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. 2024 Apr 12;58(16):6924–6933. doi: 10.1021/acs.est.3c10502

An On-Farm Workflow for Predictive Management of Paralytic Shellfish Toxin-Producing Harmful Algal Blooms for the Aquaculture Industry

Rendy Ruvindy †, Penelope A Ajani †, Sereena Ashlin ‡, Gustaaf Hallegraeff §, Kerstin Klemm ∥, Christopher J Bolch §, Sarah Ugalde §,⊥, Mark Van Asten #,∇, Stephen Woodcock †, Matthew Tesoriero †, Shauna A Murray †,*
PMCID: PMC11044886  PMID: 38608723

Abstract

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Paralytic shellfish toxins (PSTs) produced by marine dinoflagellates significantly impact shellfish industries worldwide. Early detection on-farm and with minimal training would allow additional time for management decisions to minimize economic losses. Here, we describe and test a standardized workflow based on the detection of sxtA4, an initial gene in the biosynthesis of PSTs. The workflow is simple and inexpensive and does not require a specialized laboratory. It consists of (1) water collection and filtration using a custom gravity sampler, (2) buffer selection for sample preservation and cell lysis for DNA, and (3) an assay based on a region of sxtA, DinoDtec lyophilized quantitative polymerase chain reaction (qPCR) assay. Water samples spiked with Alexandrium catenella showed a cell recovery of >90% when compared to light microscopy counts. The performance of the lysis method (90.3% efficient), Longmire’s buffer, and the DinoDtec qPCR assay (tested across a range of Alexandrium species (90.7–106.9% efficiency; r2 > 0.99)) was found to be specific, sensitive, and efficient. We tested the application of this workflow weekly from May 2016 to 30th October 2017 to compare the relationship between sxtA4 copies L–1 in seawater and PSTs in mussel tissue (Mytilus galloprovincialis) on-farm and spatially (across multiple sites), effectively demonstrating an ∼2 week early warning of two A. catenella HABs (r = 0.95). Our tool provides an early, accurate, and efficient method for the identification of PST risk in shellfish aquaculture.

Keywords: paralytic shellfish toxins (PSTs), harmful algal blooms (HABs), aquaculture industry, alexandrium spp., sxtA4 gene, molecular detection, on-farm workflow

Short abstract

This study presents a practical, three-step on-farm workflow, utilizing the sxtA4 gene for the early, efficient, and cost-effective prediction of paralytic shellfish toxin risk in shellfish aquaculture, enhancing industry preparedness and safeguarding economic loss.

1. Introduction

Paralytic shellfish toxins (PSTs) comprising saxitoxin and its analogues are a group of neurotoxic alkaloid compounds responsible for the syndrome paralytic shellfish poisoning (PSP).1 PSTs are produced by certain marine dinoflagellates, and their accumulation in shellfish has resulted in severe economic impacts through farm closures and product recalls worldwide.1−4 In Australia, a leading PST-producing species is Alexandrium catenella (group 1). This species was first implicated in a recall of blue mussels (Mytilus galloprovincialis) from Tasmania, which were contaminated with PSTs, resulting in an estimated $AUD 23 M loss.5 Since then, A. catenella blooms have reoccurred seasonally in Tasmania, with the highest concentration of PST ever recorded in 2017, at 150 mg kg–l, over 150 times above the regulatory level of 0.8 mg kg–16.6

Shellfish safety monitoring for harmful algal blooms (HABs) and their toxins as implemented by seafood safety programs worldwide commonly relies on a few main techniques. The first is the identification of harmful algal species in seawater using light microscopy, but discriminating between species requires highly specialized skills, and misidentification can result in unnecessary farm closures.7,8 The second technique is the detection of PSTs in shellfish using chemical methods such as liquid chromatography with tandem mass spectrometry (LC-MS/MS and LC-MS).9 This method is time-consuming, expensive, and not suitable for use on-farm. More rapid, cost-effective, and on-farm testing methods would therefore make on-farm shellfish harvest management simpler and faster and result in fewer closures. Methods to rapidly detect PSTs in shellfish tissue have been developed based on enzyme-linked immunosorbent assays (ELISAs), lateral flow approaches, and biosensors.10−14 However, there is known to be a time-lag of at least several days or longer between the presence of PST-producing species in the water column and a measurable toxin concentration in shellfish tissue.15 Another technique using molecular probe technology is the environmental sample processor (ESP), a deployable automated sampling device for molecular analyses in situ, yet this is still considered in its developmental infancy and expensive. Overall, there is a need for an inexpensive early warning system, which would allow sufficient time for appropriate shellfish harvesting management decisions.15

Molecular detection of toxic dinoflagellates can be performed using environmental DNA (eDNA) and quantitative polymerase chain reaction (qPCR) assays based on barcoding markers, for example, the rRNA array.16−19 Using this method, cryptic species such as A. catenella can be quantified and discriminated from other co-occurring species.7,19 However, rDNA-based qPCR assays may not always be the most appropriate means of quantification, as copy numbers of rRNA genes can vary by 3–5 orders of magnitude within a species.20−23 An assay based on a gene involved in PST biosynthesis24−26 such as sxtA4(27) may be more reliable as the variation in genomic copy numbers may be less than that of rRNA genes. While copy numbers of sxtA4 genes show variation, it is of a lesser degree, up to 1 order of magnitude.21,22 Moreover, a positive correlation has been found between sxtA4 copies per cell and the quantity of PSTs synthesized per cell, showing a dosage effect such that the quantification of sxt4A may be particularly pertinent to quantifying the likelihood of certain PST concentrations.22,28 For the purposes of seafood safety risk assessment, the identity of the species is less important than the indication of the presence of target genes linked to PST biosynthesis. For this reason, a commercial qPCR-based assay Phytoxigene DinoDtec has been developed based on the detection of a gene region, sxtA4, that is only found in dinoflagellate species that produce PSTs. While the development of the kit has been carried out in the laboratory, this study represents the first field and on-farm applications of this kit.

Phytoplankton monitoring for shellfish safety risk management using eDNA19,29−31 is currently carried out in dedicated molecular biology laboratory facilities. With the emergence of portable qPCR equipment, the quantification of target genes in situ has been made possible. To conduct qPCR on-farm, a simplified and standardized workflow is necessary. Such an early detection tool should be able to be conducted with minimal training by nonspecialists, be rapid, relatively inexpensive, use no toxic or harmful chemicals, use relatively inexpensive equipment, have minimal electricity requirements, and be able to be conducted outside of a controlled laboratory setting.

Here, we developed a simplified and standardized workflow that allows for the detection of PST-producing microalgae on site with results available within ∼2 h. The workflow included three stages: (1) water collection and filtration using the phytoxigene portable water sampler; (2) cell lysis for DNA extraction and buffer selection for sample preservation; and (3) the Phytoxigene DinoDtec lyophilized qPCR assay and data interpretation. We then evaluated this early detection tool both temporally and spatially during two HAB events of PST-producing A. catenella.

2. Materials and Methods

2.1. Cell Recovery Using a Portable Water Sampler

The gravity-operated portable water sampler (Diagnostic Technology) (Figure 1) was designed to standardize the water filtration of phytoplankton of the size class relevant for HAB analysis from 3 L seawater. The first stage of this filtration consists of a 100 μm nylon mesh as a prefilter and to remove zooplankton, debris, and larger phytoplankton, and the second stage is a 11 μm nylon fabric designed to capture and retain HAB cells. To operate, seawater is poured from the top until the chamber is full. The valve is then released to allow the seawater to flow through the 11 μm mesh. The flow-through seawater is collected with a squeeze bottle. The second-stage filter is then removed from the sampler, and the cells that have collected on the 11 μm filter are back-flushed with the filtered seawater into a collection chamber such as a 50 mL falcon tube. Finally, the water sampler should be rinsed with tap water after each sampling event. If either filter becomes clogged, then the mesh should be sprayed with diluted bleach (10%) under pressure, followed by a thorough rinse with tap water (2 min).

Figure 1.

Figure 1

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Photo and technical diagram of the gravity-operated portable water sampler showing the following components: A. a first-stage prefilter with a 100 μm mesh; B. a 3 liter water chamber; and C. a second-stage filter with an 11 μm mesh designed to capture and retain HAB cells such as dinoflagellates and all measurements. The sampler is made from PVC piping with a midline ball valve/tap.

To determine the cell recovery efficiency of the water sampler, 3 L of 5 μm filtered seawater was spiked in triplicate with 100, 1000, or 5000 cells of A. catenella (group 1 genotype, strain ATTR/F, Triabunna Tasmania, Australia). All samples were then passed through the sampler, and the sample was recovered as described above. One mL from each concentrated sample was counted using a Sedgewick–Rafter counting chamber (ProSciTech, Australia), and cell recovery was estimated from counts before and after the filtration.

2.2. Performance of Cell Lysis Method for DNA Extraction and Preservation

To develop a simple cell lysis protocol for DNA extraction from seawater samples and examine its performance, a culture of Alexandrium pacificum (strain MMWA 83) was serially diluted from an initial starting density of 170,800 cells, ten times at a 1:2 dilution rate. Diluted samples were filtered through a 25 mm Swinnex filter holding an 8 μm nitrocellulose filter (Merck Millipore, Massachusetts). Each filter was removed and inserted into a BioGX lysis tube (BioGX, Birmingham) containing ∼300 mg of sterilized glass beads and 500 μL of sterilized bead lysis buffer. Tubes were then vortexed at a maximum speed for 10 min using a Vortex-Genie 2 (Scientific Industries, New York) equipped with a 24-place adapter for 1.5–2.0 mL tubes (Qiagen, Venlo, Netherlands). Tubes were subsequently centrifuged at 1000 g for 1 min to pellet the debris. Five μL of the supernatant from each tube was transferred into the qPCR reaction plate. The DinoDtec qPCR reaction mix was rehydrated with 80 μL of nuclease-free water, and 20 μL was added to each well, resulting in a 25 μL qPCR reaction volume. The cycling conditions for qPCR were 95 °C for 2 min, followed by 45 cycles of 95 °C for 10 s and 64 °C for 45 s on a CFX96 Touch Real-Time PCR Detection System (Biorad). The fluorescence signal from the probe was quantified in the FAM channel, and the copy numbers of the gene were calculated by establishing a standard curve using the sample quantification cycle (Cq) (y-axis) and the natural log of concentration (x-axis). The percentage efficiency of each reaction was then calculated by the equation E = −1 + 10(−1/slope). A satisfactory amplification efficiency was accepted if between 90 and 110%.32 The internal amplification control (IAC), which showed whether the reaction was amplified, was measured on the HEX channel. In samples where the IAC Cq is 1.5 cycles higher than the blank or nontemplate control IAC CT, the result is considered invalid. All samples were below this threshold.

The inhibitory effect of three common DNA and tissue preservative solutions, Longmire’s buffer,33 RNALater (Sigma-Aldrich), and Lugol’s Iodine (Sigma-Aldrich) on the DNA extraction and qPCR processes, was tested using two DNA extraction methods. The inhibitory effect was visualized by examining the Cq of qPCR reactions using DNA extracted using each method. A total of 24, 8 μm nitrocellulose filters (25 mm diameter) with 31,000 cells of A. catenella strain ATTR/F on each filter were prepared by filtering a culture using syringe filtration. The samples were then split into two sets of 12. The first set (1) underwent an extraction using a FastDNA for Soil kit (MP Biomedicals) and the second underwent DNA extraction using BioGX Lysis tubes (2). Membrane filters containing A. catenella ATTR/F were inserted into either the FastDNA bead beating tubes for set 1 or in the BioGX Lysis tube for set 2, and 1 mL of each of the 3 preservative buffers was added (n = 3 per buffer and n = 3 for control with no preservative buffer). Tubes with fixed cells were then vortexed for 5 min and centrifuged at 1000 g for 1 min. The qPCR reactions were performed with the same conditions as the test of the lysis tube efficiency described previously, with the primers for sxtA4, and 1 μL of sample DNA.

2.3. Phytoxigene DinoDtec qPCR Specificity and Efficiency Test

The DinoDtec kit (Diagnostic Technology, Australia) is a commercialized qPCR assay developed based on primers targeting the sxtA4 gene (Murray et al.). To test the specificity and efficiency of the kit, qPCR reactions were compared between six non-PST-producing Alexandrium species (A. affine, A. concavum, A. leii, A. margalefi, A. fraterculus, A. pseudogonyaulax) and three PST-producing species (A. pacificum, A. catenella and A. minutum) (Table S1). DNA was extracted from 50,000 to 75,000 cells of each strain using the FastDNA Spin kit for soil and eluted in 80 μL of elution buffer. The DinoDtec mix was rehydrated with 80 μL of nuclease-free water, and a 20 μL aliquot was mixed with 2 μL of DNA and 3 μL of PCR-grade water to a total of 25 μL for each qPCR reaction. Cycling conditions and signal quantification were the same as above except this time using a MyGo Mini portable thermocycler (IT-IS LifeScience, Cork, Republic of Ireland). Phytoxigene DinoNAS standards were used to develop the standard curve for the sxtA4 copy number quantification. Phytoxigene DinoNAS is a DNA standard for the DinoDtec kit developed by the National Measurement Institute (NMI).

Efficiency tests of the DinoDtec sxtA4 qPCR assay were performed by developing standard curves from strains of A. pacificum, A. catenella, and A. minutum. Serial dilutions were carried out (1:2) on three extracts for each strain with a DNA amount equivalent to 3000 cells. qPCR reactions were carried out in triplicate, with temperature settings of 95 °C for 2 min, followed by 45 cycles of 95 °C for 10 s and 64 °C for 45 s. Standard curves were established as above, and the percentage efficiency of each reaction was calculated.

2.4. On-Farm Early Detection Tool Testing

For in situ pipeline testing, ∼10 L of seawater was collected from a depth of 5 m using a peristaltic pump from a lease offshore from Spring Bay Seafoods, Tasmania (42.59 S, 147.97 E), every Sunday between 9:30 and 11am from 22nd May 2016 to 30th October 2017. The seawater was then transported to the hatchery in Triabunna (Tasmania) and subsequently filtered using the phytoxigene portable water sampler, followed by syringe filtration and drying, as described earlier. Filters were removed, inserted into a BioGX cell lysis tube, vortexed at a maximum speed for 10 min, and centrifuged at 1000g for 1 min to precipitate cell debris. Five μL of cell lysate and 20 μL of DinoDtec were transferred and mixed in a qPCR tube, and qPCR was carried out as previously described. A negative control (5 μL of PCR-grade water) and Phytoxigene DinoNAS standard representing 5000 copies of sxtA4 was included in each qPCR run.

To infer the temporal relationship between sxtA4 copies L–1 in seawater and PSTs in mussel tissue (M. galloprovincialis), the concentration of PSTs was measured in mussels weekly across the sampling period. Approximately 10–12 individual mussels were sampled, the flesh was pooled, frozen, and couriered on ice to Symbio Laboratories Sydney, a National Association of Testing Authorities (NATA) accredited commercial laboratory for initial screening using HPLC. Shellfish flesh was homogenized, and 5 g was added to 3 mL of 1% acetic acid and boiled (100 °C) for 20 min in a water bath. The sample was allowed to cool, and centrifugation was subsequently performed for 10 min at 1000 g. The supernatant was then collected, and the remaining pellet was resuspended with 3 mL of 1% acetic acid. This prepared sample was centrifuged again at 1000 g to separate the supernatant, which was then mixed with water to get a final volume of 10 mL. An SPE C18 cartridge was used to perform the cleanup of this mixture. Standards, PST positive reference matrices, and samples were oxidized with a matrix modifier. After periodate oxidation, screening of the PST analogues including STX, GTX2, 3, C1, 2, GTX5, NEO, dcNEO, and GTX1, 4, was performed. If a positive result was reported, precolumn oxidation was used to confirm concentrations of STX, GTX2, 3, C1, 2, GTX5, dcSTX, dcGTX2, 3, NEO, dcNEO, GTX1,4, C3,4. AST (domoic acid (DA)), DSTs (OA, dinophysistoxin 1 (DTX-1), dinophysistoxin 2 (DTX-2)), and pectenotoxin 2 (PTX-2) were analyzed by liquid chromatography tandem mass spectrometry (LC-MS/MS) (AB ScieX Triple Quad 6500).34,35 Positive toxin results were reported by the Tasmanian Shellfish Quality Assurance Program as equivalent to ≥1.00 mg/kg DA (AST), ≥0.25 mg/kg OA equivalents (DSTs), and ≥0.10 mg STX eq/kg (PSTs).

Pearson correlations were conducted of sxtA4 in seawater and PST concentrations in mussel tissue with no lag time, a lag time of 1, 2, and 3 weeks for the period from May 2016 to October 2017 using GraphPad Prism 7.04 (Graphpad Software Inc.).

2.5. Workflow Case Study: Spatial Patterns of an A. catenella Bloom in Southeast Tasmania

2.5.1. Sample Collection

Seawater samples were collected from August 26 to 28, 2016, on board the RV Southern Cross, along inshore–offshore transects on the east coast of Tasmania (−42 S, 148 E) (Table S1). At each of the 18 stations, a Seabird SBE 19 PlusV2 CTD (Sea-Bird Scientific, Washington) was used to measure temperature (°C), salinity (ppt), and pH, and a 10 L Niskin bottle was used to collect water samples from each depth. From each of these 10 L samples, 3 L of seawater was subsequently collected and filtered using the phytoxigene portable water sampler as previously described. Concentrated samples were then stored in 50 mL tubes for ∼4 °C for several hours before being syringe-filtered onto triplicate filters. The filters were then transferred to 2 mL cryogenic tubes, and 1 mL of Longmire’s buffer was added to each. Samples were stored at room temperature until further downstream processing.

2.5.2. Light Microscopy Cell Identification and Counting

From each 10 L Niskin water sample collected, a 1 L subsample was collected and preserved with Lugol’s iodine (final concentration of 1%). Each of these 1 L samples was then concentrated using a sedimentation technique. For this, each sample was allowed to stand for 48 h before the supernatant was siphoned off, leaving a final volume of 10 mL. The remaining 10 mL was then well mixed and settled again in a 5 cm diameter Petri dish, and the entire contents were enumerated for A. catenella cells with an inverted light stereomicroscope at a magnification of ×200 (Leica, Wetzlar, Germany).

2.5.3. DNA Extraction and Gene Quantification

Samples concentrated using the sampler were extracted using a FastDNA Spin kit for Soil (MP Biomedicals, Solon, OH) and processed using the sampling workflow previously described before running the Phytoxigene DinoDtec qPCR assay.

3. Results and Discussion

3.1. Cell Recovery Using the Portable Water Sampler

The gravity sampler was designed to be used in the field and without electricity as it does not require the use of a pump. The sampler collects and concentrates 3 L of water, which is much more than the volume of current phytoplankton monitoring protocols used in Tasmanian monitoring marine biotoxin programs.36 This high-sample volume reduces the potential for random sampling error at early bloom development concentrations of A. catenella, which can lead to PST accumulation in mussels. The effectiveness of the sampler was demonstrated by the >90% recovery of A. catenella at all concentrations tested (100, 1000, and 5000 cells per 3 L) (Table S2). Cell recovery was lowest when the sampler filtered ∼100 cells (∼92%) and highest when recovering ∼1000 cells (∼95%). The slightly higher cell loss at low concentrations may be due to the attachment of cells to the prefilter membrane or increasingly a low precision of Sedgewick–Rafter chambers at low cell concentration.20,37,38

With a mesh size of 11 μm, the sampler also concentrates phytoplankton species responsible for diarrhetic shellfish poisoning (DSP) and amnesic shellfish poisoning (ASP), in addition to the target PSP-associated microalgae. With the availability of assays specifically designed for the detection of the toxic genera Pseudonitzschia spp.39−42 and Dinophysis spp.,43,44 the sampler device would be an effective tool for sampling other HAB taxa. However, additional work to validate species recovery would be necessary before on-farm use.

3.2. Performance of Cell Lysis for DNA Extraction and Preservation

To test the efficiency of the cell lysis in our workflow, a qPCR standard curve was developed using triplicate, 10-fold 50% serial dilutions of A. pacificum. The efficiency of the assay was 99.07%, which was deemed acceptable (Figure 2).

Figure 2.

Figure 2

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Standard curve of the DinoDtec assay using lysed cells of A. pacificum strain MMWA 83 as the template DNA, showing the quantification cycle (y-axis) versus the known cell number in log-scale (x-axis).

The Cq of the sample preserved with Longmire’s buffer was not significantly different (one-tailed t test p = 0.037, d.f = 3) from the nonpreserved sample (Table S3). The Cqs from the samples preserved with RNALater and Lugol’s were 4–6.5 units higher than those of the nonpreserved sample. Samples preserved in all three buffers did not amplify when processed using BioGX lysis tubes.

One of the main challenges in analyzing environmental samples is sample matrix effects that interfere with the quantification of the analyte of interest.45 Environmental samples contain a variable range and concentration of substances that coextract with DNA and inhibit the efficiency of the PCR amplification process. To address this issue of inhibition, a well-equipped molecular laboratory has DNA extraction protocols that can be applied to remove the inhibitory compounds from field samples.46,47 However, this is not the case with the DinoDtec assay when used in situ. The use of a simple bead lysis to break open cells, instead of conducting a full DNA extraction combined with centrifugation to precipitate and remove the cell debris before the qPCR amplification, can be an alternative method. Additionally, we have shown that Longmire’s buffer does not inhibit the qPCR reaction (see Figure 2); however, it should not be added directly to the lysis tube, but instead, the membrane filter should be removed from the buffer and transferred to a lysis tube immediately prior to qPCR. Longmire’s buffer has been shown to be an effective preservative for environmental DNA, and our study supports this as an alternative to refrigeration or freezing of environmental samples.

3.3. DinoDtec qPCR Specificity and Efficiency Test

The DinoDtec assay was found to be specific to PST-producing species, as it did not amplify DNA from any of the nonproducing species tested (Table S4). The standard curves of DNA extracted from multiple strains of three different species of PST-producing Alexandrium spp. were within an acceptable range of 90–110%, with a regression coefficient of 0.99 or higher (Table 1).

Table 1. Standard Curve Efficiencies of the Phytoxigene DinoDtec qPCR Assay with Different Species and Strains of Alexandrium.

species strain % efficiency R2
A. pacificum CS300 93.72 1.000
  CAWD44 95.75 0.999
  ACTRA02 90.74 0.999
  CS798 95.54 0.999
  CS313 106.89 0.997
  CS315 100.69 0.999
A. minutum CS324 95.45 0.991
A. catenella TRIA-E 95.31 0.999
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Previous studies have shown the specificity and utility of assays targeting sxt4A for detecting PST-producing dinoflagellate species in the marine environment.21,24,26,48−50 The DinoDtec assay is in a lyophilized form, enabling long-term storage at room temperature and transport into the field. It contains an enzyme, probe, primers, dNTP, and internal amplification control (IAC) target. In terms of phytoplankton monitoring, the impact of false positives could be significant, as it could trigger unnecessary closures and delays in the harvest of shellfish products. Our data show that the DinoDtec assay is efficient for three species of PST-producing Alexandrium, and while some variation was observed between strains of A. pacificum, the efficiency remained within acceptable limits (90–110%).

3.4. On-Farm Workflow Testing

During the 2016 A. catenella bloom, the number of sxtA4 copies L–1 began increasing at Spring Bay on 3 July 2016 (Figure 3). The sxtA4 copies of L–1 then decreased to almost zero on 17 July 17, 2016, and increased again in the following week. The PST levels in mussels started to increase during the week starting on 31st July 2016, which is approximately 4 weeks after the first sxtA4 copies L–1 were observed. The PST concentration then increased until the week starting on September 25, 2016, after which it continuously decreased (Figure 3).

Figure 3.

Figure 3

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Copy number of sxtA4 L–1 in seawater and total concentration of PST in mussel tissue (mg kg–1) during the 2016 A. catenella bloom from Spring Bay Seafoods. The inset shows a portion of the same data from June to July 2016, with sxtA4 L–1 in seawater on a log-scale, highlighting that the qPCR assay detected significant sxtA4 copies prior to the detection of toxins in mussel tissue.

In 2017, the rapid increase in sxtA4 copies L–1 number started on the week of the 21st Aug 2017, and the increase continued until 2 Oct 2017 (Figure 4). PST concentrations began to elevate during the week commencing on 5 Sept 2017. The PST concentration reached its peak on 10 Oct 2017 (140 mg kg–1), after which it gradually declined. This coincided with decreasing numbers of sxtA4 copies of L–1 during this time. As cell numbers were extremely high during the week starting on 10 Oct 2017 and 17 Oct 2017, Spring Bay Seafoods was not operational and sampling was suspended. Hence, no qPCR data were obtained.

Figure 4.

Figure 4

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Dynamics of sxtA4 L–1 copies in seawater and the total concentration of PST in mussel tissue during the 2017 A. catenella bloom from Spring Bay Seafoods.

Our correlation analyses of sxtA4 L–1 and PST mg kg–1 in mussels demonstrate the use of the DinoDtec qPCR assay as an early warning indicator of PST in Tasmanian commercial mussels, M. galloprovincialis (Table S5). Over the course of two A. catenella blooms indicates that the DinoDtec workflow provides a 2 week early warning of PST (r = 0.95, Table S5). Additionally, a Pearson correlation coefficient with a lower r value was also found when modeling the sxtA4 copies L–1 and PST mg kg–1 in mussels, suggesting that up to 3 weeks’ early warning may be possible (r = 0.7, Table S5). Our results with M. galloprovincialis are consistent with previous observations showing an approximately 2 week lag between the addition of PST-producing dinoflagellates to the water and accumulation of toxin in Mytilus spp.51

The rate of PST accumulation and depuration may also differ depending on factors such as the ploidy level of the shellfish and the ambient water temperature.15 Depuration in M. galloprovincialis is additionally influenced by size, age, soft tissue weight, and reproductive stage.52 For example, increased acidification and increased temperature could potentially cause lower maximum PST accumulation but slower depuration in M. galloprovincialis.53 Farmed mussels are grown from the same batch, which means that they have a similar size, age, and weight, potentially minimizing differences among individuals. Given the differences among species and ambient environmental factors, it is likely that the optimal lag time established between sxtA4 copies L–1 and PST mg kg –1 established in this study will be specific to mussels impacted by A. catenella during the austral winter, which, on the east coast of Tasmania, is generally characterized by water temperatures of 10–15 °C.

Toxin accumulation and detoxification dynamics of shellfish are complex and depend on many factors to modify uptake processes (e.g., filtration and feeding rate/efficiency, particle size, toxin distribution in the planktonic food web). As a result, PST uptake, PST biotransformation, and depuration rates can differ greatly between shellfish species.15,54−57 In green-lipped mussel (Perna viridis), the accumulation of PST can exceed the regulatory level within 2 days of exposure, with a significant amount of toxins removed within 3 weeks.56 Blue mussels (Mytilus edulis) accumulate toxins quickly and show limited toxin metabolism (and thus are useful indicators of the toxigenic source) and can take weeks to detoxify.54 Australian abalone uptake PST at about 10 times slower rate than mussels but with comparable depuration rates.58 In scallops, the accumulation and depuration of toxins are much slower than mussels and can take up to several months.54 In Pacific oysters (Crassostrea gigas), a PST level higher than the regulatory limit can be reached in 3 weeks when the concentration of A. minutum is between 9 and 140 cell L–1 in the surrounding water.59

3.5. Spatial Patterns of an A. catenella Bloom

During August 2016, the surface abundance of sxtA4 copies was highly variable across sites, with no consistent pattern with distance from shore in any sample zone (Figure 5). The highest sxtA4 copy abundance (in surface waters) was observed in Spring Bay (maximum abundance of 2764 sxtA4 copies L–1 at SB3), Coles Bay (max. abundance of 2017 sxtA4 copies L–1 at CB4), and Little Swanport (max. abundance 1939 sxtA4 copies L–1 at OB4).

Figure 5.

Figure 5

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Sampling stations of the RV Southern Cross survey along the east coast of Tasmania, Australia, with the concentration of surface-sxtA4 copy L–1 detected at each site during the peak (26–28 August) of the A. catenella bloom in 2016.

At the deeper (>30 m) stations of SB3 and OB1, A. catenella cell abundance was noticeably stratified, with maximum cell density observed between 5 and 10 m and decreasing below 10 m (Figure S1A,B). Salinity was uniform with depth at these sites; however, temperature stratification was evident (0.2–0.4 °C) (oceanographic data presented in Condie et al.). For stations that were shallower (<10 m), the highest number of cells were found at a depth of ∼5 m (Figure S1C,D). In estuaries (<5 m), the A. catenella cell numbers did not vary greatly between the surface and ∼3 m (Figure S1E–G). These patterns were consistent with both qPCR and light microscopy results, with the exception of two sites (CB1 and CB3), which showed variability between methods (Figure S1D).

3.6. Implications for Monitoring Strategies

During the 2016 and 2017 A. catenella blooms in Tasmania, the detection of sxtA4 copies L–1 was used to make decisions regarding mussel harvest management of Spring Bay Seafoods, prompting voluntary harvest closures prior to official PST detection and harvest closure notice from the TSQAP. Advantages of this early detection included capacity to switch harvest/production to alternative sites not impacted by PSTs and continued production throughout primary site closure periods. Declining sxtA4 copy number also provided an early indication of the bloom decline, allowing farmers to continue to source production from the alternative site until the PST level decreased to below the regulatory limit (0.8 mg kg–1).

During 2016, the decrease of sxtA4 copies L–1 occurred from the week starting on September 25, 2016 (Figure S1), while PST mg kg–1 in mussel tissue was still above the regulatory limit. While weekly samples were adequate to inform management decisions and modeling, more frequent sampling may provide information on finer-scale dynamic shifts in A. catenella and improve the resolution of the results.

The vertical profile of A. catenella cell abundance across depths during the August 2016 bloom event indicated that cell abundance was well mixed throughout the water column in estuaries but became stratified in oceanic conditions (Figure S1). At these later stations, the highest A. catenella cell abundance and sxtA4 copies were observed at a depth of 5–10 m below the sea level, decreasing in deeper samples. Such a vertical trend was also observed in bloom in the Gulf of Maine in 2001,60 where the highest concentration of cells was found in a thin layer depth of ∼10–11 m. The maximum cell abundance in the present study was linked to stratification in temperature and chlorophyll fluorescence, significant for the vertical migration of A. catenella.61 The vertical distribution of Alexandrium implies therefore that sample depth, stratification, and time of day are important considerations for cell quantification in oceanic systems.

The spatial distribution of sxtA4 copies L–1 appeared to be patchy in August 2017 (Figure 5), and no discernible pattern was observed between oceanic and estuarine stations. It is likely that factors such as upwelling and downwelling, rainfall, and current flow affected the spatial distribution of the bloom.61 As qPCR is relatively fast to implement, additional sampling stations and replicates can be used by monitoring agencies to increase accuracy and prediction in such circumstances.

To conclude, we have established a three-step tool for the detection and quantification of PST-producing dinoflagellates that is applicable to use on site. This protocol is standardized, relatively simple, and inexpensive to operate and not in need of specialized laboratory facilities. The cell recovery of the sampler was found to be high, and the lysis method for DNA extraction and the DinoDtec assay were shown to be sensitive, specific, and efficient. We successfully used this protocol to identify and describe two significant blooms of A. catenella in Tasmania, Australia, with our results suggesting that these blooms are well mixed in estuaries but appear to be subsurface in deeper, more oceanic sites, suggesting important implications for future monitoring strategies.

Acknowledgments

The authors would like to acknowledge the assistance of Dr Javier Pérez for Figure 5. This research was funded by a UTS PhD Scholarship (Dr Rendy Ruvindy), the Australian Research Council for a Future Fellowship (Murray FT120100704) and Discovery Grant (CIs Murray and John DP120103199), and the Australian Government through the Fisheries Research and Development Corporation Project No. 2014/032: Improved understanding of Tasmanian harmful algal blooms and biotoxin events to support seafood risk management (CIs Hallegraeff, Bolch, Murray, Turnbull). Finally, the authors thank Andreas Seger, David Faloon, and Justin Hulls for their field assistance.

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.est.3c10502.

  • Sampling sites/depths; results of cell recovery using a sampler; preservative agent; qPCR specificity and relationship between sxtA4 copies L –1 and PSTs (mg kg–1) in M. galloprovincialis (Tables S1–S5); and field quantification of A. catenella (Figure S1) (PDF)

The authors declare no competing financial interest.

Supplementary Material

es3c10502_si_001.pdf (123.8KB, pdf)

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