Microb Ecol DOI 10.1007/s00248-013-0348-3
ENVIRONMENTAL MICROBIOLOGY
Molecular Techniques Revealed Highly Diverse Microbial Communities in Natural Marine Biofilms on Polystyrene Dishes for Invertebrate Larval Settlement On On Lee & Hong Chun Chung & Jiangke Yang & Yong Wang & Swagatika Dash & Hao Wang & Pei-Yuan Qian
Received: 24 May 2013 / Accepted: 6 December 2013 # Springer Science+Business Media New York 2014
Abstract Biofilm microbial communities play an important role in the larval settlement response of marine invertebrates. However, the underlying mechanism has yet to be resolved, mainly because of the uncertainties in characterizing members in the communities using traditional 16S rRNA gene-based molecular methods and in identifying the chemical signals involved. In this study, pyrosequencing was used to characterize the bacterial communities in intertidal and subtidal marine biofilms developed during two seasons. We revealed highly diverse biofilm bacterial communities that varied with season and tidal level. Over 3,000 operational taxonomic units with estimates of up to 8,000 species were recovered in a biofilm sample, which is by far the highest number recorded in subtropical marine biofilms. Nineteen phyla were found, of which Cyanobacteria and Proteobacteria were the most dominant one in the intertidal and subtidal biofilms, respectively. Apart from these, Actinobacteria, Bacteroidetes, and Planctomycetes were the major groups recovered in both intertidal and subtidal biofilms, although their relative abundance varied among samples. Full-length 16S rRNA gene clone libraries were constructed for the four biofilm samples and showed similar bacterial compositions at the phylum level to those revealed by pyrosequencing. Laboratory assays confirmed that cyrids of the barnacle Balanus amphitrite preferred Lee and Chung made equal contributions to the work Electronic supplementary material The online version of this article (doi:10.1007/s00248-013-0348-3) contains supplementary material, which is available to authorized users. O. O. Lee : H. C. Chung : J. Yang : Y. Wang : S. Dash : H. Wang : P.10 % for major families)
using the clone library technique. In addition, the rank abundance curves indicated a high proportion of rare organisms in all biofilm samples (Fig. S3b). Based on pyrosequencing data, a higher microbial diversity (with more than 3,000 OTUs and estimates of up to 8,000 species phylotypes) was found in the biofilms developed in the spring than in those developed in the summer, whereas tidal level seemed to have less influence on microbial diversity in biofilms (Table 1). The Chao1 and ACE estimators were approximately two-fold higher than the number of OTUs in all samples (Tables 1 and S2), indicating that the actual number of species may be doubled. The numbers of OTUs recovered in the spring intertidal biofilms were comparable with those in deep-seawater and sponges revealed by the same technique [60]. Using the clone library technique, the bacterial diversities in our subtropical marine biofilms were substantially higher than those of biofilms from different habitats [32] but were comparable with the planktonic communities in many marine environments [53]. In general, bacterial communities in the subtidal biofilms developed in this study were slightly more diverse than those of the intertidal biofilms, whereas biofilms developed in the spring were substantially more diverse than those developed in the summer (see Shannon index, Table 1), indicating that season may have a greater influence on microbial diversity than tidal height. In contrast, a clear preference of the barnacle cyprids to settle on intertidal biofilms, regardless of the season, was observed, although the microbial diversity and
Fig. 4 16S rRNA-based phylogenetic trees showing the genetic distances among clones retrieved from the libraries for biofilms developed at different tidal levels in different seasons in reference to members of the phyla a α-Proteobacteria and b Cyanobacteria. The trees were constructed based on the neighbor-joining method. Clones from different biofilms are prefixed with the sample ID (see Table 1 for details) followed by clone number. Reference sequences from the closest relatives are retrieved from NCBI and those originated from seawater, sediment, activated sludge, corals, sponges, sea urchin, ocean crust, and biofilms are indicated by the superscripts “SW,” “SD,” “AS,” “C,” “S,” “U,” “OC,” and “BF,” respectively. Nucleotide accession numbers of the reference sequences are given in parentheses. Solid boxes show “tidalspecific” clusters, whereas dotted boxes show “season-specific” clusters. The scale bar represents percent substitutions per nucleotide position. Bootstrap values of >50 % based on 1,000 re-samplings are indicated by the numbers at the nodes
Microbial Communities in Natural Biofilms
density of the biofilms were similar, which suggests that the preference may not be related to microbial diversity or density, and that other parameters in biofilms, such as community composition, may have a greater influence.
Microbial Composition in Biofilms To determine the spatial and seasonal variations in microbial community composition of biofilms, we classified the
O.O. Lee et al.
Fig. 4 continued.
pyrosequencing tags into different phyla at a confidence threshold of 50 % using the RDP classifier. About 3–6 % of the reads could not be assigned to a known phylum, and the remaining reads were classified into 19 different phyla; Bacteroidetes, Cyanobacteria and Proteobacteria accounted for >75 % of the reads, and were the major phyla detected in all of the biofilms (Fig. 2; Table S3). Classification of the reads into lower taxonomic levels revealed extremely diverse bacterial communities in all biofilm samples, with up to 70 orders (16 of which constituted >1 % of the reads) and 500 genera (28 of which constituted >1 % of the reads) (Table S4). Although all the biofilm samples were dominated by Cyanobacteria, Proteobacteria, and Bacteroidetes, regardless of the method used, notable differences but similar trends in the proportions of these phyla (i.e., composition) were observed between the intertidal and subtidal biofilms (Figs. 2 and S4). For instance, in both seasons, Cyanobacteria accounted for more reads/clones in the intertidal biofilms than in the subtidal biofilms; conversely, more reads/clones were assigned to Proteobacteria and Planctomycetes in the subtidal
biofilms than in the intertidal biofilms. Seasonal variations in the bacterial communities were also revealed by pyrosequencing (Fig. 2; Table S3). More pyrosequencing reads were assigned to Actinobacteria, Chloroflexi, and Firmicutes in the biofilms developed in the summer than in those developed in the spring. Unifrac PCA analysis detected substantial differences among bacterial communities in the biofilms developed at different tidal levels in the summer and in the biofilms developed in different seasons for the same tidal level (Fig. S5). We constructed a heatmap to illustrate the compositional differences of Proteobacteria in the biofilms as revealed by pyrosequencing and the relative abundance of their major families was compared (Fig. 3a). The Proteobacterial communities in the biofilm samples were highly diverse with more than 60 different families revealed. Rhodobacteraceae was found to be the most dominant family in all of the biofilms, followed by Sphingomonadaceae, Alteromonadaceae, and Phyllobacteriaceae with varying proportions among the different samples (Fig. 3b). Dominance of Rhodobacteraceae in
Microbial Communities in Natural Biofilms Fig. 5 Larval settlement induction of crude extracts from the biofilms tested in a single-dish and b double-dish choice assays. Settlement-inducing protein complex (SIPC), and autoclaved 0.22-μm filtered seawater were used as the positive and negative controls, respectively. Nonpolar fractions of the biofilm extracts were coated on Petri dishes. * indicates significant variation from the null hypothesis of 25:25 distribution. Data presented are mean+1 SD (n=3) for biofilms developed in the summer. Repeated experiments with extracts from biofilms developed in the spring showed similar results
biofilms was also reported in a recent study using Phylochip analysis [8]. Cluster analysis based on the occurrence and relative abundance of different families suggested a strong similarity among the Proteobacterial communities in the intertidal biofilms developed at different seasons (Fig. 3). Compared with highly diverse Proteobacterial communities, the Cyanobacterial communities in different biofilms were less complex and were dominated by chloroplast and Family I (Fig. S6). Although the relative abundance of Families III, XI and XII was consistently higher in the intertidal biofilms than in the subtidal biofilms, cluster analysis did not group the communities according to season or tidal level. The seasonal variations in biofilm microbial communities may stem from the differences in planktonic microbial communities shaped by seasonal environmental conditions [47], whereas the spatial variations in the communities can probably be attributed to the effects of tidal action and prolonged exposure to air on the intertidal biofilms [63]. Microbial community compositions revealed by the clone library showed certain similarities with those revealed by pyrosequencing. For instance, the major phyla recovered from classifying the clones using the RDP classifier were also Proteobacteria, Cyanobacteria and Bacteroidetes, and their relative abundance was similar. However, discrepancies were also observed among these two datasets, including fewer phyla (only eight), as well as fewer orders and genera, and a non-negligible proportion of unclassified clones (Table S5).
These discrepancies may mainly be attributed to sequencing depth and resolution but may also be due to differences in classification efficiency, sequence length and the confidence threshold applied. Phylogenetic analysis revealed that a large proportion of the clones resembled sequences from uncultured bacteria (Fig. 4). These uncultured bacteria are found in a variety of environments, including seawater, sediment, activated sludge, ocean crust, coral, sponge, and sea urchin, and a few of them have also been found in biofilms. In general, clones derived from different biofilm samples were found in the same clusters, showing no clear distinction among different tidal levels or seasons. Neither did LIBSHUFF analysis detect significant differences among the four libraries, as the lower p value of the comparisons of any two of the libraries was larger than the critical p value of 0.0043 (Table S6). However, it was noted from the phylogenetic trees that two or more clone sequences from a particular tidal level or season clustered together with the same closest relatives. These clusters were labeled as ‘tidal specific’ (containing sequences from either intertidal or subtidal levels regardless of season) and ‘season specific’ (containing sequences from either spring or summer biofilms regardless of tidal level) depending on their specificity. ‘Tidal specific’ clusters were found in all of the phyla detected except Bacteroidetes and most were found in the α-Proteobacterial and Cyanobacterial trees (Fig. 4). For instance, four clones from the biofilm developed at the subtidal level in spring (Fig. 4a; 5S44, 5S60, 5S58, and 5S9)
O.O. Lee et al.
clustered with Thalassobacter stenotrophicus isolated from Mediterranean seawater, and three clones from the biofilm developed at the intertidal level in the summer (Fig. 4a; 8 M23, 8 M9, 8 M77) were closely related to a Rhodobacterales bacterium isolated from Chesapeake Bay; similarly, another three clones from the subtidal biofilms (5S52, 8S48, 8S66) and two clones from the intertidal biofilms (5 M34, 8 M69) clustered with Acrosiphonia sp. SAG127.80 and an uncultured bacterium, respectively, in the Cyanobacterial tree (Fig. 4b). ‘Season specific’ clusters were observed in the Proteobacterial and Cyanobacterial trees (Fig. 4) but not in the phyla Bacteroidetes and Planctomycetes. The largest ‘season specific’ cluster was found in the αProteobacterial tree, which comprised five clones from the biofilms developed in the spring (5 M8, 5S40, 5 M18, 5 M51, and 5 M11) and their close relative Altererythrobacter sp. KYW2 from Kwangyang Bay (Fig. 4a). Although these findings were based on comparisons among a limited number of samples, they provided indications of potential candidates for future studies. The representativeness of these clusters and the establishment of such specificity in different biofilms have yet to be investigated. It should be pointed out that our libraries may not represent the whole communities due to the inherent limitations of the technique and the steep rarefaction curves obtained; therefore, caution should be taken when drawing any conclusions based on the clone library dataset. Relationship of Major or Specific Bacterial Groups Revealed in Biofilms with Larval Settlement In our study, Proteobacteria, particularly, Rhodobacteraceae in the α-Proteobacterial lineage, was found to be the most dominant group in all the biofilm samples (Fig. 4); a similar result was obtained by Phylochip analysis of subtidal biofilms in a recent study [8]. ‘Tidal specific’ clusters were also found within clones retrieved from either intertidal or subtidal level; the bacterial clones belong to the Rhodobacteraceae family and α-Proteobacteria (Fig. 4a). Members of Rhodobacteraceae, such as those in the Roseobacter clade, are chemolitoheterotrophs which are able to use a wide spectrum of carbon sources and oxidized sulfur as an electron acceptor [12]. This may explain their prevalence in both intertidal and subtidal biofilms. Certain strains of Rhodobacteraceae have been reported to produce acyl homoserine lactone quorum-sensing molecules in subtidal biofilms [27] and these molecules can be recognized by other bacteria and hence may mediate biofilm formation and the subsequent larval settlement of marine invertebrates [26, 61]. In addition, a number of Proteobacteria isolates have been reported to inhibit larval settlement of the barnacle B. amphitrite [39]. The diverse and large population of Proteobacteria in the subtidal biofilms may comprise a number of strains that inhibit larval settlement, possibly leading to
the reduced larval settlement preference. Because concern has been raised on the classification of short V6 pyrosequencing reads into lower taxonomic levels using the RDP classifier [10], more accurate taxonomic identification of the proteobacterial reads should be obtained before a solid correlation can be made with larval settlement. In contrast, Cyanobacteria have been reported to induce larval settlement of molluscs [45], but their ability to induce or inhibit the larval settlement of barnacles has not been studied in detail. In our study, certain Cyanobacteria were specifically found in biofilms from a particular tidal level (Fig 4b) and such specificity may be related to the differential larval settlement preference in B. amphitrite. Cyanobacteria have dual nomenclature systems (bacteriological and botanical), which cause considerable confusion in their taxonomy [50] and difficulties in describing their community beyond the family level. In the present study, the chloroplasts in algae Cylindrotheca closterium and Thalassiosira eccentrica probably accounted for the proliferation of Cyanobacteria components, as indicated by Fig.4b. The presence of these algae in the biofilms and their potential role in larvae settlement need further efforts to provide evidence. Chemically Mediated Larval Settlement Preference To determine whether the settlement preference of B. amphitrite cyprids is chemically mediated, biofilms developed from both intertidal and subtidal levels in both seasons were extracted with chemical solvents and the crude extracts were tested for the induction of larval settlement. Because no appropriate absorbent was available to immobilize the polar fraction of the biofilm extracts, only the nonpolar fraction was examined. Our results indicated that the crude extracts from both intertidal and subtidal biofilms induced the settlement of the cyprids as effectively as the settlement-inducing complex in single-dish assays (Fig. 5a). However, when both intertidal and subtidal biofilm extracts were provided in a choice assay, the cyprids showed a clear preference to settle on the Petri dishes coated with extracts from the intertidal biofilms (Fig. 5b). In fact, many bioactive compounds related to the larval settlement of marine invertebrates are relatively nonpolar in nature. Kitamura et al. [35] isolated fatty acids from red algae that induced the larval settlement and metamorphosis of sea urchins. Similarly, a mixture of fatty acids induced larval settlement of the polychaete, Phragmatopoma californica [52].
Conclusions In conclusion, barcoded 16S rRNA gene pyrosequencing revealed an extraordinarily high diversity of complex microbial communities in subtropical marine biofilms, which is
Microbial Communities in Natural Biofilms
much greater than that previously estimated, with a large number of low-abundance populations and a non-negligible portion of unknown, and probably novel, bacteria. Notable differences in microbial and chemical compositions were observed in biofilms developed at different seasons and tidal levels, which support the dynamic nature of these subtropical biofilms. However, the overall dynamic changes in microbial and chemical compositions may not necessarily dictate the differential larval settlement preference of B. amphitrite. Rather, our results suggest that certain candidates in the biofilms, including Proteobacteria and Cyanobacteria, might have a crucial influence on larval settlement preference, and further studies focusing on these candidates would definitely be valuable. Although our study did not include microbial communities in the water column for comparison, it is reasonable to assume that, being the source of the microbial species in the biofilm, the differences observed in the biofilm microbial communities probably stemmed from variations in the planktonic communities. In previous studies, planktonic bacterial communities were reported to be highly diverse and environment-driven [53]. Any environmental parameters that vary seasonally or spatially can alter the planktonic communities or bacterial attachment, which in turn affect the establishment of a biofilm microbial community [16, 58]. The interrelationship among environmental parameters, the biofilm microbial community and larval settlement has still not been clearly resolved. However, the improved resolution of the biofilm community structure from this study provided a basis to direct us towards the identification of potential key players that regulate the ecological functions of biofilms. Acknowledgments This study was supported by the National Basic Research Program of China (973 Program, No. 2012CB417304), COMRA project of China (COMRRDA12SC02), and awards from the Deep Sea Institute of Science and Engineering, the Chinese Academy of Science (SIDSSE-201206) and from the King Abdullah University of Science and Technology granted to P.Y. Qian (SA-C0040/UK-C0016) and grant from HKSAR government (GRF661611) for Biofilm study.
References 1. Amann R, Ludwig W, Schleifer K (1995) Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiol Mol Biol Rev 59:143 2. Ashelford KE, Chuzhanova NA, Fry JC, Jones AJ, Weightman AJ (2005) At least 1 in 20 16S rRNA sequence records currently held in public repositories is estimated to contain substantial anomalies. Appl Environ Microbiol 71:7724–7736 3. Biddle J, Fitz-Gibbon S, Schuster S, Brenchley J, House C (2008) Metagenomic signatures of the Peru Margin subseafloor biosphere show a genetically distinct environment. Proc Natl Acad Sci U S A 105:10583 4. Bitton G (1994) Role of microorganisms in biogeochemical cycles. Wastewater microbiology. Wiley-Liss, New York, USA, pp 51–73 5. Capone D, Carpenter E (1982) Nitrogen fixation in the marine environment. Science 217:1140
6. Chao A (1984) Nonparametric estimation of the number of classes in a population. Scand J Stat 11:265–270 7. Chao A, Yang M (1993) Stopping rules and estimation for recapture debugging with unequal failure rates. Biometrika 80:193–201 8. Chung H, Lee O, Huang Y, Mok S, Kolter R, Qian P (2010) Bacterial community succession and chemical profiles of subtidal biofilms in relation to larval settlement of the polychaete Hydroides elegans. ISME J 4(6):817–828 9. Cole J, VanRaden P, O'Connell J, Van Tassell C, Sonstegard T, Schnabel R, Taylor J, Wiggans G (2009) Distribution and location of genetic effects for dairy traits. J Dairy Sci 92:2931 10. Cole JR, Wang Q, Cardenas E, Fish J, Chai B, Farris RJ, KulamSyed-Mohideen AS, McGarrell DM, Marsh T, Garrity GM, Tiedje JM (2009) The ribosomal database project: improved alignments and new tools for rRNA analysis. Nucl Acids Res 37:141–145 11. Costerton J, Cheng K, Geesey G, Ladd T, Nickel J, Dasgupta M, Marrie T (1987) Bacterial biofilms in nature and disease. Annu Rev Microbiol 41:435–464 12. Dang H, Lovell C (2002) Numerical dominance and phylotype diversity of marine Rhodobacter species during early colonization of submerged surfaces in coastal marine waters as determined by 16S ribosomal DNA sequence analysis and fluorescence in situ hybridization. Appl Environ Microbiol 68:496 13. Darwin C (1854) A monograph on the sub class Cirripedia. Trans Zool Soc London 22 14. Denef V, Mueller R, Banfield J (2010) AMD biofilms: using model communities to study microbial evolution and ecological complexity in nature. ISME J 4:599–610 15. Dobretsov S, Qian PY (2006) Facilitation and inhibition of larval attachment of the bryozoan Bugula neritina in association with mono-species and multi-species biofilms. J Exp Mar Biol Ecol 333: 263–274 16. Donlan R (2002) Biofilms: microbial life on surfaces. Emerg Infect Dis 8:881–890 17. Edgar R (2004) MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucl Acids Res 32:1792 18. Edwards R, Rodriguez-Brito B, Wegley L, Haynes M, Breitbart M, Peterson D, Saar M, Alexander S, Alexander E, Rohwer F (2006) Using pyrosequencing to shed light on deep mine microbial ecology. BMC Genom 7:57 19. Egert M, Friedrich M (2003) Formation of pseudo-terminal restriction fragments, a PCR-related bias affecting terminal restriction fragment length polymorphism analysis of microbial community structure. Appl Environ Microbiol 69:2555 20. Grosberg R (1982) Intertidal zonation of barnacles: the influence of planktonic zonation of larvae on vertical distribution of adults. Ecology 63:894–899 21. Guckert J, Antworth C, Nichols P, White D (1985) Phospholipid, ester-linked fatty acid profiles as reproducible assays for changes in prokaryotic community structure of estuarine sediments. FEMS Microbiol Lett 31:147–158 22. Hall-Stoodley L, Costerton J, Stoodley P (2004) Bacterial biofilms: from the natural environment to infectious diseases. Nat Rev Microbiol 2:95–108 23. Hansen M, Tolker-Nielsen T, Givskov M, Molin S (1998) Biased 16S rDNA PCR amplification caused by interference from DNA flanking the template region. FEMS Microbiol Ecol 26:141–149 24. Harder T, Thiyagarajan V, Qian P (2001) Effect of cyprid age on the settlement of Balanus amphitrite Darwin in response to natural biofilms. Biofouling 17:211–219 25. Holmstrom C, Rittschof D, Kjelleberg S (1992) Inhibition of settlement by larvae of Balanus amphitrite and Ciona intestinalis by a surface-colonizing marine bacterium. Appl Environ Microbiol 58:2111 26. Huang Y, Dobretsov S, Ki J, Yang L, Qian P (2007) Presence of acylhomoserine lactone in subtidal biofilm and the implication in larval
O.O. Lee et al.
27.
28.
29.
30.
31.
32.
33.
34.
35.
36. 37.
38.
39.
40.
41.
42.
43.
44.
45.
46. 47.
behavioral response in the polychaete Hydroides elegans. Microb Ecol 54:384–392 Huang Y, Ki J, Case R, Qian P (2008) Diversity and acyl-homoserine lactone production among subtidal biofilm- forming bacteria. Aquat Microb Ecol 52:185–193 Huggett M, Nedved B, Hadfield M (2009) Effects of initial surface wettability on biofilm formation and subsequent settlement of Hydroides elegans. Biofouling 25:387–399 Hung O, Thiyagarajan V, Qian P (2008) Preferential attachment of barnacle larvae to natural multi-species biofilms: does surface wettability matter? J Exp Mar Biol Ecol 361:36–41 Hung O, Thiyagarajan V, Zhang R, Wu R, Qian P (2007) Attachment of Balanus amphitrite larvae to biofilms originating from contrasting environments. Mar Ecol Prog Ser 333:229–242 Kaeberlein T, Lewis K, Epstein S (2002) Isolating “uncultivable” microorganisms in pure culture in a simulated natural environment. Science 296:1127 Kemp P, Aller J (2004) Estimating prokaryotic diversity: when are 16 S rDNA libraries large enough? Limnol Oceanogr Methods 2: 114–125 Kirchman D, Graham S, Reish D, Mitchell R (1981) Bacteria induce settlement and metamorphosis of Janua (Dexiospira) brasiliensis Grube (Polychaeta: Spirprbidae). J Exp Biol Ecol 56:153–163 Kisand V, Wikner J (2003) Limited resolution of 16S rDNA DGGE caused by melting properties and closely related DNA sequences. J Microbiol Methods 54:183–191 Kitamura H, Kitahara S, Koh H (1993) The induction of larval settlement and metamorphosis of two sea urchins, Pseudocentrotus depressus and Anthocidaris crassispina, by free fatty acids extracted from the coralline red alga Corallina pilulifera. Mar Biol 115: 387–392 Kumar C, Anand S (1998) Significance of microbial biofilms in food industry: a review. Int J Food Microbiol 42:9–27 Lane D (1991) 16S/23S rRNA sequencing. In: Stackebrandt E, Goodfellow M (eds) Nucleic acid techniques in bacterial systematics. Wiley, Chichester, UK, pp 115–175 Lau S, Thiyagarajan V, Cheung S, Qian P (2005) Roles of bacterial community composition in biofilms as a mediator for larval settlement of three marine invertebrates. Aquat Microb Ecol 38:41–51 Lau S, Thiyagarajan V, Qian P (2003) The bioactivity of bacterial isolates in Hong Kong waters for the inhibition of barnacle (Balanus amphitrite Darwin) settlement. J Exp Mar Biol Ecol 282:43–60 Lee O, Wong Y, Qian P (2009) Inter- and intraspecific variations of bacterial communities associated with marine sponges from San Juan Island, Washington. Appl Environ Microbiol 75:3513 Lewandowski Z (2000) Structure and function of biofilms. In: Evans L (ed) Biofilms: recent advances in their study and control. CRC Press, Amsterdam, The Netherlands, pp 1–17 Lozupone C, Knight R (2005) UniFrac: a new phylogenetic method for comparing microbial communities. Appl Environ Microbiol 71:8228 Maki J, Rittschof D, Samuelsson M, Szewzyk U, Yule A, Kjelleberg S, Costlow J, Mitchell R (1990) Effect of marine bacteria and their exopolymers on the attachment of barnacle cypris larvae. Bull Mar Sci 46:499–511 Miyamoto Y, Noda T, Nakao S (1999) Zonation of two barnacle species not determined by competition. J Mar Biol Assoc U K 79: 621–628 Morse A, Froyd C, Morse D (1984) Molecules from cyanobacteria and red algae that induce larval settlement and metamorphosis in the mollusc Haliotis rufescens. Mar Biol 81:293–298 Morton L, Surman S (1994) Biofilms in biodeterioration — a review. Int Biodeterior Biodegrad 34:203–221 Murray AE, Preston CM, Massana R, Taylor LT, Blakis A, Wu K, DeLong EF (1998) Seasonal and spatial variability of bacterial and
48.
49. 50.
51.
52.
53.
54. 55.
56.
57.
58.
59.
60.
61. 62.
63.
64.
65.
66.
archaeal assemblages in the coastal waters near Anvers Island, Antarctica. Appl Environ Microbiol 64:2585–2595 Muyzer G, Smalla K (1998) Application of denaturing gradient gel electrophoresis (DGGE) and temperature gradient gel electrophoresis (TGGE) in microbial ecology. Anton Leeuwenh 73: 127–141 Nicolella C, Van Loosdrecht M, Heijnen J (2000) Wastewater treatment with particulate biofilm reactors. J Biotechnol 80:1–33 Oren A (2004) A proposal for further integration of the cyanobacteria under the Bacteriological Code. Int J Syst Evol Microbiol 54:1895 Hung OS, Lee OO, Thiyagarajan V, He HP, Xu Y, Chung HC, Qiu JW, Qian PY (2009) Characterization of cues from natural multispecies biofilms that induce larval attachment of the polychaete Hydroides elegans. Aquat Biol 4:253–262 Pawlik J (1986) Chemical induction of larval settlement and metamorphosis in the reef-building tube worm Phragmatopoma californica (Sabellariidae: Polychaeta). Mar Biol 91:59–68 Pommier T, Canback B, Riemann L, Bostrom KH, Simu K, Lundberg P, Tunlid A, Hagstrom Å (2007) Global patterns of diversity and community structure in marine bacterioplankton. Mol Ecol 16:867–880 Qian P (1999) Larval settlement of polychaetes. Hydrobiologia 402: 239–253 Qian P, Thiyagarajan V, Lau S, Cheung S (2003) Relationship between bacterial community profile in biofilm and attachment of the acorn barnacle Balanus amphitrite. Aquat Microb Ecol 33:225–237 Saeed A, Bhagabati N, Braisted J, Liang W, Sharov V, Howe E, Li J, Thiagarajan M, White J, Quackenbush J (2006) TM4 microarray software suite. Methods Enzymol 411:134–193 Schloss P, Handelsman J (2005) Introducing DOTUR, a computer program for defining operational taxonomic units and estimating species richness. Appl Environ Microbiol 71:1501 Siebel MA, Charaklis W, Prieur D (1989) Seasonal variations in bacterial colonisation of stainless steel, aluminium and polycarbonate surfaces in sea water flow system. Biofouling 1:251–261 Singleton DR, Furlong MA, Rathbun SL, Whitman WB (2001) Quantitative comparisons of 16S rRNA gene sequence libraries from environmental samples. Appl Environ Microbiol 67:4374–4376 Sogin M, Morrison H, Huber J, Welch D, Huse S, Neal P, Arrieta J, Herndl G (2006) Microbial diversity in the deep sea and the underexplored “rare biosphere”. Proc Natl Acad Sci U S A 103: 12115 Steinberg P, de Nys R, Kjelleberg S (2002) Chemical cues for surface colonization. J Chem Ecol 28:1935–1951 Thiyagarajan V, Harder T, Qiu J, Qian P (2003) Energy content at metamorphosis and growth rate of the early juvenile barnacle Balanus amphitrite. Mar Biol 143:543–554 Thiyagarajan V, Lau S, Cheung S, Qian P (2006) Cypris habitat selection facilitated by microbial films influences the vertical distribution of subtidal barnacle Balanus trigonus. Microb Ecol 51:431–440 Thompson JD, Higgins DG, Gibson TJ (1994) CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res 22: 4673–4680 Tyson G, Lo I, Baker B, Allen E, Hugenholtz P, Banfield J (2005) Genome-directed isolation of the key nitrogen fixer Leptospirillum ferrodiazotrophum sp. nov. from an acidophilic microbial community. Appl Environ Microbiol 71:6319 Unabia C, Hadfield M (1999) Role of bacteria in larval settlement and metamorphosis of the polychaete Hydroides elegans. Mar Biol 133:55–64
Microbial Communities in Natural Biofilms 67. Wagner M, Smidt H, Loy A, Zhou J (2007) Unravelling microbial communities with DNA-microarrays: challenges and future directions. Microb Ecol 53:498–506 68. Wang G, Kennedy S, Fasiludeen S, Rensing C, DasSarma S (2004) Arsenic resistance in Halobacterium sp. strain NRC-1 examined by using an improved gene knockout system. J Bacteriol 186:3187 69. Wieczoreck S, Clare A, Tood C (1995) Inhibitory and facilitatory effects of microbial films on settlement of Balanus amphitrite larvae. Mar Ecol Prog Ser 119:221–228 70. Wieczorek S, Todd C (1998) Inhibition and facilitation of settlement of epifaunal marine invertebrate larvae by microbial biofilm cues. Biofouling 12:81–118
71. Zar J (1999) Biostatistical analysis, 4th edn. Prentice-Hall, Englewood Cliffs, NJ 72. Zelles L, Bai Q, Beck T, Beese F (1992) Signature fatty acids in phospholipids and lipopolysaccharides as indicators of microbial biomass and community structure in agricultural soils. Soil Biol Biochem 24:317–323 73. Zhou J (2003) Microarrays for bacterial detection and microbial community analysis. Curr Opin Microbiol 6:288–294 74. Zhou J, Bruns M, Tiedje J (1996) DNA recovery from soils of diverse composition. Appl Environ Microbiol 62:316–322 75. Zimmer R, Butman C (2000) Chemical signaling processes in the marine environment. Biol Bull 198:168
ENVIRONMENTAL MICROBIOLOGY
Molecular Techniques Revealed Highly Diverse Microbial Communities in Natural Marine Biofilms on Polystyrene Dishes for Invertebrate Larval Settlement On On Lee & Hong Chun Chung & Jiangke Yang & Yong Wang & Swagatika Dash & Hao Wang & Pei-Yuan Qian
Received: 24 May 2013 / Accepted: 6 December 2013 # Springer Science+Business Media New York 2014
Abstract Biofilm microbial communities play an important role in the larval settlement response of marine invertebrates. However, the underlying mechanism has yet to be resolved, mainly because of the uncertainties in characterizing members in the communities using traditional 16S rRNA gene-based molecular methods and in identifying the chemical signals involved. In this study, pyrosequencing was used to characterize the bacterial communities in intertidal and subtidal marine biofilms developed during two seasons. We revealed highly diverse biofilm bacterial communities that varied with season and tidal level. Over 3,000 operational taxonomic units with estimates of up to 8,000 species were recovered in a biofilm sample, which is by far the highest number recorded in subtropical marine biofilms. Nineteen phyla were found, of which Cyanobacteria and Proteobacteria were the most dominant one in the intertidal and subtidal biofilms, respectively. Apart from these, Actinobacteria, Bacteroidetes, and Planctomycetes were the major groups recovered in both intertidal and subtidal biofilms, although their relative abundance varied among samples. Full-length 16S rRNA gene clone libraries were constructed for the four biofilm samples and showed similar bacterial compositions at the phylum level to those revealed by pyrosequencing. Laboratory assays confirmed that cyrids of the barnacle Balanus amphitrite preferred Lee and Chung made equal contributions to the work Electronic supplementary material The online version of this article (doi:10.1007/s00248-013-0348-3) contains supplementary material, which is available to authorized users. O. O. Lee : H. C. Chung : J. Yang : Y. Wang : S. Dash : H. Wang : P.10 % for major families)
using the clone library technique. In addition, the rank abundance curves indicated a high proportion of rare organisms in all biofilm samples (Fig. S3b). Based on pyrosequencing data, a higher microbial diversity (with more than 3,000 OTUs and estimates of up to 8,000 species phylotypes) was found in the biofilms developed in the spring than in those developed in the summer, whereas tidal level seemed to have less influence on microbial diversity in biofilms (Table 1). The Chao1 and ACE estimators were approximately two-fold higher than the number of OTUs in all samples (Tables 1 and S2), indicating that the actual number of species may be doubled. The numbers of OTUs recovered in the spring intertidal biofilms were comparable with those in deep-seawater and sponges revealed by the same technique [60]. Using the clone library technique, the bacterial diversities in our subtropical marine biofilms were substantially higher than those of biofilms from different habitats [32] but were comparable with the planktonic communities in many marine environments [53]. In general, bacterial communities in the subtidal biofilms developed in this study were slightly more diverse than those of the intertidal biofilms, whereas biofilms developed in the spring were substantially more diverse than those developed in the summer (see Shannon index, Table 1), indicating that season may have a greater influence on microbial diversity than tidal height. In contrast, a clear preference of the barnacle cyprids to settle on intertidal biofilms, regardless of the season, was observed, although the microbial diversity and
Fig. 4 16S rRNA-based phylogenetic trees showing the genetic distances among clones retrieved from the libraries for biofilms developed at different tidal levels in different seasons in reference to members of the phyla a α-Proteobacteria and b Cyanobacteria. The trees were constructed based on the neighbor-joining method. Clones from different biofilms are prefixed with the sample ID (see Table 1 for details) followed by clone number. Reference sequences from the closest relatives are retrieved from NCBI and those originated from seawater, sediment, activated sludge, corals, sponges, sea urchin, ocean crust, and biofilms are indicated by the superscripts “SW,” “SD,” “AS,” “C,” “S,” “U,” “OC,” and “BF,” respectively. Nucleotide accession numbers of the reference sequences are given in parentheses. Solid boxes show “tidalspecific” clusters, whereas dotted boxes show “season-specific” clusters. The scale bar represents percent substitutions per nucleotide position. Bootstrap values of >50 % based on 1,000 re-samplings are indicated by the numbers at the nodes
Microbial Communities in Natural Biofilms
density of the biofilms were similar, which suggests that the preference may not be related to microbial diversity or density, and that other parameters in biofilms, such as community composition, may have a greater influence.
Microbial Composition in Biofilms To determine the spatial and seasonal variations in microbial community composition of biofilms, we classified the
O.O. Lee et al.
Fig. 4 continued.
pyrosequencing tags into different phyla at a confidence threshold of 50 % using the RDP classifier. About 3–6 % of the reads could not be assigned to a known phylum, and the remaining reads were classified into 19 different phyla; Bacteroidetes, Cyanobacteria and Proteobacteria accounted for >75 % of the reads, and were the major phyla detected in all of the biofilms (Fig. 2; Table S3). Classification of the reads into lower taxonomic levels revealed extremely diverse bacterial communities in all biofilm samples, with up to 70 orders (16 of which constituted >1 % of the reads) and 500 genera (28 of which constituted >1 % of the reads) (Table S4). Although all the biofilm samples were dominated by Cyanobacteria, Proteobacteria, and Bacteroidetes, regardless of the method used, notable differences but similar trends in the proportions of these phyla (i.e., composition) were observed between the intertidal and subtidal biofilms (Figs. 2 and S4). For instance, in both seasons, Cyanobacteria accounted for more reads/clones in the intertidal biofilms than in the subtidal biofilms; conversely, more reads/clones were assigned to Proteobacteria and Planctomycetes in the subtidal
biofilms than in the intertidal biofilms. Seasonal variations in the bacterial communities were also revealed by pyrosequencing (Fig. 2; Table S3). More pyrosequencing reads were assigned to Actinobacteria, Chloroflexi, and Firmicutes in the biofilms developed in the summer than in those developed in the spring. Unifrac PCA analysis detected substantial differences among bacterial communities in the biofilms developed at different tidal levels in the summer and in the biofilms developed in different seasons for the same tidal level (Fig. S5). We constructed a heatmap to illustrate the compositional differences of Proteobacteria in the biofilms as revealed by pyrosequencing and the relative abundance of their major families was compared (Fig. 3a). The Proteobacterial communities in the biofilm samples were highly diverse with more than 60 different families revealed. Rhodobacteraceae was found to be the most dominant family in all of the biofilms, followed by Sphingomonadaceae, Alteromonadaceae, and Phyllobacteriaceae with varying proportions among the different samples (Fig. 3b). Dominance of Rhodobacteraceae in
Microbial Communities in Natural Biofilms Fig. 5 Larval settlement induction of crude extracts from the biofilms tested in a single-dish and b double-dish choice assays. Settlement-inducing protein complex (SIPC), and autoclaved 0.22-μm filtered seawater were used as the positive and negative controls, respectively. Nonpolar fractions of the biofilm extracts were coated on Petri dishes. * indicates significant variation from the null hypothesis of 25:25 distribution. Data presented are mean+1 SD (n=3) for biofilms developed in the summer. Repeated experiments with extracts from biofilms developed in the spring showed similar results
biofilms was also reported in a recent study using Phylochip analysis [8]. Cluster analysis based on the occurrence and relative abundance of different families suggested a strong similarity among the Proteobacterial communities in the intertidal biofilms developed at different seasons (Fig. 3). Compared with highly diverse Proteobacterial communities, the Cyanobacterial communities in different biofilms were less complex and were dominated by chloroplast and Family I (Fig. S6). Although the relative abundance of Families III, XI and XII was consistently higher in the intertidal biofilms than in the subtidal biofilms, cluster analysis did not group the communities according to season or tidal level. The seasonal variations in biofilm microbial communities may stem from the differences in planktonic microbial communities shaped by seasonal environmental conditions [47], whereas the spatial variations in the communities can probably be attributed to the effects of tidal action and prolonged exposure to air on the intertidal biofilms [63]. Microbial community compositions revealed by the clone library showed certain similarities with those revealed by pyrosequencing. For instance, the major phyla recovered from classifying the clones using the RDP classifier were also Proteobacteria, Cyanobacteria and Bacteroidetes, and their relative abundance was similar. However, discrepancies were also observed among these two datasets, including fewer phyla (only eight), as well as fewer orders and genera, and a non-negligible proportion of unclassified clones (Table S5).
These discrepancies may mainly be attributed to sequencing depth and resolution but may also be due to differences in classification efficiency, sequence length and the confidence threshold applied. Phylogenetic analysis revealed that a large proportion of the clones resembled sequences from uncultured bacteria (Fig. 4). These uncultured bacteria are found in a variety of environments, including seawater, sediment, activated sludge, ocean crust, coral, sponge, and sea urchin, and a few of them have also been found in biofilms. In general, clones derived from different biofilm samples were found in the same clusters, showing no clear distinction among different tidal levels or seasons. Neither did LIBSHUFF analysis detect significant differences among the four libraries, as the lower p value of the comparisons of any two of the libraries was larger than the critical p value of 0.0043 (Table S6). However, it was noted from the phylogenetic trees that two or more clone sequences from a particular tidal level or season clustered together with the same closest relatives. These clusters were labeled as ‘tidal specific’ (containing sequences from either intertidal or subtidal levels regardless of season) and ‘season specific’ (containing sequences from either spring or summer biofilms regardless of tidal level) depending on their specificity. ‘Tidal specific’ clusters were found in all of the phyla detected except Bacteroidetes and most were found in the α-Proteobacterial and Cyanobacterial trees (Fig. 4). For instance, four clones from the biofilm developed at the subtidal level in spring (Fig. 4a; 5S44, 5S60, 5S58, and 5S9)
O.O. Lee et al.
clustered with Thalassobacter stenotrophicus isolated from Mediterranean seawater, and three clones from the biofilm developed at the intertidal level in the summer (Fig. 4a; 8 M23, 8 M9, 8 M77) were closely related to a Rhodobacterales bacterium isolated from Chesapeake Bay; similarly, another three clones from the subtidal biofilms (5S52, 8S48, 8S66) and two clones from the intertidal biofilms (5 M34, 8 M69) clustered with Acrosiphonia sp. SAG127.80 and an uncultured bacterium, respectively, in the Cyanobacterial tree (Fig. 4b). ‘Season specific’ clusters were observed in the Proteobacterial and Cyanobacterial trees (Fig. 4) but not in the phyla Bacteroidetes and Planctomycetes. The largest ‘season specific’ cluster was found in the αProteobacterial tree, which comprised five clones from the biofilms developed in the spring (5 M8, 5S40, 5 M18, 5 M51, and 5 M11) and their close relative Altererythrobacter sp. KYW2 from Kwangyang Bay (Fig. 4a). Although these findings were based on comparisons among a limited number of samples, they provided indications of potential candidates for future studies. The representativeness of these clusters and the establishment of such specificity in different biofilms have yet to be investigated. It should be pointed out that our libraries may not represent the whole communities due to the inherent limitations of the technique and the steep rarefaction curves obtained; therefore, caution should be taken when drawing any conclusions based on the clone library dataset. Relationship of Major or Specific Bacterial Groups Revealed in Biofilms with Larval Settlement In our study, Proteobacteria, particularly, Rhodobacteraceae in the α-Proteobacterial lineage, was found to be the most dominant group in all the biofilm samples (Fig. 4); a similar result was obtained by Phylochip analysis of subtidal biofilms in a recent study [8]. ‘Tidal specific’ clusters were also found within clones retrieved from either intertidal or subtidal level; the bacterial clones belong to the Rhodobacteraceae family and α-Proteobacteria (Fig. 4a). Members of Rhodobacteraceae, such as those in the Roseobacter clade, are chemolitoheterotrophs which are able to use a wide spectrum of carbon sources and oxidized sulfur as an electron acceptor [12]. This may explain their prevalence in both intertidal and subtidal biofilms. Certain strains of Rhodobacteraceae have been reported to produce acyl homoserine lactone quorum-sensing molecules in subtidal biofilms [27] and these molecules can be recognized by other bacteria and hence may mediate biofilm formation and the subsequent larval settlement of marine invertebrates [26, 61]. In addition, a number of Proteobacteria isolates have been reported to inhibit larval settlement of the barnacle B. amphitrite [39]. The diverse and large population of Proteobacteria in the subtidal biofilms may comprise a number of strains that inhibit larval settlement, possibly leading to
the reduced larval settlement preference. Because concern has been raised on the classification of short V6 pyrosequencing reads into lower taxonomic levels using the RDP classifier [10], more accurate taxonomic identification of the proteobacterial reads should be obtained before a solid correlation can be made with larval settlement. In contrast, Cyanobacteria have been reported to induce larval settlement of molluscs [45], but their ability to induce or inhibit the larval settlement of barnacles has not been studied in detail. In our study, certain Cyanobacteria were specifically found in biofilms from a particular tidal level (Fig 4b) and such specificity may be related to the differential larval settlement preference in B. amphitrite. Cyanobacteria have dual nomenclature systems (bacteriological and botanical), which cause considerable confusion in their taxonomy [50] and difficulties in describing their community beyond the family level. In the present study, the chloroplasts in algae Cylindrotheca closterium and Thalassiosira eccentrica probably accounted for the proliferation of Cyanobacteria components, as indicated by Fig.4b. The presence of these algae in the biofilms and their potential role in larvae settlement need further efforts to provide evidence. Chemically Mediated Larval Settlement Preference To determine whether the settlement preference of B. amphitrite cyprids is chemically mediated, biofilms developed from both intertidal and subtidal levels in both seasons were extracted with chemical solvents and the crude extracts were tested for the induction of larval settlement. Because no appropriate absorbent was available to immobilize the polar fraction of the biofilm extracts, only the nonpolar fraction was examined. Our results indicated that the crude extracts from both intertidal and subtidal biofilms induced the settlement of the cyprids as effectively as the settlement-inducing complex in single-dish assays (Fig. 5a). However, when both intertidal and subtidal biofilm extracts were provided in a choice assay, the cyprids showed a clear preference to settle on the Petri dishes coated with extracts from the intertidal biofilms (Fig. 5b). In fact, many bioactive compounds related to the larval settlement of marine invertebrates are relatively nonpolar in nature. Kitamura et al. [35] isolated fatty acids from red algae that induced the larval settlement and metamorphosis of sea urchins. Similarly, a mixture of fatty acids induced larval settlement of the polychaete, Phragmatopoma californica [52].
Conclusions In conclusion, barcoded 16S rRNA gene pyrosequencing revealed an extraordinarily high diversity of complex microbial communities in subtropical marine biofilms, which is
Microbial Communities in Natural Biofilms
much greater than that previously estimated, with a large number of low-abundance populations and a non-negligible portion of unknown, and probably novel, bacteria. Notable differences in microbial and chemical compositions were observed in biofilms developed at different seasons and tidal levels, which support the dynamic nature of these subtropical biofilms. However, the overall dynamic changes in microbial and chemical compositions may not necessarily dictate the differential larval settlement preference of B. amphitrite. Rather, our results suggest that certain candidates in the biofilms, including Proteobacteria and Cyanobacteria, might have a crucial influence on larval settlement preference, and further studies focusing on these candidates would definitely be valuable. Although our study did not include microbial communities in the water column for comparison, it is reasonable to assume that, being the source of the microbial species in the biofilm, the differences observed in the biofilm microbial communities probably stemmed from variations in the planktonic communities. In previous studies, planktonic bacterial communities were reported to be highly diverse and environment-driven [53]. Any environmental parameters that vary seasonally or spatially can alter the planktonic communities or bacterial attachment, which in turn affect the establishment of a biofilm microbial community [16, 58]. The interrelationship among environmental parameters, the biofilm microbial community and larval settlement has still not been clearly resolved. However, the improved resolution of the biofilm community structure from this study provided a basis to direct us towards the identification of potential key players that regulate the ecological functions of biofilms. Acknowledgments This study was supported by the National Basic Research Program of China (973 Program, No. 2012CB417304), COMRA project of China (COMRRDA12SC02), and awards from the Deep Sea Institute of Science and Engineering, the Chinese Academy of Science (SIDSSE-201206) and from the King Abdullah University of Science and Technology granted to P.Y. Qian (SA-C0040/UK-C0016) and grant from HKSAR government (GRF661611) for Biofilm study.
References 1. Amann R, Ludwig W, Schleifer K (1995) Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiol Mol Biol Rev 59:143 2. Ashelford KE, Chuzhanova NA, Fry JC, Jones AJ, Weightman AJ (2005) At least 1 in 20 16S rRNA sequence records currently held in public repositories is estimated to contain substantial anomalies. Appl Environ Microbiol 71:7724–7736 3. Biddle J, Fitz-Gibbon S, Schuster S, Brenchley J, House C (2008) Metagenomic signatures of the Peru Margin subseafloor biosphere show a genetically distinct environment. Proc Natl Acad Sci U S A 105:10583 4. Bitton G (1994) Role of microorganisms in biogeochemical cycles. Wastewater microbiology. Wiley-Liss, New York, USA, pp 51–73 5. Capone D, Carpenter E (1982) Nitrogen fixation in the marine environment. Science 217:1140
6. Chao A (1984) Nonparametric estimation of the number of classes in a population. Scand J Stat 11:265–270 7. Chao A, Yang M (1993) Stopping rules and estimation for recapture debugging with unequal failure rates. Biometrika 80:193–201 8. Chung H, Lee O, Huang Y, Mok S, Kolter R, Qian P (2010) Bacterial community succession and chemical profiles of subtidal biofilms in relation to larval settlement of the polychaete Hydroides elegans. ISME J 4(6):817–828 9. Cole J, VanRaden P, O'Connell J, Van Tassell C, Sonstegard T, Schnabel R, Taylor J, Wiggans G (2009) Distribution and location of genetic effects for dairy traits. J Dairy Sci 92:2931 10. Cole JR, Wang Q, Cardenas E, Fish J, Chai B, Farris RJ, KulamSyed-Mohideen AS, McGarrell DM, Marsh T, Garrity GM, Tiedje JM (2009) The ribosomal database project: improved alignments and new tools for rRNA analysis. Nucl Acids Res 37:141–145 11. Costerton J, Cheng K, Geesey G, Ladd T, Nickel J, Dasgupta M, Marrie T (1987) Bacterial biofilms in nature and disease. Annu Rev Microbiol 41:435–464 12. Dang H, Lovell C (2002) Numerical dominance and phylotype diversity of marine Rhodobacter species during early colonization of submerged surfaces in coastal marine waters as determined by 16S ribosomal DNA sequence analysis and fluorescence in situ hybridization. Appl Environ Microbiol 68:496 13. Darwin C (1854) A monograph on the sub class Cirripedia. Trans Zool Soc London 22 14. Denef V, Mueller R, Banfield J (2010) AMD biofilms: using model communities to study microbial evolution and ecological complexity in nature. ISME J 4:599–610 15. Dobretsov S, Qian PY (2006) Facilitation and inhibition of larval attachment of the bryozoan Bugula neritina in association with mono-species and multi-species biofilms. J Exp Mar Biol Ecol 333: 263–274 16. Donlan R (2002) Biofilms: microbial life on surfaces. Emerg Infect Dis 8:881–890 17. Edgar R (2004) MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucl Acids Res 32:1792 18. Edwards R, Rodriguez-Brito B, Wegley L, Haynes M, Breitbart M, Peterson D, Saar M, Alexander S, Alexander E, Rohwer F (2006) Using pyrosequencing to shed light on deep mine microbial ecology. BMC Genom 7:57 19. Egert M, Friedrich M (2003) Formation of pseudo-terminal restriction fragments, a PCR-related bias affecting terminal restriction fragment length polymorphism analysis of microbial community structure. Appl Environ Microbiol 69:2555 20. Grosberg R (1982) Intertidal zonation of barnacles: the influence of planktonic zonation of larvae on vertical distribution of adults. Ecology 63:894–899 21. Guckert J, Antworth C, Nichols P, White D (1985) Phospholipid, ester-linked fatty acid profiles as reproducible assays for changes in prokaryotic community structure of estuarine sediments. FEMS Microbiol Lett 31:147–158 22. Hall-Stoodley L, Costerton J, Stoodley P (2004) Bacterial biofilms: from the natural environment to infectious diseases. Nat Rev Microbiol 2:95–108 23. Hansen M, Tolker-Nielsen T, Givskov M, Molin S (1998) Biased 16S rDNA PCR amplification caused by interference from DNA flanking the template region. FEMS Microbiol Ecol 26:141–149 24. Harder T, Thiyagarajan V, Qian P (2001) Effect of cyprid age on the settlement of Balanus amphitrite Darwin in response to natural biofilms. Biofouling 17:211–219 25. Holmstrom C, Rittschof D, Kjelleberg S (1992) Inhibition of settlement by larvae of Balanus amphitrite and Ciona intestinalis by a surface-colonizing marine bacterium. Appl Environ Microbiol 58:2111 26. Huang Y, Dobretsov S, Ki J, Yang L, Qian P (2007) Presence of acylhomoserine lactone in subtidal biofilm and the implication in larval
O.O. Lee et al.
27.
28.
29.
30.
31.
32.
33.
34.
35.
36. 37.
38.
39.
40.
41.
42.
43.
44.
45.
46. 47.
behavioral response in the polychaete Hydroides elegans. Microb Ecol 54:384–392 Huang Y, Ki J, Case R, Qian P (2008) Diversity and acyl-homoserine lactone production among subtidal biofilm- forming bacteria. Aquat Microb Ecol 52:185–193 Huggett M, Nedved B, Hadfield M (2009) Effects of initial surface wettability on biofilm formation and subsequent settlement of Hydroides elegans. Biofouling 25:387–399 Hung O, Thiyagarajan V, Qian P (2008) Preferential attachment of barnacle larvae to natural multi-species biofilms: does surface wettability matter? J Exp Mar Biol Ecol 361:36–41 Hung O, Thiyagarajan V, Zhang R, Wu R, Qian P (2007) Attachment of Balanus amphitrite larvae to biofilms originating from contrasting environments. Mar Ecol Prog Ser 333:229–242 Kaeberlein T, Lewis K, Epstein S (2002) Isolating “uncultivable” microorganisms in pure culture in a simulated natural environment. Science 296:1127 Kemp P, Aller J (2004) Estimating prokaryotic diversity: when are 16 S rDNA libraries large enough? Limnol Oceanogr Methods 2: 114–125 Kirchman D, Graham S, Reish D, Mitchell R (1981) Bacteria induce settlement and metamorphosis of Janua (Dexiospira) brasiliensis Grube (Polychaeta: Spirprbidae). J Exp Biol Ecol 56:153–163 Kisand V, Wikner J (2003) Limited resolution of 16S rDNA DGGE caused by melting properties and closely related DNA sequences. J Microbiol Methods 54:183–191 Kitamura H, Kitahara S, Koh H (1993) The induction of larval settlement and metamorphosis of two sea urchins, Pseudocentrotus depressus and Anthocidaris crassispina, by free fatty acids extracted from the coralline red alga Corallina pilulifera. Mar Biol 115: 387–392 Kumar C, Anand S (1998) Significance of microbial biofilms in food industry: a review. Int J Food Microbiol 42:9–27 Lane D (1991) 16S/23S rRNA sequencing. In: Stackebrandt E, Goodfellow M (eds) Nucleic acid techniques in bacterial systematics. Wiley, Chichester, UK, pp 115–175 Lau S, Thiyagarajan V, Cheung S, Qian P (2005) Roles of bacterial community composition in biofilms as a mediator for larval settlement of three marine invertebrates. Aquat Microb Ecol 38:41–51 Lau S, Thiyagarajan V, Qian P (2003) The bioactivity of bacterial isolates in Hong Kong waters for the inhibition of barnacle (Balanus amphitrite Darwin) settlement. J Exp Mar Biol Ecol 282:43–60 Lee O, Wong Y, Qian P (2009) Inter- and intraspecific variations of bacterial communities associated with marine sponges from San Juan Island, Washington. Appl Environ Microbiol 75:3513 Lewandowski Z (2000) Structure and function of biofilms. In: Evans L (ed) Biofilms: recent advances in their study and control. CRC Press, Amsterdam, The Netherlands, pp 1–17 Lozupone C, Knight R (2005) UniFrac: a new phylogenetic method for comparing microbial communities. Appl Environ Microbiol 71:8228 Maki J, Rittschof D, Samuelsson M, Szewzyk U, Yule A, Kjelleberg S, Costlow J, Mitchell R (1990) Effect of marine bacteria and their exopolymers on the attachment of barnacle cypris larvae. Bull Mar Sci 46:499–511 Miyamoto Y, Noda T, Nakao S (1999) Zonation of two barnacle species not determined by competition. J Mar Biol Assoc U K 79: 621–628 Morse A, Froyd C, Morse D (1984) Molecules from cyanobacteria and red algae that induce larval settlement and metamorphosis in the mollusc Haliotis rufescens. Mar Biol 81:293–298 Morton L, Surman S (1994) Biofilms in biodeterioration — a review. Int Biodeterior Biodegrad 34:203–221 Murray AE, Preston CM, Massana R, Taylor LT, Blakis A, Wu K, DeLong EF (1998) Seasonal and spatial variability of bacterial and
48.
49. 50.
51.
52.
53.
54. 55.
56.
57.
58.
59.
60.
61. 62.
63.
64.
65.
66.
archaeal assemblages in the coastal waters near Anvers Island, Antarctica. Appl Environ Microbiol 64:2585–2595 Muyzer G, Smalla K (1998) Application of denaturing gradient gel electrophoresis (DGGE) and temperature gradient gel electrophoresis (TGGE) in microbial ecology. Anton Leeuwenh 73: 127–141 Nicolella C, Van Loosdrecht M, Heijnen J (2000) Wastewater treatment with particulate biofilm reactors. J Biotechnol 80:1–33 Oren A (2004) A proposal for further integration of the cyanobacteria under the Bacteriological Code. Int J Syst Evol Microbiol 54:1895 Hung OS, Lee OO, Thiyagarajan V, He HP, Xu Y, Chung HC, Qiu JW, Qian PY (2009) Characterization of cues from natural multispecies biofilms that induce larval attachment of the polychaete Hydroides elegans. Aquat Biol 4:253–262 Pawlik J (1986) Chemical induction of larval settlement and metamorphosis in the reef-building tube worm Phragmatopoma californica (Sabellariidae: Polychaeta). Mar Biol 91:59–68 Pommier T, Canback B, Riemann L, Bostrom KH, Simu K, Lundberg P, Tunlid A, Hagstrom Å (2007) Global patterns of diversity and community structure in marine bacterioplankton. Mol Ecol 16:867–880 Qian P (1999) Larval settlement of polychaetes. Hydrobiologia 402: 239–253 Qian P, Thiyagarajan V, Lau S, Cheung S (2003) Relationship between bacterial community profile in biofilm and attachment of the acorn barnacle Balanus amphitrite. Aquat Microb Ecol 33:225–237 Saeed A, Bhagabati N, Braisted J, Liang W, Sharov V, Howe E, Li J, Thiagarajan M, White J, Quackenbush J (2006) TM4 microarray software suite. Methods Enzymol 411:134–193 Schloss P, Handelsman J (2005) Introducing DOTUR, a computer program for defining operational taxonomic units and estimating species richness. Appl Environ Microbiol 71:1501 Siebel MA, Charaklis W, Prieur D (1989) Seasonal variations in bacterial colonisation of stainless steel, aluminium and polycarbonate surfaces in sea water flow system. Biofouling 1:251–261 Singleton DR, Furlong MA, Rathbun SL, Whitman WB (2001) Quantitative comparisons of 16S rRNA gene sequence libraries from environmental samples. Appl Environ Microbiol 67:4374–4376 Sogin M, Morrison H, Huber J, Welch D, Huse S, Neal P, Arrieta J, Herndl G (2006) Microbial diversity in the deep sea and the underexplored “rare biosphere”. Proc Natl Acad Sci U S A 103: 12115 Steinberg P, de Nys R, Kjelleberg S (2002) Chemical cues for surface colonization. J Chem Ecol 28:1935–1951 Thiyagarajan V, Harder T, Qiu J, Qian P (2003) Energy content at metamorphosis and growth rate of the early juvenile barnacle Balanus amphitrite. Mar Biol 143:543–554 Thiyagarajan V, Lau S, Cheung S, Qian P (2006) Cypris habitat selection facilitated by microbial films influences the vertical distribution of subtidal barnacle Balanus trigonus. Microb Ecol 51:431–440 Thompson JD, Higgins DG, Gibson TJ (1994) CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res 22: 4673–4680 Tyson G, Lo I, Baker B, Allen E, Hugenholtz P, Banfield J (2005) Genome-directed isolation of the key nitrogen fixer Leptospirillum ferrodiazotrophum sp. nov. from an acidophilic microbial community. Appl Environ Microbiol 71:6319 Unabia C, Hadfield M (1999) Role of bacteria in larval settlement and metamorphosis of the polychaete Hydroides elegans. Mar Biol 133:55–64
Microbial Communities in Natural Biofilms 67. Wagner M, Smidt H, Loy A, Zhou J (2007) Unravelling microbial communities with DNA-microarrays: challenges and future directions. Microb Ecol 53:498–506 68. Wang G, Kennedy S, Fasiludeen S, Rensing C, DasSarma S (2004) Arsenic resistance in Halobacterium sp. strain NRC-1 examined by using an improved gene knockout system. J Bacteriol 186:3187 69. Wieczoreck S, Clare A, Tood C (1995) Inhibitory and facilitatory effects of microbial films on settlement of Balanus amphitrite larvae. Mar Ecol Prog Ser 119:221–228 70. Wieczorek S, Todd C (1998) Inhibition and facilitation of settlement of epifaunal marine invertebrate larvae by microbial biofilm cues. Biofouling 12:81–118
71. Zar J (1999) Biostatistical analysis, 4th edn. Prentice-Hall, Englewood Cliffs, NJ 72. Zelles L, Bai Q, Beck T, Beese F (1992) Signature fatty acids in phospholipids and lipopolysaccharides as indicators of microbial biomass and community structure in agricultural soils. Soil Biol Biochem 24:317–323 73. Zhou J (2003) Microarrays for bacterial detection and microbial community analysis. Curr Opin Microbiol 6:288–294 74. Zhou J, Bruns M, Tiedje J (1996) DNA recovery from soils of diverse composition. Appl Environ Microbiol 62:316–322 75. Zimmer R, Butman C (2000) Chemical signaling processes in the marine environment. Biol Bull 198:168
