The impact of associated bacteria on morphology and physiology of the dinoflagellate Alexandrium tamarense

Harmful Algae 50 (2015) 65–75

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The impact of associated bacteria on morphology and physiology of the dinoflagellate Alexandrium tamarense Ce´cile Jauzein a,*, Andrew N. Evans b, Deana L. Erdner a a b

University of Texas Marine Science Institute, Port Aransas, TX, USA Department of Coastal Sciences, University of Southern Mississippi, Ocean Springs, MS, USA

A R T I C L E I N F O

A B S T R A C T

Article history: Received 12 January 2015 Received in revised form 9 October 2015 Accepted 9 October 2015

Despite their potential impact on phytoplankton dynamics and biogeochemical cycles, biological associations between algae and bacteria are still poorly understood. The aim of the present work was to characterize the influence of bacteria on the growth and function of the dinoflagellate Alexandrium tamarense. Axenic microalgal cultures were inoculated with a microbial community and the resulting cultures were monitored over a 15-month period, in order to allow for the establishment of specific algal–bacterial associations. Algal cells maintained in these new mixed cultures first experienced a period of growth inhibition. After several months, algal growth and cell volume increased, and indicators of photosynthetic function also improved. Our results suggest that community assembly processes facilitated the development of mutualistic relationships between A. tamarense cells and bacteria. These interactions had beneficial effects on the alga that may be only partly explained by mixotrophy of A. tamarense cells. The potential role of organic exudates in the establishment of these algal–bacterial associations is discussed. The present results do not support a role for algal–bacterial interactions in dinoflagellate toxin synthesis. However, variations observed in the toxin profile of A. tamarense cells during culture experiments give new clues for the understanding of biosynthetic pathways of saxitoxin, a potent phycotoxin. ß 2015 Elsevier B.V. All rights reserved.

Keywords: Algal–bacterial interactions Dinoflagellate Growth Photosynthesis Toxin

1. Introduction Phytoplankton forms the base of the food web and plays a key role in carbon cycling in marine environments. Interest in the carbon cycle has increased recently owing to problems such as climate change and coastal eutrophication. Fertilization of largescale areas of the ocean has even been proposed to mitigate climate change by stimulating the growth of phytoplankton in order to sequester carbon dioxide (Denman, 2008; Glibert et al., 2008). Impressive advances have been made in assessing environmental forcing of phytoplankton dynamics during the last decades, mainly focusing on physico-chemical controlling factors (Tian, 2006; Irwin et al., 2012). However, some mechanisms, such as biological associations between microorganisms, are still poorly understood, and their role in phytoplankton dynamics and biogeochemical cycles of marine ecosystems could be crucial.

* Corresponding author. Present address: Sorbonne Universite´s, UPMC Univ Paris 06, INSU-CNRS, Laboratoire d’Oce´anographie de Villefranche, Villefranche sur mer, France. Tel.: +33 4 93 76 38 43. E-mail address: [email protected] (C. Jauzein). http://dx.doi.org/10.1016/j.hal.2015.10.006 1568-9883/ß 2015 Elsevier B.V. All rights reserved.

The activities and fates of algae and aquatic bacteria are closely linked (Amin et al., 2012), yet the ecological significance of most naturally occurring algal–bacterial associations is still unclear. The most well-known connection is via their roles in energy fluxes and nutrient cycling, as bacteria utilize algal-derived organic matter and, in turn, regenerate inorganic nutrients (Legendre and Rassoulzadegan, 1995). There is some evidence that differences in the quality of organic matter produced by different types of phytoplankton cause shifts in the species composition of bacterial communities utilizing these algal exudates (van Hannen et al., 1999). Whilst such trophic relationships can structure microbial communities, specific interactions between algae and bacteria can alter this dynamic in subtle ways, potentially influencing bloom dynamics and the cycling of matter in the ocean (Mayali et al., 2011). Over the last several decades, coastal areas throughout the world have experienced an escalating threat associated with occurrence of ‘‘harmful algal blooms’’ (HABs). This trend comes from a global expansion of harmful algal species, aggravated by an increase in the use of the coastal zone for fisheries and recreational activities (Anderson, 2009). Among potential factors controlling outbreak and persistence of HABs, algal–bacterial interactions

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have received increasing attention in the recent past (Doucette et al., 1998; Mayali and Azam, 2004; Hasegawa et al., 2007). Algicidal marine bacteria have even been considered as potential biological control agents for regulation of HABs in natural seawaters (Doucette et al., 1999; Kim et al., 2008). A better understanding of algal–bacterial interactions could help to improve management strategies for HAB events, including mitigation and forecasting. Algal–bacterial interactions are complex, partly because they can be species-specific and vary with changes in environmental conditions (Mayali and Azam, 2004; Grossart and Simon, 2007; Danger et al., 2007). For the toxic dinoflagellate Alexandrium spp., which produces paralytic shellfish toxins (PSTs), various effects of bacteria have been reported, including promotion or inhibition of growth (Ferrier et al., 2002; Wang et al., 2012), stimulation or inhibition of encystment (Adachi et al., 1999; Adachi et al., 2002), and modulation of toxicity (Hold et al., 2001; Uribe and Espejo, 2003). The mode of interaction varies from indirect effects, through release of organic exudates (Wang et al., 2012), to symbiosis, such as intracellular bacteria within Alexandrium cells (Palacios and Marı´n, 2008). During bloom development, algal–bacterial interactions may lead to selection of specific bacteria phylotypes in the phycosphere of Alexandrium cells (Hasegawa et al., 2007), showing their importance in microbial community dynamics in the field. The aim of the present work was to characterize the influence of bacteria on Alexandrium tamarense using culture experiments in controlled conditions. In addition to analyzing the gross effects of bacteria on growth and toxin production of A. tamarense, specific biological functions, including photosynthesis, inorganic carbon (C) uptake, and phagotrophy of algal cells, were examined. Most of the laboratory studies conducted on phytoplankton– bacteria interactions are dependent on the production and maintenance of axenic cultures. Comparison of axenic and xenic clones are generally done on short time scales, e.g. after a few days or weeks of culture maintenance in newly axenic medium (e.g. Uribe and Espejo, 2003) or after inoculation with bacteria (e.g. Grossart and Simon, 2007). To our best knowledge, our study is the first to analyze these processes over the longer time scales that may be required for the establishment of specific, functional algal– bacterial associations. Experiments were run using clonal cultures of Alexandrium tamarense that have been maintained under either axenic or bacterized conditions over several years. Effects of bacteria inoculation in axenic cultures were considered over a 15 month period. The results of this study provide new insights into the influence of bacteria on biological functions of phytoplankton cells, with implications for understanding phytoplankton dynamics and organic matter cycling. In addition, our results suggest that beneficial algal–bacterial associations can contribute to the success of toxic dinoflagellates in various environments. 2. Materials and methods 2.1. Cultures and maintenance Two clones of the same strain of Alexandrium tamarense, CCMP 1493 and CCMP 1598, were obtained from the National Center for Marine Algae and Microbiota (NCMA). CCMP 1493 was isolated from the China Sea in 1991 and has been maintained under bacterized conditions since then; an axenic clone of this strain, CCMP 1598, was created a few years later and has since been grown under axenic conditions. In our laboratory, stock cultures of the two clones were grown in f/2 medium (minus Si) (Guillard and Ryther, 1962), at 16 8C, under 100 mmol photons m2 s1, with a 12:12 h light:dark cycle. All culture transfers were conducted under sterile conditions, from an initial cell concentration of 200 cells mL1. Sterility of axenic cultures was periodically

determined by direct bacteria observations with DAPI (40 ,6diamidino-2-phenylindole) staining and epifluorescence microscopy (see details below). Cultures were grown without bubbling but with a gentle stirring every other day. 2.2. Experimental design Triplicate 1L batch cultures were used to characterize growth and toxin production of the two parent and donor clones, CCMP 1598 and 1493. Analyses were conducted during exponential growth, when cell density exceeded 2500 cells mL1. Characterization of net growth was based on measurements of growth rate and cell volume. Photosynthetic health and capacity were determined from measurement of cellular chlorophyll a content and efficiency of photosystem II (Fv/Fm ratio). Autotrophic growth characterization included inorganic C-uptake capacities, from measurements of 13C-uptake rates. Mixotrophy of Alexandrium tamarense cells was assayed from observations of digestive vacuoles. Toxin profiles were determined, taking into account high potency carbamate toxins (saxitoxin [STX], neosaxitoxin [NEO], gonyautoxins [GTX1-GTX6]), low potency N-sulfocarbamoyl toxins (C1–C4) and decarbamoyl analogues (dcSTX, dcGTX2, dcGTX3). The experiment started with the initiation of ‘‘re-xenic cultures’’, axenic cells of CCMP 1598 inoculated with bacteria collected from a culture of CCMP 1493. One culture of each parent or donor clone (2L) in exponential phase was used for the creation of three re-xenic subclones. The culture of CCMP 1493 was lysed by vortexing with 0.5 mm silica–zirconium beads (BioSpec Products, Inc.). The resulting lysate was filtered through a 20 mm Nitex mesh to remove intact dinoflagellate cells. The filtrate was then centrifuged for 5 min at 3000  g to remove large cell fragments, and finally filtered through a 5.0 mm nitex mesh to remove small cell fragments. Re-xenic cultures were created by adding the resulting bacterial filtrate (500 mL of filtrate of CCMP 1493 culture) to axenic cells of Alexandrium tamarense (500 mL of CCMP 1598 culture). Re-xenic subclones were maintained under the same conditions as the parent and donor clones of A. tamarense for 15 months, by transferring cells to fresh f/2 medium when cultures reached late-log phase and without bubbling but a gentle stirring every other day. Re-xenic cultures were grown in 1L of medium in Fernbach flasks. Only one 1-L flask was simultaneously maintained for each re-xenic subclone. Cell density of A. tamarense was monitored during growth of each subculture, to determine growth rate in exponential phase and maximal cell density at plateau. When it came time to make a larger set of measurements, three replicated flasks were created for each subclone: one culture was used for maintenance of the subclone and two cultures were sacrificed for measurements. One of the three subclones was lost in less than a month, despite our best efforts to recover it. The other two subclones are designated as Re-xenic 1 and 2 in the present study. For both Rexenic 1 and 2, three replicate cultures were created after nine months and 15 months of culture maintenance and, each time, two of the replicated flasks were used for characterization of their growth and toxin production. The same parameters were monitored in the Re-xenic and parent and donor clones (CCMP 1598 and 1493). These measurements were done during the exponential phase of growth, on cultures showing a cell density higher than 2500 cells mL1. 2.3. Microscopic observations Cell abundances were determined in triplicate during the exponential growth of Alexandrium tamarense cultures, from samples preserved in Lugol’s iodine. For each sample, a minimum

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of 600 cells were counted under the microscope using a Sedgewick Rafter counting chamber. Cell concentrations were used to calculate net exponential growth rate (m, d1) as defined by Guillard (1973), using the following formula:

m¼

lnðC 2 ÞlnðC 1 Þ t 2 t 1

where m is the growth rate (d1), C1 and C2 are the cell concentrations at time 1 (t1, d) and time 2 (t2, d), respectively. Additional samples fixed in Lugol’s iodine were observed under 400 magnification in order to measure cell size of Alexandrium tamarense. Cell volume was calculated from measurements of two perpendicular cell diameters, one along the cingulum and another along the cell’s sagittal plane. The following equation was used for cell volume calculation, using an ellipsoid model for an A. tamarense cell: V¼



  2   4p D1 D2   3 2 2

where V is the cell volume (mm3), D1 is the diameter of the cingulum (mm) and D2 is the diameter of the cell’s sagittal plane (mm). For each culture condition, 360 cells were measured: 120 cells per replicated culture of parent and donor clones (CCMP 1598 and 1493, triplicate cultures) and 180 cells per culture of re-xenic subclones (duplicate cultures). The presence of bacteria, free in the culture medium or attached to Alexandrium tamarense cells, was microscopically determined from DAPI labeling of cell suspensions. One drop of formaldehyde was added to 1.5 mL of culture samples in order to facilitate dye penetration through thecal plates and membranes of A. tamarense cells. Five minutes after formaldehyde addition, DAPI (Invitrogen) was added at a final concentration of 2 mg mL1. Ten minutes incubations were performed in the dark at room temperature (RT) and stopped by filtration onto 0.2 mm dark filters (Whatman). Collected cells were washed several times with PBS. The blue staining of the nucleus by DAPI was examined under an epifluorescent microscope in order to reveal any bacteria in A. tamarense cultures. Digestive vacuoles in Alexandrium tamarense cells were observed using the acidotropic probe LysoSensor blue DND-167 (Molecular Probes, Life Technologies). This probe is almost nonfluorescent except when inside acidic organelles. Incubations were started by addition of the fluorescent probe at a final concentration of 5 mM, in 1 mL culture samples. Algal cells were visualized after 1 h of dark incubation at RT, using an epifluorescent microscope. 2.4. Indicators of photosynthetic health and capacity Culture samples (10–20 ml in triplicate) for the determination of cellular chlorophyll a content were filtered onto Whatman GF/C filters and extracted in 100% methanol for 24 h at 4 8C. The fluorescence of the extracted pigments was measured with a Turner Designs 10-AU fluorometer, and chlorophyll a concentrations were calculated using a calibration curve and correction for pheopigments according to Arar and Collins (1997) and Arar (1997). The fluorescence-based maximum quantum yield of photosystem II (Fv/Fm) was determined from the initial (Fo) and the maximum (Fm) in vivo fluorescence of culture samples dark adapted for 30 min. Fo and Fm corresponded to fluorescence measurements done before and 30 s after the addition of 3-(3,4dichlorophenyl)-1,1-dimethylurea (DCMU) (Sigma–Aldrich), respectively. These measurements were performed on 3 mL samples in triplicate, using a Turner Designs 10-AU fluorometer and a final DCMU concentration of 20 mM. Fv/Fm ratios were calculated as (Fm  Fo)/Fm (Vincent, 1980).

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2.5. C-uptake rate Incubations were performed using 40 mL aliquots of algal culture, in triplicate. Samples were spiked with a solution of 13Clabeled bicarbonate (NaH13CO3) at 78 mM-C and incubated for 1 h, under initial culture conditions. At the end of the incubation, samples were filtered through precombusted (4 h at 450 8C) A/E filters (Gelman Sciences). Filters were dried overnight and analyzed on an Integra CN elemental analysis-mass spectrometry system (PDZ Europa) for measurements of particulate carbon and 13 12 C/ C isotopic ratio. Particulate carbon values were standardized by cell density in order to obtain values of cellular C quota (Qc). Measurements of 13C enrichments were converted to net uptake rates of H13CO3 according to Collos (1987). 2.6. Toxin composition Duplicate 15 mL culture samples were transferred into centrifuge tubes, and Alexandrium tamarense cells were pelleted by centrifugation (2000  g, 5 min, RT). Cells were resuspended in 1 mL of 0.05 M acetic acid and broken by sonication. Cell lysates were frozen (20 8C) until analysis. Prior to HPLC analysis for saxitoxin and its derivatives, the samples were thawed, mixed by repetitive pipetting and centrifuged at 3000  g for 5 min. Aliquots (200 mL) of the sample extract supernatant were loaded into autosampler vials. Toxin analyses were carried out using a modification of the Oshima et al. (1989) post-column derivatization HPLC method (Anderson et al., 1994) with the following changes: 10 mL samples were injected using a Waters 717 autosampler onto either an Inertsil C8 (saxitoxin and C toxins) or an Inertsil ODS-4 (gonyautoxins) 4.6 mm  150 mm HPLC column coupled to its corresponding guard column. A Shimadzu RF-20A fluorescence detector captured the derivatized post-column signal and Millennium32 software (Waters Corporation, Milford, MA) was used for was data acquisition and analysis. Peak areas of the samples were quantified with certified reference standard solutions purchased from the National Research Council Canada (NRC – Halifax, Nova Scotia, Canada), containing toxins C1, C2, GTX1–5, dcGTX2, 3, NEO, dcSTX and STX. Toxicities (fg STX equiv cell1) were calculated from molar composition data using individual potencies in mg STX equiv mmol1 provided by the NRC as follows: C1, 2.61; C2, 41.6; GTX1, 429.4; GTX2, 155.2; GTX3, 275.6; GTX4, 313.7; GTX5, 27.8; dcGTX2, 66.5; dcGTX3, 162.7; NEO 399.3; dcSTX, 221.7; STX, 432.0. 2.7. Statistics Student’s t-test was performed to compare characteristics of parent clones. Associated results are expressed as mean  SD. Nonlinear regressions of cell size data were used to analyze their distribution. Statistical analyses were performed using Statgraphics Plus 5 (Manucistics Inc.). 3. Results The presence of bacteria in the cultures was verified from direct observations of cell suspensions stained with DAPI. No bacteria was observed in the axenic strain cultures (CCMP 1598) (Fig. 1A). The monitoring of the bacterized donor clone cultures as well as the rexenic subclones during the experiment revealed the presence of rodshaped bacteria (Fig. 1B). 3.1. Growth characteristics For parent and donor clones, cultures maintained under axenic conditions (CCMP 1598) had a significantly (t-test, p < 0.05) lower growth rate in exponential phase (0.22  0.01 d1) than cultures

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Fig. 1. Microscopic observations of cell suspensions from Alexandrium tamarense cultures after DAPI staining. Algal cells and bacteria were labeled with DAPI and exposed to UV light in order to reveal the blue staining of nucleus. Cell suspensions were sampled from axenic cultures (CCMP 1598) (A) and bacterized cultures (donor clone CCMP 1493) (B).

maintained with bacteria (CCMP 1493; 0.33  0.03 d1) (Fig. 2). For re-xenic subclones, growth rate of Re-xenic 1 was low (0.15  0.03 d1) during the 15 months of culture maintenance. Conversely, growth rate of Re-xenic 2 varied: after remaining at a low level during the first three months (0.15  0.01 d1), growth rate in exponential phase gradually increased, then stabilized after eight months of culture maintenance at 0.32  0.03 d1 (Fig. 2). After three months of subculturing, when growth rate started to increase, maximal cell density of Re-xenic 2 cultures suddenly shifted from values lower than 4000 cells mL1 to values ranging from 9000 to 12,000 cells mL1 (data not shown). 3.2. Cell size Cell size patterns from microscopic observations (Fig. 3) were better represented by a Poisson distribution (r2 ranging from 0.78 to 0.99) than a binomial one (r2 ranging from 0.66 to 0.99). Parent and donor strains showed large differences in cell size during exponential growth. The axenic clone had a lower average cell volume (10.0  0.7  103 mm3, Fig. 3A) than the xenic clone

(17.4  0.9  103 mm3, Fig. 3B). Nine months after bacterial addition, the pattern obtained for Re-xenic 1 was very similar to that of the axenic parent clone with an average cellular volume of 9.9  103 mm3 (Fig. 3C). In contrast, Re-xenic 2 showed a pattern close to that of the xenic donor clone, with an average cellular volume of 16.7  103 mm3 (Fig. 3D). After 15 months (Fig. 3E and F), the average cellular volume of Re-xenic 1 cultures increased to 13.8  103 mm3, when the cellular volume estimated for Re-xenic 2 was 16.4  103 mm3. Estimations of carbon quota per cell (Qc, Fig. 4A) gave additional information about cell size variations of Alexandrium tamarense during the experiment. In accordance with microscopic observations, Qc values for the axenic clone were significantly lower (t-test, p < 0.01) than those of the bacterized parent clone, with respective average values of 993  97 pg cell1 and 1778  143 pg cell1 obtained from replicate cultures. There was no difference between the two parent and donor clones when carbon quota was expressed per unit of cell volume; axenic and xenic cultures showed similar levels of cellular carbon density, with respective values of 0.099  0.003 pg mm3 and 0.103  0.013 pg mm3. Nine months after bacteria addition, the two re-xenic subclones showed strong differences in terms of Qc: Re-xenic 1 was similar to the axenic parent clone, while Re-xenic 2 was equivalent to the bacterized donor clone (Fig. 4A). During the next six months, a net increase in Qc values was observed for Re-xenic 1, so both the Re-xenic 1 and 2 subclones showed Qc values similar to that of the bacterized donor clone after 15 months. 3.3. Inorganic C-uptake and photosynthetic parameters

Fig. 2. Estimations of growth rate for exponentially growing cultures of Alexandrium tamarense. Data points show average values from replicate cultures. For the axenic parent clone and the xenic donor clone, data were compiled from three replicate cultures and vertical lines indicate the associated standard deviations. For the rexenic subclones (see Section 2.2 for identification), data were estimated from two replicate cultures over the course of the experiment and the dotted and solid arrows indicate average values obtained after nine months and 15 months of culture maintenance, respectively. White diamonds, grey diamonds and black diamonds indicate values obtained during months 0–5, 6–10 and 11–15, respectively.

The 13C-uptake rate was slightly lower for the axenic parent clone (CCMP 1598; 0.018  0.004 h1) compared to the bacterized one (CCMP 1493; 0.022  0.003 h1), but this difference was not statistically significant (Fig. 4B). Low 13C-uptake rates were measured for Re-xenic 1 over the course of the experiment. The uptake rate increased, however, from a value 10 times lower (0.002 h1, after nine months) to a value two times lower (0.009 h1, after 15 months) than axenic C-uptake rates (Fig. 4B). For the Re-xenic 2 subclone, 13C-uptake rate slightly increased during the course of the experiment, remaining in the range of values obtained for the two parent clones. With regards to photosynthetic parameters, similar patterns were observed for chlorophyll a content (Fig. 4C) and Fv/Fm ratio (Fig. 4D). The axenic strain showed significantly (t-test, p < 0.001) lower values for both measures than the bacterized donor clone. During maintenance of re-xenic cultures, values for Re-xenic 1 after nine months were lower than estimations characterizing the axenic parent strain. After an increase over the next six months, values observed for Re-xenic 1 were intermediate between those of the parent and donor clones. For Re-xenic 2, an increase in

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Fig. 3. Cell size distribution for exponentially growing cultures of Alexandrium tamarense. Cell volume was determined from microscopic observations for the axenic parent clone (A) and for the bacterized donor clone (B). Similar measurements were conducted for the re-xenic subclones (see Section 2.2 for identification) after nine months (Rexenic 1 (C) and Re-xenic 2 (D)) and after 15 months (Re-xenic 1 (E) and Re-xenic 2 (F)) of culture maintenance. A solid arrow indicates the average cell volume for each pattern.

chlorophyll a cellular content and Fv/Fm ratio was observed over time, and levels at the end of the experiment were close to those of the bacterized donor clone. 3.4. Characterization of acidic vacuoles Acidic vacuoles were observed during exponential growth of the xenic donor clone, as well as for both re-xenic subclones

(Fig. 5). These acidic organelles were small, no more than 4.5 mm in diameter. In most cells, acidic vacuoles accumulated on one side of the cell, often close to the plasma membrane. No cell maintained under axenic conditions showed such acidic vacuoles in their cytoplasm. The only fluorescent signal visible on axenic cells was the result of scintillons located on the cell membrane (Fig. 5A and B). Scintillons were easy to discriminate from acidic vacuoles: their blue fluorescence signal corresponded to small dots and was short-lived,

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Fig. 4. Characterization of carbon fluxes and photosynthetic parameters in exponentially growing cultures of Alexandrium tamarense. Parent and donor clones, axenic and bacterized, and re-xenic subclones 1 and 2 (see Section 2.2 for identification) were characterized according to variations in carbon quota (A), inorganic C-uptake (B), chlorophyll a content per unit of cell volume (C) and Fv/Fm ratio (D). Data points show average values obtained from replicate cultures. For the parent clones, vertical lines (delimited by bars) indicate standard deviations from three replicate cultures. For the re-xenic subclones, vertical lines represent the range of the data corresponding to two replicate cultures.

disappearing in few seconds. The Lysosensor Blue signal was more stable and showed acidic organelles located inside the cell (Fig. 5C–F). 3.5. Toxin profiles The two parent and donor clones, axenic and bacterized, exhibited similar toxin profiles. Although epimers were analytically separated, toxin profiles are presented as epimeric pairs (C1/ 2, GTX1/4) in Fig. 6 due to facile interconversion resulting from thermodynamic equilibrium. Profiles of parent and donor strains were characterized by high proportions of the N-sulfocarbamoyl toxin pair C1/2 (C2 representing 82  1 mol%), with lesser proportions of the decarbamoyl toxin dcSTX (10  1%), and traces (1– 2 mol%) of saxitoxin (STX), neosaxitoxin (neoSTX) and gonyautoxins GTX1/4 and GTX5. After bacterial addition, toxin profiles evolved similarly in both Re-xenic 1 and 2: profiles after nine months were characterized by a strong signature for dcSTX (28  3 mol% and 40  2 mol%, respectively), in addition to C1/2 (64  4 mol% and 49  2 mol%, respectively). Toxin profiles of re-xenic cultures finally converged to that of parent strains, showing similar signatures to CCMP 1598 and CCMP 1493 after 15 months of maintenance (Fig. 6). For all conditions and cultures tested, C1/2 and dcSTX represented a cumulative percentage of 92  2% of the toxin profile.

In terms of toxicity per cell, no significant difference was observed between the two parent and donor clones (t-test), with values of 3.9 pg STX equiv cell1 (0.7) and 3.4 pg STX equiv cell1 (0.4) estimated for axenic and xenic cells, respectively. Changes in toxin profiles noted for the re-xenic subclones after 9 months of culture maintenance did not induce changes in cell toxicity: re-xenic cultures showed levels of toxicity ranging from 3.2 to 3.9 pg STX equiv cell1. 4. Discussion Among various reported modes of algal–bacterial interactions, bacteria have been shown to have both beneficial and detrimental effects on phytoplankton (Grossart, 1999; Danger et al., 2007). Comparing the two parent and donor clones of Alexandrium tamarense, axenic (CCMP 1598) and bacterized (CCMP 1493), the presence of bacteria displayed beneficial effects on algal growth, as cells maintained in algal–bacterial mixed cultures had higher growth rate and larger cell size than cells grown under axenic conditions. In previous studies, bacteria were implicated in the promotion or inhibition of growth of Alexandrium cells, and even cell lysis (Ferrier et al., 2002; Pokrzywinski et al., 2012; Wang et al., 2012). These interactions are complex, vary with nutrient conditions, and may involve cell-to-cell relationships (Amaro

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Fig. 5. Microscopic observations of acidic organelles in Alexandrium tamarense cells. Algal cells were labeled with LysoSensor blue. (A–E) Algal cells were exposed to UV light, lobes of chloroplasts are visible in red. (A and B) Axenic cells (strain CCMP 1598) showing fluorescence of scintillons located on the cell membrane with a focus inside the cell (A) and on the cell membrane (B). A ring free of scintillons is visible, corresponding to the cingulum of the cell. (C and D) Xenic cells (strain CCMP 1493) showing fluorescence of acidic vacuoles. (E and F) Cell from a re-xenic subclone showing acidic vacuoles under UV light (E) or regular light (F). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

et al., 2005; Wang et al., 2010). For example, in the study of Amaro et al. (2005), bacteria isolated from A. catenella cultures released algalytic compounds when grown in high-nutrient media, but recovered symbiotic characteristics when grown with A. catenella cells. In the present work, observations of acidic vesicles in all but axenic cultures suggest that A. tamarense cells were able to complement photosynthetic growth by acquisition of organic resources under bacterized conditions. Indeed, A. tamarense cells are able to use dissolved organic compounds (Burkholder et al., 2008) as well as ingestion of bacteria (Nygaard and Tobiesen, 1993) for growth. The stimulation of algal growth by bacteria observed from present results may therefore be partly explained by trophic relationships associated with mixotrophic abilities of A. tamarense. Bacteria also appear to stimulate the photosynthetic machinery of Alexandrium tamarense cells: the bacterized donor clone showed a higher Fv/Fm ratio and chlorophyll a content than the axenic parent clone. Concerning pigment content, biophysical constraints associated with cell size would predict the reverse: larger

phytoplankton cells absorb fewer photons per pigment than smaller cells of the same shape, due to increased self-shading of pigment with increasing cell volume (Duysens, 1956; Kirk, 1994). As a result, larger cells tend to have lower intracellular pigment concentrations than smaller cells under any given irradiance regime. As bacterized cells were larger than axenic cells, measurements of pigment content in the present study show a net activation of chlorophyll a synthesis under bacterized conditions compared to axenic ones. These results are somewhat surprising because, as hypothesized by Moustafa et al. (2010), A. tamarense cells growing under axenic conditions need to optimize photosynthesis, their only energy supply. This promotion of photosynthesis in algal–bacterial mixed cultures suggests the existence of complex positive relationships between A. tamarense cells and bacteria that are broader than trophic relationships. The monitoring of re-xenic cultures shows that differences observed between the two parent and donor clones are due to presence/absence of bacteria, rejecting genetic drift associated

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Fig. 6. Toxin composition of exponentially growing cells of Alexandrium tamarense. Toxin profiles were characterized for the parent and donor clones, axenic and bacterized, and the re-xenic subclones 1 and 2 (see Section 2.2 for identification) after nine months and 15 months of culture maintenance. The different toxins are represented as follow: C1/2, grey dots; dcSTX, black filling; STX, diagonal bars; GTX1/4, white filling; GTX5, crosses; NEO, vertical bars; other toxins are pooled and shown with grey filling.

with long-term culture maintenance as a potential factor. These rexenic cultures consisted of algal cells from the axenic clone CCMP 1598 that were mixed with bacteria isolated from cultures of the xenic clone CCMP 1493. Cells first experienced a period of growth inhibition: decreases in growth rate and maximal cell density were observed in exponential and stationary phase, respectively. One subclone was even lost during the first month, when stress induced by modification of culture conditions may have facilitated overgrowth of bacteria or the proliferation of viruses. Finally, characteristics of re-xenic cultures converged towards those of the bacterized donor clone over an extended period. Growth characteristics of Re-xenic 2 (including growth rate, cell size and photosynthetic parameters) were similar to the bacterized donor strain after 15 months. Re-xenic 1 experienced a stronger and longer growth inhibition period: after nine months of maintenance, estimations of growth rate, C-uptake and photosynthetic parameters were lower than characteristics of the axenic parent clone, by up to 10-fold. This long inhibition period could result from a high abundance of free-living bacteria that were deleterious to algal growth. Indeed, bacteria can negatively affect growth of microalgae by competing for nutrients (Danger et al., 2007) and releasing algicidal compounds (Amaro et al., 2005; Wang et al., 2010; Pokrzywinski et al., 2012). However, even if rexenic subclones followed different paths on a short-term point-ofview, they all initiated a ‘‘recovery’’ period over 15 months and showed a net convergence towards characteristics of the bacterized donor strain. These patterns show that the same community assembly processes finally acted in all bacterized clones maintained in the present study. These processes could include a selective pressure and facilitated mutualistic relationships between Alexandrium tamarense cells and bacteria, promoting the establishment of functioning communities. Even if the complexity of the algal–bacterial relationship cannot be fully characterized from present results, some hypotheses can be defined, in particular regarding the role of organic exudates. First of all, the promotion of cell division, enlargement and photosynthesis of Alexandrium tamarense cells in bacterized cultures may have been induced by the synthesis and release of

phytohormones, such as auxins and cytokinins, by bacteria (Maruyama et al., 1986; Tate et al., 2013). Furthermore, the degradation of algal organic exudates by bacteria may have had indirect beneficial effects on phytoplankton growth, in addition to promotion of bacterial growth. Indeed, for Re-xenic 2, growth rate started to increase as soon as a sudden increase in maximal cell density at stationary phase was observed. Culture senescence is induced by either nutrient deprivation or accumulation of excreted deleterious compounds in culture medium (Sciandra and Ramani, 1994). Under f/2 culture conditions, such a sudden shift in maximal density of phytoplankton may be due to a change in bacterial community structure during sub-culturing: an increase in the degradation rate of organic exudates from algal cells could have delayed negative feedback due to their accumulation in culture medium. This hypothesis is supported by the fact that growth of some bacterial strains can be supported by organic matter derived from A. tamarense cells at the end of exponential growth (Simon et al., 2002). Modification of algal growth by organic exudates is probably accentuated in enclosed systems, such as culture flasks, compared to the natural environment. However, bacterial communities, in particular located in the phycosphere of algal cells (Hasegawa et al., 2007), may similarly participate in bloom development and maintenance at high algal cell concentrations. In addition to provide new insights into the influence of bacteria on phytoplankton dynamics and organic matter cycling, the present study considers impacts of algal–bacterial interactions on toxin synthesis by microalgal cells. Previous investigations of PST variation among Alexandrium species and populations have indicated that toxin profiles are genetically determined (Ishida et al., 1998) and stable enough to serve as a phenotypic marker (Cembella, 1998; Anderson et al., 1994; Alpermann et al., 2010). In the present study, axenic and xenic parent clones of Alexandrium tamarense showed similar toxin profiles, characterized by the predominance of C1/2 and dcSTX toxin types. However, transitory modifications of the toxin profile of A. tamarense cells were observed over a 15-month period, during the monitoring of rexenic cultures. Laboratory studies have shown fluctuations in toxin composition of Alexandrium cells in response to changes in physico-chemical conditions (Anderson et al., 1990; Wang et al., 2005; Etheridge and Roesler, 2005) or sudden modifications of cooccurring bacteria (Hold et al., 2001; Uribe and Espejo, 2003). However, dramatic changes in toxin composition are rare and usually reflect a simplification of the toxin profile over successive subcultures (Wang et al., 2005; Cho et al., 2008). In the present study, we observed compensatory variations between the two predominant toxins, dcSTX and C1/2, where pathways that connect these molecules involve several enzymatic steps (Fig. 7). dcSTX is sometimes called ‘‘decarbamoyl derivative of saxitoxin’’, and Kellmann et al. (2008) proposed two scenarios for its production in cyanobacteria: (i) dcSTX could be produced from hydrolytic cleavage of STX or (ii) dcSTX could be carbamoylated to form STX, as shown on Fig. 7. In other words, dcSTX could either be a derivative of or precursor to STX, and the actual sequence of final reactions leading to the complete STX molecule remains uncertain (Fig. 7). Results obtained from the monitoring of re-xenic cultures in the present study give new clues for the understanding of this biosynthetic pathway, and of the role of dcSTX in particular. Based on the current knowledge of PST biosynthesis in cyanobacteria and toxic dinoflagellates, biosynthetic pathways can be proposed for the formation of the different STX analogues observed in the present study (Fig. 7). In crude enzyme extracts from Gymnodinium catenatum, STX is converted into GTX2/3 then C1/2 (Yoshida et al., 2002). In Alexandrium tamarense, GTX2/3 is sulfated or oxidized to form C1/2 and GTX1/4, respectively (Wang et al., 2007; Oshima, 1995). Present results support the existence of an additional upstream step: the conversion of dcSTX into STX (Fig. 7). The

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Fig. 7. Proposed PSTs biosynthetic pathways in Alexandrium tamarense. Figure modified from Pearson et al. (2010). Dashed arrows represent pathways defined in cyanobacteria (Kellmann et al., 2008). Solid arrows indicate pathways identified in toxic dinoflagellates (G. catenatum, A. catenella or A. tamarense) (Oshima, 1995; Ishida et al., 1998; Sako et al., 2001; Yoshida et al., 1998; Yoshida et al., 2002; Wang et al., 2007).

transient modifications in toxin profiles observed during maintenance of re-xenic cultures could be explained by decreased enzymatic conversion of dcSTX to STX, leading to an accumulation of dcSTX to the detriment of C1/2 production. We propose that this scenario is more likely than an activation of a degradation cascade from C1/2 into dcSTX, supporting the idea of dcSTX being a precursor of STX and other STX analogues.

Whilst previous works might suggest a potential influence of bacteria on toxicity of Alexandrium cells, the present study does not support this hypothesis. Hold et al. (2001) and Uribe and Espejo (2003) investigated the potential influence of algal–bacterial interactions on PST synthesis. These studies reported decreases in toxicity of Alexandrium cells after removal of bacteria from nonaxenic cultures. These variations were concomitant with

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modifications in toxin profiles that consisted in (i) a reorientation of the last steps in synthesis of STX analogues for Alexandrium tamarense, visible through an increase in synthesis of C1/2 to the detriment of GTX1/4 (Hold et al., 2001), (ii) a halt in toxin biosynthesis pathways for Alexandrium catenella, with an overall decrease in the amount of STX and gonyautoxins per cell (Uribe and Espejo, 2003). In these previous studies, analyses were performed only after few successive rounds of subculturing or dilutions. According to our longer-term observations, modifications of toxin profiles in response to addition or removal of bacteria are more likely a transient response to stress that temporarily decreases energy input into toxin production. In conclusion, mutualistic interactions between microalgal cells and bacteria have beneficial effects on algal growth through different biological functions, including nutrition of algal cells and photosynthesis. These algal–bacterial associations could have important effects on bloom dynamics and cycling of organic matter in the ecosystem. Such associations could be a key factor explaining the success of toxic dinoflagellates in various environments, despite their poor ability to use inorganic nutrients. Acknowledgments We would like to thank Dr. D.M. Anderson and D. Kulis for providing Alexandrium tamarense cultures and for processing toxin samples. This article is a result of research funded by the National Oceanic and Atmospheric Administration Center for Sponsored Coastal Ocean Research under award no. NA09NOS4780166 to the University of Texas Marine Science Institute and by the ECOHAB program contribution no. 839.[SS] References Adachi, M., Kanno, T., Matsubara, T., Nishijima, T., Itakura, S., Yamaguchi, M., 1999. Promotion of cyst formation in the toxic dinoflagellate Alexandrium (Dinophyceae) by natural bacterial assemblages from Hiroshima Bay, Japan. Mar. Ecol. Prog. Ser. 191, 175–185. Adachi, M., Matsubara, T., Okamoto, R., Nishijima, T., Itakura, S., Yamguchi, M., 2002. Inhibition of cyst formation in the toxic dinoflagellate Alexandrium (Dinophyceae) by bacteria from Hiroshima Bay, Japan. Aquat. Microbiol. Ecol. 26, 223–233. Alpermann, T.J., Tillmann, U., Beszteri, B., Cembella, A.D., John, U., 2010. Phenotypic variation and genotypic diversity in a planktonic population of the toxigenic marine dinoflagellate Alexandrium tamarense (Dinophyceae). J. Phycol. 46 (1), 18–32. Amaro, A.M., Fuentes, M.S., Ogalde, S.R., Venegas, J.A., Suarez-Isla, B.A., 2005. Identification and characterization of potentially algal-lytic marine bacteria strongly associated with the toxic dinoflagellate Alexandrium catenella. J. Eukaryot. Microbiol. 52 (3), 191–200. Amin, S.A., Parker, M.S., Armbrust, E.V., 2012. Interactions between diatoms and bacteria. Microbiol. Mol. Biol. Rev. 76 (3), 667–684. Anderson, D.M., 2009. Approaches to monitoring, control and management of harmful algal blooms (HABs). Ocean Coast. Manage. 52, 342–347. Anderson, D.M., Kulis, D.M., Sullivan, J.J., Hall, S., 1990. Toxin composition variations of the dinoflagellate Alexandrium fundyense. Toxicon 28, 885–893. Anderson, D.M., Kulis, D.M., Doucette, G.J., Gallagher, J.C., Balech, E., 1994. Biogeography of toxic dinoflagellates in the genus Alexandrium from the northeastern United States and Canada. Mar. Biol. 120, 467–478. Arar, E.J., 1997. Method 446.0. Revision 1.2. In Vitro Determination of Chlorophylls a, b, c1+c2 and Pheopigments in Marine and Freshwater Algae by Visible Spectrophotometry. United States Environmental Protection Agency, Office of Research and Development, National Exposure Research Laboratory, Cincinnati, OH. , http://www.epa.gov/microbes/documents/m446_0.pdf. Arar, E.J., Collins, G.B., 1997. Method 445.0. Revision 1.2. In Vitro Determination of Chlorophyll a and Pheophytin a in Marine and Freshwater Algae by Fluorescence. United States Environmental Protection Agency, Office of Research and Development, National Exposure Research Laboratory, Cincinnati, OH. , http:// www.epa.gov/microbes/documents/m445_0.pdf. Burkholder, J.M., Glibert, P.M., Skelton, H.M., 2008. Mixotrophy, a major mode of nutrition for harmful algal species in eutrophic waters. Harmful Algae 8, 77–93. Cembella, A.D., 1998. Ecophysiology and metabolism of paralytic shellfish toxins in marine microalgae. In: Anderson, D.M., Cembella, A.D., Hallegraeff, G.M. (Eds.), Physiological Ecology of Harmful Algal Blooms. Volume NATO-Advanced Study Institute Series, vol. 41. Springer-Verlag, Berlin, pp. 381–403. Cho, Y., Hiramatsu, K., Ogawa, M., Omura, T., Ishimaru, T., Oshima, Y., 2008. Nontoxic and toxic subclones obtained from a toxic clonal culture of Alexandrium

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