甘肃农业大学：食品科学与工程学院（文献讲义）Selective enrichment of bacterial pathogens by microplastic biofilm

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Selective enrichment of bacterial pathogens by microplastic biofilm Xiaojian Wu a , Jie Pan b , Meng Li b , Yao Li a , Mark Bartlam c, **, Yingying Wang a, * a Key Laboratory of Pollution Processes and Environmental Criteria (Ministry of Education), Tianjin Key Laboratory of Environmental Remediation and Pollution Control, College of Environmental Science and Engineering, Nankai University, Tianjin, 300071, China b Institute for Advanced Study, Shenzhen University, Shenzhen, 518060, China c College of Life Science, Nankai University, 300071, China article info Article history: Received 28 March 2019 Received in revised form 28 June 2019 Accepted 12 August 2019 Available online 13 August 2019 Keywords: Microplastic Biofilm Metagenomics Antibiotic resistance gene Pathogen abstract Microplastics have been found to be ubiquitous in freshwater ecosystems, providing a novel substrate for biofilm formation. Here, we incubated biofilm on microplastics and two natural substrates (rock and leaf) under a controlled environment to investigate the differences of microbial community structure, antibiotic resistance gene (ARG) profiles, and ARG microbial hosts between biofilms on three types of substrates. Results from high-throughput sequencing of 16S rRNA gene revealed that microplastic biofilm had a distinctive community structure. Network analyses suggested that microplastic biofilm possessed the highest node connected community, but with lower average path length, network diameter and modularity compared with biofilm on two natural particles. Metagenomic analyses further revealed microplastic biofilm with broad-spectrum and distinctive resistome. Specifically, according to taxonomic annotation of ARG microbial hosts, two opportunisitic human pathogens (Pseudomonas monteilii, Pseudomonas mendocina) and one plant pathogen (Pseudomonas syringae) were detected only in the microplastic biofilm, but not in biofilms formed on natural substrates. Our findings suggest that microplastic is a novel microbial niche and may serve as a vector for ARGs and pathogens to new environment in river water, generating freshwater environmental risk and exerting adverse impacts on human health. © 2019 Elsevier Ltd. All rights reserved. 1. Introduction As an environmental contaminant on the global scale, microplastics are the subject of increasing scientific concern due to their elevated ecological risks and potentially adverse effects on public health. Defined as plastic particles with a size of less than 5 mm, microplastics can be generated in one of two ways: primary microplastics initially manufactured in small size for industrial purposes, such as microbeads adding to personal care products as abrasives; and secondary microplastics derived from the fragmentation of large plastic pieces by physical, chemical and biological factors, including mechanical abrasion, photooxidation and biological degradation (Rachman, 2018). Oceans have long been the focus of studies into microplastics because they are considered to be the largest sink of microplastics. Recently, however, the focus of research has expanded to include freshwater ecosystems, given that approximately 80% of microplastic contamination in marine environments originates from land and river (Rachman, 2018). Current evidence indicates that microplastics are ubiquitous in rivers on the global scale (Eerkes-Medrano et al., 2015; Klein et al., 2015; McCormick et al., 2014; Su et al., 2016), and have been discovered in freshwater biota at different trophic levels (Rachman, 2018). On the long voyage from source to sink, microplastics will be colonized by microorganisms and wrapped by biofilms after dynamic succession (Schluter et al., 2015). Colonized by diverse and metabolically complex microbial consortia, the microplastic surface is proposed to be a hotspot for horizontal gene transfer (HGT) (Schluter et al., 2015; Sorensen et al., 2005). It has been demonstrated that biofilms serve as reservoirs for pathogenic bacteria (Wingender and Flemming, 2011) and microenvironments for horizontal gene transfer (Hausner and Wuertz, 1999). Horizontal gene transfer mediates the flow of antibiotic resistance genes (ARGs) between the microorganisms in biofilm and environmental bacteria (Bengtsson-Palme and Larsson, 2015; Li * Corresponding author. ** Corresponding author. E-mail addresses: bartlam@nankai.edu.cn (M. Bartlam), wangyy@nankai.edu.cn (Y. Wang). Contents lists available at ScienceDirect Water Research journal homepage: www.elsevier.com/locate/watres https://doi.org/10.1016/j.watres.2019.114979 0043-1354/© 2019 Elsevier Ltd. All rights reserved. Water Research 165 (2019) 114979

et al., 2015; Martinez et al., 2015; Van Boeckel et al., 2015) via mobile genetic elements (MGEs) such as plasmids, transposons, bacteriophages, insertion sequences and integrons (Stokes and Gillings, 2011). Given that some opportunistic pathogens, such as Vibrio spp. have been discovered in microplastic biofilm (Foulon et al., 2016; Keswani et al., 2016; Kirstein et al., 2016), it is possible that specific pathogens in microplastic biofilm will acquire ARGs from environmental bacteria and travel with microplastics to reach remote environments. The resistance to antibiotics of pathogens harbouring ARGs make them hard to be killed by therapeutics, which poses a potential worldwide threat to ecosystem and human health. Previous studies have indicated that the biofilm communities on plastic substrates were distinctive from those in water columns or sediment (Amaral-Zettler et al., 2015; De Tender et al., 2015; McCormick et al., 2014; Oberbeckmann et al., 2014; Zettler et al., 2013). However, the link between the structure and function of the biofilm communities on microplastic is still not fully understood. Here, we investigated the differences between biofilms on microplastics and two natural substrates (rock and leaf) by comparing the microbial community structure, ARG profiles and ARG bacterial hosts of the biofilm associated with microplastics and two natural substrates. This approach allowed us to better understand the features of the microbial community on microplastics, and to provide insight into the possibility of various surfaces, both anthropogenic and naturally occurring, to spread antibiotic resistance genes via biofilms in the freshwater ecosystem. 2. Materials and methods 2.1. Experimental design and water quality parameters In order to test the effects of substrate type (anthropogenic or natural) on the associated biofilms, we used river water to culture the biofilm in bioreactor (BioFlo CelliGen 115, New Brunswick, Eppendorf, USA). The river water was collected in the Haihe River, the largest river catchment in Northern China, flowing through several cities and finally into the sea. Polyvinyl chloride (PVC) microplastic pellets (density 1.35e1.45 g cm3 , ø 3 mm) were purchased from Aladdin Biochemical Technology Co. Ltd. (Shanghai, China). Rock (quartz) was purchased from a flower shop and leaves (Platanus acerifolia) were cut into small pieces. Rocks and leaves were sieved with stainless steel laboratory grade meshes to ensure they were within the size range of 2e4 mm. All sieved particle (microplastic, rock, and leaf) were rinsed with deionized water three times and placed in the dark until they were dried at room temperature. Prior to the experiment, the bioreactor was rinsed with deionized water three times and sterilized by autoclaving. River water was continuously pumped into the bioreactor. All treated particles (microplastic, rock, and leaf) were wrapped with sterilized gauze and each type was divided into 5 independent aggregates. The 5 aggregates or each particle type correspond to the 5 replicates of the 16S rRNA gene amplicon sequencing in the following analysis. All 15 aggregates (n ¼ 5 replicates 3 types of particles) were incubated in a bioreactor with 5 L of working volume for 2 weeks (from April 18th to May 2nd, 2018). 2.2. Scanning Electron Microscope (SEM) imaging The formation of biofilms on different substrates was investigated after seven days using a field-emission scanning microscope (JEOL JSM 7800, Japan). The samples were rinsed with PBS buffer and post-fixed with 2% osmium tetroxide. Dehydrated by graded ethanol series (15 min each in 35%, 50%, 75%, 90%, followed by 45 min in 100% ethanol). The samples were then dried by Polarion E3000 Critical Point Dryer overnight. Sputter was coated with gold layer at 25 mA under Argon (Ar) atmosphere at 0.3 MPa, the samples were transferred to the conductive carbon tape mounted on the sample holder, and the morphology was characterized under the SEM. 2.3. Flow cytometry (FCM) measurement In brief, 1 g particles (microplastic, rock and leaf) were sampled and rinsed with sterile PBS. The particles were immersed in 10 mL of sterilized PBS buffer and ultrasonic treatment (B Sonopuls HD 3200, Bandelin Sonorex, Rangendingen, Germany) was used to detach the bacteria associated with the particles. The ultrasonic device included a generator (Sonopuls HD3200), ultrasonication energy transfer unit (UW 2200), booster horn (SH 213G), and needle (MS72). The settings used were: amplitude: 302 mm; cycle duration: 30 s; pulse level: 50%; power: 50%. Flow cytometry analysis was used to determine the biomass of the biofilm and the planktonic bacteria concentration every two days, based on previously described methods (Wen et al., 2015). 1 mL of sample was stained by SYBR Green I (10000 diluted, Invitrogen). Flow cytometry analysis was performed using a BD Accuri C6 Plus instrument (BD Biosciences, USA). After being mixed thoroughly with vortex and incubated in the dark for 10 min at 37 C, the emitting fluorescence signal excited by the blue laser at 488 nm was selected on FITC-PerCP tunnel (flow rate: 66 mL/min, green fluorescence tunnel: 533 nm, red fluorescence tunnel: >670 nm) and the total cell concentration (TCC) could be measured after data were processed using BD Accuri C6 Plus software as described previously (Wen et al., 2015). 2.4. Sampling and DNA extraction On day 14, the same type of particles (microplastic, wood and rock) were recovered and rinsed three times with sterilized PBS. Particles were immersed in 100 mL of sterilized PBS and treated by 30 s of ultrasonication. The detached biofilm from microplastic, rock, leaf particles (particle-associated part fraction, n ¼ 5) was collected by centrifugation (14,000 g, 10 min) and DNA was extracted using the Mobio PowerBiofilm® DNA isolation kit (Mo Bio Laboratories, Carlsbad, CA, USA). The planktonic bacteria in river water (planktonic part fraction, n ¼ 5) were collected by filtration through a sterilized mixed cellulose esters membrane with a pore size of 0.1 mm membrane (Millipore, USA). The membranes were stored at 20 C before DNA extraction. DNA was isolated using the Mobio PowerWater® DNA isolation kit (Mo Bio Laboratories, Carlsbad, CA, USA). The extraction processes followed the manufacturer's instructions. Following the extraction, agarose gel electrophoresis (2.0%) and a Qubit 2.0 Fluorometer (Invitrogen) were used to check the concentration of DNA samples, which were stored at 80 C for further study. 2.5. 16S rRNA gene sequencing and data processing Two-step PCR was conducted to amplify the 16S rRNA gene (Sutton et al., 2013). With this approach, tags and adapters were added in a second round of PCR amplification. To amplify the V3eV4 hypervariable regions of 16S rRNA gene, the primer set 338F (50 -ACTCCTACGGGAGGCAGCAG-30 ) and 806R (50 -GGACTACHVGGGTWTCTAAT-30 ) combined with adapter sequences and barcode sequences were used (Chu et al., 2015). Triplicate PCR reactions were performed in 50 mL reaction mixtures, which contained 10 mL GoTaq buffer, 0.2 mL Q5 High-Fidelity DNA Polymerase, 10 mL High GC Enhancer, 1 mL dNTP, 10 mM of each primer, 60 ng 2 X. Wu et al. / Water Research 165 (2019) 114979

genome DNA and ddH2O to make up a total volume to 50 mL. The PCR procedure conditions were as follows: an initial denaturation at 95 C for 5 min; followed by 15 cycles at 95 C for 30 s, 50 C for 30 s and 72 C for 40 s; with a final extension at 72 C for 7 min. The PCR products from the first step PCR were purified through VAHTSTM DNA Clean Beads. A second round PCR was then performed in a 40 mL reaction which contained 20 mL 2 Phmsion HF MM, 8 mL ddH2O, 10 mM of each primer and 10 mL PCR products from the first step. Thermal cycling conditions were as follows: an initial denaturation at 98 C for 30s; followed by 10 cycles at 98 C for 10s, 65 C for 30 s min and 72 C for 30 s; with a final extension at 72 C for 5 min. Finally, all PCR products were quantified by Quant-iT™ dsDNA HS Reagent and pooled together. Highthroughput sequencing analysis was performed on the purified, pooled sample using the Illumina Hiseq 2500 platform (2 250 paired ends). Sequence analysis was carried out using the QIIME pipeline (version 1.8.0) (Caporaso et al., 2010). In brief, the two pair-end sequencing data were merged into one according to the overlapping relationship using FLASH (version1.2.7) (Magoc and Salzberg, 2011). After filtering low-quality and short sequences by Trimmomatic (version 0.33) (Bolger et al., 2014) and chimera sequences by Uchime (Kozich et al., 2013), we subsampled the reads to obtain the same number of reads in each sample, which was 73,289 clean reads. The sequences were clustered into operational taxonomic units (OTUs) at 97% identity with UCLUST (Caporaso et al., 2010; Edgar, 2010). Taxonomy assignments were conducted by applying SILVA database as the reference (Caporaso et al., 2010). 2.6. Shotgun metagenomics and data processing Paired-end (2 150) metagenomic sequencing was performed on an Illumina HiSeq 2000 platform. The raw reads were dereplicated and trimmed by the quality. The clean reads were assembled into scaffolds by using IDBA-UD (version 1.1.1) (Peng et al., 2012). The open reading frames (ORFs) in scaffolds were predicted by Prodigal (version 2.6.3) (Hyatt et al., 2010) and annotated using BLASTp by applying the ARGs database (E value 105 , sequence identity 80%, query coverage 70%, alignment length 25 amino acids). The abundance of ARGs was calculated by mapping reads to the gene sequences (Ma et al., 2016) and normalized by the abundance of 16S rRNA genes (relative abundance), expressed as copies of ARGs per copy of 16S rRNA gene (Li et al., 2015), consistent with the qPCR results reported in many previous studies (Chen et al., 2017; Feng et al., 2018; Marti et al., 2018). The relative abundance of the ARGs type or subtype were calculated using the following equation (Li et al., 2015): The ORF sequences of the scaffolds carrying ARGs were annotated using RefineM (version 0.0.23) (Parks et al., 2017) and scaffolds with length greater than 500 bp were retained. All sequencing data are deposited in the National Center for Biotechnology Information (NCBI) Sequence Read Archive (accession No. SRP174395 for 16S analysis and accession No. SRP174465 for metagenomic analysis). 2.7. Data analysis Statistical analyses were performed using R (version 3.4.1, R Found Stat Comput, Vienna) and the results were visualized by Origin. The P value of microplastic biofilm > rock biofilm. The leaf biofilm possessed the highest biomass of the three biofilms, which could be the result of the fast breakdown process of the dissolved organic matter (DOM) in the leaf, thus increasing the availability of nutrients and promoting bacterial growth (Gulis and Suberkropp, 2003). During leaf decomposition, the release of a large amount of organic substances attracts colonizers to utilize the nutrients and supports the growth of the thick biofilm. The nature of the microbial community in the leaf biofilm plays an important role in the leaf utilization (McArthur et al., 1985). The varied responses of the bacterial species to components during leaf decomposition have been observed (McNamara and Leff, 2004); for example, the population of Burkholderia cepacia increased when DOM concentrations were greatest, while the population of Pseudomonas putida was inhibited when total DOM concentrations were greatest, which indicates the complexity of microbial population dynamics. Microplastics and rock do not decompose and therefore, after entering the aquatic environment, their surfaces will absorb nutrients from water and form the conditioning film (Siboni et al., 2007). Colonizers will be attracted by the conditioning film and this initial Abundance ¼ Xn 1 NARGlike sequence Lreads. LARG reference sequence N16S sequence Lreads L16S sequence ! X. Wu et al. / Water Research 165 (2019) 114979 3

community will be continuously shaped. In contrast with the rock surface that possesses fewer nutrients and lower biofilm biomass, microplastics could be used as an energy and carbon source by microorganisms producing enzymes capable of hydrolysing plastic (Yoshida et al., 2016). In addition, the variation of biofilms on the three materials could also be attributed to the discrepant consortia and microbial community structure. 3.2. Microplastic biofilm had distinctive microbial communities structure compared with rock and leaf biofilm The biofilms formed on the three materials could be divided into two categories: biofilm on anthropogenic substrates (microplastic) and natural substrates (rock and leaf). The biofilm community structure was evaluated and compared from phylogenetic composition, a-diversity (within-sample diversity), b-diversity (betweensample diversity), and differential abundance analysis of enriched/ depleted OTUs. Microplastic and rock biofilm shared the most dominant phyla, with Proteobacteria the most abundant (60%e77%), followed by Bacteroidetes (8%e15%) and Firmicutes (6%e14%). In leaf biofilm, Bacteroidetes (46%) was the most abundant (Fig. 2A). The relative proportions of Chlorobi, Acidobacteria, Gemmatimonadetes, Actinobacteria, Fibrobacteres, Planctomycetes, Hydrogenedentes, and Chlamydiae were higher in biofilms formed on microplastics than in biofilms formed on natural substrates (Table S2). Previous research has shown a lower abundance of Bacteroidetes in biofilms formed on plastic surfaces compared with native (cellulose) and inert (glass beads) particles (Ogonowski et al., 2018). In this study, the abundance of Bacteroidetes in biofilms formed on microplastics was lower than in biofilms formed on both of the natural substrates, which is consistent with previous observations. Fig. 1. Biofilm formation on the surface of microplastic, rock and leaf after 14 days of incubation. (A) Biomass of biofilm was determined by flow cytometry every two days; (B) Scanning Electron Microscope images of different substrate surface before and after the incubation of biofilm in river water (the first row: before the biofilm incubation; the second row: after the biofilm incubation). 4 X. Wu et al. / Water Research 165 (2019) 114979

To evaluate the a-diversity of the biofilm community, the Shannon-Wiener index was calculated. The a-diversity revealed a gradient of microplastic biofilm > rock biofilm > leaf biofilm. The Shannon index of the microplastic biofilm was significantly higher than those on natural substrates (Fig. 2B, Wilcoxon rank-sum test, P value < 0.05), indicating that microplastic biofilm was more diverse than the rock and leaf biofilm. The high a-diversity may suggest the resilience to perturbation (Girvan et al., 2005) and the capacity of maintaining the activities of the biofilm members (Philippot et al., 2013). Additionally, principal coordinate analysis (PCoA) was conducted to identify the separation pattern between biofilm communities. Based on a phylogenetically weighted UniFrac distance matrix, all microplastic biofilm samples were clearly clustered into one group and separated from the rock and leaf biofilm samples along the first principal coordinate, illustrating that the largest source of variation in biofilm communities is substrate type (Fig. 2C). The pattern of separation was consistent with the gradient of a-diversity from microplastic biofilm to rock biofilm to leaf biofilm. Microorganisms in river water preferentially colonize the substrate and the gradually matured biofilm selectively enriches specific microorganisms from river water. This two-way selection mutually carves the delicate architecture of biofilm. By identifying the OTUs with differential abundances in biofilm compared to in river water, i.e. significantly enriched/depleted OTUs in biofilm, the selection on the three substrates could be determined. A total of 515 OTUs were identified in all three types of biofilm and in river water. Of these, 296, 284, and 178 OTUs were significantly enriched from river water by microplastic, rock, and leaf biofilm, respectively (Fig. 3A, P value < 0.05). The majority of OTUs (282 out of 296) enriched in the microplastic biofilm also colonized rock and leaf surfaces (Fig. 3B). Microplastic biofilm was found to be more similar to rock biofilm than to leaf biofilm, given that microplastic biofilm shared 274 enriched OTUs with rock biofilm but only shared 159 enriched OTUs with leaf biofilm. Specifically, 14 OTUs were found to be uniquely enriched in microplastic biofilm, and these were distributed across 3 phyla: Proteobacteria, Gemmatimonadetes and Actinobacteria (Fig. 3C, Table S3). The OTUs uniquely enriched in the rock or leaf biofilm were distributed across 6 phyla (Bacteroidetes, Firmicutes, Saccharibacteria, Proteobacteria, Parcubacteria, and Cyanobacteria). Enriched or depleted OTUs in the microplastic biofilm revealed that colonization onto the substrates is not a passive process, and that this novel niche selects the colonizers. Previous studies have reported differences in microbial communities between microplastic biofilms and water columns, including river and ocean (Kettner et al., 2017; McCormick et al., 2014; Zettler et al., 2013). Evidence for the differences between biofilm on plastic and natural substrates (Miao et al., 2019; Ogonowski et al., 2018) has also been documented, which is consistent with our results. Miao et al. (2019) incubated biofilm on microplastic substrates (polyethylene, polypropylene) and natural substrates (cobblestone and wood) in lake water under controlled conditions. The sorting phenomenon of microbial communities between microplastic and natural substrates was observed. Ogonowski et al. (2018) exposed ambient Baltic seawater to plastic (polyethylene, polypropylene, polystyrene) and cellulose particles. The plastic associated communities were evidently different from those formed on the non-plastic substrates. In accordance with previous investigations, our study has contributed another example of the term “plastisphere” applied to freshwater ecosystems (McCormick et al., 2014). 3.3. Network structure of microplastic biofilm community was more complex and connected The biofilm community is a tightly combined network. The above analyses had proved the discrepancy of taxa composition and abundance between microplastic biofilm and natural substrates. The underlying interaction among taxa in complex communities should also be evaluated to better recognize the co-abundance pattern of microbial consortia. In order to investigate the topology of the network and the potential interactions between Fig. 2. Relative abundance, a-diversity (within-sample diversity) and b-diversity (between-sample diversity) of rock biofilm, leaf biofilm and microplastic biofilm. For each type of community (rock biofilm, microplastic biofilm, leaf biofilm and planktonic communities in river water), there were 5 replicates. (A) Histograms of phyla abundances in three types of biofilm and river water. (B) Within sample diversity (a-diversity) measurements indicate a decreasing gradient in microbial diversity from the microplastic biofilm to leaf biofilm. The horizontal bars within boxes represent the median. The tops and bottoms of boxes represent the 75th and 25th quartiles, respectively. The upper and lower whiskers extend 1.5 the interquartile range from the upper edge and lower edge of box. (C) Principal coordinate analysis (PCoA) analysis based on the weighted UniFrac distance matrix indicates a clear separation between the three types of biofilm communities and planktonic communities in river water. X. Wu et al. / Water Research 165 (2019) 114979 5

more complex than that of rock and leaf biofilms. A total of 120, 110, and 35 OTUs were identified to have a significant correlation with each other in microplastic, rock and leaf biofilm, respectively (Table S4). The microplastic biofilm had the highest number of correlated OTUs, which belonged to 12 phyla: Proteobacteria, Bacteroidetes, Firmicutes, Acidobacteria, Epsilonbacteraeota, Verrucomicrobia, Gemmatimonadetes, Patescibacteria, Tenericutes, Planctomycetes, Chlamydiae, and Nitrospirae. Shown by the co-occurring patterns between OTUs with significant correlations, the co-occurrence network depicts potential interacting or niche-overlapping relationships in which the microorganisms in biofilm could be involved (Zhou et al., 2010). The complexity of connections of the different biofilms was explored and the topological properties of the resulting networks was calculated. As can be seen from Fig. 4, the microplastic biofilm network visually possessed the highest level of complexity, whereas the rock and leaf biofilms presented a less complex network. In terms of the topology properties, the microplastic biofilm network presented the highest number of connections per node (average degree ¼ 13.835), indicating a highly connected community (Table S5). However, the average path length (2.062) and network diameter (6) for the microplastic biofilm were both lower than for the other two types of biofilms. The average path length is defined as the average number of steps along the shortest paths between each node, being a measure of efficiency on a network (Zhou et al., 2010), while the network diameter is defined as the longest distance between nodes in the network. Meanwhile, the microplastic biofilm network presented a less modular structure (modularity ¼ 0.708) than rock (0.847) and leaf biofilm (0.719), which is characterized by the presence of different groups of nodes containing high numbers of interconnections, with some degree of independency between groups (Newman, 2006). In short, the topology properties suggested that the microplastic biofilm was the community with the highest connection of nodes, but with lower average path length, network diameter and modularity. The relatively higher modularity of the leaf biofilm combined with the shortest average path length may imply a more prompt response of the microbial community to environmental perturbations (Faust and Raes, 2012), by allowing the perturbations to reach the whole network immediately so that the network structure, as well as the functions, could be altered (Zhou et al., 2010). It would appear that the microbial community attached to the leaf was not just loosely gathering around it, but could thrive and respond to environmental disturbances as a whole. 3.4. Broad-spectrum and distinctive resistome (profile of ARGs) were detected in the microplastic biofilm It is essential to evaluate the ARGs profiles in the three types of biofilm and in river water, given that information regarding of ARGs profiles in microplastic biofilm is limited compared with other environments which have been thoroughly investigated and recorded (Allen et al., 2010). The ARG profiles in all biofilm and river water samples were determined. 267 ARG subtypes belonging to 26 ARG types were identified in all samples. The diversity of ARG (number of ARG subtype) decreased, in the order of leaf biofilm- (207), microplastic biofilm- (188), rock biofilm- (185), and river water- (67) (Table S6, Table S7). The ARG abundance of biofilm was approximately 3-fold higher than that of river water (Fig. 5A), indicating that ARGs were enriched by biofilm. The dominant ARG types included multidrug-ARGs (22.03%), MLS (18.06%), bacitracin (9.59%), polymyxin (9.44%), acriflavine (6.31%), beta-lactam (6.08%), and aminoglycoside (5.81%) (Fig. 5B). Statistically, 40.6% of the total 267 ARGs were shared by all samples, such as MLS macB, multidrug mdtB, and acriflavine acrB (Fig. 5C). Our sampling site, Haihe River, is located in a high economically developed region and is a direct recipient of urban, agricultural, and industrial waste (Zhao et al., 2010). The high antibiotic emissions in the Haihe River region were verified in previous studies (Luo et al., 2010, 2011). The emission of antibiotics into the water ecosystem would contribute to the antibiotic resistance selection (Andersson and Hughes, 2011), which could explain the high diversity of ARGs in biofilms cultured with water sampled from the Haihe River. Many ARG subtypes found in the microplastic biofilm were also previously detected using PCR methods, such as ampC in wastewater (Volkmann et al., 2004), tetO, tetQ in lagoons (Smith et al., Fig. 4. OTU co-occurrence network analysis of microbial communities of rock biofilm, microplastic biofilm and leaf biofilm based on correlation analysis revealed different pattern of community topology structure. For each type of biofilm (rock biofilm, microplastic biofilm, and leaf biofilm), there were 5 replicates. Each node represents one OTU and an edge is drawn between OTUs based on 16S rRNA if they share a strong (Spearman's correlation coefficient r > 0.8) and significant (P-value < 0.01) correlation (red for positive correlation, grey for negative correlation). The OTUs were labelled at the phylum level. The size of the node is proportional to the number of connections (i.e. degree). Each edge represents the correlation between OTUs. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.) X. Wu et al. / Water Research 165 (2019) 114979 7

2004), sul1, sul2, blaTEM in river water (Jiang et al., 2013). The microplastic biofilm possessed unique ARG profiles: multidrug resistance gene smeE (0.216%), mdsC (0.041%); fluoroquinolone resistance gene qnrVC6 (0.029%); MLS resistance gene ermF (0.029%), lnuE (0.023%); beta-lactam resistance gene blaVEB-9 (0.024%); aminoglycoside resistance gene aadA13 (0.021%), APH(9)- Ia (0.009%), APH(300)-VI (0.008%); aadA16 (0.008%); fosfomycin resistance gene fosK (0.017%); rifampicin resistance gene arr-5 (0.014%); chloramphenicol resistance gene cmx (0.010%); trimethoprim resistance gene dfrA15 (0.009%) (Fig. 6A, Fig. 6B). The ARG subtype with the highest relative abundance in the microplastic biofilm belonged to the multidrug resistance type, which was also found to be dominant in other specific environments (Forsberg et al., 2012; Li et al., 2015; Zhu et al., 2013). Consistent with the results of previous studies reporting the presence of Pseudomonas in hospital wastewater treatment plants and in drinking water carrying multidrug resistant genes (Ma et al., 2016), we also found a high proportion of MDR genes in the microplastic biofilm. The microplastic biofilm possessed ARG subtypes not detected in the rock or leaf biofilms, indicating that these ARGs could not be loaded by natural particles. Specific types of ARGs may be selectively enriched by the microplastic biofilm. The extracellular substances in biofilms could be shared and the function of the cooperative interaction of biofilms would transport ARGs between the cells via the biofilm matrix (Ostrowski et al., 2011). Direct leaf litter is one of the major sources of organic matter in the freshwater system and is rapidly colonized by microorganisms when entering the river environment. Most of the leaf leachate is composed of simple carbohydrates and may provide nutrients and carbon to the bacteria on the surface. Much of the microbial biomass in the river is located in biofilms (Lock et al., 1984). As the first to encounter leachate from leaf, McArthur and colleagues Fig. 7. ARG host (the bacteria carrying ARG) detected in the three types of biofilm ARG types and ARG hosts. (A) Distributions of ARG host in the ARG types; (B) Relative proportions of the four major genera harbouring ARGs; (C) Relative proportions of the antibiotic resistance gene types of the four major harboured genera. X. Wu et al. / Water Research 165 (2019) 114979 9

showed that leaf biofilm influences the utilization of leaf leachate (McArthur et al., 1985). The highest diversity of ARG (number of ARG subtype) in the leaf biofilm reminded us that leaves may be a natural source of ARGs in the river environment. In addition to microplastics, we should also focus on the potential to transfer ARGs inside the freshwater system. A previous study based on a metagenomics analysis showed the exchange of ARGs between clinical pathogens and environmental bacteria (Forsberg et al., 2012). After verifying that the microplastic biofilm loads ARGs, it is essential to trace back the origin of the ARGs and to identify which ARGs were carried by which bacteria. 3.5. Microplastic biofilm selectively enriched opportunisitic human pathogens which carried ARGs Biofilms equip the microbial community with novel functions, including increased resistance to antibiotics (Stewart and Costerton, 2001). After verifying that the bacteria in the microplastic biofilm indeed possessed genes with resistance to antibiotics, questions remain about which bacteria possess these ARGs and whether they pose a threat to human health. To answer that question, taxonomic annotation of ARG bacterial hosts was conducted. At the genus level, we found that with the exception of the unclassified Pluralibacter (18.97%), Pseudomonas (17.78%), Leclercia (11.71%), and Pantoea (11.06%) are relatively abundant in all of the samples (Fig. 7A) (Table S8). The proportions of the four genera in biofilm on three types of substrates are shown in Fig. 7B. In both microplastic and rock biofilms, the dominant host was Pseudomonas with a proportion of 34.7% and 26.7%, respectively, while the largest proportion (20.9%) in leaf biofilm was Pantoea. Most of the hosts possessed multidrug resistance genes and aminoglycoside resistance genes (Fig. 7C). The multidrug resistance genes were carried by 50.9%, 50.2%, 46.8% of the host Pseudomonas in the microplastic biofilm, rock biofilm and leaf biofilm, respectively (Table S9). The presence of Pseudomonas in the microplastic biofilm was further verified by quantitative real-time PCR of samples from three independent bioreactors (see supplementary material for detailed data Table S10, Table S11). Next, host tracking analysis of the four major hosts (Pluralibacter, Pseudomonas, Leclercia, and Pantoea) was conducted at the species level. A number of the scaffolds carrying ARGs were annotated to plant pathogens and human pathogens. It should be noted that several pathogenic bacteria were only detected in microplastic biofilm, but not in the rock or leaf biofilms (Fig. 8). These pathogenic bacteria included: Pseudomonas monteilii, an opportunistic pathogen more frequently detected in clinical environments and the major microbial species behind the disease of hypersensitivity pneumonitis and exacerbating bronchiectasis (Aditi et al., 2017; Bogaerts et al., 2011; Ocampo-Sosa et al., 2015); Pseudomonas mendocina, an environmental bacterium causing opportunistic nosocomial infections, such as infective endocarditis and spondylodiscitis (Chi et al., 2005; Mert et al., 2007); and Pseudomonas syringae, an agriculturally important plant pathogen (Hirano and Upper, 2000). The microplastic biofilm selectively enriched these pathogens, but no enrichment was observed in either rock or leaf biofilms. As one of the most common plant pathogens, P. syringae has more than 60 pathovars and infects almost all economically important crop species (Xin et al., 2018). In a previous study, the outbreak of harmful Alexandrium taylori algae bloom was shown to result from the attachment of A. taylori cells to plastic debris (Maso et al., 2003). The plant pathogens carried by microplastics may lead to pecuniary loss. The existence of pathogens with ARGs will increase the risk of antibiotic ineffectiveness because of the difficulty in treating the infection. Organic aggregates in water are islands for microbial assemblages and sometimes for pathogens (Lyons et al., 2010). Similarly, microplastics could be considered as a much more durable island with high residence time under natural conditions. Microplastics in the river water environment may become the preferred surface for specific pathogens with other alternative natural particles existing in the same environment at the same time. Microplastics not only serve as a novel microhabitat for biofilm colonization, but they also increase the likelihood of pathogens propagating. The pathogens may have contact with humans along the river via direct exposure to the water area. A number of questions remain to be addressed by future research, including whether the architecture of the biofilm will remain complete after longer migration distances and greater geographic spread, whether the microbial community structure will be continuously shaped because of environmental factors deviation in varied water environments, and whether the pathogens in the biofilm remain hazardous and have an impact on local microbial communities. Fig. 8. Taxonomic assignment of ARG host profiles in the microplastic, rock, and leaf biofilms at the species level. The dots coloured if the host species was detected in the corresponding biofilm. The host species was coloured yellow when it is detected in the rock biofilm, red when it is detected in microplastic biofilm, green when it is detected in the leaf biofilm), grey if it was not detected. R stands for rock biofilm; MP stands for microplastic biofilm; L stands for leaf biofilm. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.) 10 X. Wu et al. / Water Research 165 (2019) 114979

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