 9). Nitrogen fixation rates were strikingly different between the two observation dates, with Sites 3 and 4 having the highest levels of nitrogenase activity on 6/17/13, and Site 5 having the highest levels on 6/24/13 (Figures 9-10). As shown in Figure 11, nitrogenase activity in general seemed to decline as temperatures increased, though it is unclear whether this was due directly to temperature; the progression of time and shifts in the species assemblage of the cultures may have confounded the results. Additionally, regressions of nitrogenase activity vs. salinity show that nitrogen fixation tended to be higher in lower-salinity cultures (Figure 12). Phase IV: Nutrient Bioassays In the Oscillatoria nutrient bioassays, there was no significant difference between the control and P treatments in terms of their chl a:phycocyanin ratio (Figure 13). The N and N+P treatments, however, were significantly different than both the control and P treatments (p=.042), with higher ratios, indicating less cyanobacterial growth. N and N+P were not significantly different from each other in terms of their chl a:phycocyanin ratios, but their chl a fluorescence levels were significantly different. Sassafras nutrient bioassays did not exhibit such clear differences, with data obscured by the high variation in gas chromatograph readings (Figure 14). No significant difference was found between any of the treatments, except for the distilled water and filtered water controls. A diel

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cycle, however, was found in the rates of N-fixation, with noticeably higher rates when cultures were exposed to light (Figure 15). Discussion The results of this research show that cyanobacterial ecology at Poplar Island is complex, affected by numerous factors including temperature, salinity, nutrients, and competing organisms. This was apparent from the beginning of the project when high variation in water quality and species composition was discovered within the same ecosystem. This could be due to a number of factors. One possibility is that wind conditions pushed filamentous cyanobacteria to the southern end of the pond, leaving fewer species in the northern side near Site 1. This could be coupled with variations in salinity and nutrients between different areas of the pond, which may have resulted from inconsistent dredging techniques and rainfall runoff affecting north and south differently. These nutrient and salinity conditions likely also affected the performance of algal competitors, which may have worked to suppress cyanobacteria in certain areas. The temperature assay results were unsurprising, considering it is widely accepted in the literature that cyanobacteria tend to perform well at high temperatures (O'Neil et al. 2012; Robarts and Zohary 1987). The absence of blooms from the sediment cultures, however, was not expected. It was hypothesized that dormant cells would rise from the sediment at higher temperatures. Blooms were not observed, possibly because of dredging that was performed in the months prior to sampling. The new sediment may not have contained any cyanobacteria and may have completely covered any dormant cells from summer 2012. The winter dredging may have also been a reason for the absence of any Microcystis blooms at Cell 6. Additionally, MES attempted to suppress Microcystis growth by placing bales of barley straw in the water before the summer bloom season began. Barley straw releases phenolic compounds as it decomposes, which are known to be toxic to Microcystis (Ball et al. 2001; Pillinger et al. 1994). This can be enhanced by fungal decomposition at higher temperatures (K. Sellner, personal communication). Since Microcystis was never observed at Poplar Island or in cultures over the course of this experiment, it is likely that Anabaena was the source of the microcystin that was measured in the toxin analysis. It should be noted, however, that the levels of microcystin observed in the lab cultures was several orders of magnitude lower than that observed at Cell 6 in the summer of 2012. The highest level measured from the cultures was 0.375 ng/L, which is well below the WHO provisional guideline value of 0.001 mg/L (World Health Organization 1998). Anabaena is also assumed to be the source of the high nitrogen fixation levels that were observed throughout this experiment, though Oscillatoria and Pseudanabaena are likely to have also contributed very small amounts. One obvious peculiarity of the ARA experiments was the high levels of nitrogenase activity at Sites 3 and 4 on 6/17/13 and the high levels at Site 5 on 6/24/13. A possible reason for this is that each site had a different concentration of Anabaena cells initially, meaning that each culture used the media’s nutrients differently over time. Although all cultures were supplied the same amount of media at intervals throughout the experiment, their consumption of nitrogen, phosphorus, and other nutrients was likely quite varied. Since nitrogen fixation is a costly process in terms of energy, it should follow that diazotrophs would benefit from consuming as much DIN as possible, before activating nitrogenase. It might be assumed that Sites 3 and 4 had reached that point on 6/17/13, while Site 5 had sufficient nutrients and did not need to begin fixing nitrogen at higher levels until

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6/24/13. Alternatively, these spikes could be due to unmeasured variables, such as bioavailable Fe, which plays an important role in nitrogen fixation. The trend of nitrogenase activity declining as temperature increased could be interpreted a number of ways. While it might appear that temperature was directly correlated with nitrogenase activity, it is more likely that a confounding variable such as addition of media or changes in the community assemblage is to blame. For example, it is possible that the first acetylene reduction test was performed at a time when the cyanobacteria had consumed most of their nutrients, while later tests may have been done after nutrients had been replenished. This would explain differences in nitrogenase activity at different temperatures, simply because elapsed incubation time coincided with incubation temperature. Alternatively, shifts in the cyanobacterial species composition of the cultures could explain the decrease in nitrogenase activitiy. Unpublished taxonomy data from A. M. Hartsig shows that over the course of the temperature experiment, concentrations of diazotrophs such as Anabaena declined, while non-diazotrophic cyanobacteria including Spirulina and Planktolyngbya limnetica were among the most common at the end of the experiment. It is unclear, however, whether this shift was due to temperature or some other factor. Nitrogenase activity also seemed somewhat diminished at higher salinities, independent of time or temperature. This trend reflects previous research showing that cyanobacteria often exhibit reduced nitrogen fixation in higher-salinity environments (Severin et al. 2012; Zhang and Feng 2008). This can either be due to increased stress on the organisms, which reduces their capacity to expend energy on nitrogenase production, or due to a shift in the community assemblage towards organisms that are more halotolerant non-diazotrophic organisms. In this case, the latter was unlikely, as the species composition between cultures of different salinities was seemingly random and extremely varied. The strikingly high levels of nitrogen used in the Oscillatoria nutrient bioassay leave little doubt that 1mM NH4 has the capacity to overwhelm cyanobacteria to the point of inhibiting growth. It is hypothesized that these high nutrient levels also caused a decrease in nitrogen fixation, though this was not tested due to equipment failure. Finally, we hypothesized that nutrient concentrations would affect nitrogen fixation levels in Sassafras River samples. Results from this experiment were not entirely clear; it appears the control treatment may have had the highest levels of nitrogen fixation, especially toward the end of the experiment, though the high variability in the data make this difficult to say conclusively. If the experiment were repeated, results might be improved by the using a more sensitive method to detect N-fixation. Also, the incubation period may have been too short to have a significant impact on nitrogen fixation. Regardless, it is apparent that even changes in nutrients as great as 10x the ambient concentration will not produce changes in nitrogen fixation that are easily measurable. Diel variations in nitrogenase activity were consistent with those expected for heterocystous cyanobacteria, where nitrogenase activity was higher in the presence of light. Previous studies have suggested that this may be due to the fact that the products of nitrogen fixation are used in photosynthesis, which only occurs during daylight hours (Capone et al. 1990). Conclusions Though one goal of this research project was to gain a better understanding of the factors that contribute to Microcystis blooms, the organism was never observed, possibly due to the addition

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of barley straw to Cell 6 waters. While a major victory for the environment, this means that our two hypotheses concerning the timing and climate of a Microcystis bloom could not be confirmed nor rejected. It was found, however, that other cyanobacteria such as Anabaena do grow well in high temperatures. Additionally, Anabaena was found to produce toxins, confirming our hypothesis. However, the levels of microcystin produced were far below WHO guidelines. Nitrogen fixation was observed in cultures from Cell 6, probably chiefly due to Anabaena. Nitrogen fixation in our cultures tended to spike suddenly at times, possibly in response to limited nutrients. However, nutrient bioassays were not able to clearly define what role nutrients play in nitrogenase activity. Thus, our hypothesis regarding nitrogen fixation in Anabaena remains unconfirmed. Acknowledgements I would like to express my gratitude to Dr. Judy O’Neil for guiding me through every step of this project, Anne Gustafson for providing assistance in the field and in the lab, Dr. Eric Schott and Kevin Meyer for assistance with toxicity analysis, and Dana Bunnell-Young for equipment troubleshooting. I also thank Mike Allen, Fredrika Moser, the staff of Maryland Sea Grant, and the National Science Foundation for making this research experience possible.

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References

Anderson, R. A. 2005. Algal Culturing Techniques. Elsevier Academic Press. Ball, A. S., M. Williams, D. Vincent, and J. Robinson. 2001. Algal growth control by a barley

straw extract. Bioresource Technology 77: 177-181. Burton, K. (n.d.). The island that almost vanished is slowly reappearing. U.S. Fish & Wildlife

Service Chesapeake Bay Field Office. Capone, D. G. 1993. Determination of nitrogenase activity in aquatic samples using the

acetylene reduction procedure, p. 621-631. Handbook of Methods in Aquatic Microbial Ecology. Lewis Publishers.

Capone, D. G., J. M. O'Neil, J. Zehr, and E. J. Carpenter. 1990. Basis for diel variation in nitrogenase activity in the marine planktonic cyanobacterium Trichodesmium thiebautii. Applied and environmental microbiology 56: 3532-3536.

Chen, J., D. Zhang, P. Xie, Q. Wang, and Z. Ma. 2009. Simultaneous determination of microcystin contaminations in various vertebrates (fish, turtle, duck and water bird) from a large eutrophic Chinese lake, Lake Taihu, with toxic Microcystis blooms. Science of the Total Environment 407: 3317-3322.

Erwin, R. M., J. Miller, and J. G. Reese. 2007. Poplar Island Environmental Restoration Project: Challenges in Waterbird Restoration on an Island in Chesapeake Bay. Ecological Restoration 25: 256-262.

Hardy, R. W. F., R. C. Burns, and R. D. Holsten. 1973. Applications of the acetylene-ethylene assay for measurement of nitrogen fixation. Soil Biology and Biochemistry 5: 47-81.

Horne, A. J., and M. L. Commins. 1987. Macronutrient controls on nitrogen fixation in planktonic cyanobacterial populations, p. 413-423.

Jacoby, J. M., D. C. Collier, E. B. Welch, F. J. Hardy, and M. Crayton. 2000. Environmental factors associated with a toxic bloom of Microcystis aeruginosa. Canadian Journal of Fisheries and Aquatic Sciences 57: 231-240.

Johnston, B. R., and J. M. Jacoby. 2003. Cyanobacterial toxicity and migration in a mesotrophic lake in western Washington, USA. Hydrobiologia 495: 79-91.

Kotut, K., and L. Krienitz. 2011. Does the potentially toxic cyanobacterium Microcystis exist in the soda lakes of East Africa? Hydrobiologia 664: 219-225.

Maryland DNR. 2013. Continuous Monitoring: Sassafras River - Budd's Landing. Maryland Environmental Service. 2013. Subcontractors Scope of Work: Environmental

Restoration Project Task 50. Maryland Environmental Service. Moisander, P. H. and others 2012. Facultative diazotrophy increases Cylindrospermopsis

raciborskii competitiveness under fluctuating nitrogen availability. FEMS Microbiology Ecology 79: 800-811.

O'Neil, J. M., T. W. Davis, M. A. Burford, and C. J. Gobler. 2012. The rise of harmful cyanobacteria blooms: The potential roles of eutrophication and climate change. Harmful Algae 14: 313-334.

Pillinger, J., J. Cooper, and I. Ridge. 1994. Role of phenolic compounds in the antialgal activity of barley straw. Journal of Chemical Ecology 20: 1557-1569.

Robarts, R. D., and T. Zohary. 1987. Temperature effects on photosynthetic capacity, respiration, and growth rates of bloom-forming cyanobacteria. New Zealand Journal of Marine and Freshwater Research 21: 391-399.

Severin, I., V. Confurius-Guns, and L. Stal. 2012. Effect of salinity on nitrogenase activity and composition of the active diazotrophic community in intertidal microbial mats. Archives of Microbiology 194: 483-491.

Smith, V. H. 1983. Low Nitrogen to Phosphorus Ratios Favor Dominance by Blue-Green Algae in Lake Phytoplankton. Science 221: 669-671.

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Thomas, R. H., and A. E. Walsby. 1986. The effect of temperature on recovery of buoyancy by Microcystis. Journal of general microbiology 132: 1665-1672.

Tonk, L. 2007. Impact of environmental factors on toxic and bioactive peptide production by harmful cyanobacteria. University of Amsterdam.

Trimbee, A. M. and E. E. Prepas.1987. Evaluation of Total Phosphorus as a Predictor of the Relative Biomass of Blue-green Algae with Emphasis on Alberta Lakes. Can. J. Fish. Aquat. Sci. Canadian Journal of Fisheries and Aquatic Sciences 44: 1337-1342.

Welch, E. B., and R. P. Barbiero. 1992. Contribution of benthic blue-green algal recruitment to lake populations and phosphorus translocation. Freshwater Biology 27: 249-260.

Whitton, B. A., and M. Potts. 2000. The Ecology of Cyanobacteria. Kluwer Academic Publishing. World Health Organization. 1998. Cyanobacterial toxins: Microcystin-LR in Drinking-water:

Background document for development of WHO Guidelines for Drinking-water Quality. WHO.

Zhang, W., and Y. Feng. 2008. Characterization of nitrogen-fixing moderate halophilic cyanobacteria isolated from saline soils of Songnen Plain in China. Progress in Natural Science 18: 769-773.

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Figure 1: Master Plan of the Poplar Island Restoration Project. Note the location of Cell 6 in the bottom right corner. (Image credit: http://www.talbotcountymd.gov)

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Figure 2: Anabaena spiroides. (Image credit: Judy O’Neil)

Figure 3: Oscillatoria spp. (Image credit: Judy O’Neil)

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Figure 4: Map of Poplar Island Cell 6 study sites. (Site 2 was not used due to inaccessibility.) NWC: Northwest Cove (Site 1). SEC: Southeast Corner (Site 3). SW16: Spillway 16 (Site 4). SWP: Southwest Peninsula (Site 5)

Site 1 3 4 5

Temperature (°C) 30 24 26 30

Salinity 12 14 14 9

DO (mg/L) 7.2 10.3 17.1 19.4

Chl a 693 1100 1177 959

Phycocyanin 5.8 23.4 26.8 22.7

Table 1: Water Quality Data by Site

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Figure 5: Temperature assay chl a fluorescence trends over time, by site.

Figure 6: Temperature assay phycocyanin fluorescence trends over time, by site.

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Figure 7: Microcystin concentration in temperature assay cultures from each site.

Figure 8: Cyanobacterial composition of each nitrogen fixation subsample in filaments per mL.

Average of all 20 cultures for each date of testing is shown. Note log scale on y-axis.

0.360  

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 1,000,000    

17Jun   24Jun  

Filaments  per  mL  

Date  

Cyanobacteria  Counts  from  N2-­‐Eixation  Experiments  

Anabaena  

Oscillatoria  

Pseudanabaena  

Spirulina  

17

Figure 9: Nitrogenase activity in subsamples from temperature assay cultures, sampled 6/17/13.

Figure 10: Nitrogenase activity in subsamples from temperature assay cultures, sampled

6/24/13.

0.00E+00  

2.00E-­‐05  

4.00E-­‐05  

6.00E-­‐05  

8.00E-­‐05  

1.00E-­‐04  

1.20E-­‐04  

0.00   5.00   10.00   15.00   20.00   25.00   30.00  Average  Instantaneous  Rate  of  Ethylene  

Production  (nmol/hour/heterocyst)  

Time  (hours)  

Nitrogenase  Activity  by  Site  (6/17/13)  

Site  1  

Site  3  

Site  4  

Site  5  

-­‐1.00E-­‐05  

0.00E+00  

1.00E-­‐05  

2.00E-­‐05  

3.00E-­‐05  

4.00E-­‐05  

5.00E-­‐05  

6.00E-­‐05  

7.00E-­‐05  

8.00E-­‐05  

9.00E-­‐05  

0.00   5.00   10.00   15.00   20.00   25.00   30.00  

Average  Instantaneous  Rate  of  Ethylene  

Production  (nmol/hour/heterocyst)  

Time  (hours)  

Nitrogenase  Activity  by  Site  (6/24/13)  

Site  1  

Site  3  

Site  4  

Site  5  

18

Figure 11: Regression of instantaneous rates of ethylene production vs. incubation temperature

at time of acetylene reduction reading.

Figure 12: Regression of instantaneous rates of ethylene production vs. salinity.

0.00E+00  

2.00E-­‐03  

4.00E-­‐03  

6.00E-­‐03  

8.00E-­‐03  

1.00E-­‐02  

1.20E-­‐02  

1.40E-­‐02  

26.50   27.00   27.50   28.00   28.50   29.00   29.50   30.00  

Instantaneous  Rate  of  Ethylene  Production  

(nmol/hour/heterocyst)  

Temperature  (C)  

Nitrogenase  Activity  vs.  Temperature  

0.00E+00  

2.00E-­‐03  

4.00E-­‐03  

6.00E-­‐03  

8.00E-­‐03  

1.00E-­‐02  

1.20E-­‐02  

1.40E-­‐02  

8   9   10   11   12   13   14   15  

Instantaneous  Rate  of  Ethylene  Production  

(nmol/hour/heterocyst)  

Salinity  

Nitrogenase  Activity  vs.  Salinity  

19

Figure 13: Chlorophyll a:Phycocyanin ratios of Oscillatoria cultures in 4 nutrient treatments:

control, nitrogen, phosphorus, and nitrogen+phosphorus.

Figure 14: Sassafras river samples’ nitrogenase activity by treatment after 4-day incubation

period.

0.0  50.0  100.0  150.0  200.0  250.0  300.0  350.0  

C   N   P   NP  Chl  a:Phycocyanin  Ratio    

Treatment  

Oscillatoria  Nutrient  Bioassay    Chl  a:PC  Ratio  

A          B      A        B  

-­‐0.100  

0.000  

0.100  

0.200  

0.300  

0.400  

0.500  

0.600  

0.700  

0.00   5.00   10.00   15.00   20.00   25.00   30.00  nmol  Ethylene  Produced  per  mL  per  hour  

Time  (hours)  

Sassafras  Nutrient  Bioassay:  Nitrogenase  Activity  by  Treatment  

DI  

Control  (Filtered)  

Control  

N  (High)  

N  (Low)  

P  (High)  

P  (Low)  

N+P  (High)  

N+P  (Low)  

20

Figure 15: Diel variation in nitrogenase activity, shown in Sassafras nutrient bioassay.

Treatments shown are (left to right): distilled water, filtered control, unfiltered control, high nitrogen, low nitrogen, high phosphorus, low phosphorus, high nitrogen+phosphorus, low

nitrogen+phosphorus.

-­‐0.100  

0.000  

0.100  

0.200  

0.300  

0.400  

0.500  

0.600  

DI      CF  

 C      NH  

   NL  

   PH  

   PL  

 NPH    NPL  

Average  nm

ol  Ethylene  Produced  per  mL  per  hour  

Treatment  

Sassafras  Nutrient  Bioassay:  Diel  Variation  in  Nitrogenase  Activity  

Light   Dark