Analysis of Cyanobacterial-Derived SaxitoxinsUsing High-Performance Ion ExchangeChromatography with ChemicalOxidation/Fluorescence Detection

John Papageorgiou, Brenton C. Nicholson, Thomas A. Linke, Con Kapralos

Australian Water Quality Centre, South Australian Water Corporation, Salisbury,South Australia, 5108, Australia

Received 2 May 2005; revised 23 June 2005; accepted 27 June 2005

ABSTRACT: A single run HPLC method utilizing ion exchange as the separation mode with a novel mobilephase system coupled to chemical postcolumn oxidation and fluorescence detection has been developedand demonstrated to be applicable to the quantitative analysis of paralytic shellfish poisons (PSPs) pro-duced by Australian cyanobacteria (Anabaena circinalis) and other cyanobacteria. Both the cyanobacterialmatrix and natural water constituents did not significantly affect the performance of this method. The dailyprecision of this method was adequate for it to be considered as a routine analytical tool for direct PSPanalysis (prePSP concentration is not required) of cyanobacterial extracts and water bodies containingPSPs (C1, C2, GTX2, GTX3, NEO, STX) in the low parts per billion concentration range (10–70 ppb).# 2005 Wiley Periodicals, Inc. Environ Toxicol 20: 549–559, 2005.

Keywords: cyanobacteria; cyanobacterial paralytic shellfish poisons (PSPs); high-performance ion ex-change chromatography (HPIC); chemical oxidation; fluorescence detection; reservoir water

INTRODUCTION

It is well established that saxitoxins or paralytic shellfish

poisons (PSPs) (Fig. 1) are produced by dinoflagellates in

the marine environment and certain cyanobacteria (e.g.,Anabaena circinalis (ACN), Aphanizomenon flos-aquae,Lyngbya wollei, Cylindrospermopsis raciborskii) in fresh

waters (Onodera et al., 1997; Lagos et al., 1999; Velzeboer

et al., 2000; Ferreira et al., 2001). Some PSP analogues are

highly neurotoxic and lethal to both animals and humans at

low levels, which has necessitated the development of ana-

lytical methods for the determination of their presence in

both shellfish (shellfish concentrate PSPs by ingestion of

toxic dinoflagellates) and water.

To date, the majority of analytical methods developed

for quantitative PSP analysis are based on reverse phase

high-performance liquid chromatography (HPLC) coupled

to chemical postcolumn oxidation and fluorescence detec-

tion of the corresponding PSP derivatives (Sullivan and

Iwaoka, 1983; Oshima et al., 1989; Oshima 1995a,b). A

major disadvantage with this mode of analysis is that

anionic and cationic ion-pairing reagents are required to

effect the adsorption and separation of cyanobacterial-

derived PSP analogues (carbamate, sulfamate, decarbamoyl

saxitoxins) because of differences in their polarity and ionic

charge. This amounts to three separate analyses being

required to determine the various PSPs. Therefore, the

Oshima HPLC method (Oshima et al.,1989) is both time-

consuming and costly for PSP analysis of multiple samples.

Recently, a novel HPLC ion exchange (HPIC) based

method was developed for the analysis of various saxitoxins in

one chromatographic run (Jaime et al., 2001). With this

method, column separation of saxitoxins was effected on anion

and cation exchange columns connected in series. Eluted

Correspondence to: J. Papageorgiou; e-mail: john.papageorgiou@

sawater.com.au

Published online in Wiley InterScience (www.interscience.wiley.com).

DOI 10.1002/tox.20144

�C 2005 Wiley Periodicals, Inc.

549

toxins were detected by mass spectrometry or fluorescence of

their electrochemically generated oxidized derivatives. HPIC-

fluorescence detection (HPIC-FD) utilizing chemical postcol-

umn oxidation was not evaluated. Chemical postcolumn oxida-

tion of PSPs is usually adopted by the water industry, in PSP

analysis. Therefore, the water industry is reluctant to substitute

existing functional chemical postcolumn oxidation equipment,

to minimize costs associated with re-training of laboratory per-

sonnel and purchase of additional equipment (electrochemical

cells/detectors or expensive mass spectrometric detectors). This

article describes a single run HPIC method with postcolumn

chemical oxidation and fluorescence detection for the quantita-

tive analysis of cyanobacterial PSPs, using a typical HPLC sys-

tem currently used in laboratories throughout the water and

shellfish industries. The emphasis on this article is predomi-

nantly from an Australian perspective.

MATERIALS AND METHODS

Solvents and Chemicals

Sodium acetate of 99% purity was purchased from Aldrich

Chemical Company and recrystallized from aqueous

Fig. 1. Structures of PSPs. Toxicity data from Oshima et al. (1989).

550 PAPAGEORGIOU ET AL.

ethanol. Ammonium acetate of analytical grade and fluores-

cent-free grade sodium acetate were purchased from

Aldrich Chemical Company and Fluka Chemical Company,

respectively. Milli-Q water (Millipore Corporation, USA)

was used in the preparation of chromatography eluents.

Chromatographic materials used for the purification of C1

and C2 toxins were purchased from Bio-Rad Laboratories,

Mississauga, ON, Canada.

Fig. 3. HPIC-FD chromatogram of a synthetic mixture that contains typical cyanobacte-rial PSPs, NEO and STX (ST1), using an ammonium acetate-based mobile phase system(Jaime et al., 2001) and chemical post-column derivatization.

Fig. 2. HPIC-FD chromatogram of a synthetic Imixture that contains typical Australiancyanobacterial PSPs, NEO and STX (ST1), using a sodium acetate-based mobile phasesystem and chemical postcolumn derivatization.

551ANALYSIS OF CYANOBACTERIAL SAXITOXINS USING HPIC-FD

PSP Toxins and Standards

Saxitoxins (STX), neosaxitoxin (NEO), decarbamoyl saxi-

toxin (dcSTX), gonyautoxins 2 and 3 (GTX2/3) that con-

tained some decarbamoyl GTX2 and 3 (dcGTX2/3), and

gonyautoxins 1 and 4 (GTX1/4) were purchased from the

National Research Council, Marine Analytical Chemistry

Standards Program (NRC-PSP-1B), Halifax, Nova Scotia,

Canada. DcGTX2/3 content present in the GTX2/3 standard

was not determined by the supplier. C1 and C2 toxins (C1/

2) were extracted from a natural Australian bloom of ACN,

using the following method: 0.5 L of the bloom material in

0.05 M acetic acid was subjected to three consecutive

freeze–thaw cycles to lyse the cells. The final thawed sus-

pension was centrifuged (10,000 g � for 20 min) and C1/2

toxins were isolated from the supernatant and purified as

described by Laycock et al. (1994). It should be mentioned

that certified C1/2 toxin standards were not commercially

available at the time that this work was carried out. Two

standard PSP stock mixtures containing the following PSP

concentrations in parts per million (ppm) were prepared

from the standards purchased and isolated as described ear-

lier; C1: 0.232, C2: 0.089, GTX2: 0.155, GTX3: 0.038,

NEO: 0.175, STX: 0.175 (ST1) and C1: 0.15, C2: 0.035,

GTX1: 0.123, GTX2: 0.198, GTX3: 0.048, GTX4: 0.054,

NEO: 0.117, dcSTX: 0.084, STX: 0.117 (ST2). Calibration

PSP standard solutions were prepared from dilutions of

ST1 with Milli-Q water.

Collection and Preparation of Natural Waterand Cyanobacterial Samples

Water samples (1 L) representing varying qualities with

respect to dissolved organic content (DOC) and total dis-

solved solids (TDS) (measured as conductivity) were col-

lected from South Australian reservoirs. Water samples

(10 mL) were passed through a 0.45 �m filter to remove

suspended matter and then 1 mL aliquots were acidified

with 2.5 M acetic acid (20 �L) and spiked with 1 mL of the

standard PSP mixture (ST1), prior to being analyzed by

HPIC-FD. A water sample contaminated with ACN was

collected from Coolmunda Dam, Warwick, Queensland,

Australia, and freeze–thawed twice and the final thawed

suspension was centrifuged (10,000 g � for 20 min) to

remove insoluble cellular material. The supernatant con-

taining the PSP mixture was diluted 1:1 with Milli-Q water

and stored at �208C. The PSP mixture was further diluted

1:10 with Milli-Q water immediately before HPIC-FD

analysis.

High-Performance Ion ExchangeChromatography Coupled toFluorescence Detection

HPIC-FD analysis was performed using a Waters HPLC

system comprising a 717 autosampler, 600 E multisolvent

delivery system, 747 scanning fluorescence detector, two

reagent manager post column pumps, post column reaction

coil, post column temperature control module, and Millen-

ium 32 software.

Chromatography was performed using a Source 15Q PE

4.6/100 anion exchange column (Pharmacia Biotech,

Uppsala, Sweden) and two Source 15S PE 4.6/100 cation

exchange columns (Pharmacia Biotech) connected in series.

Saxitoxins (5, 10, 20, or 50 �L injections) were separated

by the following gradient at 0.8 mL/min, using two aqueous

eluents (eluent A: 20 mM sodium acetate and eluent B:

450 mM sodium acetate) both adjusted to pH 6.9 with

TABLE II. Linearity and approximate LOD values of HPIC-FD coupled to chemicalpostcolumn oxidation method for PSP analysis

Toxin

Concentration

Range (ppb)

Calibration

Curve

Correlation

Coefficient (r2)Approximate LOD

(ng injected)

C1 70–260 y ¼ 1267.9x þ 0 0.995 0.05

C2 26–130 y ¼ 2530.8x þ 0 0.948 0.02

GTX-2 44–165 y ¼ 2062.7x þ 0 0.995 0.02

GTX-3 13–50 y ¼ 1644.6x þ 0 0.996 0.09

NEO 48–181 y ¼ 263.39x þ 0 0.990 0.15

STX 47–176 y ¼ 2170.2x þ 0 0.999 0.02

TABLE I. PSP mean peak heights of each group of fivereplicate injections of a PSP mixture (ST1) in Milli-Qwater and their respective CV together with overall CV(%) for the 3-day analytical period

Toxin Day 1a Day 2a Day 3a

Mean CV

over 3-Day

Period

C1 364 (2.8) 364 (7.6) 365 (4.5) 5.0

C2 93 (6.1) 99 (3.7) 91 (4.0) 4.6

GTX2 625 (2.2) 606 (1.5) 601 (2.2) 1.9

GTX3 168 (4.5) 162 (1.7) 160 (1.9) 2.7

NEO 150 (2.5) 103 (3.7) 109 (2.5) 2.9

STX 910 (2.0) 758 (1.0) 828 (1.2) 1.4

aValues in brackets are CV values for five replicate injections.

552 PAPAGEORGIOU ET AL.

acetic acid: 100% eluent A from 0 to 6 min then a linear

gradient to 100% eluent B over 25 min and holding at

100% eluent B for 20 min. A solution of periodic acid

(5 mM), ammonium formate (33 mM) and potassium dihy-

drogen phosphate (33 mM) adjusted to pH 10 (PC1) with

sodium hydroxide or to pH 7.5 (PC2) with phosphoric acid

was added at a flow rate of 0.4 mL/min to the eluted toxins.

The flowing mixture of postcolumn derivatization reagent

and column eluent was reacted at 658C and then acidified

with acetic acid (5 M, introduced at a flow rate of 0.4 mL/

min). Fluorescence detection of the oxidized PSPs was car-

ried out at excitation and emission wavelengths of 330 and

390 nm, respectively. The HPIC columns were then equili-

brated with 100% eluent A for 20 min at 0.8 mL/min prior

to the next injection.

PSPs were also separated by the following gradient at

0.8 mL/min, using two aqueous eluents (eluent A: 20 mM

ammonium acetate and eluent B: 450 mM ammonium ace-

tate) both adjusted to pH 6.9 with 25% ammonia solution

(Jaime et al., 2001): 100 % eluent A 0–5 min then a linear

gradient to 100% eluent B over 25 min and holding at 100%

eluent B for 8 min. Postcolumn oxidation (using PC1) and

detection of PSPs were performed as described earlier. The

HPIC columns were then equilibrated with 100% eluent A

for 27 min at 0.8 mL/min prior to the next injection.

RESULTS AND DISCUSSION

Comparison of Mobile Phase Systems inHPIC-FD Analysis of CyanobacterialSaxitoxins

To investigate the effect of mobile phase composition on the

performance of HPIC-FD for PSP analysis, a novel mobile

phase system was compared to the ammonium acetate sys-

tem previously reported (Jaime et al., 2001) for the analysis

of a PSP mixture. Figures 2 and 3 represent identical 20 �Linjections of a synthetic PSP standard mixture (ST1) consist-

ing of typical Australian cyanobacterial (ACN) PSPs (C1/2,

GTX2/3) (Negri and Jones, 1995; Negri et al., 1995, 1997;

Onodera et al., 1996; Velzeboer et al., 2000) and the non

usual PSPs, NEO and STX, utilizing sodium acetate and

ammonium acetate mobile phase systems, respectively. Post-

column oxidation of separated PSPs to their fluorescent

derivatives was achieved chemically rather than by electro-

chemical (Jaime et al., 2001) means. The pH of the period-

ate-based oxidation system was set in accordance to the find-

ings of Gago-Martınez et al. (2001) who demonstrated that

maximum yield of fluorescent derivatives of non-N-hydroxy-lated PSPs (common PSPs in Australian ACN) using period-

ate as the oxidant, was achieved between pH 9.0 and 10.

Figures 2 and 3 revealed that retention times for all PSPs

Fig. 4. HPIC-FD chromatogram of a South Australian reservoir water sample (3) spikedwith a PSP mixture (ST1) and acetic acid.

TABLE III. Reservoir (res) water DOC and conductivitylevels

Res Water Samplea DOC (ppm)

Conductivity

(�S/cm)

1 2.7 34

2 2.7 36

3 6.9 136

4 11.2 600

aSpiked res water samples contained the following concentrations

(ppb) of PSPs: C1, 116; C2, 44.5; GTX2, 77.5; GTX3, 19; NEO, 87.5; and

STX, 87.5.

553ANALYSIS OF CYANOBACTERIAL SAXITOXINS USING HPIC-FD

except C1 toxin were approximately 1.1 times longer, using

the ammonium acetate-based mobile phase system. Both

mobile phases resolved all PSPs examined; however, C1 and

C2 toxins were clearly better resolved using the ammonium

acetate system. Both chromatograms showed a dip in base-

line after 40 min. The use of recrystallized sodium acetate or

high-purity ammonium acetate failed to eliminate this

feature; nevertheless, it was not considered to be detrimental

to the method because it did not affect the resolution and

peak shape of the PSPs. Interestingly, PSP peak heights and

areas in Figure 2 differed from the corresponding values

observed in Figure 3. Peak height and area values for C1,

C2, GTX2, and STX in the sodium acetate-derived chroma-

togram (Fig. 2) were approximately 1.8, 1.4, 2.2, and

5.4 times higher than the corresponding values in the ammo-

nium acetate-derived chromatogram (Fig. 3). In contrast,

peak height and area values for GTX3 and NEO in the

sodium acetate-derived chromatogram were 1.4 and 2.4

times lower than the corresponding values in the ammonium

acetate-derived chromatogram. Since the pH of both mobile

phase systems was identical (pH 6.9), this indicates that

another mechanism was responsible for the observed differ-

ences in fluorescence intensity of the PSPs under different

mobile phase compositions. Fluorescence produced as a

result of complexation of nitrogen-based heterocyclic or-

ganic compounds with specific inorganic cations has been

reported (Koutaka et al., 2004). Possibly, cations such as

sodium or ammonium can complex with the guanidine group

of PSPs to form discrete PSP–cation complexes that exhibit

different oxidation kinetics and fluorescence properties. The

fact that cyanobacterial-derived PSP profiles are often domi-

nant in C1/2 toxins and that a greater yield of fluorescence

was observed for four of the six PSP toxins using our sodium

acetate-based mobile phase system prompted us to discard

ammonium acetate mobile phase-based systems in further

investigations. Also, responses of four of the five PSPs com-

mon in Australian strains of ACN are higher, with the fifth

being similar with the two mobile phases, is a significant

advantage from an Australian perspective.

Precision of HPIC-FD

To determine the within-laboratory coefficient of variation

(CV) or relative standard deviation (RSD) of this HPIC-FD

method, five consecutive replicate 20 �L injections of the

standard cyanobacterial PSP mixture (ST1) were made and

this was repeated twice more on consecutive days. Table I

shows the mean PSP peak heights for each group of five rep-

licate injections and the respective CVs together with the

overall mean CV for the 3-day period. Peak height values

were utilized instead of peak areas, since greater accuracy in

PSP quantification was achieved in the former case. This was

attributed to the closeness of retention times for the PSPs

and, in some cases, retention time variation between replicate

injections, which probably affected the accuracy of the auto-

mated peak integration process performed by our HPLC

TABLE V. Mean peak heights and CV of all reservoir water–PSP injections anddeviation from peak heights of PSPs in Milli-Q water

Toxin

Mean Peak Heights and

CV(%)a of Res Waters 1–4

Deviation (%) from Peak Heights

of PSPs in Milli-Q Water

C1 241 (2.8) þ6.3

C2 288 (5.2) þ5.3

GTX2 414 (2.8) þ2.9

GTX3 125 (4.4) þ0.7

NEO 64 (5.4) �6.7

STX 578 (4.2) �5.3

aCV values in brackets.

TABLE IV. PSP peaks heights and CV of duplicate HPLC-FD injections of PSP-spiked reservoir watersand Milli-Q water

Toxin

Mean Peak Heights and CV (%)a of Duplicate Injections

Res Water 1 Res Water 2 Res Water 3 Res Water 4 Milli-Q Water

C1 231 (2.9) 239 (0.2) 278 (4.1) 218 (4.2) 227 (1.0)

C2 289 (8.2) 245 (10.6) 265 (1.6) 354 (0.2) 274 (1.4)

GTX2 413 (1.6) 411 (8.0) 424 (1.2) 408 (0.3) 403 (1.2)

GTX3 126 (5.0) 119 (5.2) 135 (7.3) 120 (0.2) 124 (2.2)

NEO 60 (0.9) 65 (11.2) 67 (4.8) 66 (4.6) 69 (9.5)

STX 548 (2.1) 611 (9.8) 566 (4.6) 587 (0.1) 610 (2.4)

aCV values in brackets.

554 PAPAGEORGIOU ET AL.

system’s software. The data in Table I show notable differ-

ence between mean peak height values for some PSPs over

the 3-day analytical period. It is probable that slight changes

in the pH of the postcolumn oxidation solution over this

period were reflected in the difference between the oxidation

kinetics of the PSP analogues, as shown by the variability in

their peak heights. Inconsistent or troublesome performance

of chemical postcolumn reagent systems has been noted by

others (Gago-Martınez et al., 2001; Jaime et al., 2001). How-

ever, minimal variability was observed between consecutive

daily replicate PSP injections as evidenced by the CV values

obtained for each day, indicating that adequate accuracy can

be achieved for multiple samples, provided that standards are

included with each analytical run.

To further probe the precision of this method for PSP

analysis, duplicate 20-�L injections of four PSP calibration

standard solutions (prepared from dilutions of ST1 with

Milli-Q water) were made and the respective mean peak

heights were used to determine the correlation coefficients

(r2) of each PSP calibration curve (Table II). The concen-

tration range used for C1/2 and GTX2/3 was similar in

magnitude to that found in extracts of PSP-producing

Australian cyanobacteria (ACN) and that dissolved in local

natural waters. Except for C2 toxin, r2 of the calibration

curves for the cyanobacterial PSPs investigated were 0.99

or better. An approximate concentration limit of detection

(LOD) of our HPIC-FD method at a fluorescence detector

slit width of 18 nm for each cyanobacterial PSP based on a

Fig. 6. HPIC-FD chromatogram of an Australian ACN extract.

Fig. 5. HPIC-FD chromatogram of a broad spectrum cyanobacterial PSP standard (ST2)mixture.

555ANALYSIS OF CYANOBACTERIAL SAXITOXINS USING HPIC-FD

5:1 signal to noise ratio and 100 �L injections was deduced

from chromatograms of the calibration solutions (Table II).

By operating the fluorescence detector with a slit width of

40 nm, LOD values can be reduced by at least 50%; how-

ever, this setting potentially increases the risk of interfer-

ence by fluorescent nonPSP-related compounds. LOD val-

ues of our method compared favourably to those obtained

for Jaime’s (Jaime et al., 2001) HPIC-FD method.

HPIC-FD Analysis of Raw Fresh Waters

To assess the applicability of our method for the concentra-

tion determination of PSPs in local (South Australian) raw

waters, four reservoir water samples (Table III) containing

different dissolved organic carbon (DOC) and salt levels

were spiked with ST1 and 2.5 M acetic acid (see Materials

and Methods) in a 50:50:1 ratio and then subjected to

HPIC-FD analysis in duplicate.

Addition of acetic acid to the raw water–ST1 samples

prior to HPIC-FD analysis was necessary to suppress any

adverse effects of salt impurities on the chromatography of

the PSPs. Figure 4 shows a representative HPIC-FD chro-

matogram of a reservoir water–ST1–acetic acid mixture

using fluorescent-free grade sodium acetate in the mobile

phase, which was similar to that shown for ST1 in Milli-Q

water (Fig. 2).

The use of fluorescent-free grade sodium acetate mini-

mized baseline drop in HPIC-FD analysis of PSPs. There-

fore, a solution of ST1 in Milli-Q water was also reanalyzed

using fluorescent-free grade sodium acetate in the mobile

phase to better ascertain the effects of raw water contami-

nants on the performance of HPIC-FD. Table IV shows

mean peak heights and CV values of duplicate 20 and

50 �L injections of a ST1-spiked Milli-Q water and reser-

voir water samples, respectively. 50 �L injections of the

reservoir water–ST1 samples were made to compensate for

the dilution factor incurred in their preparation and to

achieve larger peaks. Therefore, peak heights from the

Milli-Q water–ST1 injections were multiplied by 1.25 for a

direct comparison. The data in Table IV showed that peak

heights of the reservoir water samples differed by up to

4.2%, 10.6%, 8.0%, 7.3%, 11.2%, and 9.8% for C1, C2,

GTX2, GTX3, NEO, and STX, respectively. In contrast,

only peak heights for NEO varied by more than 2.5%

between duplicate injections of the ST1 mixture in Milli-Q

water. The fact that reservoir water samples 3 and 4 (con-

tain highest DOC levels and salt concentrations) yielded

duplicate injections with the least variation in peak heights

indicates that total salt and DOC levels alone did not

adversely affect the performance of HPIC-FD method for

PSP analysis but rather the constituents of the DOC and

salts present. Table V shows mean peak heights and CV of

the four reservoir water–ST1 duplicate injections together

with the overall deviation (�5.3% to þ6.3%) from the cor-

responding peak heights of the PSPs in Milli-Q water. The

data showed that the matrix (Luckas et al., 2003) of local

Fig. 7. HPIC-FD chromatogram of a Brazilian Cylindrospermopsis raciborskii (T3) extract.

TABLE VI. PSP content (lg/g dry cells) in crude extractsfrom natural samples of Australian ACN and BrazilianCylindrospermopsis raciborskii (T3)

Toxin ACN T3

C1 5520 n

C2 2740 n

dcGTXs not quantitated np

GTX1 n np

GTX2 852 np

GTX3 186 np

GTX4 np np

Unidentified peak np 39 (estimate only)a

NEO np 36

dcSTX n n

STX 54 n

n, negligible; np, not present.aQuantitated against NEO.

556 PAPAGEORGIOU ET AL.

raw waters with a DOC and salt range from 2.7 to 11.2 ppm

and 34 to 600 �S/cm�1, respectively, did not significantly

affect the accuracy of our HPIC-FD method for the concen-

tration determination of PSPs present in the low to medium

ppb range (40–200 ppb). However, it is expected that salt

removal and PSP preconcentration steps would be required

prior to HPIC-FD analysis of raw waters containing very

low PSP concentrations.

HPIC-FD Analysis of Cyanobacterial Extracts

As an extension to this study, we carried out a preliminary

examination of the applicability of our HPIC method to the

analysis of PSPs extracted from both natural Australian

ACN and Brazilian Cylindrospermopsis raciborskii (T3)

samples. To facilitate our investigation, a broad spectrum

PSP standard mixture (ST2) consisting of most cyanobacte-

rial PSP variants reported to date: C1/2, dcGTX2/3 (trace),

GTX1–4, NEO, dcSTX, and STX (Mahmood et al., 1986;

Hall et al., 1990; Humpage et al., 1994; Onodera et al.,

1996; Carmichael et al., 1997; Negri et al., 1997; all rele-

vant references therein) was prepared (see Materials and

Methods). GTX5 (Negri and Jones, 1995; Negri et al.,

1995; Velzeboer et al., 2000) was not included in ST2,

since authentic standards could not be sourced at the time

of this investigation. Figures 5–7 represent HPIC-FD chro-

matograms of ST2, and extracts from ACN and T3, respec-

tively. The pH of the postcolumn reagent (PC2) used was

set at 7.5 in accordance to the findings of Gago-Martınez

et al. (2001) who demonstrated that an increase in oxidation

Fig. 9. HPIC-FD chromatogram of a Brazilian Cylindrospermopsis raciborskii (T3) extractspiked with ST1.

Fig. 8. HPIC-FD chromatogram of an Australian ACN extract spiked with ST1.

557ANALYSIS OF CYANOBACTERIAL SAXITOXINS USING HPIC-FD

yield of N-1-hydroxylated PSP variants (GTX1/4 and NEO)

and, in turn, fluorescence yield of the corresponding deriva-

tives can be achieved near this pH. Figure 5 showed that all

of the PSP variants present in ST2 were clearly separated

using our HPIC-FD procedure.

Figure 6 showed that the PSP profile (Table VI) of the

ACN extract contained significant amounts of C1/2,

dcGTX2/3 (not resolved and not quantitated), GTX2/3, a

trace amount of dcSTX, and a small amount of STX as

shown by the peaks at approximately 7, 8, 25, 27, 29, 49.5,

and 51 min, respectively. These assignments were confirmed

from HPIC-FD analysis of the ACN extract spiked with ST1

in a 1:1 ratio (Fig. 8). Since Figure 8 showed no indication

that N-hydroxylated PSPs were present in can, it was deemed

not necessary to analyze the ACN extract spiked with ST2.

Figure 8 also showed a recovery of at least 93% for each PSP

detected, indicating that the matrix of ACN did not adversely

affect the performance of our HPIC-FD method (Luckas

et al., 2003). The presence of significant amounts of

dcGTX2/3 in the ACN extract was unexpected, since PSP

profiles from typical Australian ACN samples are not domi-

nated by these variants (Velzeboer et al., 2000). Most likely,

prolonged storage of the ACN sample in the presence of

trace amounts of acid or bacteria leads to the hydrolysis or

enzymatic cleavage of the respective precursor GTX2/3 car-

bamoyl groups. Figure 7 initially indicated that the PSP pro-

file of T3 was dominated by NEO and only very low levels

of C1/2 and STX, as shown by the peaks (not labeled) at

approximately 6, 7, 45.5, and 50 min, respectively. However,

HPIC-FD analysis (Fig. 9) of T3 spiked with ST1 showed

that the peak at 6 min in Fig. 7 did not correspond to C1 and

therefore it was highly doubtful that the very small peak at

7 min in Fig. 7 corresponded to C2, since it is common

knowledge that both epimers coexist (in equilibrium). Inter-

estingly, expansion of the NEO peak in Fig. 7, as shown

in Figure 10, indicated the presence of a coeluting PSP

(shoulder) suggesting that NEO and a significant amount of

an unidentified PSP were not resolved and possibly present

in similar concentrations with respect to each other (see

Table VI for approximate PSP concentrations in T3 extract).

Further investigations utilizing HPLC-MS will be employed

in the near future to determine the identity of compounds

correlating to the peak at 6 min (Fig. 7) and the shoulder at

45.7 min (Fig. 7). PSP recovery could not be confirmed from

the T3 PSP spike experiment, since it was not possible to

confidently quantitate NEO. The fact that T3 contained sig-

nificant levels of NEO but none of the typical Australian

ACN PSPs (C1/2, GTX2/3) and the less commonly found

GTXs (Hall et al., 1990; Negri and Jones, 1995; Negri et al.,

1995) correlated well with the findings of Lagos et al. (1999)

who showed a similar PSP profile for a related Brazilian-

sourced Cylindrospermopsis raciborskii extract.

CONCLUSIONS

In conclusion, this study shows that HPIC-FD coupled to

chemical postcolumn derivatization can be successfully

used to determine not only dominant Australian cyanobac-

terial PSPs (C1/2, dcGTX2/3) but also less common var-

iants (GTX1/4, NEO, dcSTX, STX) present in natural fresh

waters and different cyanobacteria species. It is envisaged

that Australian and other potentially affected country’s

water utilities and health-related authorities would substan-

tially reduce costs associated with current lengthy reverse

phase-based ion-pairing HPLC-FD methodologies by

adopting our relatively inexpensive and rapid HPIC-FD

method for PSP analysis. From a local (Australia) perspec-

tive, HPLC systems capable of performing PSP analysis

using our HPIC-FD method can be promptly sourced from

several suppliers/manufacturers. Overall, introduction of

HPIC-FD would minimize response times of both water

Fig. 10. Expanded HPIC-FD chromatogram of a Brazilian Cylindrospermopsis raciborskii(T3) extract.

558 PAPAGEORGIOU ET AL.

utility and health-related authorities, following toxic outbreaks

of PSP-producing cyanobacteria in natural water supplies.

The authors thank Dr. Sandra Azevedo, Nucleo de Pesquisas

de Productos Naturais, CCS, Bl. H, Universidade Federal do Rio

de Janeiro, Brazil, for the provision of Cylindrospermopsis raci-borskii (T3) extract.

REFERENCES

Carmichael WW, Evans WR, Yin QQ, Bell P, Moczydlowski E.

1997. Evidence for paralytic shellfish poisons in the freshwater

cyanobacterium Lyngbya wollei (Farlow ex Gomont). Appl

Environ Microbiol 63:3104–3110.

Ferreira FMB, Soler JMF, Fidalgo ML, Fernandez-Vila P. 2001.

PSP toxins from Aphanizomenon flos-aquae (cyanobacteria)

collected in the Crestuma-Lever reservoir (Duro river, northern

Portugal). Toxicon 39:757–761.

Gago-Martınez A, Moscoso SA, Martins JML, Vazquez J-AR,

Niedzwiadek B, Lawrence JF. 2001. Effect of pH on the oxida-

tion of paralytic shellfish poisoning toxins for analysis by liquid

chromatography. J Chromatogr A 905:351–357.

Hall S, Strichartz G, Moczydlowski E, Ravindran A, Reichardt

PB. 1990. In: Hall S, Strichartz G, editors. Marine Toxins: Ori-

gin, Structure and Molecular Pharmacology. Washington, DC:

American Chemical Society. Chap. 3. p 29–65.

Humpage AR, Rositano J, Bretag AH, Brown R, Baker PD, Nich-

olson BC, Steffensen DA. 1994. Paralytic shellfish poisons from

Australian blue-green algal (cyanobacterial) blooms. Aust J

Mar Freshwat Res 45:761–771.

Jaime E, Hummert C, Hess P, Luckas B. 2001. Determination of

paralytic shellfish poisoning toxins by high-performance ion

exchange chromatography. J Chromatogr A 929:43–49.

Koutaka H, Kosuge J, Fukasaku N, Hirano T, Kikuchi K, Urano

Y, Kojima H, Nagano T. 2004. A novel fluorescent probe for

zinc ion based on boron dipyrromethene (BODIPY) chromo-

phore. Chem Pharm Bull (Tokyo) 52:700–703.

Lagos NL, Onodera H, Zagatto PA, Andrinolo D, Azevedo

SMFQ, Oshima Y. 1999. The first evidence of paralytic shell-

fish toxins in the freshwater cyanobacterium Cylindrospermosisraciborskii. Toxicon 37:1359–1373.

Laycock MV, Thibault P, Ayer SW, Walter JA. 1994. Isolation

and purification procedures for the preparation of paralytic

shellfish poisoning toxin standards. Nat Toxins 2:175–183.

Luckas B, Hummert C, Oshima Y. 2003. Analytical methods for

paralytic shellfish poisons. In: Hallegraeff GM, Anderson DM,

Cambella AD, editors. Manual on Harmful Marine Microalgae.

Paris: UNESCO.

Mahmood NA, Carmichael WW. 1986. Paralytic shellfish poisons

produced by the freshwater cyanobacterium Aphanizomenonflos-aquae NH-5. Toxicon 24:175–186.

Negri AP, Jones GJ. 1995. Bioaccumulation of paralytic shellfish

poisoning (PSP) toxins from the cyanobacterium Anabaena cir-cinalis by the freshwater mussel Alathyria condola. Toxicon33:667–678.

Negri AP, Jones GJ, Hindmarsh M. 1995. Sheep mortality associ-

ated with paralytic shellfish poisons from the cyanobacterium

Anabaena circinalis. Toxicon 33:1321–1329.

Negri AP, Jones GJ, Blackburn SI, Oshima Y, Onodera H. 1997.

Effect of culture and bloom development and of sample storage

on paralytic shellfish poisons in the cyanobacterium Anabaena

circinalis. J Phycol 33:26–35.

Onodera H, Oshima Y, Watanabe MF, Watanabe M, Bolch CJ,

Blackburn S, Yasumoto T. 1996. Screening of paralytic shell-

fish toxins in freshwater cyanobacteria and chemical confirma-

tion of the toxins in cultured Anabaena circinalis from Aus-

tralia. In: Yasumoto T, Oshima Y, Fukuyo Y, editors. Harmful

and Toxic Algal Blooms. Paris: Intergovernmental Oceano-

graphic Commission of UNESCO.

Onodera H, Satake M, Oshima Y, Yasumoto T, Carmichael WW.

1997. New saxitoxin analogues from the filamentous cyanobac-

terium Lyngbya wollei. Nat Toxins 5:146–151.

Oshima Y. 1995a. Post-column derivatisation HPLC methods for

paralytic shellfish poisons. In: Hallegraeff GM, Anderson DM,

Cambella AD, editors. Marine Microalgae. IOC Manuals and

Guides 33. Paris: Intergovernmental Oceanographic Commis-

sion of UNESCO. p 81–94.

Oshima Y. 1995b. Post-column derivatisation liquid chromato-

graphic method for paralytic shellfish toxins. J AOAC Int 78:

528–532.

Oshima Y, Sugino K, Yasumoto T. 1989. Latest advances in HPLC

analysis of paralytic shellfish toxins. In: Natori S, Hashimoto K,

Ueno Y, editors. Mycotoxins and Phycotoxins ’88. Amsterdam,

the Netherlands: Elsevier. p 319–328.

Sullivan JJ, Iwaoka WT. 1983. High pressure liquid chromato-

graphic determination of toxins associated with paralytic shell-

fish poisoning. J Assoc Off Anal Chem 66:297.

Velzeboer RMA, Baker PD, Rositano J, Heresztyn T, Codd GA,

Raggett SL. 2000. Geographical patterns of occurrence and

composition of saxitoxins in the cyanobacterial genus Ana-

baena (Nostocales, Cyanophyta) in Australia. Phycologia 39:

395–407.

559ANALYSIS OF CYANOBACTERIAL SAXITOXINS USING HPIC-FD