Download Gerardin, Y. Viral impact on microbial community composition in a

Viral impact on microbial community composition in a marine ecosystem Ylaine Gerardin Microbial Diversity 2012 [email protected] Abstract Marine environments contain a high concentration (~107 per mL) of viruses, which have been theorized to play an important role in limiting bacterial abundances. Previous experiments measuring the influence of viral-­‐mediated mortality measured changes in taxon abundance using fluorescence in situ hybridization (FISH). In this study, a sequencing-­‐based approach was used to try to find phage-­‐
limited bacterial species in a surface marine environment. Virus-­‐depleted cultures were incubated in situ and community composition analyzed by 16S RNA gene sequencing. Eight hits were identified that were preferentially enriched in virus-­‐free cultures. Analysis of communities sampled at two time points showed tight clustering of abundance-­‐weighted community composition within timepoints and divergence of these communities over time. Introduction Marine environments contain ten times the amount of viruses as bacteria. Viruses are thus likely play an important role in the microbiological community (Pernthaler, 2005). Estimates for viral mortality indicate that roughly 20% of marine bacteria are lysed by phages every day (Suttle, 2007). I hypothesize that removing viruses from a marine environment will relieve pressure on certain highly-­‐infected species that are not nutrient-­‐limited, allowing these species to increase in abundance. Previous research has been performed on culturing virus-­‐free communities. In these studies, phylogenetic change was quantified using FISH (Bouvier and del Giorgio, 2007) (Ferrera et al., 2011). However, FISH may not provide enough resolution to resolve differences in population abundances as a result of virus removal; the clades probed using FISH may contain species varying in growth rate, nutrient utilization, and phage susceptibility. In this study, a sequencing-­‐based approach was used to try to find phage-­‐limited bacterial species in a surface marine environment. Virus-­‐
depleted cultures were incubated in situ and community composition was analyzed by 16S RNA gene sequencing. 454 pyrosequencing was used to identify species preferentially enriched in the absence of viruses, and clone libraries were constructed to examine evolution of the cultures over time. Methods Virus staining Viral abundance was visualized following a published protocol (Patel et al., 2007) adapted for smaller (11 mm) diameter Anodisc filters (0.02 μm pore size, Whatman). A rubber gasket with a 9 mm diameter hole was made for use as an adaptor for the 16 mm-­‐diameter filtration columns. After centering the Anodisc filter on the 25 mm-­‐diameter support filter, the gasket was placed on top so that the edges of the hole overlapped the Anodisc filter. The filtration column was then clamped to the pedestal, sandwiching the filters and gasket to create a watertight seal. Samples were pipetted directly over the Anodisc filter at a rate slow enough to prevent overflowing the gasket hole. Dried filters were stained with SYBR-­‐Green. A 1:10 dilution in virus-­‐free deionized water (prepared by passing through a 0.025 μm polycarbonate membrane filter) was prepared from the stock solution. Each Anodisc filter was placed on a droplet of 39 μL virus-­‐free water mixed with 1 μL dilute SYBR solution and incubated for 15 minutes in the dark. After blotting the backs of the filters, the filters were then dried in the dark for at least 1 hour. The dried filters were placed on ~5 μL droplets of Anti-­‐Fade on glass slides; an additional 5 μL droplet of Anti-­‐Fade on the coverslip prevented bubbles during mounting. In situ cultures A virus-­‐free community was prepared by sequentially filtering 50 mL of Eel Pond seawater through 0.2 μm and 0.025 μm polycarbonate membrane filters (Millipore), then vortexing the 0.2 μm filter in the resulting virus-­‐free filtrate. A control community (containing viruses) was prepared by vortexing the filter in the 0.2 μm filtrate. Dialysis membrane tubes (Spectrum FloatAlyzer, 100 kDa molecular weight cutoff) were filled with 10 mL aliquots of each community or with unfiltered Eel Pond seawater as an additional control. The tubes were first washed in ethanol and virus-­‐free deionized water according to manufacterer’s directions. Twelve dialysis tubes (3 samples, 4 replicates each) were placed in a wire basket and secured to the sides of the basket with fishing line. Two foam floatation blocks were secured to the outside of the basket with rubber bands and adjusted so that the dialysis tubes could hang vertically in the water. The module was tied to a private dock in Eel Pond. Samples were harvested over the course of two weeks. DNA extraction and sequencing DNA was extracted using the PowerBiofilm kit from MoBio. 10 mL of sample (entire contents of a single dialysis tube) were passed through a 0.2 μm polycarbonate membrane filter to concentrate cells. The filter was inserted into a bead-­‐beating tube (cells facing in) and bead-­‐beaten for 30 s. Extracted DNA was PCR-­‐amplified using 8F/1492R primers for the clone library and universal barcoded primers for the 454 pyrosequencing. Sequence analysis Clone library sequences were trimmed using Abifpy with a cutoff value of p = 0.001 and converted to FASTA format. Due to poor sequence quality, the day 3 virus-­‐free culture clone library sequences were merged from two sequencing runs. Duplicates were resolved by picking the longer of the two trimmed sequences. Both clone library and 454 sequencing data were analyzed using MacQIIME. For the 454 sequencing data, reads were classified using the Silva database trimmed of the primer sequences as a training set. OTUs enriched in the virus-­‐free cultures were identified by comparing the relative abundance of each OTU (normalized to reads per tag) in the virus-­‐free versus the control samples. Hits were called if the relative abundance was at least 5x. If the OTU was not present in the control, a hit was called if the virus-­‐free sample had more than 5 reads. For the clone libraries, OTU picking (97% similarity) and taxonomy assignments were done with the default parameters. Unweighted and weighted Unifrac distances were calculated using the jackknifed beta diversity pipeline, resampling 60 sequences per run for 10 runs. CARD-­‐FISH Culture samples were fixed in 2% paraformaldehyde (9 mL + 1mL 20% PFA) for 2 hours at 4°C. Fixed samples were diluted 1:10 in PBS and 10 mL of the diluted cultures were passed through 0.2 μm polycarbonate membrane filters. Filters were dried and stored at -­‐20°C until needed. CARD-­‐FISH was performed on eighths of thawed filters as described previously (Pernthaler et al., 2002) using the probes PSA184 (Pseudoaltermonas), G V (Vibrio), OCE232 (Oceanospirillum), ROS537 (Roseobacter), and NON338 (nonsense probe). Multiple images of each filter section were taken to quantify probed clade abundance. Exposure times for DAPI and Alexa-­‐594 images were held constant throughout imaging. The two channels were saved as separate TIFF files that were then input into a custom-­‐written MATLAB function to identify bacterial cells. Images were segmented by (1) background-­‐subtraction using the MATLAB function strel with a disc diameter of 5 pixels (2) thresholding using the MATLAB function im2bw, and (3) filtering out large (>500 pixels) and small (<5 pixels) objects. An additional step of removing objects with a mean intensity below a fixed threshold was used on the Alexa 594 images to decrease the false-­‐positive rate from autofluorescence. Results and Discussion In situ culturing of marine microbiomes Sequential filtration (Figure 1A) was used to create virus-­‐free cultures of Eel Pond seawater. An experimental vessel, the Caged Aquatic Retrievable Laboratory (CARL), was created to hold twelve dialysis tubes (Figure 1B) containing either the virus-­‐free culture, the control culture, or another control of unfiltered seawater in quadruplicate. The dialysis membrane pore size (100 kDa) was chosen to prevent environmental viruses from entering the tubes while still allowing for free exchange of nutrients. Samples of each culture condition were harvested from CARL 3, 6, 8 and 13 days following inoculation and deployment. Figure 1: Preparation of virus-­free and filtration control in situ cultures. (A): Schematic diagram of sequential filtration process. Unfiltered seawater in dialysis tubes served as an additional control. (B): Dialysis tube cultures in CARL. Tubes are kept upright with foam floatation rings supplied by the manufacturer. The two sequential filtration steps were sufficient to remove cells and viruses, respectively, from the seawater (Figure 2). Unfiltered seawater contains a diversity of DNA-­‐containing particles of varying shapes and sizes. The filtrate from passage through a 0.2 μm filter contained only small round particles of sub-­‐micron diameter, presumably viruses. Passage through a 0.025 μm filter removed those particles; however, the filter shown in Figure 2A was contaminated with a few rod-­‐shaped cells. These may have come from improperly cleaned filtration equipment, despite autoclaving the columns, pedestals, and filtrate collection flasks prior to filtration. Figure 2: Sequential seawater filtrates stained with SYBR-­Green on Anodisc 0.02 μm filters. (A): Unfiltered seawater. (B): Seawater passed through a 0.2 μm nitrocellulose filter. (C): Filtrate from (B) passed through a 0.025 μm nitrocelloluse filter. Viral counts in the cultures were only measured 13 days after inoculation. At this time point, all cultures showed similar high abundances of viruses (data not shown). There are several possible sources for the viruses in the supposedly virus-­‐free culture: (1) adhesion to cells or larger sediment particles that prevented removal during filtration, (2) prophage induction, and (3) passage from the environment due to breakdown of the dialysis membrane. Possible solutions to each of these problems include, respectively (1) washing and re-­‐filtering the trapped cells on the 0.2 μm membrane, (2) treating the mitomycin-­‐C or other DNA damage agents to induce lysis of prophage-­‐containing cells, and (3) having a control consisting of virus-­‐free water incubated in the dialysis membrane in situ. Some OTUs enriched in virus-­‐free culture 454 pyrosequencing of DNA collected 3 days after inoculation was used to compare the community structure of the virus-­‐free and control cultures (Figure 3). In all three culture, the dominant taxa were Alphaproteobacteria (undetermined class), Gammaproteobacteria (undetermined class), and chloroplasts. Alphaproteobacteria contain several clades of common marine bacteria, including SAR-­‐11 and Roseobacter. Gammaproteobacteria contain the common marine genera Oceanospirillum, Alteromonas and Vibrio. The presence of chloroplasts is unsurprising, since algae was visible growing outside and inside the culture tubes (data not shown). Figure 3: Class assignments of 16S DNA collected after 3 days of growth in situ and sequenced using 454 pyrosequencing. For taxonomic assignments with indeterminant class, the next highest determined level of taxonomic classification is displayed. Eight OTUs were identified that had a relatively high abundance in the virus-­‐free culture compared to the control (Figure 4). Three of these hits were identified at the species level. Several of the hits belong to the Gammaproteobacteria, although the phylum as a whole had decreased representation in the virus-­‐free sample versus the controls (Figure 3). Figure 4: OTUs enriched in the virus-­free culture. (A): Normalized log abundance of each OTU, rank-­
ordered by the control. (B): Taxon assignments and total abundance of eight hits circled in red in (A). Cultures diverge over time Clone libraries were constructed using 16S DNA amplified from samples collected 3 and 6 days after inoculation (Figure 5). Dominant families on day 3 across all three cultures included Rhodobacteraceae, Oceanospirillaceae, Bacillariophyta (diatom chloroplasts), and Pseudoalteromonadaceae. These families are consistent with the major phyla observed in the same DNA extractions using 454 sequencing. The most obvious change from day 3 to day 6 is the massive increase in Bacillariophyta clones across all three cultures, as well as the proportion of unclassified Bacteria. Many of the unclassified sequences are singletons and may be a consequence of poor sequence quality (insufficiently stringent trimming) rather than the presence of novel bacterial species. Figure 5: Family abundances from 16S DNA collected from in situ cultures and transformed into clone libraries. The Unifrac metric was used to compare the community composition of the three culture conditions over time (Figure 6). As can be inferred from the family abundance plots in Figure 5, the weighted Unifrac metric (Figure 6A) clusters more within the day 3 and day 6 timepoints than within the different culture conditions. In addition, the day 6 timepoint for the unfiltered culture diverged from the virus-­‐
free and the control culture. Given that the communities were tightly clustered on day 3, what could have driven this divergence? The unweighted Unifrac metric, which is a measure of richness, shows that the membership in the three culture conditions (Figure 6B) was not as tightly clustered as the abundance. It could be the presence of rare species that were removed during the filtration process that caused the divergence seen in the weighted Unifrac metric. What caused the overall shift in community composition between day 3 and day 6? The communities may have still been adjusting to the confined environment of the dialysis tubes and had not yet reached steady-­‐state at the time of sampling. Another consideration is changing environmental variables. There was significant growth of algae inside and outside of the dialysis tubes between these times (data not shown). This could have led to decreasing light penetrating into the tubes. Fluctuations in ambient water temperature may also have affected community composition; it would have been interesting to compare the incubated cultures to a sample of ambient seawater. Figure 6: Similarity of day 3 (blue) and day 6 (red) communities. (A): Weighted Unifrac metric, top two principle coordinates. (B): Unweighted Unifrac metric, top two principle coordinates. Sequencing may not reflect actual abundances Control and virus-­‐free cultures were fixed 8 and 13 days after inoculation and CARD-­‐FISH (Figure 7) was performed using probes against taxa thought to be abundant based on the clone library results (Figure 5). The DAPI staining revealed many large round cells (Figure 7A) in all cultures, especially at the later time point (data not shown). These cells were probably the Bacillariophyta identified in the clone libraries, suggesting that their population continued to increase from day 8 to day 13. Large cells were filtered out of the quantification of probe staining. The CARD-­‐FISH results for day 8 are not very consistent with the clone libraries for day 6. FISH showed Roseobacter constituting 47% of bacteria in the control and 23% of the virus-­‐free cultures (Figure 7C). In comparison, Rhodobacteraceae was the most common classified bacterial family in the virus-­‐free clone library, and was also present in the control clone library (Figure 5); however, the relative abundances are the reverse of the FISH results. Oceanospirillaceae was detected in the control clone library but there were no Oceanospirilla detected for that treatment using FISH, despite the technique’s greater sensitivity. FISH showed Pseudoaltermonas and Vibrio occurring at frequencies below the sensitivity limit of the clone libraries (~1% of total bacteria and chloroplasts). The inconsistencies between the clone libraries and the CARD-­‐FISH results could be due to the two days’ growth between the samples collected for each assay, but more likely reflect amplification biases that prevent accurate quantification of species using clone libraries. Figure 7: CARD-­FISH of virus-­free and control cultures. (A): Representative epifluorescence image of cells (control, day 8) stained with DAPI (blue) and probe (Pseudoaltermonas, red). (B): Segmentation and bacterial cell identification of image in (A). (C): Quantification of CARD-­FISH results. Data for each probe taken from 4-­8 imaging fields. (*): Day 13 data not available for Roseobacter and nonsense probe. Conclusion Bacterial abundances are a balance between growth and death. Growth is limited by nutrient and other resource availability. Viruses may contribute significantly to bacterial mortality in marine ecosystems. This study presents a preliminary investigation on the role of viruses in the control of bacterial species abundances. Virus-­‐free and control seawater cultures were incubated in situ. Community analysis was done using 454 pyrosequencing, clone library construction, and CARD-­‐FISH. Eight OTUs were identified that were preferentially enriched in the virus-­‐free culture. Analysis of two timepoints shows that the community composition of the in situ cultures diverged over time. However, these results are complicated by conflicting CARD-­‐FISH data and the observation of high viral abundance inside the virus-­‐free culture after 13 days of incubation. Due to time limitations, several experiments that would have enriched the project were not performed. This include: (1) tracking viral abundance in the three culture conditions over time, (2) measuring growth rates or nutrient turnover to confirm the hypothesis that relieving viral pressure decreases growth and turnover, and (3) adding nutrients to the cultures to compare bottom-­‐up versus top-­‐down limitations on bacterial abundance. Several key open questions remain to be explored: (1) How does the community’s metabolism change in response to the removal of viruses? (2) Does predation on bacteria by protists increase when viral-­‐induced mortality is relieved? (3) How does phage-­‐mediated lysis affect nutrient cycling? More research into these questions will be needed to fully understand the role of viruses in microbial ecology. Acknowledgements Thanks to all the staff for assistance, in particular Dan Buckley (initial planning), Steve Zinder (use of his dock), Sara Kleindienst (CARD-­‐FISH), Mallory Choudoir (virus staining) and Adam Zinder (CARL construction). Special thanks go to Chuck Pepe-­‐Ranney for invaluable discussions during all phases of the project, and to team TASTY and the rest of my classmates for making this a berry fun summer. My participation in the Microbial Diversity course was funded in part by the John & Elisabeth Buck Endowed Scholarship, the Gordon & Betty Moore Foundation, and the Milton L. Shifman Endowed Scholarship. References Bouvier, T., and del Giorgio, P.A. (2007). Key role of selective viral-­‐induced mortality in determining marine bacterial community composition. Environmental Microbiology 9, 287–297. Ferrera, I., Gasol, J.M., Sebastián, M., Hojerová, E., and Koblízek, M. (2011). Comparison of growth rates of aerobic anoxygenic phototrophic bacteria and other bacterioplankton groups in coastal Mediterranean waters. Applied and Environmental Microbiology 77, 7451–7458. Patel, A., Noble, R.T., Steele, J.A., Schwalbach, M.S., Hewson, I., and Fuhrman, J.A. (2007). Virus and prokaryote enumeration from planktonic aquatic environments by epifluorescence microscopy with SYBR Green I. Nat Protoc 2, 269–276. Pernthaler, A., Pernthaler, J., and Amann, R. (2002). Fluorescence in situ hybridization and catalyzed reporter deposition for the identification of marine bacteria. Applied and Environmental Microbiology 68, 3094–3101. Pernthaler, J. (2005). Predation on prokaryotes in the water column and its ecological implications. Nat Rev Micro 3, 537–546. Suttle, C.A. (2007). Marine viruses-­‐-­‐major players in the global ecosystem. Nat Rev Micro 5, 801–812. Clone Libraries •
Ylaine1A: Eel Pond water incubated in situ in a dialysis membrane tube (Spectrum FloatAlyzer 10 mL, 100 kDa MWCO) for 3 days. Ylaine2A: ‘Virus-­‐free’ Eel Pond water incubated in situ in a dialysis membrane tube (Spectrum FloatAlyzer 10 mL, 100 kDa MWCO) for 3 days. Seawater was sequentially filtered through 0.2 um and 0.025 um filters. The 0.2 um filter was vortexed in the final filtrate. •
Ylaine3A: ‘Virus-­‐free’ Eel Pond water incubated in situ in a dialysis membrane tube (Spectrum FloatAlyzer 10 mL, 100 kDa MWCO) for 3 days. •
Ylaine1B: Eel Pond water incubated in situ in a dialysis membrane tube (Spectrum FloatAlyzer 10 mL, 100 kDa MWCO) for 6 days. Seawater was filtered through a 0.2 um filter and the filter was vortexed in the filtrate. Ylaine2B: ‘Virus-­‐free’ Eel Pond water incubated in situ in a dialysis membrane tube (Spectrum FloatAlyzer 10 mL, 100 kDa MWCO) for 6 days. Seawater was sequentially filtered through 0.2 um and 0.025 um filters. The 0.2 um filter was vortexed in the final filtrate. Ylaine3B: ‘Virus-­‐free’ Eel Pond water incubated in situ in a dialysis membrane tube (Spectrum FloatAlyzer 10 mL, 100 kDa MWCO) for 6 days. Seawater was filtered through a 0.2 um filter and the filter was vortexed in the filtrate. •
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