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Student Project

Bioremediation of eutrophic estuaries - How salinity effects the algal consumption rate of the Sydney Rock Oyster (Saccostrea glomerata)

Dominic Oberle 2019


Oysters support the largest aquaculture industry worldwide 85% of their natural reef habitat in estuarine waters is under threat due to anthropogenic effects. Oysters are bivalve, indiscriminate filter feeders. Restoration efforts are in place to use The Sydney rock oyster, Saccostrea glomerata, as a bioremediation agent to tackle eutrophication of coastal and estuarine waters. This study aims to test how salinity affects algal consumption rate of S. glomerata. A manipulative laboratory experiment was conducted testing the hourly filtration rate of 16 S. glomerate’ over 3 hours. Four salinities were tested; 0ppt, 15ppt, 30ppt and 45ppt. A spectrophotometer was used to test the optical density of the water at the start and end of the experiment. Oysters were kept in salinity treatments for 2 weeks post experiment to test post treatment survival rates. Overall, there were no significant difference of filtration rate between salinity treatments, however, all oysters subjected to 0ppt died 1-week post experiment. Slightly higher filtration averages were found in the lower salinity treatments, possibly attributed to a higher metabolic demand to osmoconform to the surrounding conditions. This study can highly influence future management planning of site selection for suitable bioremediation locations to reduce eutrophication in estuaries.


Estuaries and coastal waters are not only ecological hotspots but have been economically and culturally significant throughout history (Lotze et al. 2006). Many major cities such as, London, Sydney and Tokyo are located around highly modified estuaries. Estuaries are under immense threat from anthropogenic pressure. Centuries of habitat alteration, pollution and overexploitation has rapidly enhanced estuary degradation, loss of biodiversity and has endangered ecological stability (Adger et al. 2005; Jackson et al. 2001; Lotze et al. 2006). Recent rapid population growth of coastal cities has led to degradation of estuarine ecosystems  and water quality due to contaminants deposited into creeks and rivers flowing into estuaries(Connell 1999; Faulkner 2004; Lee et al. 2006). 

The Sydney Rock Oyster, Saccostrea glomerata (Gould, 1850), is a sessile bivalve mollusc endemic to Australia which can be found in the intertidal zones and estuaries, mainly along the East coast (Schrobback et al. 2014). They are ecosystem engineers, that when found in high densities, provide a range of ecosystem services such as global food security, filtration of water, physical coastal buffer against storms, shoreline stabilisation, carbon store and habitat for fish and invertebrates (Beck et al. 2011; Chris et al. 2018; Jonathan et al. 2012). Oysters support the world’s largest global aquaculture industry (FAO 2018). In 2018, 62% of all aquaculture revenue in NSW was generated from S.glomerata (FAO 2018; NSW Department of Primary Industries 2018).

S. glomerata
are indiscriminate filter feeders that can clear up to 3L of water hour-1 (Bayne et al. 1999). Like most bivalve molluscs they utilise modified gills that capture suspended food particles by creating a incurrent water flow (Bayne et al. 1999; Dove 2003). They remove organic and inorganic suspended particles such as toxic metal contaminates from the water column, reducing the potential for eutrophication, making them a potential bioremediator for degraded estuaries (Spooner et al. 2003). Unfortunately, over the past 130 years, roughly 85% of global oyster reef habitat has been lost to urbanisation of estuaries (Beck et al. 2011).

Oysters have been utilised in numerous bioremediation efforts to mitigate the effects of eutrophication in estuarine waters (Kellogg et al. 2014). Previous studies have explored the effects of salinity, acidification, temperature and acid sulphite soils on filtration rates on other bivalves species; filtration rates of trace metals or the effects of salinity on survival and growth rate of S. glomerata  (Chang et al. 2016; Dove 2003; Green and Barnes 2010; Nell and Holliday 1988; Parker et al. 2011). Current literature lacks a quantitative analysis on how salinity affects the filtration rate of adult Sydney rock oysters.

Through the use of a manipulative laboratory experiment, this study investigates the effect of salinity on the filtration rate of S. glomerata. I hypothesise that there will be a difference in the total algal consumption across the range of salinity levels tested. This study is of importance to support future management plans to locate suitable bioremediation sites to mitigate eutrophication in intertidal river systems and estuaries across Australia and potentially, globally.

Materials and Methods


The algae mix (25% Nanno 3600, 25% Iso 1800, 25% Pavlova 1800 and 25% TW 1200) was diluted to half strength with distilled water to reduce viscosity to improve pipetting accuracy.

As multiple algae species were utilised in the experiment, a preliminary experiment was conducted to obtain the optimal Optical Density (OD) wavelength (Adeeyinwo et al. 2013). A water algae mix was produced which would be used in the experiment. An aliquot of 1ml was placed into a cuvette.


The spectrophotometer (Beckman Coulter DU720) was “blanked” with filtered water. Before each reading, the cuvette was inverted and reverted, twice, then placed into the spectrophotometer and allowed to sit for 2 seconds. At 20nm intervals, readings were taken at wavelengths between 560nm to 740nm.

The highest OD reading of 0.0802 was recorded at 680nm (Figure 1).

Figure 1


A hemocytometer was used to count the algae cell density. 10µL of the algae mix was placed between the coverslip and chamber of each side of the hemocytometer. A random square was chosen from each side and the algae cells were counted using a microscope (Figure 2). This was repeated 5 times. A model was derived to calculate cell count from OD (Figure 3). 109 cell consumed=(OD - 0.0157)/0.0313. An average of 3.375 billion cells mL–1 was found.


Figure 2
Figure 3


30 live S. glomerata of similar length (98mm ± 3mm) were purchased from Franks seafood wholesaler in West End, Brisbane. The organisms were transported on ice from Port Macquarie, NSW. As advised from previous research, the oysters where acclimatised in the marine genomics research aquarium system (UQ, St Lucia) one week prior to the experiment (Thompson et al. 2012). They were fed algae during this period and observed to ensure they are all healthy. Five oysters died during this period. They were frozen and disposed of.

The marine genomics research aquarium system is a 2800L recirculating system which runs at 25 oC and operates on a 12:12 light:dark cycle (Challen, C., personal communication). To promote macroalgae growth, 30ml of iron was added daily (Challen, C., personal communication). Each day 10 ml of each microalgae (Nanno 3600, Pavlova 1800, TW 1200, Iso 1800 from Reed Mariculture) mixed with 500 ml seawater and is dosed hourly (Challen, C., personal communication).

During the experimental period the Marine genomics research aquarium system ran at the following parameters (Challen, C., personal communication);

  • Temp                  25.1oC
  • pH                       8.29
  • Salinity                35 ppt
  • Ammonia            0 ppm
  • Nitrite                  0 ppm
  • Nitrate                 2 ppm
  • Phosphate          3.4 ppm
  • Magnesium        1560 ppm
  • Calcium              470 ppm
  • Iron                     0 ppm
  • Redox                 ~110 mV

Four treatment levels of salinity were used, 0ppt, 15ppt, 30ppt and 45ppt. 18L of reverse osmosis (RO) water was used for the 0ppt treatment, 9L of RO water and 9L of aquarium water was used for the 15ppt treatment, 18L of straight aquarium water was used for 30ppt and 18L of aquarium water with Trophic Marine Pro Reef sea salt was added to reach 45ppt. A refractometer was used to ensure the salinity was at the desired level for each treatment.

24 hours before the experiment, the salinity treatments were created and placed into four 20L buckets. 20 oysters were measured (mm) lengthwise, assigned a number and then divided between the four buckets. Four oysters were used per treatment for the experiment; however, an extra oyster was added to each treatment in case any died during the secondary acclimatisation period. This acclimatisation period was used to negate a potential shock, where the oyster would close and not filter when placed in a new environment, this could skew data. During this 24-hour period the oysters were not fed any algae, starving the oysters, maximising feeding rates during the experiment. A bubbler was added to each bucket to ensure adequate dissolved oxygen in the water.  


16 square plastic containers (3.15L) were placed in a 4x4 grid and assigned a number 1 to 16 (Figure 4). To reduce the effect of any possible environmental factors, each treatment replicate was allocated a random number from 1-16, this corresponded to the container it was placed in during the experiment.


2L of water was added to each container from the respective treatment bucket. All water was strained through a 100µm sieve to improve accuracy of spectrophotometer readings.


A system of aerators was constructed with tubing, 16 pressure valves and air stones to ensure the oysters received sufficient dissolved oxygen and all algae would remain suspended during the experiment. One aerator was added to each container.


One oyster was added to was added to each container of respective salinity treatment. After a 20-minute acclimation period 2ml of algae was added to each container. The first optical density reading was taken. 1ml of water was aliquoted from each container and placed into individual cuvettes. All spectrophotometer readings were recorded at the same time. Before each individual reading, the cuvette was inverted and reverted twice, placed into the spectrophotometer and let to sit for 2 seconds before the OD reading was taken. This was to ensure that any large particles had not settled and that any air bubbles would rise, so that they did not interfere with the reading. A final reading was taken 3 hours after the initial reading.

Figure 4


The oysters were placed back into 20L buckets at the same salinity that they came from. Aerators were added along with 1ml algae mix every 3 days. They were monitored for 2 weeks post-treatment to measure the effect of salinity on survival rates.


The difference in OD from start to finish was taken. From the hemocytometer results we were able to calculate how many cells were consumed throughout the experiment and thus, the filtration rate per hour. A one-way ANOVA and post HOC Tukey test was conducted in R Studio to test the significance of the results.


A change in OD across all oysters was observed, illustrating all oysters filtered algae cells throughout the experiment (Figure 5). The results from the one-way ANOVA found that oysters placed in a range of salinity had no statistical significance on the rate of algae consumption [F(3,12)= 1.318, p=0.314]. A Tukey HSD further indicated that there was no significant relationship between the variables.


0ppt and 15ppt treatments had a much higher mean and standard deviation compared to 30ppt and 45ppt salinity treatments (Figure 6)


One week post-experiment found 100% mortality rate in oysters contained in 0ppt. After two weeks, 100% survival rate was observed in the oysters kept in 15ppt, 30ppt and 45ppt salinity levels.

Figure 5
Figure 6


This study determined that there was no significant difference in the filtration rate of algae consumed over the range of salinities. The hypothesis that salinity would affect the filtration rate of S. glomerata was rejected. This was surprising, considering that there was a 100% mortality of oysters kept in the 0ppt treatment after one week, confirming Holliday’s (1995) findings that S. glomerata can only withstand salinities <15ppt for short periods (Holliday 1995).


In estuarine environments, oysters are subjected to fluctuating salinities, temperature and desiccation. During high rainfall events salinities may be briefly reduced to 0ppt. Ideal growing salinities for adult Sydney rock oysters are 25-35ppt (Holliday 1995). They can tollerate salinity ranges from 0-50ppt, however they will only be able to survive environments <15ppt for approximately two weeks (Holliday 1995).


They have two coping mechanisms to avoid swelling of cells during these periods of reduced salinity (Hoyaux et al. 1976). (1) close their shells and wait for improved conditions or, (2) osmoconform to the surrounding water qualities (Hoyaux et al. 1976). The individual to matches their ionic strength and hemolymph osmotic pressure to the surrounding environment (Bishop et al. 1994; Hosoi et al. 2003). This incurs a high energetic cost (Bishop et al. 1994).


Free amino acids are released from their intracellular protein pool into the hemolymph to restore the osmotic pressure either side of their cell membrane (Hosoi et al. 2003; Parker et al. 2017). Due to these mechanisms, it was surprising that there was no significant difference in filtration rates between treatments. It was expected that the oysters subjected to 0ppt would stay closed or have much lower algal consumption. Possibly, if the experiment was conducted for a longer time period, we would see a difference between results.  


As seen in figure 6, the lower salinities of 0ppt and 15ppt had the highest mean rate of algae consumption hour–1, 24% higher than the higher salinities. 0ppt and 15ppt had a much higher standard deviation (204 x10cells hour-1 and 297 x10cells hour-1, respectively) than the higher salinities (Table 1). This indicates that oysters had variable responses to salinity. The reasoning behind the results is unclear, however one possible explanation could be: oysters at lower salinities must consume more algae to meet the higher energetic costs to osmoconform.


Chesapeake Bay has been one of the many success story regarding bioremediation to mitigate eutrophication.  In 1998, it was estimated that the native oyster, Crassostrea virginica, was at 1% of pre-colonialization levels from over harvesting throughout the 19th and early 20th century (Newell 1988; Rothschild et al. 1994). At pre-colonial levels it was  estimated that the oysters could filter the entire volume of water in the bay in 3.3 days, however, now due to depletion it would take 325 days (Newell 1988). In the 1930’s after the vast numbers of oyster were removed, eutrophication, anoxia, and hypoxia occurred due coastal development, pollution, nutrient run‐off and anthropogenic pressure (Mann and Powell 2007). The Numerous programs now operate within Chesapeake Bay to restore the bays oysters. It has been reported that restored oyster reefs have removed <1% to 15% of annual local nitrogen loads, with a maximum of 25% of daily nitrogen loads (Kellogg et al. 2014). Based on model estimates, harvesting 1 million cultured oysters from Chesapeake Bay would remove 132kg of Nitrogen, 19kg of potassium and 3823kg of carbon from the estuary (Higgins 2011).


In some cases, dense oyster aggregations have been shown to have negative effects on the local environment (Forrest and Creese 2006; Forrest et al. 2009). Physio-chemical alterations to the benthic environment have been recorded in suspended culture when overstocked and repeated culture (Forrest and Creese 2006). Biodeposits (faeces and psuedofaeces) consist of fine organic rich particles which fall and settle onto the surrounding seabed, creating an organically enriched “footprint”, leading to anoxic local conditions (Forrest and Creese 2006; Forrest et al. 2009). This can lead to a trophic shift of the benthic structure, decreasing the biodiversity of the meiofaunal assemblage, displacing macrofauna while favouring disturbance tolerant species (Forrest et al. 2009). However, benthic enrichment is considerably lower in oyster culture compared to suspended culture of fish (Karakassis et al. 2000). To avoid anoxic benthic conditions, stocking density limitations needs to be influenced by the hydrological properties. Preliminary hydrological studies should be conducted to analyse potential bioremediation sites to ensure sufficient tidal flushing is present. This has been shown to lessen the intensity but broaden the biodepositional footprint (Pearson and Black 2000).


If this study was to be repeated, it would be recommended to take three aliquots from each sample to find a mean OD value from each replicate. During the experiment it was found that it was possible to get highly variable results from each container, as suspended particles such as waste or debris can interfere with the spectrophotometer readings. This was noticed half way through the experiment and the above methodology was implemented for the last set of readings. This could explain some of the variability in the initial readings.


The findings of salinity having no significant effect on filtration rate allows us to spatially expand the range at which possible bioremediation sites could be positioned. Oyster aquaculture sites could be placed further upstream, where they would receive lower salinity levels but clean the water, minimising the effects of eutrophication in the bay. It would have to be in the intertidal zone, as mortality would occur if exposed to fresh water for extended periods. Future laboratory studies would have to be conducted to test long term filtration and mortality rates at lower salinities. In conjunction with saline mapping of a proposed estuarine environment.  If successful, field studies would be conducted at various stages upstream of river mouths to test the viability of potential bioremediation locations.


The value of oysters worldwide cannot be underestimated. They not only provide global food security as the largest aquaculture resource but provide invaluable ecosystem services; including habitat for fish and invertebrates, stabilising shorelines, provide a buffer to storms, act as a carbon store but also filter water to prevent eutrophication of our economically and culturally treasured estuarine waters. It is important to remember that oyster reef restoration should not be viewed as the key to offset current anthropogenic nutrient loads, however if implemented correctly could potentially mitigate some environmental stressors.


I would like to thank Bernie, Sandie, Chris and the tutors for their help throughout the duration of this research project.


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