Analytical methods for assessment of cyanotoxin contamination in drinking water sources

Accepted Manuscript Analytical Methods for Assessment of Cyanotoxin Contamination in Drinking Water Sources Marcela Jaramillo, Kevin E. O’Shea PII:

S2468-5844(18)30076-X

DOI:

https://doi.org/10.1016/j.coesh.2018.10.003

Reference:

COESH 78

To appear in:

Current Opinion in Environmental Science & Health

Received Date: 20 July 2018 Revised Date:

22 October 2018

Accepted Date: 23 October 2018

Please cite this article as: Jaramillo M, O’Shea KE, Analytical Methods for Assessment of Cyanotoxin Contamination in Drinking Water Sources, Current Opinion in Environmental Science & Health, https:// doi.org/10.1016/j.coesh.2018.10.003. This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

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Analytical Methods for Assessment of Cyanotoxin Contamination in Drinking Water Sources Marcela Jaramillo and Kevin E. O’Shea

Miami, FL 33199, USA Abstract:

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Department of Chemistry and Biochemistry, Florida International University, University Park,

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Contamination of drinking water sources by cyanotoxins, including microcystins (MCs), nodularins (NOD), cylindrospermopsin (CYN), anatoxin-a (ANA), homoanatoxin-a (HANA)

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and beta-N-methylamino-L-alanine (BMAA), produced during harmful algal blooms is a serious threat to human health. The detection, quantitation, and monitoring of cyanotoxins is essential to ensure public safety. A variety of indirect methods including enzyme-linked immunosorbent assay (ELISA), antibody-based techniques, and molecular approaches are available for

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cyanotoxin detection. While robust and simple, the indirect methods often lack selectivity and sensitivity. Direct detection using chromatographic techniques coupled to a mass spectrometer provide excellent selectivity and sensitivity, but require costly equipment and skilled operation.

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Matrix-assisted laser desorption ionization time of flight mass spectrometry coupled to a bioaccumulator, toxin chemical derivatization, electrochemical immunosensors, quantitative

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polymerase chain reaction of toxigenic genes, and specific aptamer recognition have also been demonstrated for cyanotoxin analysis.

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Highlights:

Cyanotoxins can cause serious drinking water contamination

Detection and quantitation can be achieved by indirect and direct methods Direct methods based on LC-MS provide high selectivity and sensitivity

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Toxic gene analyses is emerging as a forecasting tool for cyanotoxin contamination Keywords:

Introduction:

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Cyanotoxin Contamination, Water Quality, Analyses, Detection, Quantitation, Monitoring.

The presence of cyanotoxins produced by harmful algal blooms (HABs) is a serious threat to drinking water sources and an urgent global challenge [1,2]. Continued eutrophication

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and climate change have led to significant increases in HABs and the associated cyanotoxin contamination of drinking water sources [2–4]. The ingestion of potent cyanotoxins poses

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serious health hazards for humans, including increased cancer risks [5–8]. The Great Lakes, which provide drinking water for more than 35 million people, have been inundated by HABs in recent years, causing closure of public drinking water sources and unsafe conditions [9]. The state of California has been affected by numerous HAB events [2]. On June 20th 2018, a state of

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emergency was declared in eight Florida counties due to massive HABs [10]. The United States Environmental Protection Agency (USEPA) has included a number of cyanotoxins produced during HABs in the

Drinking Water Contaminant Candidate List 4 (CCL-4)[11]. For the

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common cyanotoxins, microcystins (MCs) and cylindrospermopsin (CYN), the US EPA has issued ten day health advisory (HA) values for drinking water. For children under 6 years old the

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recommendations are not to exceed 0.3 and 0.7 µg/L for MC and CYN, respectively, while for older children and adults the values are ≤ 1.6 µg /L for MC and ≤ 3.0 µg /L for CYN [12,13]. The HA for MC was developed based the specific MC variant, MC-LR, as a surrogate for all MCs [12,13].

Accurate assessment of the health risks of exposure to cyanotoxins requires unequivocal identification and quantitative analyses of their presence in drinking water sources. The methods

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employed for cyanotoxin analyses must be economical, rapid, and highly sensitive. Commercially available enzyme inhibition, enzyme linked immunoassays (ELISAs) and receptor binding assays (RBAs) allow the indirect detection of cyanotoxins [3]. These techniques are of

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modest cost, simple, and rapid, but are semiquantitative, and also suffer from matrix interferences and cross-reactivity, which can lead to inaccurate and unreliable results [3]. Quantitative polymerase chain reaction (qPCR) approaches allow the fast, reproducible and

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sensitive identification, and quantitation of target genes present in toxigenic algal species, but subsequent forecasting of cyanotoxin production in the water is often problematic [14–16]. A

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variety of molecular-based strategies, including antibodies, enzyme inhibition, aptamers, and molecularly imprinted polymers, have been developed for detection of cyanotoxins [3]. However, biosensors for freshwater cyanotoxin detection are still not commercially viable [3]. Ultraviolet (UV) or mass spectrometric (MS) detection coupled to liquid chromatographic (LC)

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separation are powerful tools for the separation and identification of a wide array of cyanotoxins [1,3]. MS detection provides detailed structural information critical to the identification of individual cyanotoxins with extremely low limits of detection (LOD), 1-10 ng L−1 [17,18]. UV

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detection is more routine and less costly compared to MS, but also less sensitive and less selective [1,3]. The lack and/or expense of reliable cyanotoxin standards is a major challenge to

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their quantitation using these techniques [19]. Herein is a summary of recent methods used to predict production, identify, and quantitate the common freshwater cyanotoxins: MCs, nodularins (NOD), CYN, anatoxin-a (ANA), homoanatoxin-a (HANA) and beta-Nmethylamino-L-alanine (BMAA) shown below, figure 1.

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Microcystin, X and Y are two variable L-amino acids O

O

COOH

N H NH Y H N

NH H2N O

N X

HN

CH2 O

O

homoanatoxin-a

O

HN

NH

O

O

BMAA

Cylindrospermopsin

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O

O O S O O

OH

O

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H 2N

O

HOOC

Red: Adda functionality O

HN

COOH

N

O

H2N

N H

O

HOOC

Anatoxin-a

H N

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O

Nodularin-R

N

NH

NH HN

O N H

OH NH2

NH O

Figure 1. Structures of Common Cyanotoxins

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Microcystins (MCs) and Nodularins (NODs): While the presence of MC and NOD has been reported in food, recreational waters, and nutritional supplements, the most common route of exposure is from drinking water [1,3]. A

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notable example of the closure of a municipal drinking water facility due to cyanotoxin contamination, occurred in Toledo, Ohio with a “do not drink order” issued to approximately

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500,000 people [9]. In Brazil, intravenous solutions tainted with MC led to the death of 60 hospital patients [5].

MCs and NOD are highly stable cyclic peptides, which all contain the unique ??-amino

acid, (2S, 3S, 8S, 9S)-3-amino-9-methoxy-2,6,8-trimethyl-10-phenyldeca-4E,6E-dienoic acid, often referred to as Adda [20]. MC, toxic at trace concentrations comparable to cyanide, is the most common class of freshwater cyanotoxins [1]. More than 120 MC variants have been

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identified which vary in potency dependent on exchange of one or two amino acids (X and Y in Figure 1), with most common and toxic variant being MC-LR [21]. The median lethal dose (LD50) of MC-LR by intraperitoneal route is 25–150 µg kg-1 of body weight in mice [22,23].

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The world health organization (WHO) guideline for MC-LR in drinking water is 1 μg L-1 [21].

MCs are often monitored by health and drinking water management agencies [24].

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Advancement of analytical techniques continue to result in the identification of new MC variants [25]. LC-MS methods allow the simultaneous detection and quantitation of MC mixtures with

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insignificant matrix interferences and low LODs (0.003 to 0.030 µg L-1) [18]. Protein phosphatase enzyme inhibition (PPI) assays are commonly employed and are also highly sensitive for detection of MCs (LOD of 0.16 µg L-1) [3]. Variations in enzyme purity and the instability of enzyme dimers in solution are the major limitations of the PPI assays [26]. Employment of an engineered phosphatase catalytic subunit [26] or the immobilization of the

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enzyme in an agarose gel have been reported to improve the accuracy of the assay while producing reproducible results [27]. PPI assays are still, however primarily used in the laboratory

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and yet to be developed for regular drinking water monitoring [3].

Antibody-based test strips provide a robust, cost-effective, and simple method for initial

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risk assessment of drinking water supplies [24]. The performance of commercially available test strips compares favorably to ELISA and PPI in the detection of MC concentrations greater than 10 µg L-1, the regulatory level for recreational waters [24]. However, for drinking water providers, the limited reliability of the test strips at lower concentrations makes them unsuitable until they can be rigorously verified [24].

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Gold nanoparticles functionalized with polypyrrole microsphere (AuNP/PPyMS) have been reported for the electrochemical immunosensor detection of MC-LR (LOD of 0.1 ng L-1) without the interference from the isomeric forms of MC-LR [28]. The precision, reproducibility

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and stability of the AuNP/PPyMS technique were reportedly acceptable for a real contaminated water source [28], but the general application may be limited by costs.

Matrix assisted laser desorption ionization time of flight MS (MALDI-TOF MS) is

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unique for the analysis of MCs in biological and environmental matrices [29]. MALDI-TOF MS was successfully applied to mussels exposed to 0.45 µg MC L-1` with accurate detection of MC

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within one day of exposure to the toxin [29]. This strategy allows for direct MC analysis in complex matrices as an early MC contamination warning system, important to water and food safety.

The concentration of unique genes of toxigenic strains of algae in drinking water sources

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determined by qPCR was used to reasonably forecast MC production during a HAB [30]. This approach allows time for local authorities to take precautionary steps by employing proactive strategies to inhibit the growth of the algal bloom and prevent the toxin from reaching drinking

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water sources and food supplies.

Anatoxin-a (ANA)

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The cyanotoxin ANA and its methylated homologue homoanatoxin-a (HANA) are

bicyclic secondary amine alkaloids (see Figure 1) [31]. ANA and HANA, highly water soluble cations, are among the smallest of the freshwater cyanotoxins [3]. ANA is a strong neurotoxin and affects the acetylcholine receptors [31]. The symptoms of ANA intoxication are sudden muscle contraction, convulsions, and rapid death by asphyxiation [31]. The WHO, Australia,

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New Zealand, Denmark, and US agencies have proposed regulatory values between 0.7 and 20 µg L−1 for ANA in drinking water [32,33]. The early detection, quantitation and monitoring of the deadly ANA is imperative. ANA

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concentrations from 0.05 to 1929 µg L−1 have been detected in US freshwaters [34,35]. ANA has been detected using gas chromatography (GC) with electron capture detection [36] and GC–MS [37]. The chemical derivatization with 4-fluoro-7-nitro-2,1,3-benzoxadiazol followed by LC

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coupled to fluorescence detector can be used to quantitate ANA and HANA in water [38]. This method was also adapted to analyze the toxins in biological matrices, fish and mussels, with

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recoveries of at least 71 % [39]. The EPA Method 545 describes the validated LC electrospray ionization tandem MS (LC-ESI-MS/MS) approach for the detection of ANA in drinking water [34].

Sample matrix effects on the electrospray ionization efficiency are a major challenge for

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the LC-MS quantitation of algal toxins [40]. The stable isotope dilution approach has been used to mitigate these matrix effects [40]. Unfortunately, isotopically labeled standards for algal toxins are expensive or not available. The proof of concept of isotope-labeling employing

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derivatization with dansyl-d6 chloride for the quantitation of ANA and HANA has been demonstrated [19]. A combination of reverse-phase and weak anion exchange chromatographic

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techniques were used to extract the derivatized ANA and HANA with LOD of 20 and 40 ng g−1, respectively [19]. The methodology may be applied to the quantitation of other algal toxins for which isotopically-labeled analogs are not available. Anatoxin-a(s) inhibits acetylcholinesterase [41]. This property has been used for

constructing biosensors for detection of anatoxin [42]. Biosensor technologies are limited by the

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sensitivity and specificity when compared to other analytical techniques [43]. These limitations were improved using an engineered acetylcholinesterase (LOD of 0.5 nM) [43]. Aptamer-based materials for molecular sensing and detection have been accomplished

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with modified gold nanoparticles (AuNPs) hybridized to a polymeric biosensor to produce an amplified “signal cascade” [44]. While still in the developmental stages, proof of principle provides an ultrahigh sensitive biosensing system with a LOD of 10−14 M, among the highest

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sensitivity reported [44].

A combination of monoclonal antibodies and magnetic Sepharose-based beads to

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preconcentrate ANA has been demonstrated [33]. The bead extract was analyzed with ion mobility spectrometry (IMS) with reported LOD and LOQ values of 0.02 and 0.08 µg L−1, respectively, and relative standard deviation less than 15 % [33]. This methodology approaches the sensitivity and selectivity of LC–MS/MS, but at a significantly lower cost [33].

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Species-specific primers were developed for the identification and cell quantification of ANA producer, Tychonema bourrellyi, in environmental samples using PCR and qPCR [14]. The assay was validated with real environmental samples. PCR, qPCR and light microscopy (LM)

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highly correlated the cell densities to ANA concentrations in water. This molecular approach allows the early, sensitive, and simple monitoring of T. bourrellyi, an appropriate indicator for

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ANA contamination risk [14].

Cylindrospermopsin (CYN)

The CYN is a tricyclic alkaloid toxin with neuro-, cyto-, dermato-, hepato-toxicities and

possible carcinogenic properties [45]. The uracil and hydroxyl functionalities are responsible for CYN toxicity (see figure 1.) [3,45]. CYN contamination of a drinking source that lead to the hospitalization of 140 children suffering from gastroenteritis in Australia in 1979, referred to as

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the Palm Island Mystery [7]. The health alert in the Australian Drinking Water Guidelines for CYN is 1 µg L-1 [46]. ELISA kits, commercially available from Abraxis and Beacon, are capable of detecting

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CYN with LODs of 0.04 and 0.1 µg L-1, respectively [47]. The LC-ESI-MS/MS EPA method 545 is commonly used for the detection of CYN [34]. CYN is zwitterionic at natural pH and not effectively concentrated by standard solid-phase extraction (SPE) [17]. However, the SPE ENVI-

analysis [17]. 15

N ammonium chloride was recently achieved

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The total synthesis of [15N5]-CYN from

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Carb cartridges are reported to successfully retain CYN for Ultra Performance (UP) LC-MS/MS

providing an isotopically labeled standard [48]. The standard was used for the precise quantification of this potent biotoxin by isotope dilution MS with LOD and LOQ of 2.5 and 8.25 ng L-1 respectively, the highest sensitivity reported to date [48]. The matrix effect was evaluated

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by using CYN free environmental water samples spiked with 100 µg L-1 CYN yielding recoveries of nearly 100 % [48]. The method was used to analyze real environmental water samples and led to the first report of cyanotoxin contamination in Lake Casitas, California [48].

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A correlation between the concentration of genes specific to CYN-producing species

quantified by Q-PCR and cell counts by microscopy was found in both water samples and

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cultured strains [49]. This assay is fast, sensitive, and promising for the monitoring of CYNproducing cyanobacteria [50] and assessing the toxicity potential of the cyanobacteria [51]. Beta-N-methylamino-L-alanine (BMAA).

BMAA, discovered approximately 50 years ago, has only recently gained attention as a

HAB product [52]. BMAA, a non-encoded amino acid, has been associated with the human neurodegenerative disease Amyotrophic Lateral Sclerosis/Parkinsonism Dementia Complex (ALS/PDC) [52]. A notorious example of the BMAA devastating neurological impact was

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reported in Guatemala, where indigenous Chamorro people regularly consume flying foxes (Pteropus mariannus) which contain high levels of bioaccumulated BMAA (up to 4 mg g -1) [8]. Mean concentrations of BMAA found in brain tissues of Chamorros who had died of ALS/PDC -1

[8]. ALS/PDC was 50-100 times more prevalent in the

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type symptoms were 6.6 µg g

Chamorro people than in another place [8]. BMAA, along with its constitutional isomers, 2,4diaminobutyric acid (DAB) and N-(2-aminoethyl) glycine (AEG), ANA and CYN were detected

metal ions which complicates the MS analysis [54].

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in natural water samples at cyanobacteria blooms sites [53]. BMAA has been proposed to chelate

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The LC MS analyses of BMAA were compared using derivatization of BMAA with 6aminoquinolyl-N-hydroxysuccinimidyl carbamate and propyl chloroformate, and underivatized BMAA employing hydrophilic interaction liquid chromatography (HILIC) [55]. These methods yield consistent BMAA detection in cyanobacteria cultures [55].

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BMAA and DAB and AEG, detection along with ANA and CYN was achieved by coupling ultra-high performance LC and a Q-exactive MS (UHPLC-HESI-HRMS) [53] equipped with a heated electrospray ionization source [53]. SPE was used for toxin purification and

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preconcentration prior LC analysis [53]. LC separation was improved by chemical derivatization with dansyl chloride [53]. The assay was validated with good linearity (R 2  > 0.998) using the

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internal calibration of DAB-d3, and LOD and LOQ from 0.007 to 0.01 µg L−1 and from 0.02 to 0.04 µg L−1, respectively, were achieved [53]. BMAA analysis in complex biological matrices was reported using

capillary

electrophoresis (CE) MS employing the stable isotope labeled (13C3,15N2) BMAA (SIL-BMAA) with LOD and LOQ of 40 nM (4.8 µg L-1) and 400 nM (48 µg L-1), respectively [56]. BMAA was detected in the in the protein hydrolyzed fractions of a biological matrix, indicating that

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BMAA is associated and/or incorporated into proteins [56]. SIL-BMAA is commercially available from Cambridge Isotope ($1500 per 10 mg). The isotope dilution method MS analysis

Multi-Class Toxin Approaches

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of BMAA in water samples using the SIL-BMAA is plausible, but has not been reported.

A number of LC-MS methods have been developed for the simultaneous detection of different cyanotoxins. An optimized SPE LC–tandem MS (LC–MS/MS) method was reported

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for detection and quantitation of CYN, ANA, NOD, twelve MC variants and two marine cyanotoxins in a single run (LODs from 1 to 10 ng L−1) [45]. The preconcentration step was

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optimized in terms of SPE cartridge, sequence of cartridges, pH, composition and volume of the elution solvent, composition of the reconstitution solvent, and sonication time of the final extract [45]. The method was successfully used to analyze real lake water samples [45]. In a similar study, UPLC–MS/MS was used after SPE preconcentration, to analyze 6 MC variants, NOD,

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CYN, ANA and a marine toxin [17]. UPLC-MS/MS was, also, reported for the simultaneous analysis of 22 cyanotoxins including ANA, CYN, NOD and MC in raw and drinking water [57]. The rapid, sensitive, and accurate identification and quantification of cyanotoxin mixtures was

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achieved by using reference standards and dynamic multiple reaction monitoring mode MS [57]. Conclusions:

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Common exposure to cyanotoxins can occur through food, recreational waters, and

nutritional supplements. However, the most common route is through consumption of contaminated drinking water. With increases in the size, frequency, and persistence of HABs leading to cyanotoxin contamination, the development of economical, highly-selective, highlysensitive, rapid, and simple analytical methods is critical for ensuring the safety of drinking water sources. Recent innovative efforts have been dedicated to improving detection and quantitation of freshwater cyanotoxins. Biological-based methods offer promising prospects for

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economical rapid detection of a number of common cyanotoxins. Although these methods are widely used in laboratory settings, real-life applications are often complicated by non-selectivity, interferences from the sample matrix and/or lack of reproducibility. Several genetic, bloom

cyanotoxin outbreaks, but suffer from reliability.

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dynamics, and PCR-type methods offer predictive options to help prevent and plan for HAB and

Many ground-breaking hybrid approaches capitalizing on the advantages of biological

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systems coupled to the sensitivity and selectivity of instrumental methods have been demonstrated, but may be cost prohibitive and require further refinement for widespread

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application. While biological based, hybrid and instrumental methods continue to evolve, the availability and cost of certified standards and isotopically labeled surrogates limit their development and economical application. To date, toxin preconcentration by means of SPE coupled to chromatographic separation with MS detection remains among the most powerful

mixtures.

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analytical tool for identification and quantitation of freshwater cyanotoxins individually and as

Acknowledgements and Disclaimers

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Marcela Jaramillo gratefully acknowledges support by the National Science Foundation Graduate Research Fellowship under Grant No. (NSF grant number: DGE-1610348). Any

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opinion, findings, and conclusions or recommendations expressed in this material are those of the authors(s) and do not necessarily reflect the views of the National Science Foundation. References

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