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Analytica Chimica Acta 675 (2010) 125–131

Contents lists available at ScienceDirect

Analytica Chimica Acta

journa l homepage: www.e lsev ier .com/ locate /aca

n amperometric method for the detection of amitrole, glyphosate and itsminomethyl-phosphonic acid metabolite in environmental waters using passiveamplers

rancisco Sánchez-Bayoa,∗, Ross V. Hyneb, Karine L. Desseilleb

Centre for Ecotoxicology, University of Technology Sydney, PO Box 123, Broadway, NSW 2006, AustraliaCentre for Ecotoxicology, Department of Environment, Climate Change and Water (NSW), PO Box 29, Lidcombe, NSW 1825, Australia

r t i c l e i n f o

rticle history:eceived 16 February 2010eceived in revised form 9 July 2010ccepted 9 July 2010vailable online 21 July 2010

a b s t r a c t

The herbicides amitrole and glyphosate, and its metabolite aminomethyl-phosphonic acid (AMPA), inwater samples have been directly analysed by high-performance liquid chromatography using an elec-trochemical (EC) detector. Limits of detection of 0.3 �g mL−1 for glyphosate, 0.05 �g mL−1 for AMPA and0.03 �g mL−1 for amitrole were comparable to those obtained by other authors using EC and also by liq-uid chromatography coupled to mass spectrometry, but the latter method requires derivatisation and

eywords:onic herbicides

onitoringolid-phase extractionMPORE® disksatural waters

pre-concentration of the sample whereas EC methods show similar sensitivity without the need of anyderivatisation. The method was specifically designed to analyse extracts from passive samplers used formonitoring of polar herbicide residues in waters. To this purpose, three types of Empore® disks weretested for their ability to adsorb and desorb these ionic, polar analytes. A procedure for their extractionfrom the membranes and reducing the interferences from other substances present in natural waters (i.e.humic acids) is described. The method is simple, does not require sophisticated equipment and is valid

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for the analysis and moni

. Introduction

Amitrole [3-amino-1,2,4-triazole, CAS-RN 61-82-5] andlyphosate [N-(phosphonomethyl)glycine, CAS-RN 1071-83-6] areroad-spectrum polar herbicides commonly used in agriculturalreas all over the world. Amitrole is typically used for clearingeeds in irrigation channels, whereas glyphosate is becoming

ncreasingly the herbicide of choice in agriculture, urban parksnd gardens due to its low toxicity to animals [1,2], no persistencend low risks to humans and the environment [3,4]. Recently, thedoption of Roundup-resistant, genetically modified crop varietiesn many countries has boosted the use of this particular herbicide

orldwide [5], so its residues are expected to appear in freshwaterystems more frequently.

The need to monitor regularly these and other broad-spectrumerbicides is three-fold: they commonly cause unintended dam-ge to nearby crops, particularly if they appear in irrigation watershat are shared over entire regions. For example, in North America

∗ Corresponding author. Centre for Ecotoxicology, Department of Environment,limate Change and Water (NSW), PO Box 29, Lidcombe, NSW 1825, Australia.el.: +61 2 9995 5091; fax: +61 2 9995 5183.

E-mail addresses: Paco.Sanchez-Bayo@environment.nsw.gov.au,anchezbayo@mac.com (F. Sánchez-Bayo).

003-2670/$ – see front matter © 2010 Elsevier B.V. All rights reserved.oi:10.1016/j.aca.2010.07.013

g of herbicides residues using passive samplers.© 2010 Elsevier B.V. All rights reserved.

herbicides are a common cause of economic injury to neighbour-ing farmers, which in the case of canola can reach up to 10% yieldlosses annually [6]. Also, their residues in water are undesirablesince they could lead to chronic depletion of some aquatic plantspecies in wetland ecosystems. In addition to these environmen-tal problems, amitrole is carcinogenic and its residues in drinkingwaters should not exceed 0.1 �g L−1 in countries of the EuropeanCommunity [7] and preferably in any country.

Numerous analytical methods have been tried to detectamitrole, glyphosate and its active metabolite, aminomethyl-phosphonic acid (AMPA), in different matrices. These includecolorimetric methods for determination of amitrole in agricul-tural products and soil [8], capillary electrophoresis [7,9], basedon their amphoteric nature, and enzyme-linked immunosorbentassays (ELISA) for glyphosate [10]. Given that chromatographicmethods are most common in analytical laboratories, and sincethese ionic, non-volatile substances cannot be analysed directlyby gas chromatography, the use of high-performance liquid chro-matography (HPLC) is preferable. However, most of these methodsendure a number of difficulties which stem from the particular

chemistry of these two compounds: their polar characteristicsmake their extraction impossible using organic solvents, whilepre-concentration of their residues requires special ion-exchangematrices and, to a large extent, it is limited [11]. To make mat-ters more difficult, the molecules of amitrole and glyphosate lack

126 F. Sánchez-Bayo et al. / Analytica Chimica Acta 675 (2010) 125–131

Table 1Some chemical characteristics of the analytes.

Properties Amitrole Glyphosate AMPA References

Formula C2H4N4 C3H8NO5P CH6NO3PMolecular weight 84.08 169.07 111.04 [39]pKa 4.2, 10.7 0.8, 2.3, 5.9, 10.9 0.9, 5.6, 10.2 [39,40]Vapour pressure (mPa) 0.00003 0.0131 No information [39]

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Water solubility (g L−1) 280 12Partition coefficient (log Kow) −0.97 −3.Half-life in water (days) 30 33–

hromophores, so they cannot be analysed by the UV detectorsommonly used in both gas and liquid chromatography. As aonsequence, the majority of chromatographic methods to dateely on derivatisation to enable fluorescence or UV detection.ypical derivatising reagents for glyphosate are o-phtalaldehyde-ercaptoethanol (OPA-Merc) in post-column derivatisation [12]

nd 9-fluorenylmethoxycarbonyl chloride (FMOC-Cl) in pre-olumn derivatisation procedures [13]. For amitrole, diazotizationith sodium nitrite [14] or derivatisation with FMOC-Cl [15] are

ften used.Liquid chromatography coupled to tandem mass spectrome-

ry (LC–MS) is perhaps the most reliable and sensitive method tonalyze the derivatives of amitrole, glyphosate and AMPA [15,16].owever, the derivatisation of environmental water samples isot recommended as it frequently generates undesirable prod-cts that may interfere with the peaks in the chromatogram.he direct detection of the analytes has been sought, and it haseen achieved for amitrole using ion-pair HPLC [17], but unfortu-ately this method is not suitable for analysis of environmentalamples because the inorganic cations present in natural watersompete with the analytes [18]. Direct detection of both amitrolend glyphosate has also been achieved using electrochemical (EC)etectors [19,20], with limits of detection in the 38–64 �g L−1 rangeor glyphosate [19] and 10–15 �g L−1 for amitrole [19,21]. Indeed,C detectors are the best choice for the direct analysis of these her-icides [22], are easy to use and much more affordable than anyS equipment. Also, EC detectors can be fitted in series with UV

nd other detectors used in HPLC, thus enabling the simultane-us detection of substances with different chemical characteristicssing several detectors.

Among the EC methods, pulsed amperometric detection (PAD)s becoming increasingly useful to analyse polar aliphatic com-ounds which are not easily detected by conventional photometricr fluorometric techniques [23]. Electrocatalytic systems are classpecific: at high pH and low potentials, virtually all carbohy-rates, amino acids, amines and sulfur compounds are adsorbedt Au or Pt electrodes and their oxidation generate a currenthich is measured against a reference electrode; the analytes

re desorbed concomitantly with formation of inert surface oxide,hich is restored by applying a negative potential in short pulses.etection of amino acids and amine compounds is carried outirectly in alkaline media [24,25] and anion-exchange chromato-raphic columns are used for separation of the compounds. Sincemitrole, glyphosate and AMPA are amphoteric compounds con-aining primary and secondary amine groups (Table 1), their

irect detection by such electrochemical system is feasible [19,20].ther authors [26] have used integrated pulsed amperometricetection (IPAD) to analyse glyphosate in blood and urine sam-les, which appear commonly among farmers and applicators27], achieving a sensitivity of 51 �g L−1 which is in the same

5.8 [39,41]−2.36 [39,40]No information [39]

range as that obtained with other EC methods and water sam-ples.

Following on the previous steps, we have developed a PADmethod to analyse residues of both amitrole and glyphosate her-bicides, as well as the AMPA metabolite, in environmental watersamples without any derivatisation procedure. The method wasspecifically designed to analyse extracts from passive samplersused for monitoring of polar herbicide residues in waters [28], as theaim of the project was to provide farm irrigators with an effectivetool to detect such residues, which usually escape detection usingstandard sampling and analytical procedures. This paper describesfirst the analytical method and then the extraction procedure toobtain amitrole, glyphosate and AMPA from three types of Empore®

disks, with the aim of assesing their suitability to be used as passivesamplers in monitoring programs.

2. Materials and methods

2.1. Reagents

Analytical grade standards of amitrole (99.9%) and glyphosate(99.2%) were purchased from Riedel-de Häen (Seelze, Germany)or from Chem Service (Alltech Associates, Sydney, Australia) andAMPA (99%) was purchased from Sigma–Aldrich (Sydney, Aus-tralia). Sodium hydroxide solution (1 M) and NaCl (99.9%) werepurchased from Ajax Finechem (Taren Point, NSW, Australia) andkept in air-tight bottles to avoid contamination by atmosphericCO2. Acetone, methanol and iso-propyl alcohol (purchased fromCrown Scientific, Minto, Australia) were all of analytical grade.Purified water was obtained from a Milli-Q system (Millipore, Mol-sheim, France). Standard stock solutions of the analytes were madein purified water and kept refrigerated (4 ◦C) no longer than a week,as glyphosate decayed under these storage conditions with a half-life of 25–30 days.

2.2. Equipment

The HPLC system consisted of the following units connected inseries: Waters 600 Controller, Waters 717plus Autosampler, anion-exchange chromatographic column (see below), Waters 2487 Dualabsorbance detector and 2465 Electrochemical detector (Waters,Milford, USA). Only the latter detector was required for the analyt-ical method described here, but the placement of the UV detectorbefore the EC detector is mentioned here because the retentiontimes are affected by this arrangement.

Separation of the analytes was performed with an IonPac AS15(4 mm × 250 mm I.D.) anion-exchange chromatographic column(Dionex, Sunnyvale, CA, USA) connected to a IonPac AG15 guardcolumn (4 mm × 50 mm I.D.). Prior to running the samples, thedetector flow cell and the column were cleaned with a mobile phase

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f pure water for 30 min followed by 0.1 M NaOH alkaline solutionpH 13) for 45 min to ensure the complete elution of any residuesetained in previous runs.

The 2465 Waters unit is a single-channel electrochemical detec-or, built in a stable Faraday-shielded oven which contains a flowell connected to three electrodes: Au working electrode (3 mmiameter), a hydrogen reference electrode (Hy-REF, Waters, Mil-ord, USA) and an auxiliary electrode that completes the circuitnd supplies the current needed to offset processes at the work-ng electrode. The working electrode is kept at virtual ground, andhe chemical oxidation takes place at very low potential (1 mV). Theolume of the flow cell in the Kel-F block was 0.3 �L (50 �m thickpacer gasket), and the temperature in the Faraday oven was setonstant at 38 ◦C. Stabilisation of the flow cell is important, sinceery small changes in pH of the mobile phase affect the currentevels at the electrode and consequently alter the baseline of thehromatograph; for this reason, freshly made solutions of NaOHhould be kept in a N2 atmosphere to avoid capturing CO2 fromhe air and formation of CO3

2− species, which lower the pH of theobile phase and affect the retention times of the analytes [23].

tabilisation of the baseline requires to run the mobile phase inAD mode for about 60 min.

Empore® SDB-RPS and SDB-XC 47 mm extraction disks (3 M,t Paul, MN, USA) were purchased from Phenomenex (Lane Cove,SW, Australia).

.3. Optimization of conditions

Optimization of the working potential for each analyte is essen-ial, and it is performed by a cyclic scanning voltammogram, whichlots the variable potential applied to the electrode against the cur-ent produced. A high ratio between the response of the mobilehase alone and that of the mobile phase plus the analyte indi-ates the best working potential. Scanning cycles were in the range0.5 to +0.7 V at a scan rate of 20 mV s−1. The NaOH mobile phases

ried varied in concentration from 0.025 to 0.25 M, correspondingo a pH range 12.1–13.02 under air-saturated conditions at normalressure and temperature in the laboratory.

The preferred mobile phase concentration and pH were choseno determine the best chromatographic conditions to separate thehree analytes with flow rates varying between 0.5 and 1 mL min−1.socratic elution is best when working with EC detectors, since gra-ients alter the pH and, therefore, destabilise the baseline of theow cell detector.

.4. Sample preparation

Fresh samples of the analytes were made in 1 mL of purifiedater, using 25 �L injections for HPLC analysis in all cases. Calibra-

ion of the stardard curves of each analyte was performed usinghe following range of concentrations: from 0.001 to 0.1 mM formitrole and AMPA, and from 0.01 to 1.0 mM for glyphosate.

Natural waters from an urban creek (South Creek, Sydney, NSW,ustralia) and Cox’s River near Sydney were filtered through 0.7 �mF/F glass microfibre filters (Whatman, Maidstone, England) to

emove any suspended particles, and then spiked with a mixture ofmitrole, glyphosate and AMPA at concentrations between 0.2 and.02 mM. Given the polar nature of the three compounds, no organicolvents are required to extract them from the water matrix, soll the samples were analysed directly, without pre-concentrationteps.

.5. Passive samplers – adsorption and extraction

Adsorbent membranes used in monitoring of polar compounds28] can concentrate the herbicides amitrole and glyphosate to a

ica Acta 675 (2010) 125–131 127

certain extent, thus lowering the detection limits obtained by directEC detection. Since each compound is retained in the membraneson a fixed proportion the time-weighted average concentration ofthe herbicides in the water sample can be estimated when deployedin a waterway over a known period of exposure [29]. For the her-bicides considered here, passive sampler monitoring has two mainadvantages over standard grab sampling of environmental waters:(i) it concentrates the residues of amitrole and glyphosate found innatural waters, allowing greater sensitivity than by direct analysis;and (ii) it provides a more realistic description of the levels of con-tamination, integrating the residual amounts found within specificperiods of time. Retention of the two herbicides and AMPA metabo-lite by such membranes, and their subsequent extraction, has notbeen measured yet, so three different Empore® disk membraneswere tried to evaluate their ability to absorb/extract these analytesfrom natural waters: a polystyrene divinyl benzene membrane(SDB-XC), a polystyrene divinyl benzene-reverse phase sulfonatedmembrane (SDB-RPS) and anion-exchange membrane made ofpolystyrene–divinylbenzene copolymer that has been modifiedwith quaternary ammonium groups.

Empore® disks were placed on DiskcoverTM-47 holders (Restek,Bellefonte, PA, USA) fitted to a vacuum manifold (Supelco, Belle-fonte, PA, USA). The disks were pre-conditioned following theinstructions from the manufacturer, i.e. adding 10 mL of acetonefollowed by 10 mL of iso-propyl alcohol, 10 mL of methanol and10 mL of purified water. Both purified and river water samples werespiked with known concentrations of the three analytes in the rangeof 0.01–0.1 mM, and then 10 mL was filtered through the disks.Experiments were performed in triplicate for each of the disk types.Extraction of the analytes adsorbed on the disks was performed bysoaking the membranes in 5 mL of 10 mM NaOH solution (pH 12)for 60 min at room temperature, and then eluting the extract undervacuum. It was found that soaking times less than 60 min result inpartial desorption of the analytes (data not shown). A subsampleof the extract (1 mL) was taken for direct analysis by HPLC and ECdetection. In order to measure the percent retention of the analytesin the membranes, a subsample (1 mL) of the purified water elutedfrom the disks was also taken for HPLC analysis.

3. Results and discussion

3.1. Voltammetric responses

The response of three independent solutions of 1 mM amit-role, glyphosate and AMPA in a 0.1 M NaOH mobile phase (pH 13),which provided the best sensitivity, is shown in the voltammogram(Fig. 1). The increasing anodic response during the positive scan forsolutions of the three compounds is higher than that of the mobilephase alone in the region between +100 and +300 mV, thus allow-ing their detection in this range of potentials. This is similar to thatreported by other authors using Cu [19] and Au electrodes [19,26].The deep peak in the reverse scan about +150 mV is due to thereduction of surface oxide (AuO) and intermediate products (AuOH)formed during the positive scan [23]. The observed response is sim-ilar to the catalytic oxidation of amines and amino acids [24,25], andresults from the adsorption of the nonbonded electron pair of theN-atom on to the oxide-free gold surface of the electrode, where it isimmediately oxidized and desorbed under alkaline conditions. Theadsorption of the analytes on the oxide-free surface is necessary forthe detection [26].

As Johnson and co-workers have demonstrated [23–25], theregeneration of the clean gold surface can be achieved by revers-ing the potential in brief pulses. Indeed, while the gold surfaceis oxidised under a high potential, the AuO formed is subse-quently reduced under a low potential, with the process being

128 F. Sánchez-Bayo et al. / Analytica Chimica Acta 675 (2010) 125–131

Fig. 1. Voltammograms for 0.1 M NaOH mobile phase (pH 13) and 1.0 mMglyphosate, AMPA and amitrole in the same mobile phase. Scan rate: 20 mV s−1. Dur-ing the positive scan, the response for solutions of the three compounds is higherti+(

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han that of the mobile phase alone in the region between +100 and +300 mV, allow-ng their detection in this range of potentials. The deep peak in the reverse scan about150 mV is due to the reduction of surface oxide (AuO) and intermediate productsAuOH) formed during the positive scan.

eversible. Unlike the gold surface, the oxidation of the analytess not reversible. Because the signal at the working electrode isroduced by the difference in current between the oxidation andeduction processes at its surface, the net signal given at the detec-or output represents the oxidation of the analyte only.

.2. Sensitivity of the detector

Since the background current at the electrode varies with the pHf the mobile phase, equal concentrations of analyte (0.1 M) wereested at various detection potentials (E1) and pH backgroundsithin the anodic range +50 to +300 mV. The preferred alkalineobile phase for all compounds was a 0.1 M NaOH solution (pH

3), as higher concentrations than this produced too much back-round noise. Best signals were obtained for amitrole at +225 mVnd for glyphosate and AMPA at +200 mV (Fig. 2), and this is con-istent with the scanning response in the voltammogram, whichhows the highest signal to noise ratio for these analytes preciselyn that range (Figs. 1 and 2).

Based on this behaviour, the PAD cycle for analytical detectionf the three compounds were set as follows: adsorption/detectionotential (E1) at +200 mV for 400 ms, during which time the signalenerated in the last 100 ms is sent to the detector output; cleaningotential (E2) at +800 mV for 300 ms; and conditioning potentialE3) at −600 mV for 300 ms to regenerate the clean gold surface.he entire pulse cycle lasts 1 s.

.3. Chromatography of the analytes in purified and naturalaters

As the 0.1 M NaOH solution (pH 13) provided the best sensi-ivity for the EC detector, it was chosen as the mobile phase for

ll HPLC analyses. Individual standard concentrations of amitrole,lyphosate and AMPA as well as their mixture in purified waterhowed sufficient separation of these compounds in the anion-xchange column for any flow rate between 0.5 and 1 mL min−1.eparation of the analytes was best achieved at low flow rates

Fig. 2. Optimization of the detection potential (E1) at the Au working electrodefor amitrole, glyphosate and its AMPA metabolite based on the peak areas of0.1 mM analyte solutions in pure water. Mobile phase: 0.1 M NaOH and flow rate0.5 mL min−1. Injection volume: 25 �L.

(0.5 mL min−1), with peaks in the range of 5.6–8.0 min, compared tothe high ones (1.0 mL min−1), which resulted in peaks in the rangeof 2.8–4.0 min. However, the lowest flow rate tried resulted in lesspeak efficiency, so a rate of 0.6 mL min−1 is probably the best com-promise as it provides enough resolution for the three compounds.Under these conditions the retention times for AMPA, amitrole andglyphosate were 4.4, 5.1 and 5.7 min respectively.

Direct analysis of filtered natural water samples produced twooverlapping peaks at 4.1 and 4.3 min that interfered with the AMPApeak in the chromatogram (Peaks A and B, Fig. 3a). These peakspossibly originate from humic acids and/or amino acids found insolution [24]. To avoid interferences with AMPA, the pH of themobile phase was modified by diluting the 0.1 M NaOH solutionwith increasing volumes of purified water in subsequent steps, i.e.90%:10%, 80%:20%, etc. Ionic compounds tend to elute at differentspeeds within the anion-exchange column depending on the pHof the mobile phase; due to the phosphonic group present in bothglyphosate and AMPA these compounds are particularly sensitivyto changes in pH, and their retention time – particularly that ofglyphosate – is delayed progressively as the pH goes lower, whereasthe amine group of amitrole does not respond much to variationsin pH (Fig. 4). It appears that a 75:25 (v/v) dilution of the origi-nal mobile phase, i.e. 75 mM NaOH and pH 12.75, is appropriate toresolve the contaminant peaks appearing in natural waters withoutcompromising those of our three analytes (Fig. 3b).

Under these conditions, the detector gave a linear response upto at least 180 �g mL−1 of glyphosate and around 50 �g mL−1 ofamitrole and AMPA (Table 2). The limits of detection for standard25 �L injections, estimated as a signal to noise ratio greater than3, were 0.32 �g mL−1 for glyphosate, 0.05 �g mL−1 for AMPA and0.03 �g mL−1 for amitrole, which are comparable to those obtainedby other authors using electrochemical detection but different typeof electrodes [19–21]. Using a similar EC detector (Au and Ag/AgClreference electrode) and same chromatographic conditions Sato etal. [26] reported a limit of detection for glyphosate in blood and

−1

urine sample seven times lower than ours (0.051 �g mL ), but theyused integrated pulsed amperometric detection (IPAD) instead ofPAD mode. It should be stressed that these EC detection limits arefor direct analysis without pre-concentration of the analytes. Directanalysis obviously limits the ability to concentrate the residual

F. Sánchez-Bayo et al. / Analytica Chimica Acta 675 (2010) 125–131 129

Table 2Optimal conditions for the analysis of amitrole, glyphosate and its metabolite AMPA by HPLC and PAD amperometric detection.

Analyte Retention time (min)a Detection limitb Range (�g mL−1) Intercept Slope r2

(�g mL−1) ng

AMPA 4.9 0.05 1.2 0.1–50 7361 20714 0.990Amitrole 5.5 0.03 0.7 0.05–40 27356 90154 0.999Glyphosate 7.7 0.32 7.8

a Mobile phase: 75 mM NaOH (pH 12.75) and flow rate 0.6 mL min−1.b Estimated for a signal to noise ratio ≥3 and 25 �L injection.

Fig. 3. Analysis of filtered river water by anion-exchange chromatography andpulsed amperometric detection (PAD) using: (a) river water; (b) river water spikedwith 0.1 mM amitrole and AMPA and 1.0 mM glyphosate; contaminant peaks (A andB). Mobile phase: 75 mM NaOH (pH 12.75) and flow rate 0.6 mL min−1. Injectionvolume: 25 �L.

Fig. 4. Optimization of the NaOH mobile phase, indicated by the pH of the solution,to resolve the peaks (retention time) of the three analytes and the contaminantsfound in natural waters (A, B and C).

1.0–180 −2693 46950 0.996

amounts by standard evaporation or solid-phase-extraction (SPE)methods, and consequently the detection limits in water samplesare those of the detector itself. The higher sensitivity of the LC/MSmethod was achieved using derivatisation and pre-concentrationof the derivatives of amitrole and glyphosate [15,16], whereas thedirect analysis of amitrole using LC/MS [11] rendered a limit ofdetection of 0.2 �g mL−1, above those reported here for direct ECdetection.

A high sensitivity may be more appropriate for analysis of verylow amitrole residues in drinking water [15], but our detection lim-its are deemed sufficient for detection of glyphosate and amitrolein environmental samples, where monitoring can be performedusing appropriate passive samplers. Typical concentrations of bothglyphosate and AMPA in effluents from urban areas are in the rangeof 0.1–2.0 �g L−1 [30], whereas in agricultural areas residue con-centrations can be as high as 8.7 �g L−1, just slightly lower thanthose found for atrazine and other commonly used herbicides [31].In order to detect such low levels of herbicide using HPLC with ECdetectors the environmental water samples must be concentrated100–500-fold, either by evaporation or by filtering large volumesof water sample through appropriate solid-phase cartridges likeAmberlite® [32]. For amitrole, residue concentrations in surfacewaters in France have been reported in the range of 2.3–190 �g L−1

[33], although they can reach 1 mg L−1 in ponds [34], and thereforeall EC methods are suitable.

3.4. Chromatography of the analytes extracted from Empore®

disks

Passive samplers are becoming common tools for monitoringpesticide residues in natural environments because they concen-trate the analytes in the membranes over time, thus offeringan integrated approach of the concentrations found in naturalwaters [35]. The difficulty with ionic herbicides like amitrole andglyphosate is to find membranes that can adsorb these analytesand allow their extraction from the polymeric matrix. Two of theEmpore® disks tried here, SDB-XC and SDB-RPS, have already beensuccessfully tested with other polar herbicides [28], while the otherone (anion-exchange) is specifically designed to capture ionic sub-stances.

To test the ability of the three membranes to adsorb theherbicides, purified water samples were spiked with nominal con-centrations of 0.1 mM glyphosate and 0.02 mM for both AMPA andamitrole, and 10 mL of the solutions were filtered through threetypes of Empore® disks as described in Section 2.4. Each experimentwas conducted several times. A subsample (1 mL) of the eluted fil-trate from spiked pure water was analysed using the conditionsdescribed above (as in Table 2) in order to estimate the percentretention of each analyte in the membranes. The analytes were sub-sequently extracted with small volumes of an appropriate alkaline

solution that matched the highest pKa of the three compounds ofinterests (between 10 and 11, Table 1) [36]. At such pH glyphosateand AMPA are strong anionic species [37] which cannot be retainedin the negatively charged polymeric membrane, and consequentlyare released from it. The same applies to amitrole. A solution of

130 F. Sánchez-Bayo et al. / Analytica Chim

Table 3Percent retention or adsorption (mean ± rsd) of amitrole, glyphosate and AMPAon three types of Empore® disks, and their recovery from the disks after 60 mindesorption with 10 mM NaOH (pH 12).

Compound/disk n Retention Extraction Recovery

GlyphosateSDB-XC 5 74.6 ± 8.3 50.8 ± 13.5 68.1SDB-RPS 6 28.2 ± 11.4 17.9 ± 3.9 63.4Anion-exchange 4 ∼100 <LODa 0

AMPASDB-XC 6 28.1 ± 19.2 32.3 ± 4.3 115.3SDB-RPS 5 29.9 ± 16.6 31.0 ± 8.0 103.9Anion-exchange 4 ∼100 <LOD 0

AmitroleSDB-XC 6 20.3 ± 11.3 21.3 ± 7.8 104.9

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SDB-RPS 8 95.7 ± 11.3 95.1 ± 9.1 99.4Anion-exchange 4 ∼100 1.4 ± 3.9 1.4

a LOD, limit of detection.

0 mM NaOH (pH 12) met these conditions for the three analytesried here (see Table 1), and it proved to work for desorbing thehree analytes when allowed to soak for 45–60 min.

All three Empore® disks retained large amounts of the targetnalytes, with the anion-exchange membranes capturing 100% inll cases (Table 3). Extraction of the analytes with the NaOH solu-ion, however, could only be performed with two of the disksSDB-XC and SDB-RPS), after soaking the membranes for 60 min tochieve a complete desorption of the analytes from the matrix – thenion-exchange membrane did not allowed the extraction of anynalyte using this method, and sonication in a water bath for 15 min

as tried as well but unsuccesfully. Therefore the anion-exchangeembranes are not suitable for passive sampler monitoring of

hese ionic herbicides using kinetic sampling when environmentaloncentrations are variable [38]. Recoveries of the analytes from

ig. 5. Analysis of extracts of amitrole, glyphosate and AMPA from SDB-XC Empore®

isks using the same chromatographic conditions as in Fig. 3: (a) standard pre-onditioning of spiked river water (0.1 mM amitrole and AMPA and 1.0 mMlyphosate); (b) after washing the disks with 10 mL of 1 M NaCl during the con-itioning step to remove the contaminant peaks (A and B).

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ica Acta 675 (2010) 125–131

the other two membranes were 104–115% for AMPA, 99–105% foramitrole and 63–68% for glyphosate (Table 3).

Direct analysis of the Empore® disk extracts produced the sameinterfering peaks (A and B) found in natural waters and sometimesan additional late contaminant (peak C) at 8.1 min (Fig. 4). How-ever, all these peaks were eliminated after washing the disks with10 mL of 1 M NaCl solution (58 g L−1) during the conditioning step,i.e. prior to filtering the water sample through the Empore® disk(Fig. 5b). This finding is important to ensure the correct preparationof the disks used in passive sampling and so avoid the interferingpeaks which could lead to false positives in the analyses.

We have also performed analyses of the three compounds bypre-column derivatisation methods, using FMOC-Cl and UV detec-tion at 264 nm, and our experience and results show the EC methoddescribed in this paper is more accurate, reliable and sensitive(considering the detection is direct, i.e. without concentrationsteps) than the derivatisation method. In fact, one of the problemsencountered with the derivatisation method was the excessivelyhigh variability in the recoveries (more then 20%) from spiked nat-ural water samples, which is probably caused by interference fromother substances present in those waters. The latter substancesare demonstrated to occur in natural waters, as shown in Fig. 3,although their identity is not known to us.

4. Conclusions

Development of a suitable HPLC analytical method for directanalysis of amitrole, glyphosate and its metabolite in environmen-tal waters and passive samplers has been achieved using a goldEC detector. A highly alkaline mobile phase (pH 12.75) allows theseparation of the three compounds through an anion-exchangecolumn, without interferences from other substances found innatural waters. The method does not require derivatisation nor pre-concentration steps, while the use of passive sampling membraneshelps improve the detection ability. Two types of membranes –Empore® disks SDB-RPS and SDB-XC – were found suitable for mon-itoring of amitrole and gyphosate in the environment using passivesamplers, since both analytes are retained in those matrices and canbe desorbed completely by soaking the membranes in an alkalinesolution (pH 12). Further testing is required to calibrate the kineticsand performance of these membranes before they can be used inpassive sampling.

Acknowledgement

The study was partially funded by a grant from Goulburn-Murray Rural Water Authority (Victoria) for salary support to F.S-B.KLD thanks the Conseil régional de Picardie (France) and the Euro-pean Union for their financial support.

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