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Impact of Polymer Colonization on the Fate of Organic Contaminants in Sediment Chen-Chou Wu, Lian-Jun Bao, Liang-Ying Liu, Lei Shi, Shu Tao, and Eddy Y. Zeng Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.7b03310 • Publication Date (Web): 21 Aug 2017 Downloaded from http://pubs.acs.org on August 21, 2017

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Environmental Science & Technology

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Impact of Polymer Colonization on the Fate of Organic

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Contaminants in Sediment

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Chen-Chou Wu,† Lian-Jun Bao,† Liang-Ying Liu,† Lei Shi,† Shu Tao,‡ and Eddy

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Y. Zeng*,†

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†

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and Guangdong Key Laboratory of Environmental Pollution and Health, Jinan University,

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Guangzhou 510632, China

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‡

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School of Environment, Guangzhou Key Laboratory of Environmental Exposure and Health,

Laboratory of Earth Surface Processes, College of Urban and Environmental Science,

Peking University, Beijing 100871, China

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ABSTRACT

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Plastic pellets and microbes are important constitutes in sediment, but the significance of

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microbes colonizing on plastic pellets to the environmental fate and transport of organic

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contaminants has not been adequately recognized and assessed. To address this issue, low-

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density polyethylene (LDPE), polyoxymethylene (POM) and polypropylene (PP) slices were

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preloaded with dichlorodiphenyltrichloroethanes (DDTs), polycyclic aromatic hydrocarbons

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(PAHs) and polychlorinated biphenyls (PCBs) and incubated in abiotic and biotic sediment

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microcosms. Images from scanning electron microscope, Lysogeny Broth agar plates and

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confocal laser scanning microscope indicated that all polymer slices incubated in biotic

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sediments were colonized by microorganisms, particularly the LDPE slices. The occurrence

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of biofilms induced higher dissipation rates of DDTs and PAHs from the LDPE slice surfaces

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incubated in the biotic sediments than in the abiotic sediments. Plastic colonization on LDPE

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slice surfaces enhanced the biotransformation of DDT and some PAHs in both marine and 1 ACS Paragon Plus Environment


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river sediments, but had little impact on PCBs. By comparison, PP and POM with unique

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properties were shown to exert different impacts on the physical and microbial activities as

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compared to LDPE. These results clearly demonstrated that the significance of polymer

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surface affiliated microbes to the environmental fate and behavior of organic contaminants

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should be recognized.

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2 ACS Paragon Plus Environment

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INTRODUCTION Plastic resin pellets, a group of man-made materials, have been ubiquitously identified

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in a variety of marine settings, ranging from populated areas to the shores of remote

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uninhabited regions.1, 2 These pellets can act as substrates for accumulating/leaching organic

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materials from/to surrounding marine environments and as platforms for microbial adhesion,

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and therefore may provide a pathway for transport of microbes and organic pollutants.3-7

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Recently we postulated that marine debris could mediate the distribution patterns of organic

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pollutants in coastal sediment.8, 9 It is also well known that any surface exposed to seawater

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would be colonized by micro-organisms, forming a thin layer of life (biofilm).5, 10 Recent

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studies indicated that the plastisphere microbes (the microbial community embedded in the

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surface of plastics) are distinct from those in the surrounding seawater and sediment.5, 10 In

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addition, some bacterium species have been demonstrated to hydrolyze plastics as the source

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of energy for growth,11, 12 although most plastics are chemically inert and resistant to

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biodegradation. However, the environmental fate and behavior of organic contaminants

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governed by the interaction between plastics and microbes (or plastic-attached microbial

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communities) have yet to be clarified,13 particularly for what occurs on the surface of plastics.

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Plastic strips, such as low-density polyethylene (LDPE) and polyoxymethylene (POM)

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sheets, have been increasingly used as passive samplers’ sorbent phases for measuring

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organic pollutants in water, sediment and air, largely thanks to their affinity and sorption

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capacity for organics.14-16 Performance reference compounds (PRCs) have been used to

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correct biofouling effects from colonization by microorganisms during field deployment,14-16

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which requires further verification by additional experimental data.17 Particularly, the

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fundamentals for the use of PRCs are related to isotropic exchange,18 but many PRCs may

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not dissipate at all during deployments. For example, Fernandez et al.19 reported that the

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dissipation rate of pre-loaded 13C-labeled p,p'-dichlorodiphenyldichloroethane (p,p'-DDD) in



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water-side LDPE deployed in the Palos Verdes Shelf was lower than that of 13C-labeled p,p'-

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dichlorodiphenyltrichloroethane (p,p'-DDT), whereas the dissipation rates were reversed in

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LDPEs exposed in sediment beds. The sediment bed-side dissipation processes were

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inconsistent with the exchange kinetics for the uptake of these compounds by passive

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sampling devices,18 because DDT has a larger polymer-water partition coefficient than

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DDD.19

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The primary goal of the present study was to investigate the sorption/desorption

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behavior of organic compounds linked to the interaction between three synthetic polymers,

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i.e., LDPE, POM and polypropylene (PP) slices, and microbes. Among the selected plastics,

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LDPE and PP are commonly encountered in the marine environment,20 while LDPE and

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POM are frequently selected as the sorbent phases in passive samplers.19, 21 The target

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analytes include dichlorodiphenyltrichloroethanes (DDTs), polycyclic aromatic hydrocarbons

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(PAHs) and polychlorinated biphenyls (PCBs), which are commonly detected on marine

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plastic debris.6, 7 The polymer slices were pre-loaded with the target analytes and immersed

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in abiotic and biotic sediments to prepare incubation microcosms (brown glass jars).

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Potential impacts of microbes on the dissipation rates of pre-loaded target compounds from

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incubated polymer slices were examined. Finally, the biofouling effects on field applications

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of passive samplers with the use of PRCs were also assessed.

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MATERIALS AND METHODS

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Materials. Authentic standards of 28 PAHs (Supporting Information (SI) List S1) and

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benzo[ghi]perylene-d12, five PCBs, i.e., PCB-18, 61, 67, 82 and 191 were purchased from

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AccuStandard (New Haven, CT). Three isotopically labeled PAHs, i.e., anthracene-d10,

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benzo[a]anthracene-d12 and benzo[a]pyrene-d12, and eight 13C-labeled PCBs congeners, i.e.,

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C-PCB-1, 15, 28, 47, 101, 153, 180 and 202, were obtained from Cambridge Isotope


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Laboratories (Andover, MA). 2-Fluorobiphenyl, p-terphenyl-d14, dibenzo[a,h]anthrancene-

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d14, naphthalene-d8, acenaphthene-d10, phenanthrene-d10, chrysene-d12 and perylene-d12 were

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purchased from Dr. Ehrenstorfer GmbH (Augsburg, Germany). Four deuterated DDTs, p,p'-

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DDT-d8, o,p'-DDT-d8, p,p'-DDD-d8 and p,p'-DDE-d8, were purchased from C/D/N Isotopes

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(Quebec, Canada).

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Low-density polyethylene (50-µm film thickness), POM (76-µm film thickness) and

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PP-type disposable food storage containers (370-µm thickness) were purchased from TRM

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Manufacturing (Corona, CA), CS Hyde Company (Lake Villa, IL) and a local supermarket,

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respectively. Before use, LDPE, POM and PP-type food containers were cut into ~ 25 mm ×

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25 mm slices, which were precleaned by dialysis in hexane twice over 48 h. The precleaned

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LDPE, POM and PP slices were immersed in water:methanol (50:50 in volume) solution

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spiked with various arrays of target compounds until use (at least for two weeks).

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Anthracene-d10 was spiked at 30 µg L−1 and all other target compounds were spiked at 3 µg

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L−1. Two loaded polymer slices were processed to determine the initial concentration of each

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pre-loaded compound, and another two unloaded slices were used as laboratory blanks to

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monitor any possible external contamination during sample processing.

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Static Microcosm Incubation. Two Hailing Bay marine sediments from sites 1 and

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16, respectively, collected in a previous study9 and one East River freshwater sediment22

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were used for microcosm incubation. Hailing Bay and East River sediments are known to

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contain abundant DDTs9 and PAHs22, respectively. The sediments were sieved, pre-

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homogenized and stored until use. Approximately 50 g of each wet sediment was added to a

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100-mL microcosm containing 30 g of Milli-Q Millipore purified water, and the slurries

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mixed by gentle agitation with a glass rod for approximately 2 min. The content in the

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microcosm was divided into two portions (abiotic and biotic sediments). The abiotic

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sediment, which also served as a control, was mixed with approximately 1 mg of sodium



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azide and sterilized by autoclave twice (2-week intervals) to inhibit bacterial activity.23 The

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biotic sediment was not subject to any treatment. One pre-loaded polymer slice was added to

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each biotic or abiotic sediment microcosm. All the microcosms were incubated at room

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temperature (25 oC) and under ambient conditions. One polymer slice pre-loaded with

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different compounds was harvested each time.

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The first batch of LDPE slices was pre-loaded with either p,p'-DDT-d8 or p,p'-DDD-d8

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and p,p'-DDT-d8 for the coastal marine sediment and PCB-61, anthracene-d10,

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benzo[a]anthracene-d12 and benzo[a]pyrene-d12 for the river sediment. Incubated LDPE

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slices were collected from the coastal marine sediment after 10, 20 and 30 days and from the

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river sediment after 7, 20 and 40 days (SI Table S2). To further examine the possibility of

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biotransformation/biodegradation of the target compounds during incubation, the second

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batch of LDPE slices was preloaded separately with p,p'-DDT-d8, p,p'-DDD-d8, p,p'-DDE-d8,

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p,p'-DDT-d8 and p,p'-DDD-d8, p,p'-DDT-d8 and p,p'-DDE-d8, and o,p'-DDT-d8 and p,p'-

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DDD-d8 for coastal marine sediment, and p,p'-DDT-d8 and p,p'-DDD-d8 also separately for

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river sediment. Loaded LDPE slices were retrieved upon 20 days of incubation (SI Table

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S2). The third batch contained POM and PP slices loaded with p,p'-DDD-d8 and p,p'-DDT-

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d8, or 13C-p,p'-DDT (only for the POM slice) for coastal marine sediment and LDPE slices

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loaded with PCB 61 and eight 13C-labeled PCB congeners or naphthalene-d8, acenaphthene-

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d10, fluorene-d10, phenanthrene-d10, pyrene-d10, chrysene-d12, perylene-d12 and

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benzo[ghi]perylene-d12 for river sediment (Table S2). For the first and second batches of

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plastic slices, 2-fluorobiphenyl, p-terphenyl-d14, dibenzo[a,h]anthrancene-d14 and PCB-82

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were used as internal standards and PCB-67, PCB-191, naphthalene-d8, acenaphthene-d10,

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phenanthrene-d10, chrysene-d12, perylene-d12 and benzo[ghi]perylene-d12 were used as

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surrogate standards. 2-Fluorobiphenyl, p-terphenyl-d14 and PCB 82 were used as internal


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standards and PCB-18, PCB-67, PCB-191, anthracene-d10 and benzo[a]anthracene-d12 were

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used as surrogate standards for the third batch of samples.

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Sample Characterization. To identify any microbial activity on polymer surfaces,

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LDPE, PP and POM slices from biotic river sediment (SI Figures S1a, S1b, S1f and S1g),

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LDPE slice from biotic (SI Figures S1c and S1h) and abiotic (SI Figures S1d and S1i) marine

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sediment and procedural blank (SI Figures S1e and S1j) after 20-day exposure were

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randomly selected. All the polymer slices were cut and fixed on copper stubs with double

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coated carbon conductive tabs. Samples were subsequently coated with platinum and imaged

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by a Zeiss Ultra 55 scanning electron microscope (SEM, Oberkochen, Germany). To further

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visualize the adherent microbes and microbial colonization on polymer surfaces, the polymer

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slices upon 10-day culturing were randomly selected, cut and rinsed with sterile phosphate

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buffered saline (Life Technology, Grand Island, NY). The traditional Lysogeny Broth agar

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plates were further used to incubate microbes adhered on polymer surfaces for 48 h at 30 ºC

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aerobically. Coatings of extracellular polymeric substance, one of the principal structural

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components of biofilms, on the surfaces of plastic strips were visualized by staining with

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fluorescent wheat germ agglutinin conjugate.24 Briefly, the polymer slices were fixed with

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4% paraformaldehyde for 1 h at 30 oC. Samples were subsequently washed once with sterile

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phosphate buffered saline and stained with 5 µg mL-1 wheat germ agglutinin Alexa Fluor 647

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fluorescent conjugate solution (Molecular Probes, Eugene, OR) and incubated for 1 h at 30 oC

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in the dark. Upon removal of unbound dyes by rinsing with sterile phosphate buffered saline,

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the samples with extracellular polymeric substance coatings were imaged by a confocal laser

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scanning microscope (LSM 700; Carl Zeiss, Germany).

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An aliquot of each sediment sample was acidified with 10% HCl overnight to remove carbonate. The samples were subsequently washed with purified water to remove chlorine



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ion and dried at 60 °C to determine TOC contents by an Element Analyzer Vario EL III

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(Elementar, Hanau, Germany).

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Sample Extraction. Prior to extraction, loaded polymer slices were rinsed with water

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and wiped with filter paper. They were cut to small pieces (∼5 × 5 mm), wrapped with filter

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paper, spiked with the surrogate standards and extracted in hexane (soaking) twice over 24 h.

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Each extract was dried with sodium sulfate, concentrated to 1 mL with a Zymark TurboVap

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500 (Hopkinton, MA) and spiked with the internal standards before instrumental analysis.

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Detailed procedures for instrumental analysis and quality assurance and quality control

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results are provided in SI Text S1.

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RESULTS AND DISCUSSION

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Characteristics of Surface of Polymer Slices. After 20-day incubation, the surface of

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LDPE slices exposed to biotic river and marine sediments became rough (SI Figures S1a, S1c

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and S1h), while the surfaces of all other samples remained smooth (SI Figures S1d and S1i).

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The Lysogeny Broth agar plates for the plastic strips from the biotic sediments were

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colonized by microbes (Figures 1a-1c), i.e., the biotic sediments indeed produced biofilms on

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the plastic strips. By comparison, the Lysogeny Broth agar plates for the plastic strips from

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the sterilized sediments remained smooth and were not colonized (Figures 1d-1f), indicating

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the effectiveness of abiotic treatments. The extracellular polymeric substance coatings only

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occurred on the slices from the biotic sediments (SI Figures S2a-S2c); particularly, LDPE

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slices (SI Figure S2a) were more colorful than PP and POM slices (SI Figures S2b and S2c).

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Among the polymer slices under investigation, POM was less susceptible to biofouling

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probably because of its smoother and denser surfaces as compared to PP and LDPE, which

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are manufactured with rough or sticky surfaces.25 All microscopic analyses suggested that


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the surfaces of polymer slices could be colonized by microorganisms, particularly the LDPE

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slices forming biofilm rapidly.

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The Retained Fractions of Pre-Loaded Target Analytes from Plastics Under

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Abiotic Conditions. The retained fractions of pre-loaded DDTs, PAHs and PCBs from

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LDPE slices after incubation were controlled by the target compounds’ physicochemical

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properties and sediment TOC. For example, the retained fractions of pre-loaded p,p'-DDD-

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d8, p,p'-DDT-d8, o,p'-DDT-d8 and p,p'-DDE-d8 in marine sediments from sites 1 and 16 were

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positively related to log Kow or log Kpew (5.08 for p,p'-DDD; 5.82 for p,p'-DDT; 6.00 for o,p'-

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DDT and 6.20 for p,p'-DDE; SI Table S1) (Figure 2a). For the river sediment, the measured

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losses of pre-loaded PAHs and PCBs after 7-, 20- and 40-day incubation of LDPE slices

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under abiotic conditions followed diffusion-mediated patterns (SI Figure S3). In addition, the

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retained fractions of p,p'-DDD-d8 and o,p'-DDT-d8 at site 1 were significantly different (p

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