Assessing the viability of microorganisms in the ballast water of vessels transiting the North Atlantic Ocean

Marine Pollution Bulletin 101 (2015) 258–266

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Assessing the viability of microorganisms in the ballast water of vessels transiting the North Atlantic Ocean Jamie L. Steichen ⁎, Antonietta Quigg Department of Oceanography, Texas A&M University, 3146 TAMU, College Station, TX 77843, United States Department of Marine Biology, Texas A&M University at Galveston, 200 Seawolf Parkway, Galveston, TX 77553, United States

a r t i c l e

i n f o

Article history: Received 8 July 2015 Received in revised form 25 September 2015 Accepted 30 September 2015 Available online 9 October 2015 Keywords: Ballast water Invasive species Harmful algal bloom Dinoflagellates Diatoms

a b s t r a c t Testing phytoplankton viability within ballast tanks and receiving waters of ballast water discharge remain understudied. Potentially harmful dinoflagellates and diatoms are transported via ballast water to Galveston Bay, Texas (USA), home to three major ports: Houston, Texas City and Galveston. Ballast water from vessels transiting the North Atlantic Ocean was inoculated into treatments representing low and high salinity conditions similar to the Ports of Houston and Galveston respectively. Phytoplankton in ballast water growout experiments were deemed viable and showed growth in low and mid salinities with nutrient enrichment. Molecular methods identified several genera: Dinophysis, Gymnodinium, Gyrodinium, Heterocapsa, Peridinium, Scrippsiella, Chaetoceros and Nitzschia. These phytoplankton genera were previously identified in Galveston Bay except Scrippsiella. Phytoplankton, including those capable of forming harmful algal blooms leading to fish and shellfish kills, are transported to Galveston Bay via ballast water, and are viable when introduced to similar salinity conditions found in Galveston Bay ports. © 2015 Elsevier Ltd. All rights reserved.

1. Introduction The introduction of invasive species to new regions via ballast water (BW) has caused detrimental impacts to coastal communities and ecosystems (Ruiz et al., 1997, 2015; Gollasch et al., 2000, 2015; Carlton and Ruiz, 2003; Muirhead et al., 2014). Organisms such as bacteria, phytoplankton, and ciliates, are frequently transported across natural barriers via BW and, if conditions are favorable may become invasive species (Zaiko et al., 2015; Burkholder et al., 2007; Drake et al., 2002; Smayda, 2002). Most notably invaders transferred via BW include: the European zebra mussel (Dreissena polymorpha) in the Great Lakes, USA (Hebert et al., 1989), the Atlantic comb jelly (Mnemiopsis leidyi) in the Black Sea (Pereladov, 1983) and the toxic dinoflagellate (Gymnodinium catenatum) in the Pacific Ocean near Tasmania, Australia (Hallegraeff and Bolch, 1992). Many studies have examined the biological community within BW while the vessels are underway (Lavoie et al., 1999; Gollasch et al., 2000; Burkholder et al., 2007). More recently research has started to focus on species viability (ex. phytoplankton) when introduced to the receiving waters (Zaiko et al., 2015; Baek et al., 2011; Kang et al., 2010; Pertola et al., 2006). Survival and subsequent success of

⁎ Corresponding author at: Department of Oceanography, Texas A&M University, 3146 TAMU, College Station, TX 77843, United States. E-mail address: [email protected] (J.L. Steichen).

http://dx.doi.org/10.1016/j.marpolbul.2015.09.055 0025-326X/© 2015 Elsevier Ltd. All rights reserved.

organisms post BW discharge, increase when donor and receiving regions are environmentally similar as seen in coastal ports located along similar latitudes (Carlton, 1996; Vermeij, 1991). BW transfer facilitates the dispersal of nonindigenous across environmental filters or barriers (ex. water circulation inter-specific competition) that would naturally prevent their distribution (Colautti and MacIsaac, 2004). Phytoplankton species taken onboard a vessel are bypassing the first natural filter of dispersal. The transport of viable cells to a new region via BW represents the bypassing of the second stage. Once organisms are discharged into new waters, surviving, adapting (to biotic and abiotic factors) and reproducing and must take place to result in a successful establishment (Ono et al., 2000). By assessing the viability of introduced species in conditions post-BW discharge, we strive to further understand the survivability of organisms which may pose invasion threat. The population in Texas (USA) is expected to double by 2050, with coastal communities experiencing the bulk of this growth (TWDB, 2007). With this development, comes an increase in impervious surfaces, more septic and/or waste water treatment plant discharge, elevated groundwater nutrients, increased atmospheric deposition from transportation sources, etc., all leading to increased runoff and subsequent elevated nutrient loading into the bays and estuaries (Quigg et al., 2009; Greene et al., 2014; Dorado et al., 2015). With this increase in urban development and changes in land use comes the challenge of managing eutrophication in Galveston Bay, the largest and most commercially and recreationally important estuary in Texas

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(Lester and Gonzalez, 2011). Changes in nutrient availability may enhance the opportunity for non-indigenous phytoplankton species to successfully establish in Galveston. In addition to the increased growth in the region, there is a predicted 78% increase in the median total BW discharge after the completion of the Panama Canal expansion in early 2016 (Muirhead et al., 2014). With this increased BW discharge there is an expected rise in the likelihood of introductions of nonindigenous species along the Gulf coast (Muirhead et al., 2014). Introduced species of phytoplankton have the potential to become invasive causing harmful algal blooms, impacting ecosystem services, including the productive oyster and fishing industries (Lester and Gonzalez, 2011; Quigg, 2011). Steichen et al. (2015) recently provided the first reporting of two dinoflagellates (Takayama and Wolozynskia) in the Bay. While it is not certain these genera were introduced to Galveston Bay via BW, similar concerns have been reported worldwide, especially in those estuaries which are home to major ports (Bax et al., 2003; Ruiz et al., 2015). Galveston Bay is home to 3 deep-water ports including: the Port of Houston which is the 10th largest port in the world and 2nd largest in the US in terms of overall waterborne tonnage in 2012, the Port of Texas City (8th largest in the US), and the Port of Galveston (see Steichen et al., 2012 for more details). Another challenge facing Galveston Bay is the expansion of the Panama Canal, set for completion in 2016; significantly larger vessels will enter the bay with greater frequency and after shorter transit times. Due to the increased carrying capacity of these Post-Panamax vessels (i.e. supertankers and larger container vessels), larger volumes of BW will be discharged per event with a corresponding increased number of propagules, increasing the potential success rate of introduced species (Casas-Monroy et al., 2015; Muirhead et al., 2014; Ruiz et al., 2015). In this study, we report that phytoplankton transported to Galveston Bay via BW were viable in a number of treatments where BW was combined with water of lower salinity. Molecular approaches were utilized to target the dinoflagellates and diatoms from the growout treatments and BW samples. Phytoplankton growth increased when introduced to waters of lower salinity and higher nutrient concentrations relative to the waters within the ballast tanks. When larger volumes of BW were introduced to our treatments (simulating increased propagule number), there was a parallel increase in the overall phytoplankton biomass.

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2. Methods 2.1. Sample collection The initial BW sample was collected by a shipping agent who collaborated with the vessel captains that volunteered to provide a BW sample. One (1) 20 L BW sample was collected from each vessel for each respective experiment. BW for all treatments within each of the three experiments was an aliquot from the initial 20 L BW sample. The sampled vessels had conducted a BW exchange in the North Atlantic Ocean while en route to the Port of Houston from the Port of Malabo, (Malabo, Equatorial Guinea, Western Africa): GE1 (14°04′N, 068°51′W), GE2 (01°37′N, 032°39′W) and GE3 (05°55′N, 021°55′W) (Fig. 1). BW age at the time of sampling ranged from GE1: 49 days, GE2: 16 days and GE3: 20 days. BW samples were collected in dark, acid-washed containers and retrieved within 24 h of collection and transported to the laboratory on ice. BW salinity was measured with a refractometer and reported using the unit-less practical salinity scale. Salinities of the BW samples ranged from 30 to 38 on a unitless scale (Table 1). Mean salinities of Port of Houston and Port of Galveston 'during the study period (2007–2009) were 10 (±5.22; n = 20) and 28 (± 4.60; n = 18) and therefore used for the low salinity (LS) and high salinity (HS) treatments respectively (see Steichen et al., 2014). 2.2. Phytoplankton growout experiments (GEs) GEs were designed to test the growth of phytoplankton in ballast tanks when exposed to changing salinity and nutrient regimes. These are analogous to nutrient addition bioassay or resource limitation bioassays performed to assess nutrient limitation (Fisher et al., 1999; Quigg, 2011). Low and high salinity treatments represent average salinities observed in the Port of Houston (avg. salinity 10) and Port of Galveston (avg. salinity 28) respectively (Steichen et al., 2014). The San Jacinto River flows directly into the Port of Houston producing higher average nutrient concentration and lower salinities compared to Port of Galveston (~25 miles south in Galveston Bay) which is more influenced by the Gulf of Mexico (Fig. 1). Gulf seawater was pumped to our facility (average salinity = 33) and filtered through a 0.22 μm Sterivex cartridge filter and autoclaved (121 °C; 40 min). Sterile distilled water was used to dilute the higher salinity gulf water for the lower and high salinity

Fig. 1. Map showing the location of ballast water exchange before BW water sample were collected from each vessel. The location where BW was exchanged include: GE1 (14°04′N, 068°51′W), GE2 (01°37′N, 032°39′W) and GE3 (05°55′N, 021°55′W). Apex of the black triangle indicates reported location of ballast exchange. The Port of Houston (29°36′39.96″N; 95°1′18.12″W) and Port of Malabo (3°46′35.4″N/8°45′19.8″E) are shown by black arrows.

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Table 1 Each treatment contains ballast water (BW), low salinity water (LS), high salinity water (HS), or a combination of these. Controls contain BW, LS or HS (not enriched with nutrients). Experimental treatments have a combination of source water with added nutrients (E) or without. Higher inoculation volumes of ballast were added to LS and HS waters with added nutrients (+). Salinity (unitless scale) is included for the corresponding treatments in GE1, GE2 and GE3. Treatment

BW BWE LS LSE BW/LS BW/LSE BW/LSE+ HS HSE BW/HS BW/HSE BW/HSE+

Volume (mL)

2.4. Analytical methods

Final salinity

Ballast

Port

GE1

GE2

GE3

1000 1000 – – 100 100 500 – – 100 100 500

– – 1000 1000 900 900 500 1000 1000 900 900 500

38 38 10 10 14 14 24 28 28 32 32 35

38 38 10 10 14 14 25 28 28 30 30 34

30 30 10 10 12 12 20 30 30 30 30 30

treatments (Table 1). To determine if phytoplankton productivity was nutrient limited, enriched treatments were prepared with nutrients −1 NH+ added at concentrations of: 30 μmol L− 1 NO− 3 , 30 μmol L 4 , −1 and 30 μmol L SiO . Nutrients were only added at 2 μmol L−1 PO3− 4 3 the beginning of the experiments; treatments with added nutrients (indicated by “E”). Control treatments contained water from one source: BW, LS and HS (Table 1). Enriched treatments include water from one source plus added nutrients: BWE, LSE and HSE (Table 1). Experimental treatments BW/LS and BW/LSE tested the physiological response of the phytoplankton within the BW when exposed to LS water without nutrients and enriched with nutrients respectively (Table 1). Treatments BW/HS and BW/HSE were conducted to measure the phytoplankton response when the BW community when introduced to HS waters without and with additional nutrients respectively (Table 1). Both BW/LSE+ and BW/HSE+ contained equal volumes of BW and lower or high salinity waters respectively (“+”; Table 1). All treatments were maintained under identical conditions (19 °C; 12:12 light: dark cycle; 130–150 μmol m−2 s−1) for 20 (GE2) to 21 (GE2) days; these conditions are known from other studies to support a large diversity of phytoplankton species. The difference in one day of length was a logistical decision not experimental. Due to available resources, biological triplicates (3) were only run for GE2. Biological triplicates were run for each of the twelve (12) treatments in GE2. The BW for each of the replicates in GE2 were an aliquot from the initial 20 L BW sample. 2.3. Fluorescence measurements A fluorescence induction and relaxation (FIRe; Satlantic Instruments) fluorometer (Gorbunov and Falkowski, 2004) was used to measure the response of the phytoplankton to each treatment over the 22 days. The FIRe system is a highly sensitive bench-top instrument that measures a range of physiological characteristics in vivo. Minimum fluorescence yield (F0) was measured every 72 h after samples were dark acclimated for 30 min at room temperature and then normalized to the gain. This was used as an indicator of phytoplankton biomass (Quigg et al., 2013a,b). The gain can be adjusted to account for differences in the phytoplankton biomass in the sample while maintaining optimum sensitivity of the instrument. F0 values were then used to calculate a Phytoplankton Response Index (PRI) (Quigg et al., 2013a) according to:

PRI ¼

The PRI was used to estimate phytoplankton growth as a ratio of the maximum biomass relative to the initial biomass in the GE experiments thereby taking into account differences in the initial biomass of samples in each treatment.

ðMaximum F 0 −Initial F 0 Þ  100 Time at Maximum F 0 ðdÞ

ð1Þ

Multivariate statistical analyses were run with Plymouth Routines in Multivariate Ecological Research (PRIMER v6) package (Clarke and Gorley, 2006). To determine differences in the minimum fluorescence yield (F0) between treatments from day 1 through day 12, we conducted a two-way cross analysis of similarities (ANOSIM) test on Bray–Curtis resemblance matrix (Clarke and Green, 1988; Clarke, 1990, 1993; Clarke and Gorley, 2006). ANOSIM is a non–parametric permutation based test to examine a priori hypotheses which provides a measure of dissimilarity between treatments in the form of an R-statistics and a p-value (9999 permutations). The R-statistic lies on a scale from −1 to 1; as R approaches 1 relative strength of differences increases (Clarke and Gorley, 2006). Typically the range is 0 to 1 although negative values close to 0 are possible and indicate higher similarity across treatments than within a treatment as shown in our results. GE2 was the only experiment where biological triplicates of each treatment were conducted and therefore the only trial in which a statistical analysis was conducted. Phytoplankton growth began to decrease on day 12 during GE2 due to nutrient limitation. The experiment was carried out until day 22 with no increase in overall growth was observed from day 13 through day 22. The statistical analysis was conducted between day 1 and 12 to capture the initial phytoplankton growth when the community was exposed to a nutrient enriched environment. 2.5. Genetic analysis To identify the phytoplankton, specifically dinoflagellates and diatoms, in the growout treatments, we conducted a genetic analysis using protocols described in detail in Steichen et al. (2014, 2015). At the end of the experiment, ~950 mL was filtered onto a 0.2 μm Sterivex cartridge filter and stored at −80 °C. Nucleic acids were extracted using a cetyltrimethylammonium-bromide (CTAB; 3%) and chloroform isoamyl-alcohol extraction followed by an isopropanol precipitation (Doyle and Doyle, 1987). The nucleic acid pellet was dried for 12 h and re-suspended in a saline buffer solution before being stored at − 80 °C for later analysis. A Nanodrop-1000 spectrophotometer V3.7 was used to estimate the nucleic acid concentrations prior to polymerase chain reaction (PCR) (Thermo Fisher Scientific, 2008). PCR amplifications were performed in 50 μL volumes containing approximately 150 ng of template DNA, 10× PCR reaction buffer with 15 mM MgCl2 (Roche Applied Science, Manheim, Germany), 50 μM of each deoxynucleotide, 0.1% bovine serum albumin 1 U Roche Taq DNA polymerase (Roche Applied Science, Manheim, Germany), 10 μM of each primer, 0.5 μL dimethyl sulfoxide. The primer set: DinoF (5′CGATTGAGTGATCCGGTGAATAA-3′) with a 40 bp GC-rich clamp and 4618R (5′-TGATCCTTCTGCAGGTTCACCTAC-3′) was used to amplify the dinoflagellate DNA (Wang et al., 2005). To amplify diatom community, the primer set 1209f: 5′-CAGGTCTGTGATGCCCTT-′3, with added 40-bp GC rich clamp (Giovannoni et al., 1988) and DiaBW8SR1: (5′-CAATGCAGWTTGATGAWCTG-3′) were used (Godhe et al., 2008). The temperature cycling conditions on an Eppendorf Mastercycler gradient thermal cycler were as follows: 1 denaturing step at 95 °C for 3 min followed by 40 cycles of 95 °C 30 s, 55 °C for 30 s, and 72 °C for 40 s, then a final extension step of 72 °C for 5 min (Oldach et al., 2000). PCR products were run through electrophoresis on a 1.5% agarose gel to verify amplification of the target region, followed by denaturing gradient gel electrophoresis performed according to Muyzer (1999) with specific protocols described in Steichen et al. (2014, 2015). DNA was then eluted from the excised bands and used as the template for a second round of PCR. The resulting DNA was run a second time (on

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DGGE) to ensure each band was unique. This final separated DNA was eluted and sequenced with DinoF (no GC-rich clamp)/4618R primers and 1209F/DiaBW8SR1 according to Big Dye Terminator v. 3.1 Sequencing Kit (Applied Biosystems, Foster City, CA). Nucleotide bases were read with a 3130 Genetic Analyzer (Applied Biosystems-Life Technologies, NY). Given sequences of the partial 18S rDNA obtained from this study were less than 200 base pairs, we did not submit to GenBank (Benson et al., 2014). The closest known genetic matches were searched for using the Basic Local Alignment Search Tool (BLAST; Altschul et al., 1997) available on the National Institute for Biotechnology Information website and maintained by the National Institutes of Health (www.ncbi.nlm. nih.gov). We only included the GenBank sequences with at least 90% similarity to the unknown sequences in our samples. Toxoplasma gondii (an Apicomplexa; close relative to the dinoflagellates) was the outgroup taxa for the dinoflagellate tree. Stramenopiles (close relative to diatoms), Bolidomonas pacifica and Bolidomonas mediterranea, were used as the outgroup for the diatom tree. All sequences were aligned using CLUSTAL W within the Molecular Evolutionary Genetics Analysis (MEGA) 5.0 program (Tamura et al., 2011) and then further aligned manually. Evolutionary distances were constructed using the Maximum Likelihood method and the NeighborJoining method with the Jukes Cantor model with algorithm (Saitou and Nei, 1987). Bootstrap values from both methods were obtained

261

from the analysis of 5000 re-samplings within the data set and values greater than 60 are shown on the branches of the phylogenetic trees (Felsenstein, 1985).

3. Results 3.1. Phytoplankton growth The minimum fluorescence yield (F0; proxy for phytoplankton biomass) was measured every three days in each of the treatments to assess changes in phytoplankton biomass (Fig. 2). GE1 did not exhibit measureable growth (data not shown) and phytoplankton were not detected with molecular analyses (data not shown). Phytoplankton growth was detectable and measured in both GE2 and GE3 (Fig. 2). In GE2 and GE3, phytoplankton response (F0) was not detected for treatments: LS, LSE, HS and HSE (data not shown). These treatments did not contain BW and were controls and/or those to which nutrients had not been added (see Table 1). These controls were conducted to ensure that there was no contamination in the low and high salinity waters which were added to BW in various treatments. Data for these treatments will not be shown and was excluded from the statistical analysis results as there was no measurable growth during the experimental time frame (Table 2).

Fig. 2. Fluorescence yield (F0) values that were normalized to the gain for treatments in GE2 (left) and GE3 (right). Treatment labels correspond to Table 1. Averages and standard deviations are shown for GE2 (calculated from biological triplicates). GE3 values correspond to a single measurement as replicates were not conducted. Colors of the bars correspond to treatment attributes: Gray and black — Nutrient enriched; White — No added nutrients.

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Table 2 Results of ANOSIM tests (R-statistic) showing correlation between treatments and fluorescence yield (F0) through day 12 (day of maximum growth in GE2). Treatments LS, LSE, HS, HSE did not contain BW did not display measurable growth and therefore are not shown here. Treatment descriptions are provided in Table 1. Significant correlations are noted in bold and denoted: n.s. for non-significant correlation and p-values for significant results: *** p b 0.001; ** p b 0.01; * p b 0.05.

BW BWE BW/LS BW/LSE BW/LSE+ BW/HS BW/HSE

BWE

BW/LS

BW/LSE

BW/LSE+

BW/HS

BW/HSE

BW/HSE+

0.093 (n.s.)

0.093 (n.s) 0.213 (n.s)

0.361 (*) 0.241 (n.s) 0.38 (*)

0.093 (n.s) 0.074 (n.s) 0.315 (*) −0.102 (n.s)

−0.176 (n.s) −0.148 (n.s) 0.065 (n.s) 0.037 (n.s) −0.019 (n.s)

0.676 (**) 0.676 (**) 0.694 (***) 0.491 (*) 0.250 (*) 0.593 (*)

0.370 (*) 0.444 (**) 0.519 (**) 0.278 (**) 0.037 (n.s) 0.241 (n.s) 0.333 (*)

Control treatments (BW, LS and HS) in GE2 had F0 values of around 0 consistent with a lack of growth (BW: Fig. 2a; LS and HS not shown). Significantly higher phytoplankton growth (F0 ) was observed in majority of the treatments where BW was combined with enriched low and high salinity waters there was (BW/LSE, p b 0.05; BW/HSE p b 0.01; BW/HSE + p b 0.05) when compared to the growth in the BW alone (Fig. 2b–c; Table 2). The only treatments with significantly higher F0 (i.e. phytoplankton growth) than the nutrient enriched BW (BWE ) was when the BW was inoculated into the enriched higher salinity water (BW/HS E ; p b 0.01) and enriched and increased volume of BW (BW/HS E +; p b 0.01) treatments (Fig. 2a, b; Table 2). The phytoplankton growth was lower in when the BW was inoculated into lower salinity water without nutrients added compared to all other treatments that contained BW in low and high salinity enriched treatments (BW/LSE, p b 0.05; BW/LSE +, p b 0.05; BW/HSE, p b 0.001; BW/HSE+, p b 0.01). The phytoplankton community had significantly higher growth in the treatments with higher salinity and nutrients (BW/HSE, p b 0.05; BW/HSE +, p b 0.01) compared to the enriched lower salinity treatments (Fig. 2b–c; Table 2). The phytoplankton growth when BW was inoculated into higher salinity enriched waters (BW/HSE ) was the only treatment that showed a significant increase compared to growth when BW was inoculated into enriched lower salinity waters (BW/LSE+, p b 0.05). Overall the treatments that include BW and enriched higher salinity waters (BW/HSE) showed significantly increased levels of phytoplankton growth compared to all treatments with the exception of when BW was combined with increased high salinity enriched water (BW/HSE+; Fig. 2b–c; Table 2). The phytoplankton growth (F0) in GE3 showed overall similar patterns to those observed in GE2 (Fig. 2). The F0for control treatments (BW, LS and HS) was approximately 0 (relative units) were consistent with GE2 results (Fig. 2). The treatments displaying the most variability compared to GE2 are those where BW was inoculated into enriched higher salinity waters (BW/HSE) and increased volume of BW (BW/HS E +). In GE3 phytoplankton growth increased in treatments with a decreased volume of BW inoculated into enriched higher salinity waters had a longer lag phase and began increasing on day 14 and then continued increasing throughout GE3 (Fig. 2d–f). The overall fluorescence was increased compared to the treatments in GE2 ads well as in the initial BW sample in GE3 (Fig. 2). The PRI was only calculated in the enriched treatments; data is not shown for the unenriched treatments (Fig. 3). In GE2, there was an increased response index of the phytoplankton in the treatments with enriched higher salinity waters (BW/HSE, BW/HSE +; PRI: 165 (±0.5) – 210 (±3.1)) and in GE3 the PRI ranged 131–174 (Fig. 3). In GE2 and GE3, enriched lower salinity treatments (BW/LSE, BW/LSE+) had overall lower PRI values compared to the higher salinity treatments (Fig. 3). The PRI in the enriched lower salinity treatments (BW/LSE, BW/LSE +) of GE2 ranged from 28 (±2.64) − 72 (±8.39) and in GE3 from 59–75 (Fig. 3). The phytoplankton response in the BW with nutrients added showed the second highest growth response increase (PRI: 205) next to the BW inoculated into enriched high salinity waters (BW/HSE+, PRI: 210; Fig. 3).

3.2. Phytoplankton assemblages Eight genera of phytoplankton were found in GE2 and GE3 (Table 3; Fig. 4 and Fig. 5). Four dinoflagellate genera (Dinophysis, Gymnodinium, Gyrodinium and Scrippsiella; Fig. 4) and two types of diatoms (Chaetoceros, family Thalassiosirales; Fig. 5) were identified in GE2. Dinoflagellates were observed in both low and high salinity treatments including BW/LSE, BW/HS and BW/HSE + (all with added nutrients) in GE2 (Table 3). Dinophysis sp. was detected in GE2 in the treatment without added nutrients (BW) and a Gymnodinum sp. was identified in BW/LSE (which had lower salinity than the initial BW sample and added nutrients). At the termination of GE3, we genetically identified four dinoflagellate genera including: Gymnodinium, Gyrodinium, Heterocapsa and Peridinium (Fig. 4) and two diatoms including Nitzschia and Thalassiosirales (Fig. 5). In GE3, dinoflagellates were observed in BW and BW/LS (without added nutrients) and BW/LSE, BW/HS and BW/HS E (with added nutrients) (Table 3). Gymnodinium sp. was identified from BW when mixed with enriched high salinity waters (Table 3; Fig. 4). Gyrodinium was identified in treatments without nutrients and higher salinity in GE2 but in enriched low salinity waters in GE3 (Table 3; Fig. 4). Heterocapsa was identified in enriched low and high salinity treatments (BW/LSE and BW/HSE+ respectively) in GE3. Peridinium sp. preferred the enriched high salinity waters. Scrippsiella was identified in GE2 within the enriched high salinity (BW/HSE).

Fig. 3. Phytoplankton response index (PRI) values (Eq. (1)) measured at the peak of growth for GE2 (black) and GE3 (white) (day 12 and 13 respectively). Standard deviations are shown as error bars for each treatment for GE2. Data is only shown for nutrient enriched treatments that contain BW. There was no measurable phytoplankton response in treatments without BW and those treatments are not shown. Refer to Table 1 for a detailed description of the composition of all treatments.

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Table 3 Dinoflagellates and diatoms genera identified from GE2 (▲) and GE3 (●) denoted in each treatment. Genera

BW

Dinophysis Gymnodinium Gyrodinium Heterocapsa Nitzschia Peridinium Scrippsiella Thalassiosirales Uncultured Chaetoceros Uncultured dinoflagellate Unknown dinoflagellate (A) Unknown dinoflagellate (B) Unknown dinoflagellate (C)

▲

BWE

BW/LS

BW/LSE ● ● ● ●

●

BW/LSE+

BW/HS

BW/HSE

BW/HSE+

▲ ▲ ● ● ●

● ● ▲

●

▲ ▲ ● ●

▲,● ● ●

▲

4. Discussion Many studies focus on the inventory of phytoplankton within ballast tanks, including at various stages of a vessel voyage (e.g., Burkholder et al., 2007; Gollasch et al., 2000; Lavoie et al., 1999); few however have considered their viability once the BW has been discharged (Baek et al., 2011). This is the first study examining the viability of

▲

dinoflagellates and diatoms within the ballast tanks of ships entering Galveston Bay. We found there was growth of phytoplankton if the BW was present in the tank for ≤ 20 days (GE2 and GE3) but not in the older BW (49d; GE1) supporting the idea that length of time within the ballast tanks also plays a role in the survival of these organisms (Burkholder et al., 2007; Sutherland and Levings, 2013). Further, with the addition of a larger volume of BW (9:1 compared to 1:1) to

Fig. 4. Maximum likelihood phylogenetic tree showing the dinoflagellate diversity of GE2 and GE3. Bootstrap values for the neighbor-joining and maximum likelihood are shown at each respective node (NJ/MLE; Bootstraps values N60%). Dashed lines represent bootstraps b60% support. The analysis included 56 nucleotide sequences with a total of 71 positions.

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Fig. 5. Maximum likelihood phylogenetic tree showing the diatom diversity of GE2 and GE3. Bootstrap values for the neighbor-joining and maximum likelihood are shown at each respective node (NJ/MLE; Bootstraps values N60%). Dashed lines represent bootstraps b60% support. The analysis included 32 nucleotide sequences with a total of 65 positions.

treatments (i.e., BW/LSE+ and BW/HSE+), there was a greater overall increase in phytoplankton biomass (Fig. 2 and Fig. 3; Table 1). This is consistent with increased propagule pressure with greater BW discharge volumes. The volume of BW discharged rather than simply the number of vessels discharging BW, plays an important role in propagule size and number, invasion supply and success rate of invaders dispersed in BW (Drake and Lodge, 2004; Ruiz et al., 2015; Simberloff, 2009; Verling et al., 2005). This study shows that species of phytoplankton transported in BW can flourish when incubated in salinities similar to those observed in the coastal ports, particularly with the addition of nutrients. The latter suggests the alleviation of nutrient limitation either in the ballast tank or in the port waters. Further studies are required to determine the impacts of various temperature and nutrient conditions on the success of phytoplankton found within BW. When BW was inoculated in higher salinity waters, there was a significant increase in the response of the phytoplankton compared to the BW introduced to the low salinity waters (Fig. 2; Table 2). This is important when considering that euryhaline

organisms may be given an advantage when discharged into coastal environments. If the coastal regions become increasingly eutrophic due to increasing development and population growth and associated impervious surfaces, septic and sewer discharges, vehicular emissions, and other nutrient-generating anthropogenic activities, BW organisms may find appropriate growth conditions and bloom upon introduction to Galveston Bay and other rapidly growing coastal regions. Previous studies conducted on the phytoplankton community in BW have also identified Gymnodinium, Gyrodinium, Heterocapsa (dinoflagellates) and Nitzschia and Thalassiosirales (diatoms) (Table 4). Gymnodinium spp. are known toxin producers and have formed harmful algal blooms after becoming introduced into a new region (Tasmania, Australia) via BW transport (Hallegraeff and Gollasch, 2006; McMinn et al., 1997). Species of Heterocapsa have also been identified as harmful and have caused mass mortality of oysters in the Seto Inland Sea, Japan (Imai et al., 2006). Scrippsiella spp. identified in these BW samples has not previously been reported in Galveston Bay (Table 3 and Table 4). Scrippsiella trochoidea, is typically a non-toxic algal bloom former and

Table 4 Dinoflagellate and diatom genera identified from GE2 and GE3 which were also identified in ballast water tanks from previous literature. In several of these reports where the specified genera were identified cells were deemed viable (indicated in “Viable” column). Genera that include toxic species are indicated in toxic column. Genera identified in this study

Toxic

Previous detection in BW Identified

Dinoflagellates Dinophysis Gymnodinium Gyrodinium Heterocapsa Peridinium Scrippsiella Diatoms Nitzschia Thalassiosirales Uncultured Chaetoceros

+ + + +

Reference(s) Viable

+ + + + + +

+ + +

Burkholder et al. (2007); David et al., 2007; Roy et al., 2012 Hallegraeff and Bolch (1992); Pertola et al. (2006) David et al., 2007; Garrett et al., 2011 Tamai, 1999; Burkholder et al. (2007); Pertola et al. (2006) Pertola et al. (2006); Burkholder et al. (2007) Casas-Monroy et al., 2011; Pertola et al. (2006)

+ + +

+ +

Klein et al., 2010; Boltovskoy et al., 2011 Kang et al. (2010) Liebich et al., 2012; Kang et al. (2010)

+

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has killed Eastern oysters (Crassostrea virginica) by physiochemical means (such as clogging of gills) during a bloom off along the Northwest Atlantic Ocean coast (Tang and Gobler, 2012). An invasion of S. trochoidea could therefore negatively impact the Galveston Bay system ecologically and economically. Galveston Bay annually oyster harvesting, dominantly C. virginica, has supported a multi-million dollar fishery important to the local economy within Galveston Bay (Lester and Gonzalez, 2011), hence its introduction to this ecosystem could have significant detrimental impacts Galveston Bay, along with Matagorda Bay were found to be “hot spots” for fish kills, with N 250 million fish deaths reported from 1961 to 2006 (Thronson and Quigg, 2008). Eutrophication has led to an increase in nuisance or toxic algal blooms in estuaries along the Texas coast (Thronson and Quigg, 2008; Quigg et al., 2009) and around the world (Anderson et al., 2012; Hallegraeff, 2015; Hallegraeff and Gollasch, 2006; Burkholder et al., 2007; Paerl et al., 2014). When organisms in ballast tanks have been limited by the decreased concentration of nutrients and are then discharged into nutrient rich port waters, the chance of their survival may increase as shown herein. The changes in the salinity conditions that we tested did not appear to suppress growth. One of the theories behind open ocean exchange of BW is that coastal organisms which require lower salinity conditions will be replaced with organism that are adapted to higher salinities, and when they are subsequently discharged in the receiving port they should perish (Pertola et al., 2006). Our results found that this is not the case, as well as those of previous studies which have observed living organisms in BW (e.g., Burkholder et al., 2007; Zaiko et al., 2015), and therefore add support to the argument that there is the need for more stringent BW treatment methods. Given that phytoplankton are small in size, globally abundant and capable of surviving inclement conditions, they are likely to survive these suboptimal conditions within BW tanks. Bacteria, phytoplankton and invertebrates are being introduced to new regions they may displace endemic populations, thereby reducing community diversity and genetic heterogeneity (Pertola et al., 2006). By sampling and monitoring BW, we may assess the frequency in which harmful organisms are transported and the potential for invasion in region of high shipping traffic. We were able to confirm that potentially harmful phytoplankton taxa are not only being transported, they have the capability to withstand the salinity differences and flourish in the new waters. The findings of this study support that if given and salinity nutrient conditions are appropriate, introduced species of phytoplankton (potentially harmful) are capable of surviving and flourishing. With the expansion of the Panama Canal set to be completed by early 2016, larger Post-Panamax vessels from the Pacific Ocean will be able to enter the Gulf of Mexico, and at an accelerated rate (i.e. shorter transit time), and capable of transporting larger cargo loads and in turn, increased volumes of ballast water (Muirhead et al., 2014). It has been predicted that within 5 years of the Panama Canal expansion, the Gulf of Mexico coast will receive ~ 80% more total ballast water discharge and a 3-fold increase in the number of vessel arrivals (Muirhead et al., 2014; Ruiz et al., 2015). For Galveston Bay, it is anticipated this will bring significant pressure on the ecosystem. Not only will there be an increased volume and frequency of BW discharge events, but from vessels which have had correspondingly shorter voyages. These large Post-Panamax vessels will be transporting cargo and potential invaders to the Gulf Coast region from Pacific Ocean including highly invaded regions (e.g., San Francisco Bay). Given over 40% of the aquatic nonindigenous species that have been introduced globally have been dispersed via shipping vessels (Gollasch et al., 2015); the biological community within the Gulf of Mexico including its coastal bays and estuaries will experience increased risk from potentially invasive species in ballast water being delivered from the Pacific Ocean (Ruiz et al., 2015). We anticipate an increase in the invasion risk and likelihood of introduction of non-native species into the Gulf of Mexico (Muirhead et al., 2014). With these changes approaching, it is strongly suggested

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that monitoring programs be put in place to be on alert for invasive organisms being delivered to the Gulf Coast regions within the estuaries and more specifically the coastal ports. Acknowledgments We would like to thank Daniel Roelke (Texas A&M University) and Robin Brinkmeyer (Texas A&M University at Galveston) and for their valuable contributions to the experimental design of this project. We would also like to express our gratitude to the various funding agencies of this project including the Advanced Research Program (0102980005-2006), Texas General Land Office—CMP (Texas Coastal Management Program GLO Contract No. 09-032-000-3349) and the Environmental Protection Agency (US EPA Cooperative agreement: MX954527). We would like to thank Fields Jackson and vessel captains that took the time to collect the BW samples; without their volunteer efforts this research would not have been possible. We extend an extra special thank you to all of the members (past, present and auxiliary) of the Phytoplankton Dynamics Laboratory at Texas A&M University at Galveston, who have assisted in the collection and processing of these samples. Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.marpolbul.2015.09.055. References Altschul, S.F., Madden, T.L., Schäffer, A.A., Zhang, J., Zhang, Z., Miller, W., Lipman, D.J., 1997. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 25, 3389–3402. Anderson, D.M., Cembella, A.D., Hallegraeff, G.M., 2012. Progress in understanding harmful algal blooms: paradigm shifts and new technologies for research, monitoring, and management. Ann. Rev. Mar. Sci. 4, 143–176. Baek, S.H., Jung, S.W., Shin, K., 2011. Effects of temperature and salinity on growth of Thalassiosira pseudonana (Bacillariophyceae) isolated from ballast water. J. Freshw. Ecol. 26, 547–552. Bax, N., Williamson, A., Aguero, M., Gonzalez, E., Geeves, W., 2003. Marine invasive alien species: a threat to global biodiversity. Mar. Policy 27, 313–323. Benson, D.A., Clark, K., Karsch-Mizrachi, I., Lipman, D.J., Ostell, J., Sayers, E.W., 2014. 352 GenBank. Nucleic Acids Res. 42, D32–37. Boltovskoy, D., Almada, P., Correa, N., 2011. Biological invasions: assessment of threat from ballast-water discharge in Patagonian (Argentina) ports. Environ. Sci. Policy 14, 578–583. Burkholder, J.M., Hallegraeff, G.M., Melia, G., Cohen, A., Bowers, H.A., Oldach, D.W., Parrow, M.W., Sullivan, M.J., Zimba, P.V., Allen, E.H., Kinder, C.A., Mallin, M.A., 2007. Phytoplankton and bacterial assemblages in ballast water of US military ships as a function of port of origin, voyage time, and ocean exchange practices. Harmful Algae 6, 486–518. Carlton, J.T., 1996. Biological invasions and cryptogenic species. Ecology 1653–1655. Carlton, J.T., Ruiz, G.M. (Eds.), 2003. Invasive Species: Vectors and Management Strategies. Island Press, Washington, USA. Casas-Monroy, O., Roy, S., Rochon, A., 2011. Ballast sediment-mediated transport of nonindigenous species of dinoflagellates on the East Coast of Canada. Aquat. Invasion 6, 231–248. Casas-Monroy, O., Linley, R.D., Adams, J.K., Chan, F.T., Drake, D.A.R., Bailey, S.A., 2015. Relative invasion risk for plankton across marine and freshwater systems: examining Efficacy of proposed international ballast water discharge standards. PLoS One 10, e0118267. http://dx.doi.org/10.1371/journal.pone.0118267. Clarke, K.R., 1990. Comparisons of dominance curves. J. Exp. Mar. Biol. Ecol. 138, 143–157. Clarke, K.R., 1993. Non-parametric multivariate analyses of changes in community structure. Aust. J. Ecol. 18, 117–143. Clarke, K.R., Gorley, R.N., 2006. PRIMER v6: user manual/tutorial PRIMER-E. PRIMER-E Ltd., Plymouth. Clarke, K.R., Green, R.H., 1988. Statistical design and analysis for a ‘biological effects’ study. Mar. Ecol. Prog. Ser. 46, 213–226. Colautti, R.I., MacIsaac, H.J., 2004. A neutral terminology to define ‘invasive’ species. Divers. Distrib. 10, 135–141. David, M., Gollasch, S., Cabrini, M., Perkovič, M., Bošnjak, D., Virgilio, D., 2007. Results from the first ballast water sampling study in the Mediterranean Sea–the Port of Koper study. Mar. Poll. Bull. 54, 53–65. Dorado, S., Booe, T., Steichen, J.L., McInnes, A.S., Windham, R., Shepard, A., Lucchese, A., Preischel, H., Pinckney, J.L., Davis, S.E., Roelke, D.L., Quigg, A., 2015. Towards an understanding of the interactions between freshwater inflows and phytoplankton communities in a subtropical estuary in the Gulf of Mexico. PLoS One http://dx.doi.org/10. 1371/journal.pone.0130931.

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