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Chemical Engineering Journal 404 (2021) 127079

Available online 23 September 2020 1385-8947/© 2020 Elsevier B.V. All rights reserved.

Engineering a microbial ‘trap and release’ mechanism for microplastics removal

Sylvia Yang Liu a, Matthew Ming-Lok Leung a, James Kar-Hei Fang a, b, *, Song Lin Chua a, c, *

a Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Kowloon, Hong Kong Special Administrative Region b Food Safety and Technology Research Centre, The Hong Kong Polytechnic University, Kowloon, Hong Kong Special Administrative Region c State Key Laboratory of Chemical Biology and Drug Discovery, The Hong Kong Polytechnic University, Kowloon, Hong Kong Special Administrative Region

A R T I C L E I N F O

Keywords: Microplastics Pseudomonas aeruginosa Biofilms Bioaccumulation Exopolymeric substances

A B S T R A C T

Plastics are discarded and accumulated in the environment at an alarming rate. However, their resistance to biodegradation allows them to persist in the environment for prolonged durations. While large plastics are easier to remove, microplastic particles from cosmetics or fragments from larger pieces are extremely difficult to remove from the environment. Furthermore, current techniques such as filters poorly retain microplastics or require harsh chemical treatments in wastewater treatment plants. Hence, microplastics enter the natural environment easily even after effluent treatments, thereby endangering aquatic life and humans who consume seafood. It is imperative to develop sustainable bioaggregation processes to trap microplastics quickly for easier removal from the environment. Here, we showed that microplastics can be trapped and aggregated in the sticky exopolymeric substances (EPS) produced by biofilms. As a proof-of-concept, we engineered a bacterial biofilm with a ‘capture-release mechanism’, whose EPS can first cause bioaggregation of microplastics for easier isola- tion, followed by an inducible biofilm dispersal mechanism that releases trapped microplastics for downstream resource recovery. We also demonstrated the potential application of the engineered biofilm in mitigating microplastics pollution in seawater samples collected in the vicinity of a sewage outfall. This capture-and-release approach should prove widely applicable to other micropollutants or biofilm-enabled catalysis.

1. Introduction

Pollution caused by plastics, especially microplastics, is a major environmental concern as the world becomes increasingly industrial- ized. Microplastics are synthetic hydrocarbon-based particles with a size range between 1 µm and 5 mm, with diverse sources from cosmetics, synthetic textile, packaging and broken pieces of larger plastic items. Their recalcitrance to biodegradation allows microplastics to persist in the natural environment, especially in water bodies. These pollutants can pass through even the most efficient water filtration systems and end up being released into water bodies. Furthermore, the highly varied composition, size and pollutants attached to the surface of microplastics pose a multitude of problems to the biota. Hence, the effects of micro- plastics on organisms could be dire, where humans or animals down the food chain could ingest them or suffer from toxic pollutants adsorbed to the microplastics [1-3].

There are multiple challenges to be addressed in the removal of

microplastics from waste and the polluted environment. Firstly, com- mon microplastics such as polyethylene (PE), polypropylene (PP) and polystyrene (PS) are positively or neutrally buoyant that exist as dispersed or suspended solid particles, making meaningful isolation for separate disposal difficult. Even with the use of harsh chemical treat- ments or expensive filters [4] in wastewater treatment processes, a significant amount of microplastics remains in the effluent, rendering wastewater treatment plants as a main source of microplastics pollution [5-7]. Next, bioremediation of plastics is currently limited to specific enzymes such as the PETase which could degrade polyethylene tere- phthalate (PET) [8], but is inefficient in removing a mixture of micro- plastics which comprise various polymer types and sizes. Lastly, incineration of plastic debris such as polyvinyl chloride (PVC) together with other waste can emit toxic dioxins [9]. Hence, PVC recovery and recycling processes are assessed to be environmentally friendlier than primary PVC production [10]. The European Commission had set new rules on PVC waste recovery and reuse in construction projects in its

* Corresponding authors at: Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Kowloon, Hong Kong Special Administrative Region.

E-mail addresses: [email protected] (J.K.-H. Fang), [email protected] (S.L. Chua).

Contents lists available at ScienceDirect

Chemical Engineering Journal

journal homepage: www.elsevier.com/locate/cej

https://doi.org/10.1016/j.cej.2020.127079 Received 27 August 2020; Received in revised form 30 August 2020; Accepted 16 September 2020

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Waste Framework Directive [11]. Bioremediation strategies have the potential to address the challenge

of microplastics contamination, but the biological processes to degrade microplastics may take a long time. There is a paucity of studies on the interactions of biological materials on microplastics. Environmental microbes could form multicellular biofilms with their self-produced exopolymeric matrix on microplastics, often with altered diversity, metabolism and function [12,13], allowing microbes to colonize on the microplastic surfaces. Metagenomic studies had been conducted to identify microbes that colonize and grow biofilms on microplastics, with the Pseudomonad genus prevalently isolated [13-16]. Furthermore, microplastics could aggregate with marine biogenic particles comprising live or dead biomatter and sink into the deep-sea sediment [17].

Biofilm formation and dispersal in many bacterial species are controlled by the intracellular c-di-GMP secondary messenger signaling. Typically, synthesis of c-di-GMP by GGDEF-containing diguanylate cy- clases (DGCs) leading to high c-di-GMP levels will promote biofilm formation, while degradation of c-di-GMP by EAL/HYP-containing phosphodiesterases (PDEs) lead to biofilm dispersal [18]. Many bacte- rial species contain multiple DGCs and PDEs, reflecting their redun- dancy is key to fine-tune metabolism and biofilm formation for survival [19]. One example of c-di-GMP signaling is the wsp chemosensory pathway in Pseudomonas aeruginosa, where WspR is a DGC involved in c- di-GMP synthesis and production of exopolysaccharides for biofilm formation [20]. The WspA is involved in sensing surfaces, leading to autophosphorylation of WspE, which in turn phosphorylates and acti- vates downstream WspR. Deletion of the wspF methylesterase gene can cause overmethylation of WspA, leading to constitutive activation of WspR and enhanced biofilm formation [20].

One potential way to remove microplastics is through bio- aggregation, which has been utilized in the gradual accumulation of toxic substances such as pesticides or metalloids by a microbe from the polluted environment [21,22]. Furthermore, convenient recovery of plastics from bioaggregation may promote the recycling of retrieved plastics, instead of choosing landfills or incineration. It is estimated that nearly 80% of plastics ended up in landfills and 12% was incinerated, but a meagre 9% was recycled [23]. Hence, recycling was ranked as the preferred choice over incineration and landfills [24], warranting the need for improved recycling efforts.

As a proof-of-concept, we engineered a ubiquitous environmental bacterium, P. aeruginosa biofilm which can efficiently aggregate microplastics within its sticky matrix and be later induced to release trapped microplastics for convenient downstream retrieval and recy- cling. This ‘trap-and-release’ bioaggregation strategy has several ad- vantages: firstly, microplastics can be aggregated irrespective of their material, size or composition, thereby circumventing the specificity issue. Next, microplastics are localized at high concentrations and can be cumulatively removed from the environment at ease. The increased total mass will promote easier removal by filtering or sedimentation in tanks. Finally, to release microplastics from the sticky exopolymeric matrix of biofilms, treatment with an inducible stimulus could disperse biofilms and release the microplastics for convenient retrieval.

2. Materials and Methods

2.1. Bacterial strains and growth conditions

The bacterial strains and plasmids used in this study are listed in Table 1. E. coli DH5a strain was used for standard DNA manipulations. For bacterial growth, LB medium was used to cultivate E. coli and P. aeruginosa strains. For experiment, P. aeruginosa strains were grown in ABTGC (ABT minimal medium supplemented with 2 g L− 1 glucose and 2 g L− 1casamino acids) [18]; artificial seawater (Instant Ocean Reef Crystals, USA) supplemented with 1 µM FeCl3, 2 g L

− 1 glucose and 2 g L− 1casamino acids; or freshwater supplemented with 1 µM FeCl3, 2 g L

− 1

glucose and 2 g L− 1casamino acids. Both glucose and amino acids were

added at environmentally-relevant levels as carbon and nitrogen sources respectively to simulate the presence of organic matter in seawater and freshwater, thus such media were routinely used in experimental studies of environmental microbes and bioremediation [25-29]. For plasmid maintenance in E. coli, the medium was supplemented with 100 μg ml− 1 ampicillin and 15 μg ml− 1 gentamicin. For marker selection in P. aeruginosa, 30 μg ml− 1 gentamicin, were used, as appropriate.

2.2. Preparation of microplastics

Microplastics of PET, polymethyl methacrylate (PMMA), nylon 6/6 and PVC were made using a Retsch CryoMill cryogenic grinder (Haan, Germany). The precooling stage lasted 7 min at 5 shakes s− 1, followed by the grinding stage for 1.5 min at 25 shakes s− 1 at –196 ◦C. The ultra- low temperature was maintain by liquid nitrogen circulated outside the grinding chamber made of zirconium oxide. To collect microplastics (<100 µm), the grinded plastics were sieved through a filter (pore size = 100 µm) before use.

2.3. Growth of biofilms on microplastics

Experimental cultivation of P. aeruginosa strains was carried out in ABTGC (ABT minimal medium supplemented with 2 g L− 1 glucose and 2 g L− 1 casamino acids) [37], artificial seawater (Instant Ocean Reef Crystals) supplemented with 2 g L− 1 glucose and 2 g L− 1 casamino acids) or freshwater supplemented with 2 g L− 1 glucose and 2 g L− 1 casamino acids).

Microplastics of varying materials and sizes were tested: PVC, nylon, PS, PMMA andPET, at 106–300 µm or < 106 µm. Bacterial cultures were grown to exponential phase in LB for 4 hrs and then diluted 1000-fold for experimental growth in 10 ml medium containing 1, 3 or 5 µg ml− 1 of microplastics in 50-ml tubes at 25 ◦C or 30 ◦C, shaken at 200 rpm, for 24 hrs until sample processing.

Table 1 Bacterial strains used in this study.

Strain/ plasmid Description Source/ Reference

P. aeruginosa PAO1 Prototypic nonmucoid wild-type strain [30] PAO1/plac-gfp Gm

r; PAO1 containing the Tn7-plac-gfp vector

[31]

PAO1/pcdrA-gfp Gm r; PAO1 containing the pcdrA-gfp vector [32]

ΔwspR wspR knockout of PAO1 constructed by allelic exchange

This study

ΔwspF wspF knockout of PAO1 constructed by allelic exchange

[33]

ΔwspF/pBAD-yhjH Gm r; ΔwspF containing the pBAD-yhjH

plasmid This study

PAO1/plac-yedQ/ pBAD-yhjH

Gmr and Carbr; OUS82 containing the plac- yedQ and pBAD-yhjH insertion vector

This study

P. putida OUS82 Prototypic nonmucoid wild-type strain [34] OUS82/plac-

yedQ/pBAD- yhjH

Gmr and Carbr; OUS82 containing the plac- yedQ and pBAD-yhjH insertion vector

This study

E. coli DH5α F–, ø80dlacZΔM15, Δ(lacZYA-argF)U169,

deoR, recA1, endA1, hsdR17(rK–, mK+), phoA, supE44, λ–, thi-1, gyrA96, relA1

Laboratory collection

Plasmid pJN105 Gmr; broad-host-range vector carrying

the araBAD promoter [35]

pBAD-yhjH Gm r; pUCP22 carrying the yhjH gene [36]

plac-yedQ Carb r; pUCP18 carrying the yhjH gene This study

Tn7-plac-gfp Apr Gm r; tn7 transposon vector carrying the

plac-gfp fusion [31]

pcdrA-gfp Apr Gm r; pUCP22 carrying the pcdrA-gfp

fusion [32]

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2.4. Quantification of dry weight mass of microplastics

Biofilms which accumulated microplastics were disrupted by vor- texing and bacterial cells were lysed by sterile ddH2O + 1% Triton-X (v/ v). The microplastics were pelleted by brief centrifugation and the su- pernatant with bacterial lysate were discarded. The microplastics were washed twice with ddH2O to remove any bacterial remnants and dried at 40 ◦C for 8 hrs. The microplastics were weighed on the analytical bal- ance with accuracy of ± 0.0001g. Experiments were performed in triplicate, and the results are shown as the mean ± s.d.

2.5. Quantification of bacterial numbers by colony-forming units (CFU)

Biofilm cells were homogenized by vigorous vortexing in 0.9% (w/v) NaCl saline solution and the cell suspension was diluted serially in 0.9% (w/v) NaCl solution. The diluted samples were then transferred to LB agar plates in 5 replicates and incubated for 16 hrs at 37 ◦C. The colonies grown on the agar plate were then enumerated, with the CFU ml− 1

tabulated by colony number X dilution factor X volume. Experiments were performed in triplicate, and the results are shown as the mean ± s. d.

2.6. Scanning electron microscopy (SEM) sample preparation and image acquisition

As previously described [38], the PAO1 and ΔwspF biofilms which contained PVC microplastics were fixed on holders by 0.1 M PBS buffer + 5% glutaraldehyde for 4 hrs. Next, the samples were dehydrated through ethanol series, 35% (v/v), 50% (v/v), 75% (v/v), 95% (v/v) once for 10 mins and absolute ethanol (greater than99% v/v) for 10 mins twice, followed by drying at room temperature for 16 hrs. Before the scanning, the samples were sputtered by gold particles (Nanoimages, MCM-200) and wiped with the conductive glue on the holders’ corners. Microscopy images were captured and acquired by using Tescan VEGA3 Scanning electron microscopy (voltage = 20 kv and the magnification = 500 × to 15000 × ). Experiments were performed in triplicate, and representative image was presented.

2.7. Epifluorescence imaging of biofilms and microplastics

As described in the previous section, PAO1/plac-gfp or PAO1/pcdrA- gfp strains were grown in 1 mg ml− 1 PVC (size < 106 μm, gray for easier observation). The biofilms which contained the microplastics were transferred carefully to an 8-well chamber (μSlide, ibiTreat, Ibidi, Ger- many). As control, planktonic PAO1/pcdrA-gfp which did not trap any PVCs were directly placed into the 8-well chamber. All microscopy images were captured and acquired by using Nikon Eclipse Ti2-E Live- cell Fluorescence Imaging System with 40 × objectives through two channels and Z-stack project at bright field and GFP fluorescence field. At least 5 images were captured for every replicate well. All the images were exported by the NIS (Nikon) program. Experiments were per- formed in triplicate, and representative image was presented.

2.8. Quantification of c-di-GMP levels in bacterial biofilms

As previously described [39], c-di-GMP were extracted from PAO1 biofilms containing trapped PVC (size < 106 μm). Briefly, samples were centrifuged at 16,000 g for 2 min at 4 ℃, and washed three times by ice- cold PBS buffer. The cell pellet was resuspended in ice-cold PBS buffer and were immediately transferred to incubator at 100℃ for 3 min. Ice- cold ethanol was then added into the sample (final concentration = 65% (v/v)) and the sample was vortexed for 15 secs. The treated samples were centrifuged at 16,000 g for 2 min, 4 ℃), and the supernatant containing extracted c-di-GMP was collected. The samples were then lyophilized and resuspended in 50 μl ddH2O. The c-di-GMP in each sample was quantified by using the c-di-GMP ELISA kit (LMAI,

Shanghai) per manufacturer’s instructions and measuring OD450nm with the microplate reader (Tecan, Infinite M1000 Pro). The c-di-GMP con- centration was normalized by protein concentration, where protein concentration (OD280nm) was measured by Nanodrop (Thermofisher, NanoDrop One, ND-One-W).

2.9. Extraction and Quantification of exopolysaccharides

As previously described [40], PAO1 planktonic cells and biofilms were collected from bacterial cultures grown in ABTGC + 1 µg ml− 1 grey PVC at 30 ◦C for 24 hrs. The PVC-containing biofilms were decanted and separated from planktonic cells in supernatant by sedimentation for 10 mins. The PVC-containing biofilms were re-suspended in 0.9% (w/v) NaCl and treated with mild water-bath sonication (Elmasonic P120H, Power = 50%, Frequency = 37 KHz, 5 mins) to separate the cells and PVC from the surface-associated matrix. The cells were then separated from the matrix by centrifugation, leaving behind the crude matrix extract.

The crude extract was then further treated by removal of eDNA by precipitation with 25% (v/v) ethanol and 0.1 M CaCl2. Extracellular proteins were then removed from the extract with 0.5 mg ml− 1 pro- teinase K at 60 ◦C for 1 hr and inactivation at 80 ◦C for 30 mins. The extract was then filtered with centrifugal filter (<3 kDa) to remove the metabolites. The extract was then lyophilized and re-suspended in sterile ddH2O. To quantify exopolysaccharide concentration, the phe- nol–chloroform assay was used and OD420nm was measured by microplate reader Infinite Pro (Tecan, Denmark). . Experiments were performed in triplicate, and the results are shown as the mean ± s.d.

2.10. Screening of DGC/PDE mutant library

As described in the previous section, DGC/PDE mutants from the Seattle Transposon Library [39] were grown in 10 ml ABTGC + 1 mg ml− 1 PVC (size < 106 μm, gray for easier observation) at 37 ℃, shaking at 200 rpm for 24 hrs. The biofilms and microplastics were transferred carefully into microscopic dishes (35 mm diameter). At least 5 micro- scopy images of each mutant were captured and acquired by stereomi- croscope (Nikon SMZ1270i, Japan) at 15 × magnification. Experiments were performed in duplicate, and representative image was presented.

2.11. Preparation of seawater samples

Surface seawater was collected from Kwun Tong, Hong Kong (22◦18′30′′N; 114◦13′11′′E) close to a sewage outfall, a sampling site which had been identified as a hotspot of microplastics [7]. Collected seawater (150 L) was sieved through 300 µm and 106 µm. Retained materials including microplastics (106–300 µm) were resuspended and concentrated in 1 L of seawater, which remained undisturbed for 3 hrs to precipitate and discard sand particles. The water column containing microplastics was used here and was well mixed and divided into three seawater samples as replicates to evaluate our bioaggregation approach. The amounts of microplastics in these samples, before and after the bacterial treatments described above, were determined using Raman microspectrometry.

2.12. Raman microspectroscopy of microplastics

Microplastics in each seawater sample were retrieved on a stainless- steel filter membrane (31 µm pore size) and assessed by a Renishaw inVia confocal Raman microspectrometer (Wotton-under Edge, En- gland) using 785 nm excitation at 10% laser power for 5 s to acquire Raman spectra (675–1767 cm− 1). The whole area coated with micro- plastics (8 mm in diameter) was scanned and mapped at a spatial res- olution of 28.4 µm to yield two-dimensional and colour-coded illustration of Raman spectra, to facilitate identification of polymer types as well as sizes and shapes of the microplastics. Baseline correction

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and smoothing of the acquired spectra were performed with the Renishaw WiRE 4.2 software. The polymer types of microplastics were identified using the Renishaw Polymeric Materials Database.

2.13. Statistical analysis

All experiments were performed in triplicates. Averages, standard deviations, and independent-group t-tests were carried out in Microsoft Excel. One-way ANOVA, followed by Tukey’s multiple comparison tests if required, were carried out in Graphpad Prism.

3. Results:

3.1. P. aeruginosa can form biofilms to bioaggregate microplastics

We first asked if microbes can perform the function of accumulating microplastics by testing the ability of P. aeruginosa to accumulate microplastics. The planktonic cells were grown with microplastics in suspension, where biofilms could start colonizing on microplastics and produce exopolymeric matrix that embed microplastic particles. We first tested a variety of microplastics of various sizes and materials. By quantifying the microplastic mass trapped in biofilms and measuring the biofilm cell number via CFU, we showed that P. aeruginosa could accu- mulate large microplastics (106–300 µm) and small microplastics (<106 µm) of varying materials at 24 hrs (Supplementary Fig. 1a-c).

Fig. 1. P. aeruginosa can form biofilms on microplastics. (a) Image of P. aeruginosa biofilm accumulating PVC (<106 µm) into a bolus (shown with black bold arrow). Scale bar (bottom right): 1 cm. (b) Mass of microplastics accumulated and lost by bio- film. (c) CFU of bacteria within and outside of biofilm. (d) Representative image of P. aeruginosa PAO1 biofilm cells on PVC particles (Scale bar: 50 µm). (e) Represen- tative electron microscopy image of P. aeruginosa biofilm cells on PVC particles (Scale bar: 1 nm). Means and s.d. from triplicate experiments are shown.

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Since bioaccumulation of microplastics by P. aeruginosa can be applied to all tested plastic types, we focused on using PVC for our downstream applications. P. aeruginosa could trap PVCs of < 106 µm diameter (grey color for easier observation) into a bolus-like aggregate (Fig. 1a). Its efficiency to trap PVCs improved over time (0, 3, 6, 12 and 24 hrs) where nearly all microplastics were accumulated at 24 hrs, so that dispersed microplastics remaining in free suspension would be reduced (Fig. 1b). The mass of trapped microplastics was also correlated to bacterial numbers (Fig. 1c), implying that P. aeruginosa biofilm was

growing and entrapping microplastics. Further examination of the ag- gregates using CLSM and SEM revealed that bacterial populations localized directly on microplastics (Fig. 1d and 1e).

3.2. C-di-GMP signaling is key to biofilm accumulation of microplastics

We next showed that P. aeruginosa employed c-di-GMP-mediated biofilm formation to bioaccumulate PVCs. This was reflected by direct quantification of c-di-GMP by ELISA (Fig. 2a) and observation of

Fig. 2. C-di-GMP signaling is key to biofilm accumulation of microplastics. (a) Quantification of c-di-GMP levels in microplastics-accumulated biofilms by ELISA. (b) Representative image of PAO1/pcdrA-gfp biofilms and planktonic bacteria on microplastics. (Scale bar: 50 µm). (c) Quantification of exopolysaccharides in microplastics-accumulated biofilms. (d) Mass of microplastics accumulated and lost by PAO1 and ΔwspR biofilm. (e) CFU of PAO1 and ΔwspR within and outside of biofilm. Means and s.d. from triplicate experiments are shown. **P < 0.01, One-Way ANOVA.

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biofilms containing a GFP-based biosensor (pcdrA-gfp reporter) whose GFP expression correlated to c-di-GMP expression [32] (Fig. 2b). Accumulation of microplastics was attributed to the production of biofilm-associated exopolysaccharides, where we observed higher polysaccharide levels in microplastic-containing biofilms than in planktonic cells from the remaining media (Fig. 2c). Mutations of exo- polysaccharides production in the ΔpelAΔpslBCD mutant [41] could lead to near-complete loss of microplastics-accumulating biofilms, confirming that exopolysaccharides are key to accumulating micro- plastics (Supplementary Fig. 2).

With the aim of enhancing the microplastics bioaccumulation pro- cess, we aimed to develop a P. aeruginosa strain with a propensity to

form biofilms and accumulate microplastics. We first screened an in- house mutant library of DGCs to identify which gene is important in PVC-mediated biofilm formation (Supplementary Fig. 3) and identified that a few mutants, especially ΔwspR (PA3702) mutant which could not accumulate microplastics. The ΔwspR mutant formed significant lesser biofilms (Fig. 2d) and accumulated lesser microplastics (Fig. 2e) as compared to the wild-type PAO1.

3.3. Overexpression of wsp operon can boost accumulation of microplastics by biofilms

To improve the capture efficiency of P. aeruginosa biofilms, we

Fig. 3. Overexpression of wsp operon can boost accumulation of microplastics by biofilms. (a) Representative electron microscopy image of ΔwspF biofilm cells on PVC particles. (b) Bacterial CFU over time. (c) Mass of microplastics accumulated by biofilms over time. (d) ΔwspF can accumulate microplastics (3 and 5 mg ml− 1) at higher efficiency than PAO1. (e) CFU of ΔwspF biofilms on microplastics (3 and 5 mg ml− 1) is higher than PAO1. Means and s.d. from triplicate experiments are shown. ***P < 0.001, One-Way ANOVA.

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engineered the expression of wsp operon in P. aeruginosa by using the ΔwspF mutant. We observed that the ΔwspF mutant could grow more biofilms (Fig. 3a-b) incorporate higher concentrations of PVC micro- plastics faster than wild-type PAO1 (Fig. 3c). By 24 hrs, the ΔwspF mutant could accumulate higher mass of PVCs than PAO1. We also tested the ability of the engineered strain to accumulate high levels of PVCs (3 and 5 mg ml− 1) and found that it could form biofilm of sufficient size to accumulate more than 90% of PVCs (Fig. 3d-e). Furthermore, the ΔwspF mutant could accumulate low-density floating microplastics (polystyrene), resulting in the sinking of polystyrene to the bottom of the container due to increased bulk (Supplementary Fig. 4). The

3.4. Engineering P. aeruginosa biofilms for ‘capture and release’ of microparticles

To incorporate the ‘release’ component in our strain, we inserted a PDE gene with an L-arabinose-inducible promoter (pBAD-yhjh plasmid) in the ΔwspF mutant. We had previously shown that the YhjH PDE could lower c-di-GMP levels and cause biofilm dispersal [42], thus addition of arabinose to the engineered ΔwspF/pBAD-yhjh strain could negate the pro-biofilm effects of wsp operon and cause biofilm dispersal. This was mediated by the production of glycosidases and proteases which can degrade the biofilm matrix [43].

To test if the engineered strain could trap and release microplastics, we first grew ΔwspF/pBAD-yhjh biofilms to accumulate microplastics in the absence of arabinose, followed by retrieval and arabinose treatment of biofilm-microplastics aggregates. In the initial growth of biofilms without arabinose, ΔwspF/pBAD-yhjh could accumulate microplastics similarly to ΔwspF, implying that the biofilm dispersal was not induced in the absence of arabinose (Fig. 4a-b). After the microplastics- containing biofilms were treated with varying concentrations of arabi- nose for 7 hrs, we quantified the recovery rate of freed microplastics from the biofilms. We found that increasing arabinose concentrations could effectively induce biofilm dispersal (Fig. 4c) and release of microplastics for recovery (Fig. 4d). To show that shear stress from shaking incubation of biofilms was not involved in biofilm dispersal, we

performed a negative control where no arabinose was added to the biofilms and found no significant changes to biofilm mass and loss of trapped microplastics (Fig. 4c-d). Furthermore, to show that arabinose was not a chemical stimulus for biofilm dispersal per se, but an inducible agent for pBAD-yhjh expression, we treated ΔwspF strain with 1% arab- inose and showed no significant changes to biofilm mass and loss of trapped microplastics (Fig. 4c-d).

3.5. Bioaggregation of microplastics from environmental samples

To show the ΔwspF/pBAD-yhjh strain could be used to aggregate microplastics from environmental samples, we collected seawater sam- ples in the vicinity of a sewage outfall where microplastics pollution was rampant and conducted our studies in bioreactors. We first tested the ability of our engineered strain to grow in freshwater and seawater supplemented with glucose and amino acids at various temperatures, and found little differences in the microplastics accumulated in the biofilms (Supplementary Fig. 5), implying that our proof-of-concept could be applied to different water sources.

Our engineered strain could form biofilms and accumulate most microplastics from seawater samples over time, with a few loose microplastics which were not trapped by biofilms (Fig. 5a). Addition of arabinose which activated the release mechanism caused dispersal of biofilms (Fig. 5b) and release of microplastics (Fig. 5c). The bio- aggregation of microplastics was not discriminatory, where the micro- plastics of different materials were recovered from the biofilms (Fig. 5d).

3.6. Bioaggregation of microplastics by Pseudomonas putida

Taking a step further for our proof-of-concept, we test its applica- bility to other bacterial species by employing the P. putida species known to colonise and form biofilms on microplastic surfaces in situ [13-16]. P. putida is an environmental bacterium which can degrade different types of pollutants and plastics [44], rendering it as another choice or- ganism for bioremediation and bioaccumulation [45,46]. Since many bacterial species do not contain the wspF mutation, we were also

Fig. 4. Trap and release mechanism of engineered ΔwspF/pBAD-yhjh strain. (a) No difference in biofilm CFU of ΔwspF/pBAD-yhjh and ΔwspF which trap microplastics. (b) No difference in mass of microplastics accumulated by ΔwspF/pBAD-yhjh and ΔwspF biofilms over time. (c) Bacterial CFU of biofilm released by arabinose in- duction of engineered strain. (d) Mass of microplastics released by arabinose induction of engineered strain. ***P < 0.001, n.s (not significant), One-Way ANOVA.

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interested in employing other biofilm-inducing mechanisms which can be conveniently applied to different microbes for trapping microplastics. One example is the use of an exogenous plasmid plac-yedQ, which encoded the YedQ DGC involved in c-di-GMP signaling [47]. We had previously employed plac-yedQ to study the induction of exopoly- saccharide production and biofilm formation by YedQ DGC in different Gram-negative bacterial species [33,48,49], which provided us the

rationale that it could be applied to accumulate microplastics in a similar fashion.

In this case, we engineered the trap mechanism in P. putida OUS82 and P. aeruginosa PAO1 by inserting plac-yedQ, and incorporated the pBAD-yhjh plasmid for release mechanism. Similar to ΔwspF/pBAD-yhjh, we first grew PAO1/plac-yedQ/pBAD-yhjh and OUS82/plac-yedQ/pBAD- yhjh biofilms to accumulate microplastics in the absence of arabinose,

Fig. 5. Biofilms can bioaccumulate micro- plastics from freshwater and seawater sam- ples. (a) Number of microplastic particles before and after microbial treatment. (b) Bacterial CFU of biofilms and dispersed cells after 8 hrs of activated release mechanism. (c) Number of microplastic particles before and after 8 hrs of activated release mecha- nism. *P < 0.05, ***P < 0.001, One-Way ANOVA. (d) material type collected and released by biofilms from environmental samples. PEHD: high-density polyethylene; PET: polyethylene terephthalate; PP: poly- propylene; PS: polystyrene.

Fig. 6. Trap and release mechanism of engineered P. putida OUS82/plac-yedQ/pBAD-yhjh and PAO1/plac-yedQ/pBAD-yhjh strain. (a) Both strains could trap micro- plastics in their biofilms. (b) Bacterial CFU of biofilm released by arabinose induction of engineered strain. (c) Mass of microplastics released by arabinose induction of engineered strains.

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followed by retrieval and arabinose treatment of biofilm-microplastics aggregates. In the initial growth of biofilms without arabinose, PAO1/ plac-yedQ/pBAD-yhjh and OUS82/plac-yedQ/pBAD-yhjh could accumulate microplastics, leaving low levels of free microplastics in the media (Fig. 6a). After the microplastics-containing biofilms were treated with varying concentrations of arabinose for 7 hrs, we quantified the recovery rate of freed microplastics from the biofilms. We found that increasing arabinose concentrations could effectively induce biofilm dispersal (Fig. 6b) and release of microplastics for recovery (Fig. 6c). This indi- cated that the trap and release mechanism could be applied flexibly to another microbial species.

4. Discussion

The United Nations’ Sustainable Development Goals (SDGs) have recently set the challenge of employing biotechnology in remediation and pollution control, so we should exploit the potential of microbial biotechnology in the removal of pollutants especially microplastics. While bacterial species with the ability to degrade microplastics have been identified, such as P. aeruginosa can degrade polystyrene and polythene [50,51], their varying plastic-degrading efficiency may pose a problem in the effective removal of microplastics from the sewage or water bodies. Furthermore, biodegradation of plastic may release toxic wastes, warranting the need to first capture microplastics efficiently, and then remove from the system (wastewater treatment plants or environment) for separate processing in isolated chambers. Hence, accumulation of microplastics is a viable way to remove microplastics safely. However, biofilms in nature also lack the bulk to cause floccu- lation or coagulation of microplastics from liquid suspension [52], thereby preventing the formation of larger aggregates which can sink to the bottom for convenient removal. While plastics such as PVCs and nylon have higher densities which allow them to sink to the bottom, low- density plastics such as polystyrene remain buoyant in liquids, which render their physical removal difficult [53].

We report the engineering of a P. aeruginosa strain with the ‘capture- and-release’ mechanism by harnessing the power of c-di-GMP signaling for biofilm and demonstrating its potential application in accumulating microplastic pollutants from polluted environmental samples. The engineered strain could accumulate microplastics at higher concentra- tions. The engineered strain could also aggregate low-density micro- plastics, which would lead to sinking and easier removal of microplastics in bioreactor tanks.

While biofilms are good at accumulating microplastics, their sticky exopolymeric matrix rendered separation of microplastics from biofilms and its recovery difficult. While it is currently not commercially feasible to recycle microplastics from the environment, improving its recovery can drive recycling efforts of plastics. Hence, we propose and incorpo- rate the release mechanism where the ΔwspF/pBAD-yhjh strain could activate its PDE activity via the addition of arabinose as stimulus. By reducing the c-di-GMP levels in biofilms, biofilms are driven towards dispersal, where glycosidases and proteases are produced to degrade the biofilm matrix [54] and release microplastics for convenient recovery of particulate plastic wastes.

This controllable biofilm development approach we have demon- strated here should prove widely applicable for other biofilm-enabled applications, such as bioremediation or biofilm-mediated biocatalysis of chemicals. Our proof-of-concept study exemplifies the potential for translation from biofilm biology to biofilm engineering for environ- mental applications. In the context of application in wastewater treat- ment plants, microplastics are difficult to remove conventionally by filters which poorly retain microplastics or harsh chemical treatment (alum salts) [4], resulting in the release of microplastics into the envi- ronment. Our work serves as proof-of-concept where microplastics are accumulated by biofilms, where they will sink to the bottom of the bioreactors for convenient removal. Upon transfer to new tanks, the release mechanism could be activated to break down biofilms into

suspensions of planktonic bacteria, and release microplastics for easier recovery. While our work may not be directly used for industrial ap- plications due to safety concerns attributing to genetically-modified bacteria, this provides the basis for future work in identifying pro- biofilm-forming isolates from sewage which can trap microplastics efficiently. The microplastic-laden biofilms could then be separated and treated with safe non-toxic anti-biofilm agents, such as nitric oxide or glycosidases [43,54,55], for biofilm disruption and microplastics release.

5. Conclusions

P. aeruginosa formed biofilms which could accumulate microplastics of varying sizes and materials within the exopolymeric matrix. Screening of biofilm mutants revealed the role of wsp operon in accu- mulating microplastics within the biofilm. Engineering a pro-biofilm strain based on wsp operon enhanced bioaccumulation of micro- plastics, while incorporating a release mechanism via dispersal pro- moted efficient release and recovery of microplastics.

Author Contributions

S.L.C and J.K.H.F designed methods and experiments. Y.S.L and M. M.L.L carried out laboratory experiments, analysed the data and inter- preted the results. All authors wrote the paper. All authors have contributed to, seen and approved the manuscript.

Declaration of Competing Interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Acknowledgements

This research is supported by the Environment and Conservation Fund, Hong Kong (ECF48/2019) and the State Key Laboratory of Chemical Biology and Drug Discovery Fund, The Hong Kong Polytechnic University (1-BBX8). We acknowledge the University Research Facility in Chemical and Environmental Analysis, and the University Research Facility in Materials Characterization and Device Fabrication, at The Hong Kong Polytechnic University for their technical assistance in Raman microspectrometry, and scanning electron microscopy, respectively.

Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi. org/10.1016/j.cej.2020.127079.

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