Introduction

It is widely accepted that harmful algal blooms (HABs) are increasing (Hallegraeff 1993; Van Dolah 2000; Anderson et al. 2002), and that certain HAB species are expanding their biogeographic ranges (e.g., Mcleod et al. 2012). Anthropogenic eutrophication (Anderson et al. 2002, 2008; Heisler et al. 2008), ballast water transportation (Hallegraeff 1993; Bolch and de Salas 2007; Smayda 2007) and potentially climate change (Moore et al. 2008; Hallegraeff 2010; Mcleod et al. 2012) are all thought to contribute to a continued global expansion of HABs. Irrespective of the proximal causes, these blooms will continue to increase environmental, and economic costs which are currently estimated at US$82 million and US$813 million annually in the US and European Union, respectively (Hoagland and Scatasta 2006), thus highlighting the need for effective management solutions. HAB management is broadly segregated by whether methods target solutions for the present or the future, and then by whether the actions are direct or indirect. Actions that aim to prevent or reduce HAB occurrences in the future are termed “prevention” approaches. The most common form of prevention aims to restrict nutrient loads to aquatic systems to reduce anthropogenic eutrophication. However, in estuarine and marine waters, successful prevention of HABs by nutrient management may be less straightforward than for freshwater HABs because of a wider variety of HAB species and environmental conditions (Smith and Schindler 2009). Furthermore, Heisler et al. (2008) note that enacting effective policies for nutrient restrictions is quite challenging and costly and would require adaptive management to account for imperfect understanding of HAB dynamics (see also Zingone and Enevoldsen 2000). Likewise, some HABs are not linked to anthropogenic eutrophication and therefore would not respond to restricted nutrient releases (e.g., Bachmann et al. 2003; Anderson et al. 2008). Recognizing these difficulties, it is imperative to pursue more immediate strategies as well. In this regard, actions that aim to minimize the impacts of HABs (e.g., aquaculture losses, human illnesses) in the present or immediate future, are termed “mitigation” approaches, and are not mutually exclusive from yielding “prevention”. Mitigation approaches are further sub-divided into either direct actions that reduce or contain a HAB population, termed “control”, or indirect/passive actions to ameliorate HAB impacts, which we term “non-control mitigation”. Non-control mitigation, such as moving finfish cages to avoid fish kills (Rensel and Whyte 2003), and monitoring to prevent the harvesting of toxic organisms (Zingone and Enevoldsen 2000; Trainer and Suddleson 2005) has proved successful. However, non-control mitigation is sometimes impractical and costly, or too passive to ameliorate certain impacts while a bloom still exists (e.g., aerosolized toxins or toxicity to wild animals). Consequently, under these circumstances, control mitigation may be more effective.

Control mitigation encompasses many different chemical, physical and biological strategies (Kim 2006; Sengco 2009). Physical controls, such as mixing (accomplished by bubbling) alleviated Microcystis sp. blooms in a lake (Jungo et al. 2001), and clay flocculation has been successful at removing a wide variety of HAB species (Sengco and Anderson 2004). Chemical controls, such as copper sulfate additions (Rounsefell and Evans 1958), and hydrogen peroxide (Matthijs et al. 2012) have also been effective. Often however, broad toxicity is cited as an obstacle to real-world use (Boylan and Morris 2003; HAB RDDTT 2008). Recent discoveries of less broadly toxic chemicals have shown promise, such as l-lysine which has been used against Microcystis spp. (Takamura et al. 2004) and thiazolidinediones (Kim et al. 2010). In addition, a number of chemicals effective against HAB species have been derived from biological origins, such as unknown compounds in barley straw extracts (Terlizzi et al. 2002), polyphenols from tea leaf extracts (Lu et al. 2013), rhamnolipid biosurfactants from Pseudomonas aeruginosa (Wang et al. 2005) and prodigiosin from the bacterium Hahella chejuensis (Jeong et al. 2005). Lastly, biological control techniques have been successful in controlling certain HAB species in controlled incubations, primarily via algicidal bacteria (Kim et al. 2008; Kang et al. 2011; Paul and Pohnert 2012) and micrograzers (Jeong et al. 2008). To date, no single approach has been truly championed, and the unique advantages of these many strategies have warranted continued examination.

Hare et al. (2005) described a bacterium, Shewanella sp. IRI-160, that had a growth inhibiting effect on three dinoflagellates (Prorocentrum minimum, Pfiesteria piscicida, and Gyrodinium uncatenum), but had no growth inhibiting effects on four non-dinoflagellate species. A recent investigation by Pokrzywinski et al. (2012) found that the algicidal activity of Shewanella sp. IRI 160 was due to a thermally stable, polar and water soluble compound or compounds secreted by the bacterium. The agent(s) accumulated in the medium, such that cell-free bacterial filtrates induced algicidal and growth-inhibitory effects, thereby negating the need to apply live bacteria. Furthermore, algicidal activity was significantly greater when applied to dinoflagellates in logarithmic growth, as compared to stationary or lag-phase cultures. The specific cellular targets, and mode of action of this secreted algicide is currently under investigation.

Testing of naturally derived algicides and allelochemicals in vitro has revealed that many act on specific components of the photosynthetic apparatus of susceptible algal cells. For example, free fatty acids disrupt membranes, resulting in the dissociation of phycobilins (but not integral chlorophylls) from the thylakoid membranes of cyanobacteria (Wu et al. 2006). Polyphenolic algicides produced by a macrophyte inhibit photosystem II (PSII) in Anabaena sp. by interfering with electron transport between the primary and secondary quinones Q A and Q B in the PSII reaction center (Leu et al. 2002). Likewise, the compounds Fischerellin A and Fischerellin B from the cyanobacterium Fischerella muscicola, target multiple sites on the electron transport chain surrounding PSII (Smith and Doan 1999), and Cyanobacterin derived from Scytonema hofmannii likely targets QA on the acceptor side of PSII (Gleason and Case 1986). Lastly, rhamnolipid biosurfactants from the bacterium Pseudomonas aeruginosa inactivated PSII in a range of HAB taxa (Gustafsson et al. 2009). Furthermore, Shi et al. (2009) found that algicidal agents produced in vivo were also capable of inhibiting the photosynthetic apparatus, showing that the algicidal bacterium Pseudomonas mendocina caused PSII inactivation in Aphanizomenon flos-aquae. In the current study, we explored whether the algicide IRI-160AA affects PSII function in a range of cultured dinoflagellates and report on the timing of membrane permeability in affected cells.

Methods

Bacterial filtrate preparation

Bacterial filtrates were prepared following Hare et al. (2005) and modifications detailed in Pokrzywinski et al. (2012). Briefly, Shewanella sp. IRI-160 was plated onto LM medium agar without antibiotics. Plates were incubated at room temperature until colony formation, whereupon a colony was transferred to 100 mL of liquid LM medium without antibiotics and incubated at 25 °C on an orbital shaker at 100 rpm for 18 h. Cultures were split into two, centrifuged, and washed twice in f/2 seawater algal medium (Guillard and Ryther 1962). The bacteria were resuspended in 80 mL fresh f/2, and incubated at 30 °C for 1 week. Finally, the culture was centrifuged, and the supernatant was filtered (0.2 μm) and stored at −80 °C until later use. The filtrate was thawed prior to use and then stored at 4 °C during subsequent applications. Following Pokrzywinski et al. (2012), we refer to this filtrate as IRI-160AA.

Algae species

Five algae species were tested for photochemical inhibition with the algicide: four dinoflagellates [Karlodinium veneficum (CCMP 2936), Gyrodinium instriatum (CCMP 2935), Prorocentrum minimum (CCMP 2233), Alexandrium tamarense (CCMP 1493)], and one cryptophyte, [Rhodomonas sp. (CCMP 757)]. Non-axenic batch cultures were grown in autoclaved 20 PSU f/2 medium at 24 °C, under 150 μmol photons m−2 s−1 on a 12:12 light/dark cycle provided by cool-white fluorescent lights. For all experiments, algae cultures were sampled during logarithmic growth, as this growth phase shows the greatest susceptibility to the algicidal and growth-inhibitory effects of IRI-160AA (Pokrzywinski et al. 2012).

Species testing

Tests comparing five species were carried out aseptically in autoclaved 15-mL glass test tubes, with independent replication. Treatments and controls were run in triplicate. Treatment cultures included addition of 4 % (v/v) IRI-160AA while controls included 4 % (v/v) f/2 medium. Logarithmic-stage cultures were inoculated mid light-cycle (of a 12:12 L/D) and were sampled for both photochemistry (see below) and cell density after 18 h of exposure and then in three subsequent 24 h intervals (=42, 66, 90 h). The 18-h sampling point was chosen because this was previously noted as a time of peak inhibition in other physiological variables (Pokrzywinski et al. submitted). Cells were fixed with Lugols Iodine solution, and later enumerated by light microscopy in a Neubauer Haemocytometer or Sedgwick-Rafter cell. Percent cell loss was calculated as: (number of cells lost/initial abundance) × 100.

Algal photochemistry

PSII function was assessed by fast repetition rate fluorometry (Kolber et al. 1998; FASTtracka II and FASTact system, Chelsea Instruments, UK). A series of 100 subsaturating LED “flashlets” at ∼1 μs intervals were applied to generate a single-turnover fluorescence transient. Fluorescence transients were used to evaluate three primary parameters: (1) the maximum quantum yield of PSII (F v/F m = F m − F o/F m), (2) the rate of primary quinone re-oxidation and electron transport out of PSII (τ; Q A- → Q A), and (3) the photochemical connectivity between PSII reaction centers (ρ) (Kolber et al. 1998; Cosgrove and Borowitzka 2010). The F v/F m ratio describes the maximum efficiency of PSII for converting absorbed light into a photochemical charge separation, and lower F v/F m values can be used to infer PSII inactivation (i.e., damage) or down-regulation of PSII reaction centers. Tau (τ) describes the time constant of electron transfer on the acceptor side of PSII, and specifically describes the time required to re-oxidize the reduced primary quinone (Q A-) measured in μs. PSII connectivity (ρ) describes the relative capacity to re-direct excitation energy from closed reaction centers, to nearby open PSII reaction centers.

Prior to fluorescence measurement, samples were dark-acclimated at room temperature for ≥20 min to re-open PSII reaction centers and relax non-photochemical quenching. The FRR sample chamber was maintained at 24 °C. The fluorescence measurement protocol consisted of three consecutive acquisitions, with each acquisition consisting of 40 repetitions of the following sequence: 100 flashlets at 1-μs intervals to reach saturation, and 50 flashlets at 49-μs intervals to record relaxation kinetics. Curve-fitting software provided with the instrument (FASTpro v3.0, Chelsea Instruments, UK) was used to derive F v/F m, τ, and ρ. All curve fits and fluorescence transients were manually inspected in real time.

Dose responses

The dose response of K. veneficum to IRI-160AA was assessed in triplicate at final concentrations of 2, 4, 6, 8, 12, and 16 % (v/v), alongside an 8 % (v/v) f/2 control. F v/F m and cell density were measured after 18 h of incubation. The control was limited to an 8 % concentration because a dilution series of f/2 concentrations was expected to yield identical results. F v/F m and cell density were converted to percent inhibition of the f/2 control ((treatment / average − control) × 100)). Prior to the conversion of cell density to percent inhibition, the average control cell densities were corrected to account for the differing dilutions used relative to the control using:

$$ \mathrm{Average}\kern0.5em 8\%\kern0.5em \mathrm{control}\kern0.5em \mathrm{cell}\kern0.5em \mathrm{density}\kern-12em -\left(\frac{\%\kern0.5em \mathrm{treatment}\kern0.5em \mathrm{concentration}-8}{100}\kern0.5em \times \mathrm{control}\kern0.5em \mathrm{cell}\kern0.5em \mathrm{density}\right) $$

The F v/F m ratio is unaffected by cell density and does not require this dilution correction.

In order to test whether IRI-160AA activity was affected by dinoflagellate cell density, a second experiment was conducted which compared IRI-160AA activity against dilutions of K. veneficum. IRI-160AA was added at a 4 % (v/v) final concentration, in triplicate to three dilutions of K. veneficum culture (no dilution, 1:3 dilution and 2:3 dilution in f/2 medium). Control cultures included addition of 4 % (v/v) f/2 medium to undiluted culture. Measurements of cell density and F v/F m were made after 18 h.

Membrane permeability

Outer-membrane permeability in relation to IRI-160AA activity was investigated in K. veneficum and G. instriatum with the fluorescent dye, SYTOX Green (Invitrogen, USA; SYTOX). SYTOX fluoresces brightly when bound to DNA but cannot pass through intact membranes, and can therefore be used to indicate the presence of dead cells in which the membrane has been compromised. Treatments included 8 % (v/v) IRI-160AA additions, and controls included 8 % (v/v) f/2. Subsamples were collected every 3 h for 24 h. SYTOX was added to subsamples in duplicate at a 1 μM final concentration and incubated for 90 m in the dark, and fluorescence was measured on a microplate reader with excitation and emission at 485 and 520 nm, respectively (FLUOstar Omega, BMG Labtech). The cellular origin of SYTOX fluorescence was verified using an epi-fluorescence microscope (Microphot-FXA, Nikon, Japan). Total SYTOX fluorescence values were blank corrected (f/2 medium + 1 μM SYTOX) and converted to SYTOX fluorescence per cell. The SYTOX fluorescence per cell of IRI-160AA treatments were converted to a percent of the average f/2 control fluorescence per cell (which did not vary significantly over 24 h, one-way ANOVA, n = 3, P > 0.05).

Dark incubation

To test how light might influence the susceptibility of K. veneficum and G. instriatum to IRI-160AA, we compared the effect of IRI-160AA incubations in the dark to IRI-160AA incubations in light/dark (L/D) cycles. Using cultures of K. veneficum and G. instriatum that were grown under 12:12 L/D cycles, two 4 % IRI-160AA treatments were tested. One of these treatments was transferred to continuous (24 h) darkness, and the other treatment was kept in the regular 12:12 light/dark cycle. Controls included addition of 4 % f/2, with one control transferred to continuous darkness, and the other kept in 12:12 light/dark cycles. Both treatments and both controls were performed in triplicate and transferred to their respective light/dark treatments immediately after IRI-160AA or f/2 addition, at approximately mid light-cycle. Cell density and photochemistry were measured at 0, 18, 42, 66, and 90 h after addition of IRI-160AA.

Statistical analyses

Statistical analyses were performed in Prism 5 (GraphPad Software Inc., USA). All experimental treatments tested were independent with small balanced sample sizes (n = 3). Different treatments were analyzed by two-way ANOVA with time and treatment as independent variables, and were followed by Bonferroni post hoc tests. The culture dilution experiment was tested by one-way ANOVA and the Tukey’s HSD post hoc test.

Results

Species comparison

All four species of dinoflagellates showed significant growth inhibition with IRI-160AA, whereas the cryptophyte, Rhodomonas sp., responded with an increased growth rate and a higher maximal abundance relative to f/2 controls (Table 1 and Fig. 1a–e). Despite all dinoflagellates showing susceptibility to IRI-160AA by growth inhibition, there were species-specific responses in the relative magnitude and timing of cell mortality, growth inhibition and recovery (Fig. 1a–d, Table 1), and photochemical inhibition and recovery (Figs. 1f–i and 2). G. instriatum and K. veneficum showed similar levels of susceptibility to the algicide (54 ± 19 % and 50 ± 20 % cell loss, respectively), with some recovery or stabilization in cell density occurring after 42 and 66 h respectively (Fig. 1, Table 1). Alexandrium tamarense exhibited a continuous decline in cell density before reaching a maximum cell loss of 7 % (±10 %), at 90 h (Fig. 1c, Table 1). P. minimum did not decline in abundance during the experiment, but did exhibit growth inhibition relative to the control (Fig. 1d, Table 1).

Table 1 Specific growth rates from Fig. 1 (algal species tested with 4 % IRI-160AA or 4 % f/2 medium). IRI-160AA growth rates and percent cell loss were calculated from T0 to the time point with the lowest cell number, except for P. minimum and Rhodomonas sp. which did not show a drop in cell density (N/a) and were calculated over 90 h. Percent cell loss was rounded to the nearest whole percentage with ±1 SD given in parentheses
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Fig. 1
figure 1

Species-specific responses in cell density (a–e) and F v/F m (f–j) among four dinoflagellates: G. instriatum (a, f), K. veneficum (b, g), A. tamarense (c, h), and P. minimum (d, i), and 1 cryptophyte, Rhodomonas sp. (e, j), after addition of 4 % IRI-160AA concentration (filled squares) compared to control cultures (open circles) after addition of 4 % f/2 medium. The species are ordered from highest to lowest response (left to right) and by relative plasma-membrane exposure, which reflects the change from naked to thecate dinoflagellates and then also by number of plates/valves (see “Discussion”). Asterisks indicate a significant difference as determined by two-way ANOVA and Bonferroni post tests (n = 3, P < 0.05). Error bars are ±1 SD of the mean

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Fig. 2
figure 2

Species-specific responses in the Q A- re-oxidation rate τ (a), and in PSII connectivity ρ (b), both expressed as relative change from the f/2 control, so that the 0 % horizontal lines represent the average f/2 control. Gi G. instriatum (open bars); Kv K. veneficum (gray bars); At A. tamarense (black bars); Pm P. minimum (striped bars); R sp. Rhodomonas sp. (fine checkered). Data are from the respective experiments shown in Fig. 1. Asterisks indicate a significant difference (n = 3, P < 0.05) determined by two-way ANOVA, and Bonferroni post tests. Error bars are ±1 SD of the mean

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G. instriatum, K. veneficum, and A. tamarense exhibited similar levels of PSII inactivation after exposure to IRI-160AA. Maximal inhibition occurred at 18 h for algicide-treated cultures of both K. veneficum and G. instriatum, and at 66 h in A. tamarense (Fig. 1f, g, h). Recovery from PSII inactivation in remaining cells in the treatment cultures occurred by 42 h in G. instriatum and by 66 h in K. veneficum (Fig. 1f, g), while A. tamarense showed negligible recovery between 66 and 90 h (Fig. 1h). Prorocentrum minimum showed no significant PSII inactivation (two-way ANOVA, P > 0.05, Fig. 1i) after exposure to IRI-160AA. Rhodomonas sp. treated with IRI-160AA exhibited no significant difference in F v/F m compared to the control (Fig. 1j).

Following the species’ responses in F v/F m, PSII re-oxidation (τ) in the algicide-treated samples took significantly longer than the f/2 controls in G. instriatum, K. veneficum, and A. tamarense (two-way ANOVA, P < 0.05) and was also observed in P. minimum (Fig. 2a). Temporally, τ was altered in G. instriatum and K. veneficum only after 18 h of exposure, and recovered to control levels (0 % horizontal line) by 42 h. In A. tamarense, τ remained significantly slower as compared to the controls until the end of the experiment (Fig. 2a), and in P. minimum τ was inhibited after 90 h only. Connectivity (ρ) was significantly lower only in K. veneficum and A. tamarense (two-way ANOVA, P < 0.05), and recovered by 42 h in K. veneficum, and did not recover in A. tamarense (Fig. 2b). Conversely, ρ was significantly higher relative to the control in P. minimum after 90 h when exposed to the algicide (two-way ANOVA, P < 0.05, Fig. 2b). In Rhodomonas sp. treated with IRI-160AA, τ was significantly faster at 42 and 90 h, and ρ was significantly lower than the control at 90 h.

Dose response

K. veneficum exhibited a typical dose response in photochemical inhibition and cell density with increasing IRI-160AA concentrations from 2 to 16 % (Fig. 3). The percent inhibition using F v/F m values was significantly lower than for cell density only at a 4 % concentration (two-way ANOVA, n = 3, P < 0.05). The average EC-50 calculated from the fitted dose response curves was 7.0 and 8.7 % (v/v) IRI-160AA when calculated using cell density and F v/F m respectively. Final IRI-160AA concentrations of 12 to 16 % resulted in near to 100 % photochemical and cellular inhibition relative to the control after 18 h. The time required for recovery in growth and F v/F m was also dose-dependent (i.e. longer at higher doses), as was relative membrane permeability as determined by SYTOX fluorescence (higher SYTOX fluorescence was noted at higher doses; data not shown). In contrast to the effects of IRI-160AA concentration, modifying algal cell density by dilution had no effect on the relative inhibition in photochemistry or cell density after 18 h exposure to IRI-160AA at 4 % (v/v) concentration (Fig. 3 inset). Cell densities and F v/F m in all IRI-160AA treatments were, however, significantly lower than the control in the algal dilution experiment (n = 3, one-way ANOVA and Tukey test, P < 0.05).

Fig. 3
figure 3

Dose response based on cell density (filled triangles) and F v/F m (open circles) in K. veneficum expressed as % inhibition relative to the f/2 control, after 18 h. Variable slope dose response curves are fitted to the data. Error bars are ±1 SD. Percent inhibition based on cell density and F v/F m are significantly different at the 4 % concentration (two-way ANOVA and Bonferroni post tests, n = 3, P < 0.05). The inset figure shows relative inhibition in F v/F m (filled bars) relative to the f/2 control and cell density (open bars) relative to initial densities in culture dilutions treated with 4 % IRI-160AA. Full no dilution, 2/3 33 % dilution, 1/3 66 % dilution. No significant differences were observed among dilutions for either cellular inhibition, or photochemical inhibition (one-way ANOVA and Tukey test). However, IRI-160AA treatments all differed significantly from f/2 controls (n = 3, P < 0.05)

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Membrane permeability

When photochemistry was followed with greater temporal resolution, K. veneficum exhibited moderate PSII inactivation after 18 h exposure to the algicide (Fig. 4a), whereas G. instriatum showed large and immediate (<3 h) PSII inactivation (Fig. 4b). Growth for K. veneficum appeared to slow by 6 h after addition of IRI-160AA and cell density was significantly lower than the controls between 18 and 24 h (Fig. 4c), whereas a large immediate decline in cell density occurred within the first 6 h of exposure to the algicide for G. instriatum (Fig. 4d). By 9 h, there was a significant increase in cell membrane permeability (detected by SYTOX green) in algicide-treated K. veneficum, which peaked abruptly after 24 h (Fig. 4e). In comparison, significant cell permeability was detected in G. instriatum immediately after IRI-160AA addition, which then peaked earlier near 12 h (Fig. 4f). For G. instriatum, the timing and relative changes in SYTOX fluorescence correlated well with the relative changes in F v/F m and cell density. In contrast, the main decline in F v/F m in K. veneficum at 18 h preceded the maximum permeability by 6 h, but did coincide with the commencing decline in cell density between 15 and 24 h.

Fig. 4
figure 4

Twenty-four-hour incubations with K. veneficum (left panes) and G. instriatum (right panes). Cultures were incubated with either 8 % v/v IRI-160AA (filled squares) or 8 % v/v f/2 medium (open circles). F v/F m (a, b), cell density (c, d) and SYTOX fluorescence (e, f; relative to the f/2 control) are presented. Grayed areas behind the data represent the dark period. Asterisks indicate time points with a significant difference, as determined by two-way ANOVA and Bonferroni post tests (P < 0.05, n = 3). Error bars represent ±1 SD

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Darkness

K. veneficum and G. instriatum showed different responses to darkness both in the presence and absence of IRI-160AA. Compared to light controls, dark K. veneficum controls showed photochemical inhibition and stunted growth by 42 h, and precipitous declines in both photochemistry and cell number by 90 h (Fig. 5a, b). Contrarily, dark G. instriatum controls showed a small but consistent down-regulation of PSII photochemistry after 18 h and showed inhibited growth from 42 h, but neither photochemistry nor cell density showed the precipitous declines by 90 h as in K. veneficum. Dark IRI-160AA treatment of K. veneficum induced greater PSII inactivation after 18 h (Fig. 5a), and a larger loss in cell density after 42 h (Fig. 5b) as compared to light IRI-160AA treatments. Neither cell density nor PSII inactivation recovered in K. veneficum treated with IRI-160AA and kept in continuous darkness. Contrary to the dark response of K. veneficum, G. instriatum showed no significant increase in cellular mortality in dark algicide applications relative to those kept in light/dark cycles (Fig. 5d). However, G. instriatum did show significantly more PSII inactivation by 42 h in dark IRI-160AA treatments, and like K. veneficum failed to fully recover from the initial PSII inactivation (Fig. 5c) in the dark IRI-160AA treatments.

Fig. 5
figure 5

Effects of shifting treated cultures into continuous darkness immediately upon addition of 4 % (v/v) IRI-160AA in K. veneficum (a, b) and G. instriatum (c, d). Light treatments are indicated by open symbols, dark treatments are indicated by filled symbols, circles indicate controls and squares are IRI-160AA treatments. Asterisks indicate a significant difference only between IRI-160AA light and IRI-160AA dark treatments (two-way ANOVA and Bonferroni post tests, P < 0.05, n = 3). Error bars are ±1 SD of the mean

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Discussion

A large proportion of algicides and allelochemicals derived from marine and freshwater microalgae appear to target competitor phytoplankton cell membranes, often simultaneously inactivating PSII and photosynthetic electron transport, and inhibiting growth (Smith and Doan 1999; Legrande et al. 2003 and references therein). Here, we have employed analysis of single-turnover chlorophyll fluorescence to evaluate PSII inhibition in dinoflagellates under algicidal treatment, using a previously described filtrate from the bacterium Shewanella sp. IRI-160 (Hare et al. 2005; Pokrzywinski et al. 2012).

General dynamics of photobiology and cell number

With the exception of P. minimum, we found a significant decline in the maximum quantum yield of PSII (F v/F m) during algicide incubations, and observed declines in cell density and inhibition of growth in all dinoflagellates tested. Significant loss in PSII photochemistry was accompanied by inhibition in τ (Q A- re-oxidation) and ρ (PSII connectivity). Broadly, these results describe the disruption of photosynthetic electron transport (PET) through PSII. While we cannot ascertain temporal loss in carbon fixation from this data, such reduction in PET would most likely lead to a substantial loss in ATP and NADPH necessary for carboxylation and other cellular processes. Although the relative magnitude of PSII inactivation showed some variability among experiments, we are confident in our species-specific differences because repeated experiments returned the same trends on average (data not shown). We believe some of this variability may stem from natural variations in the production and accumulation of the bioactive compound(s) in each batch of Shewanella sp. IRI-160.

The recovery of F v/F m back to control levels after incubation with 4 % IRI-160AA occurred within 90 h in all but A. tamarense, and was considered to originate from a small population of less effected cells, while the majority were lysed and were no longer contributing to the active chlorophyll a fluorescence signal. In addition to the photochemical recovery seen in the residual population that avoided mortality at lower concentration algicide treatments, these cells were also capable of cell division. Clearly, the dynamics of any population re-growth would be complicated further if these experiments were carried out in natural parcels of water that contained other competing phytoplankton as well as micro and mesograzers.

Species specificity and membrane exposure

Pokrzywinski et al. (2012) concluded that thecate dinoflagellates exhibited reduced levels of algicidal activity compared with naked dinoflagellates. The results presented here corroborate this pattern, with K. veneficum and G. instriatum exhibiting similar declines in cell density and photochemistry, whereas the thecate dinoflagellate A. tamarense exhibited reduced susceptibility to IRI-160AA with a lag in the photochemical response. However, the lack of any photochemical response with algicide treatment yet significant reduction in cell growth noted for P. minimum provides evidence that the algicidal mode of action across these different dinoflagellates is not ubiquitous. Furthermore, the thecal morphology of A. tamarense and P. minimum are very different, with A. tamarense having many similarly sized thecal plates (Fukuyo 1985), and P. minimum comprised primarily of two large valves (Faust et al. 1999). It is possible that plasma-membrane access is required for the algicidal compound(s) to be effective against chloroplast function or that P. minimum is only susceptible to the algicide by an as yet unknown cellular pathway. Several examples in the literature of algicide resistance in organisms with physical barriers lend support to the thecae protection hypothesis. For example, reduced cytotoxicity from fatty acids in Anabaena P-9 heterocysts compared to vegetative cells implicated a protection mechanism of the heterocyst’s thick cell wall (Wu et al. 2006). Secondly, the cellulose plates of the dinoflagellates, Cochlodinium polykrikoides, P. minimum, and Prorocentrum micans, were thought to reduce the speed and magnitude of algicidal effects by peptides, compared with the high activity on the relatively exposed plasma membranes of raphidophytes (Park et al. 2011). Thirdly, gram-positive bacteria were susceptible to a rhamnolipid biosurfactant, but a gram-negative bacterium was not, leading Sotirova et al. (2008) to suggest that lipopolysaccharides in the outer membrane formed a barrier to surfactant activity. Further, the results of Sotirova et al. (2008) led Gustafsson et al. (2009) to propose that the complex cell wall of Microcystis aeruginosa (a gram-negative harmful cyanobacteria) may contribute to this species’ resistance to a rhamnolipid biosurfactant as well.

Another consideration is the different plastid origins of the dinoflagellates tested in this study, with a view to identifying possible links driving the species-specific susceptibilities and photosynthetic responses observed. G. instriatum, P. minimum and A. tamarense share the putatively ancestral type plastid, with the photosynthetic pigment peridinin, a form II RuBisCo, and a highly reduced mini-circle genome, while K. veneficum has a the more derived plastid with fucoxanthin, form I RuBisCo, and a full circular plastid genome (Daugbjerg et al. 2000; Yoon et al. 2005; Wisecaver and Hackett 2011). In consequence, the fewer plastid encoded genes, and possibly less redox regulation of remaining plastid genes in peridinin containing dinoflagellates, require a higher degree of nuclear-encoded plastid-bound protein translocation and are therefore possibly more susceptible to plastid damage if inter-organelle signaling and plastid protein targeting is impaired by IRI-160AA. However, this relationship between plastid genome reduction and algicide susceptibility is likely complex, as evidence suggests that K. veneficum has a chimeric proteome derived from nuclear-encoded secondary (peridinin-based) and tertiary (fucoxanthin-based) endosymbioses (Patron et al. 2006). If there is an influence of plastid origin, then the importance of the plastid would likely depend upon the extent of endosymbiotic gene transfer, and in particular, how many and which genes have been transferred to the nucleus.

Dose responses

The small but significantly greater inhibition based on cell density as compared to F v/F m at 4 % (Fig. 3) is difficult to interpret, but may be due to highly susceptible cells lysing quickly, with little to no time spent in an inhibited-state, such that photoinhibitory signals are not readily observed. The broad similarity between the dose response curves based on photochemistry and cell density however suggests that the increased inhibition based on F v/F m values is likely due to a larger number of affected cells, rather than from increased inhibition in a subset of affected cells. In further support of this, we observed that under a low IRI-160AA concentration (4 %), a proportion of the population (found at the surface of the culture medium) retains a significantly higher photosynthetic competency than the integrated population as a whole (data not shown). Likewise, this also explains why photochemistry appears to recover by 42 h following treatment with 4 % IRI-160AA (Fig. 1). It is important to stress here, however, that this recovery represents a substantially smaller population of cells than at the start of the experiment, and the number of residual photochemically active cells is dose-dependent (Fig. 3). Further testing is needed in order to discern if multiple applications of a low concentration of IRI-160AA may eliminate these residual cells as well.

In a second dose response experiment, serial dilutions of algal culture were treated with 4 % IRI-160AA to examine the possibility that the algicidal activity is dependent on cell density. Here, the degree of mortality and the decline in F v/F m did not vary between culture dilutions (Fig. 3 inset). This response differs from the results described in Mu et al. (2009), in which enhanced algicidal activity was observed by the algicidal bacterium, Ochrobactrum sp. FDT5, when inoculated into cultures with lower chlorophyll a concentrations (Mu et al. 2009). Given the dose response elicited by increasing IRI-160AA concentration as described above, where higher concentrations induced higher mortalities and larger decreases in PSII efficiency, we conclude that IRI-160AA activity is independent of cell density within the range tested here.

Membrane permeability and F v/F m

When photochemistry and membrane permeability were compared in tandem, the two dinoflagellate species tested responded differently to IRI-160AA addition. G. instriatum had a simultaneous decline in F v/F m and cell viability, while K. veneficum lost chloroplast function before plasma membrane integrity, with peak PSII inactivation 6 h prior to the peak loss in cell viability. Two hypotheses could explain this discrepancy between the species. Firstly, that the location, concentration, and susceptibility of algicide target sites differ between the two dinoflagellates (Pokrzywinski et al. submitted). Alternatively, Pokrzywinski et al. (2012) identified three mass ions within the bacterial filtrate (with masses of 239.2, 301.2, and 470.1 Da), which may each contribute to algal mortality and/or growth inhibition. Given that multiple targets and multiple compounds are a common feature of allelopathy (Einhellig 1999), an alternative hypothesis is that differing susceptibility to each component may explain the differences observed between the two dinoflagellates. Confirmation of such hypotheses cannot be resolved without the isolation and identification of the bioactive compound(s) and subsequent identification of the specific cellular target sites.

Darkness

There is a range in dark tolerance across phytoplankton taxa, including species known to cause harmful algal blooms (Smayda and Mitchell-Innes 1974; Peters and Thomas 1996; Furusato et al. 2004; Popels et al. 2007), thus it is not surprising that G. instriatum displayed a much higher degree of general dark tolerance than K. veneficum. However, it is important to note the significant difference between the responses of these two species when treated with the algicide under prolonged darkness, and how their response correlates to what appears to be their basal tolerance to prolonged dark exposure. While this area of research is limited, Mayali et al. (2007) have noted that the bacterial exposure while in prolonged darkness increased the rate of ecdysis in the dinoflagellate Lingulodinium polyedrum. Mayali et al. (2007) suggested that darkness may reduce the ability of L. polyedrum to produce antibiotics, and although not directly applicable here since live bacteria were not used, an analogous mechanism involving antagonistic/protective molecule production could be a possibility for K. veneficum as well. The more rapid loss of PSII photochemistry noted for IRI-160AA-treated K. veneficum in the dark could be due to either a direct amplification of chloroplast dysfunction or may point to an accelerated loss of cellular function at another location. In this regard, disruption in mitochondrial activity and/or synthesis of chloroplast-targeted proteins in the cytoplasm could result in a similar decline in PSII activity as noted here. It is well known that homeostatic chloroplast protein repair requires some light and complete darkness can lead to greater inactivation (Bergo et al. 2003), however, our data may point to indirect disruption of chloroplast function as a better explanation. Notably, Pokrzywinksi et al. (submitted) have shown that IRI-160AA exposure in dinoflagellates induces several cellular markers indicative of programmed cell death (PCD), including extracellular release of hydrogen peroxide, caspase-like activity, and phosphatidylserine inversion of cellular membranes. Likewise, Segovia and Berges (2005) have noted that PCD in the chlorophyte Dunaliella tertiolecta was accelerated when cells were placed in prolonged darkness in the presence of the cytoplasmic protein synthesis inhibitor cycloheximide. Given that maintenance respiration in dinoflagellates can represent over half of the total dark respiration rate in some species, due in part to high protein turnover (Jauzein et al. 2011), and that disruption in mitochondrial activity may lead to down-stream loss in photochemistry (Saradadevi and Raghavendra 1992), it appears that IRI-160AA may affect K. veneficum mitochondrial activity to a greater degree than G. instriatum, and this leads to an even faster disruption in chloroplast electron transport activity in K. veneficum.

Conclusion

Dinoflagellate responses to IRI-160AA were found to be species-specific, but included PSII inactivation, loss in membrane integrity, inhibition of growth, and mortality. In agreement with Pokrzywinski et al. (2012), thecae appeared to provide protection from algicidal attack, and also appeared to limit PSII inactivation. Due to differences in susceptibility between A. tamarense and P. minimum, we suggest that the thecae-armoring hypothesis could be extended to include relative plasma-membrane exposure. In comparing the effects of IRI-160AA on K. veneficum and G. instriatum we found that peak PSII inactivation was decoupled from peak SYTOX permeability in K. veneficum but not in G. instriatum, and that darkness affected K. veneficum substantially more than G. instriatum. These discrepancies led us to speculate that physiological differences can substantially alter susceptibility to IRI-160AA. Ultimately, the specificity of this algicide for certain dinoflagellates remains promising from an applied standpoint and further work on the mode of action, the efficacy of the algicide in situ, and identification of the bioactive compounds are currently underway.