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.pdf

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 <0.05 was regarded as statistically significant (Wilcoxon Rank Sum test). To understand the inter associations among microorganisms in the whole community, we used network analysis (Barberan et al., 2012). After calculating the pairwise Spearman's correlation coefficients (r), a matrix was constructed to explore the potential relationships in the microbial community. The correlation between two nodes were considered as statistically significant for r 0.8 and P value 0.01 and the correlation network was formed. The analysis result was performed using R (version 3.4.1) and visualized by the Gephi (version 0.9.2). 3. Results and discussion 3.1. Biomass of microplastic biofilm was more than rock biofilm but less than leaf biofilm Measured by flow cytometry every two days, the biomass of microplastic biofilm constantly increased and reached a peak of 3.1 108 cells g1 on day 8 (Fig. 1A, Table S1). The planktonic cells in the surrounding water maintained a relatively steady concentration of 2.7 107 cells mL1 . The highest biomass of microplastic was 2.4 times that of the rock biofilm and 0.14 times that of leaf biofilm. Based on the SEM micrographs, the occurrence of the biofilm after a short period of time compared with the pristine surface of the materials was further demonstrated (Fig. 1B, Fig. S1). The outline of bacterial cells in microplastic and leaf biofilm were both evident. The cell surface of microplastic biofilm was smooth while the cells of leaf biofilm were rough and small amount of filiform extracellular polymeric substance (EPS) could be observed. The observed biomass was in the order leaf biofilm > 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. 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