SOILS, SEC 3 & REMEDIATION ANDMANAGEMENT OF CONTAMINATED OR DEGRADED LANDS & RESEARCH ARTICLE

Metal removal from and microbial property improvementof a multiple heavy metals contaminated soil by phytoextractionwith a cadmium hyperaccumulator Sedum alfredii H.

Wenhao Yang & Taoxiang Zhang & Siliang Li & Wuzhong Ni

Received: 25 October 2013 /Accepted: 18 February 2014# Springer-Verlag Berlin Heidelberg 2014

AbstractPurpose Effects of phytoextraction by Sedum alfredii H., anative cadmium hyperaccumulator, on metal removal fromand microbial property improvement of a multiple heavymetals contaminated soil were studied under greenhouseconditions.Materials and methods A rhizobox experiment with an an-cient silver-mining ecotype of S. alfredii natively growing inZhejiang Province, China, was conducted for remediation of amultiple heavy metals contaminated soil. The rhizobox wasdesigned combining the root-shaking method for the separa-tion of rhizospheric vs near-rhizospheric soils andprestratifying method for separation of sublayers rhizosphericsoils (0–10mm from the root) and bulk soil (>10 mm from theroot). Soil and plant samplings were carried out after 3 and6 months of plant growth.Results and discussion Cadmium (Cd), zinc (Zn), and lead(Pb) concentrations in shoots were 440.6, 11,893, and91.2 mg kg−1 after 6 months growth, and Cd, Zn, and Pbremoved in the shoots were 0.862, 25.20, and 0.117 mg/plant.Microbial biomass C, basal respiration, urease, acid phospha-tase, and invertase activities of the rhizospheric soils weresignificantly higher than that of unplanted soils after 6 monthsgrowth. Microbial biomass carbon (MBC) of 0–2 mm andbasal respiration (BR) rate of 0–8 mm sublayer rhizosphericsoils were significantly higher than that of bulk soil after6 months growth. So were the three enzyme activities of 0–4 mm sublayer rhizospheric soils. BR rate and urease weresignificantly negatively correlated with soluble Cd, so were

MBC, acid phosphatase, and intervase activities with solubleZn, MBC, BR rate, and three enzyme activities with solublePb.Conclusions Harvesting shoots of S. alfredii could removeremarkable amounts of Cd, Zn, Pb, and lower water-solubleCd, Zn, and Pb concentrations in the rhizospheric soils. MBC,BR rate, and enzyme activities of the metal polluted soil,especial ly the rhizospheric soi ls increased withphytoextraction process, which is attributed to the stimulationof soil microbes by planting as well as the decrease in soil-soluble metal concentration.

Keywords Basal respirationrate .Enzymeactivity .Microbialbiomass carbon . Phytoextraction . Rhizospheric . SedumalfrediiH.

1 Introduction

Soil heavy metal contamination has become a widespreadenvironmental problem. Inmany cases, soils are contaminatedwith more than one heavy metal (Keltjens and Van Beusichem1998), which is multiple heavy metals contamination (Zhouet al. 2007; Moreno-Jiménez et al. 2009). Excessive heavymetals are toxic to organisms including soil microorganisms,especially under multiple contamination (Khan et al. 2010;Pan and Yu 2011; Hu et al. 2013). Adverse effects of heavymetal contamination on soil microorganisms, which play cen-tral roles in organic matter decomposition and nutrient cy-cling, can cause the degradation of soil quality (Belyaeva et al.2005; Zhang et al. 2010). With the increasing emphasis onsustainable fertility and environmental benefits, alleviating theeffects of heavy metals contamination on soil microbes andrestoring the microbial properties of contaminated soils areextremely important.

Responsible editor: Peter Schroeder

W. Yang : T. Zhang : S. Li :W. Ni (*)College of Environmental and Resource Sciences, MOE KeyLaboratory of Environment Remediation and Ecosystem Health,Zhejiang University, Hangzhou 310058, People’s Republic of Chinae-mail: wzni@zju.edu.cn

J Soils SedimentsDOI 10.1007/s11368-014-0875-7

Chemical, physical, and phytological processes have beendeveloped for the remediation of metal contaminated soils.Phytoextraction, removing metals by accumulating in plantshoots or harvested organs with moderate impact on the soiland environment, is particularly suggested as the other alter-natives are expensive and invasive (Vangronsveld et al. 2009).Identifying the hyperaccumulators, which can accumulatemetals to a certain degree of concentration in their shoots, iscritical for the success of phytoextraction. Baker and Brooks(1989) have proposed that hyperaccumulators should havecapabilities of accumulating more than 1,000 mg/kg for As,Pb, Cu, and Ni, 10,000mg/kg for Zn, and 100mg/kg for Cd intheir shoots. Although more than 500 kinds of plants havebeen cited in the literature as hyperaccumulators, few plantspecies have been utilized for phytoextraction in practice sofar. For efficient phytoextraction, plants need to grow vigor-ously and healthy, which is not so straightforward in pollutedsoils (Fellet et al. 2007). Except soil properties, local climateconditions also have an influence on phytoextraction efficien-cy. Growing native plants for phytoextraction is a usefuloption, as these plants are often better in terms of survival,growth, and reproduction under the practical environmentthan that introduced from other climate regions (Yoon et al.2006).

Although evaluating the capacity of hyperaccumulators touptake heavy metal is essential, the assessment of thephytoextraction on soil properties is critical as well. The goalof phytoextraction is not only to remove heavy metals fromcontaminated soils but, more importantly, to restore the ca-pacity of soils to perform or function according to their po-tential (Hernandez-Allica et al. 2006). Soil microbial proper-ties such as soil microbial biomass, basal respiration, andenzyme activities have been used for assessing soil quality(Mora et al. 2005; Clemente et al. 2007; Jusselme et al. 2013)and the degree of restoration in metal contaminated soils(Jiang et al. 2010; Hernandez-Allica et al. 2006). The rhizo-sphere is a dynamic region surrounding root where multipleinteracting processes take place (Darrah et al. 2006) and is amicrobiosphere with different chemical, physical, and biolog-ical properties vs bulk soil (Zhang et al. 2012). Rhizosphere-associated microbial processes are of special interest in thephytoremediation since soil microorganisms exert profoundinfluence on soil fertility. Several reports have indicated thatrhizosphere microbial process was an important factor invegetation reestablishment and soil restoration (Garcia et al.2005; Sinha et al. 2009; Zhang et al. 2012). Rhizosphere soilis commonly separated from plant root surface or segment bygentle shaking the roots (Ortas 1997), and this root-shakingmethod was widely used to study the rhizosphere characteris-tics of hyperaccumulators (Gonzaga et al. 2009; Gremionet al. 2004; Wenzel et al. 2003). Rhizosphere is not restrictedto a specific volume but rather characterized by gradients ofproperties, from the root surface to the surrounding bulk soil

(Aviani et al. 2006). Rhizobox is a useful tool to study thegradients of rhizosphere, especially the stratified rhizoboxwhich separates the soils from different distances of the root.Description of chemical gradients in the rhizosphere was thepurpose of a large number of studies (Puschenreiter et al.2005; Kim et al. 2010a). However, the microbial gradientsin the rhizosphere of the hyperaccumulators being used forphytoextraction of contaminated soil, especially multiplemetals contaminated soil, were much less investigated.

The ancient silver-mining (ASM) ecotype of Sedumalfredii Hance, as a perennial plant natively growing inZhejiang Province, China, was identified as a Cdhyperaccumulator in our previous study (Yang et al. 2013).Using stratified rhizobox, combining the root-shaking methodand the stratified method, the objectives of this study were to(1) assess the capacity of S. alfredii to uptake Cd, Zn, and Pbfrom soil and (2) investigate the effects of phytoextraction byS. alfredii on microbial properties (microbial biomass carbon,basal respiration, and enzyme activities) of a Cd, Zn, and Pbmultiple-contaminated soil, especially the rhizospheric soils.The relationships between soil microbial properties and solu-ble metal concentrations in rhizosphere were also discussed.

2 Materials and methods

2.1 Soil and plant preparation

The experimental soil was collected from a mining site inZhejiang Province, Southeast China. After collection, soilsamples were air-dried, ground, and sieved <1 mm. Thegeneral properties of the soil were as follows: pH 6.6; totalC, 17.4 g kg−1; total N, 1.55 g kg−1; total P, 2.8 g kg−1;hydrolyzable N, 80.1 mg kg−1; available P, 22.46 mg kg−1;total Cd, 73.9 mg kg−1; total Zn, 9,055 mg kg−1; and total Pb,18,447.9 mg kg−1. The shoots of S. alfredii H. were obtainedfrom an ancient silver-mining site in Zhejiang Province. Theequal-sized plant shoots were selected and re-rooted in thegreenhouse using a basic nutrient solution: 2.00 mmol L−1

Ca(NO3)2 4H2O, 0.10 mmol L−1 KH2PO4, 0.50 mmol L−1

MgSO4 7H2O, 0.10 mmol L−1 KCl, 0.70 mmol L−1 K2SO4,10.0 μmol L−1 H3BO3, 0.50 μmol L−1 MnSO4 H2O,1.0 μmol L−1 ZnSO4 7H2O, 0.20 μmol L−1 CuSO45 H2O,0.01 μmol L−1 (NH4)6MoO24, and 100 μmol L−1 Fe–EDTA.Solution pHwas adjusted daily to 5.8 with 0.1 mol L−1 HCl or0.1 mol L−1 NaOH. The nutrient solution was aerated contin-uously and replaced every 4 days.

2.2 The structure of the specially prestratified rhizobox

The specially prestratified rhizobox has been designed andused in our former work (Yang et al. 2013). The rhizobox wasvertically divided into two compartments, i.e., the upper zone

J Soils Sediments

and the lower zone using nylon net (300 meshes) (Fig. 1). Theupper zone, into which plant roots were allowed to penetrate,was with the height of 30 mm. The nylon net between theupper zone and lower zone prevented roots from entering intothe lower zone, and a horizontal root mat was formed alongthis nylon net. The soils in the upper zone were separated intorhizospheric and near-rhizospheric soils by shaking roots. Thelower zone (50 mm) was divided into six sublayers by theprestratified nylon net, e.g., 0–2, 2–4, 4–6, 6–8, 8–10, and>10 mm from the root mat. Different layers were stratified bynylon net (300 meshes) fixed on a stainless steel ring (diam-eter, 170 mm; thickness, 2 mm). The soils from 0 to 10 mm ofthe root mat were treated as sublayer rhizospheric soils and the>10 mm soil was regarded as the bulk soil. This rhizoboxcombined the shaking method for the separation ofrhizospheric vs near-rhizospheric soils and the prestratifyingmethod for the separation of sublayers of the rhizosphericsoils and bulk soil as the root mat method.

2.3 Rhizobox experiment

The rhizobox experiment had a completely randomized de-sign that had two treatments such as unplanted (control) andplanted in the rhizobox. Each treatment had six pots at thebeginning of the experiment (three for 3-month samples andthree replicates for 6-month samples).The rhizosphere effectswere studied in the upper zone and lower zone separately. Therhizospheric soil and near-rhizospheric soils sampled in theupper zone and were compared with the unplanted control. Inthe lower zone, the microbial properties of soils in the different

layers were determined to assess the intensity of plant growtheffects.

For the planted treatments, four re-rooted S. alfredii seed-lings were transplanted into each rhizobox. The rhizoboxexperiment was conducted under greenhouse conditions.The average temperature was 20/25 °C night/day, and relativehumidity was about 70 %. Soil moisture content was adjustedto 60–70 % of water holding capacity by weighing therhizobox weight and adding deionized water needed. Duringthe experiment, two samplings of plant and soil were carriedout. The first sampling was done 3 months after the transplan-tation of the plants with three replications, and the secondsampling was done 6 months after the transplantation of theplants with three replications as well. In the first sampling, itwas noted that the roots formed a mat on the nylon mesh in theplant-grown compartment but did not pass through it.Harvesting involved the sequential dismantling of eachrhizobox, separating the layers of each soil zone of therhizobox and removing the plants from the plant-grown com-partment. In each time, unplanted soil samples were alsocollected with three replications. The soils in the upper zonewere separated into rhizospheric and near-rhizospheric soils.According to the method reported by some studies for thesampling of rhizospheric soils (Garcia et al. 2005; Sinha et al.2009), the soil strongly adhering to roots and within the spaceexplored by roots was considered as rhizospheric soil in ourresearch. According to the method for the sampling of near-rhizospheric soil (Huang et al. 2007; Yang et al. 2013), theother proportions of soils in the upper zone were treated asnear-rhizospheric soils. The soils from 0 to 10 mm distancesof the root plane and the bulk soil (>10 mm from the root

Upper zone soil

Bulk soil

30mm

10mm

Root mat

40mm

Sub-layer

rhizospheric soils

170mm

Nylon net (300 mesh)

Stainless Steel Ring

(diameter: 170 mm; thickness:2 mm)

Nylon net

Fig. 1 Diagram of the rhizobox.The lower zone is divided into sixsublayers by the nylon net, e.g.,0–2, 2–4, 4–6, 6–8, 8–10, >10(bulk soil) mm from the root mat

J Soils Sediments

plane) were sampled following the prestratified sublayers. Thesoil samples from each layer were homogenized separatelybefore analysis. After sampling, one portion of fresh soilsamples was passed through a 2-mm sieve, sealed in a plasticbag, and stored at 4 °C with its real moisture status, andmicrobial analysis was carried out within 2 weeks after sam-pling (Mora et al. 2005). For analysis of water-soluble heavymetals, soils were air-dried and sieved through 1 mm. At eachsampling, the plants were harvested. Roots and shoots weremanually separated from soils and washed with deionizedwater, and then roots and shoots were oven-dried at 70 °Cfor 48 h to obtain dry weights. The dried plant samples wereground into fine powder using a mortar mill for heavy metalanalysis.

2.4 Soil, plant, and microbial analysis

The soil chemical properties were analyzed according to themethods described by Lu (2000). The plant tissues weredigested with HNO3/HClO4 (4:1, v/v). Briefly, the groundplant samples (0.25 g) were placed in the digestion tube andmixed with 4 mL HNO3 and 1 mL HClO4. Then, they werecovered with glass stopper and left overnight at room temper-ature. The tubes were then placed on a temperature-controlleddigestion block and heated until the solution became clear.Finally, digested material was dissolved with a 0.5 mol/LHNO3 solution into a 50-mL volumetric flask and broughtup to volume with the same solution. The water-soluble heavymetal was extracted according to the method described in theliterature (Séguin et al. 2004). A soil/water ratio of 1:5 (m/v)was used to obtain sufficient solution for analysis. After 2 hequilibration on the shaker, the sample solutions were centri-fuged (4,000×g, 10 min), and the supernatant was passedthrough 0.45-μm membrane filters. The concentrations ofCd, Zn, and Pb in the digestive and extractive solutions weredetermined using atomic adsorption spectrophotometer.

Soil microbial biomass carbon was measured by the fumi-gation–extraction method (Vance et al. 1987). Briefly, two soilsamples, one was fumigated for 24 h at 28 °C in the dark withchloroform, and the other was incubated without fumigated.The fumigated and nonfumigated samples were extracted with50 mL of 0.5 mol L−1 K2SO4. The concentration of organic Cin the extractant was determined using an automated totalorganic carbon analyzer. Microbial biomass C was calculatedas the difference between organic C extracted from fumigatedand nonfumigated soil extracts.

Soil basal respiration rate was determined bymeasuring theCO2 evolved during 24 h (Vogeler et al. 2008). Briefly, soilwas placed in a hermetically sealed glass bottle with a vialcontaining 0.1 mol L−1 NaOH in the dark at 22 °C. Afterincubation for 24 h, the vial was taken out, and the remainingNaOH was titrated with 0.05 mol L−1 HCl. Soil basal

respiration rate was expressed as milligrams CO2–C per kilo-gram per hour.

Soil urease activity was determined with a method accord-ing to previous literature (Cang et al. 2009). Fresh soil samplewas mixed with 10 % urea (with citric acid buffer at pH 6.7).The soil solution was incubated at 37 °C for 24 h in the dark.The released ammonium was quantified colorimetrically in aspectrophotometer at 578 nm. Soil urease activity wasexpressed as micrograms ammonia released per gram perhour. For the measurement of acidic phosphatase activity,fresh soil was mixed with substrate (disodium phenyl phos-phate solution, pH5.0). The soil solution was incubated at37 °C for 2 h in the dark. After incubation, the suspensionwas filtered and treated with 4-aminoantipyrine and potassiumferricyanide for colorimetric analysis. The phenol releasedwas determined colorimetrically in a spectrophotometer at510 nm (Cang et al. 2009). Phosphatase activity wasexpressed as micrograms phenol released per gram per hour.Invertase activity was assayed with the method as describedby Guan (1986). Fresh soil was incubated at 37 °C for 24 hwith 8 % sucrose (with phosphate buffer at pH 5.5). Theglucose released by invertase reacted with 3,5-dinitrosalicylicacid, and then was measured at 508 nm. Results wereexpressed as milligrams glucose released per gram per 24 h.

2.5 Statistical analysis and data treatment

All statistical analyses were carried on the software SPSS forWindows. Multiple comparisons of significant differencesbetween means were made using the Duncan’s test using alevel of significance of P<0.05. The relationships between thesoil microbial properties and water-soluble Cd, Zn, and Pbwere explored using Pearson’s correlation coefficients. Basedon metal concentrations in shoots, roots, and soils,bioconcentration factor (BF=shoot metal concentration/soilmetal concentration) and translocation factor (TF=shoot met-al concentration/root metal concentration) were calculated.

3 Results

3.1 Plant growth and heavy metal uptake

During the experiment, the ancient silver mining ecotype ofS. alfredii had a continuous growth and biomass of shoots androots were significantly (P<0.05) higher after 6 monthsgrowth than that of the 3 months growth (Table 1). The Cdconcentrations in shoots were 477.6 and 440.6 mg kg−1 aftergrowing for 3 and 6 months, respectively; Zn were 12,642 and11,893 mg kg−1; and Pb were 302.2 and 91.2 mg kg−1

(Table 1). The BFs achieved by S. alfredii for Cd, Zn, andPb were 6.46, 1.40, and 0.016 after 3 months growth, andwere 5.96, 1.31, and 0.005 after 6 months growth (Table 2).

J Soils Sediments

The TF achieved by S. alfredii for Cd, Zn, and Pb were 4.80,2.12, and 0.100 after 3 months growth, and were 5.65, 2.15,and 0.041 after 6 months growth (Table 2). After 3 monthsgrowth, 0.174 mg of Cd, 4.60 mg of Zn, and 0.105 mg of Pbwere accumulated in the shoots per plant. After 6 monthsgrowth, 0.862 mg of Cd, 25.20 mg of Zn, and 0.117 mg ofPb were accumulated in the shoots per plant, which weresignificantly (P<0.05) larger than that of 3 months growth(Table 1).

3.2 Effects of S. alfredii growth on concentrations of soilwater-soluble heavy metals

In the upper zone, the water-soluble Cd, Zn, and Pb concen-trations of the near-rhizospheric soil were significantly(P<0.05) higher than that of the unplanted soils after growingfor 3 and 6 months (Fig. 2). The water-soluble Cd, Zn, and Pbconcentrations of the rhizospheric soil were significantly(P<0.05) lower than that of the near-rhizospheric andunplanted soils after 6 months growth (Fig. 2). After 6 monthsgrowth, the water-soluble Zn concentration of the rhizosphericsoil were significantly (P<0.05) lower than that of 3 monthgrowth (Fig. 2). In the lower zone, the results of theprestratified rhizobox showed that the water-soluble Cd andZn concentrations of 0–6 mm and the water-soluble Pb con-centrations of 0–4 mm sublayer rhizospheric soils were sig-nificantly (P<0.05) lower than that of the bulk soil after3 months growth (Table 3). After 6 months growth, thewater-soluble Zn concentration of the 0–2 mm sublayerrhizospheric soil was significantly (P<0.05) lower than that

of other sublayer rhizospheric soils and the bulk soil. Thewater-soluble Pb concentrations of the 0–4 mm sublayerrhizospheric soils were significantly (P <0.05) lower than thatof the bulk soil (Table 3). After 6 months growth, the water-soluble Zn concentrations of the sublayer rhizospheric soils(0–10 mm) were significantly (P<0.05) lower than that ofafter 3 months growth (Table 3).

3.3 Effects of S. alfredii growth on soil microbial biomasscarbon and basal respiration rate

In the upper zone, soil microbial biomass carbon (MBC)contents of the rhizospheric and near-rhizospheric soils weresignificantly (P<0.05) higher than that of the unplanted soilafter transplanting S. alfredii for 3 and 6 months. And theMBC content of the rhizospheric soil was significantly(P<0.05) higher than that of the near-rhizospheric soil after6 months growth (Fig. 3). Soil MBC content of therhizospheric soil after transplanting S. alfredii for 6 monthswas significantly (P<0.05) higher than that of 3 monthsgrowth (Fig. 3). In the lower zone, the MBC contents of the0–2 and 2–4 mm sublayer rhizospheric soils were significant-ly (P <0.05) higher than that of the bulk soil after 3 monthsgrowth (Table 4). After 6 months growth, the MBC content ofthe 0–2 mm sublayer rhizospheric soil was significantly(P<0.05) higher than that of 6–8 and 8–10 mm sublayerrhizospheric and bulk soils (Table 4). The MBC content ofthe 0–2 mm soil after 6 months growth was significantly(P<0.05) higher than that of 3 months growth (Table 4).

Table 1 Shoot and root biomass, metal concentrations, and accumulations of S. alfredii after 3 and 6 months growth

Growth time Dry weight Heavy metal concentration (mg kg−1) Heavy metal accumulation

(g/pot) Cd Zn Pb Cd (mg/plant) Zn (mg/plant) Pb (mg/plant)

Shoot

3 months 1.46±0.12 b 477.6±28.8a 12,642±410a 302.2±3.7a 0.174±0.015b 4.60±0.24b 0.105±0.007a

6 months 7.37±0.63 a 440.6±32.5a 11,893±1106a 91.2±6.4b 0.862±0.027a 25.20±0.05a 0.117±0.001a

Root

3 months 0.31±0.02 b 99.8±3.5a 5,991±322a 3,027±97a 0.008±0.001b 0.47±0.05b 0.219±0.003b

6 months 0.75±0.01 a 78.1±2.6b 5,518±317a 2,187±108b 0.015±0.001a 1.03±0.07a 0.409±0.024a

Mean value (n=3)±standard error. Data followed by different letters are significantly different at P<0.05

Table 2 BF and TF of S. alfredii for Cd, Zn, and Pb after 3 and 6 months growth

Growth time/months Bioconcentration factor Translocation factor

Cd Zn Pb Cd Zn Pb

3 6.46±0.39a 1.40±0.05a 0.016±0.000a 4.80±0.31a 2.12±0.08a 0.100±0.001a

6 5.96±0.44a 1.31±0.12a 0.005±0.001b 5.65±0.43a 2.15±0.08a 0.041±0.015b

Mean value (n=3)±standard error. Data followed by different letters are significantly different at P<0.05

J Soils Sediments

The basal respiration (BR) rates of the rhizospheric andnear-rhizospheric soils were significantly (P<0.05) higherthan that of the unplanted soil after transplanting S.alfredii

for 3 and 6 months (Fig. 4). The BR rate of the rhizosphericsoil was significantly (P<0.05) higher than that of the near-rhizospheric soils after 3 and 6months growth. After 6 monthsgrowth, the BR rates of the rhizospheric and near-rhizosphericsoils were significantly (P<0.05) higher that that of 3 monthsgrowth (Fig. 4). In the lower zone, the BR rates of the soilsfrom 0–2 and 2–4 mm of the root mat were significantly(P <0.05) higher than that of the 8–10 mm and the bulk soilafter 6 months growth and that of 4–6 and 6–8 mm were alsosignificantly (P<0.05) higher than that of the bulk soil(Table 4). After 6 months growth, the BR rates of the sublayerrhizospheric soils (0–10 mm) were significantly (P<0.05)higher than that of after 3 months growth (Table 4).

3.4 Effects of S. alfredii growth on soil enzyme activities

In the upper zone, the activities of the three hydrolytic enzymein the rhizospheric soil were significantly (P<0.05) higherthan that in the near-rhizospheric and unplanted soils after 3and 6 months growth (Fig. 5), and the three enzyme activitiesof the near-rhizospheric soil were significantly (P <0.05)higher than that of the unplanted soil after 6 months growth(Fig. 5). The three enzyme activities of the rhizospheric andnear-rhizospheric soils after 6 months growth were signifi-cantly (P<0.05) higher than 3 months growth (Fig. 5). In thelower zone, urease activities of the 0–2 and 2–4 mm sublayerrhizospheric soils were significantly (P<0.05) higher than thatof the bulk soil after 3 months growth (Table 5). After6 months growth, urease, acid phosphatase, and invertaseactivities of the 0–2 and 2–4 mm soils were significantly(P<0.05) higher than that of the bulk soil (Table 5). The acidphosphatase and invertase activities of the 0–2 mm sublayerrhizospheric soil after 6 months growth were significantly(P<0.05) higher than that of after 3 months growth (Table 5).

3.5 Correlationships between soil microbial propertiesand soil water-soluble heavy metals

The correlations between soil microbial properties and water-soluble Cd, Zn, and Pb in the lower zone after 6 monthsgrowth were shown in Table 6. The results showed that soilBR rate and urease activity were significantly (P<0.05) neg-atively correlated with water-soluble Cd content. Soil MBCcontent, acid phosphatase, and intervase activities were sig-nificantly (P<0.05) negatively correlated with water-solubleZn content. All of the soil microbial properties measured weresignificantly (P<0.05) negatively correlated with water-soluble Pb content. The results of correlation analysis alsoshowed that acid phosphatase and intervase activities weresignificantly (P<0.05) positively correlated with soil MBCcontent. Urease, acid phosphatase, and intervase activitieswere significantly (P<0.05) positively correlated with soilBR rate.

0

0.03

0.06

0.09

0.12

3 months 6 months

Sampling time

Wat

er s

olu

ble

Cd

(mg

kg-1

)Unplanted soil Near-rhizospheric soil Rhizospheric soil

b

b

a

b

a

c

0

1

2

3

4

5

3 months 6 months

Sampling time

Wat

er s

olu

ble

Zn

(mg

kg-1

)

Unplanted soil Near-rhizospheric soil Rhizospheric soil

b

b*

a

b

a

c

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

3 months 6 months

Sampling time

Wat

er s

olu

ble

Pb

(mg

kg-1

)

Unplanted soil Near-rhizospheric soil Rhizospheric soil

b

b

a

b

a

c

Fig. 2 Water-soluble Cd, Zn, and Pb concentrations of the unplanted,near-rhizospheric, and rhizospheric soils after 3 and 6 months growth ofS. alfredii. Bars represent standard errors (SE). Different letters indicatesignificant differences among soil layers at P<0.05. *P<0.05, significantdifferences between 3 and 6 months growth

J Soils Sediments

4 Discussion

4.1 Plant heavy metal uptake and effects of phytoextractionon soil water-soluble metals

The Cd, Zn, and Pb concentrations of the studied soil exceedthe thresholds that are considered toxic to normal plants (Cd,8 mg kg−1; Zn, 400 mg kg−1; Pb, 400 mg kg−1) (Kabata-Pendias and Pendias 1984). During the experiment, the ASMecotype of S. alfredii Hance grew well and showed no visualsymptoms of phytotoxicity throughout the experiment, show-ing a high tolerance to Cd, Zn, and Pb. Compared with thenotional criterion of hyperaccumulator (Cd, 100 mg kg−1; Pb,1,000 mg kg−1; Zn, 10,000 mg kg−1 in shoot) (Baker andBrooks 1989), Cd and Zn concentrations in shoots of the ASMecotype of S. alfredii reached the hyperaccumulating level. Aplant’s ability to uptake metal from soils and translocate itfrom roots to shoots can be measured using the BF and TF

(Yoon et al. 2006).The BF is more important than shootconcentration when considers the potential of phytoextractionfor a given species (Zhao et al. 2003). Plants exhibiting BFand TF values greater than one are suitable for phytoextraction(Fitz and Wenzel 2002). In this study, the BF and TF ofS. alfredii for Cd and Zn were greater than one, which con-firms the ability of S. alfredii to uptake Cd and Zn from the soiland efficiently translocate to shoots. In addition, S. alfrediiwas able to accumulate certain amounts of Cd and Zn in theirshoots (Table 1). In a similar phytoextraction research,1,483.3 μg/plant of Cd and 24.9 mg/plant of Zn were accu-mulated in Thlaspi caerulescens shoots after 5 months growth(Epelde et al. 2008). Compared to the T. caerulescens, theASM ecotype of S. alfredii showed a remarkable capacity totake up Cd and Zn from polluted soils and accumulate them inthe shoots. According to Zhao et al. (2003), the potential formetal phytoextraction depends on three variables: (i) plantbiomass, (ii) the bioconcentration factor, (iii) the soil massthat requires remediation. It is difficult to predict the numberof years required for phytoextraction to be completed because

Table 3 Water-soluble heavy metals concentrations (mg kg−1) in sublayer rhizospheric soils (0–10mm) and bulk soil(>10 mm) of the lower zone after 3and 6 months growth of S. alfredii

Distances (mm) Cd Zn Pb

3 months 6 months 3 months 6 months 3 months 6 months

0–2 0.029±0.003b 0.029±0.001a 0.482±0.021d* 0.339±0.039b 0.280±0.064b 0.342±0.079b

2–4 0.029±0.001b 0.029±0.001a 0.650±0.003c* 0.491±0.038a 0.354±0.128b 0.388±0.037b

4–6 0.028±0.006b 0.030±0.004a 0.814±0.043b* 0.492±0.028a 0.437±0.063ab 0.504±0.047ab

6–8 0.031±0.005ab 0.032±0.003a 0.868±0.010ab* 0.476±0.048a 0.455±0.006ab 0.461±0.046ab

8–10 0.031±0.003ab 0.035±0.006a 0.864±0.015ab* 0.513±0.018a 0.618±0.061a 0.483±0.026ab

>10 0.041±0.001a 0.043±0.006a 0.950±0.072a* 0.550±0.011a 0.675±0.082a 0.620±0.051a

Mean value (n=3)±standard error. Data followed by different letters are significantly different at P<0.05

*P<0.05

0

50

100

150

200

250

3 months 6 months

Sampling time

Mic

rob

ial c

arb

on(m

g k

g-1)

Unplanted soil Near-rhizospheric soil Rhizospheric soil

b

a a

a*

b

c

Fig. 3 Microbial biomass carbon contents of the unplanted, near-rhizospheric, and rhizospheric soils after 3 and 6 months growth ofS. alfredii. Bars represent standard errors (SE). Different letters indicatesignificant differences among soil layers at P<0.05 level. *P<0.05,significant differences between 3 and 6 months growth

Table 4 Microbial biomass carbon content and basal respiration rate ofthe soils in sublayer rhizospheric soils (0–10mm) and bulk soil (>10mm)of the lower zone after 3 and 6 months growth of S. alfredii

Distances(mm)

Microbial carbon (mg kg−1) Basal respiration(mg CO2–C kg−1 h−1)

3 months 6 months 3 months 6 months

0–2 70.8±5.8ab 151.4±21.4a* 3.67±0.74a 6.25±0.01a*

2–4 79.6±2.2a 112.3±15.6ab 2.84±0.24a 6.46±0.46a*

4–6 62.5±5.2bc 97.7±17.7ab 2.72±0.64a 5.46±0.10ab*

6–8 62.6±2.5bc 77.2±25.4b 3.11±0.15a 5.49±0.37ab*

8–10 61.9±0.5bc 73.6±28.5b 2.56±0.27a 4.49±0.56bc*

>10 55. 5±3.1c 69.9±21.4b 2.80±0.15a 3.95±0.31c*

Mean value (n=3)±standard error. Data followed by different letters aresignificantly different at P<0.05

*P<0.05

J Soils Sediments

conditions in pot experiments, including the volume of soilexplored by the plant roots and the growth conditions, aremuch different from field conditions. However, we have esti-mated the number of years based on the conditions of thepresent experiment. In this study, the amounts of Cd and Znaccumulated in the shoots were 3.45 and 100.80 mg/pot.Based on the metal concentrations and soil mass, the amountsof Cd and Zn in the upper plant zone were about 59.12 and7,244 mg. If we hypothesize that the phytoextraction processare able to remove the same amounts of Cd and Zn in con-secutive harvests; then, it would take 17 and 72 consecutiveharvests (or 8.5 and 36 years assuming two harvests per year)to clean up all the Cd and Zn. Then, under our hypotheticalestimations, the polluted soil in this study could indeed beremediated by the ASM ecotype of S. alfredii in a reasonablenumber of years.

Soluble metals are the most readily available form andmost hazardous for the biota (Séguin et al. 2004). Variationsof soluble metals as affected by plant growth forphytoextraction are considerable. Hyperaccumulators can up-take heavy metals as well as enhance metal solubility via rootexudation (Fitz andWenzel 2002). The concentration of ion insoil depends on the balance between the solubilization fromsoils and uptake by plants (Gonzaga et al. 2009). In this study,the increase in water-soluble metals of the near-rhizosphericsoil was as the results of the balance of metal activatation toplant uptake. This was consistent with previous studies, whichshowed that the soluble metals were enriched due to plantgrowth (Séguin et al. 2004; Puschenreiter et al. 2005;Courchesne et al. 2008). In contrast, water-soluble metalconcentrations of rhizospheric soil were significantly lowerthan that of the unplanted soils. This indicated that the rate ofmetal uptake was higher than the activating rate in the

rhizospheric soil. Kim et al. (2010b) and Liu et al. (2011)reported water-soluble metal depletion in the rhizosphere ofother hyperaccumulators. The results of prestratified rhizoboxmethod showed that soluble metals in 0–6 mm sublayer soilsdecreased significantly. Similar depletion zones have been

0

4

8

12

16

3 months 6 months

Sampling time

Bas

al r

espi

rati

on(m

g C

O2-

C k

g-1h-1

)Unplanted soil Near-rhizospheric soil Rhizospheric soil

a

b

cc

b*

a*

Fig. 4 Basal respiration rates of the unplanted, near-rhizospheric, andrhizospheric soils after 3 and 6 months growth of S. alfredii. Barsrepresent standard errors (SE). Different letters indicate significant differ-ences among soil layers at P<0.05 level. *P<0.05, significant differencesbetween 3 and 6 months growth

0

10

20

30

40

50

3 months 6 months

Sampling time

Ure

ase(

µg

amm

onia

g−

1 h−

1)

Unplanted soil Near-rhizospheric soil Rhizospheric soil

a

a*

b b c

b*

0

200

400

600

800

3 months 6 months

Sampling time

Ph

osp

hat

ase(

µg

ph

enol

g−1

h−1)

Unplanted soil Near-rhizospheric soil Rhizospheric soil

bb

ab*

c

a*

0

10

20

30

40

3 months 6 months

Sampling time

Inve

rtas

e(m

g gl

uco

se g

-1 2

4h-1

)

Unplanted soil Near-rhizospheric soil Rhizospheric soil

c

b

ab*

c

a*

Fig. 5 Urease, acid phosphatase, and invertase activities of theunplanted, near-rhizospheric, and rhizospheric soils after 3 and 6 monthsgrowth of S. alfredii. Bars represent standard errors (SE).Different lettersindicate significant differences among soil layers at P<0.05 level.*P<0.05, significant differences between 3 and 6 months growth

J Soils Sediments

observed in the rhizosphere of many crops for nutrient ionssuch as phosphate, potassium, and nitrate (Hinsinger 1998;Jungk 2002). Fitz et al. (2003) studied arsenic depletion in therhizosphere of Pteris vitata and showed that the arsenic flux(labile pools) was about 55 % reduced in a zone within 3 mmfrom the root surface, followed by a steep gradient to bulk soil.

4.2 Effects of S. alfredii growth on soil microbial properties

Metals have frequently been reported to have adverse effectson soil microbial mass and activities as well as biochemicalprocesses (Belyaeva et al. 2005; Gao et al. 2010; Khan et al.2010). Thus, the recovery of microbial properties ofremediated soils might be a key issue for the success ofremediation (Kumpiene et al. 2009). In this study, S. alfrediigrowth increased soil MBC contents, BR rates, and enzymesactivities, which have key functions on the cycling of C, N,and P in the upper zone. Epelde et al. (2009) and Jiang et al.(2010) reported that phytoextraction could improve soil mi-crobial properties. In our study, these beneficial effects were

more accentuated with the extension of growth time. Rootexudates have been considered as the most important factorfor microbial changes around roots (Bais et al. 2006).Microorganism growth in soil is based on organic carbonsources. Plants can supply organic carbon for the organisms,through litter material, dead roots, and rhizodeposits(Grayston et al. 1996). Therefore, the higher microbial bio-mass and activities in the planted soils can be attributed to theinput of readily decomposable nutrients exudated from plantroot. However, changes of soil microbial properties in heavymetal polluted soils during phytoextraction were due to thecomprehensive effects of metals, plants, and microbes. In ourstudy, the reducing of soluble metals in the rhizospheric soildue to plant growth may also contribute the increase of bio-mass and activity of soil microorganism. Enhancement ofmicrobial activities with the decrease of metal concentrationswas in agreement with previous studies (Mench et al. 2006;Zhang et al. 2008; Jiang et al. 2010). Interestingly, thoughsoluble metals of the near-rhizospheric soils increased, themicrobial activities increased as well. This may due to the

Table 5 Urease, acid phosphatase and invertase activities of the soils in sublayer rhizospheric soils (0–10 mm) and bulk soil(>10 mm) of the lower zoneafter 3 and 6 months growth of S. alfredii

Distances (mm) Urease Phosphatase invertase

(μg ammonia g−1 h−1) (μg phenol g−1 h−1) (mg glucole g−1 24 h−1)

3 months 6 months 3 months 6 months 3 months 6 months

0–2 47.9±3.0a* 34.9±0.8a 48.3±4.1a 74.3±3.5a* 3.20±0.03a 4.66±0.51a*

2–4 38.9±6. 8ab 35.4±1.5a 47.4±3.6a 63.4±2.1ab 4.14±0.07a 4.09±0.29a

4–6 34.9±5.3abc 32.7±3.3ab 46.5±4.1a 49.7±3.9bc 3.51±0.32a 3.06±0.02b

6–8 28.2±7.6bc 30.9±2.5ab 46.5±1.0a 49.8±0.9bc 4.62±0.81a 3.21±0.01b

8–10 25.3±5.0bc 23.3±7.6ab 46.5±1.1a 49.3±6.0bc 4.01±0.80a 2.71±0.01b

>10 17.4±2.4c 18.5±5.7b 41.4±2.2a 43.2±6.9c 4.32±0.95a 2.66±0.02b

Mean value (n=3)±standard error. Data followed by different letters are significantly different at P<0.05

*P<0.05

Table 6 Correlative coefficients (p) amongmicrobial properties and water-soluble Cd, Zn, and Pb of the sublayer rhizospheric soils (0–10mm) and bulksoil (>10 mm) of the lower zone after 6 months growth

Microbialcarbon

Respiration Urease Phosphatase Invertase Cd Zn Pb

Microbial carbon 1.000 0.796 (0.058) 0.750 (0.086) 0.958a (0.003) 0.971a (0.001) −0.656 (0.157) −0.893a (0.017) −0.819a (0.046)Respiration 1.000 0.976a (0.001) 0.828a (0.042) 0.879a (0.021) −0.910a (0.012) −0.680 (0.137) −0.899a (0.015)Urease 1.000 0.742 (0.091) 0.803 (0.054) −0.964a (0.002) −0.662 (0.152) −0.850a (0.032)Phosphatase 1.000 0.979a (0.001) −0.666 (0.149) −0.881a (0.021) −0.915a (0.011)Invertase 1.000 −0.691 (0.128) −0.845a (0.034) −0.875a (0.023)Cd 1.000 0.624 (0.186) 0.850a (0.032)

Zn 1.000 0.812a (0.050)

Pb 1.000

a Correlation is significant at the 0.05 level

J Soils Sediments

promotion of the plant growth is stronger than the inhibition ofthe increased heavy metals on the soil microbial biomass andactivities. Martínez-Iñigo et al. (2009) showed that plantgrowth limited the adverse effects of metals on β-galactosidase and rendered it higher than the original level inthe unpolluted soil. Courchesne et al. (2008) found that bothof water-soluble metals and microbial biomass and enzymaticactivity in the rhizosphere soil were higher than that of bulksoil.

Soil microorganisms are sustained by a small area of soilssurrounding root, which cannot be restricted to a specialvolume but characterized by gradients of properties from theroot surface to the bulk soil (Liu et al. 2012). The results ofprestratified rhizobox method used in our study showed thatsoil microbial properties decreased with increase in the dis-tances from the root mat in general. The distance-dependentchanges of soil microbial properties in the rhizosphere causedby phytoextraction weremore obvious after 6months growth.,which were similar to the previous studies (Xu and Xia 2009;Xie et al. 2012; Yang et al. 2013). The plant influences therhizosphere mainly through the root exudates, but the rootexudates diffuse into the soil where they gradually disappeardue to the increasing radius from root surface and microbialconsumption (Xie et al. 2012). The distance-dependent deple-tion of soil MBC contents, BR rates, and enzymes activities inthe sublayer rhizospheric soils may due to the gradient deple-tion of root exudates.

4.3 Correlationships between soil microbial propertiesand soil water-soluble heavy metals

Metals can affect the growth, morphology, and metabolism ofsoil microorganisms through functional disturbance, proteindenaturation, and destruction of cell membrane integrity(Leita et al. 1995). The negative correlations between soilmicrobial biomass and activities (BR rates and enzymes ac-tivities) and soluble metals found in this study were consistentwith the previous findings. Mora et al. (2005) showed thatmicrobial biomass, β-glucosidase, arylsulphatase, and dehy-drogenase activities were negatively correlated with solubleCd, Cu, and Zn. Negative correlations were also found be-tween soil microbial biomass, phosphatase activity, and water-soluble heavy metals (Wang et al. 2007a, b). Thus, in ourstudy, the reduction of water-soluble heavy metals contributedto the increasing in soil microbial biomass and activities in thesublayer rhizospheric soils. The positive correlations betweensoil enzyme activities and microbial biomass carbon and basalrespiration found in this study indicated that soil microorgan-ism play an important role in retaining soil enzyme activities.Similar results on these relationships can be found in theliteratures (Mora et al. 2005; Vogeler et al. 2008; Yang et al.2013).

5 Conclusions

The ancient silver-mining ecotype of S. alfredii H., nativelygrowing in Zhejiang Province, could hyperaccumulate Cd andZn under the multiple metals polluted soil conditions and beregarded as Cd and Zn hyperaccumulator. Phytoextraction byS alfredii enhanced soil MBC content, BR rate, and activitiesof three soil enzymes, which have key functions on the cy-cling of C, N, and P, especially for the longer plant growthperiod. The improvement of soil microbial properties byphytoextraction was due to the stimulatory effects of plantgrowth on soil microbes and the decrease in soluble metalconcentration. Phytoextraction by the natively growingS. alfredii can be suggested for the remediation of multipleheavy metals polluted soil.

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