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Ex situ determination of freely dissolved concentrations of hydrophobic organic chemicals in sediments and soils: basis for interpreting toxicity and assessing bioavailability, risks and remediation necessity

Nature Protocols volume 15, pages 1800–1828 (2020)Cite this article

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Abstract

The freely dissolved concentration (Cfree) of hydrophobic organic chemicals in sediments and soils is considered the driver behind chemical bioavailability and, ultimately, toxic effects in benthic organisms. Therefore, quantifying Cfree, although challenging, is critical when assessing risks of contamination in field and spiked sediments and soils (e.g., when judging remediation necessity or interpreting results of toxicity assays performed for chemical safety assessments). Here, we provide a state-of-the-art passive sampling protocol for determining Cfree in sediment and soil samples. It represents an international consensus procedure, developed during a recent interlaboratory comparison study. The protocol describes the selection and preconditioning of the passive sampling polymer, critical incubation system component dimensions, equilibration and equilibrium condition confirmation, quantitative sampler extraction, quality assurance/control issues and final calculations of Cfree. The full procedure requires several weeks (depending on the sampler used) because of prolonged equilibration times. However, hands-on time, excluding chemical analysis, is approximately 3 d for a set of about 15 replicated samples.

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Fig. 1: Photograph of different passive samplers.

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No datasets were generated or analyzed during the current study.

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Acknowledgements

M.T.O.J. acknowledges financial support from the European Chemical Industry Council’s Long-range Research Initiative program (Cefic-LRI), under contracts ECO22 and ECO43. F.S. acknowledges support by the Czech Ministry of Education, Youth, and Sports (LM2018121) and the European Structural and Investment Funds, Operational Program Research, Development, and Education (CZ.02.1.01/0.0/0.0/16_013/0001761). R.L. acknowledges support from SERDP ER-2538. This publication represents U.S. Environmental Protection Agency ORD-033094.

Author information

Authors and Affiliations

  1. Institute for Risk Assessment Sciences, Utrecht University, Utrecht, the Netherlands

    Michiel T. O. Jonker

  2. Atlantic Coastal Environmental Science Division, Office of Research and Development, U.S. Environmental Protection Agency, Narragansett, RI, USA

    Robert M. Burgess

  3. Department of Chemical, Biochemical, and Environmental Engineering, University of Maryland, Baltimore County, Baltimore, MD, USA

    Upal Ghosh

  4. RM Parsons Laboratory, Department of Civil and Environmental Engineering, Massachusetts Institute of Technology, Cambridge, MA, USA

    Philip M. Gschwend

  5. Geotechnics and Environment, Norwegian Geotechnical Institute, Oslo, Norway

    Sarah E. Hale

  6. Graduate School of Oceanography, University of Rhode Island, Narragansett, RI, USA

    Rainer Lohmann

  7. Center for Fisheries, Aquaculture and Aquatic Sciences, and Department of Zoology, Southern Illinois University, Carbondale, IL, USA

    Michael J. Lydy

  8. Chemistry Department, Southern California Coastal Water Research Project Authority, Costa Mesa, CA, USA

    Keith A. Maruya

  9. Civil, Environmental and Construction Engineering, Texas Tech University, Lubbock, TX, USA

    Danny Reible

  10. Research Centre for Toxic Compounds in the Environment (RECETOX), Faculty of Science, Masaryk University, Brno, Czech Republic

    Foppe Smedes

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  1. Michiel T. O. Jonker
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  4. Philip M. Gschwend
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  7. Michael J. Lydy
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  8. Keith A. Maruya
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  10. Foppe Smedes
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Contributions

M.T.O.J. wrote the manuscript. F.S. drafted Box 1 and the section on interpretation of PRC results and helped fine-tune several conceptual and methodological aspects. R.L. drafted the ‘PRC’ section in the ‘Experimental design’ section. R.M.B., U.G., P.M.G., S.E.H., M.J.L., K.A.M., and D.R. contributed to improving the manuscript by providing comments and edits.

Corresponding author

Correspondence to Michiel T. O. Jonker.

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Key reference using this protocol

Jonker, M. T. O. et al. Environ. Sci. Technol. 52, 3574–3582 (2018): https://doi.org/10.1021/acs.est.7b05752

Integrated supplementary information

Supplementary Figure 1 Weighing polymer strips.

Weighing of polymer strips is most convenient when using a (cleaned) support device (e.g., an upside down aluminum cryotube, as shown). This prevents the sampler from contacting the possibly contaminated surface of the balance and facilitates picking-up the sampler with tweezers.

Supplementary Figure 2 Cutting SPME fibers.

Cutting SPME fibers can be performed using a glass plate, underneath which a ruler is fixed. The image also shows a magnifying glass, a razor blade, a 20 mL vial for the fibers, and a fiber roll.

Supplementary Figure 3 Adding sample to the equilibration system.

Use a glass funnel placed in the mouth of either the bottle (polymer strips) or 15 mL vial (SPME fibers) when adding the sediment or soil sample.

Supplementary Figure 4 Application of aluminum foil on 120 mL bottles.

When using 120 mL equilibration bottles, apply a 5x5 cm piece of thick acetone-cleaned laboratory aluminum foil in order not to expose the sample to the plastic cap. Place the foil on the mouth of the bottle, with the dull side facing the inside of the bottle. Carefully crimp the foil around the neck, making sure that the foil touches the bottle mouth completely, showing no creases.

Supplementary Figure 5 Overall setup for collecting polymer strips.

Overview of needed materials and setup for collecting and cleaning polymer strips.

Supplementary Figure 6 Overall setup for collecting SPME fibers.

Overview of needed materials and setup for collecting and cleaning SPME fibers.

Supplementary Figure 7 Collecting a sampler from a sediment or soil suspension.

Using a tea sieve placed on a beaker is a convenient way of collecting a polymer strip or short SPME fiber from an equilibration bottle/vial.

Supplementary Figure 8 Placing polymer strips in extraction vials.

Stick the cleaned polymer strip in the mouth of the corresponding autosampler vial and cut into multiple pieces of ≤ 7 mm, such that the pieces will fit in the flat position at the bottom of the vial.

Supplementary Figure 9 Cleaning SPME fibers.

Hold the SPME fiber between the fore/middle finger and thumb and wipe with a damp tissue, held in the other hand.

Supplementary Figure 10 Cutting SPME fibers.

Stick the fiber into the mouth of the corresponding autosampler vial and cut with a clean wire cutter into pieces with such a length that these will be submerged in the solvent (e.g., ≤ 1.3 cm pieces in the case 300 µL inserts are used).

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Supplementary Figs. 1–10, Supplementary Methods and Supplementary Table 1.

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Jonker, M.T.O., Burgess, R.M., Ghosh, U. et al. Ex situ determination of freely dissolved concentrations of hydrophobic organic chemicals in sediments and soils: basis for interpreting toxicity and assessing bioavailability, risks and remediation necessity. Nat Protoc 15, 1800–1828 (2020). https://doi.org/10.1038/s41596-020-0311-y

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