Multi-species biofilm formation from seagrass-associated bacterial communities Melissa Kardish PhD Candidate, Population Biology Graduate Group, University of California, Davis Microbial Diversity Course 2017 Submitted: August 22, 2017 Abstract While broad successional patterns are widely studied in ecological communities, much less is known about the fine-scale interactions that drive their development. To elucidate these patterns, I studied the quick growing members of bacterial biofilms associated with seagrass. By isolating fast growing members and then assessing biofilm production via a crystal violet assay, I identified species combinations that produced expected, greater than expected, and less than expected sized biofilms based on their individual biofilm production. These interactions are likely important in marine environments where forming a biofilm may give better or more consistent access to resources. Further experiments will investigate the microscale arrangement and potential mechanisms for inhibitory interactions or facilitations between organisms. Introduction The species composition and spatial structure of communities throughout ecological succession is fundamental to determining the dynamics of a community. While much is known about the different stages succession and the associated compositional and biogeochemical changes, teasing apart key players and interactions in these systems and broader ecological patterns is less well understood. However, knowing different paths or speeds of ecological succession based on different players could be key to understanding long term diversity patterns, species composition, and stability in these communities.

Bacterial biofilms are an excellent model for thinking about how communities assemble and structure themselves.1 They are tractable systems that assemble on a surface and can be measured at a given location2 Single species biofilms have been studied at length in for negative medicinal implications3 as well as aquatic environments to control growth in wastewater.4

Study of biofilms in the ocean have primarily looked at broad community composition during succession and cannot detect finer scale species interactions that may drive how these communities succeed or fail.5,6 To detect these interactions, one would have to look at species-level interactions and scale up to get total community function working from the microcosms up to the field where 16S type data could elucidate how changes in these smaller settings play out in slightly longer succession.

Seagrasses are no exception to being covered in biofilms. Seagrasses accumulate bacterial,7 fungal, and algal communities on their leaf surfaces.8 Seagrass microbial communities are correlated with decreased disease.9 This community structure, driven by biofilm assembly, may play a role in larger ecosystem and/or plant health. Understanding the early establishment of biofilms on these surfaces could help describe effects on ecosystem-level health.

Here, I investigate the properties of polymicrobial biofilms from communities associated with seagrasses. I selected fast growing members of these communities, as well as a few bacteria isolated from adjacent seawater, to compare biofilm formation and potential assembly mechanisms driving early colonization of unoccupied surfaces. I hypothesize that different initial communities will lead to different speeds of biofilm formation – which might result in increased

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fitness for an organism or increase potential for nutrient acquisition when its attached to a surface. Additionally, I hypothesize that there will be identifiable key players in these early biofilms that can drastically alter their stability, speed of formation, and overall composition. Methods Sample Collection Initial samples were collected from Stony Beach (41.529˚ E, 70.674˚ N) at 8 a.m. on July 26, 2017 during the low tide. Water was collected from the center of the beach and on the east end next to the jetty. One shoot of Zostera marina (eelgrass) was collected as well. All samples were stored in seawater, and plated later that day.

Later eelgrass samples were collected with the help of the MRC from Devil’s beach (41.522˚E, 70.681˚N). At 1:45pm on August 7, 2017, several shoots were collected along with seawater during a low tide and cloudy weather. These shoots were transported back stored in seawater, transported back to the lab, and used for FISH imaging. Cultures Samples collected from seawater were cultured through a dilution series on seawater complete (SWC) plates (Appendix 1). Samples from eelgrass were prepared by either directly plating the leaf on the plate surface then streaking isolated from the lawn that grew u- or by vortexing a leaf segment vigorously for 30 seconds in SWC and then plating the solution in a dilution series.

All selected colonies were streaked for isolation for further assays. Liquid cultures of samples were frozen in 25% glycerol stocks and stored at -80˚C. I extracted DNA for 16S sequencing by taking a small sample of a colony and boiling it for 15min in 20 µl ALP (alkaline PEG200). I then did a 50 µl PCR with 0.2 µl 10 mM 8F and 1391R primers. Samples were then sequenced with the 16S 1200R primer. Sequences were classified to genus using the RDP Classifier (https://rdp.cme.msu.edu/classifier/classifier.jsp). Biofilm Assays Many combinations of biofilms were assessed in this study. To determine which species to study further, an initial assay measured the biofilm production of biofilms measured biofilms for 7 timepoints (0, 2.75, 4, 5.25, 6.5, 7.75, and 9 hours) which allowed for determination of the optimal window to study these fast growing biofilms in. All other plates assessing biofilm formation were incubated with time points taken at 2.5, 4, 5.5, and 7 hours after inoculation.

These other plates consisted of pairwise, three bacteria, four bacteria, and five bacteria combinations. Combinations tested are listed in Table 2. During each test, individual bacterial strains were tested as a control for consistency of biofilm growth. Two sets of 5 bacteria isolated were used for these combinatorics tests, with one bacteria, L, present in both tests.

The biofilm assay itself consisted of placing a 1:100 dilution of an overnight culture in a well of a 96 well plate. Plates were placed in a humidified chamber at 30˚C until the time when biofilm production was measured. At that time, all liquid media/culture is first shaken, then rinsed with DI water out of each plate. Then 125 µl of 0.1% crystal violet solution was added to each well. After staining for 10 minutes, wells were washed 4 times with DI water. All remaining liquid was removed and let plates were let dry 10,11.

One to two days later, after completely drying, biofilms were solubilized by adding 125µl of 30% acetic acid. After 10 minutes, wells were mixed and transferred to optically clear plates.

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On a Promega GloMaxDiscover, I measured absorbance at 560nm to assess amount of crystal violet as a proxy of total biofilm formation.

Expected biofilm production was calculated based on the fraction of species present in the biofilm assay initially.12 In the combinations of five bacteria, each isolate was 1/5 of the initial community, I expect that one fifth of its biofilm production alone will be in the final community. The same fraction method was used with other combinations (1/4, 1/3, 1/2). Each plate had internal controls of isolated species individually to control for experimental error/exceptions. Mono-FISH staining A series of FISH staining protocols were used to image the natural surfaces of seagrass leaves. Fixation of leaves began within an hour of collection of plant. Leaves were cut into small segments and sorted by approximate age. They were then incubated at room temperature with 2% formamide for 1 hour. Afterwards, samples were rinsed in 1xPBS three times before storing in 1xPBS at 4˚C. Various probes were attempted and used, but here I am only presenting data on the all bacteria probe. To hybridize the probe, I prepared 10 µl hybridization buffer with 9 µl working solution and 1 µl EUB probe (EUB338I-III). A small sample of seagrass leaf was dipped in the hybridization buffer and then incubated for 2 hours in a 35% formamide humidified chamber at 46˚C. Leaf segments were then washed, first for 10 minutes in washing buffer, then 5 minutes in PBS, then MilliQ water before drying. Once dry, leaves were stained with DAPI for 10 minutes before two one-minute rinses in PBS. Leaf surfaces were dried at room temperature again before being dipped into a small amount of 4:1 mix of Citifluor:Vectashield in a petri dish with coverslip on the bottom. A 1% agarose pad was placed on top to keep the sample against the surface of the coverslip for imaging. Microscopy and Microscopy Analysis FISH samples were viewed on an Axio1 (Zeiss) and on a confocal laser scanning microscope (LSM 800; Zeiss). All subsequent image processing enhanced contrast in images to allow for better comparisons between fields was performed in ImageJ/Fiji. Results FISH visualization While there were high levels of autofluorescence from the leaf itself as well as from algae growing on its surface, there was overlap between EUB probe and DAPI stain that indicated presence of bacteria and similar levels of fluorescence were not observed in untreated slides (Fig 1 top left). Additionally, there was an increase in autofluorescent algae with increased age. While harder to identify bacteria with the autofluorescence, there are signs of bacteria wrapping around other organisms on the surface of eelgrass (Fig 1 top right). Relative levels of epiphytic biofilm cover are pictured in Fig 1 lower panels. Colonies isolated A total of 25 colonies were isolated successfully from seawater or leaf surfaces. These included 17 Vibrio isolates, three Pseudoaltermonas, two Tenacibaculum, one Cobetia, and one Shewanella (Table 1). Formation of biofilms

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All isolated colonies formed biofilms in crystal violet test plates (Fig. 2). These biofilms peaked at different rates. Colonies that formed some, but consistent and not overwhelming amounts of biofilms were selected for further studies (Fig. 3). This included one Cobetia, one Tenacibaculum, one Shewanella, one Pseudoaltermonas, and 5 Vibrio. Based on this information, while I took multiple timepoints of all biofilms as internal controls moving forward, biofilms presented in the later figures are all at approximately 5.5 hours in (equivalent to T3 in Fig. 3). Pairwise Comparisons I first tested pairwise comparisons (Figure 4). Here there are many combinations (e.g., bacteria V and bacteria L) that had exactly or close to the expected biofilm production based on their biofilm production in monocultures. A few combinations greatly exceeded expected biofilm production (bacteria D and L produced a biofilm that was nearly four times the expected biofilm production). Finally, a few combinations seemed inhibitory to biofilm production. Bacteria N grows very little biofilm, while bacteria V grows a substantial amount. However, when they were co-cultured there was almost no biofilm production. Other bacteria cultured with bacteria N grew the expected biofilm (with L or Ea) or more than the expected biofilm (with Eb). Mixed cultures Combining sets of these communities resulted in stronger biofilm formation than was expected based on the expected proportions contributed in both sets tested and as high or higher than the greatest biofilm produced by any individual isolate (Fig 5). Dropout experiments From each of the sets of five bacteria, I also tested combinations of three (Fig. 6) and four (Fig. 7) bacteria to see changes in biofilm formation. Interestingly, both combinations of four bacteria grew near-equivalent biofilms to the 5 isolate combos, except when they were each missing a specific player. In the EaEbLNV combination, if bacteria Eb (a Vibrio that formed substantial biofilms) was missing, the total biofilm in the set of four was less than the expected, while all other combinations of four isolates produced more. In the DJKLO combination, if bacteria D (a Vibrio that does not form substantial biofilms) was missing, the total biofilm was less than the expected biofilm production rather than more as was seen in all other combinations of four. Results from additional experiments can be found in Appendix B. In Appendix C, I include a discussion of day-to-day variation in the biofilms of these organisms. Biofilm production by the same organisms on different days was largely statistically identical with some notable exceptions that do not affect the results described above. Discussion Understanding the polymicrobial-interactions that govern biofilm assembly and production can yield insight into the dynamics of their communities. Studying a system such as this that examines quickly growing bacteria on a substrate, provides an opportunity to study the early successional dynamics.

The FISH images shown, along with literature,13 suggest that bacteria are among the initial founding members of populations and might play roles in the establishment of later

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colonizing communities as they establish quite quickly. This supported these attempts to identify early interactions as bacteria seem important to understand later successional phenomenon.

These combinations themselves, even in lab biofilm assays, showed interesting cross-species interactions. A few of these combinations may specifically be worth following up in further depth because of evidence of interesting cross genera interactions. When grown together, bacteria L (a Pseudoaltermonas) and bacteria D (a Vibrio) produce a much greater than expected biofilm. However, when they are co-cultured with bacteria J (a Tenacibaculum), they produced the expected biofilm, one similar to bacteria J’s biofilm production alone. However, if one looks at the J’s biofilm production with L it is lower than J’s. Adding D recovers biofilm production to J’s original levels. Additionally, there may be a mechanism of suppression from bacteria N (a Vibrio) on bacteria V (a Cobetia). However, more work would have to be done to elucidate the mechanism of each of these interactions. One potential approach would be to visualize biofilm structure under a microscope and examine cell density, closeness to other cell morphologies, etc. This approach would be especially possible if a GFP plasmid was inserted into one of the strains which would allow easy identification of bacterial type in biofilms (attempted in Appendix B).

In the experiments where one of the five isolates was dropped, it is interesting that two different Vibrio isolates with two different morphologies produced the same reduction in community biofilm when dropped out. They may have a shared mechanism that enhances community biofilm production as they are both Vibrio; however, there may also be an unconserved mechanism indicating that Vibrio strains have multiple mechanisms by which they can enhance biofilm production. Teasing apart which of these possibilities is true would require further experiments including mutagenesis or potentially genomic comparison.

Further follow-up to this experiment in an ecological context would require extending assessment of biofilm formation to different environments. As these samples were isolated from a seagrass leaf in a tidal zone, testing the differences in different environments with higher or lower flow during biofilm formation would be relevant for attachment.

Additionally, I have isolated lytic phages from many of these hosts (Appendix B.1). Introducing these viruses during biofilm production could identify broad and narrow host ranges

Ultimately, based on experiments like these, determination of patterns of later stage assembly could be determined, potentially by forming these initial biofilms and then exposing them to natural communities to see the speeds of change in structure and biofilm formation of these communities. Conclusions There are clearly beneficial and antagonistic interactions with respect to biofilm formation. This biofilm formation is likely important in a marine environment as it would allow bacteria to anchor themselves in a desirable, potentially high nutrient environment. Through closely studying the interactions in a laboratory setting we can get a better understanding of the natural interactions of microbial communities in complex environments. Acknowledgements I would like to thank the Microbial Diversity Course. Special thanks to Kyle Costa, Dianne Newman, George O’Toole for a ton of support during this project. Much thanks also to Viola Krukenberg and Georgia Squyres for their support with the FISH imaging, as well as Jessica Mark Welch for discussions on approaches for seagrass imaging. Thanks also to the other faculty and TAs that I would not have been able to do this project, or learn nearly as much from the

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course, without.

Funding for me has come from the NSF, UC Davis, and the S.O. Mast Fellowship. This work would not have been possible without their support. Thanks also to the support of family, friends, and the Stachowicz lab for continued support as I dropped off the face of the earth to dive headfirst into microbiology. Finally, many, many thanks to Lowell West for logistical support, sanity checks, and copy editing. References 1. Cordero, O. X. & Datta, M. S. Microbial interactions and community assembly at

microscales. Curr. Opin. Microbiol. 31, 227–234 (2016). 2. Donlan, R. M. Biofilms : Microbial Life on Surfaces. 8, 881–890 (2002). 3. Percival, S. L., Suleman, L., Vuotto, C., Donelli, G. & Percival, S. L. Healthcare-

associated infections , medical devices and biofilms : risk , tolerance and control. 323–334 (2017). doi:10.1099/jmm.0.000032

4. Romaní, A. M., Guasch, H. & Balaguer, M. D. Aquatic Biofilms. (2016). 5. Dang, H. & Lovell, C. R. Bacterial Primary Colonization and Early Succession on

Surfaces in Marine Waters as Determined by Amplified rRNA Gene Restriction Analysis and Sequence Analysis of 16S rRNA Genes. 66, 467–475 (2000).

6. Sathe, P. & Dobretsov, S. in Aquatic Biofilms 47–62 (2016). 7. Marhaeni, B., Radjasa, O. K., Bengen, D. G. & dan Richardus F. Kaswadji. SCREENING

OF BACTERIAL SYMBIONTS OF SEAGRASS Enhalus sp. AGAINST BIOFILM-FORMING BACTERIA. 13, 126–132 (2010).

8. Borowitzka, M. A., Lavery, P. S. & van Keulen, M. in Seagrasses: Biology, Ecology, and Conservation 441–461 (2007).

9. Lamb, J. B., Water, J. A. J. M. Van De, Bourne, D. G. & Altier, C. No Title. 1–4 10. O’Toole, G. A. Microtiter Dish Biofilm Formation Assay. 10–11 (2011).

doi:10.3791/2437 11. O’Toole, G. A. et al. in METHODS IN ENZYMOLOGY 310, 91–109 (1999). 12. Traverse, C. C., Mayo-smith, L. M., Poltak, S. R. & Cooper, V. S. Tangled bank of

experimentally evolved Burkholderia bio fi lms re fl ects selection during chronic infections. 110, (2012).

13. Kurilenko, V. V, Ivanova, E. P. & Mikhailov, V. V. Zonal Distribution of Epiphytic Microorganisms on the Eelgrass Zostera marina. 70, 427–428 (2001).

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FiguresFigure 1. At left is an image of a young seagrass l. Both images are stained with DAPI (blue) and hybridized with a EUB FISH probe (green). As leaves age, the surface community accumulates a variety of micro and macro epiphytes. Below are larger images of seagrass displaying relative epiphyte covers.

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Figure 2. Biofilm growth was assessed with a crystal violet assay as described in the methods. This resulted in staining where biofilms occurred in the well. This mostly occurred at the air/liquid interface, but also was present elsewhere. For example, Eb is a Vibrio that grows best at the air liquid interface, but also elsewhere throughout the well.

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Figure 3. The biofilm production over time of all bacteria tested in these studies. Those in black are labeled with their code for Table 1 and were used in later experiments. Most bacteria eventually grew some amount of biofilm.

Figure 4. The pairwise comparisons of all biofilms studied here. Two sets of five initial strains were used. Bacteria L was used in both studies. The expected biofilm production is calculated based on the proportion of each bacteria in the mixed culture and the biofilm production each organism made on its own. Some combinations produced the expected amount of biofilm, some produced more and some produced less.

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Figure 5. When all five species were combined they built larger than expected biofilms. Expected biofilm production is calculated the same as previously.

Figure 6. Combinations of three different isolates. Expected values are the same as described in other figures. Note: not all combinations were tested.

Figure 7. All combinations of four different isolates.

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Tables Table 1. Characteristics of species isolated and used in experiments. Rows in dark grey were used in later experiments. Rows in light grey are simply for aesthetic distinction.

Sample Code

Inoculation source

Colony morphology

RDP classification

Picture

Aa Stony beach, east; leaf on plate

Small clear colonies, uneven round edges

Vibrio (100%)

Ab Stony

beach, east; leaf on plate

Small, slightly yellow colonies

Vibrio (100%)

Ba Stony

beach, east; leaf on plate

Large cream colonies, smells

Vibrio (100%)

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Bb Stony beach, east; leaf on plate

Small white colonies with smooth edges

Vibrio (56%)

C Stony

beach, central, seawater 1:100 dilution plate

Small round white colonies, isolated enveloping D

Vibrio (100%)

D Stony

beach, central, seawater 1:100 dilution plate

Small almost clear colonies, isolated being enveloped by C

Vibrio (100%)

Ea Stony

beach, east; leaf on plate, center of inoculation

Small, white, glistening, translucent colonies

Shewanella (100%)

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Eb Stony beach, east; leaf on plate, center of inoculation

Large flat white colonies, fast growing

Vibrio (100%)

F Stony

beach, east; leaf vortexed in SWC, 1:100 dilution

Hard to isolate, foggy edges, opaque center, white

Vibrio (100%)

G Stony

beach, east; leaf vortexed in SWC, 1:100 dilution

Hard to isolate, white foggy edges, thick center

Vibrio (100%)

H Stony

beach, east; leaf vortexed in SWC, 1:100 dilution

Orange small round colonies with smooth edges

Pseudoaltermonas (100%)

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I Stony beach, east; leaf vortexed in SWC, 1:1000 dilution

Small, round white/clear colonies

Vibrio (100%)

J Stony

beach, east; leaf vortexed in SWC, 1:1000 dilution

Iridescent, edges slightly unclear

Tenacibaculum (100%) *for additional information on this strain, please see Nadia Herrera’s report

K Stony

beach, east; leaf vortexed in SWC, 1:100 dilution

Small, clear colonies with foggy edges, isolated next to L

Vibrio (100%)

L Stony

beach, east; leaf vortexed in SWC, 1:100 dilution

White, medium sized round colonies, isolated next to K

Pseudoaltermonas (100%)

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M Stony beach, east; leaf vortexed in SWC, 1:100 dilution

White, completely opaque, small, very round; isolated with N and O

No high quality sequence

N Stony

beach, east; leaf vortexed in SWC, 1:100 dilution

Small, white, round colonies; isolated with M and O

Vibrio (100%)

O Stony

beach, east; leaf vortexed in SWC, 1:100 dilution

Small, white, slightly translucent colonies that spread out where streaked; isolated with N and M

Vibrio (100%)

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P Stony beach, east; leaf vortexed in SWC, 1:1000 dilution

Small white, mostly opaque colonies

Pseudoaltermonas (100%)

Q Stony

beach, east; leaf vortexed in SWC, 1:1000 dilution

Very very small white colonies

Vibrio (100%)

N/A

R Stony beach, central, seawater 1:10 dilution plate

Large glistening hard to isolate colony, seems to spread out where streaked, foggy edges

Vibrio (100%)

S Stony

beach, east; seawater 100ul on plate

Iridescent, small colonies with unclear edges; grow quickly along streak lines

Tenacibaculum (100%)

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T Stony beach, east; seawater 100ul on plate

Small, but not tiny, round, white colonies

Vibrio (100%)

U Stony

beach, east; seawater 100ul on plate

Larger white flat colonies, some expanding along streak line

Vibrio (100%)

V Stony

beach, east; seawater 1:10 dilution plate

Small, white, evenly-colored colonies with smooth edges

Cobetia (100%)

Appendices . Appendix A Media for the main experiments described above was all done on seawater complete (SWC) plates or liquid media. The recipe for 1 liter follows.

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1 L seawater base NaCl 20g MgCl26H2O 3 g CaCl26H2O 0.15 g

KCl0.5 g 5g tryptone 1g yeast, 3ml glycerol, 5 ml 1M MOPS, pH 7.2

1.05g 0.25 ml 5M NaOH add H2O to 5ml total volume after adjusting pH to 7.2 with HCL or NaOH

15g agar (for plates) Appendix B : Supplemental Experiments Two additional experiments were begun to isolate viruses and to visualize the biofilms made. B.1 Isolation of viruses In collaboration with Fatima Hussain, I isolated lytic phages associated with 7 of the strains used in these later experiments using incubated, 0.2um filtered seawater with SWC and the 9 isolates used in the majority of experiments above. These were streaked once for isolation and stored after being spun down and filtered with a 0.2um filtered. B.2 Failed experiments attempting to visualize a biofilm With the interesting interactions between bacteria L, D, and J described above in biofilm formation, I hoped to visualize some of the interactions above. This unfortunately resulted in failed, but attempted experiments, as I attempted to electroporation competent cells of each strain with a plasmid (PGB5; DKN #27) containing Tet and Amp resistance using low concentrations of tetracycline. Unfortunately, I did not have time to optimize this protocol and was unable to get a GFP plasmid in which would have enabled clearer distinction of different strains of bacteria. Appendix C In addition to the differences described above I examined differences between days in these biofilm cultures to test that these were done. Not all of these were taken at the exact same time after starting inoculation, but all were taken approximately 5.5 hours after the experiment began. Most differences are not significant (comparing within groups using ANOVA and Tukey’s post-hoc test). Surprisingly, however, bacteria J has dramatic differences in its ability to form biofilms which varied across days and tests (p-values < 0.01). This suggests there are some initial conditions necessary for J that may have been met on some days and not others. Alternatively this may indicate variation in density of overnight cultures. See the 2017 Microbial Diversity course reports of E. Perry and N. Herrera for further discussion about differences in timing of Tenacibaculum colonies in development.

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