Zn Isotope Fractionation in the Oyster Crassostrea hongkongensis and

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Environmental Measurements Methods

Zn isotope fractionation in the oyster Crassostrea hongkongensis and implications for contaminant source tracking L Ma, Yunlong Li, Wei Wang, Nanyan Weng, Robert Douglas Evans, and Wen-Xiong Wang Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.8b06855 • Publication Date (Web): 18 Mar 2019 Downloaded from http://pubs.acs.org on March 18, 2019

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Environmental Science & Technology

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Zn isotope fractionation in the oyster Crassostrea hongkongensis and implications for contaminant source tracking

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Lan Maa,b, Yunlong Lib, Wei Wanga, Nanyan Wengb, R. Douglas Evansc, Wen-Xiong Wangb*

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a Environment & Life Sciences Graduate Program, Trent University, 1600 West Bank Drive, Peterborough, ON, K9L 0G2, Canada

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b Department of Ocean Science, The Hong Kong University of Science and Technology (HKUST), Kowloon, Hong Kong, and Marine Environmental Laboratory, Shenzhen Research Institute, HKUST, Shenzhen 518000, China

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c Water Quality Centre, Trent University, 1600 West Bank Drive, Peterborough, ON, K9L 0G2, Canada

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*Corresponding author: [email protected]

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ACS Paragon Plus Environment


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Abstract Variations in stable isotope ratios have been used to trace the sources of contaminants as well

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as their biogeochemical pathways in the environment. In the present study, we investigated the

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influences of internal redistribution among tissues and ambient water conditions on Zn isotope

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fractionation in oysters. There was no significant difference in Zn isotope ratios during in vivo

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Zn transportation among various oyster tissues. Estuarine oysters were exposed to additional Zn

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either at different salinities or at different Zn concentrations, following which the Zn isotope

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ratios in the oysters were measured. Results showed no significant difference in δ66/64Zn values

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in the oysters exposed at different salinities. Tissue Zn accumulation increased with increasing

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Zn levels in water over the 30 days exposure. Within this period there was a nearly 0.3‰

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difference in averaged δ66/64Zn values in the exposed oysters compared to the initial δ66/64Zn

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values in the oysters prior to exposure. However, there was no evidence of significant difference

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in δ66/64Zn values in oysters exposed at different Zn levels, with post-exposure signatures similar

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to the δ66/64Zn values of the Zn solution added. Our results suggested that the δ66/64Zn values

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measured in the oysters were approaching the δ66/64Zn values of the ‘source’ faster with

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increasing Zn concentrations added in the water. This study highlighted the absence of Zn

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isotope fractionation during Zn internal distribution and in vivo transport in oysters. Calculation

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of the contributions of different Zn sources demonstrated that oysters can be a sentinel animal for

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Zn source tracking in marine environments.

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Keywords: Zinc, oysters, isotopic fractionation, source tracking, bioaccumulation

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TOC

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Introduction Oysters and other bivalves are used frequently as sentinel organisms in aquatic systems to

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monitor environmental contamination because of their outstanding ability to accumulate

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contaminants such as Zn 1–3. Their tissue concentrations are generally considered to be

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representative of spatial and time-integrated bioavailable metals at the collection site 4–8. Lu et al.

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9

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(including the Pearl River Estuary, PRE), to determine the bioavailable metals from the collected

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environments. Pan and Wang 10 showed that the high levels of metals in bivalve tissues were

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proportional to the levels of bioavailable metals in the exposure environments. Many laboratory

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experiments have also been conducted to examine the influences of various physico-chemical

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conditions (e.g., metal concentration, speciation and exposure conditions such as temperature,

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salinity, metal mixture) on metal uptake, accumulation and elimination in these oysters 11,12.

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Variations in the levels of metals including Cu, Zn, Ni and Cd have been observed among

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different organs in oysters, resulting from the differences in the physiological roles of organs or

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tissue-specific metal bioaccumulation patterns 11.

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measured the metal concentrations in oysters at >30 sites in the Chinese coastal waters

Recent studies suggest that Zn stable isotope ratios can be used for source determination

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and possibly tracing Zn biogeochemical pathways in the environment 13,14. Oysters are known to

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accumulate extraordinarily high concentrations of Zn in their soft tissues, and in some

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contaminated estuaries, concentrations could be as high as 6% of the tissue dry weights 7.

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Previous studies have attempted to use Zn isotope ratios in oysters to attribute sources, for

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example, in Desolation Sound and Barkley Sound of British Columbia, Canada 15,16, in the

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fluvial Gironde Estuary, France 17 and in Sepetiba Bay, Brazil 18. The Zn isotope signature

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observed in the biota may be due to either the signature(s) of the source(s) or isotopic

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fractionation resulting from any physical, biological or chemical changes within the organism

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itself, or a result of all these factors combined. Weiss et al. 19 found that in a soil-plant system,

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enrichment of heavy Zn isotopes occurred in the roots compared to the exposure solution,

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suggesting that plant uptake of Zn was an important process affecting the isotopic fractionation.

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In mammals such as sheep and mice, Zn isotopes also fractionated between organs, body fluids,

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diets and feces, suggesting cellular fractionation 20. In oysters, previous studies focussed on

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either identifying the source contribution of Zn 15–18 or assessing the impact of other factors (e.g.,

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salinity, temperature, metal concentrations) on Zn concentrations 11,12. No attempt has been made 4


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to determine the isotopic composition of Zn among various tissues, in contrast to other metals

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(e.g., Hg) in other biota 21,22. Nevertheless, prior to the use of isotope ratios in sentinel organisms

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to identify and partition the sources, it is critical to test the effect of internal fractionation on

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isotope signatures in oysters for future application of Zn isotope in source tracking.

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In the present study, Zn isotope ratios in the estuarine oyster Crassostrea hongkongensis

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(C. hongkongensis) obtained from the PRE, where concentrations of 2,000 - 4,900 µg Zn /g were

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previously reported8, were analysed in several tissues to better understand the effects of Zn

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distribution among tissues on Zn isotope fractionation. Two controlled laboratory experiments

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were also conducted by exposing uncontaminated oysters to different levels of Zn at one salinity

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or to one Zn level but at different salinities. Zn isotope ratios were then analyzed to obtain better

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insights into the effects of environmental factors (i.e., salinity and Zn levels) on Zn isotope

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fractionation in these oysters. To better constrain the experimental conditions, we employed only

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one single ‘source’ of Zn (i.e., constant Zn isotope signature in the exposure solution). The

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degree of Zn isotope fractionation was determined by comparing either the δ66/64Zn values

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among different tissues or the δ66/64Zn values of the ambient Zn source to the δ66/64Zn values in

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oysters. Finally, we constructed a mixing model to quantify the sources of Zn accumulation in

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the oysters under the laboratory-controlled conditions.

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Materials and Methods

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Depuration of oysters

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Oysters (average tissue dry weight of 0.6 g) with high Zn levels in their soft tissues were

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collected from the Pearl River Estuary (PRE), China, in December 2016, and brought back to the

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laboratory within 6 h. Oyster shells were cleaned by using a plastic brush to gently remove the

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particulates on the shells. Seawater of a salinity of 20 psu was prepared for oyster depuration by

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mixing filtered (< 1 µm) seawater (salinity of 30 psu) collected from the eastern side of

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Shenzhen, China (22°31'47'' N 113°29'21''E) with high purity water (HPW 18 MΩ cm) at a ratio

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of 2:1. Three oysters were allowed to depurate in aerated seawater at a salinity of 20 psu for 3

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days, as described in Pan and Wang10. Meanwhile, the other three individuals were kept frozen

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before further processing. After 3 days, the soft tissues of all six oysters were gently separated

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from the shells, rinsed with HPW and then were dissected into four parts, i.e., mantle, gill, 5



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digestive gland, and adductor muscle, and placed into individual acid-washed centrifuge tubes.

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All the tubes were kept in plastic zip bags and then freeze-dried at -80 °C until further

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processing. Dry weights of the soft tissues were recorded.

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Oyster exposure at different Zn concentrations

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Oysters (C. hongkongensis, average tissue dry weight of 0.7 g) used in the exposure

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experiment were collected from Beihai, Guangxi, China (21°25'32.0''N 109°17'33.9''E), a site

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remote from any urban and industrial activities, in June 2017. In the laboratory, oyster shells

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were gently cleaned with a plastic brush and then they were acclimated for 4 days in seawater

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with a salinity of 20 psu (the same as the Beihai collection site) at 20 °C. Similar to the seawater

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preparation for oyster depuration, exposure water (i.e., with a background Zn concentration of 3

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µg L-1) used in the current study was also prepared by mixing seawater collected from the

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eastern side of Shenzhen with HPW at a ratio of 2:1, to obtain a salinity of 20 psu. Water was

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aerated and changed every two days and the oysters were fed with algal powder (ORI-GO,

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Skretting) at a feeding rate of approximately 2% of oyster soft tissue dry weight every two days.

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To prepare the exposure water at different Zn levels, 0.01 g of Zn (ZnCl2 powder, Riedel-

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de Haën) was weighed into a clean centrifuge tube and 10 mL of 1% HNO3 diluted from double-

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distilled (duoPUR system, Milestone) trace metal grade HNO3 (VWR International) was added to

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obtain the Zn stock solution, which was used for the entire exposure period. Twelve 15 L plastic

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tanks were washed with 2% HNO3, then rinsed three times with HPW and filled with 14 L

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seawater (20 psu). Appropriate amounts of the original Zn solution were then added to the

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ambient seawater in each tank to obtain the desired added (nominal) Zn concentrations (i.e., 0,

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10, 20 and 100 g Zn L-1, respectively). Since the volume of the exposure seawater (14 L) was

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much larger than the volume of the Zn solution added to the seawater (e.g., maximum 7×10-4 L

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of stock solution were added to obtain 100 g Zn L-1), neither pH (the Zn stock solution was kept

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in 1% HNO3) nor the total seawater volume in each tank was influenced by the original Zn

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solution added. The water in each tank was stirred manually and then allowed to equilibrate for 1

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h at 20 °C before transferring 12 oysters of similar shell lengths into each tank. The water was

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continuously aerated and renewed every two days in order to keep approximately constant Zn

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levels in the exposure tanks during the entire experimental period. While the seawater was being

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renewed, the oysters were transferred to separate containers with seawater at 20 psu without Zn 6


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solution added and fed algal powder at a rate of approximately 2% of oyster soft tissue dry

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weight. The plastic tanks were then rinsed with HPW three times and then filled with 14 L of 20

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psu seawater, followed by different amounts of Zn stock solution as required for each exposure

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concentration. After 1.5 h, oysters were moved back to the specific tank for continuing the

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exposure.

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Oysters from each tank were harvested after 30 days of exposure and depurated in seawater

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at 20 psu for 2 h in order to remove any Zn carried in oysters from the exposure water before

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further processing. Two individuals were randomly chosen from each tank and their soft tissues

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were gently separated from the shell, rinsed with HPW and placed in a clean centrifuge tube.

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Wet weights were recorded and then soft tissues were freeze-dried at -80 °C, following which

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dry weights were recorded. The contents of the two centrifuge tubes, i.e. the two individuals

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sampled, from the same tank were pooled together to provide enough material for total Zn and

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Zn isotope ratio determinations. Mortality was less than 4% (i.e., 4 individuals out of a total of

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108) after 30 days and there was no apparent toxicity arising from exposure at higher metal

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levels.

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Six of the oysters selected before the start of exposure were also dissected and processed

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for Zn concentrations and δ66/64Zn values determination. Their soft tissues were also gently

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separated from the shell and stored in separate centrifuge tubes. The soft tissues were then

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freeze-dried at -80 °C and the dried tissues from two centrifuge tubes were pooled together

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(resulting in n=3) for further total Zn and Zn isotope ratio determination. These values

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represented the background Zn concentration and isotope signatures of oysters in this exposure

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experiment (BG).

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Oyster exposure at different salinities

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In another laboratory experiment, oysters were exposed to a single Zn concentration at

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different salinities following which the Zn concentrations and the δ66/64Zn values in the oysters

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were determined. Exposure water of four salinities, i.e., 5, 12, 20, 28 psu was prepared by mixing

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the HPW with seawater of 30 psu. Similar sized oysters (C. hongkongensis) (averaged tissue dry

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weight of 0.3 g) were collected from the Beihai, Guangxi, China with salinity of 20 psu and then

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acclimated sequentially to these 4 levels by increasing/decreasing salinity by 4 psu every two

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days. After being acclimated under the laboratory conditions for 15 days, 23 oysters were 7



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transferred to each of 12 (i.e., 4 salinity exposures x 3 replicates) 60 L plastic tanks. Density of

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oysters used in this experiment was lower than that used in the Zn concentration experiment

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primarily due to the longer time of exposure. Zinc was added at a concentration of 5 g L-1. The

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exposure water was changed every 2 days and oysters were removed from the exposure tanks,

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placed into separate containers with seawater at the appropriate salinities, without adding Zn, and

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fed the algal powder for 1.5 h. The oysters were then transferred back to the exposure waters,

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which were continuously aerated. At the end of 6 weeks exposure, oysters were harvested, and

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one individual was collected randomly from each tank. Soft tissue samples were separated from

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the shell and prepared for analysis as described above.

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Sample processing and Zn isotope ratio analysis

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Freeze-dried oyster tissues were ground to a fine powder and mixed homogenously.

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Approximately 0.2 g of dried oyster tissue were digested with 10 mL of 15.6 M HNO3 at 100

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bars and 110 °C using a microwave digestion system (Milestone Srl). Solid samples were

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completely digested after one hour and the clear liquid was transferred into 30 mL PTFE vessels.

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Samples were then dried on a hot plate at 90 °C and re-dissolved in 10 mL of 0.5 M HNO3 for

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further analysis. Certified Reference Material oyster tissue 1566a (NIST), and USGS rock

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materials (i.e., BHVO-2 and AGV-2) were subjected to the same digestion procedures for

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digestion batch to monitor the recovery and the rock materials were used for further comparison

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of accuracy of Zn isotope ratio analysis.

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A 0.05 mL subsample from each sample was diluted with 0.5 M HNO3 according to

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anticipated total Zn concentrations in the samples. Total Zn concentrations were measured by

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Triple Quadrupole Inductively Coupled Plasma Mass Spectrometry (ICP-MS) (Agilent 8800,

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Agilent Technologies) at the Water Quality Center (Trent University, Canada); a high purity

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ICP-MS multi-element standard solution (SCP Scientific, Canada) was used for calibration.

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Standard reference materials were also for the calibration.

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In order to achieve the precision and accuracy of measurement of Zn isotope ratios

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necessary for comparison among treatments, ion exchange chromatography was utilized for the

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separation of Zn from the sample matrix and a double-spike protocol and standard-sample

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bracketing were used for Zn isotope ratios determination 23. In brief, a 67Zn-70Zn double spike 8


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solution was added to all the samples and Zn isotope standards (IRMM-3702) with a

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spike:sample ratio of 1:2, which was calculated from an optimal spike-to-sample mixing

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proportion algorithm developed by Rudge 24. One hundred ng of Zn was then prepared for

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further Zn isotope ratio analysis by mixing approximately 67 ng Zn from the sample with 33 ng

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Zn from the Zn double-spike solution. The mixed sample/spike was then kept in a Teflon vessel

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and dried on a hotplate at 100 °C. One mL of 2 M HCl diluted from double-distilled (duoPUR

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system, Milestone) trace metal grade HCl (VWR International) was added after dryness and the

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sample was dried again. The dried sample was re-dissolved in 1 mL of 7 M HCl before

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separation of Zn from the matrix.

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Zn was separated from the sample matrix using an ion exchange chromatography

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procedure modified from Weiss et al. 25 to minimize the spectral interferences during mass-

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spectrometric analysis and/or significant matrix effects on signal intensities. Acid washed AG

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MP-1 100-200 mesh anion exchange resin (1.6 mL, Bio-Rad Laboratories, Mississauga, Canada)

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was loaded into a 10 mL plastic chromatography column (Bio-Rad Laboratories). The resin was

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rinsed with 10 mL of 0.5 M HNO3 and 2 mL of HPW alternatively, three times, after which 6

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mL of 2 M HCl was loaded onto the column for column conditioning. The 1 mL sample in 7 M

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HCl was then loaded onto the column. The major matrix elements were firstly eluted with 30 mL

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of 7 M HCl and 10 mL of 2 M HCl. Then Zn was eluted from the column with 12 mL of 0.5 M

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HNO3 and collected into clean Teflon vessels. New resin was used for each sample. The eluent

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containing the Zn was evaporated to dryness, then 2 mL of 0.5 M HNO3 was added and the

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samples were refluxed for 4 h (i.e., frequently stirred by gently moving the vessels) to ensure

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complete dissolution. In addition, a Zn standard solution (AA-ETH Zn) obtained from ETH

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Zürich (Switzerland) was also doped with 67Zn-70Zn double spike with a sample:spike ratio of

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2:1 and the Zn isotope ratio was determined for inter-calibration.

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PTFE vessels used for sample digestion as well as sample Zn isotope ratio measurement

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were cleaned in a PTFE beaker in a solution of boiling 6 M HNO3 + 6 M HCl for 3 days, and

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then cleaned again with 50% HNO3 for a day before storage in 10% HNO3. Vessels were rinsed

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with HPW before use. All the vessel cleanings and sample preparations were conducted in a

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Class 10000 metal free clean room and Zn isotope ratios in the samples were measured by multi-

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collector (MC) ICP-MS (Nu Plasma II) at the Water Quality Center, Trent University, Canada. A 9



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dsn-100 desolvating nebulizer was used for sample introduction (CETAC Technologies, USA).

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Zn double-spike isotope data collected from the mass spectrometric analyses were further

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processed and double-spike data reduction was performed offline by applying the methods

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outlined by Siebert et al. 26, in order to obtain the mass bias corrected Zn isotope composition in

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un-spiked samples. All the delta Zn values (units of ‰) were calculated relative to an averaged

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Zn standard reference material IRMM3702 (i.e. 3702-ave) with standard-sample-standard

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bracketing (Eq. 1).

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δ66/64Zn = ((66Zn/64Zn)sample / (66Zn/64Zn)3702-ave -1) × 1000

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where (66Zn/64Zn)sample is the value of the measured ratio in a sample and (66Zn/64Zn)3702-ave is the

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average value of the ratio for IRMM 3702 before and after that sample.

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The δ66/64Zn value in the Zn solution used in the exposure experiments was also measured by

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following the same analyzing procedure as the oyster samples.

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Statistics

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(1)

Statistical analyses were carried out using SPSS 16.0 with α = 0.05. Multiple ANOVAs

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and Tukey’s honestly significant differences post hoc test were performed to examine the

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differences in either total Zn concentrations or δ66/64Zn values in oysters from different

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treatments. Sigmaplot 12.0 was used to create graphs.

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Results and Discussion

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Quality assurance (QA)/Quality control (QC)

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The recovery of the total Zn was 100-103%, 86%, and 87% relative to the certified values

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of NIST 1566a (oyster tissue), CRMs BHVO-2 (rock), and AGV-2 (rock) (http://georem.mpch-

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mainz.gwdg.de), respectively. The contributions of Zn from the digestion procedures, AG MP-1

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resin only, and instrument background were 3%, 3.2% and 0.05%, respectively, indicating that

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the contribution of Zn from sample preparation was negligible and the observed Zn isotope ratios

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among the samples were not unduly influenced by Zn introduced by the analytical procedures.

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The NIST 1566a oyster reference material was determined to have a δ66/64Zn value of -0.46 ±

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0.07‰ (1sd, n=4). The measured δ66/64Zn values for the rock CRMs were 0.08‰ (n=1) for 10


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BHVO-2 and 0.02 ± 0.001 ‰ (1sd, n=2) for AGV-2, which agreed well with the values reported

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in the literature 18 after normalization to the JMC standard. The δ66/64Zn values in AA-ETH Zn

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solution relative to IRMM 3702 were -0.01 ± 0.05‰ (1sd, n=2), which was in agreement with

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the values reported by Archer 27, validating the δ66/64Zn values determined in the present study.

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Zn isotope fractionation in oysters during depuration and tissue partitioning The total Zn concentrations in oysters with or without 3 days depuration was 7,057 ± 313

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µg g-1 (1sd, n=2) and 6,898 ± 410 µg g-1 (1sd, n=3), respectively, showing there were no

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significant differences (p > 0.05) in the total Zn amount in oyster before and after 3 days of

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depuration at 20 psu. However, there were significant changes in the total Zn concentrations

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among tissues, with a 56% reduction in the Zn concentration in gills but nearly 1.98 times

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increase in Zn in the mantle after depuration (bars, Fig. 1). The changes in these two Zn pools

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kept the Zn mass balance in the oysters (i.e., 60.8% of Zn lost from the gill vs. 61.4% of Zn

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increase in mantle), indicating in vivo Zn re-distribution during this depuration process. Weng et

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al. 28 visualized the subcellular distribution of Zn as well as the types of cells which accumulated

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Zn in gill and mantle in oysters (C. hongkongensis), and showed the variations in metal-rich sites

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and diversity of metal complexations in gill and mantle. The rapid drop of the Zn concentration

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in the gill and increase of Zn concentration in the mantle were apparently caused by Zn

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transportation between these organs.

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Unlike total Zn concentrations in the oyster tissues, no significant difference (p > 0.05) in

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the δ66/64Zn values was found in oyster tissues with or without 3 days of depuration (circles, Fig.

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1), consistent with the insignificant change in total Zn concentration in oysters after three days.

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Pan and Wang 10 reported an efflux rate as low as 0.006 d-1 of Zn in oysters collected from a

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location remote from contamination, showing insignificant changes in the total Zn concentrations

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in oysters for typical depuration times. However, it is difficult to predict whether significant Zn

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isotope fractionation and redistribution would occur in oysters over a longer depuration time. In

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addition, similar δ66/64Zn values were observed in the four tissues even when the Zn

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concentrations in the adductor muscle were only 17% of the Zn in the other three organs (Fig. 1,

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bars). Similarity of the δ66/64Zn values in the four tissues suggested no significant Zn isotope

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ratio fractionation during the Zn in vivo transportation in oysters. 11



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Zn isotope fractionation at different salinities Zinc concentrations and the δ66/64Zn values in oyster tissues were determined after

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exposure to Zn at different salinities (Fig. 2). Results showed that there was no significant

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difference in Zn concentrations in oysters exposed at different salinities (p > 0.05; Fig. 2 left

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panel), although slightly higher average Zn concentrations were found in oysters at 12 psu and

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lower levels at 28 psu (i.e., 1978 µg g-1 and 1304 µg g-1, respectively, Table 1). The effects of

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salinity on the physiology as well as metal uptake by oysters were studied previously. For

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example, Olive et al. 29 determined the effects of salinity (15 to 45 psu) on the filtration rates of

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juvenile tropical oysters (Crassostrea iredalei), and the oysters displayed the lowest filtration

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rate at 15 or 45 psu but the highest filtration rate 35 psu. Such a difference in filtration rate in

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bivalves 30 may result in different Zn uptake by oysters at different salinities 12. Also, Turner et

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al. 31 noted from laboratory experiments that a larger amount of Zn was complexed with chloride

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with increasing salinity, forming chloro-complexes 32. However, salinity did not cause significant

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differences in Zn tissue concentrations in oysters at different salinities under the current

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experimental conditions.

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In addition, there were no significant differences in δ66/64Zn values measured in the oysters

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exposed to 5 µg Zn L-1 at the four salinities (p > 0.05, Fig. 2 right panel), suggesting the absence

300

of significant influence of salinity on the δ66/64Zn values in oysters. However, because the Zn

301

concentrations in oysters were the same at different salinity, it was difficult to conclude whether

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the smaller variations in δ66/64Zn values in oysters were caused by the similar Zn concentrations

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in oysters or due to the lack of salinity effect on the δ66/64Zn values. Moreover, the δ66/64Zn value

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in the Zn solution used in the exposure experiment was 0.06 ± 0.02‰ (1sd, n=2), which differed

305

by 0.20‰ to 0.38‰ from the averaged δ66/64Zn values in the oysters. Such difference suggested

306

the possibility of using this system to partition Zn isotopes in the oysters compared to the Zn

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isotopes in water (‘source’) (discussed below).

308 309 310 311

Effects of Zn exposure levels on Zn isotope fractionation As expected, higher concentrations of Zn exposure led to higher Zn bioaccumulation in the oysters, with a significant linear correlation between the Zn concentration in oysters versus the 12


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Zn levels added in the exposure water (r2= 0.99, p < 0.05, data not shown). All the Zn

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concentrations in oysters after exposure were significantly (p < 0.01) higher than the initial Zn

314

concentrations in the background oysters (i.e., BG, hatched bar, Fig. 3 left panel). The measured δ66/64Zn values in the oysters were -0.11±0.12‰, 0.02±0.03‰ and

315 316

0.03±0.12‰ at 10, 20 and 100 µg Zn L-1, respectively (Table 1). These δ66/64Zn values were not

317

significantly different from each other, showing that there was no significant effect of Zn

318

exposure levels on the δ66/64Zn values in oysters despite large differences in Zn concentrations in

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the oysters (e.g., oysters exposed at 100 µg L-1 displayed a 2x higher Zn concentration than those

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at 20 µg L-1, Table 1). In both exposure experiments (salinity and Zn levels), there was no

321

correlation between the Zn concentration in oysters and the δ66/64Zn values (Fig. 4). Thus, similar

322

δ66/64Zn values in the oysters exposed at 4 salinities were not caused by the similar Zn

323

concentrations, but were instead due to the lack of salinity influence.

324

In this experiment, a nearly 0.03‰ to 0.17‰ difference in the averaged δ66/64Zn values in

325

the oysters was observed from that in the Zn solution. The δ66/64Zn values in oysters exposed at

326

10, 20 and 100 µg Zn L-1 were plotted against the Zn concentrations added to water (Fig. 5,

327

triangle up in pink, green, yellow, respectively). Interestingly, the trend in δ66/64Zn values

328

followed closely with the Zn levels in the water. We also calculated the uptake rate of Zn by

329

oysters (averaged dry weight of 0.7 g) exposed at different Zn levels at 20 psu (∆Zn/g/d =

330

([Zn]oyster - [Zn] BG oyster)/0.7/30). The estimated ∆Zn concentration at 5 µg Zn L-1 was 291.5 µg g-

331

1,

332

the same exposure time from the salinity experiment (averaged dry weight of 0.17 g) at 5 µg Zn

333

L-1 and 20 psu. Given the similar Zn uptake by oysters between the two exposure experiments,

334

the δ66/64Zn values in oysters exposed at 5 µg Zn L-1 and 20 psu were also plotted against the Zn

335

concentrations added to water (Fig. 5, open triangle up). The regression curve showed that the

336

δ66/64Zn values in oysters probably reflected the δ66/64Zn values of Zn ‘source’ at specific Zn

337

water concentrations.

which was comparable to the ∆Zn concentrations in oysters (i.e., 290.5 µg g-1) calculated for

338 339 340 341

Source contributions Neither salinity nor Zn exposure level significantly influenced the δ66/64Zn values in oysters in the current study. It is worth noting that in the Zn exposure experiment, there was a 13



Page 14 of 27

342

significant change in Zn isotope composition in oysters before (i.e., BG oysters, a δ66/64Zn value

343

of -0.38±0.11‰, Table 1) and after the exposure (i.e., -0.04±0.09‰, -0.11±0.12‰, 0.02±0.03‰

344

and 0.03±0.12‰ at 0, 10, 20 and 100 µg Zn L-1 added, Table 1). Thus, although the primary

345

source of Zn post-exposure was the Zn added in our exposure tanks, the measured δ66/64Zn in

346

oysters after the exposure period resulted from a mixture of different Zn sources, i.e., Zn in

347

ambient water, Zn solution added, Zn contained in the algal powder, and finally the background

348

Zn in oysters prior to exposure (BG oyster). The ambient seawater was a mixture of HPW (non-

349

detectable Zn concentration) and the seawater at 30 psu salinity at a ratio of 1:2 (as outlined in

350

the methods), which had a dissolved Zn concentration of 5 µg L-1 and a δ66/64Zn value of 0.15‰

351

(unpublished data). Consequently, the concentration of the ambient exposure water was 3 µg L-1.

352

The Zn concentration and the δ66/64Zn values for the algal powder were 31.7± 3.16 µg/g d.w.

353

(1sd, n=5) and -0.23 ± 0.04 ‰ (1sd, n=5), respectively. It was not possible to determine the exact

354

amount of algal powder ingested by each oyster. However, we can calculate the maximum

355

amount of Zn taken up by the oysters by assuming a 100% feeding and a 100% assimilation of

356

Zn from the algae by the oysters. Our calculation suggested that a maximum of 3% of Zn

357

accumulated by the oysters may be derived from the potential feeding of algae. As a result, Zn

358

accumulation from algae can be ignored and the relative contributions of the three remaining

359

sources of Zn to the δ66/64Zn values in the oysters from the Zn exposure experiments can be

360

calculated. Therefore, it is possible to use a simple mixing equation to estimate the proportions

361

of δ66/64Zn originating from each source to the oysters, using the following equations (2)-(4)

362

modified from Chen et al. 33:

363

δ66/64Zn Oyster = XA δ66/64Zn A + XB δ66/64Zn B + XC δ66/64Zn C

(2)

364

XA + XB + XC =1

(3)

365

XB /XC = (CZn) B/(CZn) C

(4)

366 367

where A represents the BG oysters, B stands for the spiked Zn in seawater, and C is the ambient

368

Zn in the exposure seawater. XA, XB and XC are the proportional contributions from each δ66Zn

369

value to the oysters. In equation (4), (CZn) B represents the different Zn exposure concentrations,

370

and (CZn) C is 3 µg L-1 in the ambient seawater.

371 372

In oysters exposed to seawater without Zn solution added , the contribution to δ66/64Zn values from BG in the oysters (A) was 64%, with 36% originating from the Zn present in the 14


Page 15 of 27


373

ambient seawater (C) (Fig. 6, triangle up in blue color). The contribution to δ66Zn from the

374

spiked Zn increased with increasing Zn concentrations in the water. At (CZn) B = 100 µg L-1, for

375

example, up to 90% of the observed δ66/64Zn value came from the spiked Zn with only 3% from

376

the ambient seawater (C) and 7% from the BG oyster (Fig. 6, triangle up in yellow color). These

377

calculations showed that measured δ66/64Zn values in oysters can be explained by the mixing of

378

the various Zn sources. These calculations were only applicable to the oysters exposed to

379

different Zn levels because we lack information on the δ66/64Zn values in either of the BG oyster

380

or the seawater used in the salinity experiment. However, based on the regression line (Fig. 5) by

381

combing the δ66/64Zn values in oysters exposed at 20 psu obtained from the two different

382

experiments, the δ66/64Zn values determined in the oysters were caused mainly by different

383

contributions from the ‘source’. The present study therefore demonstrates the insignificant

384

influences of water salinity and Zn exposure levels on Zn isotope fractionation in the oysters,

385

which predominantly reflected the δ66/64Zn values of the exposure source. It is thus possible to

386

use oysters as a sentinel organism for contamination source evaluation by analyzing the Zn

387

isotope ratios.

388 389 390

Implications The δ66/64Zn values in oyster tissues collected from the field (i.e., PRE) were analyzed.

391

Although after depuration for three days, the total Zn concentrations in the gills dropped by 50%

392

but increased by nearly 1.7 times in the mantle, there was no significant difference in δ66/64Zn

393

values between the gill and mantle, indicating that Zn transport, storage and detoxification

394

caused no Zn isotope fractionation. In addition, variations of salinity and Zn exposure levels did

395

not have a major impact on Zn isotope fractionation, suggesting that the isotopic composition of

396

Zn in oysters is primarily determined by the signature of the exposure ‘source(s)’. As a result,

397

differences in δ66/64Zn values in our oysters before and after exposures (i.e., background and

398

newly accumulated) are a mixture of δ66/64Zn values from the different contributing sources (i.e.,

399

ambient seawater and spiked Zn solution), primarily related to the amount of Zn from each

400

source. Our study suggests that Zn isotope ratios in oysters can be used as a tracer of sources of

401

Zn in aquatic systems, since there was little evidence of Zn fractionation during Zn uptake or

402

transport in the oysters under various conditions. 15



403 404

Acknowledgement

405

We thank the anonymous reviewers for their comments on this work. This research was funded

406

by a NSERC Canada (Natural Sciences and Engineering Research Council) Discovery Grant to

407

R.D.E., and a TUYF fund (19SC01) to W.-X. W. We would like to thank Hayla Evans for

408

helping revise the manuscript.

16


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409

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503 504 505


Table 1. Amount of Zn added to exposure water at different levels and salinities, measured total Zn concentrations (µg g-1) and δ66/64Zn (‰) values in oysters collected after the exposures. Mean ± 1 standard deviation (n=3).

506

g-1)

Oyster δ66/64Zn (‰)

245.4±55.4 560.5±212.9 577.0±71.3 745.8±169.5 1339.8±181.0

-0.38±0.11 -0.04±0.09 -0.11±0.12 0.02±0.03 0.03±0.12

Salinity experiment 5 psu 5 1593.1±226.5 12 psu 5 1978±437.5 20 psu 5 1490.5±208.3 28 psu 5 1304.5±197.9 BG: background oysters before exposure to different Zn levels

-0.14±0.08 -0.16±0.08 -0.32±0.12 -0.28±0.09

Zn added (µg L-1)

[Zn] (µg

Zn level experiment BG* Control 10 20 100

507

0 10 20 100

508 509 510

21



511 512 513 514 515 516

Fig. 1. Total Zn concentrations (bars) and δ66/64Zn values (circles) in gill, mantle, digestive gland and adductor muscle tissues of oysters collected from the Pearl River Estuary. Solid bars and circles represent oysters without depuration (i.e., 0 days), while open bars and circles represent oysters after depuration for 3 days. Symbols presented as averaged values ± 1sd (n=2 for each tissue).

22


Page 22 of 27

Page 23 of 27


517 518

519 520 521 522

Fig. 2. Total Zn concentrations (left panel) and δ66/64Zn values (right panel) in oysters exposed to Zn at different salinities. Symbols and error bars indicate average ± 1sd (n=3) for each salinity treatment.

523

23



524

525 526 527 528 529 530

Fig. 3. Total Zn concentrations (left panel) and δ66/64Zn values (right panel) in oysters exposed at different Zn levels for 30 days (hatched bar/triangle up: oysters before the exposure; open bars/circles: oysters exposed to different levels of Zn in water). Symbols and error bars indicate average ± 1sd (n=3) for each water Zn level.

24


Page 24 of 27

Page 25 of 27


531 532

533 534 535 536 537 538 539

Fig. 4. Plot of total Zn concentration versus δ66/64Zn value in oysters from experiments of different Zn levels exposure experiments (i.e., solid triangle up in blue, pink, green and yellow colors represent oysters exposed to water with 0 µg Zn L-1, 10 µg Zn L-1, 20 µg Zn L-1, 100 µg Zn L-1 added at 20 psu, respectively) and different salinities (i.e., open diamond, square, triangle up and circle represent oysters exposed to water at 5 psu, 12 psu, 20 psu and 28 psu, respectively with 5µg Zn L-1 added). Symbols and error bars indicate average ± 1sd (n=3).

25



540

541 542 543 544 545 546 547 548 549

Fig. 5. δ66/64Zn values in oysters versus Zn added in water from both salinity and Zn level experiments. Open triangle up represents the oysters exposed to 5 µg L-1 at 20 psu obtained as part of the salinity exposure experiment, while the triangles up in pink, green and yellow colors represent the oysters exposed to 10, 20 and 100 µg Zn L-1, respectively, at 20 psu obtained from the different Zn exposure levels experiment. Error bars indicate 1sd (n=3). Black solid line is the regression of δ66/64Zn values in oysters versus Zn added in exposure water. Blue solid line is the 95% confident interval while the red solid line is the 95% prediction interval of the regression line.

26


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Page 27 of 27


550 551

552 553 554 555 556

Fig. 6. Ternary plot of the contributions of the three Zn sources to the δ66/64Zn values in the oysters from the Zn exposure experiments. Symbols in blue, pink, green and yellow colors represent the oysters exposed to seawater at 20 psu with 0 µg Zn L-1, 10 µg Zn L-1, 20 µg Zn L-1, 100 µg Zn L-1 added, respectively.

27


Zn Isotope Fractionation in the Oyster Crassostrea hongkongensis and

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