Environ Sci Pollut Res DOI 10.1007/s11356-014-2848-1

RESEARCH ARTICLE

Phytomanagement of Cd-contaminated soils using maize (Zea mays L.) assisted by plant growth-promoting rhizobacteria Helena Moreira & Ana P. G. C. Marques & Albina R. Franco & António O. S. S. Rangel & Paula M. L. Castro

Received: 22 September 2013 / Accepted: 28 March 2014 # Springer-Verlag Berlin Heidelberg 2014

Abstract Zea mays (L.) is a crop widely cultivated throughout the world and can be considered suitable for phytomanagement due to its metal resistance and energetic value. In this study, the effect of two plant growth-promoting rhizobacteria, Ralstonia eutropha and Chryseobacterium humi, on growth and metal uptake of Z. mays plants in soils contaminated with up to 30 mg Cd kg−1 was evaluated. Bacterial inoculation increased plant biomass up to 63 % and led to a decrease of up to 81 % in Cd shoot levels (4– 88 mg Cd kg−1) and to an increase of up to 186 % in accumulation in the roots (52–134 mg Cd kg−1). The rhizosphere community structure changed throughout the experiment and varied with different levels of Cd soil contamination, as revealed by molecular biology techniques. Z. mays plants inoculated with either of the tested strains may have potential application in a strategy of soil remediation, in particular short-term phytostabilization, coupled with biomass production for energy purposes.

Responsible editor: Elena Maestri H. Moreira : A. P. G. C. Marques : A. R. Franco : A. O. S. S. Rangel : P. 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 e-mail: [email protected] H. Moreira e-mail: [email protected] A. P. G. C. Marques e-mail: [email protected] A. R. Franco e-mail: [email protected] A. O. S. S. Rangel e-mail: [email protected]

Keywords Zea mays . Soil . PGPR . Phytomanagement . Cadmium . Biomass production . Remediation

Introduction Soil pollution by heavy metals is one of the main ecological problems worldwide. Cadmium is among the main metal contaminants and is one of the most toxic substances to living organisms, affecting plants, animals, and humans (Glick 2010; Kirkham 2006). This toxic metal has been released into the soil due to several anthropogenic processes, namely mining activity, production of batteries and fertilizers, and disposal of contaminated waste (Cui and Wang 2006; Lagriffoul et al. 1998). In plants particularly, high levels of this nonessential element can induce several negative effects, diminishing their growth and nutrient uptake and inhibiting photosynthesis (Prasad 1995; Toppi and Gabbrielli 1999). Depending on total concentration levels of Cd in soil, polluted areas are impaired for several uses, namely for residential, commercial, and, particularly, agricultural purposes, which have the lowest acceptable threshold level. Remediation techniques are therefore required to reduce or attenuate environmental risks. Phytoremediation, the use of plants to restore heavy metal-contaminated land, has arisen as a promising alternative technology to the use of classical chemical or physical cleaning methods (Juwarkar et al. 2010; Marques et al. 2009) in not severely polluted areas (Kavamura and Esposito 2010; Khan 2005). Although this technique is less expensive and less harmful to soil microbial diversity (Glick 2010), a long time is required to achieve acceptable metal levels. A soil remediation strategy known as phytomanagement can compensate such a time by adding value to the plants used for remediation and preventing movement of contaminants through the environment while reducing contamination (Fässler et al. 2010). This kind of

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management of contaminated soil also contributes to enhance soil quality and productivity. Fast-growing plants with high yield, biomass, and metal tolerance such as maize (Zea mays L.), a cadmium-tolerant plant, have been explored as alternatives in soil remediation applications (Meers et al. 2005, 2010; Thewys et al. 2010). While mitigating contamination, these plants can be used for biomass production, which is considered one of the most promising renewable energy options (Meers et al. 2010; Mleczek et al. 2010; Ruiz et al. 2009). However, there are some restrictive issues concerning the remediation process of metal-contaminated soils using plants, such as the bioavailability of the metals in soil, which is often a limiting factor for their extraction by plants (Li and Ramakrishna 2011). An alternative to chemical amendments, namely the use of chelating agents (Gunawardana et al. 2010), is the use of metal-resistant rhizosphere bacteria which may also enhance metal bioavailability (Rajkumar et al. 2010). The limited growth of plants exposed to high Cd levels, which can induce several toxic symptoms (Prasad 1995), and the Cd content on the plant harvestable parts (McKendry 2002) are also problems in plant-based remediation strategies. Plant growth-promoting rhizobacteria (PGPR) have been reported to reduce heavy metal stress and to promote phytoremediation and biomass production in contaminated soils (Ma et al. 2011; Marques et al. 2013; Miransari 2011; Saharan and Nehra 2011). Growth-promoting mechanisms include the production of phytohormones (auxins, cytokinins, and gibberellins), the siderophores, the enzyme 1-aminocyclopropane-1-carboxylate (ACC) deaminase, and the fixation of atmospheric nitrogen (Glick 2010; Ma et al. 2011; Saharan and Nehra 2011). There are some studies involving Z. mays in phytoremediation (Li et al. 2009; Meers et al. 2005; Thewys et al. 2010; Wang et al. 2007; Wójcik and Tukiendorf 2005) and biomass production strategies (Dhugga 2007). Studies to elucidate the benefits of PGPR for growth of maize in Cd-contaminated soil towards achieving a phytomanagement/remediation strategy are valuable. In the present study, two bacterial strains, Ralstonia eutropha and Chryseobacterium humi (a novel species) (Pires 2010), isolated from a heavy metal-contaminated site, which had shown to promote the growth of Z. mays plants in agricultural soil (Marques et al. 2010), were tested. The effects of these bacterial strains on the growth and metal accumulation of Z. mays plants grown in Cd-contaminated soils, as well as changes in the rhizosphere community, were investigated.

Materials and methods Soil preparation Agricultural sandy loam soil with no detectable Cd and average pH of 6.71, 3.1 % organic content, 4.8 % water content,

1,736 mg kg−1 of N, 2,600 mg kg−1 of P, and 10,600 mg kg−1 of K (soil dry weight (DW)) was sampled randomly from a depth of 0–20 cm (other soil properties have been previously described in Marques et al. 2013). Root and litter materials were removed, and samples were air dried, passed through a 2-mm sieve, and mixed uniformly. Cadmium-treated soil was prepared by autoclaving soil (120 °C for 70 min in two consecutive days), drying in an oven at 40 °C for 4 days, and mixing with CdCl2 to achieve concentrations of 0, 10, 20, and 30 mg Cd kg−1 to mimic different contamination levels. Soil moisture content was maintained at 60 % of the water holding capacity by adding deionized sterile water. Spiked soil was subjected to three cycles of wet and dry processes for approximately 4 weeks in the greenhouse and was mixed once a week so that Cd was evenly dispersed in the soil (Blaylock et al. 1997). Experimental plan Two PGPR strains, R. eutropha (B1) and C. humi (B2), have been isolated from sediment samples collected from an industrially contaminated site in northern Portugal and shown to be able to tolerate Cd concentrations up to 500 mg L−1 in liquid cultures (Pires 2010). A pot experiment was conducted in a controlled growth room (12-h photoperiod, 450 μmol m−2 s−1 photosynthetically active radiation, 18–21 °C temperature range, 50–60 % relative humidity range) in order to test the effect of the selected PGPR on Z. mays plants grown in Cd-treated soil. Treatments in the greenhouse included assays with non-spiked soil and soil spiked with Cd at three concentrations, 10, 20, and 30 mg kg−1. Each treatment was subjected to a different type of inoculation: control (no bacteria), B1 (R. eutropha), and B2 (C. humi) strains. Four replicates were used for each Cd level/ inoculation type treatment. Z. mays seeds (variety Aveline, purchased from Lusosem, Portugal) were surface sterilized with 0.5 % (v/v) NaOCl for 10 min and were rinsed several times with deionized sterile water. Seeds were germinated in plastic pots (8-cm diameter and 10-cm height) with 400 g of the testing soils. Each pot received six seeds which were placed at 2-cm depth. Pots were randomly rearranged in the greenhouse every 2 weeks during the experiment. After germination, seedlings were reduced to four per pot. Ten milliliters of each bacterial strain suspension (108 CFU mL−1) in nutrient broth was used for the inoculation, by spraying soil surfaces (Marques et al. 2010), 2 weeks after germination. To the control pots, 10 mL of sterile nutrient broth was added. Plants were harvested after 12 weeks, separated in roots and shoot and washed with tap water, followed by washes with a 0.1 M HCl solution, and deionized sterile water. Shoot and root biomass was determined after oven drying at 70 °C for 48 h, grinding, and sieving to <1 mm.

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These samples were then digested at high temperature in a PerkinElmer microwave (Waltham, USA), using the method of USEPA 3052, and the Cd content in the samples was determined using flame atomizer-atomic absorption spectroscopy (FA-AAS) (Wallinga et al. 1989) in a Unicam 960 spectrophotometer (Waltham, USA) with a detection limit for Cd of 0.0033 mg L−1. Rhizosphere soil of each pot was collected at the end of the experiment for metal bioavailability analysis. Ethylenediaminetetraacetic acid (EDTA) (exchangeable) and ammonium acetate (NH4-Ac) (extractable) Cd fractions (Thomas 1982) were determined for the soils using, respectively, 1:10 soil 0.05 M EDTA (Houba et al. 1995) and 1:5 soil 1 M NH4-Ac (De Koe 1994). The resulting solutions were incubated for 2 h at 20 °C and then filtered through a 0.45-μm cellulose acetate filter, and the Cd content in the samples was determined using the FA-AAS. DNA extraction, PCR amplification, and DGGE Rhizosphere soil (0.25 g) of each treatment was collected 1 day after bacterial inoculation (i) and at the end of the experiment (f) and used to extract total DNA, which was purified by the Power Soil DNA Isolation Kit (MO BIO Laboratories, Inc., USA). Primers 338F-GC (338F primer with a GC clamp to stop total denaturation of DNA during denaturing gradient gel electrophoresis (DGGE)) and 518R (Muyzer et al. 1993) were used for the amplification of the V3 region of bacterial 16S rRNA gene from the purified DNA. The polymerase chain reaction (PCR) amplification was performed in 50-μL reaction mixtures containing 1× PCR buffer, 3 mM MgCl2, 5 % dimethyl sulfoxide, 200 μM of each nucleotide, 0.6 μM of each primer, 2 U Taq polymerase (Promega, USA), and 1–20 ng of purified DNA. The PCR temperature profile was adapted from Henriques et al. (2006): initial denaturation at 94 °C for 5 min; 30 cycles of denaturation at 92 °C for 30 s; annealing at 55 °C for 30 s; extension at 72 °C for 30 s; and final extension at 72 °C for 30 min. The reactions were performed in a Bio-Rad iCycler Thermal Cycler (Bio-Rad Laboratories, Richmond, CA, USA). Electrophoresis in 1.5 % (w/v) agarose gel was performed to analyze the DNA amplification products which were stained with SYBR® Safe DNA Gel Stain. DGGE was performed using a DCodeTM Universal Mutation Detection System (Bio-Rad Laboratories, Richmond, CA, USA). PCR products (20 μL) containing ca. 300 ng of DNA were loaded onto 8 % (w/v) polyacrylamide gels (37.5:1, acrylamide/bisacrylamide) in 0.5× TAE buffer (20 mM Trisacetate, pH 7.4, 10 mM sodium acetate, 0.5 mM Na2EDTA), using a denaturing gradient ranging from 35 to 70 % for 16S 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 ImagerTM BlueLight Transilluminator (Invitrogen TM , USA) and a MicroDOCTM gel documentation system (Cleaver Scientific Ltd, UK). Analysis of DGGE banding pattern Gel images showing bacterial community profiles in the different treatments were analyzed with the Bionumerics® software (version 6.6, Applied Maths, St-Martens-Laten, Belgium). Every gel contained two lanes (HyperLadderTM 100 bp—Bioline) with a standard of 12 bands for 16S rRNA gene for internal and external normalization as an indication of the quality of the analysis. Band’s matching position was set at 1 % tolerance, with an optimization of 0.47 %. DGGE sample profiles were compared using Jaccard’s similarity coefficient with 1 % tolerance and clustered according to the Ward method. Branch quality was assessed using a cophenetic correlation. Banding patterns were converted onto a binary matrix of presence/absence, and a Canonical correspondence analysis (CCA) was carried out in order to correlate with plant parameters (biomass and Cd accumulation). Monte Carlo randomization test with 999 interactions was used to detect a significant correlation (P<0.05). Both tests were performed using PC-ORD, version 5, MJM Software (Teer Braak 1986). Statistical analysis Statistical analysis was performed using the SPSS program (SPS Inc., Chicago, IL, USA, version 17.0). Analysis of variance (ANOVA) and the Duncan test (P<0.05) were performed to compare treatments. Chemical reagents The chemicals used were analytical grade. Liquid reagents were obtained from Pronalab (Sintra, Portugal) and Promega (USA) and solid reagents from Sigma-Aldrich (Missouri, USA) and Merck (Darmstadt, Germany). Deionized water had a specific conductance less than 0.1 μS cm−1.

Results Effects of Cd and of PGPR on biomass and metal accumulation, bioconcentration and translocation in Z. mays plants With increasing Cd concentrations, both shoot and roots biomasses of Z. mays plants decreased significantly (P<0.05), although the differences between the bacterial treatments and the control varied among the Cd concentrations (Fig. 1). On

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different from that found for the roots. In the roots, accumulation of Cd significantly increased (P<0.05) with PGPR (except for the 20 mg Cd kg−1), whereas in the shoots, it significantly decreased (except for 30 mg Cd kg−1). Metal accumulation in roots was higher than in shoots; while in the belowground tissues, Cd accumulation ranged from 25 to 138 mg kg−1, and in the aerial part, it ranged from 4 to 88 mg kg−1. Inoculation with either strain decreased significantly shoot accumulation in plants grown in all Cd-spiked soils, and for B1, it was always significant (P<0.05). However, the opposite effect was induced on root Cd accumulation by the PGPR treatment with both strains increasing it at all soil concentrations (although not significantly (P<0.05) at 20 mg Cd kg−1). Bioconcentration factors (BCFs) were determined as the ratio between the metal concentration in plant organs to that in the soil (Fig. 2a, b). Values of BCF for Z. mays plants shoot ranged from 0.4 to 2.9 and were generally significantly

the contrary, the plant–bacteria associations showed increased shoot and roots biomass, although not always significantly (P<0.05), depending on the Cd concentration (Fig. 1). The B2 strain contributed to significantly (P<0.05) improve the biomass of shoots and roots, particularly at the 10 mg Cd kg−1 level—17 and 47 %, respectively—whereas strain B1 only increased plants’ biomass at a soil concentration of 20 mg Cd kg−1 (17 and 63 % for shoots and roots, respectively). However, at the highest Cd level of 30 mg kg−1, none of the bacterial strains had a significant influence on Z. mays plants’ biomass (Fig. 1). The cadmium accumulation in shoots and roots of Z. mays plants (Table 1) was significantly related to increasing Cd concentrations in the soil (P<0.05) (Cd was not detected in plants grown in non-spiked soil). However, at each soil concentration individually, the aboveground organs of plants associated with PGPR showed a Cd accumulation pattern

*** F(Cd)3,36 = 111.033

a

*** F(B)3,36 = 13.651

25

* F(CdxB)6,36 = 3.165

shoots biomass (g)

20 15

a

a a

b

b

a,b

b

a

a

b

a

a

10 5 0 Cd 0

Cd 10

Cd 20

Cd 30

[Cd] soil (mg kg-1)

b

8

a

*** F(Cd)3,36 = 32,623

roots biomass (g)

7

*** F(B)3,36 = 6,400

6

NS F(CdxB)6,36 = 1,352

a a ,

5 4

b

a

3

b a,b

a ,

b a ,

2

a ,

a ,

1 0 Cd 0

Cd 10

Cd 20

Cd 30

[Cd] soil (mg kg-1)

Fig. 1 Shoot (a) and root (b) biomass (g/pot) of plants exposed to different Cd concentrations (mg kg−1) in soil, with or without PGPR. 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 on shoot and root biomass. The test results are shown with the test statistic for each case (Cd soil metal concentration; B bacterial treatment; CdxB metal × bacterial treatment interaction) and as NS nonsignificant 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 one-way ANOVA are F2, 9 =9.24 (P<0.01), F2, 9 =6.452 (P<0.05), F2, 9 =4.447 (P<0.05), and F2, 9 =1.729 (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 =10.552 (P<0.01), F2, 9 =3.334 (P>0.05), F2, 9 =7.332 (P<0.05), and F2, 9 =0.736 (P>0.05), respectively, for 0, 10, 20, and 30 mg Cd kg−1 spiked soils

Environ Sci Pollut Res Table 1 Cd accumulation (mg kg−1) in Zea mays roots and shoots Treatment

Cd (mg kg−1) Roots 10

Control B1 B2

Shoots 20

24.97±6.05b 78.43±4.63a 52.25±2.23a 83.61±3.92a a 71.34±23.43 82.18±7.01a ***F(Cd)2, 27 =17,576 ***F(B)2, 27 =203,286 ***F(CdxB)4, 27 =4.781

30

10

20

118.15±7.20b 135.36±9.23a 137.94±8.37a

21.45±4.29a 37.67±4.68a 6.52±1.85b 29.65±2.16b b 4.08±0.74 21.32±1.79c ***F(Cd)2, 27 =1,172.781 ***F(B)2, 27 =48.226 ***F(CdxB)4, 27 =6,472

30 88.20±5.27a 73.54±6.56b 82.43±0.63b

No Cd accumulation was detected for plants grown in non-spiked soil. Therefore, results are not included in the table. The results are expressed as means ±SD (n=4). One-way ANOVA was performed for each Cd concentration. Means in the same column with different superscript 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 non-significant at the level P<0.05 and significant at the levels P<0.05 and P<0.001, respectively. Two-way ANOVA was performed to determine the influence of Cd concentration in soil and of bacterial treatment in plant accumulation. The test results are shown with the test statistic for each case (Cd—soil metal concentration; B—bacterial treatment; CdxB—metal × bacterial treatment interaction) and as non-significant at the level P<0.05 and significant at the levels P<0.05, P<0.01, and P<0.001 NS non-significant *P<0.05; **P<0.01; ***P<0.001, significance level

increased (P<0.05) by the increasing Cd concentration in the soil. Bacterial inoculation significantly (P<0.05) decreased the shoot BCFs (with the exception of B2 inoculation of plants grown at 30 mg Cd kg−1). The highest decrease in the BCF values was recorded for plants grown in 10 mg Cd kg−1 spiked soil, with reductions of 70 and 81 % for B1- and B2-treated plants, respectively. Plants grown in soils spiked with 20 and 30 mg Cd kg−1 also presented reductions in the BCFs when treated with PGPR. In Z. mays plant roots, BCF values ranged from 2.5 to 7.1, with the highest increase of 185 % observed in plants grown in 10 mg Cd kg−1 spiked soil in the presence of PGPR. Translocation factors (TF) were determined as the ratio of Cd concentration in the shoot tissue over Cd concentration in the root tissues (Fig. 2c). All treatments presented TF lower than 1, with the highest values registered in plants grown in 30 mg Cd kg−1 spiked soil. Nevertheless, inoculation with either bacterial strains induced a significant (P<0.05) reduction in TF values with B2 leading to more marked decreases in the TF values at the lower Cd concentrations (10 and 20 mg Cd kg−1). Mobilization of Cd in soil In order to obtain information about Cd availability to plants, NH4-Ac (available) and EDTA-extractable (exchangeable) Cd forms in rhizosphere soils were determined at the end of the experiment. Values ranged from 0.02 to 6.1 mg Cd kg−1 and from 0.33 to 27 mg Cd kg−1, respectively, for NH4-Ac and EDTA-extractable forms, at the different Cd soil concentrations (Table 2). Strain B1 always increased the available and

exchangeable forms of this metal in soil fractions, although not always significantly; in non-spiked soil, inoculation with strain B1 increased the available fraction by 1,250 %. In Cdtreated soils, strain B1 increased up to 12 and 9 % the available and exchangeable fractions, respectively, although not always in a significant way. In soils inoculated with strain B2, a general and significant (P<0.05) increase of Cd mobility was noticed (except for plants exposed to 20 mg Cd kg−1, when the effect was neutral). The increment on Cd bioavailability was higher in soils inoculated with this bacterial strain rather than with strain B1, with Cd in the available fraction of inoculated treatments being 60 % higher in non-spiked soils, as compared with the non-inoculated control; an increase of up to 59 % was also observed for soils spiked with 10 mg Cd kg−1. The exchangeable fraction was also increased but only up to 22 % at 30 mg Cd kg−1. Rhizosphere community Bacterial rhizosphere community profiles at the beginning (i) and at the end (f) of the experiment were analyzed by DGGE. A total of 52 bands were detected and assigned with an average of 18 bands in each DGGE profile. As shown in Fig. 3a, generally samples collected immediately after the inoculation had more visible bands, suggesting the presence of more bacterial taxa, than samples collected at the end of the experiment. Clustering of the DGGE profiles showed two main clusters with a correlation of 74 %, the first including samples collected from non-spiked soil (initial and final) and from soil spiked with 10 mg Cd kg−1 (initial) and the second including samples

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a

*** F(Cd)3,36 = 209.736

3.5

*** F(B)3,36 = 70.684

a

BFC- shoots

3.0

a

***F(CdxB)6,36 = 17.649

a

b

2.5 a

2.0 b

Control

1.5

B1

c b

1.0

B2 b

0.5 0.0 Cd 10

Cd 20

Cd 30

[Cd] soil (mg kg-1)

b

10

NS F(Cd)3,36 = 1.003

a

9

*** F(B)3,36 = 24.363

8

*** F(CdxB)6,36 = 14.090

BCF- roots

7 6

a

5 4

a

a

a a

a

b

b

3 2 1 0 Cd 10

Cd 20

Cd 30

[Cd] soil (mg

c

1.2

kg-1) *** F(Cd)2,27 = 50.900

a

***F(B)2,27 = 98.015

1.0 * F(CdxB)4,27 = 28.883

a

0.8 TF

Fig. 2 Shoot (a) and root (b) bioconcentration factors (BCF) and translocation factors (TF) (c) of plants exposed to different Cd concentrations (mg kg−1) in soil, with and without PGPR. 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 on shoot and root BCF and TF. The test results are shown with the test statistic for each case (Cd soil metal concentration; B bacterial treatment; CdxB metal × bacterial treatment interaction) and as NS nonsignificant 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 BCF of shoots, the F values of one-way ANOVA are F2, 9 =47.387 (P<0.001), F2, 9 =27.108 (P<0.001), and F2, 9 = 9.146 (P>0.01), respectively, for 10, 20, and 30 mg Cd kg−1 spiked soils. For the BCF of roots, the F values of one-way ANOVA are F2, 9 =19.019 (P<0.001), F2, 9 = 1.025 (P<0.05), and F2, 9 =7.723 (P<0.05), respectively, for 10, 20, and 30 mg Cd kg−1 spiked soils. For the translocation factors, the F values of one-way ANOVA are F2, 9 =79.054 (P<0.001), F2, 9 = 13.931 (P<0.01), and F2, 9 = 12.273 (P<0.01), respectively, for 10, 20, and 30 mg Cd kg−1 spiked soils

b

a

b

0.6 b

0.4

b b

0.2

b

0.0 Cd 10

Cd 20 [Cd] soil (mg

collected from soil spiked with 10 (final), 20, and 30 mg Cd kg−1 (initial and final) (Fig. 3b). Within both clusters, samples generally grouped together according to the collection time and metal concentration in the soils. DGGE band pattern showed bands appearing more frequently in samples (initial and final) collected from the rhizosphere of non-spiked soil and soil spiked with 10 mg Cd Zn kg−1 and others appearing more frequently in samples collected from 20 and 30 mg Zn kg−1.

Cd 30 kg-1)

The CCA (Fig. 4) showed that the final samples from the rhizosphere of the control and 10 mg Cd kg−1 spiked soil distributed in the first and fourth quadrant, respectively, while samples from 20 and 30 mg Cd kg−1 spiked soil distributed in second and third quadrants. Results from CCA showed that all samples clustered according to metal concentration in soil and not to bacterial treatment (data not shown). Cadmium accumulation in root and shoot which presented an intraset correlation of, respectively, −0.917 and −0.911 with axis 1,

Environ Sci Pollut Res Table 2 The NH4-Ac and EDTA-extractable Cd levels in soils (mg kg−1) Treatment

Cd (mg kg−1) 0

10

20

30

NH4-Ac

EDTA

NH4-Ac

EDTA

NH4-Ac

EDTA

NH4-Ac

EDTA

0.02±0.03c 0.25±0.05b 0.4±0.01a ***(F2,9=97)

0.33±0.09b 0.37±0.1b 0.51±0.05a ***(F2,9=23)

1.7±0.5b 1.9±0.3b 2.7±0.2a ***(F2,9=39.6)

6.6±0.3b 7.2±0.1a 7.3±0.2a ***(F2,9=49.8)

3.6±0.2b 4.0±0.0a 3.4±0.3b ***(F2,9=6.43)

14.0±1.0ab 15.1±0.5a 13.0±1.0b **(F2,9=2.24)

5.3±0.5b 5.5±0.4b 6.1±0.1a ***(F2,9=129)

22.1±1.0b 25.0±2.0b 27.0±3.0a *** (F2,9=6.98)

Control B1 B2

Results are expressed as means ± SD (n=4). One-way ANOVA was performed for each d concentration. Means in the same column with different superscript 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*, ***Significant at the level P < 0.05 and P < 0.001, respectively. Two-way ANOVAwas performed to determine the influence of Cd concentration in soil and of bacterial treatment in plant accumulation. The test results are ***F(Cd)3,36= 21169, ***F(B)2,36=351, ***F(CdxB)6,36=128 and ***F(Cd)3,36= 11823, ***F(B)2,36=81.4, ***F(CdxB)6,36=69.6 for available and exchangeable fractions respectively (Cd – soil metal concentration; B – bacterial treatment; CdxB - metal x bacterial treatments interaction and 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).

contributed to separate samples collected from 30 mg Cd kg−1 spiked soil from the others. On the other hand, root and shoot biomass, which presented an intraset correlation of, respectively, −0.589 and −0.713 with axis 2, contributed to delineate the group composed of samples collected from non-spiked soil.

Discussion Z. mays plants have high biomass and are tolerant to heavy metals, including Cd, Zn, and Pb (Wuana and Okieimen 2010). Plant growth-promoting rhizobacteria are known to have a beneficial effect on plant growth, enhancing nutrient uptake and diminishing the deleterious effects of heavy metals in plants grown in contaminated soils (Khan 2005; Wójcik and Tukiendorf 2005). In this work, two heavy metal-tolerant PGPR, namely R. eutropha (B1) and C. humi (B2) (Marques et al. 2010), were used as bioinoculants for Z. mays in Cdcontaminated soils. These bacteria exhibit high indole acetic acid, hydrogen cyanide, ammonia, and siderophore production and have been shown to increase biomass of Z. mays plants grown in non-contaminated soil in a greenhouse experiment (Marques et al. 2010). Both bacteria have also ACC deaminase activity and exhibit tolerance to Cd up to 500 mg L−1 (Pires 2010) and seem thus appropriate for the inoculation of the plant in the establishment of phytoremediation strategies. Cadmium has no known function in plants (Lagriffoul et al. 1998; Naees et al. 2011). Unpolluted soils, according to Pierzinsky et al. (2000), contain total soil Cd concentration below 1 to 2 mg kg−1, and hazardous levels are reached for values up to 30 mg kg−1. The metal levels chosen in this study

were over the values considered as of concern by the Dutch standards (targets of 2 mg Cd kg−1 and intervention levels of 12 mg Cd kg−1) or by the Canadian Soil Quality Guidelines (1.4 and 22 mg Cd kg−1). In such conditions, survival is only guaranteed for plants that evolved some resistance mechanisms to high metal levels (Kavamura and Esposito 2010). In the present work, Z. mays plants were exposed to phytotoxic Cd levels, and increasing Cd contamination levels generally reduced plant biomass, although down to a level that was essentially stabilized up to 30 mg kg−1 (Fig. 1). Moreover, such a reduction was more intensive for roots than for shoots, when compared to the respective controls, likely reflecting the higher accumulation of the metal observed for the former. The study of Shi and Cai (2009) has also shown similar results for eight different energy crops grown with increasing Cd treatments (up to 200 mg kg−1). In the present work, metal accumulation in plant tissues was generally increased with increasing Cd concentration, in accordance to what has been previously reported by Wang et al. (2007). In their study, they have reported that concentrations of Cd ranging from 1 to 100 μM have inhibitory effects on root growth of the exposed maize seedlings at 100 μM, but Cd uptake and accumulation at this concentration reached the highest values at the end of the experiment, with an average of 1,954.5 mg kg−1 dry weight. Lagriffoul et al. (1998) have also reported similar results, i.e., Z. mays plants treated with increasing Cd concentrations, maximum of 25 μM, show an increase in plant Cd content up to 304 mg kg−1 and a significant decrease in shoot and leaf dry biomass at this metal concentration. Sunflower plants (Helianthus annuus) also show a positive relation between the levels of the metal in the soil (0, 10, 20, and 30 mg Cd kg−1) and its accumulation by the plants (Marques et al. 2013). To mitigate the metal toxicity effects, some species exclude metals from plant tissues and/or retain them in roots (Kirkham

Environ Sci Pollut Res Fig. 3 Analysis of the bacterial community in rhizosphere soil samples of Zea mays exposed to Cd: DGGE profiles of the samples collected in non-spiked soil and 10 kg−1 contaminated soil (Cd 10) (a); DGGE profiles of the samples collected in 20 mg kg−1 (Cd 20) and 30 mg kg−1 (Cd 30) contaminated soils (b); similarity of the rhizosphere soil samples based on Ward clustering method (c) (C control; B1 sample inoculated with strain B1; B2 sample inoculated with strain B2; i beginning of the experiment; f end of the experiment; M ladder)

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2006; Wang et al. 2007). Previous reports have classified Z. mays as a root accumulator (Li et al. 2009; Mench and Martin 1991), which was also observed in the present study. Taken all these information together, results did suggest that maize plants tend to retain excess of Cd in the roots, possibly in the attempt to prevent/reduce deleterious effects of the metal on carbon assimilating apparatus of the aerial parts of the plant. Cadmium can enter root cells as Cd2+ or as Cd chelates and reaches the stele through an intracellular pathway (symplasmic)—loaded from the symplasm into the xylem and subsequently being sequestered in the vacuole inside plant cells. The metal can also reach the xylem via an extracellular pathway (apoplasmic)—being immobilized in the cellular

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walls and intercellular spaces—which is restricted to the extreme root tip and to the regions where lateral roots initiate (Lux et al. 2011). The endodermis with its Casparian bands represents a barrier to metal movement through the apoplasm, avoiding thus the transport to the shoot tissues (Ranathunge et al. 2005). Schreiber et al. (1999) have shown that exposure of maize to Cd results in an increase in the production and changes in the composition of endodermal impregnation materials, reducing apoplasmic movement of Cd and its consequent transport to the shoots, which seems to be in accordance to what occurred in the present study. Plant growth-promoting bacteria can influence the growth of plants through different mechanisms (Ma et al. 2011;

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Axis 1 Fig. 4 Canonical correspondence analysis (CCA) of the DGGE bacterial community from the rhizosphere samples of Zea mays exposed to Cd. The figure shows the relationship among samples from the DGGE profiles and the tested treatments (white square (0) unspiked soil, white circle (10) 10 mg Cd kg−1 spiked soil; black up-pointing triangle (20) 20 mg Cd kg−1 spiked soil; black circle (30) 30 mg Cd kg−1 spiked soil; C control; B1 sample inoculated with strain B1; B2 sample inoculated with strain B2). The first axis accounted for 19.1 % and the second axis accounted for 12.5 % (P=0.007) of the variance for the rhizosphere samples, respectively. Vectors from joint biplot represent strength and direction of environmental data (Cd root Cd accumulation in root; Cd shoot Cd accumulation in shoot; Bm root root biomass; Bm shoot shoot biomass)

Saharan and Nehra 2011). Inoculation of maize plants with the two metal-resistant bacteria under study (obtained from a heavy metal-contaminated soil (Marques et al. 2010; Pires 2010)) increased root and shoot biomass not only in nonspiked soil but also in Cd-spiked soils, showing a general tendency of improvement of plant growth, one of the main characteristics described for these bacteria (Naees et al. 2011). These results suggest that these strains can be possibly used as biofertilizers (Babalola 2010; Marques et al. 2010). However, this is not always the trend, as the same B1 and B2 strains used with sunflower growing at similar Cd soil concentrations, no effect on shoot biomass and a general decrease of root biomass of the plant can be observed (Marques et al. 2013). In the present study, Z. mays plants were able to accumulate Cd in shoots and roots to levels much higher than the usual toxicity threshold for plant tissues (5–10 mg kg−1 plant tissue) (Förstner 1995), without visible toxicity symptoms. Generally, both tested strains had a significantly positive effect on root

Cd uptake. Taken together with the fact that maize is a crop known for its low metal shoot accumulation abilities (Meers et al. 2005), an even more reduced shoot metal accumulation occurred in the plants grown in bacterial inoculated soils, which appears to be an enhanced metal exclusion strategy that suggests some shielding effect provided by the bacteria. A similar effect has also been reported by Jiang et al. (2008), showing a reduction in shoot accumulation in Z. mays plants inoculated with Burkholderia sp.. Nevertheless, other studies conducted to investigate the role of PGPR in plant metal uptake have also shown the opposite effect for other metals. Sheng et al. (2012) have found a significant increase of total Cu content in roots and shoots of Z. mays plants grown in a Cu-contaminated soil (total Cu of 1,068 mg kg−1) induced by Bacillus megaterium JL35. Similarly, rhizosphere inoculation with Bacillus subtilis strain SJ-101 as seed bioinoculant increases Ni accumulation in plant tissues of Brassica juncea var. Pusa Bold (Zaidi et al. 2006). In sunflower, Marques et al. (2013) have shown that the PGPR bacteria used in the present study have no effect on shoot metal accumulation while showing different effects on roots. Strain B1 has no significant influence and B2 shows ability to promote a decrease on root’s Cd accumulation, unlike what was observed in this report with maize. A good indicator of metal accumulation capacity is the BCF, as it takes into account the total Cd concentration in the soil to which the plant is exposed. In the present study, all plants showed BCF values higher than 1.0 in roots; generally, for Z. mays plant shoots, the values were lower. Root bioconcentration was significantly affected by bacterial inoculation (with the exception of plants grown in 20 mg Cd kg−1treated soils), and shoot BCF was positively related with soil Cd concentration and negatively related with bacterial inoculation, with both B1 and B2 decreasing BCF. This comes to support the above proposition that these PGPR provided some protective effect against Cd accumulation in maize shoots. By the same token, translocation factor evaluates the plant ability of transferring the metal from roots to shoots. In the present study, for B1- and B2-treated plants, translocation increased with increasing soil Cd concentration. The results indicate that Cd was not easily translocated in Z. mays plants, as the values shown for all the treatments were lower than 1 (Fig. 2c). Similar results have been obtained for H. annuus and for Glycine max grown in Cd-treated substrates (50 up to 200 mg Cd kg−1) for 28 days (Shi and Cai 2009). An (2004) has also reported a significant Cd immobilization by the sweet corn roots exposed to increasing Cd soil concentrations (40, 80, 160, 320, and 640 mg Cd kg−1 soil DW). Additionally, when compared with the respective controls, TF values at all soil Cd levels were decreased by inoculation with both bacterial strains. These results, coupled with the high BCF shown for the root tissues, did indeed indicate that maize plants inoculated with these PGPR might have a potential use for

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Cd phytostabilization and biomass production. According to McKendry (2002), a lower shoot accumulation of the contaminant metal is an important characteristic for an energy crop intended to be used for phytomanagement. Additionally, a high biomass production and low nutrient requirements should be present in energy crop characteristics, grown in polluted areas. The variant of maize used in this study (Aveline) is a hybrid that grows up to 3.2 m in height and is known for its high productivity, thus having the potential to be used for this purpose. This variant coupled with bacterial inoculation may improve Cd short-term phytostabilization abilities of maize in the field. Metal bioavailability is the extent of the Cd available for the plant uptake (Kirkham 2006). Total Cd content of soil is not sufficient to assess its real ecotoxicological action to plants and microorganisms, since heavy metals are mostly present in non-mobile forms (Violante et al. 2010). Therefore, the determination of mobile forms is important to perceive the extension of the availability of metals to plants. Soil extraction with NH4-Ac allows access to the exchangeable metal ions, retained by weak electrostatic interaction, while EDTA extraction promotes the release not only of the exchangeable metal ions but also of the Cd bound to carbonates and organic complexes (Podlesakova et al. 2001). Previous studies have reported that PGPR can significantly promote heavy metal bioavailability (Marques et al. 2013; Sheng and Xia 2006; Sheng et al. 2012), which was observed in the present work, as inoculated soils had generally increased available Cd levels (Table 2). The increase of the Cd bioavailability enabled the metal root uptake by the maize plants (Table 1 and Fig. 2b), although the metal accumulation in shoots was reduced (Table 2 and Fig. 2a)—which could be explained by the bacteria biosorption and accumulation capacities that may reduce metal toxicity for the plants’ shoot (Kavamura and Esposito 2010; Violante et al. 2010). This behavior increases the phytostabilization potential (defined as the metal immobilization within the roots and root zone of plants and the consequent prevention of metal dispersion by establishment of a green cover (EUGRIS 2014)) of the studied associations and renders their application for energy production purposes a stronger possibility. From the DGGE patterns, it was observed that variations in the Cd soil concentration affected soil bacterial community structure, with lower diversity in contaminated soils, as already reported in other studies (Frostegård et al. 1996; Kozdrój 1995; Lorenz et al. 2006; Marques et al. 2013). This may be explained by the metal toxicity (Bruins et al. 2000), which may induce the establishment of a tolerant but less rich community (Frostegård et al. 1996; Khan 2005). A general decrease in the number of DGGE bands of the rhizosphere samples from the beginning to the end of the experiment was also noticed, with the initial community being more similar within samples from the same Cd concentration, possibly

denoting a longer term effect of metal exposure. Canonical correspondence analysis also highlighted that the variations between the communities of the samples collected from the different soil treatments at the end of the experiment were globally driven by the increasing cadmium soil concentration. Cadmium shoot and root accumulation was higher in plants grown in 30 mg Cd kg−1 contaminated soil while biomass was higher for plants collected from non-spiked soil. Time may have also interfered in the shift of the microbial community as already reported in other studies (Khan et al. 2010; Sheng et al. 2012). Additionally, the complex interaction of the rhizosphere-established community and the nutrient requirements to the development of the maize plants may have reduced the availability of nutrients supplied to the bacterial community (Khan et al. 2010). There are several reports of increasing phytoremediation abilities for plants inoculated with PGPR (Glick 2010; Naees et al. 2011; Saharan and Nehra 2011). However, bacteria may have different effects depending on the plant species and on the type of metal contamination. Marques et al. (2013) have shown different outcomes for the same bacteria with sunflower grown in Cdcontaminated soils. Similar results have been reported by Li and Ramakrishna (2011) in copper-contaminated soil where a Pseudomonas strain stimulates the growth of maize but has no significant effect on the increase of sunflower shoot biomass. Grandlic et al. (2008) have also shown that some strains of PGPR increase significantly the growth of buffalo grass (Buchloe dactyloides) in a 0 % compost treatment but have no significant effect when tested on quailbush (Atriplex lentiformis). On the other hand, in 10 % compost treatment, these bacteria have no significant effect on buffalo grass while other strains increase significantly quailbush biomass. The effect of PGPR appears to be plant specific, and further investigation is needed to understand the mechanisms and environmental conditions through which these bacteria may activate their beneficial traits to plants and therefore improve their feasibility to assist phytoremediation.

Conclusions Plant growth promotion abilities of R. eutropha and C. humi were shown in the present study, and thus, these metalresistant bacteria may improve Cd short-term phytostabilization abilities of Z. mays. The maize plants can stabilize the soil during its cycle of growth, as metals are mainly retained in the root area. The increase in shoot biomass observed for inoculated plants grown in the Cd-contaminated soils potentiates the application of the produced biomass for energy purposes after harvest of the aboveground tissues. The roots will remain in the soil, adding organic matter and nutrients to it, which can help further immobilizing the metals when a new crop round is installed. Simultaneously, an even

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with low translocation to the shoots, small amounts of the metal would be extracted from the soil, rendering this soil less polluted and nutrient depleted. The subsequent green covers established at every maize cycle of growth would also prevent erosion and the consequent dispersion of the metal. Acknowledgments This work was supported by Fundação para a Ciência e a Tecnologia and Fundo Social Europeu (III Quadro Comunitário de apoio), research grants to Helena Moreira (SFRH/BD/ 64584/2009), Ana Marques (SFRH/BPD/34585/2007), and Albina Franco (SFRH/BD/47722/2008), and by national funds through FCT— Fundação para a Ciência e Tecnologia under the project PEst-OE/EQB/ LA0016/2011.

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