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Characterization of bacterial community in biofilm and sediments of wetlands dominated by aquatic macrophytes
Ecological Engineering 97 (2016) 242–250
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Characterization of bacterial community in biofilm and sediments of wetlands dominated by aquatic macrophytes Si Pang a , Songhe Zhang a,∗ , XiaoYang Lv a , Bing Han a , Kaihui Liu a , Changhao Qiu a , Chao Wang a , Peifang Wang a,∗ , Harry Toland b , Zhenli He c a Ministry of Education Key Laboratory of Integrated Regulation and Resource Development on Shallow Lakes, College of Environment, Hohai University, Nanjing 210098, China b Geography & Earth Sciences, Aberystwyth University, Llandinam Building, Penglais Campus, Aberystwyth, Wales, SY23 3DB, United Kingdom c University of Florida, Institute of Food and Agricultural Sciences, Indian River Research and Education Center, 2199 South Rock Road, Fort pierce, FL 34945, USA
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Article history: Received 27 September 2015 Received in revised form 16 July 2016 Accepted 5 October 2016 Keywords: Biofilm Macrophytes Sediment Nitrogen cycle
a b s t r a c t Though aquatic macrophytes play an important role in wetlands, their effects on bacteria community structures in biofilms and sediments are far from clear. In the present study, bacterial communities were investigated in biofilm attached to leaves, stems and roots of aquatic macrophytes (Myriophyllum verticillatum, Nymphoides peltatum and Trapa japonica) and in vertical sediment cores from vegetated and unvegetated areas in a wetland located in Lake Hongze. The densities of microbes and epiphytic algae in biofilms were higher on leaves of M. verticillatum than those of two floating macrophytes. Phyla Proteobacteria, Bacteroidetes, Chloroflexi, Firmicutes and Verrucomicrobia were detected in both biofilms and sediments. As revealed by cluster analysis and principal component analysis, differences in structures of microbial communities were detected between biofilms and sediments and between vegetated and unvegetated sediments. The potential roles of nitrifying- and denitrifying- bacteria in sediments with respect to those in biofilms were discussed. These results highlight that the restoration of aquatic macrophytes can increase bacteria diversity and the surface and quantity of biofilms and therefore bacteria diversity. These data provide useful information for further understanding the role of aquatic macrophyte-biofilm system in wetlands in future. © 2016 Elsevier B.V. All rights reserved.
1. Introduction Aquatic macrophytes, key components of aquatic ecosystems, can absorb nutrients, heavy metals and other inorganic or organic contaminants from the water column and the sediments (Dhote and Dixit, 2009). Aquatic macrophytes are widely grown in wastewater treatment wetlands for improving water quality (Thorén, 2007). However, due to the improper anthropogenic activities aquatic macrophytes have declined significantly and biodiversity in freshwater has decreased, this will be a major driver of future ecosystem change (Hooper et al., 2012). Biofilms (known as epiphytic microbes) can form on the gasliquid and solid-liquid interfaces, and potentially constitute an important step in the integration of biogeochemical cycles and
∗ Corresponding authors. E-mail addresses: [email protected] (S. Zhang), [email protected] (P. Wang). http://dx.doi.org/10.1016/j.ecoleng.2016.10.011 0925-8574/© 2016 Elsevier B.V. All rights reserved.
dynamics of microbes to ecosystem function (Battin et al., 2003). Aquatic plants are key components in spatial heterogeneity, which is essential for the establishment and development of biofilms in aquatic ecosystems. Aquatic plants represent a special substrate for biofilms, as they can release oxygen, which is beneficial to aerobic bacteria attached to the plant, and promotes the transformation of nitrogen in water (Eriksson, 2001; Thorén, 2007). In fact, plants can also affect the function and structure of bacterial community in sediments, especially in rhizosphere (Herrmann et al., 2008). Despite the important ecological roles of aquatic plant-epibiotic bacteria system, in aquatic ecosystems, the community ecology of these bacteria is far from being understood (He et al., 2014). The functions of biofilms are always determined by the biodiversity and species of bacteria forming it. The epiphytic bacterial communities were reported to be diverse and host-specific (Crump and Koch, 2008; He et al., 2012), while the growth status and secretions of macrophytes can affect the bacterial composition in biofilms and sediments (Hempel et al., 2008; Jensen et al., 2007). By using high-through sequencing methods (454 pyrosequenc-
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ing), He et al. (2014) found that bacteria community structures in epiphytic biofilm are different from those in surrounding water column. These references provided valuable information that biofilms can be formed on aquatic macrophytes surface and have roles in nitrogen removal. However, to the present authors’ knowledge, minimum information is available about the structure and functions of microbial communities on aquatic plants, nor in the rhizosphere in Lake Ecosystems, at the meta-genomic level. Lake Hongze, the fourth largest freshwater lake of China, is a moderately eutrophic freshwater Lake with an average water depth of 1.4 m. In recent years, many wetlands were constructed by restoration of abundant floating and submersed macrophytes in lake area to improve the water quality and biodiversity. In order to investigate structure of bacterial communities in biofilms from different organs of common floating and submersed macrophytes and in vegetated and unvegetated vertical sediments and to compare nitrifying and denitrifying bacteria composition in biofilms to those in the sediments, 454 pyrosequencing methods were employed to detect the microbial communities in biofilm attached to the leaves, stems (except for Myriophyllum verticillatum) or roots of M. verticillatum, Nymphoides peltatum and Trapa japonica, and in sediments from vegetated or unvegetated area in Lake Hongze. The results were expected to improve our understanding the role of aquatic macrophytes in the functions of wetlands. 2. Materials and methods 2.1. Study sites and sampling M. verticillatum, N. peltatum and T. japonica were the dominant aquatic macrophytes in the sampling sites of a wetland located in Lake Hongze. Plant samples and sediment samples were collect in July 2013 because aquatic plants always thrive in summer. Plants were collected without brushing, and floating macrophyte N. peltatum and T. japonica samples were separated into leaves, stems and roots, while submersed macrophyte M. verticillatum samples were separated into leaves and roots. The root samples were washed with double distilled H2 O. Three replicates were collected for each plant sample and each replicate contained at least 50 plants. The M. verticillatum is rooted in sediments while roots of N. peltatum and T. japonica always suspend in water column (Supplementary Fig. S1). Surface water (2 L each site, three replicates were collected) was sampled from the macrophytes dominated area. Undisturbed vertical sediment cores were sampled using a Beeker sampling device (Eijkelkamp Agrisearch Equipment, Netherlands). The average depth of sediments in this lake was approximately 30 cm around these sampling sites. Three sediment cores (30 cm) were collected from sampling site without plants (water depth, 0.6 m) or with high density of M. verticillatum (water depth, 1.2 m). Core vertical sediment samples were divided into nine layers (3 cm per layer), and the segment layers of 0–3 cm, 10–12 cm and 21–24 cm were selected to represent the surface, middle and bottom layer of the sediment, respectively. The sediment samples were stored in aseptic plastic bags, and all samples were kept in an ice box and were transported to the lab within 24 h. Nutrient concentrations in sediments and surface water were provided as Supplementary materials (Table S1). 2.2. Sample pre-treatment Approximately 50 g of fresh plant material were transferred into a sterile 500 mL polyethylene bottle containing 400 mL of 50 mM phosphate buffered saline (PBS, pH = 7.4) solution. Epiphytic microbes were detached after 3 min of ultra-sonication, 30 min of
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shaking (225 r/min) and subsequent ultra-sonication for 3 min (He et al., 2012). The surface area of leaves was calculated by using image software Adobe Photoshop CS3 (Adobe Systems Software Ireland Ltd, American). For bacteria counting, formaldehyde was added in the 5 mL of eluents or surface water samples to a final concentration of 2% formaldehyde, while 5 mL eluent and water samples were added with Lugo’s reagent for algae identification. Plant leaves for microscopic observation were fixed in a final concentration of 2% formaldehyde in 50 mM PBS solution. 2.3. Measurements of the bacteria and algae 100 L eluent or water sample was further mixed with 700 L 4 ,6-diamino-2-phenylindole (DAPI, 10 g/mL) and incubated in the dark for 30 min. The samples were filtered through a 0.22 m incubated filter. The number of bacteria on the black membrane was counted under a fluorescence microscope (ZEISS, Germany) and for each slide, thirty random fields were enumerated. 100 L eluent or water sample was dropped in the middle of the plankton counting chamber (Beijing Purity instrument CO., LTD, China) and then were observed under a fluorescence microscope (ZEISS, Germany). Fifty random fields were counted for each sample. The algae were identified at genus level based on morphological parameters. Triplicates were performed for each sample. 2.4. Scanning electron microscopy (SEM) analysis Plant leaves were cut into small pieces (5mm × 5 mm). After being dehydrated through a series concentration of ethanol (in sequence 30%, 50%, 70%, 80% and 90%) for 15 min at each concentration, these small pieces were immersed in 100% ethanol twice for 15 min each and then transferred to a freeze-drier. The dried samples were visualized by using a scanning electron microscopy after being sputter coated with gold (S3400, Hitachi, Japan). 2.5. DNA extraction, PCR amplification and 454 pyrosequencing The biofilm solution or the sediment samples were mixed with ethyl alcohol at 1:2 ratio (v:v) and then centrifuged at 8000 rpm for 10 min. All the condensed samples were collected and stored at −80 ◦ C for DNA extraction. Biofilm and sediment DNA (three replicates per sample) were extracted with the PowerBiofilm DNA Isolation kit and the PowerSoil DNA Isolation kit (MoBio Laboratories, USA) according to manufacturer’s protocols, respectively. Three extracted DNA solutions from the same sample were combined for the following analysis. The 16S rRNA gene of bacteria was amplified with the primer 342F (5 -CTACGGGGGGCAGCAG-3 ) and 806R (5 -GGACTACCGGGGTATCT-3 ) (Mori et al., 2014). PCR reactions were performed in a 20 L mixture, containing 4 L of 5 × FastPfu Buffer, 2 L of 2.5 mM dNTPs, 0.8 L of each primer (5 M), 0.4 L of FastPfu Polymerase, and 10 ng of template DNA. The amplification program consisted of an initial denaturation step at 95 ◦ C for 2 min, followed by 25 cycles, where 1 cycle consisted of 95 ◦ C for 30 s (denaturation), 55 ◦ C for 30s(annealing) and 72 ◦ C for 30 s (extension), and a final extension at 72 ◦ C for 5 min. Before sequencing, each PCR product was purified with the AxyPrep DNA Gel Extraction Kit (Axygen Biosciences, Union City, CA, U.S.) and quantified using QuantiFluorTM −ST (Promega, U.S.). Amplicons from different samples were then mixed to achieve equal mass concentrations in the final mixture, which were used for pyrosequencing on a Roche 454 GS FLX+ Titanium platform (Roche 454 Life Sciences, Branford, CT, U.S.) according to standard protocols. The low quality sequences with average quality score <20 over a 50 bp sliding window and sequences shorter than 200 bp, with homopolymers longer than six nucleotides, and con-
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taining ambiguous base calls or incorrect primer sequences were removed from the pyrosequencing-derived data sets. The high-quality sequences were assigned to samples according to barcodes. Sequences were aligned in accordance with SILVA alignment (Pruesse et al., 2007) and clustered into operational taxonomic units (OTUs). OTUs that reached 97% similarity level were used for diversity (Claeson et al., 2013), richness (Ace), Good’s coverage, and Rarefaction curve analysis by using Mothur (version 1.5.0 http://www.mothur.org/wiki/Schloss SOP#Alpha diversity) (Schloss et al., 2011). The phylogenetic affiliation of each 16S rRNA gene sequence was analyzed by RDP Classifier (http://rdp.cme.msu.edu/) against the silva (SSU115) 16S rRNA database using confidence threshold of 70% (Amato et al., 2013). The pyrosequencing reads have been deposited at the GenBank under accession number SRX1114590, SRX1114639, SRX1114640, SRX1114797, SRX1115235, SRX1115260, SRX1115270, SRX1115275, SRX1115276, SRX1115279, SRX1115282, SRX1115309, SRX1115330, SRX1115340, in the bioproject accession number PRJNA289748. 2.6. Statistical analysis One-way ANOVA with Dunnett’s multiple comparisons test (p < 0.01) was employed to analyze the microbe and algae densities. The microbial community barplot (phylum and genus) was used to compare bacterial community structures across all the samples, while the cluster analysis and principal component analysis (PCA) were used to measure the similarity between the samples. Bray–Curtis indices were calculated and represented in a heat map format to depict the similarity and dissimilarity between bacterial communities. Other data were analyzed with Origin 8.0. 3. Results 3.1. Characterization of bacteria and algae in surface water and biofilm Fig. 1 Biofilm morphology on leaves of three aquatic plants was monitored by using SEM with a magnification factor of 2,000 and 10,000. Abundant algae and bacteria inhabited the interface between leaves and water (Fig. 1). Cocconeis, Surirella, Synedra and Bacillus were detected on M. verticillatum leaves (Fig. 1A and 1D). Cocconeis, Cymbella and Bacillus were observed on N. peltatum leaves (Fig. 1B and 1E) and Nitzschia, Bacillus, cocci and filamentous bacteria (Fig. 1C and 1F) were noted on T. japonica leaves. Fig. 2 The planktonic bacteria and algae densities in water column were 3.26(± 0.06) × 105 and 2.16(± 0.11) × 107 cells/L, respectively. The density of epiphytic bacteria was approximately 105 -106 cells cm−2 on leaves of three plants (Fig. 2). The densities of epiphytic bacteria and algae on leaves of M. verticillatum were higher than that of N. peltatum and T. japonica. A total of 33 algae genera were detected in water and biofilm samples from Lake Hongze. In surface water 24 algae genera were observed, while 20, 15 and 17 algae genera were observed on the leaves of M. verticillatum, N. peltatum and T. japonica, respectively (Supplementary Fig. S2). Chlorella and Microcystis were dominant algae genera in all the four samples. 3.2. Characterization of bacterial community in biofilm and sediments Fig. 3A total of 152,181 high-quality sequences (with 427 bp of average length) trimmed from 170,280 valid sequences were used for further analysis (Supplementary Table S3). The richness indices of ACE ranged from 684.76 to 2661.84 in fourteen samples. The Shannon index ranged from 3.18 (stem of T. japonica) to 5.74
(root of M. verticillatum) in biofilm samples and from 5.56 (middle layer of vegetated sediment) to 6.49 (surface layer of unvegetated sediment) in sediment cores. The Good’s coverage (94–98%) and r- Shannon curve showed these trimmed sequences can represent the species richness of each sample (Supplementary Fig. S5). In total, 42 bacterial phyla were detected in the fourteen samples (Supplementary Table S4). Proteobacteria was the most dominant phylum in biofilm (36.49–83.21%) and sediments (26.81–35.55%), followed by Bacteroidetes (7.79–37.80%), Chloroflexi (1.98–9.00%), Firmicutes (0.99–16.01%) and Verrucomicrobia (0.80–12.67%) in biofilm and by Chloroflexi (16.00–21.79%), Acidobacteria (5.01–19.64%), Bacteria unclassified (1.61–17.87%), Firmicutes (2.20–7.88%), Bacteroidetes (1.58–7.62%) and Verrucomicrobia (1.02–6.42%) in sediments. The percentages of unclassified bacteria and Candidate division OP8 phylum were higher in sediment samples than in the biofilm samples, especially in the vegetated sediment samples (Supplementary Table S4). In phylum Proteobacteria, Alphaproteobacteria (9.31-29.99%), Betaproteobacteria (2.02-15.84%) and Gammaproteobacteria (11.41–71.76%) dominated in the biofilm samples while (0–1.41%) and Deltaproteobacteria Epsilonproteobacteria (0.07–2.12%) occurred at a low frequency. In sediment samples, the percentage of Betaproteobacteria (7.28–14.09%), Gammaproteobacteria (6.64–13.73%), Epsilonproteobacteria (0.01–7.86%) and Deltaproteobacteria (5.53–8.31%) was higher than that of Alphaproteobacteria (0.25–2.16%) (Supplementary Table S5). It should be noted that class Epsilonproteobacteria was higher in vegetated sediments than that in unvegetated sediments and biofilm samples. The mean frequencies of Sphingobacteriia, Flavobacteriia, Verrucomicrobiae and Spartobacteria class were higher but that of Acidobacteria, Anaerolineae, Chloroflexi uncultured, Candidate division OP8 norank, OPB35 soil group and KD4-96 were lower in biofilms than those in sediments (Supplementary Table S5). Fig. 4 At the genus level, 921 genera were detected in all the samples, but only 15 were shared by all the 14 samples. A total of 100 genera were commonly shared by 8 epiphytic bacterial samples, while 149 genera were shared by 6 sediment samples. A total of 48 genera that ranked at the top 10 abundant genera in at least one of the fourteen samples were selected to construct a heat map (Fig. 4). Based on the heat map, most of these genera occurred at a high frequency in biofilms or sediments and only a few genera in both biofilms and sediments. Frequencies of several genera differed obviously among the three plants and among the leaves, stems and roots of the same macrophyte. For example, the frequencies of genera Caldilineaceae uncultured, Chryseobacterium and Cloacibacterium were higher in biofilm from leaves than those from roots of M. verticillatum. Frequency of Rhodobacter was higher in biofilm from M. verticillatum than that from other plants, while genus Enterobacteriaceae unclassified was lower in biofilm from M. verticillatum than those from other plants (Supplementary Fig. S6 and Table S6). The proportions of Subgroup 19 norank, Candidate division OD1 norank, Verrucomicrobium and Kazan-3B09 norank were higher in the unvegetated sediments than in the vegetated sediments, while the relative abundance of BD211 terrestrial group norank, Candidatus Acetothermus, Prosthecomicrobium, SubsectionI FamilyI norank and vadinBC27 wastewatersludge group were higher in the vegetated sediments than in the unvegetated sediments. 3.3. Cluster and PCA analysis Fig. 5 As revealed by cluster analysis, at genus level, bacterial community structures in biofilm were different from those in sediments (Fig. 5A) and four sub-groups were generated. Sed-
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Fig. 1. Scanning electron micrographs of the epiphytic biofilm on the leaves of M. verticillatum (a and d), N. peltatum (b and e) and T. japonica (c and f) under 2000 (a, b and c) and 10000 (d, e and f).
Fig. 2. Density of epiphytic bacteria (a) and epiphytic algae (b) on different macrophytes. Different lowcase letters indicates significant differences (p < 0.01).
iment samples from vegetated areas and unvegetated areas were clustered into group I and group II, respectively. The bacterial community structures in biofilms from floating plants N. peltatum and T. japonica were clustered into group III, while those from submersed plant M. verticillatum were in group IV. For the same plants the bacterial community structures from the leaves were more similar to those from stems than from roots, and for the sediments, bacterial community structures in bottom layer were more similar to those from middle layer than from surficial layer. Four groups (Fig. 5B) generated from PCA, which expressed 59.97% of the total variability. Members in each group were the same as those in groups from cluster analysis (Fig. 5A). It should be noted that samples from the unvegetated sediments almost overlapped together with those from the vegetated sediments.
3.4. Microorganisms associated with nitrogen cycle Table 1 The percentages of nitrifying bacteria and denitrifying bacteria in the nitrogen cycle bacteria were selected (Table 1). The nitrifying bacteria including Nitrospina, Nitrospira and Nitrosococcus
occurred in 0.01-3.90% of total bacteria reads. The relative abundance of nitrifying bacteria in the unvegetated sediments was five times higher than in the vegetated sediments. The denitrifying bacteria were found in phyla Bacteroidetes, Firmicutes, Nitrospinae and Proteobacteria, and most of the denitrifying bacteria belonged to Proteobacteria. The relative abundance of the denitrifiers ranged from 6.85% to 19.23% in the biofilm samples, but averaged only 2.86% in the sediment samples.
4. Discussion 4.1. Floating- and submersed-macrophytes provide special niches for microbes Biofilm is an irreplaceable component of primary production in macrophyte-dominated aquatic ecosystem. In the present study, the algae composition in the surrounding water was different from those in biofilm attached to plants (Supplementary Fig. S4). A recent report (He et al., 2014) showed a marked divergence in the community structures between epibiotic bacteria and bacterioplankton.
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Fig. 3. Bacterial composition of the different communities. Relative read abundance of different bacterial phyla within the different communities. Sequences that could not be classified into any known group were assigned as ‘Bacteria unclassified’. ‘Others’ refer to the taxa with their maximum abundance <1% in any sample.
They suggested that leaves of submerged macrophytes might serve as ‘concentrators’ of bacterial communities, by supplying more nutrients and micro-niches for the migrating bacteria cells. Abundant bacteria and algae were detected in biofilms attached to leaves of three aquatic plant plants (Fig. 1 and 2), supporting that bacteria and algae are major components of epiphytic microbes on the macrophytes (He et al., 2014; Hempel et al., 2008). The epiphytic algae density on the leaves of submersed macrophyte M. verticillatum was significantly higher than that of floating macrophyte N. peltatum and T. japonica (Fig. 2). Neif et al. (2013) found that the mean density of epiphyton was higher on submerged macrophyte E. najas than that on emergent macrophyte E. azurea. Accumulated data showed that submersed macrophytes have a complex architecture and a larger accessible surface area for epiphytic microbes than the floating plants, offering different opportunities for the epiphyton establishment (Cattaneo et al., 1998). Allelopathically active and plant exudates (such as polyphenols) may be another reason for the difference in epiphyte biomass and microbial composition (Hempel et al., 2008). Taken together, these data suggest that the remediation of aquatic plants in water system can increase the microbial biodiversity and quantity of biofilms, which play a key role in energy transduction and uptake or retention of nutrients in aquatic ecosystems (Battin et al., 2003).
4.2. Bacteria composition appears to be diverse and somewhat host-specific The mean alpha diversities parameters (Ace index and Shannon index) of bacterial communities overall were lower in biofilm than in sediments (Supplementary Table S3). The alpha diversities of biofilm in this study were higher than those on leaves of Potamogeton crispus reported by He et al. (2014), who found that bacteria in biofilm from P. crispus exhibited higher alpha diversity than bacterioplankton. Similar to phyla on leaves of lettuce (Rastogi et al., 2012) and P. crispus (He et al., 2014), Proteobacteria, Bacteroidetes, Chloroflexi, Firmicutes, Actinobac-
teria and Verrucomicrobia were the most abundantly represented phyla on the three aquatic plants (Fig. 3 and Supplementary Table S4). The compositions of dominant classes (Cytophaga, Flavobacteria, Bacteroidetes, Alphaproteobacteria, Betaproteobacteria, Gammaproteobacteria and Actinomycetes) in biofilm of the present study were different from those revealed by using FISH (Fluorescence in situ hybridization) method in M. spicatum and macroalgae (Hempel et al., 2008) but somewhat similar to that by using 454 pyrosequencing method in P. crispus (He et al., 2014). Bacterial classes Alphaproteobacteria and Gammaproteobacteria dominated in biofilms in the present study were also found to dominate on other macrophytes (Burke et al., 2011; He et al., 2014; Tujula et al., 2010). He et al. (2012) compared the microbial communities by using T-RFLP and found Betaproteobacteria was the most abundant on V. natans and H. verticillata plants, while Cyanobacteria and Gammaproteobacteria was the second largest group on V. natans and H. verticillata, respectively. In the present study Cyanobacteria accounted for only 0.30% on average (Supplementary Table S4). Epiphytic bacterial communities on macrophytes appear to be diverse and host-specific, except for the samples from the roots of two floating plants (Fig. 5). Similar phenomena were also found in aquatic (Crump and Koch, 2008; He et al., 2012) and terrestrial plants (Rastogi et al., 2012). Differences were observed in the bacterial communities among four species of aquatic angiosperm plant and between leaves and roots (Crump and Koch, 2008). The difference in bacterial communities may be attributed to the spatial variation. The leaves of N. peltatum and T. japonica always float on the surface of the water, while the whole plant of M. verticillatum is submersed in the water (Supplementary Fig. S2). The different growth environments may exhibit differences in water flow, illumination conditions and nutrient concentration and therefore lead to the difference in bacterial communities (He et al., 2012). Additionally, microbes in biofilm attached to roots inhabit particular niches, where nutrient source and environmental parameters can be affected by plant roots, surface water, sediment composition and even aquatic animals. Therefore, the species richness and diversity
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Fig. 4. Heat map of top 10 genera in each sample. The top 10 abundant genera in each sample were selected (a total of 48 genera for all 14 samples) and compared with their relative abundances (percentages) in other samples. The color intensity (log scale) in each panel shows the percentage of a genus in a sample, referring to color key at the bottom.
of the epiphytic bacteria were commonly higher on roots than those on leaves and stems (Supplementary Table S3). 4.3. Aquatic macrophyte roots potentially alter the bacterial community structure of sediments Bacteria are involved in the transformation of complex organic compounds and minerals in sediments. The microbial communities in biofilms were different from those in sediments (Fig. 4
and 5). For example, percentages of class Alphaproteobacteria on average were lower in sediments (1.19% ± 0.81%) than in biofilm (18.61 ± 6.69%), while Epsilonproteobacteria (3.12 ± 3.42%) and Deltaproteobacteria (6.87 ± 1.18%) were mainly detected in sediments (Supplementary Table S5). In general, the dominant bacteria phyla in vertical sediments in this study were similar to those in sediments from Lake Taihu, China (Chen et al., 2015) and Dianchi, China (Bai et al., 2012). However, at the genus level, bacterial community composition always varied in different locations of the
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Fig. 5. Cluster (a) and principal component analysis (b) of bacterial communities. S-VS: vegetated surface sediment; M-VS: vegetated middle sediment; B-VS: vegetated bottom sediment; S-US: unvegetated surface sediment; M-US: unvegetated middle sediment; B-US: unvegetated bottom sediment; L-MV: leaves of M. verticillatum; R-MV: roots of M. verticillatum; L-NP: leaves of N. peltatum; S-NP: stems of N. peltatum; R-NP: roots of N. peltatum; L-TJ: leaves of T. japonica; S-TJ: stems of T. japonica; R-TJ: roots of T. japonica.
same lake, different lakes, and even different seasons (Chen et al., 2015). A recent report (Bai et al., 2012) showed that in sediments of Lake Dianchi, the most abundant classified bacterial genera in the high organic carbon sediment were different from those in low organic carbon sediments. These differences can be ascribed to the differences in environment parameters (i.e. water flow, dissolved oxygen, disturbance) and nutrient level. The structures of bacterial communities in the vegetated sediments were different from those in the unvegetated sediments (Fig. 4). Though it was impossible to provide enough evidence to confirm whether these differences were caused by the activity of aquatic plant roots alone, as revealed by PCA (Figue 5b), bacterial community composition was obviously different among the three vertical layers of the sediments from vegetated areas as compared to those from the unvegetated areas. Jensen et al. (2007) also found a significant difference between the bacterial communities associated with the seagrass roots and the surrounding bulk sediment. These differences may be ascribed to the differences in organic material accumulation, oxygen and chemical exudates in the rhizosphere zone in the vegetated area (Faulwetter et al., 2013). Therefore, the roots of these plants may have altered the bacterial community in the surrounding sediments. 4.4. Bacteria community in nitrogen-cycle Microorganisms, including nitrifying bacteria and denitrifying bacteria, play important roles in the nitrogen cycles of various ecosystems through nitrification and denitrification processes (Hayatsu et al., 2008). Though nitrification activity and nitrifying bacteria were reported on the surface of submersed macrophytes (Trias et al., 2012), detailed information on these bacteria remains unclear. In the present study, three genera of nitrifiers Nitrosospira, Nitrospina and Nitrosococcus were detected; Nitrosospira was detected only in biofilm, while Nitrospina and Nitrosococcus were detected only in sediments (Table 1). Nitrifiers prefer high dissolved oxygen concentration areas, but percentages of nitrifiers were lower in the vegetated sediments (0.62–0.72%) than in unvegetated sediments (3.22–3.90%) (Table 1). Herrmann et al. (2008) found a slightly lower relative abundance of ammonia-oxidizing bacteria in the rhizosphere of L. uniflora than in the unvegetated sediments. In the present study, the unvegetated sediment sam-
ples were taken from shallower areas, which is beneficial for the nitrifying bacteria (Flood et al., 2015). However, nitrifying bacteria occurring at 0.01-0.06% in biofilm may be ascribed to the low ammonium concentration in the water column (Table S1). In fact, the abundances of ammonia- and nitrite- oxidizing bacteria usually represent less than 1% of the total bacterial population, for example, in activated sludge of full-scale wastewater treatment plants, but they are extremely important for nitrogen removal. Denitrifying bacteria reduce nitrates to molecular nitrogen or gaseous nitrogen. A total of eighteen genera containing these bacteria, or identified as denitrifiers were summarized (Table 1). Rhodobacter was the primary denitrifier in biofilm from the leaves (11.32%) and roots (7.50%) of M. verticillatum, while Acinetobacter and Pseudomonas dominated in most of the macrophyte samples (accounting for 6.71% on average). Nitrospinaceae was the most dominant denitrifier in all sediment samples, yet the proportions of other denitrifying genera were extremely low. The potential dentrifiers Pseudomonas, Comamonas, Achromobacter, Alcaligenes and Flavobacterium were also detected in the previous studies (Drysdale et al., 1999; Gumaelius et al., 2001; Hayatsu et al., 2008; Lai and Shao, 2008). In the present study, occurrence of denitrifying bacteria in the bacterial community ranged from 1.95% to 3.70% in sediments and from 6.85% to 19.23% in biofilm (Table 1). Eriksson and Weisner (1997) found that the level of denitrification can be relatively higher in epiphytic microbial communities than in those in sediments. Abovementioned results highlight the import role of submersed macrophyte-biofilm system on nitrogen cycles. Plants can release oxygen through photosynthesis system at the daytime, while plants and most organisms in biofilm consume oxygen through respiration in their lifetime. Therefore, aerobic and low oxygen micro-environment can possibly be formed at the interface between plant and water during the day and the night, respectively, and a possible aerobic-anaerobic micro-environment may be formed at the rhizosphere-sediment interface. These micro-environments are beneficial to nitrifying and denitrifying organisms. However, the data are still not adequetly robust explain the role of aquatic plants in modifying the ntrogen-cycle in freshwater the column, because ntrogen-cycle related archaea and fungus may also exist in the biofilms (Hayatsu et al., 2008). For example, epiphytic ammonia-oxidizing bacteria and archae
Bottom
2.69 – 1.21 3.90 0.05 – – 0.11 – 0.83 – 0.06 0.64 – – – 0.01 1.72 – – – 0.09 3.51
Middle
1.95 – 1.27 3.22 0.04 – – 0.04 0.01 1.19 – 0.04 0.70 – 0.06 – 0.01 1.47 – – – 0.14 3.70 2.55 – 1.29 3.84 0.02 – – 0.04 – 0.80 – – 0.42 – 0.07 – – 1.46 – – 0.01 0.20 3.02
Surface
5. Conclusions Total microbe and algae concentrations on leaves of submersed macrophyte were higher than those of two floating macrophytes in this wetland system. The structures of bacterial communities in sediments are markedly different from those in the biofilms attached on the three aquatic macrophytes, and there is an obvious difference in bacterial community between the vegetated and unvegetated sediments. These results suggest that three aquatic macrophytes provided special niche for microbes in water column and sediments. The relative abundance of nitrifying bacteria were higher in sediment than in biofilm, while that of denitrifying bacteria were markedly higher in biofilm than in sediments. However, the role of epiphytic microbial communities in nitrogen removal should be furtherly investigated in the future.
0.54 – 0.15 0.69 0.03 – – – – 0.33 – 0.04 0.03 – – – – 1.55 – – – – 1.98
Bottom
0.46 – 0.26 0.72 – – – – – 0.38 – 0.02 0.05 – 0.01 – – 2.53 – – – 0.01 3.00 – 0.06 – 0.06 0.05 4.34 – 0.12 0.54 0.12 0.04 – 3.37 – 0.06 0.43 0.31 0.05 2.68 0.20 1.66 – 13.97
Middle Surface Root
– 0.01 – 0.01 – 13.19 – 0.01 0.43 – – – 0.47 0.01 – 0.04 0.13 – 0.70 0.39 1.27 – 16.64 – 0.02 – 0.02 – 2.61 – 0.03 0.26 – 0.05 0.61 3.06 0.03 – 0.31 0.04 0.06 2.56 0.20 0.43 – 10.25 – 0.06 – 0.06 – 3.15 – 0.08 0.06 – 0.05 0.07 3.18 0.08 – 0.43 0.12 – 7.10 0.83 0.41 0.01 15.57
Stem Leaf Stem Leaf
Conflict of interest The authors declare no conflict of interest.
– 0.02 – 0.02 – 4.36 – – 0.62 0.10 0.01 – 1.86 0.08 – 0.29 0.23 – 0.90 0.73 1.75 – 10.93
This study was, in part, supported by Grants from the National Natural Science Foundation of China (Grant No. 51379063 and 51579075), Science Fund for Creative Research Group of the National Natural Science Foundation of China (51421006), and Innovation Project from Ministry of Education of China (IRT13061), the Fundamental Research Funds for the Central Universities (2016B06714), Jiangsu water resource technology project (2014076 and 2015085) and a Project Funded by the Priority Academic Program Development of Jiangsu Higher Education Institutions.
– 0.04 – 0.04 – 3.41 – 0.01 0.20 0.01 0.34 – 2.45 0.32 0.01 0.53 1.37 0.01 1.20 1.87 7.50 – 19.23
– 0.03 – 0.03 – 0.89 0.14 0.05 0.20 – 0.04 0.15 0.90 – – 0.33 0.07 0.01 2.14 0.14 1.79 – 6.85
Root
– – – – – 3.70 – – 0.76 0.04 – – 0.26 0.01 – 0.08 0.60 0.02 0.76 0.89 11.32 – 18.44
Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.ecoleng.2016.10. 011. References
Total
Achromobacter Acinetobacter Alcaligenes Azospira Bacillus Comamonas Dechloromonas Desulfovibrio Flavobacterium Flexibacter Halomonas Hydrogenophaga Hyphomicrobium Nitrospinaceae uncultured Pseudomonas Rhizobium Rhodobacter Sphingobacterium Total Denitrifying bacteria
Nitrifying bacteria
Nitrospina (NOB) Nitrospira (NOB) Nitrosococcus (AOB NOB)
Leaf
Root
Acknowledgements
’–’ Indicates genus was not detected.
Vegetated sediment T. japonica N. peltatum M. verticillatum Genus
Table 1 The abundance of bacteria associated with nitrogen cycling in each sample (percentages).
249
were detected on marine macroalgae (Trias et al., 2012), and the abundance of ammonia-oxidizing archaea was higher than ammonia-oxidizing bacteria in the rhizosphere of aquatic plants (Chen et al., 2008; Herrmann et al., 2008). In addition, the function of abundant bacteria remains unclear, especially for those unclassified bacteria (Supplementary Table S6). Therefore, further study is demanded to explore this subject in future.
0.46 – 0.16 0.62 – – – 0.01 – 0.25 – – 0.04 – 0.01 – – 1.63 – – – 0.01 1.95
Unvegetated sediment
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Ecological Engineering journal homepage: www.elsevier.com/locate/ecoleng
Characterization of bacterial community in biofilm and sediments of wetlands dominated by aquatic macrophytes Si Pang a , Songhe Zhang a,∗ , XiaoYang Lv a , Bing Han a , Kaihui Liu a , Changhao Qiu a , Chao Wang a , Peifang Wang a,∗ , Harry Toland b , Zhenli He c a Ministry of Education Key Laboratory of Integrated Regulation and Resource Development on Shallow Lakes, College of Environment, Hohai University, Nanjing 210098, China b Geography & Earth Sciences, Aberystwyth University, Llandinam Building, Penglais Campus, Aberystwyth, Wales, SY23 3DB, United Kingdom c University of Florida, Institute of Food and Agricultural Sciences, Indian River Research and Education Center, 2199 South Rock Road, Fort pierce, FL 34945, USA
a r t i c l e
i n f o
Article history: Received 27 September 2015 Received in revised form 16 July 2016 Accepted 5 October 2016 Keywords: Biofilm Macrophytes Sediment Nitrogen cycle
a b s t r a c t Though aquatic macrophytes play an important role in wetlands, their effects on bacteria community structures in biofilms and sediments are far from clear. In the present study, bacterial communities were investigated in biofilm attached to leaves, stems and roots of aquatic macrophytes (Myriophyllum verticillatum, Nymphoides peltatum and Trapa japonica) and in vertical sediment cores from vegetated and unvegetated areas in a wetland located in Lake Hongze. The densities of microbes and epiphytic algae in biofilms were higher on leaves of M. verticillatum than those of two floating macrophytes. Phyla Proteobacteria, Bacteroidetes, Chloroflexi, Firmicutes and Verrucomicrobia were detected in both biofilms and sediments. As revealed by cluster analysis and principal component analysis, differences in structures of microbial communities were detected between biofilms and sediments and between vegetated and unvegetated sediments. The potential roles of nitrifying- and denitrifying- bacteria in sediments with respect to those in biofilms were discussed. These results highlight that the restoration of aquatic macrophytes can increase bacteria diversity and the surface and quantity of biofilms and therefore bacteria diversity. These data provide useful information for further understanding the role of aquatic macrophyte-biofilm system in wetlands in future. © 2016 Elsevier B.V. All rights reserved.
1. Introduction Aquatic macrophytes, key components of aquatic ecosystems, can absorb nutrients, heavy metals and other inorganic or organic contaminants from the water column and the sediments (Dhote and Dixit, 2009). Aquatic macrophytes are widely grown in wastewater treatment wetlands for improving water quality (Thorén, 2007). However, due to the improper anthropogenic activities aquatic macrophytes have declined significantly and biodiversity in freshwater has decreased, this will be a major driver of future ecosystem change (Hooper et al., 2012). Biofilms (known as epiphytic microbes) can form on the gasliquid and solid-liquid interfaces, and potentially constitute an important step in the integration of biogeochemical cycles and
∗ Corresponding authors. E-mail addresses: [email protected] (S. Zhang), [email protected] (P. Wang). http://dx.doi.org/10.1016/j.ecoleng.2016.10.011 0925-8574/© 2016 Elsevier B.V. All rights reserved.
dynamics of microbes to ecosystem function (Battin et al., 2003). Aquatic plants are key components in spatial heterogeneity, which is essential for the establishment and development of biofilms in aquatic ecosystems. Aquatic plants represent a special substrate for biofilms, as they can release oxygen, which is beneficial to aerobic bacteria attached to the plant, and promotes the transformation of nitrogen in water (Eriksson, 2001; Thorén, 2007). In fact, plants can also affect the function and structure of bacterial community in sediments, especially in rhizosphere (Herrmann et al., 2008). Despite the important ecological roles of aquatic plant-epibiotic bacteria system, in aquatic ecosystems, the community ecology of these bacteria is far from being understood (He et al., 2014). The functions of biofilms are always determined by the biodiversity and species of bacteria forming it. The epiphytic bacterial communities were reported to be diverse and host-specific (Crump and Koch, 2008; He et al., 2012), while the growth status and secretions of macrophytes can affect the bacterial composition in biofilms and sediments (Hempel et al., 2008; Jensen et al., 2007). By using high-through sequencing methods (454 pyrosequenc-
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ing), He et al. (2014) found that bacteria community structures in epiphytic biofilm are different from those in surrounding water column. These references provided valuable information that biofilms can be formed on aquatic macrophytes surface and have roles in nitrogen removal. However, to the present authors’ knowledge, minimum information is available about the structure and functions of microbial communities on aquatic plants, nor in the rhizosphere in Lake Ecosystems, at the meta-genomic level. Lake Hongze, the fourth largest freshwater lake of China, is a moderately eutrophic freshwater Lake with an average water depth of 1.4 m. In recent years, many wetlands were constructed by restoration of abundant floating and submersed macrophytes in lake area to improve the water quality and biodiversity. In order to investigate structure of bacterial communities in biofilms from different organs of common floating and submersed macrophytes and in vegetated and unvegetated vertical sediments and to compare nitrifying and denitrifying bacteria composition in biofilms to those in the sediments, 454 pyrosequencing methods were employed to detect the microbial communities in biofilm attached to the leaves, stems (except for Myriophyllum verticillatum) or roots of M. verticillatum, Nymphoides peltatum and Trapa japonica, and in sediments from vegetated or unvegetated area in Lake Hongze. The results were expected to improve our understanding the role of aquatic macrophytes in the functions of wetlands. 2. Materials and methods 2.1. Study sites and sampling M. verticillatum, N. peltatum and T. japonica were the dominant aquatic macrophytes in the sampling sites of a wetland located in Lake Hongze. Plant samples and sediment samples were collect in July 2013 because aquatic plants always thrive in summer. Plants were collected without brushing, and floating macrophyte N. peltatum and T. japonica samples were separated into leaves, stems and roots, while submersed macrophyte M. verticillatum samples were separated into leaves and roots. The root samples were washed with double distilled H2 O. Three replicates were collected for each plant sample and each replicate contained at least 50 plants. The M. verticillatum is rooted in sediments while roots of N. peltatum and T. japonica always suspend in water column (Supplementary Fig. S1). Surface water (2 L each site, three replicates were collected) was sampled from the macrophytes dominated area. Undisturbed vertical sediment cores were sampled using a Beeker sampling device (Eijkelkamp Agrisearch Equipment, Netherlands). The average depth of sediments in this lake was approximately 30 cm around these sampling sites. Three sediment cores (30 cm) were collected from sampling site without plants (water depth, 0.6 m) or with high density of M. verticillatum (water depth, 1.2 m). Core vertical sediment samples were divided into nine layers (3 cm per layer), and the segment layers of 0–3 cm, 10–12 cm and 21–24 cm were selected to represent the surface, middle and bottom layer of the sediment, respectively. The sediment samples were stored in aseptic plastic bags, and all samples were kept in an ice box and were transported to the lab within 24 h. Nutrient concentrations in sediments and surface water were provided as Supplementary materials (Table S1). 2.2. Sample pre-treatment Approximately 50 g of fresh plant material were transferred into a sterile 500 mL polyethylene bottle containing 400 mL of 50 mM phosphate buffered saline (PBS, pH = 7.4) solution. Epiphytic microbes were detached after 3 min of ultra-sonication, 30 min of
243
shaking (225 r/min) and subsequent ultra-sonication for 3 min (He et al., 2012). The surface area of leaves was calculated by using image software Adobe Photoshop CS3 (Adobe Systems Software Ireland Ltd, American). For bacteria counting, formaldehyde was added in the 5 mL of eluents or surface water samples to a final concentration of 2% formaldehyde, while 5 mL eluent and water samples were added with Lugo’s reagent for algae identification. Plant leaves for microscopic observation were fixed in a final concentration of 2% formaldehyde in 50 mM PBS solution. 2.3. Measurements of the bacteria and algae 100 L eluent or water sample was further mixed with 700 L 4 ,6-diamino-2-phenylindole (DAPI, 10 g/mL) and incubated in the dark for 30 min. The samples were filtered through a 0.22 m incubated filter. The number of bacteria on the black membrane was counted under a fluorescence microscope (ZEISS, Germany) and for each slide, thirty random fields were enumerated. 100 L eluent or water sample was dropped in the middle of the plankton counting chamber (Beijing Purity instrument CO., LTD, China) and then were observed under a fluorescence microscope (ZEISS, Germany). Fifty random fields were counted for each sample. The algae were identified at genus level based on morphological parameters. Triplicates were performed for each sample. 2.4. Scanning electron microscopy (SEM) analysis Plant leaves were cut into small pieces (5mm × 5 mm). After being dehydrated through a series concentration of ethanol (in sequence 30%, 50%, 70%, 80% and 90%) for 15 min at each concentration, these small pieces were immersed in 100% ethanol twice for 15 min each and then transferred to a freeze-drier. The dried samples were visualized by using a scanning electron microscopy after being sputter coated with gold (S3400, Hitachi, Japan). 2.5. DNA extraction, PCR amplification and 454 pyrosequencing The biofilm solution or the sediment samples were mixed with ethyl alcohol at 1:2 ratio (v:v) and then centrifuged at 8000 rpm for 10 min. All the condensed samples were collected and stored at −80 ◦ C for DNA extraction. Biofilm and sediment DNA (three replicates per sample) were extracted with the PowerBiofilm DNA Isolation kit and the PowerSoil DNA Isolation kit (MoBio Laboratories, USA) according to manufacturer’s protocols, respectively. Three extracted DNA solutions from the same sample were combined for the following analysis. The 16S rRNA gene of bacteria was amplified with the primer 342F (5 -CTACGGGGGGCAGCAG-3 ) and 806R (5 -GGACTACCGGGGTATCT-3 ) (Mori et al., 2014). PCR reactions were performed in a 20 L mixture, containing 4 L of 5 × FastPfu Buffer, 2 L of 2.5 mM dNTPs, 0.8 L of each primer (5 M), 0.4 L of FastPfu Polymerase, and 10 ng of template DNA. The amplification program consisted of an initial denaturation step at 95 ◦ C for 2 min, followed by 25 cycles, where 1 cycle consisted of 95 ◦ C for 30 s (denaturation), 55 ◦ C for 30s(annealing) and 72 ◦ C for 30 s (extension), and a final extension at 72 ◦ C for 5 min. Before sequencing, each PCR product was purified with the AxyPrep DNA Gel Extraction Kit (Axygen Biosciences, Union City, CA, U.S.) and quantified using QuantiFluorTM −ST (Promega, U.S.). Amplicons from different samples were then mixed to achieve equal mass concentrations in the final mixture, which were used for pyrosequencing on a Roche 454 GS FLX+ Titanium platform (Roche 454 Life Sciences, Branford, CT, U.S.) according to standard protocols. The low quality sequences with average quality score <20 over a 50 bp sliding window and sequences shorter than 200 bp, with homopolymers longer than six nucleotides, and con-
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taining ambiguous base calls or incorrect primer sequences were removed from the pyrosequencing-derived data sets. The high-quality sequences were assigned to samples according to barcodes. Sequences were aligned in accordance with SILVA alignment (Pruesse et al., 2007) and clustered into operational taxonomic units (OTUs). OTUs that reached 97% similarity level were used for diversity (Claeson et al., 2013), richness (Ace), Good’s coverage, and Rarefaction curve analysis by using Mothur (version 1.5.0 http://www.mothur.org/wiki/Schloss SOP#Alpha diversity) (Schloss et al., 2011). The phylogenetic affiliation of each 16S rRNA gene sequence was analyzed by RDP Classifier (http://rdp.cme.msu.edu/) against the silva (SSU115) 16S rRNA database using confidence threshold of 70% (Amato et al., 2013). The pyrosequencing reads have been deposited at the GenBank under accession number SRX1114590, SRX1114639, SRX1114640, SRX1114797, SRX1115235, SRX1115260, SRX1115270, SRX1115275, SRX1115276, SRX1115279, SRX1115282, SRX1115309, SRX1115330, SRX1115340, in the bioproject accession number PRJNA289748. 2.6. Statistical analysis One-way ANOVA with Dunnett’s multiple comparisons test (p < 0.01) was employed to analyze the microbe and algae densities. The microbial community barplot (phylum and genus) was used to compare bacterial community structures across all the samples, while the cluster analysis and principal component analysis (PCA) were used to measure the similarity between the samples. Bray–Curtis indices were calculated and represented in a heat map format to depict the similarity and dissimilarity between bacterial communities. Other data were analyzed with Origin 8.0. 3. Results 3.1. Characterization of bacteria and algae in surface water and biofilm Fig. 1 Biofilm morphology on leaves of three aquatic plants was monitored by using SEM with a magnification factor of 2,000 and 10,000. Abundant algae and bacteria inhabited the interface between leaves and water (Fig. 1). Cocconeis, Surirella, Synedra and Bacillus were detected on M. verticillatum leaves (Fig. 1A and 1D). Cocconeis, Cymbella and Bacillus were observed on N. peltatum leaves (Fig. 1B and 1E) and Nitzschia, Bacillus, cocci and filamentous bacteria (Fig. 1C and 1F) were noted on T. japonica leaves. Fig. 2 The planktonic bacteria and algae densities in water column were 3.26(± 0.06) × 105 and 2.16(± 0.11) × 107 cells/L, respectively. The density of epiphytic bacteria was approximately 105 -106 cells cm−2 on leaves of three plants (Fig. 2). The densities of epiphytic bacteria and algae on leaves of M. verticillatum were higher than that of N. peltatum and T. japonica. A total of 33 algae genera were detected in water and biofilm samples from Lake Hongze. In surface water 24 algae genera were observed, while 20, 15 and 17 algae genera were observed on the leaves of M. verticillatum, N. peltatum and T. japonica, respectively (Supplementary Fig. S2). Chlorella and Microcystis were dominant algae genera in all the four samples. 3.2. Characterization of bacterial community in biofilm and sediments Fig. 3A total of 152,181 high-quality sequences (with 427 bp of average length) trimmed from 170,280 valid sequences were used for further analysis (Supplementary Table S3). The richness indices of ACE ranged from 684.76 to 2661.84 in fourteen samples. The Shannon index ranged from 3.18 (stem of T. japonica) to 5.74
(root of M. verticillatum) in biofilm samples and from 5.56 (middle layer of vegetated sediment) to 6.49 (surface layer of unvegetated sediment) in sediment cores. The Good’s coverage (94–98%) and r- Shannon curve showed these trimmed sequences can represent the species richness of each sample (Supplementary Fig. S5). In total, 42 bacterial phyla were detected in the fourteen samples (Supplementary Table S4). Proteobacteria was the most dominant phylum in biofilm (36.49–83.21%) and sediments (26.81–35.55%), followed by Bacteroidetes (7.79–37.80%), Chloroflexi (1.98–9.00%), Firmicutes (0.99–16.01%) and Verrucomicrobia (0.80–12.67%) in biofilm and by Chloroflexi (16.00–21.79%), Acidobacteria (5.01–19.64%), Bacteria unclassified (1.61–17.87%), Firmicutes (2.20–7.88%), Bacteroidetes (1.58–7.62%) and Verrucomicrobia (1.02–6.42%) in sediments. The percentages of unclassified bacteria and Candidate division OP8 phylum were higher in sediment samples than in the biofilm samples, especially in the vegetated sediment samples (Supplementary Table S4). In phylum Proteobacteria, Alphaproteobacteria (9.31-29.99%), Betaproteobacteria (2.02-15.84%) and Gammaproteobacteria (11.41–71.76%) dominated in the biofilm samples while (0–1.41%) and Deltaproteobacteria Epsilonproteobacteria (0.07–2.12%) occurred at a low frequency. In sediment samples, the percentage of Betaproteobacteria (7.28–14.09%), Gammaproteobacteria (6.64–13.73%), Epsilonproteobacteria (0.01–7.86%) and Deltaproteobacteria (5.53–8.31%) was higher than that of Alphaproteobacteria (0.25–2.16%) (Supplementary Table S5). It should be noted that class Epsilonproteobacteria was higher in vegetated sediments than that in unvegetated sediments and biofilm samples. The mean frequencies of Sphingobacteriia, Flavobacteriia, Verrucomicrobiae and Spartobacteria class were higher but that of Acidobacteria, Anaerolineae, Chloroflexi uncultured, Candidate division OP8 norank, OPB35 soil group and KD4-96 were lower in biofilms than those in sediments (Supplementary Table S5). Fig. 4 At the genus level, 921 genera were detected in all the samples, but only 15 were shared by all the 14 samples. A total of 100 genera were commonly shared by 8 epiphytic bacterial samples, while 149 genera were shared by 6 sediment samples. A total of 48 genera that ranked at the top 10 abundant genera in at least one of the fourteen samples were selected to construct a heat map (Fig. 4). Based on the heat map, most of these genera occurred at a high frequency in biofilms or sediments and only a few genera in both biofilms and sediments. Frequencies of several genera differed obviously among the three plants and among the leaves, stems and roots of the same macrophyte. For example, the frequencies of genera Caldilineaceae uncultured, Chryseobacterium and Cloacibacterium were higher in biofilm from leaves than those from roots of M. verticillatum. Frequency of Rhodobacter was higher in biofilm from M. verticillatum than that from other plants, while genus Enterobacteriaceae unclassified was lower in biofilm from M. verticillatum than those from other plants (Supplementary Fig. S6 and Table S6). The proportions of Subgroup 19 norank, Candidate division OD1 norank, Verrucomicrobium and Kazan-3B09 norank were higher in the unvegetated sediments than in the vegetated sediments, while the relative abundance of BD211 terrestrial group norank, Candidatus Acetothermus, Prosthecomicrobium, SubsectionI FamilyI norank and vadinBC27 wastewatersludge group were higher in the vegetated sediments than in the unvegetated sediments. 3.3. Cluster and PCA analysis Fig. 5 As revealed by cluster analysis, at genus level, bacterial community structures in biofilm were different from those in sediments (Fig. 5A) and four sub-groups were generated. Sed-
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Fig. 1. Scanning electron micrographs of the epiphytic biofilm on the leaves of M. verticillatum (a and d), N. peltatum (b and e) and T. japonica (c and f) under 2000 (a, b and c) and 10000 (d, e and f).
Fig. 2. Density of epiphytic bacteria (a) and epiphytic algae (b) on different macrophytes. Different lowcase letters indicates significant differences (p < 0.01).
iment samples from vegetated areas and unvegetated areas were clustered into group I and group II, respectively. The bacterial community structures in biofilms from floating plants N. peltatum and T. japonica were clustered into group III, while those from submersed plant M. verticillatum were in group IV. For the same plants the bacterial community structures from the leaves were more similar to those from stems than from roots, and for the sediments, bacterial community structures in bottom layer were more similar to those from middle layer than from surficial layer. Four groups (Fig. 5B) generated from PCA, which expressed 59.97% of the total variability. Members in each group were the same as those in groups from cluster analysis (Fig. 5A). It should be noted that samples from the unvegetated sediments almost overlapped together with those from the vegetated sediments.
3.4. Microorganisms associated with nitrogen cycle Table 1 The percentages of nitrifying bacteria and denitrifying bacteria in the nitrogen cycle bacteria were selected (Table 1). The nitrifying bacteria including Nitrospina, Nitrospira and Nitrosococcus
occurred in 0.01-3.90% of total bacteria reads. The relative abundance of nitrifying bacteria in the unvegetated sediments was five times higher than in the vegetated sediments. The denitrifying bacteria were found in phyla Bacteroidetes, Firmicutes, Nitrospinae and Proteobacteria, and most of the denitrifying bacteria belonged to Proteobacteria. The relative abundance of the denitrifiers ranged from 6.85% to 19.23% in the biofilm samples, but averaged only 2.86% in the sediment samples.
4. Discussion 4.1. Floating- and submersed-macrophytes provide special niches for microbes Biofilm is an irreplaceable component of primary production in macrophyte-dominated aquatic ecosystem. In the present study, the algae composition in the surrounding water was different from those in biofilm attached to plants (Supplementary Fig. S4). A recent report (He et al., 2014) showed a marked divergence in the community structures between epibiotic bacteria and bacterioplankton.
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Fig. 3. Bacterial composition of the different communities. Relative read abundance of different bacterial phyla within the different communities. Sequences that could not be classified into any known group were assigned as ‘Bacteria unclassified’. ‘Others’ refer to the taxa with their maximum abundance <1% in any sample.
They suggested that leaves of submerged macrophytes might serve as ‘concentrators’ of bacterial communities, by supplying more nutrients and micro-niches for the migrating bacteria cells. Abundant bacteria and algae were detected in biofilms attached to leaves of three aquatic plant plants (Fig. 1 and 2), supporting that bacteria and algae are major components of epiphytic microbes on the macrophytes (He et al., 2014; Hempel et al., 2008). The epiphytic algae density on the leaves of submersed macrophyte M. verticillatum was significantly higher than that of floating macrophyte N. peltatum and T. japonica (Fig. 2). Neif et al. (2013) found that the mean density of epiphyton was higher on submerged macrophyte E. najas than that on emergent macrophyte E. azurea. Accumulated data showed that submersed macrophytes have a complex architecture and a larger accessible surface area for epiphytic microbes than the floating plants, offering different opportunities for the epiphyton establishment (Cattaneo et al., 1998). Allelopathically active and plant exudates (such as polyphenols) may be another reason for the difference in epiphyte biomass and microbial composition (Hempel et al., 2008). Taken together, these data suggest that the remediation of aquatic plants in water system can increase the microbial biodiversity and quantity of biofilms, which play a key role in energy transduction and uptake or retention of nutrients in aquatic ecosystems (Battin et al., 2003).
4.2. Bacteria composition appears to be diverse and somewhat host-specific The mean alpha diversities parameters (Ace index and Shannon index) of bacterial communities overall were lower in biofilm than in sediments (Supplementary Table S3). The alpha diversities of biofilm in this study were higher than those on leaves of Potamogeton crispus reported by He et al. (2014), who found that bacteria in biofilm from P. crispus exhibited higher alpha diversity than bacterioplankton. Similar to phyla on leaves of lettuce (Rastogi et al., 2012) and P. crispus (He et al., 2014), Proteobacteria, Bacteroidetes, Chloroflexi, Firmicutes, Actinobac-
teria and Verrucomicrobia were the most abundantly represented phyla on the three aquatic plants (Fig. 3 and Supplementary Table S4). The compositions of dominant classes (Cytophaga, Flavobacteria, Bacteroidetes, Alphaproteobacteria, Betaproteobacteria, Gammaproteobacteria and Actinomycetes) in biofilm of the present study were different from those revealed by using FISH (Fluorescence in situ hybridization) method in M. spicatum and macroalgae (Hempel et al., 2008) but somewhat similar to that by using 454 pyrosequencing method in P. crispus (He et al., 2014). Bacterial classes Alphaproteobacteria and Gammaproteobacteria dominated in biofilms in the present study were also found to dominate on other macrophytes (Burke et al., 2011; He et al., 2014; Tujula et al., 2010). He et al. (2012) compared the microbial communities by using T-RFLP and found Betaproteobacteria was the most abundant on V. natans and H. verticillata plants, while Cyanobacteria and Gammaproteobacteria was the second largest group on V. natans and H. verticillata, respectively. In the present study Cyanobacteria accounted for only 0.30% on average (Supplementary Table S4). Epiphytic bacterial communities on macrophytes appear to be diverse and host-specific, except for the samples from the roots of two floating plants (Fig. 5). Similar phenomena were also found in aquatic (Crump and Koch, 2008; He et al., 2012) and terrestrial plants (Rastogi et al., 2012). Differences were observed in the bacterial communities among four species of aquatic angiosperm plant and between leaves and roots (Crump and Koch, 2008). The difference in bacterial communities may be attributed to the spatial variation. The leaves of N. peltatum and T. japonica always float on the surface of the water, while the whole plant of M. verticillatum is submersed in the water (Supplementary Fig. S2). The different growth environments may exhibit differences in water flow, illumination conditions and nutrient concentration and therefore lead to the difference in bacterial communities (He et al., 2012). Additionally, microbes in biofilm attached to roots inhabit particular niches, where nutrient source and environmental parameters can be affected by plant roots, surface water, sediment composition and even aquatic animals. Therefore, the species richness and diversity
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Fig. 4. Heat map of top 10 genera in each sample. The top 10 abundant genera in each sample were selected (a total of 48 genera for all 14 samples) and compared with their relative abundances (percentages) in other samples. The color intensity (log scale) in each panel shows the percentage of a genus in a sample, referring to color key at the bottom.
of the epiphytic bacteria were commonly higher on roots than those on leaves and stems (Supplementary Table S3). 4.3. Aquatic macrophyte roots potentially alter the bacterial community structure of sediments Bacteria are involved in the transformation of complex organic compounds and minerals in sediments. The microbial communities in biofilms were different from those in sediments (Fig. 4
and 5). For example, percentages of class Alphaproteobacteria on average were lower in sediments (1.19% ± 0.81%) than in biofilm (18.61 ± 6.69%), while Epsilonproteobacteria (3.12 ± 3.42%) and Deltaproteobacteria (6.87 ± 1.18%) were mainly detected in sediments (Supplementary Table S5). In general, the dominant bacteria phyla in vertical sediments in this study were similar to those in sediments from Lake Taihu, China (Chen et al., 2015) and Dianchi, China (Bai et al., 2012). However, at the genus level, bacterial community composition always varied in different locations of the
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Fig. 5. Cluster (a) and principal component analysis (b) of bacterial communities. S-VS: vegetated surface sediment; M-VS: vegetated middle sediment; B-VS: vegetated bottom sediment; S-US: unvegetated surface sediment; M-US: unvegetated middle sediment; B-US: unvegetated bottom sediment; L-MV: leaves of M. verticillatum; R-MV: roots of M. verticillatum; L-NP: leaves of N. peltatum; S-NP: stems of N. peltatum; R-NP: roots of N. peltatum; L-TJ: leaves of T. japonica; S-TJ: stems of T. japonica; R-TJ: roots of T. japonica.
same lake, different lakes, and even different seasons (Chen et al., 2015). A recent report (Bai et al., 2012) showed that in sediments of Lake Dianchi, the most abundant classified bacterial genera in the high organic carbon sediment were different from those in low organic carbon sediments. These differences can be ascribed to the differences in environment parameters (i.e. water flow, dissolved oxygen, disturbance) and nutrient level. The structures of bacterial communities in the vegetated sediments were different from those in the unvegetated sediments (Fig. 4). Though it was impossible to provide enough evidence to confirm whether these differences were caused by the activity of aquatic plant roots alone, as revealed by PCA (Figue 5b), bacterial community composition was obviously different among the three vertical layers of the sediments from vegetated areas as compared to those from the unvegetated areas. Jensen et al. (2007) also found a significant difference between the bacterial communities associated with the seagrass roots and the surrounding bulk sediment. These differences may be ascribed to the differences in organic material accumulation, oxygen and chemical exudates in the rhizosphere zone in the vegetated area (Faulwetter et al., 2013). Therefore, the roots of these plants may have altered the bacterial community in the surrounding sediments. 4.4. Bacteria community in nitrogen-cycle Microorganisms, including nitrifying bacteria and denitrifying bacteria, play important roles in the nitrogen cycles of various ecosystems through nitrification and denitrification processes (Hayatsu et al., 2008). Though nitrification activity and nitrifying bacteria were reported on the surface of submersed macrophytes (Trias et al., 2012), detailed information on these bacteria remains unclear. In the present study, three genera of nitrifiers Nitrosospira, Nitrospina and Nitrosococcus were detected; Nitrosospira was detected only in biofilm, while Nitrospina and Nitrosococcus were detected only in sediments (Table 1). Nitrifiers prefer high dissolved oxygen concentration areas, but percentages of nitrifiers were lower in the vegetated sediments (0.62–0.72%) than in unvegetated sediments (3.22–3.90%) (Table 1). Herrmann et al. (2008) found a slightly lower relative abundance of ammonia-oxidizing bacteria in the rhizosphere of L. uniflora than in the unvegetated sediments. In the present study, the unvegetated sediment sam-
ples were taken from shallower areas, which is beneficial for the nitrifying bacteria (Flood et al., 2015). However, nitrifying bacteria occurring at 0.01-0.06% in biofilm may be ascribed to the low ammonium concentration in the water column (Table S1). In fact, the abundances of ammonia- and nitrite- oxidizing bacteria usually represent less than 1% of the total bacterial population, for example, in activated sludge of full-scale wastewater treatment plants, but they are extremely important for nitrogen removal. Denitrifying bacteria reduce nitrates to molecular nitrogen or gaseous nitrogen. A total of eighteen genera containing these bacteria, or identified as denitrifiers were summarized (Table 1). Rhodobacter was the primary denitrifier in biofilm from the leaves (11.32%) and roots (7.50%) of M. verticillatum, while Acinetobacter and Pseudomonas dominated in most of the macrophyte samples (accounting for 6.71% on average). Nitrospinaceae was the most dominant denitrifier in all sediment samples, yet the proportions of other denitrifying genera were extremely low. The potential dentrifiers Pseudomonas, Comamonas, Achromobacter, Alcaligenes and Flavobacterium were also detected in the previous studies (Drysdale et al., 1999; Gumaelius et al., 2001; Hayatsu et al., 2008; Lai and Shao, 2008). In the present study, occurrence of denitrifying bacteria in the bacterial community ranged from 1.95% to 3.70% in sediments and from 6.85% to 19.23% in biofilm (Table 1). Eriksson and Weisner (1997) found that the level of denitrification can be relatively higher in epiphytic microbial communities than in those in sediments. Abovementioned results highlight the import role of submersed macrophyte-biofilm system on nitrogen cycles. Plants can release oxygen through photosynthesis system at the daytime, while plants and most organisms in biofilm consume oxygen through respiration in their lifetime. Therefore, aerobic and low oxygen micro-environment can possibly be formed at the interface between plant and water during the day and the night, respectively, and a possible aerobic-anaerobic micro-environment may be formed at the rhizosphere-sediment interface. These micro-environments are beneficial to nitrifying and denitrifying organisms. However, the data are still not adequetly robust explain the role of aquatic plants in modifying the ntrogen-cycle in freshwater the column, because ntrogen-cycle related archaea and fungus may also exist in the biofilms (Hayatsu et al., 2008). For example, epiphytic ammonia-oxidizing bacteria and archae
Bottom
2.69 – 1.21 3.90 0.05 – – 0.11 – 0.83 – 0.06 0.64 – – – 0.01 1.72 – – – 0.09 3.51
Middle
1.95 – 1.27 3.22 0.04 – – 0.04 0.01 1.19 – 0.04 0.70 – 0.06 – 0.01 1.47 – – – 0.14 3.70 2.55 – 1.29 3.84 0.02 – – 0.04 – 0.80 – – 0.42 – 0.07 – – 1.46 – – 0.01 0.20 3.02
Surface
5. Conclusions Total microbe and algae concentrations on leaves of submersed macrophyte were higher than those of two floating macrophytes in this wetland system. The structures of bacterial communities in sediments are markedly different from those in the biofilms attached on the three aquatic macrophytes, and there is an obvious difference in bacterial community between the vegetated and unvegetated sediments. These results suggest that three aquatic macrophytes provided special niche for microbes in water column and sediments. The relative abundance of nitrifying bacteria were higher in sediment than in biofilm, while that of denitrifying bacteria were markedly higher in biofilm than in sediments. However, the role of epiphytic microbial communities in nitrogen removal should be furtherly investigated in the future.
0.54 – 0.15 0.69 0.03 – – – – 0.33 – 0.04 0.03 – – – – 1.55 – – – – 1.98
Bottom
0.46 – 0.26 0.72 – – – – – 0.38 – 0.02 0.05 – 0.01 – – 2.53 – – – 0.01 3.00 – 0.06 – 0.06 0.05 4.34 – 0.12 0.54 0.12 0.04 – 3.37 – 0.06 0.43 0.31 0.05 2.68 0.20 1.66 – 13.97
Middle Surface Root
– 0.01 – 0.01 – 13.19 – 0.01 0.43 – – – 0.47 0.01 – 0.04 0.13 – 0.70 0.39 1.27 – 16.64 – 0.02 – 0.02 – 2.61 – 0.03 0.26 – 0.05 0.61 3.06 0.03 – 0.31 0.04 0.06 2.56 0.20 0.43 – 10.25 – 0.06 – 0.06 – 3.15 – 0.08 0.06 – 0.05 0.07 3.18 0.08 – 0.43 0.12 – 7.10 0.83 0.41 0.01 15.57
Stem Leaf Stem Leaf
Conflict of interest The authors declare no conflict of interest.
– 0.02 – 0.02 – 4.36 – – 0.62 0.10 0.01 – 1.86 0.08 – 0.29 0.23 – 0.90 0.73 1.75 – 10.93
This study was, in part, supported by Grants from the National Natural Science Foundation of China (Grant No. 51379063 and 51579075), Science Fund for Creative Research Group of the National Natural Science Foundation of China (51421006), and Innovation Project from Ministry of Education of China (IRT13061), the Fundamental Research Funds for the Central Universities (2016B06714), Jiangsu water resource technology project (2014076 and 2015085) and a Project Funded by the Priority Academic Program Development of Jiangsu Higher Education Institutions.
– 0.04 – 0.04 – 3.41 – 0.01 0.20 0.01 0.34 – 2.45 0.32 0.01 0.53 1.37 0.01 1.20 1.87 7.50 – 19.23
– 0.03 – 0.03 – 0.89 0.14 0.05 0.20 – 0.04 0.15 0.90 – – 0.33 0.07 0.01 2.14 0.14 1.79 – 6.85
Root
– – – – – 3.70 – – 0.76 0.04 – – 0.26 0.01 – 0.08 0.60 0.02 0.76 0.89 11.32 – 18.44
Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.ecoleng.2016.10. 011. References
Total
Achromobacter Acinetobacter Alcaligenes Azospira Bacillus Comamonas Dechloromonas Desulfovibrio Flavobacterium Flexibacter Halomonas Hydrogenophaga Hyphomicrobium Nitrospinaceae uncultured Pseudomonas Rhizobium Rhodobacter Sphingobacterium Total Denitrifying bacteria
Nitrifying bacteria
Nitrospina (NOB) Nitrospira (NOB) Nitrosococcus (AOB NOB)
Leaf
Root
Acknowledgements
’–’ Indicates genus was not detected.
Vegetated sediment T. japonica N. peltatum M. verticillatum Genus
Table 1 The abundance of bacteria associated with nitrogen cycling in each sample (percentages).
249
were detected on marine macroalgae (Trias et al., 2012), and the abundance of ammonia-oxidizing archaea was higher than ammonia-oxidizing bacteria in the rhizosphere of aquatic plants (Chen et al., 2008; Herrmann et al., 2008). In addition, the function of abundant bacteria remains unclear, especially for those unclassified bacteria (Supplementary Table S6). Therefore, further study is demanded to explore this subject in future.
0.46 – 0.16 0.62 – – – 0.01 – 0.25 – – 0.04 – 0.01 – – 1.63 – – – 0.01 1.95
Unvegetated sediment
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