Science of the Total Environment 425 (2012) 262–270

Contents lists available at SciVerse ScienceDirect

Science of the Total Environment

j ourna l homepage: www.e lsev ie r .com/ locate /sc i totenv

Poplar clones of different sizes, grown on a heavy metal polluted site, are associatedwith microbial populations of varying composition

Elisa Gamalero a, Patrizia Cesaro a, Angela Cicatelli b, Valeria Todeschini a, Chiara Musso a,Stefano Castiglione b, Arturo Fabiani c, Guido Lingua a,⁎a Università del Piemonte Orientale “Amedeo Avogadro”, Dipartimento di Scienze e Innovazione Tecnologica, viale T. Michel 11, 15121, Alessandria, Italyb Università degli Studi di Salerno, Dipartimento di Chimica e Biologia, via Ponte don Melillo, 84084, Fisciano (SA), Italyc Centro per la Ricerca e la Sperimentazione in Agricoltura, Centro di ricerca per l'Agrobiologia e Pedologia (ABP), Piazza Massimo d'Azeglio 31, 51121 Firenze, Italy

⁎ Corresponding author. Tel.: +39 0131 360 233; faxE-mail address: guido.lingua@mfn.unipmn.it (G. Lin

0048-9697/$ – see front matter © 2012 Elsevier B.V. Alldoi:10.1016/j.scitotenv.2012.03.012

a b s t r a c t

a r t i c l e i n f o

Article history:Received 27 October 2011Received in revised form 7 March 2012Accepted 7 March 2012Available online 3 April 2012

Keywords:Heavy metalsSoilPoplarPhytoremediationCulturable bacteriaDGGE

We performed a field trial to evaluate the response of different poplar clones to heavy metals. We found thatpoplar plants of the same clone, propagated by cuttings, had a marked variability of survival and growth indifferent zones of the field that were characterized by very similar physical–chemical prosperities. Sincemetal uptake and its accumulation by plants can be affected by soil microorganisms, we investigated soil mi-crobial populations that were collected in proximity to the roots of large and small poplar plants. We usedmicrobiological and molecular tools to ascertain whether bacterial strains or species were associated withlarge, or small poplars, and whether these were different from those present in the bulk (without plants) soil.We found that the culturable fraction of the bacteria differed in the three cases (bulk soil, small or large pop-lars). While some taxa were always present, two species (Chryseobacterium soldanellicola and Variovoraxparadoxus) were only found in the soil where poplars (large or small) were growing, independently fromthe plant size. Bacterial strains of the genus Flavobacteriumwere prevalent in the soil with large poplar plants.The existence of different microbial populations in the bulk and in the poplar grown soils was confirmed bythe DGGE profiles of the bacterial culturable fractions.Cluster analysis of the DGGE profiles highlighted the clear separation of the culturable fraction from thewhole microbial community. The isolation and identification of poplar-associated bacterial strains from theculturable fraction of the microbial community provided the basis for further studies aimed at the combineduse of plants and soil microorganisms in the remediation of heavy metal polluted soils.

© 2012 Elsevier B.V. All rights reserved.

1. Introduction

Phytoremediation can be defined as the use of plants for the vola-tilization, stabilization, degradation and/or extraction of pollutantsfrom contaminated soils, while preserving or restoring the biologicaland physical structure of the soil itself (Pilon-Smits, 2005). Poplarshave been shown to possess the main useful traits for phytoremedia-tion, such as the ability to grow on nutrient-poor soils, a deep andwide-spreading root system, metal resistance or tolerance and a fastrate of growth (Baldantoni et al., 2011; Di Baccio et al., 2003;Sebastiani et al., 2004; Soudek et al., 2004). In addition, poplarsshow a large clonal variability in their heavy metal (HM) toleranceor accumulation (Dos Santos Utmazian et al., 2007; Kopponen et al.,2001; Laureysens et al., 2004, 2005; Zalesny et al., 2005).

For this reason, in a previous study, a large number of Populus albaL. and Populus nigra L. clones, growing in an area highly polluted by

: +39 0131 360 243.gua).

rights reserved.

copper and zinc, were screened for HM tolerance; two white poplarand four black poplar clones were selected for their high survival per-centage, and were characterized for their metal accumulation abilityand polyamine leaf concentration (Castiglione et al., 2009).

In the present study we describe the composition of the microbialpopulations observed after the establishment of a new experimentalfield in the same industrial site. The experimental field was arrangedin randomized blocks consisting of the previously selected clones andsome commercially available ones (namely Jean Pourtet andBeaupré). The hybrid poplar clone I-214 (Populus×canadensisMoench) [registered to the RNCF (Registro Nazionale Cloni Forestali)]was used as a spacer between the cuttings of each different clone. Theinitial purpose of this supplementary field trial was to confirm previ-ous results and investigate the soil reclamation after some years ofgrowth. Some unexpected observations, however, prompted differentanalyses on unplanned topics. While the overall survival of theplanted clones was consistent with the previous data, we observedthat cuttings of the same poplar clone showed a marked variabilityof survival and growth in different zones of the field, even thoughthe chemical features of the soil in the various areas were not

263E. Gamalero et al. / Science of the Total Environment 425 (2012) 262–270

significantly different. As metal phytoextraction and accumulation inplants can be affected by soil microorganisms (Shilev et al., 2001;Abou-Shanab et al., 2008; Solìs-Domìnguez et al., 2011), we decidedto study by means of microbiological and molecular tools the soil mi-crobial populations collected in proximity of the roots of large andsmall poplar plants. Bacterial viability, activity and density are inhibitedby heavy metals. However, under this environmental pressure, somebacterial populations can adapt to the pollutant, become dominantand, attracted by root exudates, colonize the roots of the plants. Thesebacterial populations can support plant growth either by relievingtoxicity by decreasing metal bioavailability (releasing chelators, modi-fying the soil pH or the redox potential), or by increasing plant toleranceto metals by various mechanisms (reviewed by Gamalero et al., 2009).

Little is known about the whole microbial communities associatedwith poplar roots (van der Lelie et al., 2009; Gottel et al., 2011). Veryrecently, the bacterial and fungal communities living on and withinthe root of Populus deltoides Bartram ex Marsh. in mature naturaltrees were described (Gottel et al., 2011).While the poplar rhizosphereis dominated by Acidobacteria and Alphaproteobacteria, Gammaproteo-bacteria (54%) and Alphaproteobacteria (23%) are prevalent among theendophytes. Since poplar is frequently exploited for phytoremediationpurposes, attention was first focused on the isolation and characteriza-tion of culturable bacterial endophytes that were potentially useful forplanning assisted phytoremediation (Van Aken et al., 2004; Doty et al.,2005; Moore et al., 2006) and only later on the characterization ofwhole endophytic communities (Ulrich et al., 2008).

In contrast, information regarding the bacterial communities asso-ciated with the roots of poplar plants grown in heavy metal pollutedsoil is lacking. To our knowledge, this is the first study aimed at de-scribing the bacterial communities associated with the roots of poplarclones grown in a heavy metal polluted soil, under open field condi-tions. Large and small poplars were analyzed independently because:(1) according to Castiglione et al. (2009), we know that in the samefield, larger poplars have higher survival chances; (2) some bacterialtraits important in plant responses to environmental stress can resultin increased plant growth: this is the case, for instance, for the abilityto synthesize 1-aminocyclopropane-1-carboxylic acid deaminase orindole-3-acetic acid synthesis (Gamalero et al., 2009).

The aim of this study was to isolate and identify bacterial strainsthat are unique to large or small poplars and are different fromthose present in the bulk (without plants) soil.

2. Materials and methods

2.1. Soil characteristics and description of experimental field

The experimental field was established in a long abandoned agri-cultural area near the town of Serravalle Scrivia (Alessandria) innorthern Italy (UTM coordinates: 32T, 4955850 N, 487340 E). Thefield is located close to an industrial plant manufacturing semi-finished copper and zinc alloy products, whose fumes have heavilypolluted the surrounding area. The soil is loam (USDA: sand 31%, silt46%, clay 23%), with a slightly acidic pH (6.3 on average, but rangingfrom 5.0 to 7.4) and organic matter is 4.34%. N, P and K are 42.02,17.20 and 57.07 mg kg−1 DW, respectively. Mean soil Zn concentrationis about 950 mg kg−1 DW in the first 20 cm of soil; in the second layer(20–40 cm) it is almost identical, while it decreases slightly in the thirdlayer (40–60 cm). Cu concentration decreases with increasing depth,from ca 1300 mg kg−1 DW in the first layer, to about 1200, and1100 mg kg−1 DW in the second and third layers, respectively. Amore detailed description of the site and of the methods used for itschemical characterization is given in Ravera et al. (2006) and inCastiglione et al. (2009). Before the establishment of the experimentalfield, the vegetation was largely dominated by Dog's Tooth Grass[Cynodon dactylon (L.) Pers.], and large patches of ground had no vege-tation at all.

In March 2004, a plantation was established with unrooted cut-tings of two P. alba clones, named AL35 and AL22, four P. nigra clones,named NG12, NG19, SN26 and SN56, all of them selected on the basisof their survival percentages on polluted soil as described byCastiglione et al. (2009), and with some registered clones: P. alba(clone Villafranca), P. nigra (clone Jean Pourtet), Populus×generosaHenry (clone Beaupré).

Prior to planting, the field was leveled and cleared of the largeststones and other debris; a rotor tiller was used for final soil prepara-tion. The clones were planted as 20-cm long unrooted cuttings afterbeing soaked in running tap water for 24 h. Cuttings were plantedmanually in a single-row design with an inter-row distance of0.75 m, and a space of 0.35 m between cuttings, to a depth of 18 cm,leaving just one bud above the soil surface. A randomized block de-sign was used with 8 rows of 28–32 cuttings, for a total of over 250cuttings. To avoid the border effect, one row of the registered cloneI-214 (P.×canadensis) was planted along the borders of the experi-mental plot. Clone I-214 was also used as a spacer between theother poplar clones. The experimental field was irrigated every15 days from May to the end of August. Mechanical weeding was car-ried out twice during the growing season. A commercial organicslow-release fertilizer (total organic N 4.5%, organic C 22%, P2O5 3%;Grenagro Medio Plus, Grena, S. Bonifacio, Verona, Italy) was appliedtwice (200 kg ha−1).

2.2. Evaluation of clone performance

In October 2004, six months after setting the experimental field,percentage survival and the size of the poplars were estimated; thelatter was evaluated on a 1–3 scale, where: 1, b20 cm; 2, >20 andb50 cm; 3, >50 cm high. The same data collection was repeated thefollowing year (October 2005, second growth season).

For the sampling of soil and the following microbiological analy-ses, we choose to investigate I-214 plants of different size (largeand small), as these were the most abundant ones in the experimen-tal field; furthermore they were evenly distributed in the field usedfor the trial.

2.3. Soil sampling and chemical analysis

The bulk soil (where plants were not introduced) and the soil as-sociated with the roots of small (I-214S) or large (I-214L) poplars,five per each kind, were sampled at a depth of 30 cm, after removingthe surface layer (3.0–5.0 cm). Three soil cores were taken in theproximity of the poplar rhizosphere, therefore a total of fifteencores were taken for each kind of soil; the soil was then pooled andthe following parameters were analyzed in the three soils followingthe methods established by the Italian law (G.U. 248/99): moisturecontent, pH, total organic carbon (TOC), soil organic matter (SOM),bioavailable and total P. Determination of Cu and Zn concentrationsin the soils was performed following the EPA (Environmental Protec-tion Agency) method no.6020a (http://www.epa.gov/epaoswer/hazwaste/test/pdfs/6020a.pdf) by using an ICP-MS (InductivelyCoupled Plasma-Mass Spectrometry, Agilent Technologies Inc., Paloalto, CA, mod. 4500) technique with terbium (159Tb) and rhodium(103Rh) as internal standards (100 μg/ml each). For biological pur-poses, an aliquot of each soil sample was aseptically collected,mixed into one pool and maintained at −20 °C and −80 °C for DNAand RNA extraction, respectively, while for the culturable fractionanalysis, soil samples were immediately processed.

2.4. Determination of the plant copper and zinc concentration

Approximately 0.5 g DW each of leaves, stems and roots was usedfor the determination of Cu and Zn concentration, separately. Sampleswere weighed and then digested in 10 ml concentrated HNO3 in a

264 E. Gamalero et al. / Science of the Total Environment 425 (2012) 262–270

CEM MARS 5 microwave digestor (Cologno al Serio, BG, Italy). Thedigested material was filtered through 45-μm filters, and then deio-nized water was added to a final volume of 100 ml. Metal concentra-tion was assessed by means of a calibration curve, after measurementby inductively coupled plasma optic emission spectrometry (ICP-OES)using an IRISAdvantage ICAPDUOHR series (Thermo Jarrell Ash, Franklin,MA, USA) spectrometer. Certified standards (BCR 062, 100, 129 and145R, by the Institute for Reference Materials and Measurements,Ratieseweg, Belgium), of known element concentration, were analyzedwith the samples in order to confirm the accuracy of the procedure.

2.5. Soil enzymatic activities

Total microbial activity was measured using the fluorescein (FDA)diacetate hydrolysis assay following the method of Adam and Duncan(2001). Acid phosphomonoesterase was measured according to Eivaziand Tabatabai (1977). Alkaline and neutral phosphomonoesteraseswere evaluated following Alef and Nannipieri (1995). Cellulase activityon (CMC) was evaluated as previously described by Schinner and vonMersi (1990). β-Glucosidase activity was evaluated in agreement withEivazi and Tabatabai (1988). Urease was evaluated according toBremner andMulvaney (1978). All the determinations of enzymatic ac-tivities were replicated five times and the values were expressed as theaverages of the five spectrophotometrical measurements.

2.6. Soil metabolic fingerprinting

Ten grams of fresh soil from each site was suspended in 90 ml of0.18% sodium pyrophosphate and shaken vigorously for 2 h. Soil ex-tracts were serially diluted in 0.1 M MgSO4, to the representative di-lution and centrifuged at 1000 rpm for 10 min; 150 μl of thesupernatant was inoculated in each well of the GN (Gram negative)Biolog microplates (Biolog, Hayward, CA, USA). Three plates for eachsite were incubated at 25 °C for 7 days, read manually and digitizedevery day in order to follow the microbial activity over time. The inten-sity of the color developed in each well was expressed as a numericalvalue: 2 for a positive reaction, 1 for a borderline reaction and 0 for noreaction.

2.7. Assessment of the culturable bacterial density

Fifty grams of fresh soil from each site was added to 450 ml of so-dium pyrophosphate (0.18%), shaken at 25 °C, 180 rpm for 2 h andleft to settle for 30 min. Serial dilutions were performed in sulfatebuffer and 100 μl of each suspension was distributed in triplicate on1/10 Trypticase Soy Agar (TSA-Fluka, Milan, Italy) added with100 μg ml−1 of cycloheximide (MP Biomedicals, France). Bacterialdensity was recorded after 2 and 7 days of incubation at 28 °C andexpressed as cfu g−1 of dry soil.

Thirty-four colonies from a representative dilution were selectedand isolated from 1/10 TSA plates. Cultures of bacterial cells, grownin Trypticase Soy Broth were stored at −80 °C in 50% glycerol.

2.8. Identification of culturable bacteria

Total DNA was extracted from single bacterial colonies usingREDExtract-N-Amp Plant PCR KIT (Sigma-Aldrich, Milano, Italy). Bac-terial cells were added to 20 μl of Extraction Solution. After incubationat 95 °C for 20 min, 20 μl of Dilution Solution was added to dilute DNAExtraction solution.

16S rDNA genes were amplified by PCR using REDExtract-N-AmpPCR ReadyMix, included in the kit and the specific primer pairs(Gentry et al., 2004; Wilson et al., 1990). PCR condition for 16SrDNA amplification was: 1 min denaturation at 94 °C, 1 min annealingat 55 °C, extension 2 min at 72 °C, for 35 cycles. Initial denaturationand final extension were 3 min at 94 °C and 7 min at 72 °C. Amplified

DNA was purified using a PCR purification kit (GE Healthcare, Milan,Italy) following the supplier instructions and quantified by agarose gelelectrophoresis (1.0% in TAE buffer). The purified DNA products weresequenced using ABI Big Dye Terminator version 3.1 Cycle SequencingReady Reaction Kit (Applied Biosystems, Milano, Italy) and PCR se-quencing products were run on ABI 310 Automated Genetic Analyzer(Applied Biosystems, Milan, Italy).

For sequencing PCR products, the same primer pairs used for am-plification of 16S rDNA regions (338F, 5′CTCCTACGGGAGGCAGCAG3′;1492R, 5′GGTTACCTTGTTACGACTT3′) and two internal primers (27F,5′ AGAGTTTGATCCTGGCTCAG3′; 784R, 5′ GGACTACCAGGGTATC-TAATCC3′) were employed to fully cover the entire 16S rDNA se-quence. The thermal cycling profile of the sequencing reaction was:10 s denaturation at 94 °C, 5 s at 55 °C, extension 4 s at 64 °C for25 cycles. The chromatograms were assembled using SequencingAnalysis Software version 5.1 (Applied Biosystems, Milan, Italy) andtransformed in a text file for further bioinformatics analysis. DNAsequences were compared against all bacterial 16S rDNA referencesequences available at the NCBI World Wide Web database (http://www.ncbi.nlm.nih.gov/BLAST/).

2.9. DNA extraction from culturable bacteria and soil

The DNA was extracted from cumulative pellets of the culturablebacterial heterotrophic fraction by the CTAB method (Ausubel et al.,1987) and directly from 1 g of soil using the UltraClean™ Soil DNAKit (MO BIO Laboratories, Inc., Carlsbad, CA) following the manufac-turer's instructions. Extractions were performed in triplicate.

2.10. PCR amplification of 16S rDNA for DGGE analysis

For DGGE analysis of the culturable or total Eubacterial communi-ty, V6–V8 regions of 16S rDNA were amplified with primers GC986fand Uni1401r as described by Felske et al. (1998). Each reaction mix-ture (50 μl) contained 25 ng of amplified soil DNA, 250 μM of dNTPs,1.5 mM MgCl2, 1× Buffer (67 mM Tris–HCl pH 8.8; 16.6 mM(NH4)2SO4; 0.01% Tween-20; MgCl2 5 mM), and 2.5 U of Taq DNAPolymerase (Polymed, Florence, Italy). The reaction was performedin a Techne thermocycler (Bibby Scientific, Riozzo di Cerro al Lambro,Italy) with the following thermal protocol: one initial cycle at 94 °Cfor 1 min and 30 s, 56 °C for 30 s, 72 °C for 45 s, subsequently 33 cy-cles at 95 °C for 20 s, 56 °C for 30 s and 72 °C for 45 s, followed by afinal extension step at 72 °C for 5 min. Each sample was amplifiedthree times and the amplicons were pooled together before DGGEanalysis.

2.11. DGGE analysis of the Eubacterial community

The analysis was performed with the INGENYphorU-2 System(Ingeny International BV, Goes, The Netherlands) on a 6% polyacryl-amide gel (acrylamide/bis ratio 37.5:1), under denaturating condi-tions (urea, 7 M; 40% formamide with a denaturing gradient rangingfrom 45 to 65%). The gels were run in 1× TAE buffer at 80 V for17 h at a temperature of 60 °C and stained with SYBR® Gold (dilution1:10,000 in 1× Tris-acetate–EDTA—TAE—buffer) for 30 min in thedark. Visualization and digital pictures were performed using aChemiDoc System (BIORAD, Segrate, Italy).

2.12. Diversity indices calculation

Electrophoretic patterns, as well as band intensity, were used tocalculate the Shannon–Wiener H′ diversity index, the evennessindex EH, and the Simpson dominance index D by using Gel CompareII Software v4.6 (Applied Maths, Sint-Martens-Latem, Belgium).

For each lane H′=−SΣi=1 pi ln pi, where S is the total number ofbands and pi is the relative abundance of the i band calculated as the

Table 1Percentage of dead or alive plants (of all clones) in each line of the experimental field.

Line 1 2 3 4 5 6 7 8

a a b a a b a c

% dead 19.3 12.5 35.4 12.9 14.3 34.6 25 53.6% alive 80.6 87.5 64.5 87.1 85.7 65.4 75 46.4

Different letters indicate statistically significant distribution of dead–alive plants inlines, following a χ2 test (pb0.05).

265E. Gamalero et al. / Science of the Total Environment 425 (2012) 262–270

ratio between i band intensity and the sum of the intensities of all thebands in a lane; EH=H′/H′max where H′max=lnS; D=−SΣi=1 pi2.

2.13. Dendrogram construction and bacterial diversity determination

The similarity of the microbial populations was determinedthrough pair-wise comparisons of band presence/absence and inten-sity on the fingerprinting pattern of each soil sample by Gel CompareII software. A matrix of the similarity values was obtained using theDice Coefficient. This matrix was used to construct a dendrogram,according to the unweighted pair group method arithmetic average(UPGMA) cluster analysis, which also considers band intensity.

2.14. Sequencing of DGGE bands

Selected DGGE bands were excised from the gel, resuspended in20 μl of sterile TE (0.1 mM EDTA) and stored at −30 °C. The DNAextracted from the DGGE bands was re-amplified as previously de-scribed with the bacterial primers GC986f and Uni1401r and thePCR products were cloned into the pCR4-TOPO vector by using theTOPO TA cloning kit for sequencing according to the recommenda-tions of the manufacturer (Invitrogen, Milano, Italy), and chemicallycompetent Escherichia coli TOP 10 cells (Invitrogen, Milano, Italy)were transformed with the plasmid. The presence of cloned insertswas checked by PCR using the T3 (5′-ATTAACCCTCACTAAAGGGA-3′)and T7 (5′-TAATACGACTCACTATAGGG-3′) primers directly on thebacterial colonies diluted in water and lysed at 100 °C for 5 min. Thereactions were performed in a final volume of 25 μl containing 2.5 μlof 10× PCR buffer (Finnzymes, Keilaranta, Finland), 1.5 mM MgCl2,125 mM (each) dNTPs, 500 nM (each) primers, and 0.5 U of Taq poly-merase (Finnzymes). An aliquot (2 μl) of five diluted bacterial colo-nies was added to a 23 μl PCR mix. The reaction was performed in aTechne thermocycler (Bibby Scientific, Riozzo di Cerro al Lambro,Italy) with the following thermal protocol: 94 °C for 5 min, followedby 94 °C for 1 min, 56 °C for 1 min, and 72 °C for 1 min (30 cycles)and finally 10 min at 72 °C. For each DGGE bands three T3/T7 PCRfragments were sequenced.

PCR products were sequenced using the universal T3 and T7primers (see above) and the conditions for PCR mixture amplificationwere the same used for 16S rDNA sequencing (see the aboveIdentification of culturable bacteria section).

2.15. Statistical analyses

A χ2 test was used to compare the survival of the clones (dead vs.alive) in each line of the experimental field.

Since bacterial populations approximate an exponential normaldistribution, bacterial density values were logarithmically trans-formed before analysis. Data on enzymatic activities obtained fromfive replications were expressed as mean values. Non-transformedand transformed values were submitted to analysis of variance andto Fisher's least significant test (p≤0.05) using Statview 5.0 statisticspackage (SAS Institute Inc., Cary, USA).

Hierarchical clustering analysis, classifying a set of observationsdescribed by numerical variables in groups, was performed on soilmetabolic fingerprinting using Biodiversity Pro software (http://www.sams.ac.uk/dml/projects/benthic/bdpro/downloads.htm)based on the single linkage technique. The similarity among the siteswas calculated by the same software as Jaccard index (Ji=J/r 100,where J is the number of species found in both samples and r is thenumber of species found in one, or the other sample).

Electrophoretic patterns, as well as band intensity, were used tocalculate the Shannon–Wiener H′ diversity index, the evennessindex EH, and the Simpson dominance index D by using Gel CompareII software v4.6 (Applied Maths, Saint-Martens-Latem, Belgium).

3. Results

3.1. Performance and metal accumulation in poplar plants

According to the percent survival and average size of each poplarclone (including the spacer clone, I-214), Beaupré was the clonewith the highest survival percentage (92%), closely followed byAL35, Villafranca and NG12 (87%), while Jean Pourtet (57%), AL22and SN26 (both at 60%) were the worst performers. Intermediate sur-vival percentage values were measured for NG19 and I-214 (73%). Re-garding the average size of each clone, clones AL22 and AL35produced the largest plants (2.00 and 1.92, respectively), while thesmallest plants were those belonging to clones I-214 (0.97), JeanPourtet (1.13) and NG19 (1.18). Clones Baupré, NG12 and Villafrancashowed intermediate size classes (1.45, 1.54 and 1.62, respectively).

Because of the very different size shown by plants belonging to thesame clone and because some results contrast with previous observa-tions (Castiglione et al., 2009), survival percentages and size werealso analyzed by lines, comparing I-214 to the pool of all the otherclones (i.e. all clones except I-214). Considering all the plants together,dead plants weremore frequent within some lines and aχ2 test, con-sidering only dead vs. alive plants per line, indicated significant differ-ences comparing different lines. Furthermore, plotting the survival bylines of I-214 poplars and that of the pool of all the other clones (andthis was done in order to compare two populations of the same sizeand because each clone was not present in large numbers in all thelines), the patternswere very similar (Table 1), suggesting the existenceof a position effect. Therefore, six I-214 plants, three of large (I-214L)and three of small (I-214S) size, were chosen for further investigations.Chemical analysis of their root, stem (woody branches) and leaf sam-ples showed that Cu and Zn accumulation were higher in the largeplant (root metal concentration being almost three times that of thesmaller one, and significantly different—Table 2). Root samples weremicroscopically analyzed for the presence of mycorrhizal fungi, butnone was detected (data not shown).

3.2. Chemical and biochemical characterization of the soil

The soil moisture was lowest in the bulk soil (11.8%), while itincreased in the soil with plants (16.1% and 15.3% for I-214S and I-214L, respectively). Soil organic matter was always around 4% of thedry soil (4.06%, 4.64% and 3.97% for the bulk soil, I-214S and I214L, re-spectively), while TOC was 5.13% in the bulk soil and slightly lower inthe soil with poplars (4.39% for I-214S and 4.17% for I-214L). Thevalues for pH ranged from 7.77 (I-214L) to 7.97 (bulk soil) and 7.98(I-214S). While the concentration of total P was comparable in thethree soils, the amount of bioavailable P in the soil associated withthe poplar was higher than in the bulk soil (Table 3). Moreover, theconcentration of Cu in the three sites was higher than the guidelinevalues for the industrial area, while the amount of Zn was higherthan the guideline values established for the green area (Table 3).

Microbial activity, evaluated as cleavage of FDA with the concomi-tant release of fluorescein, was about three times higher in I-214Lthan in the bulk soil and I-214S. Phosphomonoesterases were foundin the three sites. Alkaline phosphomonoesterase was highest in I-214L and the acid ones in I-214L were about twice that measured in

Table 4Quantification of the enzymatic activities, metabolic versatility and the density of cul-turable bacteria in the three soils.

Bulk soil I-214S I-214L

FDA hydrolysis 1 7.98±0.42 a 6.54±0.33 a 21.57±0.95 bAlkalinephosphomonoesterase2

570.33±44.93 c 363.96±51.56 a 641.89±37.53 b

Neutralphosphomonoesterase2

0.00±0.00 a 135.17±10.21 b 107.74±8.34 b

Acidphosphomonoesterase3

5328.57±95.76 a

5476.70±102.86 a

9536.28±124.65 b

Urease4 21.32±1.22 c 6.21±0.44 b 0.00±0.00 aβ-Glucosidase3 0.00±0.00 a 11.00±0.93 b 0.00±0.00 aCellulase5 13.42±0.67 b 10.83±0.88 b 0.00±0.00 aMetabolic versatility6 5.21 31.25 46.87Culturable bacterialdensity7

6.86±0.17 a 6.93±0.35 ab 7.17±0.18 b

Values designated with the same letters are not significantly different (pb0.05)according to Fisher's LSD.

1 μg fluorescein g−1 of dry soil h−1.2 μg phenol g−1 of dry soil h−1.3 μg p-nitrophenol g−1 of dry soil h−1.4 μg ammonium g−1 of dry soil h−1.5 μg glucose g−1 of dry soil h−1.6 Percentage of positive wells of the Biolog GN microplate after 7 days of incubation.7 log CFU/g of dry soil.

Table 5Relative abundance of microbial species in each soil sample.

Bulk soil I-214 S I-214L

Arthrobacter niigatensis 0.21Arthrobacter oryzae 0.14 0.11 0.08Arthrobacter sp. 0.11 0.07 0.04Bacillus cereus 0.07 0.07 0.04Bacillus megaterium 0.04Bacillus mycoides 0.11 0.04Bacillus pumilus 0.04Bacillus simplex 0.04 0.04 0.08Bacillus sphaericus 0.04

Table 2Copper and zinc concentration (mg kg−1 DW) in roots, stems and leaves of large (I-214L)and small (I-214S) poplars.

Roots Stem Leaves

Cu Zn Cu Zn Cu Zn

I-214S 149±16 a 186±14 A 10±1 a 84±7 A 59±4 a 495±32 AI-214L 405±40 b 590±51B 21±2 b 106±9 A 91±8 b 645±48 B

Comparisons between each metal concentration in the same organ of large and smallplants resulting in statistically significant differences are indicated with differentletters (small letters for Cu, capital for Zn—pb0.05, N=5).

266 E. Gamalero et al. / Science of the Total Environment 425 (2012) 262–270

bulk soil and I-214S. Urease activity was three times higher in bulk soilthan in I-214S, while it was absent in I-214L. Cellulase and β-glucosidase, as enzymes involved in C cycling, were both found, al-though at low level, in I-214S. Finally, metabolic versatility, measuredas percentage of positive reaction obtained after inoculating Biolog GNmicroplates with soil extracts, followed an increasing gradient fromthe bulk soil to I-214S and I-214L (Table 4). More in detail, 11.6% ofthe reactions were exclusive of one site, while 23.1% of the reactionswere shared among the sites. Sites I-214S and I-214L, sharing 31.6% ofthe reactions, showed the highest similarity (62% of similarity). On thecontrary, only 8.4% of the reactions were shared between the bulk soiland I-214L (26% of similarity).

3.3. Quantification and identification of culturable bacteria

Density of culturable bacteria was comparable in bulk soil and I-214S. Nevertheless, bacterial density was higher in I-214L than inbulk soil (Table 4). A total of 100 bacterial strains were randomly se-lected, isolated, and characterized for their cellular and colony mor-phology (data not shown) and identified by 16S rDNA sequencing(Table 7, supplementary material). The culturable fraction differedaccording to the site. Five taxa (Arthrobacter oryzae, Arthrobactersp., Bacillus cereus, Bacillus simplex and Streptomyces) were commonto the sites. Two species (Chryseobacterium soldanellicola and Vario-vorax paradoxus) were preferentially associated with the soil wherepoplars were present, independently from the plant size. Finally,strains belonging to the genus Flavobacterium were prevalent in I-214L (Table 5).

The culturable fraction of the bulk soil was represented exclusive-ly by Gram positive bacteria; among them, the genera Arthrobacter,Bacillus, and Streptomyces were dominant and covered the 92.86% ofthe culturable bacteria identified in this site (Table 5). The abundanceof Gram positive strains in I214S (57.1%) and I214L (30.8%) was lessthan that of the bulk soil. The prevalent species in I-214S and I-214Lwas V. paradoxus (Table 5).

3.4. DGGE analysis of the culturable bacterial fraction

The DGGE profiles of bacterial culturable fractions were quite dif-ferent among the three sites (Fig. 1A). In particular, the soil I-214Lwas characterized by a completely different pattern compared to

Table 3Chemical and physical characterization of the three soil samples.

Bulk soil I-214S I-214L

Bioavailable phosphorus(% dry soil×10−4)

50±3 a 270±31 b 100±11 c

Total phosphorus (% dry soil×10−4) 570±48 a 650±55 a 560±53 aCopper (mg kg−1 of dry soil) ⁎ 886±85 a 996±98 a 1045±112 aZn (mg kg−1 of dry soil) ⁎⁎ 773±72 a 780±76 a 742±71 a

Different letters indicate statistically significant differences (pb0.05, N=5).⁎ The guideline values for the green and industrial area established by the Italian Law

(D. Lgs 152/06) are 120 and 600, respectively.⁎⁎ The guideline values for the green and industrial area established by the Italian Law(D. Lgs 152/06) are 150 and 1500, respectively.

the others and clustered alone (Fig. 1B). On the contrary, the bulksoil and I-214S showed 58% of similarity and clustered together(Fig. 1B). While the biodiversity indices were similar in bulk soiland in I-214S, the sample I-214L showed the highest number ofbands and value of Shannon–Wiener index (Table 6).

3.5. DGGE analysis of the total bacterial community

The DGGE analysis of the bacterial 16S rDNA (Fig. 1A) showedcharacteristic profiles for each sample with remarkable differencesin band position as well as band intensity among the samples.

In fact, the UPGMA analysis (Fig. 1B) indicated a low degree ofsimilarity between the three soils, with I-214S and I-214L being themost similar (54% of similarity) and the bulk soil clustering alone.

The number of bands in the DGGE patterns differed among thesoils. In detail, the sample I-214S showed the highest number ofbands and value of biodiversity indices. On the other hand, the bulk

Bacillus subtilis 0.04Bacillus sp. 0.04Chitinophaga ginsengisegetis 0.04Chryseobacterium soldanellicola 0.11 0.12Flavobacterium columnare 0.16Flavobacterium johnsoniae 0.04Flavobacterium sp. 0.16Paenibacillus lautus 0.04 0.07Paenibacillus sp. 0.04 0.04Streptomyces canus 0.04Streptomyces diastachromogenes subsp. Luteus 0.04Streptomyces lincolnensis 0.04Streptomyces globisporum 0.04Streptomyces sp. 0.07 0.04 0.04Variovorax paradoxus 0.25 0.24

Bulk soil culturable

I-214S culturable

I-214L culturable

Bulk soil

I-214S

I-214L

M 1 2 3 4 5 6 MA)

B)

12

3

4

5

678

910

11

121314

1516

1718

1920

21

2223

24

25

26

27

29

3031

dgge

30 35 40 45 50 55 60 65 70 75 80 85 90 95 100

Fig. 1. (A) DGGE banding pattern of the 16SrDNA from culturable and the total bacteria(M=marker; 1st lane bulk soil; 2nd lane bulk soil culturable; 3rd lane I-214S; 4th laneI-214S culturable; 5th lane I-214L; 6th lane I-214L culturable. B) The dendrogram of hier-archical cluster analysis was based on UPGMA and Dice's coefficient and generated by GelCompare II software.

267E. Gamalero et al. / Science of the Total Environment 425 (2012) 262–270

soil was characterized by a lower biodiversity (2.61 for the bulk soil,vs. 3.14 and 2.95 for I-214S and I-214L, respectively) and evenness(0.92 for the bulk soil, vs. 0.95 and 0.94 for I-214S and I-214L, respec-tively) compared to I-214S and I214L. In addition, the value of theSimpson's dominance index in the bulk soil was about twice thatmeasured in the soils associated with poplar (Table 6).

3.6. Sequence analysis

Overall, 31 bands have been excised from the DGGE of the wholebacterial communities. After re-amplification and examination fortheir uniqueness by DGGE, they were cloned and at least three clonesfor each band were sequenced. The putative phylogenetic affiliationof the sequences is reported in Table 9 (supplementary material).From this analysis, Acidobacteria were the most represented in thethree samples (20% in bulk soil, 23.3% in I214S and 48.5% in I214L).The frequency of bands corresponding to α- and γ-Proteobacteria inthe bulk soil (10% for both the taxonomic groups) and in the soil

Table 6Biodiversity indices of the different soils calculated from the DGGE banding pattern.

Site Numberof bands

BiodiversityShannon–Wiener

DominanceSimpson

EvennessPielou

Bulk soil 17 2.61 0.09 0.92Bulk soil culturable 11 2.12 0.14 0.88I-214S 27 3.14 0.05 0.95I-214S culturable 11 2.24 0.12 0.94I-214L 23 2.95 0.06 0.94I-214L culturable 18 2.53 0.10 0.88

associated with poplar (6.7% and 10% in I214S, respectively; 9% and12.1% in I214L, respectively) was similar, while that of β-Proteobacteria in I214S (23.3%) and I214L (18.2%) were twice thatmeasured in bulk soil (10%). On the contrary, bands correspondingto δ-Proteobacteria and Nitrospira were found only in the bulk soil, al-though with a low frequency (3.3% and 6.6%, respectively). Sequencesbelonging to the Actinobacteria were detected only in bulk soil (10%)and in I214S (16.7%), while Gemmatimonadetes were present in allthe samples, with a higher frequency in bulk soil (13.3%) than inI214S (3.3%) and I214L (3%).

4. Discussion

In general, all the clones selected during the previous survey con-firmed their good survival, with rates always exceeding 50% of thecuttings initially set in place. The clones AL22, AL35, NG12, SN26showed percentages of survival that are very close to those reportedby Castiglione et al. (2009); on the other hand, registered clones(Beaupré, Villafranca and Jean Pourtet) had much higher survival per-centages than in the previous experiments (Castiglione et al., 2009).In particular, Beaupré had the highest survival percentage (92%),while Jean Pourtet had the worst performance, with only 57% of theinitial cuttings still alive at the end of the first growing season. Nodata from previous experiments were available for the registeredclone I-214, which had an intermediate survival percentage (73%) incomparison with the other clones used in this study. Consideringthe average size of the various clones at the end of the first growingseason, again their performances were usually comparable to thosereported by Castiglione et al. (2009), with the exception of AL35and Villafranca on one side (they were larger in respect with the pre-vious field trial 2003) and NG19 on the other side, which presentedmuch smaller plants.

Visual inspection of the poplar field trial immediately suggestedthat the same clone was growing in a different way in the variouszones of the field according to the row position; moreover, their sur-vival percentage seemed to be particularly low in some patches of thefield. Therefore, we plotted the survival of the plants by lines, clusteringthem into two groups: I-214 and all the other clones, in order to havetwo populations of comparable size. In this way, we expected that anydifference that depended on some soil factors should lead to a similarsurvival percentage of the two populations and this was the case, asshown in Table 1. Given the broad genetic diversity of the Salicaceaefamily (Aravanopoulos et al., 1999; Smulders et al., 2008; Castiglioneet al., 2010), different responses of the poplar clones could be expectedas previously reported (Laureysens et al., 2004; Zalesny et al., 2005;Castiglione et al., 2009).

In general, chemical analyses of the soil failed to indicate any rele-vant difference in the composition and/or amount of nutrients, or pol-lutants. Phosphorus concentration, in fact, was quite constant in allcases, but there were differences in the amount of bio-available P: in-deed, the ratio between the available and total P was 2.0 and 4.7times higher in I-214L and I-214S, respectively, than that measured inthe bulk soil. This fact, which is consistent with the higher phospho-monoesterase activity found in I-214S and I-214L soils compared tothe bulk soil, suggested that the differential plant survivalmight dependupon the presence of some enzymatic activity. Hence, we collected soilsamples close to the trunk of some large and small I-214 poplars and inan areawhere cuttings had not been planted (bulk soil).We then inves-tigated the bacterial biodiversity by means of culture-dependent andindependent methods (Gamalero et al., 2003).

It is worth mentioning that, while the pattern of accumulation ofmetals in planta was consistent with previous reports, i.e. with mostCu retained in the roots (Kopponen et al., 2001; Borghi et al., 2007;Todeschini et al., 2007) and Zn transferred to the epigeous part andespecially to the leaves (Kopponen et al., 2001; Laureysens et al.,2004; Franchin et al., 2007; Unterbrunner et al., 2007; Lingua et al.,

268 E. Gamalero et al. / Science of the Total Environment 425 (2012) 262–270

2008; Todeschini et al., 2011), I-214L presented significantly higherconcentrations of the HMs than I-214S, as if their ability to tolerate(and accumulate) metals was increased. This fact is of special interestfor various reasons:

i) in the first place, it might be expected that plants with highermetal concentration in their organs should be smaller, as theywould have to deal with a larger amount of toxic substances;

ii) the fact that this did not happen would suggest that somemechanism prevented the higher metal concentration frombeing toxic;

iii) the observed differences in size did not depend on the plantgenome, as these plants were clones;

iv) the differences did not depend on the chemical features of thesoil;

v) if the factors inducing an increased tolerance to HMs could beisolated and manipulated, they could be especially useful forphytoremediation practices.

In addition, the concentration of Cu and Zn in the I-214S and I-214Lsoil samples was comparable, while their accumulation in the plant or-gans (root, stem and leaves) was higher in I-214L than in I-214S, indi-cating a higher bioavailability, or a greater extraction capacity. Thedensity of the culturable fraction of the microflora, its activity and itsmetabolic versatility tended to increase from the bulk soil to I-214L,clearly indicating the presence of a “rhizosphere effect” (Hartmann etal., 2008). This effect corresponded to greater microbial community ac-tivities in the rhizosphere compared to those in the bulk soil, due to theexudates released by roots, in spite of the high and homogeneous con-centration of HMs recorded in these soils. With the aim of selecting andidentifying culturable bacteria that are able to support plant growthunder this stressful condition, and therefore could be used as inoculantsin further phytoremediation studies, about 100 bacterial strains fromthe bulk soil and the I-214S and I-214L sample soils were isolated andidentified. Our results showed that the culturable fraction differedaccording to the field site where the soil was collected. In the bulksoil, in fact, only Gram positive bacteria were detected, represented byboth sporulating (Bacillus sp.) and non-sporulating (Arthrobacter sp.and Streptomyces sp.) species. The absolute dominance of Gram-positive bacteria in soil has been related to highly stressed and selectiveenvironmental conditions in which only bacteria able to produce resis-tance structures can survive (Fabiani et al., 2009). In agreement withother studies (Curl and Truelove, 1986; Marilley and Aragno, 1999;Söderberg et al., 2004), the abundance of Gram-negative strains washigher in soil associated with the plants than in the bulk soil. In fact,under conditions of sufficient supply of nutrients, Gram-negative bacteriashow high proliferation and activity rate, providing them a competitiveadvantage in the colonization of the rhizosphere. Interestingly, two spe-cies, C. soldanellicola and V. paradoxus, were preferentially associatedwith the poplar soils, with V. paradoxus being the prevalent species inI-214S and I-214L. C. soldanellicola was isolated from roots of sand-dune plants and described for the first time in 2006 (Park et al., 2006),but it was never reported to be associated with any other plant species,or as being tolerant to HMs. On the contrary, V. paradoxus, which wasisolated and described for the first time by Willems et al. (1991), wasshown to possess marked degradative capabilities against pollutants(Vallaeys et al., 1998; Smith et al., 2005; Nishino and Spain, 2006), toproduce NAHLs (N-Acyl-homoserine lactones), the signal molecules in-volved in cell-to-cell communication (Leadbetter and Greenberg,2000), and to promote plant growth under stressful conditions via thesynthesis of the enzyme ACC (1-aminocyclopropane-1-carboxylate)deaminase (Belimov et al., 2001; 2005). In addition, Flavobacteriumwas the prevalent genus in I-214L. Flavobacteria are spread in soil andrhizosphere where they play an important role in the biodegradationof recalcitrant polymers (Xu et al., 2010), tolerate high levels of majorpollutants, including HMs (Piotrowska-Seget et al., 2005; Kuffner etal., 2008), may oxidize and dehalogenate common organic pollutants

(Xun et al., 1992;Whiteley and Bailey, 2000) and promote plant growthunder natural and stressed conditions (Kuffner et al., 2008; He et al.,2010).

Culture-independent methods provided information regardingthe biodiversity of the whole and the culturable microflora, the latterbeing part of the whole microbial community. The cluster analysisperformed on the basis of the DGGE profiles highlighted a clear sepa-ration of the culturable fraction from the whole microbial community.According to the rDNA band profiles of the gel, the bulk soil patternsconsisted of few deep and several less intense bands, indicating thatin bulk soil only one or two bacterial populations dominated, whileseveral other populations, less prevalent, seemed to be equally abun-dant. In contrast, the I-214S and I-214L DDGE pattern consisted ofseveral intense bands and of a lower number of weak bands. This isin agreement with the highest dominance index calculated for thebulk soil, especially for the whole microflora.

The culturable fraction of the bacteria isolated from the I-214S soiland from the bulk soil, having the same number of bands and a similarShannon–Wiener index, showed the highest similarity, while the I-214Lsoil, characterized by the highest biodiversity, clustered separately. Onthe other hand, the band pattern of the whole microflora, accountingfor the quality of the microorganisms and not for their number alone,revealed a higher similarity between the I-214S and I-214L soils, withthe bulk soil clustering separately and showing the lowest biodiversityindex. The higher genetic biodiversity measured for the whole bacterialcommunity and the culturable fraction in the poplar-associated soilsamples, indicate that the poplar plants select the associated microflorathrough the release of root exudates. This is consistent with the obser-vation that some specific microbial groups and populations, that arebetter adapted than others to the rhizospheric environment, are prefer-entially associated with the roots (Mavingui et al., 1992; Lemanceau etal., 1995; Edel et al., 1997). Typically, distinct bacterial communities areassociated with different plant species (Latour et al., 1996; Grayston etal., 1998; Smalla et al., 2001) and even to single genotypes (VanOverbeek and Van Elsas, 2008; Micallef et al., 2009). However, sincethe band pattern of the I-214S and I-214L soils was similar but did notfully overlap, the size of the plant and the physiological stage could berelevant factors affecting the biodiversity of the associated microflora.In fact, the developmental stage of the plantmay be one of themain fac-tors affecting rhizobacterial community structure (Herschkovitz et al.,2005; Lerner et al., 2006; Mougel et al., 2006).

Band sequencing provided information about the whole bacterialcommunity composition, and relevant indications about the specieswhich were typically associated with the bulk soil or to the I-214soils. A high frequency of rDNA sequences corresponding to Acido-bacteria was found in the whole bacterial communities of all thethree collected soils. Acidobacteria are ubiquitous in bulk soils,where they represent the second most abundant phylum after theProteobacteria (Janssen, 2006), and in rhizosphere soils (Lee et al.,2008). Their high diversity and abundance suggest that this groupplays a key role for the soil functioning and rhizosphere ecosystemsand it may be linked to plant health and fitness.

Despite their high abundance in soil, an ecological understandingof this important bacterial phylum is limited because most of itsmembers cannot be maintained in culture and few molecular investi-gations have focused exclusively on this group. Nevertheless, it hasbeen observed that the abundance of Acidobacteria is negatively cor-related with soil carbon availability (Fierer et al., 2007) and stronglyregulated by soil pH, with the highest presence found in environ-ments with the lowest pH (Fierer et al., 2007; Lauber et al., 2008;Jones et al., 2009). In this framework, it should be noted that al-though at the sampling time the three soil samples showed an alka-line pH, measurements performed during the four years ofphytoremediation revealed a slightly acidic pH ranging from 5.0 to7.4, which could be favorable to the selection of bacteria belongingto this phylum.

269E. Gamalero et al. / Science of the Total Environment 425 (2012) 262–270

In addition, contrasting results regarding the sensitivity of Acido-bacteria to pollutants have been obtained. While Acidobacteria areconsidered to be sensitive to various pollutants and tend to disappearas a consequence of the contamination (El-Fantroussi et al., 1999;Saul et al., 2005; George et al., 2009), they were reported to representa significant portion of the bacterial strains in metal-contaminatedsoils, although they are rarely found in the active microbial com-munity (Gremion et al., 2003; Linton et al., 2007; Fabiani et al.,2009).

α- and γ-Proteobacteria were equally found in the three soils, in-dicating that these groups are not affected by the presence of theplant roots. On the other hand, β-Proteobacteria were more frequentin I-214S and I-214L soils suggesting that this group is typically asso-ciated with the poplar roots.

rDNA sequences belonging to the Actinobacteria (high G+C con-tent Firmicutes) were detected in bulk soil and in I-214S, while theywere absent in the I-214L soil. Although there is increasing evidencethat Gram-positive bacteria might be more dominant in the rhizo-sphere than previously expected (Normander and Prosser, 2000;Smalla et al., 2001; Duineveld et al., 2001), no other sequences ofGram-positive bacteria were found in all three collected soils.

5. Conclusions

This study confirms once more the importance of undertakingfield studies. In spite of the problems and limitations arising fromthis kind of approach, some features of the field soil, like the char-acteristics of the largely diverse microbial communities, cannotfully be understood by means of experiments under controlledconditions.

The analysis of the whole microbial community provided the de-scription of the differences between the bulk soil and the soilswhere poplars were grown (and indeed, the DGGE results indicatedthat I-214S and I-214L clustered together). At the same time, theanalysis of the culturable fraction, with its higher resolution in theidentification of the microbial taxa, allowed the identification ofsome strains that were associated with the poplars. C. soldanellicolaand V. paradoxus were associated with the poplar, irrespective ofthe plant size, while the genus Flavobacterium was dominant in theI-214L soil. The characterization of the active fraction of the wholebacterial community is in progress. Moreover, the assessment of theeffects induced by bacterial strains identified as Flavobacterium spp.,C. soldanellicola and V. paradoxus on poplar plants cultivated in Cuor Zn polluted soil, will provide further insight into the role of thesemicrobial species in promoting plant growth under stressfulconditions.

Acknowledgments

The authors wish to thank M. Sobrero, D. Vigani, D. Romiglio andA. Peterson for technical help during plant growth and experiments,and management and staff of KME Italy (especially M. Sterpi andB. Ciarlo) for assistance.

The research was financed by Associazione Ambiente-Territorio-Formazione to GL, and by the Italian Ministry for Education, Universi-ty and Research (PRIN 2005055337) to SC and GL.

The Authors thank Prof. Elizabeth Illingworth for the revision ofEnglish.

Appendix A. Supplementary data

Supplementary data to this article can be found online at doi:10.1016/j.scitotenv.2012.03.012.

References

Abou-Shanab RA, Ghanem K, Ghanem N, Al-Kolaibe A. The role of bacteria on heavy-metal extraction and uptake by plants growing on multi-metalcontaminatedsoils. World J Microbiol Biotechnol 2008;24:253–62.

Adam G, Duncan H. Development of a sensitive and rapid method for the measurementof total microbial activity using fluorescein diacetate (FDA) in a range of soils. SoilBiol Biochem 2001;33:943–51.

Alef K, Nannipieri P. Enzyme activities. In: Alef K, Nannipieri P, editors. Methods in ap-plied soil microbiology and biochemistry. London: Academic Press; 1995.p. 311–73.

Aravanopoulos FA, Kim KH, Zsuffa L. Genetic diversity of superior Salix clones selectedfor intensive forestry plantations. Biomass Bioen 1999;16:249–55.

Ausubel FM, Brent R, Kingstone RE, Moore DD, Seidman JG, Smith JA, et al. Current pro-tocols in molecular biology, section 2.4.2. New York, N.Y: Wiley; 1987.

Baldantoni D, Bellino A, Cicatelli A, Castiglione S. Artificial mycorrhization does not in-fluence the effects of iron availability on Fe, Zn, Cu, Pb and Cd accumulation inleaves of a heavy metal tolerant white poplar clone. Plant Biosys 2011;145:236–40.

Belimov AA, Safronova VI, Sergeyeva TA, Egorova TN, Matveyeva VA, Tsyganov VE, et al.Characterization of plant growth-promoting rhizobacteria isolated from pollutedsoils and containing 1-aminocyclopropane-1-carboxylate deaminase. Can J Micro-biol 2001;47:642–52.

Belimov AA, Hontzeas N, Safronova VI, Demchinskaya SV, Piluzza G, Bullitta S, et al.Cadmium-tolerant plant growth-promoting bacteria associated with the roots ofIndian mustard (Brassica juncea L. Czern.). Soil Biol Biochem 2005;37:241–50.

Borghi M, Tognetti R, Monteforti G, Sebastiani L. Responses of Populus × euramericana(P. deltoides × P. nigra) clone Adda to increasing copper concentrations. EnvironExp Bot 2007;61:66–73.

Bremner JM, Mulvaney RL. Urease activity in soils. In: Burns RG, editor. Soil enzymes.New York: Academic Press; 1978. p. 150–96.

Castiglione S, Todeschini V, Franchin C, Torrigiani P, Gastaldi D, Cicatelli A, et al. Clonaldifferences in survival capacity, copper and zinc accumulation, and correlationwith leaf polyamine levels in poplar: a large-scale field trial on heavily pollutedsoil. Environ Pollut 2009;157:2108–17.

Castiglione S, Cicatelli A, Lupi R, Patrignani G, Fossati T, Brundu G, et al. Genetic struc-ture and introgression in riparian populations of Populus alba L. Plant Biosys2010;144:656–68.

Curl EA, Truelove B. The rhizosphere. Berlin: Springer; 1986.Di Baccio D, Tognetti R, Sebastiani L, Vitagliano C. Responses of Populus deltoides ×

Populus nigra (Populus × euramericana) clone I-214 to high zinc concentrations.New Phytol 2003;159:443–52.

Dos Santos Utmazian MN, Wieshammer G, Vega R, Wenzel WW. Hydroponic screeningfor metal resistance and accumulation of cadmium and zinc in twenty clones ofwillows and poplars. Environ Pollut 2007;148:155–65.

Doty SL, Dosher MR, Singleton GL, Moore AL, Van Aken B, Stettler RF, et al. Identifica-tion of an endophytic Rhizobium in stems of Populus. Symbiosis 2005;39:27–35.

Duineveld BM, Kowalchuk GA, Keijzer A, van Elsas JD, van Veen JA. Analysis of bacterialcommunities in the rhizosphere of chrysanthemum via denaturing gradient gelelectrophoresis of PCR-amplified 16S rRNA as well as DNA fragments coding for16S rRNA. Appl Environ Microbiol 2001;67:172–8.

Edel V, Steinberg C, Gautheron N, Alabouvette C. Populations of nonpathogenic Fusariumoxysporum associated with roots of four plant species compared to soilborne popula-tions. Phytopathol 1997;87:693–7.

Eivazi F, Tabatabai MA. Phosphatases in soils. Soil Biol Biochem 1977;9:167–72.Eivazi F, Tabatabai MA. Glucosidases and galactosidases in soil. Soil Biol Biochem

1988;20:601–6.El-Fantroussi S, Verschuere L, Verstraete W, Top EM. Effect of phenylurea herbicides on

soil microbial communities estimated by analysis of 16S rRNA gene fingerprintsand community-level physiological profiles. Appl Environ Microbiol 1999;65:982–8.

Fabiani A, Gamalero E, Castaldini M, Cossa GP, Musso C, Pagliai M, et al. Microbiologicalpolyphasic approach for soil health evaluation in an Italian polluted site. Sci Tot En-viron 2009;407:4954–64.

Felske A, Wolterink A, Van Lis R, Akkermans ADL. Phylogeny of the main bacterial16SrDNA sequences in Drentse A grassland soils (The Netherlands). Appl EnvironMicrobiol 1998;64:871–9.

Fierer N, Bradford M, Jackson R. Toward an ecological classification of soil bacteria.Ecology 2007;88:1354–64.

Franchin C, Fossati T, Pasquini E, Lingua G, Castiglione S, Torrigiani P, et al. High con-centrations of Zn and Cu induce differential polyamine responses in micropropa-gated white poplar (Populus alba L. ‘Villafranca’). Physiol Plant 2007;130:77–90.

Gamalero E, Lingua G, Berta G, Lemanceau P. Methods for studying root colonization byintroduced beneficial bacteria. Agronomie 2003;23:407–18.

Gamalero E, Berta G, Lingua G, Glick BR. Effects of plant growth-promoting bacteria andAM fungi on the response of plants to heavy metal stress. Can J Microbiol 2009;55:501–14.

Gentry TJ, Wang G, Rensing C, Pepper IL. Chlorobenzoate-degrading bacteria in similarpristine soils exhibit different community structures and population dynamics inresponse to anthropogenic 2-, 3-, and 4-chlorobenzoate levels. Microbial Ecol2004;48:90-102.

George IF, Liles MR, HartmannM, LudwigW, Goodman RM, Agathos SN. Changes in soilAcidobacteria communities after 2,4,6-trinitrotoluene contamination. FEMS Micro-biol Lett 2009;296:159–66.

Gottel NR, Castro HF, Kerley M, Yang Z, Pelletier DA, Podar M, et al. Distinct microbialcommunities within the endosphere and rhizosphere of Populus deltoides rootsacross contrasting soil types. Appl Environ Microbiol 2011;77:5934–44.

270 E. Gamalero et al. / Science of the Total Environment 425 (2012) 262–270

Grayston SJ, Wang S, Campbell CD, Edwards AC. Selective influence of plant species onmicrobial diversity in the rhizosphere. Soil Biol Biochem 1998;30:369–78.

Gremion F, Chatzinotas A, Harms H. Comparative 16 S rDNA and 16 S rRNA sequenceanalysis indicates that Actinobacteria might be a dominant part of the metabolical-ly active bacteria in heavy metal-contaminated bulk and rhizosphere soil. EnvironMicrobiol 2003;5:896–907.

Hartmann A, Rothballer M, Schmid M, Hiltner Lorenz. a pioneer in rhizosphere micro-bial ecology and soil bacteriology research. Plant Soil 2008;312:7-14.

He CQ, Tan GE, Liang X, Du W, Chen YL, Zhi GY, et al. Effect of Zn-tolerant bacterialstrains on growth and Zn accumulation in Orychophragmus violaceus. Appl SoilEcol 2010;44:1–5.

Herschkovitz Y, Lerner A, Davidov Y, Rothballer M, Hartmann A, Okon Y, et al. Inocula-tion with the plant growth promoting rhizobacterium Azospirillum brasilensecauses little disturbance in the rhizosphere and rhizoplane of maize (Zea mays).Microb Ecol 2005;50:277–88.

Janssen PH. Identifying the dominant soil bacterial taxa in libraries of 16S rRNA and16S rRNA genes. Appl Environ Microbiol 2006;72:1719–28.

Jones RT, Robeson MS, Lauber CL, Hamady M, Knight R, Fierer N. A comprehensive sur-vey of soil acidobacterial diversity using pyrosequencing and clone library analy-ses. ISME J 2009;3:442–53.

Kopponen P, Utriainen M, Lukkari K, Suntioinen S, Kärenlampi L, Kärenlampi S. Clonaldifferences in copper and zinc tolerance of birch in metal-supplemented soils. En-viron Poll 2001;112:89–97.

Kuffner M, Puschenreiter M, Wieshammer G, Gorfer M, Sessitsch A. Rhizosphere bacte-ria affect growth and metal uptake of heavy metal accumulating willows. Plant Soil2008;304:35–44.

Latour X, Corberand T, Laguerre G, Allard F, Lemanceau P. The composition of fluores-cent pseudomonad population associated with roots is influenced by plant and soiltype. Appl Environ Microbiol 1996;62:2449–556.

Lauber CL, Strickland MS, Bradford MA, Fierer N. The influence of soil properties on thestructure of bacterial and fungal communities across land-use types. Soil Biol Bio-chem 2008;40:2407–15.

Laureysens I, Blust R, De Temmerman L, Lemmens C, Ceulemans R. Clonal variation in heavymetal accumulation and biomass production in poplar coppice culture: I. Seasonalvariation in leaf, wood and bark concentrations. Environ Pollut 2004;131:485–94.

Laureysens I, De Temmerman L, Hastir T, Van Gysel M, Ceulemans R. Clonal variation inheavy metal accumulation and biomass production in a poplar coppice culture: II.Vertical distribution and phytoextraction potential. Environ Pollut 2005;133:541–51.

Leadbetter JR, Greenberg EP. Metabolism of acyl-homoserine lactone quorum-sensingsignals by Variovorax paradoxus. J Bacteriol 2000;182:6921–6.

Lee S-H, Ka J-O, Cho J-E. Members of the phylum Acidobacteria are dominant and meta-bolically active in rhizosphere soil. FEMS Microbiol Lett 2008;285:263–9.

Lemanceau P, Corberand T, Gardan L, Latour X, Laguerre G, Boeufgras J-M, et al. Effect oftwo plant species, flax (Linum usitatissinum L.) and tomato (Lycopersicon esculentumMill.), on the diversity of soilborne populations of fluorescent pseudomonads. ApplEnviron Microbiol 1995;61:1004–12.

Lerner A, Herschkovitz Y, Baudoin E, Nazaret S, Moënne-Loccoz Y, Okon Y, et al. Effectof Azospirillum brasilense on rhizobacterial communities analyzed by denaturinggradient gel electrophoresis and automated intergenic spacer analysis. Soil BiolBiochem 2006;38:1212–8.

Lingua G, Franchin C, Todeschini V, Castiglione S, Biondi S, Burlando B, et al. Arbuscularmycorrhizal fungi differentially affect the response to high zinc concentrations oftwo registered poplar clones. Environ Pollut 2008;153:137–47.

Linton PE, Shotbolt L, Thomas AD. Microbial communities in long-term heavy-metalcontaminated ombrotrophic peats. Water Air Soil Poll 2007;186:97-113.

Marilley L, Aragno M. Phylogenetic diversity of bacterial communities differing in de-gree of proximity of Lolium perenne and Trifolium repens roots. Appl Soil Ecol1999;13:127–36.

Mavingui P, Laguerre G, Berge O, Heulin T. Genetic and phenotypic diversity of Bacilluspolymyxa in soil and in the wheat rhizosphere. Appl Environ Microbiol 1992;58:1894–903.

Micallef SA, Shiaris MP, Colon-Carmona A. Influence of Arabidopsis thaliana accessionson rhizobacterial communities and natural variation in root exudates. J Exp Bot2009;60:1729–42.

Moore FP, Barac T, Borrernans B, Oeyen L, Vangronsveld J, van der Lelie D, et al. Endo-phytic bacterial diversity in poplar trees growing on a BTEX-contaminated site: thecharacterisation of isolates with potential to enhance phytoremediation. Syst ApplMicrobiol 2006;29:539–56.

Mougel C, Offre P, Ranjard L, Corberand T, Gamalero E, Robin C, et al. Dynamic of the ge-netic structure of bacterial and fungal communities at different development stages ofMedicago truncatula Geartn. Cv. Jemalong line J5. New Phytol 2006;170:165–75.

Nishino SF, Spain JC. Biodegradation of 3-nitrotyrosine by Burkholderia sp. strain JS165and Variovorax paradoxus JS171. Appl Environ Microbiol 2006;72:1040–4.

Normander B, Prosser JI. Bacterial origin and community composition in the barleyphytosphere as a function of habitat and pre-sowing conditions. Appl EnvironMicrobiol 2000;66:4372–7.

Park MS, Jung SR, Lee KH, Lee M-S, Do JO, Kim SB, et al. Chryseobacterium soldanellicolasp. nov. and Chryseobacterium taeanense sp. nov., isolated from roots of sand-duneplants. Int J Syst Evol Microbiol 2006;56:433–8.

Pilon Smits EAH. Phytoremediation. Ann Rev Plant Biol 2005;56:15–39.

Piotrowska-Seget Z, Cycon´ M, Kozdro´j J. Metal-tolerant bacteria occurring in heavilypolluted soil and mine spoil. Appl Soil Ecol 2005;28:237–46.

Ravera M, Ciccarelli C, Gastaldi D, Rinaudo C, Castelli C, Osella D. An experiment in theelectrokinetic removal of copper from soil contaminated by the brass industry.Chemosphere 2006;63:950–5.

Saul DJ, Aislabie JM, Brown CE, Harris L, Foght JM. Hydrocarbon contamination changesthe bacterial diversity of soil from around Scott Base, Antarctica. FEMS MicrobiolEcol 2005;53:141–55.

Schinner F, von Mersi W. Xylanase, CM-cellulase and invertase activity in soil: an im-proved method. Soil Biol Biochem 1990;22:511–5.

Sebastiani L, Scebba F, Tognetti R. Heavy metal accumulation and growth responses inpoplar clones Eridano (Populus deltoides × maximowiczii) and I-214 (P. × eurameri-cana) exposed to industrial waste. Environ Exp Bot 2004;52:79–88.

Shilev S, Ruso J, Puig M, Benlloch M, Sancho ED. Rhizospheric bacteria promote sun-flower (Helianthus annuus L.) plant growth and tolerance to heavy metals. MinervaBiotec 2001;13:37–9.

Smalla K, Wieland G, Buchner A, Zock A, Parzy J, Kaiser S, et al. Bulk and rhizospheresoil bacterial communities studied by denaturing gradient gel electrophoresis:plant-dependent enrichment and seasonal shifts revealed. Appl Environ Microbiol2001;67:4742–51.

Smith D, Alvey S, Crowley DE. Cooperative catabolic pathways within an atrazine-degrading enrichment culture isolated from soil. FEMS Microbiol Ecol 2005;53:265–73.

Smulders MJM, Cottrell JE, Lefevre F, van der Schoot J, Arens P, Vosman B, et al. Struc-ture of the genetic diversity in black poplar (Populus nigra L.) populations acrossEuropean river systems: consequences for conservation and restoration. For EcolManag 2008;255:1388–99.

Söderberg KH, Probanza A, Jumpponen A, Bååth E. The microbial community in the rhi-zosphere determined by community-level physiological profiles (CLPP) and directsoil- and cfu-PLFA techniques. Appl Soil Ecol 2004;25:135–45.

Solìs-Domìnguez FA, Valentìn-Vargas A, Chorover J, Maier RM. Effect of arbuscular my-corrhizal fungi on plant biomass and the rhizosphere microbial community struc-ture of mesquite grown in acidic lead/zinc mine tailings. Sci Total Environ2011;409:1009–16.

Soudek P, Tykva R, Vanĕk T. Laboratory analyses of 137Cs uptake by sunflower, reedand poplar. Chemosphere 2004;55:1081–7.

Todeschini V, Franchin C, Castiglione S, Burlando B, Biondi S, Torrigiani P, et al. Responsesof two registered poplar clones to copper, after inoculation, or not, with arbuscularmycorrhizal fungi. Caryologia 2007;60:146–55.

Todeschini V, Lingua G, D'Agostino G, Carniato F, Roccotiello E, Berta G. Effects of highzinc concentration on poplar leaves: a morphological and biochemical study. Envi-ron Exp Bot 2011;71:50–6.

Ulrich K, Ulrich A, Ewald D. Diversity of endophytic bacterial communities in poplargrown under field conditions. FEMS Microbiol Ecol 2008;63:169–80.

Unterbrunner R, Puschenreiter M, Sommer P, Wieshammer G, Tlustos P, Zupan M, et al.Heavy metal accumulation in trees growing on contaminated sites in Central Eu-rope. Environ Pollut 2007;148:107–14.

Vallaeys T, Albino L, Soulas G, Wright AD, Weightman AJ. Isolation and characterizationof a stable 2,4-dichlorophenoxyacetic acid degrading bacterium, Variovorax para-doxus, using chemostat culture. Biotechnol Lett 1998;20:1073–6.

Van Aken B, Peres CM, Doty SL, Yoon JM, Schnoor JL.Methylobacterium populi sp. nov., anovel aerobic, pink-pigmented facultatively methylotrophic, methane-utilizingbacterium isolated from poplar trees (Populus deltoides_nigra DN34). Int J SystEvol Microbiol 2004;54:1191–6.

van der Lelie D, Taghavi S, Monchy S, Schwender J, Miller L, Ferrieri R, et al. Poplar andits bacterial endophytes: coexistence and harmony. Crit. Rev. Plant Sci. 2009;28:346–58.

Van Overbeek L, Van Elsas JD. Effects of plant genotype and growth stage on the struc-ture of bacterial communities associated with potato (Solanum tuberosum L.).FEMS Microbiol Ecol 2008;64:283–96.

Whiteley AS, Bailey MJ. Bacterial community structure and physiological state withinan industrial phenol bioremediation system. Appl Environ Microbiol 2000;66:2400–7.

Willems A, De Ley J, Gillis M, Kersters K. Comamonadaceae, a new family encompassingthe acidovorans rRNA complex, including Variovorax paradoxus gen. nov., comb. nov.,for Alcaligenes paradoxus (Davis 1969). Int J Syst Bacteriol 1991;41:445–50.

Wilson KH, Blitchington RB, Greene RC. Amplification of bacterial 16S ribosomal DNAwith polymerase chain reaction. J Clin Microbiol 1990;28:1942–6.

Xu L, Teng Y, Li Z-G, Norton JM, Luo YM. Enhanced removal of polychlorinated biphe-nyls from alfalfa rhizosphere soil in a field study: the impact of a rhizobial inocu-lum. Sci Tot Environ 2010;408:1007–13.

Xun LY, Topp E, Orser CS. Confirmation of oxidative dehalogenation of pentachlorophe-nol by a Flavobacterium pentachlorophenol hydroxylase. J Bacteriol 1992;174:5745–7.

Zalesny Jr RS, Bauer EO, Hall RB, Zalesny JA, Kunzman J, Rog CJ, et al. Clonal variation insurvival and growth of hybrid poplar and willow in an in situ trial on soils heavilycontaminated with petroleum hydrocarbons. Int J Phytorem 2005;7:177–97.