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Solanum nigrum L. weed plants as a remediation tool for metalaxyl-pollutedeffluents and soils

Jorge Teixeira a,!, Alexandra de Sousa a, Manuel Azenha b, José Tiago Moreira a, Fernanda Fidalgo a,A. Fernando Silva b, Joaquim L. Faria c, Adrián M.T. Silva c

a Center for Biodiversity, Functional & Integrative Genomics (BioFIG), Departamento de Biologia, Faculdade de Ciências, Universidade do Porto,rua do Campo Alegre s/n, 4169-007 Porto, Portugalb CIQUP, Departamento de Química e Bioquímica, Faculdade de Ciências, Universidade do Porto, rua do Campo Alegre 687, 4169-007 Porto, Portugalc Laboratório de Catálise e Materiais, Laboratório Associado LSRE/LCM, Departamento de Engenharia Química, Faculdade de Engenharia, Universidade do Porto,Rua Dr. Roberto Frias s/n, 4200-465 Porto, Portugal

a r t i c l e i n f o

Article history:Received 22 January 2011Received in revised form 8 June 2011Accepted 10 June 2011Available online 7 July 2011

Keywords:Black nightshadeXenobioticPhytoremediationGlutathione-S-transferasePeroxidaseMetalaxyl

a b s t r a c t

In this work, the phytoremediation potential of metalaxyl, a commonly used persistent, mobile and lea-chy fungicide, by Solanum nigrum L. plants was studied. The study revealed that this plant species can beused as an excellent metalaxyl phytoremediation tool, thus providing a cost effective and environmen-tally friendly clean technology for the decontamination of sites and effluents. As it can be sowed directlyin the remediation site, is able to complete its life cycle without suffering major stress. Because it accu-mulates high amounts of the fungicide in the aboveground tissues, enables its concentration and properdisposal by cutting off the corresponding plant part. The study also suggests that the tolerance to metal-axyl is due to a suitable antioxidant response comprising proline accumulation and guaiacol peroxidaseand glutathione-S-transferase enhanced activities, that reduce oxidative damage to the plant organs.

! 2011 Elsevier Ltd. All rights reserved.

1. Introduction

Pesticide application has become an integral part of agricultureworldwide. Pesticide drift outside the target area is economicallywasteful and potentially hazardous for nearby non-targeted plants,animals or other organisms. Soil, surface areas, groundwater, andsediments become contaminated with pesticides because of spills,accidents, misapplication, and/or runoff and soil erosion from crop-lands (Karthikeyan et al., 2004; Henderson et al., 2006). To mini-mise the impact of this pollution it is important to developinnovative and cheap technologies to clean these contaminatedsites and waters.

Phytoremediation is an emergent technology that employsplants and their associated microbiota to remove, contain or renderharmless environmental contaminants (Bindu et al., 2008). Phyto-remediation is gaining a lot of importance in recent times since it isa cost effective, promising and environmentally friendly cleantechnology. Plants can be used to clean up the contaminated envi-ronment with minimum disturbance, as they serve as solar-driven

pumps, which extract and concentrate elements from the soil andwater (Eapen and D’Souza, 2005).

It is generally recognised that plants can remediate organic pol-lutants by, per example, direct root uptake and consequent accu-mulation of nonphytotoxic metabolites in plant tissues (seereferences in Wilson et al., 2001). Enzymatic degradation of a xeno-biotic in plants usually involves several enzymatic steps before adetoxified product of some stability is formed. Depending on thenature of the compound, it may directly react with biogenic conju-gation partners, or the molecule may be transformed (activated)into a substrate that might become susceptible to biomoleculecoupling. The latter reactions correspond to phase I, or chemicalactivation, and usually involve hydrolysis or oxidation, leading tothe formation of reactive groups within the xenobiotic molecule(Coleman et al., 1997; Schroder and Collins, 2002).

In phase II, conjugation reactions deactivate the xenobiotics, orthe phase I-activated molecules, by covalent linkage to an endoge-nous hydrophilic molecule, such as malonate, glucose or glutathi-one (GSH), to form a water-soluble conjugate (Coleman et al.,1997; Schroder and Collins, 2002), where malonyl- and glucos-yltransferases and glutathione-S-transferases (GST) are involved(Pilon-Smits, 2005). Phase II products are either non-toxic, or lesstoxic than the parent compound. Phase II is therefore an importantprotective phase in the detoxification process. The inactive water-

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! Corresponding author. Tel.: +351 220402729; fax: +351 220402709.E-mail addresses: agteixei@fc.up.pt, jteixeira@fc.up.pt (J. Teixeira).

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soluble conjugates are then exported from the cytosol to the vacu-ole or the apoplast by membrane-located transport proteins, theABC-type ATPases, thus initiating phase III, the compartmentationand processing part of the detoxification process (Coleman et al.,1997; Schroder and Collins, 2002; Pilon-Smits, 2005).

In this work metalaxyl (C15H21NO4, methyl N-(methoxyacetyl)-N-(2,6-xylyl)-D-alaninate, CAS No. 57837-19-1) (PAN PesticideDatabase, 2011) was chosen as a model compound because of itscommon use in vineyards and several horticulture industriesworldwide, and due to its physiochemical properties. This systemicfungicide is soluble in water (8.4 g L!1), and is available in severalcommercial formulations (Wilson et al., 2001; Syngenta, 2009).The primary routes of dissipation of metalaxyl in surface soil ap-pear to be aerobic soil metabolism, leaching and plant uptake.The compound is stable to hydrolysis, photolysis in water and soil,and does not volatilise appreciably. Both laboratory and field stud-ies indicate that metalaxyl is persistent and mobile and will leachin many soils. Although metalaxyl does not pose a threat to non-target species, it does exceed the Level of Concern (LOC) forgroundwater quality based on the Environmental ProtectionAgency’s criteria for mobility and persistence. Metalaxyl is slightlytoxic to wild mammals, freshwater fish and aquatic invertebrates(EPA, 1994). Furthermore, N-(2,6-dimethylphenyl)-N-(methoxy-acetyl) alanine is also prone to naturally form in soils from metal-axyl, being considered dangerous to the environment (PANPesticide Database, 2011). The reasons just stated further justifythe need to research possible strategies for its removal from theenvironment.

To date very few reports were published regarding metalaxylphytoremediation (Wilson et al., 2001; Turgut, 2005), none focus-ing on ubiquitous plants, such as Solanum nigrum L. (black night-shade), an annual or perennial weed, widely growing from theVariable Zone to the Torrid Zone in Europe, America and Asia(Wang et al., 2008), being therefore well adapted to several cli-matic conditions. It has a fast growth rate (completing its life cyclein about 2–3 months, depending on the environmental conditionssurrounding it) and large shoot biomass (Wang et al., 2008), whichare elite characteristics for its use as a phytoremediation tool(Pilon-Smits, 2005). Furthermore, it has been reported to remedi-ate polychlorinated biphenyls (Rezek et al., 2007), hence revealinga strong potential as a remediation tool for organic pollutants.

The present study aims to determine the removal efficiency of S.nigrum towards metalaxyl for its use as a phytoremediation tool toclean up metalaxyl-contaminated environments. Considering theimportance of the antioxidant system in plant metabolism and instress protection, to assess metalaxyl-induced oxidative stress inS. nigrum, and to evaluate the relative importance of the stress pro-tective mechanisms to cope with the produced Reactive OxygenSpecies (ROS), lipid peroxidation, free proline content, photosyn-thetic-related pigments, as well as morphological changes in rootsand shoots of seedlings and fully grown S. nigrum plants exposed toincreasing metalaxyl concentrations for 4 weeks, were analysed.

2. Material and methods

2.1. Effect of metalaxyl on seed germination and seedling development

In order to determine if metalaxyl could be deleterious to, orexert a negative effect on S. nigrum seed germination and seedlingdevelopment, thus preventing the direct seeding of the remedia-tion site, seeds were surface sterilised by vigorous shacking in70% ethanol for 3 min and subsequently dried on filter paper in alaminar flow hood. Seeds were then distributed evenly throughdifferent sterile containers baring sterile Hoagland solutionembedded in filter paper, supplemented with the following

metalaxyl concentrations (supplied as a pesticide solutioncommercially available as Ridomil" by Syngenta, Portugal): 0.0(control), 6.25, 12.5, 25.0, 50.0, 75.0 and 100.0 mg L!1 (or ppm –parts per million).

After 2-d incubation period at 4 #C in the dark (for seed germi-nation stratification) seeds were incubated for 4 weeks in a growthchamber at 23 #C with a 16 h/8 h (light/dark) photoperiod. Seedgermination percentage was determined at the end of 1–2 weeksof treatment, while seedling weight and shoot and root size weredetermined at the end of the 4-weeks exposure period.

2.2. Effect of metalaxyl on plant growth

Four weeks old control-grown seedlings were grown under theconditions already described, in pots containing a substrate mix-ture of vermiculite:perlite (2:1), and treated with 0.0 (control),6.25, 12.5, 25.0 and 50.0 mg L!1 metalaxyl (Ridomil"). Biometricparameters were accessed after 4 weeks of exposure. Becauseplants treated with 6.25 mg L!1 did not show any morphologic dif-ferences when compared to the control plants, suggesting that theydid not suffer any toxicity, and because those treated with25 mg L!1 metalaxyl did not produce sufficient biomass for all ex-pected assays, only the shoots and roots from control and12.5 mg L!1-treated plants were frozen in liquid N2 for stress bio-markers and enzyme activity analysis. Plant organs derived fromthis treatment were also oven-dried and used for water contentand metalaxyl levels determinations.

2.3. In planta metalaxyl levels

Metalaxyl levels were determined using a procedure similar toone previously described (Viñas et al., 2008). Mainly, oven driedroots or shoots were homogenised with a pestle and 0.03–0.05 gwere subjected to solid–liquid extraction with 5 mL of ethylace-tate. After filtration, the extracts were evaporated under a nitrogenstream and the residue reconstituted in water. The pesticide resi-dues were then subjected to direct-immersion solid-phase mic-roextraction (SPME, PDMS/DVB fiber, Supelco, Bellefont, CA) atroom-controlled temperature (25 #C) during 15 min. The pesticideresidues were finally desorbed at a GC–MS port for analysis. A Var-ian GC–MS instrument comprising an ion-trap Saturn 2100T detec-tor, a 1177 split/splitless injector and a Varian VF-5 ms capillarycolumn (20 m length, 0.15 mm internal diameter, 0.15 lm filmthickness) was used. The injector was operated at 250 #C, splitlessmode (2 min) plus 2 additional min with the split valve on. The col-umn flow was kept at 0.7 mL min!1 (Helium with 99.9999% pur-ity). Temperature program: 80 #C (2 min) followed by a ramp15 #C min!1 up to 300 #C. Detection was performed at the lSISion preparation mode with isolation window of 1 m z!1 units (se-lected m z!1 for metalaxyl: 206). The quantification relied on exter-nal calibration with matrix-matching standards prepared fromcontrol plants containing undetectable amounts of metalaxyl. Suchtype of standards allowed method recoveries within 95–105%.

2.4. Photosynthetic pigments analysis

For the extraction of photosynthetic pigments frozen shootsfrom control and 12.5 mg L!1-treated plants were grinded in 80%acetone with quartz sand using a pre-chilled mortar and pestle.The extracts were centrifuged at 2200g for 15 min and the contentsof chlorophylls and carotenoids were quantified (Lichtenthaler,1987).

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2.5. Determination of H2O2, lipid peroxidation and free proline

Hydrogen peroxide was extracted from 100 mg of frozen tissueusing liquid N2 and macerated with 1 mL of phosphate buffer(50 mM, pH 6.5). The homogenate was centrifuged at 6000g for25 min. To determine the H2O2 levels, 0.1 mL of extracted solutionwas mixed with 1 mL of 0.1% (w/v) titanium sulphate in 20% H2SO4,and the mixture was then centrifuged at 6000g for 15 min. Readingsof the supernatant were made at 410 nm (Shimadzu UV-240 spec-trophotometer). H2O2 levels were calculated using a molar extinc-tion coefficient 2.8 mM!1 cm!1 (Domínguez et al., 2009).

The determination of thiobarbituric acid reacting substances(TBARS), as an indicator of membrane lipid peroxidation, was mea-sured in terms of malondialdehyde (MDA) content. Shoots and rootswere grinded to a fine powder in liquid N2 with a mortar and pestle.Subsequently about 200 mg of powder were homogenised in 2 mL of0.1% (w/v) trichloroacetic acid (TCA), and a centrifugation at 10 000gfor 5 min was followed. For every 250 lL of the supernatant, 1 mL of20% (w/v) TCA containing 0.5% (w/v) thiobarbituric acid was added.The mixture was heated at 95 #C for 30 min and then quickly cooledin an ice bath for 10 min. After cooling the mixture was centrifugedat 10 000g for 15 min and the absorbance of the supernatant wasmeasured at 532 nm. Measurements were corrected for the non-specific turbidity by subtracting the absorbance at 600 nm. The con-centration of MDA was calculated using the extinction coefficient155 mM!1 cm!1 (Heath and Packer, 1968).

Free proline was extracted from frozen tissue samples in 3% (w/v) sulphosalicylic acid with quartz sand at 4 #C. After centrifugation(500g for 10 min at 4 #C) the level of proline was quantified in thesupernatant (Bates et al., 1973).

2.6. Glutathione-S-transferase (GST) and guaiacol peroxidase (GPX)assays

For the GST assays 1 g of the frozen aliquots were homogenisedwith 4 mL of grinding buffer (50 mM Tris–HCl, 1 mM EDTA, 1 mMPMSF, 1 mM DTT and 0.05% (w/v) PVPP, pH 7.5) at 4 #C. Homoge-nates were centrifuged at 20 000g for 25 min at 4 #C, the superna-tant recovered and protein content determined by the Bradfordmethod (Bradford, 1976). The enzymatic assay consisted on50 mM phosphate buffer (pH 7.5), 1 mM 1-Chloro-2,4-Dinitroben-zene (CDNB) and 100 lL of supernatant. The reaction started uponthe addition of GSH (to a final concentration 1 mM) and the in-crease in absorbance at 334 nm was followed spectrophotometri-cally for one minute. The non-enzymatic conjugation of CDNBwas determined by replacing the protein extract by grinding buf-fer. The latter DAbs334nm/min was subtracted to the first value toestimate the real GST activity using the extinction coefficient9.6 mM!1 cm!1 (Li et al., 1995).

For accessing GPX activity, frozen tissue aliquots were homog-enized, at 4 #C, with 3 mL phosphate buffer (100 mM, pH 7.3) pergram of tissue. Homogenates were centrifuged at 10 000g for10 min at 4 #C. The supernatant was recovered and protein contentquantified by the Bradford method (Bradford, 1976). GPX activitywas measured by the oxidation of Guaiacol in the presence ofH2O2 within 1 min. Each assay contained 50 mM phosphate buffer(pH 7.3), 60 mM Guaiacol, 1 mM H2O2 and 20 lL of extract. Thereaction was started by the addition of H2O2 and followed spectro-photometrically at 470 nm. Activities were calculated consideringthe extinction coefficient 26.6 mM!1 cm!1 (Amako et al., 1994).

2.7. Fisher and t-tests

All assays and measurements were performed at least in tripli-cate (n P 3). Variance analysis was performed by Fisher test and

the means were statistically analysed using a two-sided t-test. Sta-tistical significance is assumed at P < 0.05.

3. Results and discussion

3.1. Seed germination and seedling development

Because seed germination was not affected by the exogenousmetalaxyl concentrations used (data not shown) it is possible toconclude that direct seeding of the remediation site can be donein order to implement its colonisation by S. nigrum.

Seedling dry weight significantly decreased along with theincreasing metalaxyl concentrations used (Fig. 1), starting fromthe 25 mg L!1 treatment, having been registered 1.7 and 2.0-folddecreases for the seedlings grown exposed to 25 and 50 mg L!1,respectively, 2.3-fold for the 75 mg L!1 and 4.4-fold for the100 mg L!1 treatments.

Seedling shoot height started to decrease with the 50 mg L!1

treatment, and this tendency paralleled the increasing metalaxylconcentrations, being only significant for the 100 mg L!1 treatment(1.5-fold) (Fig. 2a), revealing metalaxyl toxicity to seedlings at veryhigh concentrations.

Root length significantly decreased with metalaxyl concentra-tions equal or superior to 12.5 mg L!1, being more pronouncedfor higher pesticide concentrations (1.4, 1.6, 1.6, 1.8 and 3.0-foldfor the 12.5, 25, 50, 75 and 100 mg L!1 treatments, respectively)(Fig. 2b). The described data reveals that seedling growth is com-promised starting from the 12.5 mg L!1 concentration, mainly asa consequence of the root size reduction. In fact, root elongationmay be considered a test to assess the influence of organic com-pounds on plants’ early developmental stages (Kummerová et al.,2008). As the other biometric parameters were not affected bythe 12.5 mg L!1 treatment, and because the recommended dosein crop production (600 g a.i ha!1) corresponds to 0.8 mg kg!1 soil(assuming soil depth 5 cm and density 1.5 g mL!1) (Sukul, 2006)and the expected metalaxyl concentrations in runoffs range fromaround 5 to 47 lg L!1 (Wilson et al., 2001), thus largely inferiorto the concentrations used in this study, one can conclude that S.nigrum seedlings are capable to develop into plants when in con-tact to metalaxyl concentrations up to 12.5 mg L!1. In practicalterms, these results further support the possibility of direct seedingfor the colonisation of the phytoremediation site with S. nigrum,this way decreasing the need of more man-power for manuallyplanting it.

3.2. Metalaxyl effects on plant growth

Metalaxyl was deleterious to plant growth starting with the12.5 mg L!1 solution, as visible and significant macroscopic

Fig. 1. S. nigrum seedling dry weight (mg) in response to the increasing exogenousmetalaxyl concentrations after 4 weeks of exposure. Columns represent mean ± SD(n P 3). Dotted columns are statistically different from the control (P < 0.05).

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differences started to be observed with this treatment. Minimalplant growth occurred with the 50 mg L!1 treatment (data notshown) and the lowest concentration used (6.25 mg L!1) did notexert any negative effect on plant morphology. As a consequence,metalaxyl concentrations of 50 mg L!1 and above were not consid-ered in the following.

Shoot height and root length decreased significantly when themetalaxyl was added to the substrate mixture at concentrationshigher or equal to 12.5 mg L!1, as observed in Fig. 3. Shoot heightdecreased around 2.0 and 3.6-fold in the 12.5 mg L!1 and25 mg L!1 treatments, respectively. Root length decreased approx-imately 1.2 and 1.8-fold respectively in the 12.5 mg L!1 and25 mg L!1 treatments.

Only the shoots and roots collected from the 25 mg L!1 treat-ment showed a significant 11-fold decrease in fresh weight

(Fig. 4a and b, respectively). No significant variations on dry weightcould be observed for all treatments (data not shown).

The reduction in plant biomass, as well as other deleterious ef-fects, in response to a 7-d exposure period to metalaxyl in concen-trations equal or superior to 25 mg L!1 has been reported forseveral plant species (Wilson et al., 2001). Nevertheless, the datacollected in the present study was obtained after a 4-weeks expo-sure period, which corresponds approximately to the full life cycleof this plant species (floral buds were visible at the end of this per-iod of time in all treatments up to 25 mg L!1). This demonstratesthat treated plants can complete their life cycle when exposedup to 25 mg L!1 of metalaxyl, this way allowing the remediationsite to be naturally colonised by the produced seeds. As so, S. ni-grum reveals to be an excellent candidate for the remediation ofmetalaxyl-contaminated sites as it withstands being exposed tothis compound for a large period of time.

These results show that S. nigrum development becomes com-promised when exposed to metalaxyl concentrations equal orsuperior to 12.5 mg L!1. Because plant biomass derived from the25 mg L!1 exposure was too low and because such concentrationsare not expected to occur in either runoffs or metalaxyl-containingeffluents (Wilson et al., 2001), this treatment was not consideredfor future determinations.

3.3. In planta metalaxyl levels

As shown in Fig. 5, it is possible to verify that 12.5 mg L!1-ex-posed S. nigrum plants were able to extract and accumulate metal-axyl in their tissues. Shoots corresponded to the plant part wheremost of the metalaxyl extracted by the roots was accumulated,being this value about eight times higher then the value deter-mined in roots, which corresponds to a bioaccumulation factor ofabout 25 for shoots and of 3 for roots.

Fig. 2. S. nigrum seedling shoot height (a) and root length (b), expressed in cm, inresponse to the increasing exogenous metalaxyl concentrations after 4 weeks ofexposure. Columns represent mean ± SD (n P 3). Dotted columns are statisticallydifferent from the control (P < 0.05).

Fig. 3. S. nigrum shoot and root size (cm) reduction in response to increasingmetalaxyl concentrations in the substrate mixture after 4 weeks of exposure.Columns represent mean ± SD (n P 3). Asterisks indicate statistical differences fromthe control (P < 0.05).

Fig. 4. S. nigrum shoots (a) and roots (b) fresh weight, expressed in grams, after4 weeks of exposure to increasing metalaxyl concentrations in the rhizosphere.Columns represent mean ± SD (n P 3). Dotted columns are statistically differentfrom the control (P < 0.05).

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These results show that this plant species is adequate to phyto-extract metalaxyl from contaminated sites and solutions, as moremetalaxyl is accumulated in shoots than in roots. The differencesin metalaxyl levels between shoots and roots may be explained be-cause metalaxyl has a log KOW of 1.64 (see references in (Wilsonet al., 2001)), meaning that it is phloem- and xylem-mobile (Sch-roder and Collins, 2002), and therefore it may be transported tothe shoots via the transpiration stream.

Information on plant metabolism of metalaxyl or metalaxyl-M(enantiomers) can be found essentially on the reports elaboratedby organisations such as at the Joint FAO/WHO Meetings on Pesti-cide Residues (JMPR) (Metalaxyl-M, first draft prepared by DenisHamilton, JMPR, 2002) or the European Commission (Review re-port for the active substance Metalaxyl-M, SANCO/3037/99-final,European Commission, 2002). These reports are solely based onunpublished studies performed under the responsibility of theindustrial producers. Although the employed analytical methodsare briefly described, detailed information on metabolite identifi-cation, namely the electron-impact (EI) fragmentation mass spec-tra are not provided.

The FAO report presents results of the residue compositionfound in tubers and foliage of potato plants, closely related to S. ni-grun. The parent compound, metalaxyl, was a major component inthe tubers and foliage, but significant metabolites, especially N-(2-hydroxymethyl-6-methylphenyl)-N-(methoxyacetyl) alaninemethyl ester and its glucose conjugate were also found in foliage.This metabolite is simply obtained by the hydroxylation of amethyl group connected to the aromatic ring of metalaxyl. Onthe basis of gas chromatography performance characteristics, thesetwo structures are distinctive enough to be resolved by the chro-matographic method employed in the present work and so theeventual presence of the metabolite would be detectable, whichwas not the case, therefore indicating its absence in the plant or-gans analysed.

Even though the mass spectrum is not available in our NISTdatabase a fairly good idea of what it should look like can be drawnfrom the knowledge of the routes of fragmentation of commonfunctional groups. The benzyl alcohol structure is known to haveas base peak its molecular cation (parent ion, non-fragmented)which indicates that this sub-structure of the metabolite is quitestable to EI fragmentation, leading to the expectation that a visiblefragment of molecular mass at m z!1 295 would be found since thatof metalaxyl at m z!1 279 is found. The stability introduced by thebenzyl alcohol group would also allow finding some of the mostabundant fragments present in the metalaxyl mass spectrum butbearing m z!1 incremented in 16 units (mass difference betweenmetabolite and metalaxyl due to oxygen addition). No chromato-graphic peaks bearing such characteristics or others that might

make us suspect of the presence of a metabolite of metalaxyl werefound, both in root and shoot samples. One possible explanationlies on the glucose conjugate form of the metabolite. This formdoes not seem likely to be amenable to GC, it is probably thermallydegraded at the desorption chamber, at 250 #C, to unknown prod-ucts that may never leave the microextraction fibre or reach themass analyser. A more detailed study would have to be setup,employing other analytical means, such as LC–MS, and preferablyin possession of analytical standards of all the compounds of inter-est (not available commercially), if metabolite identification/quan-tification would be a major expected outcome of the present work.

According to the nature of the metalaxyl quantification proce-dure used, it is possible to state that most of the metalaxyl reachedthe shoots in its native form, i.e., suffering minimal chemical trans-formation by the plant. So, the obtained results show that by con-centrating this unchanged pesticide in its aboveground tissues,black nightshade allows the convenient disposal of the pollutantby cutting the shoots off. Since after cutting off the shoots theunderground part of the plant remains in the site, the newbranches thus formed will continue to accumulate the pollutant,allowing a continuous and efficient metalaxyl removal. Further-more, by also accumulating metalaxyl in roots, this plant speciesalso contributes for its phytostabilisation, this way preventing itsrelease to the environment, plus favouring its degradation by therhizosphere microbial community (Pilon-Smits, 2005).

3.4. Biochemical determinations

Hydrogen peroxide levels were always higher in shoots than inroots, independently of the experimental situation analysed, a sit-uation easily understandable by the fact that shoots are photosyn-thetic organs, where electron transfer reactions are constantlyoccurring leading to ROS formation (Taiz and Zeiger, 2010). The12.5 mg L!1 metalaxyl treatment induced a significantly 1.2-foldincrease in the H2O2 levels in shoots, as observed in Fig. 6a, whileproline levels significantly increased by 1.4-fold in roots from thetreated plants, as depicted in Fig. 6b.

Hydrogen peroxide levels increases in response to oxidativestress imposed by organic pollutants exposure have already beenreported in Brassica (Singh et al., 2006) and the increase in H2O2

and proline levels detected in this study in response to metalaxylexposure indicate that plants suffered mild oxidative stress whenexposed to 12.5 mg L!1 (Pernia et al., 2008). The amino acid prolineis known to be involved in the response to several biotic and abi-otic stresses (Skopelitis et al., 2006), preventing damage to plantcells. The data obtained in this work suggests that the high prolinelevels in the roots exert an important protective role there, as rootscorrespond to the plant organ that permanently is in close contactwith metalaxyl, and therefore, more prone to cellular damage.

No significant changes in the photosynthetic pigments levels(chlorophyll a, chlorophyll b and carotenoids), as well as in the li-pid peroxidation, were detected in the described treatment (datanot shown), indicating that the oxidative stress plants sufferedwhen exposed to metalaxyl was not severe (Amaya-Chávez et al.,2006; Pernia et al., 2008) and/or that other protective mechanisms,simultaneously to proline accumulation, were put to action like in-creased glutathione levels, known to occur in response to an oxida-tive burst (Edwards et al., 2005), this way reducing the harmfuleffects of the produced ROS.

3.5. Enzyme activities

Protection against xenobiotic-induced damage is primarilyexhibited by: (a) direct biotransformation of the xenobiotic tomore polar and less toxic compounds, suitable for export fromthe cell (Edwards et al., 2005), having also as result a decreased

Fig. 5. Metalaxyl levels in shoots and roots of the 12.5 mg L!1-treated plantsexpressed as mg kg!1 dry weight. Columns represent mean ± SD (n P 3).

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biological half-life (Coleman et al., 1997) and (b) cleaning up toxicintermediates formed from reactive oxygen intermediates genera-tion (Edwards et al., 2005).

Metalaxyl is readily activated by ester hydrolysis in phase I, atleast in sunflower plants, the major metabolites being the corre-sponding carboxylic acids (Zadra et al., 2002) that will be involvedin phase II reactions, but other activation reactions may occur dueto the relatively high diversity in chemical functional groups pres-ent in this molecule.

Glutathione (GSH) is an important metabolite that acts both asa reducing agent, protecting cells against oxidative stress, and as anucleophile by covalently binding to xenobiotics inhibiting chem-ical toxicity (Coleman et al., 1997). Furthermore, one of the mostcommonly conserved responses to oxidative burst in plants is theenhanced synthesis of GSH along with the accumulation of GSH-dependent proteins, such as GST (Teisseire and Vernet, 2001; Ed-wards et al., 2005).

Measured GST activities were always higher in roots than inshoots, independently of the experimental situation analysed.GST activity in roots significantly increased by 7-fold in12.5 mg L!1-treated plants when expressed by protein content. Inshoots from the same plants this significant increase was evenhigher: 14-fold (Fig. 7a). The observed increases on a protein basissuggest de novo synthesis of more GST. Following these results, itis possible that the enhanced GST activities registered in both plantorgans make some metalaxyl nontoxic, thus playing some protec-tive role in the detoxification process (Coleman et al., 1997). Thehigher GST activities in roots suggest that these plant organs corre-spond to the major site where the xenobiotic may be partially con-jugated to GSH, while the increase in the activity in shoots may bea reflection of the higher metalaxyl levels present in this plant part,as a possible way to decrease its toxic effects to the plant.

As this xenobiotic may also be conjugated to other moleculesbesides GSH both in roots as in shoots, it is an indication that S. ni-grum plants are able to extract more metalaxyl from the environ-ment then that quantified in their tissues (although conjugatedforms were not detected by the quantification method used), fur-ther emphasising its use as a powerful tool for metalaxyl-contam-inated sites remediation. However, and as already stated, a moredetailed study would have to be setup, employing other analyticalmeans, for metabolite identification/quantification to further dis-sect the apparently little metalaxyl plant metabolism.

Accordingly to the increased H2O2 levels in response to metal-axyl exposure, the highest GPX activities were also found in the12.5 mg L!1-treated plants, being detected in roots and shoots a2 and 6-fold significant increased activity, respectively, as observedin Fig. 7b.

Because the technique used in this study does not allow the dis-crimination between ascorbate peroxidases (APX) and GPX, thevalues presented are the reflection of both peroxidase activities(Amako et al., 1994). The data obtained in this work suggests thatthe increased peroxidase activities contributed for plant protectionfrom the deleterious effects of the elevated H2O2 levels in bothplant parts, similarly to what happens when duckweed is exposedto folpet (Teisseire and Vernet, 2001).

4. Conclusions

This work clearly shows that S. nigrum plants are capable ofremediating metalaxyl-contaminated sites (effluents or soils),when present in concentrations up to 12.5 ppm.

S. nigrum may be directly seeded at the remediation site orplanted at an initial developmental stage after germination.

Fig. 6. (a) Hydrogen peroxide levels detected in shoots and roots of control and12.5 mg L!1 metalaxyl-treated plants for 4 weeks expressed as nmole per gram offresh weight. (b) Proline levels determined in shoots and roots of control andmetalaxyl-treated plants expressed as lg proline per gram of fresh weight. Columnsrepresent mean ± SD (n P 3). Asterisks indicate statistical differences from thecontrol (P < 0.05).

Fig. 7. (a) GST activities detected in shoots and roots of control and 12.5 mg L!1

metalaxyl-treated plants for 4 weeks expressed as nkat per mg of protein. (b) GPXactivities detected in shoots and roots of control and 12.5 mg L!1 metalaxyl-treatedplants for one month, expressed as nkat per mg of protein. Columns representmean ± SD (n P 3). Asterisks indicate statistical differences from the control(P < 0.05).

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Author's personal copy

Although the exposure to 12.5 mg L!1 metalaxyl concentrations in-duces a mild oxidative stress to plants, antioxidant defences suchas proline production and H2O2-scavenging enzymes appear toeffectively protect the plant and decrease the H2O2-related delete-rious effects, allowing plants to thrive until the end of their lifecycle.

Finally, S. nigrum plants have proven to be adequate to phytoex-tract metalaxyl, as it accumulates mainly in the aboveground tis-sues, thus allowing its proper removal by cutting off these plantparts.

Acknowledgments

The authors gratefully acknowledge the University of Porto forfinancial support (Project PhotoWeed, IJUP 2009/10) with the con-tribution of Santander Totta.

AMTS acknowledges financial support from POCI/N010/2006.

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