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ARTICLE Impacts of different salinities on bacterial biofilm communities in fresh water Lei Zhang, Guang Gao, Xiangming Tang, and Keqiang Shao

Abstract: Natural and anthropogenic salinization continuously impacts inland aquatic ecosystems. Associated bacterial biofilms respond rapidly to environmental conditions and are potential bioindicators for changes in water quality. This study evaluates the effects of different salinity concentrations (0.3‰–10‰) on bacterial biofilms communities grown in fresh water from Lake Bosten. Bacterial communities associated with biofilms were analyzed using terminal restriction fragment length polymorphism and clone library analyses of 16S rRNA genes. Results indicated that the attached bacterial community composition (ABCC) changed over several weeks of biofilm growth, but all followed similar bacterial successional trends in the different salinity groups. Detailed analysis showed the following. (i) ABCC did not differ (P > 0.05) in the low-salinity groups (0.3‰–3.5‰), which may be related to the lower osmotic pressure and the shorter time scale (weeks) of their present habitats. (ii) There were significant differences between the oligosaline (3.5‰) and saline (10‰) groups (P < 0.05). In particular, genus Flavobacterium became dominant in attached bacterial communities in the saline groups. The higher abundance of genus Flavobacterium was possibly due to the biological and metabolic characteristics of the bacteria. (iii) Some bacterial taxa can maintain the higher abundance within attached bacteria in the entire process of biofilms growth, such as the genera Hydrogenophaga and Methyloversatilis in Betaproteobacteria and the family Sphingomonadaceae in Alphaproteobacteria. These data suggested that the bacterial successional trends within biofilms seem almost unaffected by salinity (0.3‰–10‰), but ABCC in saline groups (10‰) are notably changed. Key words: bacterial community composition, biofilms, salinity. Résumé : La salinisation naturelle et anthropogène affecte sans relâche les écosystèmes des eaux intérieures. Les biofilms bactériens qui y sont associés répondent rapidement aux conditions environnementales et seraient des indicateurs biologiques aptes a` révéler des changements de la qualité de l’eau. La présente étude évalue l’incidence de diverses salinités (0,3–10 ‰) sur les communautés bactériennes des biofilms cultivés dans de l’eau douce du lac Bosten. On a analysé les communautés bactériennes associées aux biofilms par polymorphisme de la longueur des fragments de restriction terminaux et par l’analyse de banques clonales de gènes d’ARNr 16S. Les résultats ont indiqué que la composition de la communauté bactérienne attachée (CCBA) changeait nettement tout au long de la croissance des biofilms s’étalant sur plusieurs semaines, mais toutes ont suivi des tendances de succession qui se regroupaient selon les divers groupes de salinité. Une analyse détaillée a montré que (i) les CCBA n’étaient pas différentes (P < 0,05) dans les groupes a` faible salinité (0,3–3,5 ‰), ce qui s’expliquerait par la pression osmotique plus basse et la brève longévité (s’exprimant en semaines) de leurs habitats présents; (ii) il y a eu des différences significatives entre les groupes oligosalins (3,5 ‰) et salins (10 ‰) (P < 0,05). Tout particulièrement, le genre Flavobacterium est devenu dominant dans les communautés bactériennes attachées des groupes salins. L’abondance supérieure du genre Flavobacterium pourrait tenir des caractéristiques biologiques et métaboliques des bactéries; et (iii) certains taxons bactériens peuvent demeurer hautement abondants au sein des bactéries attachées tout au long du processus de développement des biofilms, dont les genres Hydrogenophaga et Methyloversatilis chez les Betaproteobacteria et la famille des Sphingomonadaceae chez les Alphaproteobacteria. Ces données indiquent que les tendances de succession des bactéries que l’on observe au sein des biofilms semblent n’être presque pas affectées par la salinité (0,3–10 ‰), tandis que les CCBA des groupes salins (10 ‰) sont nettement altérés. [Traduit par la Rédaction] Mots-clés : composition de la communauté bactérienne, biofilms, salinité.

Introduction Global warming and intense growth in human activities cause water salinization to continuously impact natural inland aquatic ecosystems (Walpole and Simmons 1996), especially in arid and semiarid regions. For example, Lake Bosten, formerly the largest inland freshwater lake in the arid regions of northwest China, has changed from a freshwater to an oligosaline lake over the past 50 years (Xie et al. 2011). Catchment changes and anthropogenic disturbances to hydrological cycles increase salt loads to water

bodies, and bacteria are thought to be a sensitive sentinel to environmental changes (Newton et al. 2011). Most bacteria in fresh water are found growing as biofilms on surfaces of submerged substrata or sediments (Costerton et al. 1987; Blenkinsopp and Costerton 1991; Bartrons et al. 2012). Attached bacteria are a crucial component in periphyton and play an important role in the composition of aquatic microbial food webs and the formation and decomposition of organic matter. Previous studies on attached bacteria mainly addressed

Received 11 November 2013. Revision received 2 April 2014. Accepted 3 April 2014. L. Zhang. State Key Laboratory of Lake Science and Environment, Nanjing Institute of Geography and Limnology, Chinese Academy of Sciences, 73 East Beijing Road, Nanjing, 210008, People’s Republic of China; University of Chinese Academy of Sciences, Beijing 100049, People’s Republic of China. G. Gao, X. Tang, and K. Shao. State Key Laboratory of Lake Science and Environment, Nanjing Institute of Geography and Limnology, Chinese Academy of Sciences, 73 East Beijing Road, Nanjing, 210008, People’s Republic of China. Corresponding author: Guang Gao (e-mail: [email protected]). Can. J. Microbiol. 60: 319–326 (2014) dx.doi.org/10.1139/cjm-2013-0808

Published at www.nrcresearchpress.com/cjm on 4 April 2014.

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bacterial functional or metabolic characteristics as biofilms development, such as bacterial biomass, production, cell density, and extracellular enzyme activity (Lowe et al. 1996; Sabater and Romaní 1996; Sobczak 1996). With the development and application of modern molecular biology techniques to microbial ecology (Giovannoni et al. 1990; Muyzer et al. 1993; Liu et al. 1997), research focus is shifting to attached bacterial community composition (ABCC), the differences of community structure, or structure– function relationships (Gehrke et al. 2001; Lehman et al. 2001; Worm et al. 2001; Lyautey et al. 2005; Despland et al. 2012). The bacterial communities within biofilms respond rapidly to changing environmental conditions; therefore, bacterial community composition of artificially grown and field-grown biofilms has previously been used as a bioindicator for water quality in freshwater and estuarine water bodies (Nocker et al. 2007; Campbell et al. 2011). Salinity is considered the major environmental determinant of microbial community composition on a global scale, according to Lozupone and Knight (2007). It can directly affect abundance, growth, and activity of bacterial communities and presents a physiological barrier for certain bacterial groups (Ben-Dov et al. 2008; Caporaso et al. 2011). Several studies have indicated that the bacterioplankton community composition changes along a salinity gradient (Wu et al. 2006; Pommier et al. 2007; Silveira et al. 2011). In contrast, little is known about the impacts of varying salinity on the composition of biofilms bacterial communities. What is not clear is whether attached bacteria follow similar successional trends with varying salinity, or how the ABCC in biofilms responds to changing salinity conditions. In our study, the same freshwater community subjected to different salinities without needing to deal with competition by salt-adapted species might be considered as a case for freshwater bacteria ending up in the sea.

Materials and methods Experimental design We conducted a mesocosm experiment for 30 consecutive days (1–30 August 2012) in a laboratory at the Institute of Lake Bosten using levels of salinity that encompassed a range from fresh water to saline water. Each of 12 glass containers (60 cm × 50 cm × 30 cm, open only at the top) was filled with 80 L of water and artificial substrates for biofilms growth (Bio-cords). Water used for the experiment was obtained from the macrophyte-covered area of Lake Bosten in Xinjiang, China. Four experimental treatments were used, and each group had 3 replicates. In the control groups (salinity: 0.3‰), the containers were not supplemented with sterile crystallized salt. In the salt-addition groups (brackish, oligosaline, and saline groups), containers were supplemented sterile crystallized salt to salinity concentrations of 1.5‰, 3.5‰, and 10‰, respectively. We simulated the attached bacterial succession in different levels of salinity from the early Lake Bosten (fresh water, 0.3‰), to the present Lake Bosten (brackish, 1.5‰), to the future Lake Bosten (e.g., another oligosaline lake in this region: Lake Ulungur, 3.5‰), to a saline lake (10‰). Sampling, measurement of biotic and abiotic variables The Bio-cords were collected on days 5, 10, 15, 20, and 30. Within 2 h after sampling, the biofilms material of Bio-cords was carefully scraped into 300 mL of sterile water using a sterile brush. Additionally, 200 mL of the water for 16S rRNA gene analysis was collected on 0.2 ␮m pore-size polycarbonate filters (Millipore) in the field by using a hand-driven vacuum pump. The filters were stored at –80 °C prior to nucleic acid extraction. Another subsample (46 mL) was transferred into autoclaved PP-tubes containing 4 mL of prefiltered (pore size 0.2 ␮m) glutaraldehyde (final concentration 2% (v/v)) and then refrigerated at 4 °C for bacterial counts. On each sampling day, we sampled 500 mL of water from the containers at 1000 hours every morning for immediate chem-

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ical analysis according to standard methods (Clesceri et al. 1992), including total nitrogen, total phosphorus, and chemical oxygen demand. The concentrations of dissolved oxygen, chlorophyll a, total dissolved solids, and salinity, as well as pH and water temperature in the experimental glass containers were determined using a multiparameter water quality sonde (YSI 6600V2, USA). The bacterial abundance in water samples was determined by epifluorescence direct counting method after DAPI (4=,6diamidino-2-phenylindole) staining (Porter and Feig 1980). DNA extraction, polymerase chain reaction (PCR) amplification, and terminal restriction fragment length polymorphism (T-RFLP) analysis Total nucleic acids for filtered microorganisms was extracted and purified using proteinase K, sodium dodecyl sulfate, and cetyltrimethylammonium bromide concomitant with phenol– chloroform extraction and isopropanol precipitation according to the protocol of Zhou et al. (1996). Bacterial 16S rRNA genes were amplified using the Cy5-labeled forward primer 8F (5=- AGAGTTTGATCCTGGCTCAG-3=) and the unlabeled reverse primer 926R (5=CCGTCAATTCCTTTGAGTTT-3=) (Liu et al. 1997). The purified DNA from mixed triplicate extraction was used as PCR template. PCR amplification was carried out in a thermocycler (Applied Biosystems Veriti Thermal Cycler) using a touchdown program: denaturation at 95 °C for 5 min, 20 cycles of denaturation at 95 °C for 30 s, annealing at 65 °C for 30 s (the temperature was decreased by 0.5 °C every cycle until the temperature of 55 °C was reached), and extension at 72 °C for 1.5 min. An additional 15 cycles were carried out at an annealing temperature of 55 °C, followed by a final extension at 72 °C for 10 min. The triplicate PCR products were purified and concentrated using the E.Z.N.A. Cycle-Pure kit. Later these products were digested with QuickCut Hha I (Takara) at 37 °C for 10 min, then purified, separated, and analyzed with a Beckman Coulter Ceq. 8000 Genetic Analysis System (Beckman Coulter, Fullerton, California, USA). After electrophoresis, T-RFLP data were preprocessed using the online tool T-REX (Culman et al. 2009). Analysis of similarity (ANOSIM) was performed to statistically test the effects of habitats on ABCC using the PRIMER 6 package (PRIMER-E, Ltd., UK) based on the distance indices of T-RF peak area data. Cloning, sequencing, phylogenetic analysis, and assignment of T-RFs Eight clone libraries were generated with mixed bacterial 16S DNA templates in triplicate extractions retrieved at days 5 and 30 from the control groups and the salt-addition groups. The general bacterial primers 8F (5=-AGAGTTTGATCCTGGCTCAG-3=) and 1492R (5=-GGTTACCTTGTTACGACTT-3=) (Weisburg et al. 1991) were used in PCR amplifications. The PCR protocols were the same as described above. The PCR products were purified immediately with the E.Z.N.A. Cycle-Pure kit, and the PCR products were ligated into the pGEM-T Easy Vector (Promega) according to the manufacturer’s instructions. The presence of the 16S rRNA gene in positive colonies was checked by PCR amplification using vector primers (M13F and M13R) and randomly selected for sequencing, which was carried out by Invitrogen (Shanghai, China). Raw sequences were edited manually using the software BioEdit (Hall 1999). The high-quality sequences (without chimeric artifacts) were clustered in operational taxonomic units (OTUs) with a 0.03 cut-off value (equivalent to 97% similarity) using the software Mothur (Schloss et al. 2009). The coverage index, Chao1 richness estimator, Shannon diversity index (H=), and reciprocal Simpson index were also calculated by the software Mothur. Principal coordinate analysis based on weighted UniFrac distances was performed using the function provided by the UniFrac web interface (Lozupone et al. 2006). The sequence of each OTU was chosen as a representative and was blasted against the NCBI (http://www.ncbi.nlm.nih.gov/BLAST/) and the Ribosomal Published by NRC Research Press

Zhang et al.

Database Project II (RDP II; http://rdp.cme.msu.edu/) to assign a phylogenetic affiliation to the 16S rRNA gene sequences. To test the phylogenetic assignments based on in silico T-RF analysis, randomly selected clones were analyzed by in vitro T-RF by finding the first HhaI enzymatic digestion site downstream from 8f. The differences between 8 constructed clone libraries were statistically assessed using the 兰-LIBSHUFF command in the Mothur software. The Cramér – von Mises statistic was calculated to determine the probability that the observed difference among the libraries was due to chance. With an experiment-wise error rate of 0.05, and taking into account a Bonferroni correction due to the multiple comparisons, the libraries were considered significantly different if the P value was less than 0.0017 (Singleton et al. 2001; Schloss et al. 2004).

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Fig. 1. Changes of the number of terminal restriction fragments (T-RFs), Shannon’s diversity index (H=), and attached bacterial abundance in different saline habitats over time.

Nucleotide sequence accession number The partial 16S rRNA gene sequences determined in this study have been submitted to the GenBank database under the accession Nos. KF826960–KF827424.

Results Changes in bacterial abundance and diversity over time With increasing succession times, changes in the bacterial abundance, H= values, and the number of T-RFs were all apparent in different treatment groups (Table S11), but all showed similar trends, such as an initial increase at the first few days of submersion, followed by a slight decrease and a subsequent increase at the later stage (Fig. 1). Diversity estimators and comparison of selected clone libraries To identify the succession of bacterial communities in different treatment groups, 8 clone libraries were retrieved from the control, brackish, oligosaline, and saline groups at days 5 and 30. This process generated 465 bacterial 16S rRNA gene sequences >1400 bp long from the 8 water samples (Table 1). These sequences were classified into 253 unique OTUs at a similarity level of ≥97% by Mothur. Estimated richness (Chao1) varied from 27.9 to 111.1, with a median of 66.8 (Table 1). The reciprocal Simpson index was much lower in the 4 day 5 libraries (C-5, B-5, O-5, and S-5) than in the day 30 libraries (C-30, B-30, O-30, and S-30), revealing a more uneven distribution of the phylotypes (OTUs) for the early stage of bacterial succession. The lower H= values representing species richness and abundance of the former 4 libraries confirmed the Simpson index analysis (Table 1). A comparison with LIBSHUFF statistics also indicated that ABCC differed significantly between the early and later stages of succession process. We obtained P < 0.0017 for the comparisons of each day 5 to each day 30 library and also for the comparison among all early or later sequences. Nevertheless, the later libraries differed, whereas the majority of early libraries were statistically similar (P > 0.0017) (Table S21). A scatterplot of the first 2 principal coordinates (PC) by the UniFrac analysis (Fig. 2) indicated that PC1 and PC2 explained 24.6% and 15.5% of the data variation, respectively. The early (day 5) libraries were separated from the later (day 30) libraries in the PC1 plot. This result corroborates the LIBSHUFF analysis. Phylogenetic analysis To further explore bacterial phyla composition in these different communities, we classified sequences from each library with BLAST and the RPD classifier tools. The resulting 465 clone sequences were distributed among 12 different major bacterial groups. The majority of clones retrieved from the 8 samples at different succession stages were affiliated with Alphaproteobacteria, Betaproteobacteria, Gammapro-

1

teobacteria, Bacteroidetes, Planctomycetes, while a minor portion belonged to Actinobacteria, Firmicutes, Acidobacteria, Verrucomicrobia, and Cyanobacteria (Fig. 3). Clones belonging to Proteobacteria, Bacteroidetes, and Planctomycetes were dominant in all stages of the successional process of different treatment groups. Over time, however, the proportion of each phylum was significantly different among the early and later succession stages. At the early succession stage (day 5), clones belonging to Betaproteobacteria (70.5% ± 9.8%) were dominant in all treatments. Over time, clones belonging to Alphaproteobacteria (25.7% ± 7.0%), Gammaproteobacteria (11.6% ± 3.6%), Bacteroidetes (11.9% ± 10.5%), and Planctomycetes (15.8% ± 4.2%) increased significantly at the later succession stage (day 30), whereas clones belonging to Betaproteobacteria markedly decreased (23.1% ± 8.9%) (Fig. 3). Succession of bacterial community revealed by T-RFLP The terminal fragment sizes were compared in silico with lengths of restricted 16S rRNA genes of the clones obtained in our study. The dominant T-RFs (≥5% of total area) were obtained, and the matching bacterial species detected in each clone library were recorded (Table S31). Distinct patterns of T-RFLP fingerprints were observed in different treatments over time (Fig. 4 and Fig. S11). In the control groups (freshwater salinity) at day 5, the T-RFs at 62 bp (genus Pasteuria), 67 bp (genus Methyloversatilis), 82 bp (family Sphingomonadaceae), 147 bp (genus Hydrogenophaga), 206 bp (family

Supplementary data are available with the article through the journal Web site at http://nrcresearchpress.com/doi/suppl/10.1139/cjm-2013-0808. Published by NRC Research Press

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Table 1. Richness and diversity measures for the clone libraries of control and salt addition groups by the Chao1 richness estimator, Shannon's diversity index (H'), and Reciprocal Simpson index (RSI). Library

Clones

OTUs

Chao1

H'

RSI

C* (%)

Control T=5 days (C-5) Control T=30 days (C-30) Brackish T=5 days (B-5) Brackish T=30 days (B-30) Oligosaline T=5 days (O-5) Oligosaline T=30 days (O-30) Saline T=5 days (S-5) Saline T=30 days (S-30)

58 58 56 60 59 57 60 57

24 43 29 45 22 35 20 35

44.0 105.0 71.8 111.1 33.3 77.2 27.9 63.9

2.59 3.65 3.04 3.68 2.81 3.38 2.36 3.40

9.2 77.8 18.6 68.1 16.9 40.3 7.1 43.1

72.9 45.8 66.1 41.7 83.3 60.3 81.7 61.4

Note: All statistics are for operational taxonomic units (OTUs) defined at the species level (0.03% distance). *C, Coverage index.

Fig. 2. Results of principal coordinate analysis with a UniFrac distance matrix comparing the 8 samples. The scatterplot is of principal coordinate1 (PC1) vs. principal coordinate 2 (PC2). C-5, B-5, O-5, and S-5 — day 5 libraries; C-30, B-30, O-30, and S-30 — day 30 libraries.

Table 2. Analysis of similarity (ANOSIM) comparisons of attached bacterial communities in various salinity habitats. Comparison

Sample statistic R

P

Different treatments Control vs. brackish groups Control vs. oligosaline groups Control vs. saline groups Brackish vs. oligosaline groups Brackish vs. saline groups Oligosaline vs. saline groups

0.191* −0.076 0.052 0.228 −0.020 0.332 0.536*

0.012 0.802 0.246 0.063 0.460 0.056 0.024

*P < 0.05.

abundance of >10% (Fig. 4). Furthermore, T-RFs of 91 bp (genus Flavobacterium) became the most abundant (with relative abundances of 29.0%, 22.6%, and 23.8%). At day 30, the structure of the bacterial community had changed again. The 82 bp (family Sphingomonadaceae) and 567 bp (genus Limnobacter) T-RFs dominated the bacterial populations with relative abundances of >20% (Fig. 4). In the present study, there were marked temporal variations in the percentage of dominant T-RFs in each treatment. Analysis of similarity also showed that differences in bacterial community structure among the 4 treatments were significant (R = 0.191, P < 0.05) (Table 2). Furthermore, the pair-wise comparisons in bacterial community structure showed insignificant differences among the control, brackish, and oligosaline groups (P > 0.05), whereas significant differences were found between oligosaline and saline groups (R = 0.536, P < 0.05) (Table 2).

Discussion Burkholderiaceae), and 567 bp (genus Limnobacter) with a relative abundance of >5% dominated the bacterial populations (Fig. 4 and Table S31). The T-RF of 82 bp was a clear dominant population (relative abundance, 37.1%). At days 10, 15, and 20, the relative abundance of most of these dominant T-RFs had changed little and remained the most abundant within the bacterial T-RFLP profile. At day 30, new T-RFs (142, 205, 227, 340, 357, 365, and 398 bp) with a relative abundance of >1% were detected (Fig. S11). In the brackish and oligosaline groups at days 5, 10, and 15, most of the T-RFs identified were simultaneously recorded in the control groups (Fig. 4). At days 20 and 30, more accompanying species with a relative abundance of >1% were detected (Fig. S11). In the saline groups at day 5, distinct bacterial T-RFs were identified. These T-RFs all showed a relative abundance of >1%, and those of 82, 91, 147, and 567 bp exhibited a relative abundance of >10% (Fig. S11). Among them, the T-RF of 91 bp that was first detected was assigned to genus Flavobacterium in Bacteroidetes. At days 10, 15, and 20, the structure of the bacterial community showed similar patterns. T-RFs of 62, 67, 82, 91, and 147 bp were dominant in the bacterial populations (Fig. S11). These T-RFs all showed a relative abundance of 5%, and those of 62 bp (genus Pasteuria) and 147 bp (genus Hydrogenophaga) exhibited a relative

Different solid media, such as rocks (Jackson et al. 2001; Tank and Dodds 2003; Anderson-Glenna et al. 2008), suspended particles (Tang et al. 2010), aquatic plant (Crump and Koch 2008), and artificial substrate (Jones et al. 2007; Witt et al. 2011), in aquatic ecosystems facilitate the attachment of many species of bacteria with various metabolic patterns to gradually form biofilms, which facilitate bacterial species growth as well as other biotic communities. In our study, attached bacterial succession in habitats of different salinity readily occurred during the development of biofilms. Salinity is a determining factor in the distribution of bacterial communities in aquatic ecosystems (Logares et al. 2009). Generally, based on ecological theory, species diversity should decline with increasing salinity, such as for the floral and faunal species in inland lakes (Hammer and Heseltine 1988; Newton et al. 2011). When applied to bacterial communities, however, this theory seems less applicable (Herlemann et al. 2011; Wang et al. 2011; Zhang et al. 2013). Our results showed that attached bacterial abundance, H= values, and the number of T-RFs of different salinity treatments did not decrease gradually with increasing salinity but had an initial increase at the early stage, followed by a slight decrease and a subsequent increase at the later stage (Fig. 1). The effect of different salinity concentrations on bacterial community structure in freshwater biofilms was determined using T-RFLP and cloning and sequencing of 16S rRNA genes. Published by NRC Research Press

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Fig. 3. Bacterial biofilm community composition of the 8 samples along a salinity gradient (from fresh water to saline water) in the control and salt-addition groups.

Fig. 4. Relative abundances of major attached bacterial genera or families (≥5% abundance in all samples) recovered from samples with a salinity gradient over time.

We observed obvious temporal variation of ABCC in habitats of different salinities. Within hours, an organic film forms on the surface, which can facilitate the arrival and adhesion of bacteria from the water column (Costerton et al. 1987). Subsequently, rapid colonization of surfaces by bacterial populations may have occurred within the few days of submersion (Jackson et al. 2001; Martiny et al. 2003).

During the development of biofilms, resource competition drives the development of the attached bacterial community. Additional biofilms formation provides a more favorable environment for secondary colonization of bacteria (Jefferson 2004). In this process, better resource or space competitors may exclude less competitive bacteria, which might manifest as a decrease Published by NRC Research Press

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in richness (Fig. 1). Despite a decrease in species diversity, the relative abundance of most of these dominant bacterial taxa had changed little, so they were still among the most abundant taxa within attached bacteria; for example, Betaproteobacteria was always a dominant taxa in the attachment process (Fig. 3). Betaproteobacteria have been reported as dominant in freshwater systems, such as oligotrophic lakes (Alfreider et al. 1996), humic lakes (Buck et al. 2009), and drinking water biofilms (Kalmbach et al. 1997). Their dominance in attached bacteria in this study might be related to the Proteobacteria in Lake Bosten belonging mainly to Betaproteobacteria (Tang et al. 2012). Further analysis at the genus level found many clone sequences in Betaproteobacteria related to Hydrogenophaga (147 bp) and Methyloversatilis (62 bp). The genus Hydrogenophaga occur in different habitats degrading a variety of organic compounds (Brenner et al. 2005). Members of the genus Hydrogenophaga are often described as being “knallgas” bacteria, facultative autotrophs capable of fixing CO2 while utilizing molecular hydrogen as the electron donor and oxygen as the terminal electron acceptor for growth (Aragno and Schlegel 1999; Kampfer et al. 2005; Gan et al. 2011). The genus Methyloversatilis affiliated with the order Rhodocyclales has only recently been discovered and has been shown to be important for denitrification (Kalyuzhnaya et al. 2006; Baytshtok et al. 2008; Kittichotirat et al. 2011). The ability to utilize nitrogen might confer ecological advantages upon Methyloversatilis for growth in biofilms. Alphaproteobacteria were also an important class in attached bacteria in all treatment groups (Fig. 3). Most Alphaproteobacteria sequences were closely related to the family Sphingomonadaceae (82 bp) (Fig. 4 and Table S31). Members of this family are strictly aerobic chemoheterotrophs and widespread in nature, occurring in soils, water, plant surfaces, and biofilm of pipes (White et al. 1996; Cavicchioli et al. 1999; Koskinen et al. 2000). More important, they can adapt to man-made environments. Most notably, these bacteria can degrade xenobiotic compounds (Stolz 2009) and can survive in chlorinated waters, possibly because of their oligotrophic character and production of biofilms (Furuhata et al. 2007; Hong et al. 2010; Yim et al. 2010), which is further proof of their remarkable capacity to adapt to new habitats; hence, the dominance of Sphingomonadaceae in all treatment groups was not unexpected. During the later stage of bacterial succession, an additional increase in bacterial species richness as a more mature biofilm with complex spatial structure (the creation of new spatial and ecological niches) may facilitate greater diversity through increased variation in habitat and available resources. In our study, the relative abundances of members of the classes Alphaproteobacteria and Gammaproteobacteria and phyla Bacteroidetes and Planctomycetes were significantly increased at the later stage of bacterial succession (Fig. 3). In particular, members of Planctomycetes formed a dominant fraction in attached bacteria, accounting for an initial occurrence of 1.7% ± 0.1% (day 5) to a later increase of 15.8% ± 4.2% (day 30). Previous studies have demonstrated that this phylum is abundant in river biofilms (Brümmer et al. 2000), in river snow (Böckelmann et al. 2000), and in particle-associated communities (Allgaier and Grossart 2006). Most Planctomycetes members can form small flagellated swarmer cells, which swim before attaching and reproducing, when they begin aerobic decomposition of the macromolecule substrate (Fuerst 1995). In addition, members of Planctomycetes play a key role in carbon and nitrogen cycles under anaerobic conditions (Damste et al. 2005; Woebken et al. 2007); therefore, the higher abundance of Planctomycetes members in later successional stages might be related to its biological and metabolic character. Our results may be incomplete because our experimental systems did not include sediments, which are known to harbor dormant bacteria as well as spores. These inactive microorganisms may become active or dominant if the proper conditions are

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given. In addition, as for many molecular techniques (e.g., DGGE, ARDRA), actual bacterial diversity may be masked and underestimated because of the limited throughput of T-RFLP (the dominant members of a community) and the clone library (Claesson et al. 2010). Hence, bacterial taxa initially detected may still be present in later biofilms but accompanied by rapid growth of dominant taxa. Our results contrast with the limited studies of bacterial communities within a bioreactor or on leaf surfaces where no discernible pattern was indicated in the changes in bacterial community richness during biofilms succession (van der Gast et al. 2008; Redford and Fierer 2009), possibly because of a discontinuous successional sequence, frequently influenced by environmental conditions (Redford and Fierer 2009). Our results (i) are more consistent with the findings of Jackson et al. (2001), Martiny et al. (2003), and Lyautey et al. (2005), who observed predictable changes in bacterial diversity, with the higher levels of bacterial diversity in the early and late stages of biofilms development, and (ii) better fit the model of biofilms succession proposed by Jackson et al. (2001). Although similar successional trends of bacterial biofilms communities were observed in all treatment groups (Fig. 1), the ANOSIM showed that ABCC among low-salinity habitats (0.3‰– 3.5‰) did not differ (P > 0.05); however, significant differences were found between the saline and oligosaline groups (P < 0.05). In the low-salinity habitats, attached bacteria may require intergenus adaptations to cope successfully with the fluctuation of low osmotic pressure. Another explanation may be shifts in gene expression and regulation rather than ABCC on the shorter time scale (weeks) (Edmonds et al. 2009). In the saline groups, bacterial communities are currently subjected to an unexpected test in waters enriched with salts. While successional trends are similar to those in low-salinity habitats, acute bacterial response to high salinity resulted in significant shifts in ABCC. For instance, genus Flavobacterium was dominant in attached bacterial communities in the saline groups (Fig. 4). The genus Flavobacterium is the type genus of the family Flavobacteriaceae and has the following main characteristics: Gram-negative rods that are motile by gliding, are chemoorganotrophs and aerobes, and decompose several polysaccharides but not cellulose (Bernardet et al. 1996; Bernardet and Nakagawa 2006). Members of the genus Flavobacterium are widely distributed in soil, freshwater, and saline habitats and tolerate a wide range in salinity (Bernardet and Bowman 2006). As expected, genus Flavobacterium dominated attached bacteria in the saline groups in our study. In conclusion, this study suggests that the impacts of different salinities (0.3‰–10‰) on freshwater ecosystems do not change the successional trends of bacterial biofilms communities; therefore, these trends also provide further evidence for the model of biofilms succession proposed by Jackson et al. (2001). Owing to the limited time scale (weeks) and osmotic pressure, no significant differences among ABCC were found in low-salinity habitats (0.3‰–3.5‰) during the development of biofilms. However, as salinity further increased (10‰), ABCC obviously changed (e.g., genus Flavobacterium). Our study represents a relatively detailed analysis of ABCC response in fresh water to increasing salt concentrations, which may serve as an indicator for future water salinity changes in arid regions.

Acknowledgements We thank staff at the Environmental Monitoring Station of the Environmental Protection Bureau of Bayingolin Mongolia Autonomous Prefecture for helping with sample collection and water chemical analysis. This work was supported by the National Natural Science Foundation of China (grants 41171388 and 31270505), the National Water Pollution Control and Management of Science and Technology Major Projects (grant 2013ZX07104-004) and the Special Environmental Research Funds for Public Welfare of the State Environmental Protection Administration (grant 201309041). Published by NRC Research Press

Zhang et al.

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