Inoculating Helianthus annuus (sunflower) grown in zinc and cadmium contaminated soils with plant growth promoting bacteria – Effects on phytoremediation strategies

Chemosphere 92 (2013) 74–83

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Inoculating Helianthus annuus (sunflower) grown in zinc and cadmium contaminated soils with plant growth promoting bacteria – Effects on phytoremediation strategies Ana P.G.C. Marques, Helena Moreira, Albina R. Franco, António O.S.S. Rangel, Paula M.L. Castro ⇑ CBQF – Centro de Biotecnologia e Química Fina - Laboratório Associado, Escola Superior de Biotecnologia, Universidade Católica Portuguesa/Porto, Rua Dr. António Bernardino de Almeida, 4200-072 Porto, Portugal

h i g h l i g h t s  The tested strains reduced sunflower biomass losses in metal contaminated soils.  Bacteria decreased up to 67% and 64% Zn and Cd accumulation in sunflower tissues.  Bacterial community diversity decreased with increasing metal pollution in soils.  Inoculation with bacteria decreased the reduction of bacterial diversity up to 83%.  Sunflower inoculation is effective in enhancing its phytoremediation potential.

a r t i c l e

i n f o

Article history: Received 22 October 2012 Received in revised form 1 February 2013 Accepted 17 February 2013 Available online 9 April 2013 Keywords: Phytoremediation Helianthus annuus PGPR Zn Cd Microbial dynamics

a b s t r a c t Plant growth promoting bacteria (PGPR) may help reducing the toxicity of heavy metals to plants in polluted environments. In this work the effects of inoculating metal resistant and plant growth promoting bacterial strains on the growth of Helianthus annuus grown in Zn and Cd spiked soils were assessed. The PGPR strains Ralstonia eutropha (B1) and Chrysiobacterium humi (B2) reduced losses of weight in metal exposed plants and induced changes in metal bioaccumulation and bioconcentration – with strain B2 decreasing up to 67% Zn accumulation and by 20% Zn bioconcentration factor (BCF) in the shoots, up to 64% Zn uptake and 38% Zn BCF in the roots, and up to 27% Cd uptake and 27% Cd BCF in plant roots. The impact of inoculation on the bacterial communities in the rhizosphere of the plant was also assessed. Bacterial community diversity decreased with increasing levels of metal contamination in the soil, but in rhizosphere soil of plants inoculated with the PGPR strains, a higher bacterial diversity was kept throughout the experimental period. Inoculation of sunflower, particularly with C. humi (B2), appears to be an effective way of enhancing the short term stabilization potential of the plant in metal contaminated land, lowering losses in plant biomass and decreasing aboveground tissue contamination. Ó 2013 Elsevier Ltd. All rights reserved.

1. Introduction Pollution by heavy metals (HMs) due to anthropogenic activities is extensive (Barceloux, 1999). Cadmium is an element with no known biological function and one of the most toxic heavy metals (Courbot et al., 2004). Zinc, although an essential trace element, is toxic at high levels (Hao et al., 2012). Soil contamination with these HM is a widespread problem and causes a considerable threat to the environment. Metal polluted land is not adequate

⇑ Corresponding author. Tel.: +351 22 558 00 59; fax: +351 22 509 03 51. E-mail addresses: [email protected] (A.P.G.C. Marques), hmoreira@ porto.ucp.pt (H. Moreira), [email protected] (A.R. Franco), arangel@porto. ucp.pt (A.O.S.S. Rangel), [email protected] (P.M.L. Castro). 0045-6535/$ - see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.chemosphere.2013.02.055

for food or feed crops cultivation and such soils require remediation to reduce hazardous risk. The application of ecological remediation methods, such as phytoremediation appears as an excellent low cost option. The use of metal tolerant or accumulating species that are good energy crops in such process is becoming a very promising opportunity as renewable energy source, avoiding the utilization of farmland for the production of such non-edible biomass (Mlezeck et al., 2010). Helianthus annuus (sunflower), one of the most important crops worldwide, is a plant not only with food and energy value, but also with phytoremediation potential. Sunflower is a documented metal accumulator (Cindy et al., 2006; Niu et al., 2007; Fassler et al., 2010; Rojas-Tapias et al., 2012) and its growth on contaminated land for simultaneous remediation and further energy production has been studied (Madejon et al., 2003).

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Heavy metal contaminated soils are not only a source of important tolerant or even metal accumulator plant species, but also of metal-tolerant microorganisms (Batty, 2005). Amongst these organisms, plant growth promoting rhizobacteria (PGPR) are bacteria colonizing the plant root capable of promoting plant growth by various mechanisms – namely through the fixation of atmospheric nitrogen, utilization of 1-aminocyclopropane-1-carboxylic acid (ACC) as a sole N source, production of siderophores and antipathogenic substances, production of plant growth regulators (phytohormones, such as auxins), and also through the transformation of nutrient elements (Glick et al., 1999). Several studies suggest that phytoremediation can be more effective after inoculation of selected plant species with PGPR, with these enhancing plant growth and survival in metal contaminated soils (Wu et al., 2006; Grandlic et al., 2008), with a reduction in the requirement for amendments and its associated costs (BecerraCastro et al., 2012). Some of the mechanisms suggested in the amelioration of HM toxicity in plants by PGPR may be the reduction in metal uptake by the plant (Vivas et al., 2006) or in the amounts of detrimental ethylene induced stress (Rajkumar et al., 2006). Bacteria associated with plant roots can thus be considered as an important component of the phytoremediation technology (Glick, 2003). The aims of this work were to examine the effects of two metal resistant and PGPR bacterial strains on plant growth and on Zn and Cd uptake and accumulation by H. annuus exposed to different metal levels, and to assess the impact of the contamination and the selected strains on the bacterial communities in the rhizosphere of the plant.

Luria–Bertani broth (MLB) (Tiago et al., 2004), supplemented with 15% (v/v) glycerol, confirmed by plating the cultures into tryptic soy agar (TSA).

2.2. Preparation of the substrate The soil used in this study was an agricultural soil from the North of Portugal (soil properties are shown in Table 1). Soil was collected randomly in the selected area, to a 20 cm depth. The soil was milled to <2 mm, sterilized at 120 °C for 70 min in two consecutive days and dried in an oven at 40 °C for 4 d. The soil was then amended with Cd or Zn as CdCl2 and ZnCl2 respectively. Solutions of the salt were prepared with deionized water and applied to the soil to obtain concentrations of 0, 10, 20, and 30 mg Cd kg 1 and 0, 100, 500, and 1000 mg Zn kg 1 dry soil. The treated soils were wetted for 1 week by adding deionized water to maintain 60% of the water holding capacity; the soil was then dried in the greenhouse for approximately 2 weeks. This spiked soil was subjected to three cycles of wet and dry processes, favoring the distribution of the metals throughout the entire soil mass and allowing the soil to reach equilibrium between available and organic matter absorbed fractions (Blaylock et al., 1997). The exchangeable – ethylenediaminetetraacetic acid (EDTA) extractable, and available – ammonium acetate (NH4-Ac) extractable – Cd and Zn fractions were determined for the soils at the end of this process (Thomas, 1982).

2.3. Experimental design 2. Materials and methods 2.1. Bacteria isolation and characterization Selected bacterial species were indigenous to a metal contaminated site. The site has a long history of metal contamination, due to the industrial activity in the surrounding area (Marques et al., 2007) but is prolific in vegetation. Bacterial isolation was performed from sediments collected at the site. The strains selected for the present work were identified as B1 (Ralstonia eutropha) and B2 (Chryseobacterium humi) (Pires, 2010; Pires et al., 2011). These strains, identified through 16S rRNA, showed tolerance to Cd and Zn up to levels of 500 mg L 1 in liquid culture (Pires, 2010), revealed in vitro plant growth promoting traits, and were reported to promote plant growth in vivo (Marques et al., 2010) and were therefore selected to carry the present study. They were maintained as pure cultures in modified

The experiment consisted of a factorial design with two metals (Cd and Zn), four matrix Cd levels (0 – control non-spiked soil – 10, 20, and 30 mg Cd kg 1), three matrix Zn levels (0 – control nonspiked soil – 100, 500, and 1000 mg Zn kg 1) and three bacterial treatments (B0 – no bacteria, B1 and B2). Each treatment was replicated four times. Helianthus annuus seeds (variety IBERICO, purchased from LusoSem, Portugal) were surface sterilized with 0.5% (v/v) NaOCl for 10 min and were subsequently washed with sterilized deionized water. Seeds were germinated in plastic pots with about 300 g sterilized of the agricultural soil. Each pot received 10 seeds. Pots were randomized on the greenhouse, process that was repeated every 2 weeks during the experiment. After sowing, seedlings were reduced to three per pot. The pots were inoculated with 10 mL of a solution of each bacterial strain (108 CFU mL 1) (Vivas et al., 2006) pre-grown in nutrient broth medium – tryptic soy broth (TSB) – for

Table 1 Soil properties. Total As (mg kg 1) Extractable B (mg kg 1) Extractable Ca (mg kg 1) Total Cd (mg kg 1) Total Co (mg kg 1) Total Cr (mg kg 1) Total Cu (mg kg 1) Extractable Fe (mg kg 1) Total Hg (mg kg 1) Extractable K (mg kg 1) Extractable Mg (mg kg 1) Extractable Mn (mg kg 1) Total N (mg kg 1) Total Ni (mg kg 1) Total P (mg kg 1) Total Pb (mg kg 1) Total Zn (mg kg 1)

<5 (L.D.) (aqua regia) 0.32 ± 0.09 (diantrimide) 1629 ± 33(ammonium acetate) <1.8 (L.D.) (aqua regia) 15 ± 3 (aqua regia) 14 ± 2 (aqua regia) 33 ± 2 (aqua regia) 130 ± 9 (Lakanen-EAA) <0.01 (L.D.) (aqua regia) 106 ± 124 (Egner-Rhien) 109 ± 6 (ammonium acetate) 55 ± 2 (Lakanen-EAA) 1736 ± 50 (Kjedhal) 21 ± 7 (aqua regia) 2600 ± 23 (colorimetric-ascorbic acid) 158 ± 12 (aqua regia) 32 ± 4 (aqua regia)

pH

6.71 ± 0.08 (potenciometric)

Organic content (%) CEC (cmol kg

1

)

3.1 ± 0.2 (Walkey–Black) 13.6 ± 0.5 (Mehlich)

Texture: loam Clay Sand

(hydrometer) 16 ± 5 9.7 ± 0.8 76 ± 8

Water content (%)

4.84 ± 0.03 (gravimetric)

Values are shown as average ± standard deviation (n = 3). Methods used in each case are shown parenthetically.

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24–48 h at 28 °C (B1 and B2). Ten millilitre of sterile nutrient TSB medium (B0) was also added to the control treatment pots. The plants were maintained in a controlled growth room (12 h photoperiod, 450 lmol m 2 s 1 photosynthetically active radiation, 18–38 °C temperature range, 16–71% relative humidity range), and were watered daily. Harvest occurred 20 weeks after the beginning of the experiment. 2.4. Plant analysis Entire plants were washed with tap water, followed by washing with HCl 0.1 M, and with de-mineralized water, separated in roots and shoots, after which root elongation and shoot length were registered. The biomass of the plants was determined after shoots and roots were oven dried at 70 °C for 48 h. Plants were then grinded and sieved to <1 mm and shoot and root plant samples were digested at high temperature in a PerkinElmer MicroWave using method 3052 USEPA. Cadmium and Zn content was determined using FA-AAS of the digests (Wallinga et al., 1989) in a Unicam 960 spectrophotometer (Waltham, USA). 2.5. Microbial community analysis Rhizospheric soil samples (up to 5 cm depth, 2.5 cm radius from the plant stem) were taken at sowing, after bacterial inoculation (i) and at the end of the experiment (f) from each pot, and for each treatment a compost sample was prepared. DNA extraction. The genomic DNA extraction of rhizosphere samples was performed using the Power Soil DNA Isolation Kit (MO BIO Laboratories, Inc., USA) according to the manufacturer’s instructions. The extracted DNA was kept at 20 °C until its use for DGGE. 2.5.1. 16S rRNA polymerase chain reaction (PCR) conditions The primers 338F-GC and 518R were used for the amplification of the highly variable V3 region of bacterial 16S rRNA gene fragments (Muyzer et al., 1993). The PCR amplification was carried out in 50 ll reaction mixtures containing 1 PCR buffer (Promega, US), 3 mM MgCl2, 5% dimethylsulfoxide, 200 lM of each nucleotide, 30 pmol of each primer, 2 U Taq polymerase (Promega, US), and 1–20 ng of purified DNA. The PCR temperature profile was as described by Henriques et al. (2006), changing the final extension step to 30 min at 72 °C. The reactions were performed in a BioRad iCycler Thermal Cycler (Bio-Rad Laboratories, Richmond, CA, USA). 2.5.2. DGGE PCR-amplified fragments were separated by DGGE using a DCode™ Universal Mutation Detection System (Bio-Rad Laboratories, Richmond, CA, USA). The PCR products containing ca. 300 ng of DNA were loaded onto 8% (w/v) polyacrylamide gels for 16S rRNA gene (37.5:1, acrylamide:bisacrylamide) in 0.5 TAE buffer (20 mM Tris–acetate, pH 7.4, 10 mM sodium acetate, 0.5 mM Na2EDTA) using a denaturing gradient ranging from 35% to 70% for rRNA gene. Electrophoresis was performed at 60 °C in 1 TAE buffer, initially at 20 V (15 min) and then at 75 V (960 min). The gels were stained in a 10 GelGreen Nucleic Acid Stain solution (Biotium Inc., USA) in 0.1 M NaCl. The DGGE images were acquired using a Safe Imager™ Blue-Light Transilluminator (Invitrogen™, USA) and a microDOC gel documentation system (Cleaver Scientific Ltd., UK). The initial (i) and final (f) bacterial community profiles in the different treatments were analyzed using BionumericsÒ software (version 6.6, Applied Maths, St.-Martens-Laten, Belgium). Every gel contained two lanes with a standard of six bands for 16S rRNA gene for internal and external normalization and as an indication

of the quality of the analysis. Band matching position was set at 1% of tolerance, with an optimization of 0.47%. DGGE sample profiles were compared using Jaccard similarity coefficient with 1% tolerance and clustered according to the Ward method. Branch quality was assessed using a cophenetic correlation. Band patterns were converted onto a binary matrix of presence/absence, and correlated with plant growth parameters, through a Canonical Correspondence Analysis (CCA) using PC-ORD (version 5, MJM Software) (Braak, 1986). Monte Carlo randomization test with 1000 interactions was used to detect a significant correlation (p < 0.05). 2.6. Statistical analysis Each plant treatment comprised four replicates. Statistical analysis was performed using the SPSS program (SPSS Inc., Chicago, IL Version 17.0). The data were analyzed through analysis of variance (ANOVA). To detect the statistical significance of differences (P < 0.05) between means, the Duncan test was performed. 3. Results 3.1. Soil properties Characteristics of the agricultural soil are described in Table 1. The exchangeable and available Cd and Zn fractions were determined for the control and spiked soils and were, respectively: non-detectable for non-spiked soil (<0.15 mg Cd kg 1), 2.9 ± 0.1 and 0.7 ± 0.1 mg Cd kg 1 for the 10 mg Cd kg 1, 16 ± 3 and 3.5 ± 0.4 mg Cd kg 1 for the 20 mg Cd kg 1, and 21 ± 2 and 6.0 ± 0.3 mg Cd kg 1 for the 30 mg Cd kg 1 spiked soils; 19 ± 2 and 0.7 ± 0.3 mg Zn kg 1 for nonspiked soil, 52 ± 5 and 10.0 ± 0.4 mg Zn kg 1 for the 100 mg Zn kg 1, 421 ± 52 and 84 ± 10 mg Zn kg 1 for the 500 mg Zn kg 1, and 815 ± 17 and 272.0 ± 28 mg Zn kg 1 for the 1000 mg Zn kg 1 spiked soils. 3.2. Plant biomass and metal uptake: shoots and roots Plants grown in 1000 mg Zn kg 1 presented early toxicity signs and did not evolved from seedling. Therefore, it was not possible to consider this treatment in further analysis and it is thus not included in the presented data. 3.2.1. H. annuus shoots Biomass of H. annuus shoots decreased with increasing Zn concentrations in a significant (P < 0.05) way (Fig. 1A), but not always for Cd exposed plants (Fig. 2A). Bacteria inoculation had no significant (P < 0.05) effect in shoot biomass production for plants grown at the same metal level. Increasing metal concentration in the soil had a positive significant (P < 0.05) effect on metal accumulation in H. annuus shoots for both Cd and Zn. Zinc shoot accumulation ranged from 45 mg Zn kg 1 in plants grown in control soil to 468 mg Zn kg 1 in shoots of plants exposed to 500 mg Zn kg 1 spiked soil (Table 2). Cadmium was not detected in plants grown in the control soil (<0.45 mg Cd kg 1) and ranged from 193 mg Cd kg 1 in plants grown in 10 mg Cd kg 1 spiked soil to 310 mg Cd kg 1 in shoots of plants exposed to 30 mg Cd kg 1 spiked soil (Table 2). Inoculation with strain B2 decreased significantly (P < 0.05) shoot accumulation of Zn (Table 3) but had no effect on Cd accumulation (Table 2), for all tested soil concentrations. In general, strain B1 had no significant (P < 0.05) effect on shoot metal levels. Bioconcentration factors (BCF), defined as the ratio between the metal (Zn or Cd) concentration in the shoots and in soil, were determined for H. annuus shoots (Table 3) – values ranged from

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(A)

12

a

*** F(Cd)3,36 = 12.3

*** F(Zn) 2,27 = 187

a a

10

(A) 12

NS F(B)2,27 = 0.331

a

a

a

*** F(B)3,36 = 1.03

a a

a

a

a

NS F(ZnxB) 4,27 = 1.52

a

a

10

a a

8

a

a

a

a

6

8 B0 B1 B2

4

(g)

(g)

*** F(CdxB)6,36 = 2.00

a

a

B0

6

B1 B2

4

2

2

0 0

100

500 -1

0

Zn soil (mg kg )

0

(B)

20

30

-1

Cd soil (mg kg ) a

*** F(Zn) 2,27 = 63.3

0.7

(B)

*** F(B) 2,27 = 8.72

a

0.6

1

0.8

0.5 b 0.4

b

a

*** F(Cd)3,36 = 16.4

0.9

* F(ZnxB) 4,27= 3.81

ab

(g)

10

0.8

*** F(B) 3,36 = 41.4

a

a

*** F(CdxB)6,36 = 19.5

0.7

b B0

0.6

0.3

B2

(g)

B1

b

ab

0.5

B0 b

b

a

b

a

0.2

a a

0.4

B1 B2

b b

0.1

0.3

c

0.2

0

0.1 0

100

500 -1

Zn soil (mg kg )

0 0

10

20

30

-1

Fig. 1. Helianthus annuus shoot (A) and root (B) biomass (g per pot) when exposed to different Zn concentrations (mg kg 1) in the soil. The error bar represents the SD (n = 4). Two-way ANOVA was performed to determine the influence of soil Zn concentration and of bacterial treatment (B0- no bacteria, B1 – Ralstonia eutropha and B2- Chryseobacterium humi) in shoots and roots biomass. The test results are shown with the test statistic for each case (Zn – soil metal concentration; B – bacterial treatment; Zn  B – metal  bacterial treatments interaction) and as: NS – Non-significant at the level P < 0.05; ⁄Significant at the level P < 0.05; ⁄⁄Significant at the level P < 0.01; ⁄⁄⁄Significant at the level P < 0.001. One-way ANOVA was performed for each Zn concentration in the soil. Means for the same metal concentration with different letters are significantly different from each other (P < 0.05) according to the Duncan test. For the shoot tissues, the F-values of oneway ANOVA are F2,9 = 2.39 (P > 0.05), F2,9 = 0.277 (P > 0.05) and F2,9 = 0.853 (P > 0.05) respectively, for 0, 100 and 500 mg Zn kg 1 spiked soils. For the root tissues, the F-values of one-way ANOVA are F2,9 = 6.05 (P < 0.05), F2,9 = 3.51 (P > 0.05) and F2,9 = 8.27 (P < 0.01) respectively, for 0, 100 and 500 mg Zn kg 1 spiked soils.

0.65 to 2.0 in the case of Zn treatments, while in Cd treatments much higher values were observed, ranging from 10.1 to 20. Bacterial inoculation only had a significant (P < 0.05) effect on metal concentration from soil in plants grown in 500 mg Zn kg 1, with both B1 and B2 decreasing BCF in comparison to non-inoculated plants. 3.2.2. H. annuus roots Generally, root biomass of H. annuus decreased with increasing metal concentrations, and this was significant (P < 0.05) for both Zn (Fig. 1B) and Cd (Fig. 2B) exposed plants. Bacteria inoculation also had a significant (P < 0.05) effect in root biomass production, although it was not possible to establish a clear trend; it seems

Cd soil (mg kg ) Fig. 2. Helianthus annuus shoot (A) and root (B) biomass (g per pot) when exposed to different Cd concentrations (mg kg 1) in the soil. The error bar represents the SD (n = 4). Two-way ANOVA was performed to determine the influence of soil Cd concentration and of bacterial treatment (B0- no bacteria, B1 – Ralstonia eutropha and B2- Chryseobacterium humi) in shoots and roots biomass. The test results are shown with the test statistic for each case (Cd – soil metal concentration; B – bacterial treatment; Cd  B – metal  bacterial treatments interaction) and as: NS – Non-significant at the level P < 0.05; ⁄Significant at the level P < 0.05; ⁄⁄Significant at the level P < 0.01; ⁄⁄⁄Significant at the level P < 0.001. One-way ANOVA was performed for each Cd concentration in the soil. Means for the same metal concentration with different letters are significantly different from each other (P < 0.05) according to the Duncan test. For the shoot tissues, the F-values of oneway ANOVA are F2,9 = 2.39 (P > 0.05), F2,9 = 2.83 (P > 0.05), F2,9 = 0.088 (P > 0.05) and F2,9 = 3.02 (P > 0.05) respectively, for 0, 10, 20 and 30 mg Cd kg 1 spiked soils. For the root tissues, the F-values of one-way ANOVA are F2,9 = 6.05 (P < 0.05), F2,9 = 101 (P < 0.001), F2,9 = 23.1 (P < 0.001) and F2,9 = 1.28 (P > 0.05) and respectively, for 0, 10, 20 and 30 mg Cd kg 1spiked soils.

though that at low metal concentration, biomass of roots was decreased in the presence of bacteria when compared to non-inoculated plants, while at higher Zn (500 mg kg 1) or Cd (20 and 30 mg kg 1) concentrations that effect was not observed or an opposite effect was observed, as in the case of the inoculation with B2 strain (Figs. 1B and 2B). Increasing metal concentration in the soil had a positive significant (P < 0.05) effect on metal accumulation in H. annuus roots for both metals tested. Zinc root accumulation ranged from 51 mg Zn kg 1 in plants grown in control soil to 443 mg Zn kg 1 in roots of plants exposed to 500 mg Zn kg 1 spiked soil (Table 2).

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Table 2 Metal accumulation (mg kg Treatment

1

) in Helianthus annuus roots and shoots.

Zn (mg kg

1

)

Roots

No bacteria B1 B2

Shoots

0

100

500

0

100

500

141 ± 21a 90 ± 35b 51 ± 22b ** (F2,9 = 11.4) *** F(HM)2,27 = 201 *** F(B)2,27 = 21.0 NS F(HMxB)4,27 = 1.68

213 ± 30a 161 ± 35b 134 ± 12b ** (F2,9 = 8.34)

443 ± 80a 430 ± 43a 312 ± 20b * (F2,9 = 7.22)

132 ± 40a 160 ± 11a 45 ± 37b ** (F2,9 = 14.2) *** F(HM)2,27 = 272 *** F(B)2,27 = 15.8 *** F(HMxB)4,27 = 12.0

198 ± 17a 195 ± 17a 174 ± 2b * (F 2,9 = 1.36)

468 ± 33a 326 ± 39b 374 ± 31b *** (F 2,9 = 17.6)

30

10

20

30

667 ± 57a 683 ± 1a 554 ± 42b ** (F2,9 = 11.6)

196 ± 11a 195 ± 8a 193 ± 15a NS (F2,9 = 0.101) *** F(HM)2,27 = 6.94 NS F(B)2,27 = 1.90 NS F(HMxB)4,27 = 1.58

273 ± 7a 272 ± 13a 266 ± 19a NS (F2,9 = 0.310)

310 ± 17a 302 ± 12a 305 ± 14a NS (F2,9 = 0.368)

Cd (mg kg

1

)

10

20

261 ± 54ab 529 ± 20ab 324 ± 70a 653 ± 90a 196 ± 25b 385 ± 90b ** * F2,9 = 5.80) (F2,9 = 13.1) *** F(HM)2,27 = 30.3 *** F(B)2,27 = 40.0 *** F(HMxB)4,27 = 35.7

No bacteria B1 B2

Results are expressed as means ± SD (n = 4). One-way ANOVA was performed for each metal concentration. Means in the same column with different letters are significantly different from each other (P < 0.05) according to the Duncan test. The test results are shown with the test statistic and as: NS – Non-significant at the level P < 0.05; Two-way ANOVA was performed to determine the influence of metal concentration in soil and of bacterial treatment in plant accumulation. The test results are shown with the test statistic for each case (HM – soil metal concentration; B – bacterial treatment; HM  B – metal  bacterial treatments interaction) and as: NS – Non-significant at the level P < 0.05. * Significant at the level P < 0.05. ** Significant at the level P < 0.01. *** Significant at the level P < 0.001.

Table 3 Bioconcentration factor (BCF) in roots and shoots of Helianthus annuus exposed to metals. Treatment

Zn (mg kg

1

)

Roots

Shoots

100 No bacteria B1 B2

500 a

500

0.9 ± 0.2 0.86 ± 0.09a 0.62 ± 0.04b * (F2,9 = 7.22) *** F(HM)1,18 = 241 * F(B)1,18 = 3.55 NS F(HMxB)2,18 = 1.91

2.0 ± 0.3 2.0 ± 0.3a 1.74 ± 0.8a NS (F2,9 = 1.37)

0.94 ± 0.07a 0.65 ± 0.08b 0.75 ± 0.07b *** (F2,9 = 17.6)

10

20

30

10

20

30

26 ± 5ab 32 ± 7a 20 ± 2b * (F2,9 = 5.80) * F(HM)2,27 = 6.64 *** F(B)2,27 = 21.4 NS F(HMxB)4,27 = 1.83

26 ± 1b 33 ± 4a 19 ± 4c ** (F2,9 = 13.1)

22 ± 2a 22.77 ± 0.04a 18 ± 1b ** (F2,9 = 11.6)

20 ± 1a 19.5 ± 0.7a 19 ± 2a NS (F2,9 = 0.101) *** F(HM)2,27 = 359 NS F(B)2,27 = 0.0221 NS F(HMxB)4,27 = 0.253

13 ± 1a 13.7 ± 0.7a 13.6 ± 0.4a NS (F2,9 = 0.310)

10.3 ± 0.6a 10.1 ± 0.4a 10.2 ± 0.5a NS (F2,9 = 0.368)

2.1 ± 0.3 1.6 ± 0.4b 1.3 ± 0.1b ** (F2,9 = 8.34) *** F(HM)1,18 = 111 *** F(B)1,18 = 12.5 * F(HMxB)2,18 = 13.91 Cd(mg kg

No bacteria B1 B2

100 a

a

1

)

Results are expressed as means ± SD (n = 4). One-way ANOVA was performed for each metal concentration. Means in the same column with different letters are significantly different from each other (P < 0.05) according to the Duncan test. The test results are shown with the test statistic and as: NS – Non-significant at the level P < 0.05. Two-way ANOVA was performed to determine the influence of metal concentration in soil and of bacterial treatment in BCF. The test results are shown with the test statistic for each case (HM – soil metal concentration. B – bacterial treatment; HM  B – metal  bacterial treatments interaction) and as: NS – Non-significant at the level P < 0.05. * Significant at the level P < 0.05; ** Significant at the level P < 0.01; *** Significant at the level P < 0.001.

Cadmium accumulation was not detected in plants grown in the control soil (<0.45 mg Cd kg 1), and ranged from 196 mg Cd kg 1 in plants grown in 10 mg Cd kg 1 spiked soil to 683 mg Cd kg 1 in roots of plants exposed to 30 mg Cd kg 1 spiked soil (Table 2). Inoculation with strain B2 always decreased significantly (P < 0.05) root accumulation of Zn and Cd (Table 2), for all tested soil concentrations. Strain B1 had similar effect only in plants exposed to low levels of Zn (0 and 100 mg kg 1).

Bioconcentration factors (BCFs) defined as the ratio between the metal (Zn or Cd) concentration in the roots and in soil, were also calculated for H. annuus roots (Table 3) – values ranged from 0.62 to 2.1 in the case of Zn treatments, while in Cd treatments much higher values were observed, ranging from 18 to 33. Similarly to what happened with metal accumulation, strain B2 always decreased significantly (P < 0.05) BCF in the roots of H. annuus.

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Fig. 3. Analysis of the composition of the bacterial community in rhizosphere soil samples of Helianthus annuus exposed to Zn (A) and Cd (B): DGGE profiles of the samples (I).

3.3. Microbial communities dynamics In order to test the effects of the different treatments on the structure of the bacterial community in the rhizosphere soils, the DGGE patterns of extracted DNA were analyzed (Fig. 3).

A decrease in diversity, translated by a reduction of the number of visible bands, was observed from the initial (i) to the final (f) DGGE profiles in the rhizosphere samples. Reductions of 29%, 4% and 12% were observed for non-spiked soil respectively when non-inoculated, inoculated with B1 and B2; no variations occurred

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for 100 mg Zn kg 1 spiked soil when inoculated with B1 and B2; and 53%, 22% and 4% for 500 mg Zn kg 1 spiked soil respectively when non-inoculated, inoculated with B1 and B2 (Fig. 3A-I). Similarly decreases of diversity from the beginning to the end of the experiment of 85%, 23% and 59% were observed for 10 mg Cd kg 1 spiked soil respectively when non-inoculated, inoculated with B1 and B2; 75%, 45% and 53% for 30 mg Cd kg 1 spiked soil respectively when non-inoculated, inoculated with B1 and B2; and 80%, 27% and 21% for 30 mg Cd kg 1 spiked soil respectively when non-inoculated, inoculated with B1 and B2 (Fig. 3B-I). A decrease on the number of bands was also evident when comparing samples from contaminated and non-spiked soil, especially in the final samples, for both metals, indicating a relation between reduction in diversity and Cd and Zn exposure (Fig. 3). Clustering of the profiles indicated some differences among the rhizosphere soils. For Zn treatments, the profiles of the rhizosphere samples from all soils spiked with 500 mg Zn kg 1 and of B1 and B2 inoculated soils spiked with 100 mg Zn kg 1 clustered apart from the rhizosphere samples of non-spiked soil and non-inoculated 100 mg Zn kg 1, with a correlation of 73%; from the latter group, profiles of control soil samples clustered apart from 100 mg Zn kg 1 samples, with a correlation of 90% (Fig. 3A-II). For Cd treatments the profiles of the rhizosphere samples from all soils spiked with 20 and 30 mg Cd kg 1 collected at the end of the experiment clustered apart from all the other rhizosphere samples, with a correlation of 75%; in the latter group, profiles of the samples generally clustered apart according to their metal concentration (Fig. 3B-II). Canonical correspondence analysis (CCA) showed that the treatments clustered together according to the concentration in the soil (Fig. 4) but not with bacterial inoculation (data not shown). In Zn treatments, samples from the rhizosphere of non-spiked soil distributed in the first and fourth quadrants, samples from the rhizosphere of 100 mg Zn kg 1 spiked soil distribute in the first and third quadrants and samples from the rhizosphere of 500 mg Zn kg 1 spiked soil distributed in the second and third quadrants (Fig. 4A). In Cd treated soils, samples from the rhizosphere of non-spiked soil distributed in the first and fourth quadrants, samples from the rhizosphere of 10 mg Cd kg 1 spiked soil distributed in the third quadrant, samples from the rhizosphere of 20 and 30 mg Cd kg 1 spiked soil distributed in the second and third quadrant (Fig. 4B).

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Axis 1 The common total metal content in unpolluted-soils is below 1 mg kg 1 for Cd and between 10 and 300 mg kg 1 for Zn (Kabata Pendias and Pendias, 1992). The soils used in the present work were spiked to levels of severe metal contamination, to values of 30 mg Cd kg 1 and 500 mg Zn kg 1 and can be considered as recently-contaminated, with high levels of Cd and Zn in soluble forms, as seen by their available and extractable metal fractions. The metal levels chosen in this study are within the range of those considered as of concern by the Dutch standards (targets of 2 mg Cd kg 1 and 140 mg Zn kg 1 and intervention levels at 12 mg Cd kg 1 and 720 mg Zn kg 1) or by the Canadian Soil Quality Guidelines (1.4–22 mg Cd kg 1 and 200–360 mg Zn kg 1). The contamination of soils with such toxic metal concentrations can impair plant growth on such land, causing disturbances in nutrient uptake and other metabolic and physiological processes of plants and consequently growth inhibition (Toppi and Gabbrielli, 1999; Hao et al., 2012). In general, increasing metal concentration in the soil exerts a severe effect on root growth and function, resulting in a diminished uptake of water and nutrients and in ensuing reduction in weight (El-Tayeb et al., 2006). In the present work, this growth inhibition with increasing metal concentrations in

Fig. 4. Canonical Correspondence Analysis (CCA) of the DGGE bacterial community from the rhizosphere samples of H. annus exposed to Zn (A) and Cd (B). The figure shows the relationship among the bands in the DGGE profiles (represented by numbers in the figure) and the tested treatments (j unspiked soil, d 100 mg Zn kg 1 spiked soil; N 500 mg Zn kg 1 spiked soil). The first axis accounted for 18.7%, the second axis accounted for 15.1% (p = 0.038) of the variance for the rhizosphere samples, respectively. The figure shows the relationship among the bands in the DGGE profiles (represented by numbers in the figure) and the tested treatments (j unspiked soil, d 10 mg Cd kg 1 spiked soil; N 20 mg Cd kg 1 spiked soil;  30 mg Cd kg 1 spiked soil). The first axis accounted for 18.4%, the second axis accounted for 11.6% (p = 0.002) of the variance for the rhizosphere samples, respectively.

the soil occurred for Zn exposed plants, but not for plants grown in Cd treated soil. For Cd exposure, root and shoot growth was hardly affected for soil concentrations up to 20 mg kg 1, which is similar to the results of Simon (1998), who reported no effect of Cd concentration on sunflower biomass for Cd soil levels up to 10 mg kg 1 and with the results of Ullah et al. (2011), who found that levels of Cd of 20 mg kg 1 had no significant effect on shoot dry biomass. Cadmium and Zn levels in the plant tissues reported in this work exceeded the phytotoxic concentrations proposed by

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Kabata Pendias and Pendias (1992) – 100 to 400 mg Zn kg 1 and 5 mg Cd kg 1 – and are similar or higher than the values registered for sunflower in other studies. Meers et al. (2005) reported high levels of Zn and Cd accumulation – ca. 250 and 5 mg kg 1 respectively, for sunflower growing in metal contaminated soil (575 mg Zn kg 1 and 6 mg Cd kg 1); in the work of Hao et al. (2012) H. annuus presented accumulation levels of up to 314 mg Zn kg 1 and 1.8 mg Cd kg 1 (when grown in soil contaminated with 1353 mg Zn kg 1 and 2.91 mg Cd kg 1). Nehnevajova et al. (2008) reported levels of up to 401 and 274 mg Zn kg 1 and 1.58 and 0.67 mg Cd kg 1 in sunflower leaves and stems grown in soil contaminated with average levels of 759 mg Zn kg 1 and 0.81 mg Cd kg 1; when H. annuus was genetically manipulated its extraction abilities were 616% and 505% of that in non-modified plants (Nehnevajova et al., 2009). Madejon et al. (2003) observed accumulation levels of up to ca. 75 and 52 mg Zn kg 1 respectively in roots and stems of sunflower plants grown in an area affected by a toxic metal spill (209 mg Zn kg 1 dry soil), with no toxicity signs. In the present studies values as high as 443 mg Zn kg 1 and 683 mg Cd kg 1 were found in plant tissues. The importance of rhizobacteria in plant HM tolerance and their ability to promote growth in metal contaminated soils renders them as a good choice for microbial assisted phytoremediation (Rajkumar et al., 2008). Bacteria having the characteristics of producing ACC deaminase, siderophores and hydrocyanide (HCN), IAA, and of nutrient solubilization (ex. phosphate solubilization) can stimulate plant growth and protect plants from metal toxicity (Glick, 2010; Zhang et al., 2011). A bacteria may affect plant growth and establishment in degraded environments using one or more of these mechanisms,– that is the case of the inoculated bacterial strains, as shown in previous work of Marques et al. (2010). In non-inoculated plants reductions of 19% and 82% in root growth were observed in 100 and 500 mg Zn kg 1 spiked soils, while in the presence of strains B1 and B2 the reductions of growth for plants grown at the highest Zn concentration were lower, 66% and 58% and of 6% and 3% in H. annuus roots exposed to 100 and 500 mg Zn kg 1. When H. annuus was exposed to 20 and 30 mg Cd kg 1 non-inoculated plants reduced growth in 28% and 49%, whereas plants inoculated with B1 showed 25% and 6% and plants inoculated with B2 showed 50% and 18% reductions, respectively. A similar effect of root protection to metal exposure was observed by Rajkumar et al. (2008) for sunflower plants inoculated with the PGPR Bacillus weihenstephanensis in the presence of Zn, Ni and Cu. Despite the fact that neither of the tested PGPR strains induced increases in H. annuus biomass, although that effect was observed for these strains in other plant species, Zea mays (Marques et al., 2010), a protection of the roots under metal exposure was ensured, with strain B2 – C. humi – being more effective across the range of metal exposure. Inoculation with strain B2 decreased significantly shoot and root Zn accumulation and of root Cd levels, being effective at all tested concentrations, while strain B1 was only capable of inducing effects on metal accumulation at low matrix metal levels, and only in the root tissues. This reduction in metal accumulation was previously observed in sunflower plants inoculated with PGPR, namely for Ni (Rajkumar et al., 2008), and in other plants such as barley, tomato, Brassica juncea and Brassica campestris (Belimov et al., 1998; Burd et al., 1998, 2000; Nie et al., 2002; Madhaiyan et al., 2007). It is possible that bacteria can contribute to reduce metal toxicity and accumulation by sharing the metal load with the plant, as many strains have important abilities of bioaccumulation, preventing metal accumulation by plants (Delorme et al., 2001). Microbial mechanisms responsible for decreasing plant metal uptake can also comprise the binding of the metal to functional groups of the selected bacteria, or its chelation by bacterial extracellular polymers, siderophores and

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organic acids (Ma et al., 2011), as well as the capacity to change metal bioavailability by producing ammonia or organic bases that will induce the formation of metal precipitates in the rhizosphere or via metal reduction/oxidation (Chen and Cutright, 2003). R. eutropha and C. humi have shown to produce siderophores and ammonia (Marques et al., 2010), which may have contributed to the decreases observed in metal uptake by H. annuus when inoculated with these strains. However, the opposite effect has also been registered in many plant species – suggesting that organic acids produced by bacteria facilitate metal availability and mobilization for the plant (Whiting et al., 2001). He et al. (2010) reported increased Zn accumulation in PGPR inoculated Orychophragmus violaceus; Sheng and Xia (2006) showed the improvement of Cd uptake through bacteria inoculation in Brassica napus. Accumulation and consequently bioconcentration patterns varied with the level of metal exposure. Accumulation and exclusion are the two basic strategies by which plants respond to high levels of heavy metals in the soil (Vogel-Mikus et al., 2005). In the case of Cd root accumulation was always significantly higher than in the shoots, while for Zn exposed plants root accumulation was generally similar to shoot accumulation Additionally, translocation factors (the ratio between metal accumulation in the shoots and roots), lower than 1 in most cases, were obtained for both metals, appearing thus that H. annuus adopts a tolerance/immobilization strategy, as suggested before (Madejon et al., 2003). Inoculation in the rhizosphere did not change this profile. Bioconcentration factors for Cd were much higher than those shown for Zn. This is coherent with the generally accepted pattern of high Cd transfer coefficient between soil and plant (Kloeke et al., 1984). Metal uptake may be influenced by electro-negativity of the metal ions (Cd = 1.69 and Zn = 1.65, according to the Pauling scale) being positively related to the uptake capacity (Millaleo et al., 2010), which is coherent with the results obtained in the present work. Heavy metals contamination can cause serious changes in the composition of bacterial communities and its activity in soil (Gomes et al., 2010). In this work, this was translated by a decrease in microbial diversity in the metal contaminated soils, indicating a toxic effect of metals in the establishment of the bacterial community throughout time of exposure. Similar profiles were reported by Khan et al. (2010), who also showed that this reduction on microbial diversity increased consistently in soils with increasing Cd and Pb concentrations. Frostegard et al. (1995) also reported similar trends, proposing mortality of the sensitive organisms of the community in the initial stages of the experiment. In spite of this, the inoculation with either B1 and B2 decreased the reduction of the number of bands observed in the DGGE profiles from the initial (i) and final (f) rhizosphere samples, which seems to indicate that the inoculated PGPR prevented the loss of variability of the bacterial community throughout the time of the experiment and had some kind of protection effect from the loss of microbial diversity under metal exposure. In fact, DGGE analyses have shown previously that PGPR could change the bacterial colonization of metal contaminated soils, by inducing the proliferation of other bacteria on the rhizosphere (de-Dashan et al., 2010), as root secretion induced by the PGPR may stimulate the proliferation of other bacteria (Epelde et al., 2010). Nevertheless, in the present work, DGGE profiling and CCA showed that the inoculation with strains R. eutropha (B1) and C. humi (B2), had no visible significant clustering effect – no significant effect of the bio inoculation by PGPR on the microbial community structure have also been reported previously (Mahaffe and Kloepper, 1997; Lottman et al., 2000). However, changes in bacterial composition induced by PGPR inoculation may not be desirable if important native species are lost, thus affecting subsequent crops in a field situation (Roesti et al., 2006). On the other hand, it was observed that variations in concentration of both metals significantly changed the soil

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bacterial community, as previously reported by the study Gomes et al. (2010) for Cd and Zn exposure, and it is possible that the effect of metal exposure was stronger, hiding all possible effects of bacterial inoculation. It seems that various mechanisms of action on phytoremediation by PGPR can occur and that these are both plant and substrate dependent (Grandlic et al., 2008). Additionally, environmental conditions to which an inoculant is exposed will certainly influence whether or not certain plant growth promotion traits are activated (Becerra-Castro et al., 2012), and these processes may be delayed by the high concentrations of HM in the soils (Dell’Amico et al., 2005) – as it appears to have happened in the present work. Therefore, microbial-assisted phytoremediation needs further investigation. 5. Conclusions Although neither of the tested PGPR strains induced increases in plant biomass when the same metal level was analyzed, both reduced losses of weight in metal exposed plants when compared to the control and induced changes in metal bioaccumulation – with strain B2 (C. humi), decreasing significantly shoot and root Zn accumulation and root Cd uptake. Bacterial community diversity was affected by the level of metal pollution in soils and inoculation with both R. eutropha (B1) and C. humi (B2) decreased the reduction of bacterial diversity from the beginning to the end of the experiment, seeming to protect the community from the toxic effects of the metals throughout the time of exposure. It seems thus that the inoculation with sunflower with these bacteria, and particularly with C. humi (B2), appears to be an effective way of enhancing the stabilization abilities of the plant in metal contaminated land, decreasing losses in biomass with increasing metal exposure and ensuring a protection of the roots from metal uptake. This strategy of short term phytostabilization, as H. annuus is an annual species, in soils with degrees of contamination similar to those presented in this study will allow the decrease of metal dispersion and further environmental risks posed by such polluted land. The application of this crop can make the long period of remediation economically and environmentally viable through the valorization of the produced biomass for added value. However, the use of the produced biomass, which presents some degree of metal contamination, should be further studied to understand its suitability for energy purposes. Acknowledgements This work was supported by FCT – Fundação para a Ciência e a Tecnologia and Fundo Social Europeu (III Quadro Comunitário de apoio), research grant of Ana Marques (SFRH/BPD/34585/2007), Helena Moreira (SFRH/BD64584/2009), and Albina Franco (SFRH/ BD/47722/2008) and by National Funds from FCT through project PEst-OE/EQB/LA0016/2011. References Barceloux, D.G., 1999. Zinc. J. Toxicol. Clin. Toxicol. 37, 279–292. Batty, L.C., 2005. The potential importance of mine sites for biodiversity. Mine Water Environ. 24, 101–103. Becerra-Castro, C., Monterroso, C., Prieto-Fernandez, A., Rodriguez-Lamas, L., Loureiro-Vinas, M., Acea, M.J., Kidd, P.S., 2012. Pseudometallophytes colonising Pb/Zn mine tailings: a description of the plant microorganism rhizosphere soil system and isolation of metal tolerant bacteria. J. Hazard. Mater. 217–218, 350–359. Belimov, A.A., Kunakova, A.M., Vasilyeva, N.D., Kovatcheva, T.S., Dritchko, V.F., Kuzovatov, S.N., Trushkina, I.R., Alekseyev, Y.V., 1998. Accumulation of radionuclides by associative bacteria and the uptake of 134Cs by the inoculated barley plants. In: Malik, K.A., Mizra, M.S., Ladha, J.K. (Eds.), Nitrogen Fixation With Non-Legumes. Kluwer Academic Publishers, Dordrecht, pp. 275–280.

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