In the co-cultures, we calculated B12 cell quotas based on pB12 and cell abundances for L. polyedrum ranging from 5 × 10−18 to 2 × 10−17 mol cell−1 (**Table 2**). These cell quotas are comparable to 4 × 10−22 and 1.7 × 10−18, the values reported for dinoflagellates by Tang et al. (2010; **Table 2**). Our measured cell quotas show between one and two orders higher cellular B12 content for L. polyedrum than for both types of D. shibae phenotypes, attached and free-living. In our results, L. polyedrum had lower Ks but higher cell quotas compared with other dinoflagellate species (Tang et al., 2010); the former might provide an ecological advantage to L. polyedrum or partially compensate for the disadvantage of a high cell quota. The cell quota concentrations based on cell volumes are 2–3 orders of magnitude lower for L. polyedrum cells than for D. shibae cells (**Table 1**). Panzeca et al. (2009) quantified the concentration of B12 during a L. polyedrum bloom in a coastal, ranging from 2 to 61 pM, concentrations that should not be limiting for L. polyedrum. L. polyedrum abundance was not quantified at the time of B12 measurements, but at nearby locations at a later date the abundance was shown to be high, ranging from 1 × 106 to 6 × 106 (cell L−1 ) (Peña-Manjarrez et al., 2009).

According to Croft et al. (2005), a minimum of 10 ng L−1 of B12 is required for growth in culture by most phytoplankton species, while Tang et al. (2010) showed that HAB species need higher concentrations than non-bloom forming species. Most of the B12 quantification in the field or in culture studies measured the dissolved form, and only few studies related to dinoflagellates in the field (Carlucci, 1970; Gobler et al., 2007; Panzeca et al., 2009) or in cultures (Croft et al., 2005; Tang et al., 2010; Cruz-López and Maske, 2016). Recently Suffridge et al. (2017), quantified the particulate fraction for several B-type vitamins in the field and cultured marine bacteria, given previous cell quotas for marine phytoplankton or heterotrophic bacteria are based on uptake rates (reviewed in Droop, 2003; Bonnet et al., 2010).

To quantify the B12 in the different fractions, we used a highly sensitive enzyme-linked immunosorbent assay (ELISA) able to recognize all B12 forms (cyanocobalamin, methylcobalamin, hydroxocobalamin, and coenzyme B12), but unable to distinguish them individually. The cyanocobalamin produced by D. shibae can be used directly by the algal auxotroph without post-modifications (Helliwell et al., 2016). A likely explanation for our co-culture experimental results is that

L. polyedrum acquired B1 and B12 vitamins from D. shibae in sufficient quantity to sustain the maximum growth over many generations; however, it had not been shown what D. shibae receives in return from L. polyedrum. For D. shibae to grow together with L. polyedrum, it would need organic substrate for biomass formation but in addition also specific compounds because D. shibae is auxotroph for B7, niacin and 4-aminobenzoic acid (Biebl et al., 2005). The production and release of niacin and 4-amino-benzoic acid by a dinoflagellate host was previously suggested by Wagner-Döbler et al. (2010) and Wang et al. (2014a) to explain their co-cultures of Prorocentrum minimum CCMP1329 with D. shibae in medium lacking B12 or any of the other essential vitamins mentioned above. However, the dinoflagellate P. minimum CCMP1329 is B7 auxotroph (Wagner-Döbler et al., 2010) and therefore B7 had to be supplied to the medium. In the case of L. polyedrum, B7 is not required (Cruz-López and Maske, 2016) and the culture medium was not supplemented with B7, niacin or 4-amino-benzoic acid. We assume that these organic growth factors could not be provided by the prepared seawater medium. We suggest that these three compounds were provided by L. polyedrum in the form of exudates in sufficient quantity to support the growth of the D. shibae population. In return, L. polyedrum thus promoted the production of B1 and B12. Due to technical limitations, we only measured B7 in the particulate fraction, but not in the dissolved fraction. Future research should quantify B7, niacin or

4-amino-benzoic acid in the dissolved fraction to better define the mutualistic relationship between D. shibae and L. polyedrum (**Figure 4**).

On the other hand, the release of B7 from different phytoplankton species has been documented previously (Bednar and Holm-Hansen, 1964; Carlucci and Bowes, 1970a,b; Aaronson et al., 1971, 1977), cell quotas for dinoflagellates are based on uptake rates (Tang et al., 2010). Our pB7 results showed no pB7 in the L. polyedrum axenic culture, whereas in co-culture with D. shibae, L. polyedrum we found from 4.74 × 10−19 to 9.41 × 10−19 mol cell−1 . Adjusting to pmol cell−1 , our results are in the range (4.74 × 10−7 to 9.41 × 10−7 ) for the cell quotas reported for Gymnodinium instriatum L3, and two orders of magnitude higher than those reported for P. minimum CCMP696 and P. minimum PB3 (Tang et al., 2010).

For marine heterotrophic bacteria, the B7 production has been inferred, based on their available genomes, or in situ transcriptional studies (Luo and Moran, 2014; Sañudo-Wilhelmy et al., 2014; Gómez-Consarnau et al., 2018). Among the Roseobacter clade, of the 52 Roseobacter genomes analyzed by Luo and Moran (2014), 30 species were found to be auxotrophs for B7, including D. shibae (Biebl et al., 2005; Wienhausen et al., 2017; Gómez-Consarnau et al., 2018). Of these 30 Roseobacter species, 13 produce B1 and B12, and 17 only B12 (**Table 4**). In the case of D. shibae in co-culture with L. polyedrum, we could only detect B7 in the attached D. shibae phenotype, with cell quotas ranging from 1.72 × 10−7 to 1.9 × 10−7 pmol cell−1 .

In a recent study, Suffridge et al. (2017), developed a method to quantify B-type vitamins in the dissolved and particulate fraction using cultures of marine bacteria. They obtained cell quotas of B7 for Synechococcus strain CC9311 (9.9 × 10−6 ± 8.1 × 10−6 mol cell−1 ), and for Vibrio AND4 (2.47 × 10−6 ± 1.36 × 10−6 mol cell−1 ). In our cultures, surprisingly the free-living TABLE 4 | Select genes for B-vitamin biosynthesis in 30 B7 auxotroph Roseobacter isolate genomes.

Presence √ , ; absence X, .

B7, biotin synthase; B1, thiamine synthase; B12, cobalamin synthase (extracted from Luo and Moran, 2014). Isolation source extracted from www.roseobase.org and https://genome. jgi.doe.gov. n. a., Not available.

D. shibae did not contain pB7. As was shown by Luo and Moran (2014), the B7 auxotrophy is a widespread phenomenon among the Roseobacter clade, although this auxotrophy is not always related to a symbiotic lifestyle with phytoplankton, which needs to be clarified with further culture and genomic studies.

### D. shibae Frequency of Attachment on L. polyedrum

Eukaryotic phytoplankton and heterotrophic bacteria are interacting to fulfill the need for vitamins of the former and the need of excreted organic substrate of the latter. Different modes of exchange of vitamins and organic substrate are possible; vitamin-producing bacterial epibionts can be attached to the vitamin-auxotroph cells, or the vitamin-producing and consuming cells can be freely suspended, and the exchange of the dissolved compounds is maintained by turbulent and molecular diffusion at sufficiently high rates to supply the metabolic demand of the auxotrophs (Croft et al., 2005; Droop, 2007).

We used vitamin-replete and vitamin-limited cultures of L. polyedrum to produce contrasting host-bacterial culture conditions assuming that the V-L condition would promote the cellular attachment of bacteria and host. The frequency of D. shibae attached to L. polyedrum was higher under V-L condition compared to V-R condition (**Figure 2**; **Table S3**; Mann-Whitney, p < 0.005). When V-L cultures returned to V-R condition, the frequency of attachment reduced to average abundance. Higher attachment frequency under V-L condition suggests a more intense exchange of metabolites under growth culture conditions where this exchange would be essential to the growth of either population.

Because no carbon substrates were added to the culture medium that could have supported the growth of D. shibae, our results suggest that D. shibae was able to use dinoflagellate photosynthates as a carbon source in return for supplying the host with vitamin B1 and B12. As pointed out above L. polyedrum might have provided more specific growth enhancing substrates to D. shibae. Although in media without added vitamin D. shibae was found to be attached more frequently to L. polyedrum cells suggesting a synergy between both species, it leaves the question open about the mechanism of delivery. Vitamin could have either been exchanged in the dissolved form originating from freeliving partners or by a more direct exchange of metabolites when both cell types are physically attached. Interestingly, in similar experiments but with a natural community of microbes instead of D. shibae, the frequency of attachment of microbes to L. polyedrum did not increase but the abundance of freeliving bacteria in the co-culture increased (Cruz-López and Maske, 2016). For co-cultures of L. polyedrum with bacterial communities, the vitamin exchange and balance is more difficult to analyze because an unknown part of the bacterial community is also B12 auxotroph.

During the co-culture of L. polyedrum and D. shibae, we quantified the free-living and attached D. shibae cells in V-R and V-L cultures, and observed that the abundance of free-living D. shibae in the V-R culture was lower than in V-L culture (**Figure 3B**; ANOVA, p > 0.05), and that L. polyedrum in the V-R culture was lightly colonized in comparison with the V-L culture (**Figure 2**; **Table S3**; Mann-Whitney, p < 0.005). These two alternative lifestyles of D. shibae have been documented before (Biebl et al., 2005; Wagner-Döbler et al., 2010; Wang et al., 2014a), and are controlled by quorum sensing (QS) via CtrA signal transduction protein (Patzel et al., 2013; Wang et al., 2014b). Note, however, that increased bacterial attachment frequency under vitamin limitation coincided with an increased abundance of free-living bacteria, further work will be required to confirm that this phenotype is driven by nutrient exchange requirements.

### CONCLUSION

Our data show that the B1 and B12 limiting concentrations for L. polyedrum growth are lower than typical concentrations found in coastal waters. In vitamin-depleted cultures of L. polyedrum, the limitation could be overcome by co-culture with D. shibae,

### REFERENCES

because the bacterium was likely able to provide sufficient quantities of B1 and B12 to support the growth of L. polyedrum. In return, D. shibae was likely provided sufficient exudates by L. polyedrum to support its growth. Also, the mutualism between both partners may have extended beyond the supply of B1 and B12 and dissolved organic carbon. Since D. shibae is auxotrophic for B7, niacin and 4-amino-benzoic acid, L. polyedrum needed to provide sufficient quantities of these compounds to overcome D. shibae's auxotrophy. This mutualism was probably helped by the phenotypic ability of D. shibae to choose between a free-living or attached lifestyle, where the attached lifestyle could serve as a reservoir of B7 within this isogenic population.

### AUTHOR CONTRIBUTIONS

RC-L conceived, performed, and wrote the article. HM conceived and wrote the article. KY and NH performed and wrote the article.

### ACKNOWLEDGMENTS

This work was partially supported by the National Council of Science and Technology (CONACyT-México) project CB-2008-01 106003 (to HM), a Ph.D. scholarship, and a Postdoctoral fellowship for overseas to RCL (CONACyT-México). Additionally, we would like to thank Avery O. Tatters (University of Southern California) for kindly providing the L. polyedrum strain and Prof. Carl J. Carrano (San Diego State University) for allowing us to use his laboratory to prepare samples. We would like to show our gratitude to Sharon and Christopher A. Rhodes, the owners of Drug Delivery Experts LLC., for allowing us to use their facility to generate LC-MS data. Finally, we would like to thank the reviewers for constructive comments and suggestions.

### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fmars. 2018.00274/full#supplementary-material

variants of vitamin B12. Curr. Biol. 26, 999–1008. doi: 10.1016/j.cub.2016. 02.041

and cell division modes in Dinoroseobacter shibae. ISME J. 7, 2274–2286. doi: 10.1038/ismej.2013.107

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Cruz-López, Maske, Yarimizu and Holland. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Iron and Harmful Algae Blooms: Potential Algal-Bacterial Mutualism Between *Lingulodinium polyedrum* and *Marinobacter algicola*

### Kyoko Yarimizu1 , Ricardo Cruz-López 1,2 and Carl J. Carrano1 \*

1 Department of Chemistry and Biochemistry, San Diego State University, San Diego, CA, United States, 2 Department of Biological Oceanography, Centro de Investigación Científica y de Educación Superior de Ensenada, Baja California, Mexico

Phytoplankton blooms can cause acute effects on marine ecosystems either due to their production of endogenous toxins or due to their enormous biomass leading to major impacts on local economies and public health. Despite years of effort, the causes of harmful algal blooms (HAB) are still not fully understood. Our hypothesis is that bacteria that produce photoactive siderophores may provide a bioavailable form of iron to commensally associated phytoplankton, which could in turn affect algal growth and bloom dynamics. Here we report a laboratory-based study of binary cultures of the dinoflagellate Lingulodinium polyedrum, a major HAB species, with Marinobacter algicola DG893, a phytoplankton-associated bacterium that produces the photoactive siderophore vibrioferrin. Comparing binary cultures of L. polyedrum with both the wild type and the vibrioferrin minus mutant of M. algicola shows that bacteria are necessary to promote dinoflagellate growth and that this growth promotion effect is at least partially related to the ability of the bacterium to supply bioavailable iron via the siderophore vibrioferrin. These results support the notion of a carbon for iron mutualism in some bacterial-algal interactions.

Keywords: *Lingulodinium polyedrum*, *Marinobacter*, iron, algal-bacterial interactions, binary culture

# INTRODUCTION

Phytoplankton blooms are a frequent phenomenon in the coastal regions of every continent in the world. Certain phytoplankton algae occurring in mass proliferations produce toxins. Such harmful algal blooms (HABs) can, directly or indirectly, impact marine ecosystems with repercussions on local economies and public health (Lewitus et al., 2012). Direct effects on human health are often related to the consumption of shellfish that have ingested toxic phytoplankton, resulting in the accumulation of toxins and, consequently upon human consumption, paralytic, diarrheic, and neurotoxic poisoning which can sometimes be fatal (Anderson, 1994; Honner et al., 2012). HABs also have indirect impacts such as impairment of water quality leading to losses in the tourism and recreational sectors. Even phytoplankton blooms that do not produce toxins can be detrimental and lead to ecological impacts such as the displacement of indigenous species, habitat alteration, or oxygen depletion (Glibert et al., 2005). The economic effects caused by HABs in the U.S. alone were estimated at \$82 million per year in 2006 (Hoagland and Scatasta, 2006). Current strategies to reduce health-related impacts due to HABs are based on frequent coastal monitoring and early

### *Edited by:*

Stanley Chun Kwan Lau, Hong Kong University of Science and Technology, Hong Kong

### *Reviewed by:*

Marius Nils Müller, Universidade Federal de Pernambuco, Brazil Simon M. Dittami, UMR8227 Laboratoire de Biologie Intégrative des Modèles Marins, France

> *\*Correspondence:* Carl J. Carrano<EMAIL_ADDRESS>
### *Specialty section:*

This article was submitted to Aquatic Microbiology, a section of the journal Frontiers in Marine Science

*Received:* 27 February 2018 *Accepted:* 07 May 2018 *Published:* 05 June 2018

### *Citation:*

Yarimizu K, Cruz-López R and Carrano CJ (2018) Iron and Harmful Algae Blooms: Potential Algal-Bacterial Mutualism Between Lingulodinium polyedrum and Marinobacter algicola. Front. Mar. Sci. 5:180. doi: 10.3389/fmars.2018.00180 detection of HAB species and toxin levels. Nevertheless, the factors leading to the origins of HABs are still not well understood. Consequently, there is no strategy for their ultimate prevention.

There is an ongoing debate whether the factors leading to HABs are natural or anthropogenic. Oceanographic factors and global climate change including changes in upwelling and or relaxation and reversal of winds may be the major drivers initiating phytoplankton blooms (Roegner et al., 2002; Tweddle et al., 2010). HABs have also been proposed to be the consequence of human activities including eutrophication, changes in land use and agriculture, overfishing, and ballast water discharge (Glibert et al., 2005). Others consider that better analytics result in increased detection of HAB species (Burkholder, 1998).  While numerous studies concentrate on the causes of HABs, few have considered that bacterial species coexisting with microalgae could contribute to their development. However since Bell and Mitchell reported that specific bacterial communities occur and microbial activity was altered in the so-called phycosphere (Bell and Mitchell, 1972; Bell and Lang, 1974), more and more studies suggest that such algalbacterial interactions are very specific and important (Azam and Malfatti, 2007; Amin et al., 2015; Bertrand et al., 2015; Ramanan et al., 2016; Seymour et al., 2017). A plausible hypothesis suggests that the mutualistic association of some phytoplankton and bacteria is based on nutrient exchange. Nitrogen and phosphorus are the best studied in this regard, although the molecular nature of interactions involving these nutrients is not well understood (Hallegraeff and Gollasch, 2006). Alternatively specific bacteria may affect algal growth and bloom dynamics by their control of iron, a trace element which is often growth limiting to phytoplankton in the marine environment (Maldonado et al., 2005; Miethke and Marahiel, 2007; Croot and Heller, 2012).

Iron is an essential element for all living organisms including phytoplankton and bacteria due to its involvement in photosynthesis and respiration. Despite iron being the fourth most abundant element on the Earth, its bioavailability in the marine environment is extremely low due to its poor solubility under the mildly alkaline aerobic conditions present in the ocean (Martin and Fitzwater, 1988; Wu and Luther, 1994). To overcome this low bioavailability, bacteria and fungi have evolved sophisticated systems to produce high-affinity ironchelating compounds called siderophores to acquire, transport, and process this essential metal ion (Sandy and Butler, 2009). The major role ofsiderophores is to bind mineral phases of iron and to deliver the iron siderophore complex to specific outer membrane receptors on microbial cells. Several hundred siderophores have been isolated and extensively studied with respect to their synthesis, structures and transport mechanisms over the last three decades (Yamamoto et al., 1994; Challis, 2005; Sandy and Butler, 2009; Raymond et al., 2015). While much research has been done on terrestrial siderophores, the study of marine siderophores is less extensive and only a relatively few have been fully elucidated (Vraspir and Butler, 2009). Nevertheless one of the two major attributes that seem to distinguish marine siderophores from those of terrestrial origin is the tendency of the former to contain an α- and β-hydroxy acid group in their iron-binding domain (Barbeau et al., 2001, 2002; Küpper et al., 2006). The significance of the presence of these functional groups in siderophores is in their ability to make the ironsiderophore complex photoreactive. The chelated Fe(III) in such siderophores is reduced in the presence of sunlight via an internal redox process to release Fe(II), a more soluble form of iron (Amin et al., 2009b). It has been proposed that sunlight-driven reduction of the Fe(III) would transiently produce Fe(II) which might be utilized not only by the siderophore-producing bacteria themselves but also by non-siderophore producing bacteria and other organisms such as phytoplankton (Maldonado et al., 2005; Naito et al., 2008; Amin et al., 2009a, 2012a).

While iron acquisition by bacteria is well understood, that of phytoplankton remains less so. There are data to support the possibility of a variety of iron uptake mechanisms being operative simultaneously depending on the species involved. These include an iron-reductive route via a cell surface reductase, a direct xenosiderophore-mediated mechanism, and cell surfaceenhanced processes (Sutak et al., 2012; McQuaid et al., 2018). However as of yet there are no well documented examples, with some exceptions among the cyanobacteria, where phytoplankton have been shown to produce their own siderophores for iron acquisition (Raven, 2013). Thus how phytoplankton effectively acquire this metal from the low concentration iron environment is still unclear.

Our hypothesis is that algal growth and bloom dynamics may be affected by the increased bioavailability of iron engendered by the presence of photoreactive siderophores produced by mutualistic bacteria. Previously, we studied a bloom of the dinoflagellate, Lingulodinium polyedrum, at the Scripps Pier (San Diego, CA, USA) in 2011. L. polyedrum is one of the HAB species known to produce Yessotoxins, a group of polyether toxins which can accumulate in shellfish and show high toxicity to mice via intraperitoneal injection (Tubaroa et al., 2004). As the first step to search for an association of phytoplankton with specific bacterial photoreactive siderophore producers (producing petrobactin, aerobactin, and vibrioferrin siderophores), we monitored both the population of L. polyedrum and the bacterial siderophore producers before, during, and after the 2011 bloom. The results showed that both L. polyedrum and the bacterial siderophore producers simultaneously increased and decreased during the bloom period (Yarimizu et al., 2014). At the L. polyedrum bloom maximum, the total number of bacterial producers of photoreactive siderophore reached their maximum accounting for roughly 9% of the total bacterial population suggesting that such high abundance of photoreactive siderophore producers could potentially provide bioavailable iron to the phytoplankton in this environment. Furthermore, when the PCR-derived amplicons were sequenced, a phylogenetic tree constructed from the sequencing results showed that the community of siderophore producers in pre-bloom was statistically significantly different than those found during and after the bloom by UniFrac analysis suggesting that this particular bacterial community could be involved in bloom initiation.

As a proof of concept, the present study was performed using laboratory culture data to correlate the association of the vibrioferrin siderophore producing bacterium, Marinobacter algicola DG893 and the dinoflagellate L. polyedrum with iron as a nutrient. It is hoped that finding an algal-bacterial mutualism based on iron availability may be useful for a more thorough understanding of the mechanisms of HAB formation.

# MATERIALS AND METHODS

### Trace Iron Cleaning Procedures

The container cleaning technique was adopted from Bruland and Franks (1979). All plastic and glass containers were washed with pure water and soaked in 3 N hydrochloric acid for at least 2 weeks at ambient temperature. The acid-washed containers were rinsed with Milli-Q water and dried in a laminar-flow air bench. For those experiments requiring sterile conditions, acid-washed containers were placed in pouches and autoclaved at 121◦C for 30 min followed by a 45 min dry cycle. Alternatively, purchased sterile containers NuncTM Cell Culture Treated Flasks with Filter Caps were used.

### Growth Media

Of eight sources of natural waters screened, Pacific Ocean oligotrophic water was initially selected for our use due to its extremely low dissolved iron concentration (pM). This was replaced later by Scripps Pier seawater because of its easier accessibility and comparable L. polyedrum growth patterns observed in the two sources of seawater. Seawater from Scripps Pier (32.153◦N, 117.115◦W, San Diego, CA) was collected (pH 8.2–8.4, 980 mOsm/Kg H2O, total dissolved iron 3–4 nM) and filtered immediately through 0.22 mm pore size membrane. The filtered seawater was mixed with approximately 0.005% metal free hydrochloric acid, autoclaved for 30 min, cooled at ambient temperature for a day, and stored in 5◦C until use. The pH of the autoclaved seawater was ensured to be between 8.0 and 8.2 at ambient temperature. L1 nutrient was added to sterile seawater per manufacturer's instruction (L1 medium). For those cultures requiring controlled iron concentrations in media, L1 trace element solution without FeCl3 and Na2EDTA was prepared in house (Guillard and Hargraves, 1993) and added along with other L1 nutrients(nitrate, phosphate, silicate, vitamins) to sterile seawater (L1-Fe medium). Serial dilutions were made on L1 medium with L1-Fe medium to prepare growth medium with defined total iron concentrations, [Fe]T.

# Artificial Sea Water (ASW)

The component solutions were prepared in three separate containers, one with 15 g NaCl, 0.75 g KCl, 1 g NH4Cl in approximately 500 mL water, one with 12.4 g MgSO4 **.** 7H2O in small volume of water, and one with 3.0 g of CaCl2 **.** 2H2O in a small volume of water. The three solutions were mixed and 0.1 g disodium ß-glycerol phosphate was added. The pH was adjusted to 8.0–8.2 and volume was adjusted to 1 L. The solution was autoclaved at 121◦C for 30 min.

# Algal Maintenance and Growth Monitoring

The L. polyedrum strain used in this study was isolated from Venice Beach, California, and kindly provided to us by Avery Tatters (University of Southern California). L. polyedrum cells were maintained in sterile 1L Erlenmeyer flasks containing L1 medium and capped with a ventilation sponge. Cultures were exposed to 100 µmol photons m−2 s −1 on a 12-h alternating light and dark cycle at a temperature of 20 ± 2 ◦C (standard growth condition). The upper portion of a culture containing healthy cells was diluted 1/5 with L1 medium every 3 weeks. Algal growth was monitored by direct cell count using an inverted microscope (Nikon Eclipse TE2000-U, 4× objective). To prepare cells in controlled iron medium, cultures containing 104 -105 cells per mL was placed in a 15-mL Falcon tubes and centrifuged for a few seconds to pellet cells which were gently washed three times and re-suspended to an appropriate volume with controlled iron medium.

# "Axenic" *L. polyedrum* Preparation

Non-axenic L. polyedrum cells were grown to 105 cells/mL in L1 medium. Antibiotics 0.1% (v/v) (Penicillin 5 units/mL, Streptomycin 5µg/mL, Neomycin 10µg/mL) were added to a non-axenic culture and incubated for 24 h under the standard growth condition. Antibiotic-treated cultures were placed in 15 mL Falcon tubes and gently centrifuged for a few seconds. The pelleted cells were washed three times and re-suspended with L1 medium containing 0.05% (v/v) antibiotics in a sterile culture flask to appropriate volume. Antibiotics 0.05% (v/v) was added every 3 days to maintain the culture "axenic." The absence of cultureable bacteria in the antibiotic treated L. polyedrum was tested by spreading a 10 µL sample of the culture on a marine broth plate (5 g/L peptone, 1 g/L yeast extract, 15 g/L agar in 75% seawater) followed by incubation at 25◦C for 2–3 days. The absence of any visible bacterial colonies was considered indicative of an "axenic" culture of L. polyedrum.

### Bacteria Strains and Growth Monitoring

The model bacterium used was Marinobacter algicola DG893 (GenBank code NZ\_ABCP00000000.1) since it has been isolated by Green et al. (2004) and well characterized as a producer of the photoactive siderophore vibrioferrin (Amin et al., 2007). The mutant 1pvsAB-DG893 was made as described by Amin et al. (2012a) by 872 base pair deletion from the wild type to knock out two siderophore biosynthesis genes, pvsA and pvsB. To monitor bacterial growth, two methods were used: Optical density at 600 nm was applied to apparently turbid samples which generally contained ≥107 /mL of bacterial cells. Enumeration of bacteria by serial dilution was applied for samples containing lower numbers of bacteria cells (<107 /mL). For enumeration techniques, serial dilution was made on those samples with sterile seawater, each diluted sample was spread on marine broth plates, and the number of colony forming units (CFU) recorded from the lowest diluted sample plate. CFU in original sample per mL was calculated by (CFU in diluted sample)/(volume plated in mL) × dilution factor.

### Stock Bacteria Solution

One liter of marine broth was prepared (5 g/L peptone, 1 g/L yeast extract, 15 g/L agar in 75% seawater, pH 8.2, sterile) and dispensed into sterile tubes with a 5 mL volume. A few colonies of bacteria picked from stock plates were transferred into the marine broth tubes and shaken at 25◦C for 1–2 days until they reached log phase. The cells were collected by centrifugation, washed three times and re-suspended in sterile medium to an optical density (OD600) of 0.3 (stock bacteria solution).

### CAS-Dye Assay

The method was adopted from Alexander and Zuberer (1991) to detect siderophore production by DG893. Using acid washed glassware, the following stock solutions were prepared: 10 mM HDTMA in water, 2 mM CAS in water, 1 mM FeCl3.6H2O in water, 50 mM piperazine anhydrous buffer (pH 5.6), and 0.2 M 5-sulfosalicylic acid in water. CAS solution was prepared by pouring a mixture of 15 mL of CAS and 3 mL of FeCl3.6H2O slowly into 12 mL of HDTMA followed by addition of 50 mL of piperazine. The final volume was adjusted to 200 mL with water. CAS shuttle solution was prepared by adding 20 µL of 5-sulfosalicylic acid to every mL of CAS solution (used within 1 day of preparation). Samples to be tested were centrifuged to remove particles and 0.5 mL supernatant was mixed with 0.5 mL of CAS shuttle solution followed by incubation in the dark at ambient temperature for 10 min. A color change from dark purple to clear red indicated siderophore production.

### Materials and Reagents

The following materials were purchased from Sigma-Aldrich: Chrome azurol S (CAS, C-1018, MW605.28), antibiotics (Penicillin/Streptomycin/Neomycin, P4083- 100 mL), peptone (P-1265), iron(III) chloride hexahydrate (FeCl3 **.** 6H2O, 10025-77-1, MW270.30), manganese(II) chloride tetrahydrate (MnCl2 **.** 4H2O, 13446-34-9, MW197.91), hexadecyltrimethylammonium bromide (HDTMA, H6269- 100G, MW364.45), 1,4-diazacyclohexane, diethylenediamine (Piperazine, Sigma-Aldrich, P45907-100G, MW 86.14), 5-sulfosalicylic acid (390275, MW218. 18), zinc sulfate heptahydrate (ZnSO4 **.** 7H2O,<PHONE_NUMBER>, MW287.54), cobalt(II) chloride hexahydrate (CoCl2 **.** 6H2O,<PHONE_NUMBER>, MW129.83), copper(II) sulfate pentahydrate (CuSO4 **.** 5H2O, 209198-5G, MW249.69), sodium molybdate dehydrate (Na2MoO4 **.** 2H2O, 331058-5G, MW241.95), selenous acid (H2SeO3, 211176-10G, MW128.97), nickel(II) sulfate hexahydrate (NiSO4 **.** 6H2O, 227676-100G, MW262.85), sodium orthovanadate (Na3VO4, 450243-10G, MW183.91), potassium chromate (K2CrO4, 216615-100G, MW194.19), salicylaldoxime (SA, 84172-100G). The following materials were purchased from Fisher Scientific: trace metal-free hydrochloric acid (Optima, A466-250), yeast extract (BP9727-500), agar (BP1423-500), Casamino Acids (BP1424-500), NuncTM Cell Culture Treated Flasks with Filter Caps (12-565-57), ethylenediaminetetraaceticacid, tetrasodium salt dihydrate (Na4EDTA**.** 2H2O, BP121-500, MW416.2), boric acid (AAJ67202A1). Other materials were purchased as follows: 0.22µm filter membrane (Millipore, MillexGV, SLGV033RS), L1 medium kit (Bigelow, MKL150L), pure water (Barnstead water system, 18.2 mΩ). Vibrioferrin (VF, MW434.35) was isolated and purified in house according to Amin et al. (2007).

# RESULTS

### Bacterial Growth

The growth of DG893 and mutant 1pvsAB-DG893 in L1 medium containing different total iron concentrations [Fe]T (1,000, 100, 10, and 0 nM) was monitored. All media were divided in two parts, one maintaining [EDTA] at 10,000 nM (L1 nutrient level) with variable [Fe]T, and the other reducing [EDTA] along with [Fe]T to maintain molar ratio of Fe: EDTA of 1:1. All media were prepared with and without sterile Casamino Acids. In each medium, ca. 106 cells/mL of DG893 or mutant 1pvsAB-DG893 were added and shaken in the dark at 30◦C. The bacterial growth in each medium was monitored for 5 days by optical density at 600 nm. Neither WT DG893 nor the mutant grew in simple seawater or L1-Fe medium without a carbon source (**Figure 1A**). Addition of iron to seawater or L1-Fe medium also did not maintain growth (**Figure 1B**). However upon addition of Casamino Acids, both DG893 and the mutant grew and their growth increased with increasing [Fe]T in media (**Figure 1C**). There was no remarkable difference in growth pattern between DG893 and the mutant, consistent with the report by Amin et al. (2012a). In summary, both DG893 and the mutant require a carbon source to grow to a detectable level in seawater medium.

# Siderophore Production and Total Iron Concentration [Fe]T

The ability to produce the siderophore vibrioferrin (VF) by DG893 but not by the mutant was confirmed by the CASdye assay. Since siderophore biosynthesis is generally turned on under low iron concentrations and turned off under high concentrations (Braun, 1995; Braun et al., 1998; Miethke and Marahiel, 2007), WT was grown in L1 medium containing various [Fe]T with 0.3% Casamino Acids at 30◦C in dark for 24 h to find the approximate [Fe]T under which DG893 produces VF. The WT did not produce VF in the media with [Fe]T ≥10µM but did so with a [Fe]T of 1µM or less. The mutant 1pvsAB-DG893 did not show signs of siderophore production under any of the conditions tested. The results suggest that under normal seawater conditions, where [Fe]T is estimated to be nM to pM, that siderophore biosynthesis in DG893 is likely turned on.

# Axenic Culture Preparation

The difficulty in preparing and maintaining axenic cultures of many marine algae has been reported in the past (Green et al., 2004; Jauzein et al., 2015; Liu et al., 2017). In fact, Lupette et al. (2016) noted that completely axenic cultures of a model green algal species could not be maintained despite the use of antibiotic treatment protocols. We were partially successful in our efforts to prepare axenic L. polyedrum. Initially, non-axenic L. polyedrum cultures containing 105 cells/mL were treated with antibiotics (1% v/v Penicillin 50 units/mL, Streptomycin 50µg/mL, Neomycin 100µg/mL) for 24 h under the normal growth conditions. The treated cells were washed and resuspended with sterile L1 media and while the "axenicity" of the culture at this point was confirmed, bacterial colonies reappeared after only a few days. An attempt was made to keep 1% antibiotics in the culture longer than 24 h, however,

FIGURE 1 | added carbon source. The black bold line, gray bold line, gray wide dotted line, and gray narrow dotted line represent L1 media with 1,000, 100, 10, 0 nM [Fe]T, respectively. The mutant DG893 showed a similar pattern. (C) DG893 incubated in L1 media containing 10 mg/mL Casamino Acids and different [Fe]T. The black bold line, gray bold line, gray wide dotted line, and gray narrow dotted line represent L1 media with 1,000, 100, 10, 0 nM [Fe]T, respectively.

dinoflagellate cells began to disintegrate after 2 days suggesting that 1% antibiotics must be removed from the culture within 24 h. The fact that the negative control and sterile L1 medium without phytoplankton cells remained bacteria-free for weeks ruled out a possibility of culture contamination. We believe that the antibiotics are most likely eliminating only free-living bacteria leaving attached bacteria unaffected. The next effort of preparing axenic cultures was made by immediate dilution of antibiotic-treated cultures. The rationale of this attempt was to minimize attached bacteria on L. polyedrum cells by dilution. The antibiotic treated L. polyedrum culture was diluted 200-fold with sterile L1 medium and incubated under standard growth conditions. The diluted culture remained bacteria-free for the first 10 days, however microbial growth was observed on the 12th day as the number of L. polyedrum cells also increased. Although this dilution method suggested that the culture stayed bacteria free for up to 10 days, the L. polyedrum cell count was too dilute to be useful for further study. The next attempt at axenic culture preparation was made with three sequential antibiotic treatments. The non-axenic culture was treated with 1% antibiotics for 24 h, washed three times, and re-suspended with sterile L1 medium. The cells were allowed to recover under normal growth conditions for a day and then again treated with 1% antibiotics. This procedure was repeated three times. This procedure also failed as the dinoflagellate cells began to disintegrate. Finally, using a similar procedure but reducing the concentration of antibiotics to 0.1% for the first treatment and adding 0.05% subsequently every 2–3 days resulted in L. polyedrum cultures without visible cell damage that appeared to be free of free-living bacteria for more than a month.

### *L. polyedrum* Growth Screening

A matrix containing thirty cultures of L. polyedrum in L1 medium containing different numbers of DG893 cells in the initial inoculum (107 , 106 , 105 , 104 , 103 , 0 CFU/mL) and different [Fe]T (104 , 103 , 102 , 101 , and 0 nM) was set up. The same matrix was set up with the mutant 1pvsAB-DG893. Bacterial growth was monitored by enumeration of bacteria by serial dilution and phytoplankton growth was monitored by direct cell counts under the microscope. Regardless of the starting bacterial cell number, both DG893 wild type and mutant grew in all the binary culture matrices over 28 days to eventually reach a bacterial population of ca. 105 CFU/mL. The group of media containing 1,000 nM [Fe]T showed the greatest growth of L. polyedrum regardless of the starting bacterial inoculum (**Figure 2A**). The group of media containing ≤10 nM [Fe]T showed very slow L. polyedrum growth regardless of starting bacterial inoculum. No significant differences in L. polyedrum growth co-cultured with DG893 or the mutant were observed, which left a question as to whether VF secreted by DG893 would have a significant effect on L. polyedrum growth. It was speculated that under these growth conditions, the amount of VF being produced by DG893 was insufficient to see detectable changes in L. polyedrum growth. This was later confirmed (vide infra) when excess VF artificially added to the culture produced pronounced L. polyedrum growth. Finally, the growth pattern of axenic L. polyedrum was similar to that of a non-axenic culture. This observation left open another question as to whether DG893 and L. polyedrum were commensally associated. However this also was confirmed by experiments described below where after subsequent culturing the "axenic" L. polyedrum ceased to grow while binary L. polyedrum DG893 kept growing exponentially (**Figure 3**).

### Subsequent Batch Culture Growth of *L. polyedrum* in DG893 Binary Culture

L. polyedrum grown in L1 media containing various [Fe]T and DG893 (**Figure 2A**) for 28 days were diluted with appropriate media to adjust L. polyedrum to 102 cells/mL in the 2nd subsequent batch culture. The L. polyedrum growth in this 2nd subsequent culture was monitored for another 28 days under each growth condition. The result showed that the 2nd subsequent batch culture L. polyedrum in the higher two [Fe]T (10,000 and 1,000 nM) maintained their exponential growth while those in the lower three [Fe]T did not grow (**Figure 2B**). Although the cells under the latter conditions did not completely

FIGURE 2 | L. polydrum co-cultured with a starting inoculum of DG893 at 103 cells/mL in media with various [Fe]T. (A) First generation growth of L. polyedrum in the presence of DG893. The starting inoculum of DG893 was 103 cells/mL which reached 105 cells/mL after 28 days incubation under the standard growth conditions. A similar pattern was observed in all other groups: i.e., media containing a starting inoculum of DG893 of 107 , 106 , 105 , 104 , 103 , 0 cells/mL and [Fe]T of 104 , 103 , 102 , 101 , or 0 nM. (B) Second generation growth of L. polyedrum in the presence of DG893. After 28 days the cultures in (A) were diluted to a L. polyedrum concentration of 102 cells/mL with appropriate media and incubated under the standard growth condition for an additional 28 days.

FIGURE 3 | Rescue of L. polyedrum by addition of DG893. (A) First generation of L. polyedrum growth maintained in L1 media either "axenic" or co-cultured with DG893. (B) Growth of L. polyedrum in the second subculture either "axenic" or co-cultured with DG893. After the second subculture of "axenic" L. polyedrum was incubated under the standard growth conditions for 21 days, an inoculum containing approximately 102 cells/mL of DG893 was added and incubation continued for an additional 14 days.

die out, the few cells that survived had their swimming activity under the microscope clearly reduced.

### Axenic *L. polyedrum* Culture and Rescue

The growth of "axenic" and binary cultures of L. polyedrum with DG893 in L1 was compared over several subsequent batch cultures. No significant difference in growth was observed in the 1st culture, however, a clear difference was observed in the 2nd subsequent batch culture of L. polyedrum with and without DG893. While L. polyedrum with DG893 grew fully in both the 1st and 2nd subsequent batch cultures, "axenic" L. polyedrum ceased to grow (**Figure 3**). When DG893 was added to the 2nd subsequent "axenic" batch culture, L. polyedrum cells began to regrow exponentially demonstrating that DG893 could rescue "axenic" L. polyedrum.

### Non-axenic *L. polyedrum* Growth and Rescue

The non-axenic L. polyedrum was maintained in L1 medium and assumed to contain not only siderophore-producing bacteria but also other unknown bacteria. The non-axenic culture was divided in two parts, one diluted to 1/5 with L1 medium and other diluted to 1/5 with L1-Fe medium (batch subculture 1). When the cell count reached approximately 104 cells/mL, both cultures were further diluted with appropriate medium. Therefore, one culture maintained [Fe]T at L1 level throughout the subsequent batch cultures while the other contained successively reduced [Fe]T. The L. polyedrum growth was monitored for 28 days for each subculture. After the 6th subculture where [Fe]T in the medium was estimated to be 2 nM, L. polyedrum stopped growing completely (**Figure 4**). On 29th day of the 6th subculture, an iron supplement to give a [Fe]T of 5,850 nM was added to the culture and growth was continued to be monitored for additional 28 days. The results indicate that the L. polyedrum culture which stopped growing under 2 nM [Fe]T, could be rescued by addition of iron after 28 days (**Figure 5**).

# *L. polyedrum* Growth in DG893 Supernatant

Earlier it was stated that when L. polyedrum was co-cultured with either DG893 or the mutant 1pvsAB-DG893, little difference was observed in its growth pattern. We speculated that this may have been due to the small amount of VF produced by the equilibrium population of DG893 in the binary culture. To test this idea, samples of the supernatant of the media from the WT DG893 grown under ideal conditions and containing detectable amounts of VF by CAS-dye assay, as well as the mutant 1pvsAB-DG893 which did not, respectively, were added to L. polyedrum cultures and growth under these different conditions was compared. It should be noted that while we traditionally grow DG893 and the mutant in marine broth containing Casamino Acids, these were found to be toxic to L. polyedrum cells. In this experiment, we instead used ASW with succinic acid (at from 0.001 to 0.1%) as a carbon source (pH 8.2) to grow DG893 and the mutants prior to their supernatant being added to L. polyedrum cultures. L. polyedrum growth was enhanced the most by the addition of the DG893 supernatant. The mutant supernatant also showed a slightly positive effect on L. polyedrum growth compared to media without bacterial supernatant, however the degree of growth was not as large as that induced by the DG893 supernatant (**Figure 6**). These results suggest that VF has potential influence on L. polyedrum growth, but that other extracts from bacteria (vitamin B12?) may also have a potential effect on their growth. Further experiments were performed using purified VF itself. When purified VF at 57µM was directly added to a L. polyedrum culture in L1 media containing 100 nM [Fe]T, L. polyedrum reached cell counts approximately 1.5 times as large as those in media without VF.

### DG893 Growth in *L. polyedrum* Supernatant

"Axenic" L. polyedrum cells were resuspended in L1 media containing 100, 1,000, 10,000 nM [Fe]T and incubated under the standard growth conditions for 2 days to collect their organic extracts. The L. polyedrum cultures were divided in two parts, one of which was filtered through a 0.22µm membrane to collect only organics and the other was kept without filtration to keep phytoplankton cell debris. The L. polyedrum supernatant with and without cells was added to the stock DG893 containing 0.3% Casamino Acids, and the mixture shaken at 30◦C for several days. The control was prepared in the same manner without L. polyedrum supernatant and cells. In the media containing 1,000 and 10,000 nM [Fe]T, DG893 growth was enhanced by the presence of L. polyedrum supernatant and was even more pronounced with the supernatant containing cell debris (**Figure 7**). In media with 100 nM [Fe]T, only a subtle difference was observed in DG893 growth with and without L. polyedrum supernatant.

# DISCUSSION

Before proceeding to a discussion of the results presented here it is important to operationally define what we mean by [Fe]T and "axenic" cultures. Here we used the term total iron concentrations [Fe]T to mean the total analytical amount of iron added to culture and includes all phases of iron present in the solution. This is different from the total dissolved iron concentration which we routinely measure for environmental seawater samples (and which is approximately 2–6 nM for Scripps Pier water for example) as determined by cathodic stripping voltammetry (CSV). Total dissolved iron in such environmental samples are processed by filtration through a 0.22µm membrane followed by acidification. As a result, total dissolved iron in a sample is much lower than the total analytical iron concentration. We use this operational definition as it is not possible to measure total dissolved iron in culture media using the CSV method because equilibrium between the various iron oxo/hydroxo species potentially present in seawater at pH 8.2 is reached only extremely slowly and thus concentrations of total dissolved iron are constantly changing over the time course utilized in these experiments. Using a straight forward expression of total analytical iron concentration is thus a potentially more

reproducible approach with which to compare the effects of iron on bacterial and algal growth in a laboratory setting.

Secondly we use the term "axenic" in quotation marks for describing cultures of L. polyedrum, to mean the absence of any observable cultureable bacteria based on the absence of visual bacterial colonies when cultures were placed on solid media and incubated for several days. We were only partially successful making the culture "axenic" by treating the culture with low concentration of antibiotics and sequential addition of the antibiotics to the culture over extended times. Using this method the culture appeared to remain bacteria free for at least a month. However when the sequential addition of antibiotics to

the culture was stopped, bacteria growth was observed to return within a few days. Thus clearly the cultures were not truly axenic. Nevertheless our main concern was that the dominant species of bacteria growing in our binary culture should be DG893 and that "axenic" cultures should contain insignificant numbers of unknown bacteria. We confirmed the former by qPCR and the latter was further supported by presence of few if any bacteria seen via DAPI staining. To our knowledge, there is at present no general protocol for preparing a completely axenic culture of L. polyedrum hence our use of the operational definition of "axenic" described here.

We initially investigated conditions under which the bacterium Marinobacter DG893 and the dinoflagellate L. polyedrum could grow independently. As expected, in the absence of an added carbon source DG893 could not grow in natural unamended seawater or in L1 supplemented ASW irrespective of the iron concentration. In the presence of an adequate artificial source of carbon however bacterial growth was controlled by the available concentration of iron. Under idealized conditions, both DG893 and the VF null mutant reached a final concentration of around 105 CFU/mL independent of the initial inoculum. However even without any externally added carbon source DG893 grew well in the presence of L. polyedrum indicating that the dinoflagellate leaked enough dissolved organic carbon (DOC) to fully support the growth of the bacterium. The notion that DG893 could utilize L. polyedrum as a carbon source was further supported by the observation that growth of DG893 was stimulated when it was incubated in the media containing the supernatant from a filtered L. polyedrum culture which contains non-specific organics secreted from L. polyedrum. These may possibly include dimethylsulfoniopropionate (DMSP) a labile carbon and sulfur source known to "leaked" from dinoflagellates (Caruana and Malin, 2014).

In the case of both "axenic" and nonaxenic cultures of the photosynthetic dinoflagellate L. polyedrum, iron proved

FIGURE 6 | L. polyedrum growth in media with DG893 supernatant, mutant supernatant, and without bacterial supernatant. The black lines, gray lines, and dotted lines represent L. polyedrum growth in media with added DG893 supernatant, with added VF null mutant supernatant, and without any bacterial supernatant, respectively.

to be a growth limiting nutrient. Thus maximum growth occurred at total iron concentrations of 100–1,000 nM while higher concentrations proved toxic and lower concentrations (0–10 nM) were growth limiting. In the first culture, both "axenic" and non-axenic cultures grew equally well. However, their growth showed a remarkable difference in the second batch subculture. Here the non-axenic L. polyedrum culture continued to grow exponentially although now only at the higher [Fe]T. When the iron concentrations were kept high, L. polyedrum cultures continued to thrive up to at least 6 subcultures. However when the iron concentrations were allowed to be reduced by dilution in subsequent subculturing, the growth gradually ceased. Poorly growing subcultures in low iron media could however be rescued by the addition of additional iron to the culture. Remarkably "axenic" L. polyedrum subcultures ceased to grow regardless of [Fe]T but could be rescued by the addition of DG893. These results emphasize the importance of both bacteria and iron to L. polyedrum growth.

Our results further suggest that the L. polyedrum growth promoting effects of the bacterium Marinobacter DG893 were in fact not unspecific but at least partially related to its production of the photoactive siderophore vibrioferrin. Thus only the supernatant of DG893 that contained sufficient amounts of vibrioferrin facilitated L. polyedrum growth. While the addition of the supernatant from a culture of the VF null mutant of DG893 also improved growth of L. polyedrum the effect was much less than that from the WT. This observation was further confirmed by direct addition of purified VF into

FIGURE 7 | DG893 growth in media containing different [Fe]T with and without presence of L. polyedrum supernatant and cell debris. The solid black lines represent growth of DG893 in the presence of L. polyedrum supernatant with their cell debris, the gray lines are DG893 growth in L. polyedrum supernatant without their cell debris, and the dotted lines are DG893 without any L. polyedrum supernantant. DG893 was grown in media containing (A) 10,000 nM [Fe]T, (B) 1,000 nM [Fe]T, and (C) 100 nM [Fe]T.

the L. polyedrum culture which growth increased 1.5 times more than those without VF. These results imply that the presence of the siderophore vibrioferrin clearly influences L. polyedrum growth, but that other factors such as vitamin B12 (Cruz-Lopez and Maske, 2015) provided by bacteria may also be important in promoting this apparent bacterial-algal mutualism.

Here we have focused on a bacteria that produces the photoreactive siderophore, vibrioferrin. Vibrioferrin was chosen for more detailed study for a number of reasons. First vibrioferrin has been isolated from several different bacteria which are known to be closely associated with HAB species (Amin et al., 2007) and we found that bacteria that had the ability to synthesize vibrioferrin were by far the most numerous as compared to the producers of other photoreactive siderophores which we followed during the bloom of L. polyedrum at Scripps Pier in 2011 (Yarimizu et al., 2014). We initially supposed that vibrioferrin was a good candidate for a siderophore that could provide bioavailable iron for both the bacterial producers and their algal partners as it is the most rapidly photolyzed of the photoactive siderophores tested and that photolyzed vibrioferrin has no further affinity for iron. This feature is unusual in that most other photoactive siderophores retain the ability to strongly bind Fe(III) even after photolysis (Amin et al., 2009b). However we have recently shown that while vibrioferrin was the best source of iron for L. polyedrum of the ones we tested (Yarimizu et al., 2017), it was not due to its photolysis since iron uptake from VF was the same in the dark as in the light. We therefore attribute its ability to provide bioavailable iron to the dinoflagellate to its relatively weak iron binding properties (with respect to other more traditional siderophores) as it lacks the sixth donor group required to complete the octahedral coordination geometry preferred by Fe(III) (Amin et al., 2009b). This in turn leads to a less negative reduction potential (that falls within the biological range) so that the iron can readily be released from it by the cell surface reductases known to be present in L. polyedrum.

As a final note, while this laboratory study strongly supports a carbon for iron mutualism between the dinoflagellate L. polyedrum and Marinobacter DG893 as previously proposed (Amin et al., 2009a), it is unclear at present how, or indeed if, such a mutualism will operate in the field. To this end we have collected data from several field studies involving both HAB and non-HAB bloom events and are currently searching for relationships between phytoplankton, vibrioferrin producing bacteria, and available iron. We hope that both these laboratory and field studies will complement each other and advance our knowledge of the potential role of bacterialalgal interactions in understanding the mechanisms of HAB formation.

### AUTHOR CONTRIBUTIONS

KY, RC-L, and CC designing the study, executing the experiments, analyzing the data, interpreting the data, reviewing the data for accuracy and integrity, drafting the work, and revising this report.

### ACKNOWLEDGMENTS

We thank Avery Tatters (University of Southern California) for providing the L. polyedrum strain and helping us to set up an algal culture laboratory. We thank Professor Michael Latz (Scripps Institute of Oceanography) for providing us L. polyedrum. We also thank Professor Katharine Barbeau (Scripps Institute of

### REFERENCES

Oceanography) for providing us Pacific Ocean oligotrophic water and advice on iron measurement techniques as well as Professor Frithjof C. Küpper (University of Aberdeen) for a critical reading of the manuscript. A National Science Foundation grant to CJC (CHE-1664657) is gratefully acknowledged for partial support of this work.

and okadaic acid in mice. Toxicon 43, 439–445. doi: 10.1016/j.toxicon.2004. 02.015

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Yarimizu, Cruz-López and Carrano. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# The Bacterial Symbiont *Phaeobacter inhibens* Shapes the Life History of Its Algal Host *Emiliania huxleyi*

Anna R. Bramucci 1†, Leen Labeeuw1†‡, Fabini D. Orata1‡, Elizabeth M. Ryan2 , Rex R. Malmstrom2 and Rebecca J. Case1 \*

### *Edited by:*

Matthias Wietz, University of Oldenburg, Germany

### *Reviewed by:*

Rurun Wang, Merck, United States Hui Wang, Helmholtz-Zentrum für Infektionsforschung (HZI), Germany

> *\*Correspondence:* Rebecca J. Case<EMAIL_ADDRESS>
†These authors have contributed equally to this work.

### *‡Present Address:*

Leen Labeeuw, Plant Functional Biology and Climate Change Cluster (C3), University of Technology Sydney, Ultimo, NSW, Australia Fabini D. Orata, Department of Chemical and Materials Engineering, University of Alberta, Edmonton, AB, Canada

### *Specialty section:*

This article was submitted to Microbial Symbioses, a section of the journal Frontiers in Marine Science

*Received:* 17 February 2018 *Accepted:* 09 May 2018 *Published:* 29 May 2018

### *Citation:*

Bramucci AR, Labeeuw L, Orata FD, Ryan EM, Malmstrom RR and Case RJ (2018) The Bacterial Symbiont Phaeobacter inhibens Shapes the Life History of Its Algal Host Emiliania huxleyi. Front. Mar. Sci. 5:188. doi: 10.3389/fmars.2018.00188 1 Department of Biological Sciences, University of Alberta, Edmonton, AB, Canada, 2 Department of Energy Joint Genome Institute, Walnut Creek, CA, United States

Marine microbes form host-associated biofilm communities that are shaped by complex interactions between bacteria and their host. The roseobacter Phaeobacter inhibens exploits both symbiotic and pathogenic niches while interacting with its microalgal host Emiliania huxleyi. During co-cultivation over extended periods with E. huxleyi, we show that P. inhibens selectively kills two host cell types, the diploid calcifying strain and the haploid flagellated strain. Meanwhile, various non-calcifying diploid strains are resistant to this pathogen or the pathogen is avirulent to this cell type. This differential pathogenesis has the potential of dramatically altering the composition of E. huxleyi blooms, which are typically dominated by the roseobacter-susceptible calcifying strain. This cell type makes calcite plates, which are an important sink in the marine carbon cycle and forms part of the marine paleobotanic record. P. inhibens kills the haploid cells, which have been proposed as critical to the survival of the algae, as they readily escape both eukaryotic predation and viral infection. Consequently, bacteria such as P. inhibens could influence E. huxleyi's life history by selective pathogenesis, thereby altering the composition of cell types within E. huxleyi populations and its bloom-bust lifestyle.

Keywords: coccolithophore, roseobacter, phytoplankton pathogen, marine pathogens, pathogen ecology, cell type, phytoplankton life history, bacterial–algal interactions

# INTRODUCTION

On a microscopic scale, the marine environment is a heterogeneous mixture of nutrient "hotspots" formed by plankton and marine snow (Azam, 1998). Marine microbes take advantage of this aquatic array of nutrient gradients and "hotspots" by preferentially occupying specific niches (Hunt et al., 2008; Stocker, 2012). Motile microbes capable of directly associating with phytoplankton thereby expose themselves to a continuous stream of metabolites leaked from their algal hosts (Sapp et al., 2007; Geng and Belas, 2010; Sule and Belas, 2013). The concentrations of these metabolites are at their highest at the cell surface and within the phycosphere (Bell and Mitchell, 1972), the area immediately surrounding an algal cell, suggesting that algal-associated bacteria experience the greatest nutrient benefit and most likely to exchange communication and bioactive molecules.

While many bacteria directly benefit from algal metabolites exuding out of leaky or dying algae, some bacteria have fine-tuned their ability to sense and respond to host molecules by expressing symbiotic or pathogenic traits (Miller et al., 2004; Tripp et al., 2008; Seymour et al., 2010; Case et al., 2011; Wang et al., 2014; Amin et al., 2015) or releasing bioactive molecules that in turn alter the host's behavior or survival (Seyedsayamdost et al., 2011b; Labeeuw et al., 2017). Some bacteria engage in symbiotic relationships with their algal host by producing small molecules, such as vitamins or growth hormones, that are beneficial or even required by the alga (Bolch et al., 2011).

The haptophyte Emiliania huxleyi, which dominates coccolithophore blooms (Baumann et al., 2008), has recently been described as a host for bacteria in the marine environment (Seyedsayamdost et al., 2011b; Segev et al., 2016). It can quickly form dense populations (>106 cells·L −1 ) (Rhodes et al., 1995; Tyrrell and Merico, 2004) over vast expanses of the upper ocean (>250,000 km2 ) (Holligan et al., 1993). Its bloom-bust life cycle dramatically restructures the marine ecosystem, as it becomes a habitat-forming species for bacteria and viruses in the open ocean. Bloom formation has been associated with a variety of environmental factors (Tyrrell and Merico, 2004), while bloom collapse is frequently attributed to viral infection or microzooplankton grazing (Wolfe et al., 1994, 1997; Wilson et al., 2002). The ubiquity and abundance of E. huxleyi in the oceans, as well as its production of important intermediates in the carbon and sulfur biogeochemical cycles, have made it an important model phytoplankton species.

The E. huxleyi species complex, with an average diameter of 5–7µm, has three distinct cell types: non-motile non-calcifying cells (N-type, diploid), non-motile coccolith-bearing cells (Ctype, diploid), and motile scale-bearing cells (S-type, haploid) (Klaveness, 1972; Rhodes et al., 1995; Frada et al., 2012; von Dassow et al., 2015). E. huxleyi blooms are comprised of complex mixtures of C and S cells (Klaveness, 1972), though the fast growing calcifying C cells make up the majority of the bloom (Bidle et al., 2007; Baumann et al., 2008). It is not presently understood how these three cell types interact or alternate to shape the alga's life history, but each cell type is capable of asexual proliferation (Klaveness, 1972). However, the ways in which these different cell types impact the ecology, evolution, interactions, and metabolites of the E. huxleyi species complex is largely unknown, except for its interaction with viruses (EhVs).  Once an EhV infects a diploid cell, it proliferates and produces the viral glycosphingolipids that kill the algal cell (Vardi et al., 2009). Viral infection triggers caspaselike activity in E. huxleyi (associated with an upregulation of algal metacaspases), suggesting that the virus is hijacking algal programmed cell death (PCD) machinery to kill the host (Bidle et al., 2007).

Although viral infection and grazers have been studied (Wolfe et al., 1994; Wilson et al., 2002), the association of bacteria with E. huxleyi is largely unexplored (Seymour et al., 2010; Curson et al., 2011). We have investigated how E. huxleyi interacts with the marine α-proteobacteria Phaeobacter inhibens DSM 17395, previously named Phaeobacter gallaeciensis BS107 (Seyedsayamdost et al., 2011b). P. inhibens is a member of the roseobacter clade that is frequently identified within E. huxleyi blooms (González et al., 2000; Green et al., 2015; Segev et al., 2016). This bacterium produces a number of novel bioactives, including the antibiotic tropodithetic acid (TDA) (Geng et al., 2008; Berger et al., 2011; Thole et al., 2012) and potent algaecides called roseobacticides (Seyedsayamdost et al., 2011b, 2014) and roseochelins (Wang and Seyedsayamdost, 2017). These bioactives might allow P. inhibens to live a duplicitous lifestyle as both a beneficial symbiont and as a pathogen. TDA has been implicated in P. inhibens' chemical defense of various hosts (D'Alvise et al., 2012; Prol García et al., 2013; Beyersmann et al., 2017). Indeed, this bacterium is a symbiont of Ulva australis, chemically defending this ubiquitous seaweed from colonization (Rao et al., 2007). It can also act as a probiotic for turbot cod larvae, protecting it against Vibrio anguillarum infections (Planas et al., 2006). It also produces roseobacticides that could facilitate pathogenesis, as they have specific activity at very low concentrations against certain non-calcifying strains of E. huxleyi, causing these cells to lyse (Seyedsayamdost et al., 2011b). Additionally, a recent report suggested that P. inhibens uses the plant growth hormone indole-3-acetic acid (IAA) to kill calcifying (C-type) E. huxleyi (Segev et al., 2016), although the dynamics of this interaction have yet to be elucidated.

We co-cultured an axenic representative strain of the three E. huxleyi cell types (N diploid, C diploid, and S haploid) with P. inhibens and monitored photosynthetic health, as well as the cell dynamics of both the algae and bacterium. We demonstrate that P. inhibens selectively causes the precipitous populationwide death of aged E. huxleyi coccolith-bearing C cells as well as the motile scale-bearing S cells, while various non-calcifying diploid N populations were not killed. Algal death directly benefits the bacterium, which increases its own population size during its host's decline. Finally we utilized a P. inhibens mutant library to demonstrate that the potent roseobacticides produced by P. inhibens, which were previously shown to kill N cells at nanomolar concentrations (Seyedsayamdost et al., 2011b), are not required for P. inhibens pathogenesis of C or S cells. These findings suggest that alternative, as of yet unknown, pathogenicity factor(s) may play equally critical roles in the pathogenesis of P. inhibens toward E. huxleyi.

# MATERIALS AND METHODS

### Algal and Bacterial Strains

All E. huxleyi strains were obtained from the National Centre for Marine Algae and Microbiota (East Boothbay, ME, USA). This includes coccolith-producing (C-type) strain CCMP3266, scalebearing haploid (S-type) strain CCMP3268, and non-calcifying diploid (N-type) strains CCMP370, CCMP372, CCMP374, CCMP379, and CCMP2090. The N-type strains are all from unique geographical locations spanning the biogeographical range of all available axenic E. huxleyi cultures (**Table 1**). They were maintained in L1-Si medium (Guillard and Hargraves, 1993) at 18◦C in a diurnal incubator (8:16 h dark-light cycle) with 41.51 ± 11.15 µmol·m−2 ·s −1 of light during the light period. Algal cultures and medium were checked for bacterial contamination by microscopic observations and by inoculation onto half-strength marine agar (½ MA) (18.7 g Difco Marine

\*Calcifying diploid (C), non-calcifying diploid (N), and scale-bearing haploid (S).

†Co-culture experiments were observed over 30 days without precipitous algal death. ‡The axenic form of the polymicrobial non-calcifying diploid strain CCMP1516.

Broth 2216 and 9 g NaCl, supplemented with 15 g Difco agar in 1 L) followed by incubation at 30◦C for 2 days. E. huxleyi was inoculated with a 10−1 dilution, grown statically for 5 days to 105 cells·mL−1 (early-log) prior to all experiments.

The wild type P. inhibens DSM 17395 (Frank et al., 2014), was maintained at 30◦C on ½ MA (see Supplementary Methods for identification of the bacterium). Colonies were then transferred to half-strength marine broth (18.7 g Difco Marine Broth 2216 and 9 g NaCl in 1 L). P. inhibens was grown to stationary phase in a shaking incubator at 30◦C, 160 rpm, for 24 h and then subsequently re-cultured in the same conditions prior to experimentation. Transposon mutants of P. inhibens DSM 17395 (Wetmore et al., 2015) were grown under the same conditions with the addition of 200 µg·mL−1 kanamycin (Sigma-Aldrich).

### *P. inhibens* Transposon Mutants

Transposon mutants in TDA biosynthesis have been identified previously as deficient in TDA and roseobacticide production and consequently have white colonies, as opposed to the typical brown colonies of the wild type (Wang et al., 2016). Mutants screened all produced white colonies and were in two genes in the tda gene cluster of a P. inhibens plasmid pPGA1\_262: tdaB (PGA1\_262p00970) and paaZ2 (PGA1\_262p00800); as well as two genes in the paa gene cluster encoded on the chromosome: paaA (PGA1\_c04080) and patB (PGA1\_c00860) (Wetmore et al., 2015; Wang et al., 2016).

Gene knockout strains of P. inhibens DSM 17395 were obtained from a previously generated library created through molecularly barcoded transposon mutagenesis (Wetmore et al., 2015; Price et al., 2016), and specific mutants were recovered following the approach outlined by Cole et al. (2017). Briefly, the P. inhibens mutant library was spread on MA plates with 50 µg·mL−1 kanamycin and incubated overnight at ∼22◦C. Colonies were picked using Qpix460 (Molecular Devices), arrayed into 384-well plates containing half-strength YTSS media (2 g yeast, 1.25 g tryptone, and 20 g sea salts in 1 L) with 7.5% glycerol and 50 µg·mL−1 kanamycin, and incubated overnight at ∼22◦C. Next, 25 nL of each well was collected using an Echo525 liquid handler (Labcyte) as part of a multiplexing strategy involving pooling of rows, columns, and plates. These pools were subject to PCR amplification and sequencing of the molecular barcodes that identified the transposon insertion site of each mutant strain (Wetmore et al., 2015). The well location of each mutant was determined from amplicon sequencing results using an in-house script. Successful interruption of the targeted genes was confirmed by sequencing over the mutation site. Genomic DNA was extracted using GeneJet Genomic DNA Purification Kit (Thermo Scientific). Gene-specific primers were designed, and PCR was performed using the Phusion High-Fidelity DNA polymerase (Thermo Scientific). PCR products were either column- or gel-purified using QIAquick PCR Purification Kit or MinElute Gel Extraction Kit (QIAGEN), respectively. Amplicon sequencing was performed using the Sanger dideoxy method (Applied Biosystems 3730 Genetic Analyzer).

### Bacterial and Algal Co-cultivation

Bacterial-algal co-cultivation was performed as previously described (Bramucci et al., 2015). Briefly, stationary phase bacterial cultures were washed twice by centrifugation and resuspended in L1-Si medium before further centrifugation and resuspension to the target cell concentration in colony forming units (CFU)·mL−1 . E. huxleyi and the bacteria were mixed 1:1 (volume:volume) with a final concentration in the co-culture of 102 CFU·mL−1 bacteria and 105 cells·mL−1 algae, then 1 mL of this co-culture was aliquoted in 48-well plates (Becton Dickinson). The same cell densities of the bacterium and the alga were inoculated as monocultures in L1-Si medium and aliquoted into the microtitre plate. All controls/co-cultures were performed in triplicate. The microtitre plates were incubated in a diurnal incubator (8:16 h dark-light cycle) at 18◦C for all experiments.

Co-cultivation of CCMP3266 or CCMP3268 with P. inhibens transposon mutants were performed as described, with control and co-cultures amended with kanamycin to a final concentration of 100 µg·mL−1 to keep selective pressure for the mutants (Wetmore et al., 2015). E. huxleyi was grown with various concentrations of kanamycin (0, 10, 50, 100, 200 µg·mL−1 ) to determine that 100 µg·mL−1 kanamycin did not adversely affect growth of photosynthetic yield (Supplementary Figure S1). The cell density of mutants was enumerated on ½ MA with and without 200 µg·mL−1 kanamycin throughout the co-cultivation with algae to ensure the transposon mutation had not been lost from P. inhibens.

### Roseobacticide B Inhibition Assay

Roseobacticide B was obtained from Dr. Mohammad Seyedsayamdost (Princeton University, NJ, USA) and dissolved in methanol as previously described (Seyedsayamdost et al., 2011b). The compound was then added to senescent (i.e., cultures were declining after reaching their maximal cell fluorescence) E. huxleyi (CCMP3266 and CCMP3268) at half the maximal inhibitory concentration (IC50) previously identified for the sensitive N strain (Seyedsayamdost et al., 2011b), as well as ten-fold lower and higher (final concentrations of 0.019, 0.19, and 1.9µM). A final concentration of 2% methanol was added to experimental and control cultures. The samples were incubated in microtiter plates in the same conditions previously used for bacterial co-culture experiments for 24 h before obtaining pulse-amplitude-modulation (PAM) fluorescence measurements.

### Fluorescence Measurements

A PAM fluorometer (WATER-PAM, Heinz Walz) was used to measure chlorophyll fluorescence (Schreiber et al., 1986; Bramucci et al., 2015). All samples were taken at the midpoint of the dark cycle (4 h) and diluted in sterile L1-Si medium to within the detection range of the PAM fluorometer. Samples were maintained at 18◦C throughout handling. A dark adaption period of 3 min was determined, after which a saturating pulse was applied and the fluorescence readings were taken in triplicate at intervals of 1 min 30 s to calculate the minimal dark fluorescence (F0), the maximum dark fluorescence (Fm), and the photosystem II (PSII) maximum efficiency (Fv/Fm), Fv/Fm = (Fm−F0)/Fm (Schreiber et al., 1986; Baker, 2008). Triplicate readings of each sample were averaged and the three microtitre wells were treated as replicates to determine the maximum quantum efficiency.

# Flow Cytometry

Subsamples from algal controls and co-cultures were fixed for flow cytometry, incubated in the dark for 10 min with 0.15% glutaraldehyde (Sigma-Aldrich), flash-frozen in liquid nitrogen, and stored at −80◦C until flow cytometry was performed using a FACSCalibur (Becton Dickinson). A 488 nm laser was used for excitation. Samples were then run using chlorophyll fluorescence (emission = 670 nm) for detection and cell counting (cells·mL−1 ).

# Bacterial Enumeration

Phaeobacter inhibens population density grown alone and in coculture was enumerated by counting CFU on ½ MA after 2 days of incubation at 30◦C. Five replicate counts from each well were averaged, and the triplicate wells were used as experimental replicates for analysis. Although P. inhibens attaches to E. huxleyi and itself, aggregated cells were not observed microscopically after 5 min of vigorous vortexing.

### Microscopy

Brightfield images of algal controls and co-cultures were obtained using a 63× Axio Scope.A1 objective lens (Zeiss), equipped with an Optronics digital camera and PictureFrame software v2.3 (Zeiss). Epifluorescence images were obtained using a 100× Axio Imager.M2 microscope objective lens (Zeiss), equipped with a monochrome camera (AxioCam 506 mono). Epifluorescence microscopy was also used to assess algal chlorophyll auto-florescence and to visualize algal and bacterial DNA when stained with DAPI (4′ ,6-diamidino-2 phenylindole dihydrochloride) (DNA–DAPI complex: excitation = 364 nm; emission = 454 nm) (Life Technologies). Unfixed E. huxleyi control and co-culture aliquots were stained with DAPI according to manufacturer's instructions (30◦C, 20 min) then immediately pelleted by centrifugation (5,000 × g, room temperature, 2 min). Cells were gently washed twice in sterile L1-Si medium and analyzed immediately on the epifluorescence microscope. Images were acquired simultaneously for three different channels and overlaid using Zen 2 Blue Edition software v2 (Zeiss). The differential interference contrast channel was overlaid with 1) algal chlorophyll auto-fluorescence (red: excitation = 610–650 nm; emission = 670–720 nm) and 2) DNA– DAPI complex fluorescence (blue: excitation = 350–400 nm; emission = 417–477 nm).

### Data Processing and Statistical Analysis

Flow cytometry data were processed using FlowJo v9.2 (Tree Star Inc.). Quantitative data from all other experiments were processed using SigmaPlot 12.0 (Systat Software). Statistical significance was determined using a one-way ANOVA and Tukey HSD test.

# RESULTS AND DISCUSSION

### *P. inhibens* Kills Select *E. huxleyi* Cell Types

This study involves the bacterium previously identified as P. gallaeciensis BS107 that produces roseobacticides, which have a specific algaecidal effect on E. huxleyi (Seyedsayamdost et al., 2011b); however, there is evidence that there were differences in strains of the bacteria submitted to various repositories (Buddruhs et al., 2013). Therefore, we sequenced the genome of our strain and its identity was confirmed to be P. inhibens DSM 17395 (Frank et al., 2014) based on both 100% average nucleotide identity and percent (in silico) DNA–DNA hybridization to the published DSM 17395 genome (Thole et al., 2012; Supplementary Figure S2).

To determine if P. inhibens has a host cell type preference in its interactions with E. huxleyi, we co-cultured P. inhibens with five axenic N strains from distinct geographical locations, one axenic C strain (CCMP3266), and one axenic S strain (CCMP3268) (**Table 1**). The C and S strains were both killed by P. inhibens, and all tested N type strains survived, regardless of geographic origin (**Table 1**). To further investigate the differential pathogenesis of P. inhibens on various E. huxleyi cell types, an axenic N [CCMP2090, derived from the polymicrobial calcifying CCMP1516 (Orata et al., 2016; Zhang et al., 2016)], C (CCMP3266), and S [CCMP3268, derived from a single cell isolation from CCMP3266 (von Dassow et al., 2015)] strains (**Figure 1**) were investigated in more depth. Each strain was grown alone and in co-culture with P. inhibens for 14 days and monitored for PSII maximum quantum efficiency (Fv/Fm), which is affected by cellular stress and/or loss of functional PSII centers (**Figure 2**), as well as algal and bacterial cell density (**Figure 3**).

When grown alone, all three algal cell types had a stable PSII maximum quantum efficiency (Fv/Fm > 0.5) throughout the experiment, although they entered a senescent stage, represented by reduced fluorescent health after 6–10 days (**Figures 2A–C**). The algal cultures grown alone did not die in this experiment and have been shown to live for > 60 days in microtitre plates (Bramucci et al., 2015). Both C and S cultures grown alone reached their peak density between 6 and 10 days followed by a characteristic slow decline during senescence (**Figures 3B,C**), typified by gradual losses of cell numbers (Franklin et al., 2012). It has been suggested that P. inhibens stimulates E. huxleyi's growth rate in the early growth phases (Segev et al., 2016); however, no growth stimulation was detected for the N, C, or

S cell types in the co-cultures, with no significant difference between E. huxleyi's cell density for any of the cell types in co-culture or the control in the first 6 days of the experiment (**Figure 3**).

The N strain grown in co-culture with the bacterial symbiont maintained chlorophyll health, functional PSII systems (**Figure 2A**), and cell density (**Figure 3A**) comparable to the axenic cultures throughout the experiment, despite P. inhibens being found attached to its surface (**Figures 2D,E**). This algal strain, alone or in co-culture with the bacterium, grows to a higher cell density than C- or S-type grown alone from 12 days onwards (P < 0.05) (**Figure 3**).

In contrast, the C and S cultures grown in co-culture with P. inhibens experienced an accelerated and premature decline in photosystem health (8–10 days) (**Figures 2B,C**). This population-wide decline is associated with a concurrent decrease in individual chlorophyll content in the algal cells cocultured with the bacterium (**Figures 2F–I**). When co-cultured with P. inhibens, C-type cell densities are much lower than the control by 14 days (P < 0.05) compared to the axenic control (**Figure 3B**). A similar trend is observed for the Stype strain (**Figure 3C**). All S cells lost their photosynthetic ability in the co-culture with P. inhibens by day 12 with a detectible decrease in photosynthetic health initiated on day 8 (**Figure 2C**). This timing for rapid decline triggered by a bacterial pathogen coincides closely with the onset of senescence in the algal host.

The rapid decline of PSII efficiency when co-cultured with P. inhibens is intriguing, as it has not been observed in starved senescent E. huxleyi cells (Franklin et al., 2012), but occurs during viral infection and lysis of E. huxleyi cells by EhVs (Bidle et al., 2007). These findings suggest that the algaecidal activity of P. inhibens against its microalgal host, E. huxleyi, is dependent on algal cell type, and that the targeted cell types are different to EhVs which are only known to kill N- and C-type cells (Frada et al., 2008; Mordecai et al., 2017). Similarly, the closely related roseobacter pathogen, Ruegeria sp. R11 (Fernandes et al., 2011) (also known as Nautella italica R11; Vandecandelaere et al., 2009; Rodrigo-Torres et al., 2016), kills C and S cells and not N cells (Mayers et al., 2016). While the mechanism of P. inhibens pathogensis is not known, clues can be found in EhVs, which upregulate metacaspase activity in calcifying diploid cells, inducing alga PCD (Bidle et al., 2007). EhVs induce an autophagy-like PCD event in the algal host (Schatz et al., 2014). Autophagy, or the genetically programed lysosomal degradation of cellular constituents (Kroemer et al., 2009), is a vital part of the host immune response and can infer protection against bacterial pathogens by ensuring the rapid degradation of virulence factors (Cemma and Brumell, 2012). Because this viral-induced PCD response in E. huxleyi results in a loss of PSII function directly before death due to viral lysis (Bidle et al., 2007), it might follow that a similar mechanism is activated by the bacterial pathogen. Environmental stress is a factor in gradual loss of photosynthetic health, but the sudden complete loss of photosynthetic health has only so far been seen in the C cells undergoing viral-induced apoptosis. The activation of algal caspase-like molecules resulting in apoptosis-like PCD has already been described in the pathogenic interaction between Ruegeria sp. R11 and the C- and S-type culture CCMP3266 (Mayers et al., 2016).

This resistance of N-type cells to pathogenic roseobacters has been previously reported for the N-type strain, CCMP2090, cultured with Ruegeria sp. R11 (Mayers et al., 2016). Together, these findingssuggest that, although the N cell used in the current study (CCMP2090) was recently derived from a polymicrobial calcifying parent strain (CCMP1516), calcifying and noncalcifying diploid strains have important biological differences, which might for instance confer widespread resistance to roseobacter pathogenesis. There are at least two possible explanations as to why E. huxleyi N cells are widely resistant to P. inhibens pathogenesis, or that P. inhibens does not interact with N cells. The first possibility is that N cells escape the pathogen, similar to the haploid escape from EhVs (Frada

et al., 2008; Mordecai et al., 2017). The haploid S-type is thought to be resistant to viral lysis, and it was postulated that the haploid's scaly coverings or that gene loss or mutation might infer viral resistance to the haploid cell type (Frada et al., 2008). Supporting this theory, an increase in culture temperature confers temperature-induced resistance to EhVs by altering the outer sphingolipids of representative calcifying and non-calcifying diploid E. huxleyi strains, impeding viral recognition and infection of target cells (Kendrick et al., 2014). By analogy, it is possible that P. inhibens does not interact with E. huxleyi N cells due to differential recognition and/or attachment to the alga, as was suggested for Ruegeria sp. R11 (Mayers et al., 2016). However, P. inhibens does attach to N cells (**Figure 1B**) and so N cell resistance, or P. inhibens lack of virulence, must have another mechanism. A second possibility for N cell escape is that there might be genetic differences between C and N cells. There are two prevalent theories as to how N cells are generated: (1) through a series of mutations resulting in, among other differences, a malformed coccolith-forming vesicle (van der Wal et al., 1983) and (2) due to prolonged lab domestication (Zhang et al., 2016). To our knowledge, only one axenic C and S culture exist, and it is commonly reported that coccoliths were lost through subsequent culturing. For example, the polymicrobial CCMP1516 was a C-type culture that recently became an N-type culture; it was a C-type culture when it produced its axenic daughter strain CCMP2090, which is N-type (Zhang et al., 2016). Future genomic effort should focus on sequencing parent (C) and daughter (N) cultures to elucidate if a genetic basis for the phenotypic switch can be identified as it could elucidate resistance mechanisms to the bacterial pathogen and the molecular basis of coccolithogenesis.

### *P. inhibens* Benefits From Growth With *E. huxleyi* Before and After its Death

Bacterial population dynamics when grown alone and in coculture were monitored using CFU (**Figures 3D–F**). Under the experimental conditions used (algal medium L1-Si from seawater, with no additional carbon), P. inhibens was able to grow to a maximum cell density of 105 CFU·mL−1 without E. huxleyi at 2–4 days (**Figures 3D–F**), after which P. inhibens cell density declined. Prolonged monitoring of P. inhibens grown without a host showed that all bacteria died by 25 days. However, its growth was greatly enhanced by the presence of E. huxleyi, reaching

100–1,000 times higher densities when grown with an algal host (**Figures 3D–F**).

The growth benefit conveyed to P. inhibens is greater in the short term (14 days and less) when grown with C- and S-type cells of E. huxleyi than the N-type (**Figures 3D–F**). The bacterial cell density in the C cell co-culture appears to benefit from 2 to 6 days (P < 0.01) and 10 days (P < 0.05) when compared to the N culture (**Figures 3D,E**). Directly after the C-type culture suffers a decline in both PSII health and numbers of chlorophyll containing cells (day 12), P. inhibens cell density increases 8.5 fold (P < 0.01), compared to those grown in co-culture with Ntype, which does not die (day 12) (**Figures 3A,B,D,E**). P. inhibens co-cultured with S-type also has a significant (P < 0.01) benefit in terms of increased cell counts compared to the N-type coculture between 4 and 10 days (**Figures 3D,F**), the time when the algae has reduced PSII maximum quantum efficiency as well as declining cells per mL in the co-culture (**Figures 2C**, **3C**). Additionally, the benefit to P. inhibens persisted after the death of the algal host (12–14 days). In fact, P. inhibens grown with C- and S-type maintained high cell counts until at least 30 days, when both co-cultures had 106 and 107 CFU·mL−1 , respectively, despite death of its host. P. inhibens grown in co-culture with N-type maintained high cell counts until at least 30 days (108 CFU·mL−1 ).

This increased persistence and higher population density of P. inhibens when co-cultured with E. huxleyi is likely the result of nutrients made available by algal exudate (Borchard and Engel, 2012). Phytoplankton constantly leak sugars and oxygen from photosynthesis, as well as other chemicals such as dimethylsulfoniopropionate (DMSP) and amino acids, which are chemoattractants for roseobacters (Mitchell et al., 1985; Baines and Pace, 1991; Miller and Belas, 2004; Miller et al., 2004). Furthermore, DMSP-degrading microbes, such as P. inhibens, benefit directly from being able to efficiently assimilate the sulfur from DMSP directly into bacterial amino acids (Curson et al., 2011). These results demonstrate a population wide benefit for P. inhibens to being grown in co-culture with an algal host, regardless of host cell type. P. inhibens then increases its fecundity and persistence by killing certain cell types of its host, presumably because algal cell lysis could provide it with nutrients (Kolb et al., 2013). This interaction makes P. inhibens an opportunistic pathogen, not a parasite (Seyedsayamdost et al., 2011a), as

methanol. The final concentration of methanol solvent added to control and experimental cultures was 2% of the final volume. P. inhibens transposon mutants deficient in roseobacticide synthesis were co-cultured with (C) CCMP3266 and (D) CCMP3268. Minimal algal fluorescence of the population (F0) was measured on day 10. Triplicate wells that represent independent experiments were sacrificially sampled; error bars represent ±1 standard error.

parasites reduce the health and fecundity of their host and benefit from prolonging the host's life, while P. inhibens causes C- and S-type cells to die prematurely.

### Are Roseobacticides Responsible for Death of C- and S-Type Cells?

The rapid decline in maximum quantum yield and loss of chlorophyll observed in the C- and S-type cultures at 8–10 days (**Figure 2**) is consistent with the physiological response of E. huxleyi to roseobacticides, which causes cell membrane blebbing, chloroplast loss, and lysis of N-type cells (Seyedsayamdost et al., 2011b). Roseobacticides are produced by P. inhibens when supplied with p-coumaric acid (pCA) (Seyedsayamdost et al., 2011b) and other lignin precursors (Wang and Seyedsayamdost, 2017). pCA is produced by E. huxleyi and hypothesized to be made during senescence (Seyedsayamdost et al., 2011b) where it is an intermediary of the lignin and flavonoid pathways (Labeeuw et al., 2015).  To determine if roseobacticides were responsible for decreases in quantum yield and chlorophyll loss, roseobacticide B was added to senescent C and S cultures of E. huxleyi on day 9—the day when those E. huxleyi cultures typically decline when grown with P. inhibens—at IC50 of 0.19µM determined for another N-type strain, CCMP372 (Seyedsayamdost et al., 2011b). Roseobacticide B concentrations 10-fold higher and lower than the IC50 were also tested. Surprisingly, no changes in PSII maximum quantum yield were observed at any roseobacticide concentrations (**Figures 4A,B**). Furthermore, the characteristic declines in C and S cultures were reproduced when co-cultured with four different P. inhibens mutants whose roseobacticide production genes had been disrupted (**Figures 4C,D**). These results suggest roseobacticides were not responsible for the death of C- and S-type cells, and that P. inhibens must be producing additional algaecidal compounds or virulence factors to kill E. huxleyi. In addition, non-calcifying diploid strains surviving co-culture with P. inhibens included CCMP372 (**Table 1**), which were previously shown to be killed by roseobacticides (Seyedsayamdost et al., 2011b).

Phaeobacter inhibens produces many different compounds with roles in microbe-microbe interactions. Segev et al. (2016) has proposed that P. inhibens uses IAA to kill E. huxleyi.  However, Segev et al. (2016) also showed that P. inhibens did not produce IAA at high enough concentrations to kill E. huxleyi CCMP3266 (0.4–10 nM). In addition, E. huxleyi CCMP3266 produces IAA itself as a cell-cell signal at a higher concentration (200 µM) in the presence of tryptophan (Labeeuw et al., 2016). Finally, the Ntype strain, CCMP2090, is susceptible to IAA when co-cultured at lower concentrations (10–100µM), exhibiting morphological responses and reduced health similar to those seen in terrestrial plants (Labeeuw et al., 2016) that we did not observe in the P. inhibens–CCMP2090 co-culture. Taken together, these data show IAA is probably not the bioactive molecule causing E. huxleyi's decline when grown with P. inhibens, and that some other molecule or mechanism is responsible.

Tryptophan, the precursor to IAA, was also found to be lethal at the high concentration (1,000µM) required to produce sufficient IAA to kill E. huxleyi (Labeeuw et al., 2016). Tryptophan was also shown to be released by CCMP3266, and the addition of exogenous tryptophan causes faster killing of CCMP3266 by both P. inhibens (Segev et al., 2016) and another roseobacter, Ruegeria sp. R11 (Labeeuw et al., 2016). This indicates that tryptophan may be a signal or metabolite for the bacteria to become virulent.

### CONCLUSION

Phaeobacter inhibens interaction with E. huxleyi is dependent on the cell type of its algal host. This differential pathogenesis is also observed for EhVs, although the targeted cell types

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differ. Currently, N-type cells are heavily relied upon for cell biology studies of E. huxleyi because they are readily maintained axenically and avoid many of the problems associated with working with an autofluorescent mineral (i.e., coccoliths). However, our findings suggest that we should not assume that N-type cells will have similar biological interactions to their calcifying counterparts. The ability of P. inhibens to target coccolith-bearing (C) diploid cells and scale-bearing (S) haploid cells, while not killing non-calcifying diploid cells is a unique role, likely representing differentiation from EhVs, which kills both diploid cell types but not the haploid cell type (Frada et al., 2008). We have now shown two roseobacters, P. inhibens and Ruegeria sp. R11, can kill populations of C- and S-type cells (Mayers et al., 2016). This may have wide-reaching implications, as S cells are resistant to EhV lysis, which was postulated as a mechanism for the algal population to regenerate the dominant C cell population following viral-induced bloom collapse (Frada et al., 2008). Given the present findings, it is possible that the roseobacter's rapid pathogenesis of S cells could limit the alga's ability to reseed C cell populations. Additionally, roseobacter killing of C and S populations, while N populations evade bacterial induced death, could have important implications for the marine carbon cycle and formation of the paleobotanical record (Coolen, 2011), as well as the overall distribution of E. huxleyi blooms.

# AUTHOR CONTRIBUTIONS

AB, LL, and RC conceived of the experiments. AB and LL carried out the experiments. FO carried out the whole-genome sequencing and analysis. ER and RM constructed the P. inhibens mutants. AB, LL, FO, and RC drafted the manuscript. All authors have read and approved the manuscript.

### ACKNOWLEDGMENTS

We would like to thank Dr. Mohammad Seyedsayamdost (Princeton University) for providing the roseobacticide B and Albert Rosana (University of Alberta) for assistance with sequence confirmation of the barcoded transposon mutants. This work was supported by the Natural Sciences and Engineering Research Council of Canada (grant 402105) to RC. The work conducted by the U.S. Department of Energy Joint Genome Institute, a DOE Office of Science User Facility, is supported under Contract No. DE-AC02-05CH11231.

### SUPPLEMENTARY MATERIAL

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**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Bramucci, Labeeuw, Orata, Ryan, Malmstrom and Case. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Novel ssDNA Viruses Detected in the Virome of Bleached, Habitat-Forming Kelp Ecklonia radiata

Douglas T. Beattie1 \*, Tim Lachnit 1, 2, Elizabeth A. Dinsdale3 , Torsten Thomas 1 and Peter D. Steinberg1, 4

1 Centre for Marine Bio-Innovation and School of Biological, Earth and Environmental Sciences, University of New South Wales, Sydney, NSW, Australia, 2 Zoological Institute, Christian-Albrechts-University Kiel, Kiel, Germany, 3 Department of Biology, San Diego State University, San Diego, CA, United States, 4 Sydney Institute of Marine Science, Mosman, NSW, Australia

Kelp forests provide essential habitats for organisms in temperate rocky shores. Loss of kelp forests has occurred over large areas in a number of temperate regions, including in Australia, where the dominant kelp Ecklonia radiata has been lost from substantial areas of the shoreline. Loss of E. radiata has been associated with environmental stressors, including increased temperature and anthropogenic contaminants, as well as biological factors, such as herbivory. Disease may also play a role, but there is little information on the role of disease in the loss of kelp from coastal ecosystems or on the potential role of pathogenic microorganisms, such as viruses. E. radiata across much of its distribution in Australia can develop a "bleached" phenotype, which may be a disease. To investigate whether the phenotype was associated with a potential viral agent, we shotgun sequenced viral particles that were isolated from kelp with normal (healthy) and bleached phenotypes. Each virome consisted of ∼380,000 reads, of which ∼25% were similar to known viruses. All samples were dominated by bacteriophages, but novel ssDNA virus sequences were detected that were almost exclusively in viromes from the bleached kelp phenotype. These ssDNA viruses are covered by 11 contigs that contained complete capsids and characteristic rep genes that were 30–60% similar to those of circular, Rep-encoding ssDNA viruses (CRESS-DNA viruses). CRESS-DNA viruses have not previously been described from macroalgae, and the rep genes were similar to CRESS-DNA viruses from marine water samples, snails, crabs, anemones, but also dragonflies. This raises the interesting possibility that the kelp could be a vector of the CRESS-DNA viruses to other organisms that are associated with the bleached state.

Keywords: circovirus, microbial ecology, virome, kelp, marine disease, metagenome, CRESS-DNA, viral ecology

### INTRODUCTION

Ecklonia radiata is a large, stipitate brown alga (class Phaeophyceae, order Laminariales, or commonly, kelp), and the dominant primary habitat-forming organism over 71,000 km2 of the temperate rocky coastlines of Australia (Bennett et al., 2016). Kelps, such as E. radiata, provide habitat and nutrients that are essential to coastal food webs (Steneck et al., 2002), and the importance of E. radiata habitats in temperate Australia has led to them being described as

### Edited by:

Matthias Wietz, University of Oldenburg, Germany

### Reviewed by:

Karen Dawn Weynberg, The University of Queensland, Australia Simon M. Dittami, UMR8227 Laboratoire De Biologie Intégrative Des Modèles Marins, France

\*Correspondence: Douglas T. Beattie<EMAIL_ADDRESS>
### Specialty section:

This article was submitted to Microbial Symbioses, a section of the journal Frontiers in Marine Science

Received: 20 November 2017 Accepted: 22 December 2017 Published: 12 January 2018

### Citation:

Beattie DT, Lachnit T, Dinsdale EA, Thomas T and Steinberg PD (2018) Novel ssDNA Viruses Detected in the Virome of Bleached, Habitat-Forming Kelp Ecklonia radiata. Front. Mar. Sci. 4:441. doi: 10.3389/fmars.2017.00441 "The Great Southern Reef " (Bennett et al., 2016). Fisheries, tourism, and other industries taking place within these habitats are estimated to be worth AU\$10 billion annually to the Australian economy (Bennett et al., 2016).

In recent years there has been a loss of kelp cover in temperate regions of Australia from locations across much of the range of E. radiata, from Perth on the West coast, along the Southern coast, up to the Solitary Islands on the East coast (Connell et al., 2008; Ling and Johnson, 2012; Vergés et al., 2016; Wernberg et al., 2016). Loss of kelp results in major losses in biodiversity, ecosystem function, and associated fisheries (Andrew and O'Neill, 2000; Bishop et al., 2010; Burge et al., 2014; Vergés et al., 2016; Wernberg et al., 2016), and in Australia loss of E. radiata has been associated with a number of environmental stressors. This includes stressors associated with urbanization, such as increased nutrients and sedimentation of the coastal region (Connell et al., 2008), and ocean warming, through both direct physiological effects of temperature as well as indirect effects through range extensions of herbivorous sea urchins or fishes, which consume large areas of kelp forests (Wernberg et al., 2011; Ling and Johnson, 2012; Vergés et al., 2016). Though relatively little studied in seaweed systems, microbial disease may also play a role in loss of habitat-forming seaweeds (Egan et al., 2014). Disease in marine systems appears to be increasing globally, including in communities that are dominated by primary habitat-forming organisms, such as coral reefs and kelp forests (Rosenberg et al., 2007; Vega Thurber et al., 2009; Krediet et al., 2013), and disease in seaweeds has been linked to anthropogenic stressors, such as ocean warming (Campbell et al., 2011; Case et al., 2011).

Microbial communities are integral to the health of all organisms, such that hosts and their associated microbial communities (microbiome) are increasingly being treated as a holistic system, or holobiont (Egan et al., 2013). This approach is now common for understanding the interaction between marine habitat-forming organisms, such as corals, seaweeds, or sponges (Bourne et al., 2009; Egan et al., 2013; Webster and Thomas, 2016). Disturbances to the microbiome of these organisms are associated with stress and disease of the host (Morris et al., 2016). For example, disruption of the microbiome of the Australian macroalga Delisea pulchra leads to bleaching and reduced reproductive capacity (Egan et al., 2013; Campbell et al., 2014; Wahl et al., 2015), and microbial pathogens cause bleaching of D. pulchra in both the field (Campbell et al., 2011) and in laboratory inoculation experiments (Kumar et al., 2016). Disruption of the holobiont and resultant disease is increasingly seen as critical for other marine habitat forming organisms.

We have recently identified a bleaching phenotype in E. radiata, which occurs widely across its biogeographic range and which is characterized by bleached areas of the thallus with reduced photosynthetic capacity (Marzinelli et al., 2015). The bacterial components of the microbiome of bleached vs. nonbleached E. radiata were distinct, suggesting that the microbiome of the algae undergoes a shift associated with the disruption of normal function (e.g., photosynthesis; Marzinelli et al., 2015). However, specific bacterial pathogens, which induce bleaching, have not yet been extensively characterized. Other components of the microbiome may also affect the health of hosts. In particular, viruses are likely ubiquitous components of marine holobionts and are now emerging as pathogens or potential pathogens in a diverse range of marine organisms, including archaea (Rohwer and Thurber, 2009), microalgae (Brussaard, 2004; Weynberg et al., 2017), sea stars (Hewson et al., 2014), and macroalgae (Lachnit et al., 2015).

Viruses consist of two major groups, bacteriophages that infect bacteria and viruses that infect eukaryotes. The tailed bacteriophages with dsDNA genomes, Caudovirales, include the families Siphoviridae, Myoviridae, and Podoviridae. Bacteriophages also include Inoviridae, which possess ssDNA genomes, and together these bacteriophages form a major component of marine viral assemblages (Mann, 2005; Dinsdale et al., 2008b; Roux et al., 2012; Xue et al., 2012). Abundant eukaryote viruses detected in algae are from the family Phycodnaviridae, which have a dsDNA genome of 100–550 kb and infect both unicellular and multicellular algae (Wilson et al., 2009). Phycodnaviridae have been investigated in the filamentous brown-alga Ectocarpus siliculosus, a model organism, and a few genomes have been sequenced (Van Etten et al., 1991; Müller and Frenzer, 1993; Easton et al., 1997; Kapp et al., 1997; Maier et al., 1998; Delaroque and Boland, 2008). However, few studies have been conducted on macroalgae from the field (e.g., viruses in Laminaria digitata McKeown et al., 2017) and none on E. radiata.

ssDNA viruses, which have small circular genomes (1– 5 kb), are an emerging group of viruses that are implicated in marine disease and mass mortalities across a number of marine phyla, including fishes, molluscs, echinoderms, and crustaceans (Hasson et al., 1995; Friedman et al., 2005; McLoughlin and Graham, 2007; Lang et al., 2009). ssDNA viruses include a subgroup which possess circular genomes, and a particular replication initiation gene,rep. These are therefore called Circular Rep-Encoding ssDNA (CRESS-DNA) viruses (Rosario et al., 2009; Delwart and Li, 2012; Labonté and Suttle, 2013; Simmonds et al., 2017). These include the Geminiviridae, Nanoviridae, Circoviridae, and Genomoviridae, which employ rolling-circle replication (RCR), facilitated by the Rep protein (Rosario et al., 2012b). In addition to the Rep protein, CRESS-DNA viruses have a characteristic stem-and-loop secondary structure, which facilitates RCR. The loop region contains a conserved nonanucleotide motif, NANTATTAC (Labonté and Suttle, 2013). The Circoviridae and Nanoviridae are diverse in aquatic and marine environments, including ocean, freshwater, wastewater, deep-sea vents, Antarctic lakes and ponds, and hot springs (Angly et al., 2006; Rosario et al., 2009, 2012a; Cassman et al., 2012; Liu et al., 2012; Labonté and Suttle, 2013).

Here we investigate the viruses associated with bleached and non-bleached E. radiata from Sydney, Australia. First, virus particles were extracted from kelp tissue. DNA was extracted from these and amplified using whole-genome amplification (WGA), which is biased toward small and circular genomes (Sabina and Leamon, 2015), but thus aids in detection of potentially rare and small genomes in metagenomic samples (Kim et al., 2008; Cheval et al., 2011; Rosario et al., 2012a; Fahsbender et al., 2015). This was followed by shotgun sequencing and metagenomic analysis. We describe the general features of the kelp virome, and test whether components of the virome differ between the non-bleached and bleached phenotype, in order to explore whether viral pathogens may be a possible cause of the bleaching phenotype. Our results show that circoviruses were nearly absent in the non-bleached kelp, but in high abundance in the bleached kelp, and we describe these novel viruses in detail.

### METHODS

### Sampling

Individual thalli of E. radiata were sampled from Long Bay in Sydney, Australia (−33.97, 151.26) during June 2014 at a depth of 3 m. The water temperature at the time of sampling was 16◦C. Tissue samples of blades were taken from three bleached individuals and three non-bleached, healthy, thalli. These phentoypes are described in Marzinelli et al. (2015). Viral particles were isolated from kelp tissue as described in Lachnit et al. (2015).

### DNA Extraction and Metagenomic Sequencing

Viral DNA was extracted from virus particles in SDSextraction buffer (1% SDS, 200 mM Tris, 50 mM EDTA, pH 7.5) in the presence of 1% (w/v) polyvinylpyrrolidone (MW 40,000), 1% 2-mercaptoethanol and proteinase K (0.5 mg ml−1 ) at 37◦C for 30 min, followed by 56◦C for 15 min. Double the volume of DNA-extraction buffer (2.6 M NaCl, 150 mM Tris, 75 mM EDTA, 2.5% cetyltrimethylammonium bromide (CTAB), 1% 2-mercaptoethanol, pH 7.5) was added and samples were incubated for another 15 min at 56◦C. An equal volume of chloroform:isoamyl alcohol (24:1) was added to the solution, mixed and centrifuged at 12,000 RCF at room temperature for 5 min. The chloroform:isoamyl alcohol extraction of the supernatant was repeated three times to remove contaminating polysaccharides and proteins. The supernatant was transferred into a new tube and the nucleic acids were precipitated by the addition of 0.7 volume of isopropanol. After an incubation step at 20◦C for 2 h nucleic acids were pelleted by centrifugation at 14,000 RCF at 4◦C for 20 min. The pellet was washed with 75% (v/v) ethanol, air dried and dissolved in 30 µl molecular grade water.

All samples underwent whole genome amplification steps using a REPLI-g Mini Kit (QIAGEN), where the reaction was conducted using 20 µL of denatured DNA, incubating the reaction mix at 30◦C for 16 h. DNA was frozen and submitted to the Ramaciotti Centre at UNSW for library preparation using a TruSeq Low Input kit (Illumina). Shotgun metagenomic sequencing was conducted using an Illumina MiSeq sequencer with 2 × 250 bp chemistry. A total of six viral metagenomes (or viromes) were sequenced, three from kelp with the bleached phenotype and three from the non-bleached phenotype. The reads were processed using Prinseq (Schmieder and Edwards, 2011) by trimming the ends where the quality score was below 20, and removing exact duplicates. The paired-end reads were assessed using default settings on PEAR (Zhang et al., 2014) to determine if they could be joined to form longer sequences.

### Virome Profile Analysis

Several bioinformatics approaches were used to identify the types of phage and eukaryotic viruses in the kelp viromes. First, the reads were aligned against the NCBI non-redundant protein database (NR), downloaded on May 05 2017. Alignment was performed using BLASTx (Altschul et al., 1990) with an e-value of 10−5 . Alignment matches were screened to remove all non-viral similarities found. Alignment results were analyzed using MEGAN (Huson et al., 2016) to infer best taxonomic and/or functional assignments. The Least Common Ancestor (LCA) parameters was used with a minimum score of 50, max expected e-value 10−5 , top 10%, LCA 100%, minimum complexity 0. To detect significant differences in the relative abundances of assignments to each virus family, STAMP (Parks et al., 2014) was used to perform two tailed Welch's t-test (α = 0.05) with Bonferroni correction to compare the bleached and non-bleached kelp viromes.

To assess possible contamination of the viromes by host genomic or bacterial DNA, the entire set of similarities found, including similarities to viruses and non-viruses, was considered. Sequences, which had higher alignment bitscores to non-viruses, were aligned against all the brown-alga sequences in NCBI (downloaded on May 05 2017), using BLASTn with an e-value of 10−5 . The sequences with <95% similarity to Ecklonia spp. suggested that they are not part of the brown algae. This threshold was chosen as essential genes (e.g., rbcL) of different species of Eckonia, and even some distally related members of the brown algae are >95% similar (Rothman et al., 2015). Decontamination programs (e.g., DeconSeq Schmieder and Edwards, 2011) were not employed to automate the removal of potential bacterial contamination, as genes are often shared between bacteriophage and their hosts. Instead, more information on the origin of the reads was obtained by assembly and assessment of the genomic content (see below), and contamination was retrospectively removed from the results and analyses.

The individual metagenomes were assembled to provide longer contigs using SPAdes 3.9.0 (using default parameters; Bankevich et al., 2012). SPAdes was chosen as it performs better than other assemblers on virome data (Hesse et al., 2017; Papudeshi et al., 2017; Vollmers et al., 2017). Reads were mapped to the contigs using Bowtie 2.2.5 (Langmead and Salzberg, 2012) and visualized using Tablet (Milne et al., 2009) to assess the evenness of read coverage and to identify possible chimeric assemblies, which were removed from further analysis. To investigate the distribution of viruses between the bleached and non-bleached kelp, the contigs were aligned to the NR database as described above. The contigs and their component reads were considered contamination and removed from the analysis and results if (a) the alignment bitscore to a virus reference was less than to a non-virus sequence, and (b) the alignment matches did not contain core genes of the assigned virus taxon e.g., phage-tail genes of Caudovirales.

# Viral Genome Reconstruction

All six metagenomes were pooled and cross-assembled with SPAdes 3.9.0 to obtain longer contigs and/or complete genomes. Assessment and alignments of the cross-assembled contigs were performed as described above. At this stage of the analysis, circoviruses were identified as the only part of the eukaryotic DNA virus community that varied in abundance between bleached and non-bleached kelp, and therefore were investigated in more detail. Open reading frames (ORFs) on the circoviral contigs were detected using Prodigal (Hyatt et al., 2010) with a length of > 50 amino acids. Predicted protein sequences were aligned against one another, and their reference sequences using the MUSCLE aligner with default settings (Edgar, 2004). To visualize the diversity of the ORFs a phylogenetic tree was produced using the Jukes-Cantor genetic distance model and the neighborjoining algorithm with 100 bootstraps. Circoviridae contigs were submitted to Mfold (Zuker, 2003) to determine their stem-and-loop secondary structure. The loop motif was compared across the Circoviridae group through alignment using MUSCLE.

# RESULTS

### Overall Composition of the Virome of Ecklonia radiata

The six E. radiata viromes contained an average of 386,000 reads with an average length of 149 ± 2SD bp. (Supplementary information S1). No reads were removed through quality control, although 14% of the reads were trimmed up to 10 bp where the read quality was <20. Most paired reads could not be joined, and so forward-reads were only used in alignment searches. Analysis of alignment results found 81,256 ± 22,106 virus-like reads per metagenome (∼20% of the sequences were similar to the NR database). The majority (86–94%) of virus-like reads were assigned to bacteriophage in the order Caudovirales (**Figure 1**). In order of abundance, similarities were detected to the viral groups Siphoviridae (average of 48% of virus-assigned reads), Myoviridae (16%), unclassified bacteriophages (12%), Inoviridae (8%), Podoviridae (7%), followed by Circoviridae (7%). Within the Caudovirales, reads similar to Siphoviridae were more abundant (adjusted p-value = 0.003) in the non-bleached kelp phenotype, however the total number of bacteriophage reads within the Caudovirales did not significantly differ between non-bleached and bleached kelp viromes. In the eukaryotic viral fraction, reads similar to Phycodnaviridae and other large dsDNA viruses were lower in abundance (<2% of reads assigned) compared to the bacteriophages and did not differ in abundance between viromes of non-bleached and bleached kelp (adjusted p-value = 0.785). Sequences of ssDNA viruses belonging to the Circoviridae were rare overall (0.48% of total reads), but were almost exclusively in bleached kelp viromes. Six reads were assigned to Circoviridae from a single non-bleached kelp virome, whereas 6,403 reads were assigned from bleached kelp viromes.

Assembly of all reads per individual metagenome provided an average of 4,011 ± 351 contigs of 38,709 ± 6,392 bp length. The taxonomic assignment of the contigs mirrored that of the reads (Supplementary information S2). Bacteriophage possessed the greatest amount of read coverage in the contigs (60%), followed by the Phycodnaviridae (8%) and Circoviridae (6%). The Circoviridae contigs (n = 42) were assembled only in the bleached kelp viromes, with the exception of a single contig from one non-bleached kelp virome. Similar to the read data, Circoviridae contigs were the only eukaryotic viral group that varied in coverage between the bleached and non-bleached kelp phenotypes (adjusted p-value = 0.002).

Cross-assembly of all six metagenomes yielded 7,561 contigs, the longest being 246,893 bp (N50 20,791 bp, N95 246 bp). When aligned against an NCBI virus-only protein database, 847 contigs showed similarity to viruses (**Figure 2**). In descending order of abundance these were assigned to Siphoviridae (97 contigs), environmental viruses from metagenomes (92), Myoviridae (59)**,** Podoviridae (48), unclassified bacteriophages (47), Circoviridae (26), Phycodnaviridae (21), Mimiviridae (15), Inoviridae (9) and other virus families with very low abundances (total of 434).

We assessed contamination by regarding the contig similarities to virus and non-viruses. Two thousand two hundred and sixty-seven contigs of the total 7,561 were given a taxonomic assignment through alignment to NR (Supplementary information S3). Eight hundred and forty-seven contigs had similarity to viruses. Of these, 146 contigs had no similarity to other references, 576 contigs were also similar to bacteria and 125 contigs were also similar to eukaryotes. The majority (36%) of contigs similar to both viruses and eukaryotes were similar to E. siliculosus Virus-1. Four contigs had both higher alignment bitscores to brown-alga sequences, and did not include characteristic virus genes or conserved domains and were thus assumed to be eukaryotic. The contigs with similarity to both viruses and bacteria (576) were examined further, finding 113 contigs with higher bitscores to bacterial references and no characteristic bacteriophage genes, and assumed to be from bacterial cells. Contigs and their constituent reads that were considered eukaryotic or bacterial contamination were removed from the analyses and results of virus abundances and gene content. Contigs that were assigned as circoviruses had no other similarities to references in NR, and as the circoviruses were the only eukaryotic virus that showed variation between kelp phenotypes, these were analysed in more detail.

# Investigation of Circoviridae-Like Contigs

The Circoviridae contigs (n = 26) were 739–5,420 bp long, and contained up to two ORFs. The first ORF was 855 bp, and the only translated similarity between these and reference proteins were two ORFs with similarity to putative CRESS-DNA virus capsid genes from environmental metagenomes. These showed 30% pairwise identity to putative capsid genes of a circular virus associated with a sea anemone (Genbank accession: YP\_009163900.1) and a circular ssDNA virus isolated from an Antarctic ice shelf pond (Genbank accession: YP\_009047127.1).

The second ORF was 305–336 amino acids and was the characteristic rep genes of CRESS-DNA viruses. The rep genes were found in three different lengths (915 bp, 924 bp, and 1,008 bp) suggesting three genotypes, which we refer to from hereon as A-C. There were three contig configurations of the circovirus sequences: Type-A (8 contigs), Type-B (2 contigs) and Type-C (single contig; **Figure 3**). Type-A contained both the capsid and repA, in reverse orientation with a conserved intergenic region (IR) between the two ORFs. The IR of the eight contigs shared a pairwise identity of 70%. Type-B contained repB and the IR. Type-C contained only repC and no capsid sequences or the IR. The origin of replication (ori), included the characteristic nonanucleotide stem and loop structures involved in rolling circle replication of CRESS-DNA viruses and was located upstream of the rep gene in all contigs (**Figure 3**). The nonanucleotide motif in each contig was TAGTATTAC. The other 15 of the 26 circovirus-like contigs that did not contain

complete ORFs were found to contain fragments of rep and capsid genes of circoviruses, suggesting incomplete assembly.

The rep genes from the 11 contigs and the closest reference sequences in the database were aligned. The rep genes formed three clusters with each cluster showing >95% similarity. Between all rep genes in our dataset, and their most similar reference sequences, the rep sequences were 27–60% similar. The references with greatest similarity to the detected circoviruses were from viruses found in association with aquatic samples or invertebrates, including marine snails and hermit crabs, but also dragonflies (**Figure 4**).

# DISCUSSION

We characterized the virome of both bleached and nonbleached individuals of the kelp E. radiata. Sequences similar to bacteriophages in the Siphoviridae were detected in greatest abundance overall, followed by Myoviridae, unclassified phages, Inoviridae, and Podoviridae. For most viral taxa there was little difference in the relative abundance in reads between the two kelp phenotypes. However, eukaryote circoviruslike sequences differed significantly in abundance between bleached and non-bleached kelp, with bleached viromes containing the vast majority (>99%) of circovirus-like reads. The circoviruses sequences from E. radiata included three phylogenetic groups, which all contained the characteristic rep gene of CRESS-DNA viruses, and were most similar to recently described circoviruses from pelagic water samples and marine invertebrates.