Phytotoxicity and bioaccumulation of ZnO nanoparticles in Schoenoplectus tabernaemontani

Chemosphere 120 (2015) 211–219

Contents lists available at ScienceDirect

Chemosphere journal homepage: www.elsevier.com/locate/chemosphere

Phytotoxicity and bioaccumulation of ZnO nanoparticles in Schoenoplectus tabernaemontani Dongqing Zhang a,⇑, Tao Hua a, Fei Xiao b, Chunping Chen b, Richard M. Gersberg c, Yu Liu a, David Stuckey d, Wun Jern Ng a, Soon Keat Tan a a Advanced Environmental Biotechnology Centre, Nanyang Environment & Water Research Institute, School of Civil and Environmental Engineering, Nanyang Technological University, 1 CleanTech Loop, #06-10, Singapore 637141, Singapore b School of Chemical and Biological Engineering, N1.2-B1-03, Nanyang Technological University, Singapore c Graduate School of Public Health, San Diego State University, Hardy Tower 119, 5500 Campanile, San Diego, CA 92182-4162, USA d Department of Chemical Engineering, Imperial College London, London SW7 2AZ, UK

h i g h l i g h t s  S. tabernaemontani showed significant inhibition for ZnO treatment at level of 1000 mg L  The levels of Zn in the roots under ZnO NP treatment were 402–36 513 lg g

1

.

1

.  S. tabernaemontani showed clear potential to accumulate ZnO NPs in the roots.  Translocation of ZnO NPs from root to shoot was limited.  ZnO NPs could penetrate the cell wall and be taken up by plant cell of S. tabernaemontani.

a r t i c l e

i n f o

Article history: Received 18 April 2014 Received in revised form 15 June 2014 Accepted 16 June 2014

Handling Editor: Tamara S. Galloway Keywords: ZnO nanoparticles Bioaccumulation Phytotoxicity Schoenoplectus tabernaemontani

a b s t r a c t The rapid development of nanotechnology will inevitably result in an increasing release of engineered nanoparticles (NPs) to wastewaters. In this study we investigated the fate and toxicity of ZnO NPs in aquatic plant mesocosms, as well as the potential for root accumulation and root-to-shoot translocation of these Zn NPs in the wetland plant Schoenoplectus tabernaemontani exposed to ZnO NPs. The growth of S. tabernaemontani in these hydroponic mesocosms was significantly inhibited by ZnO NPs (1000 mg L 1) compared to a control. Levels of Zn in the plant roots for the ZnO NP treatment ranged from 402 to 36 513 lg g 1, while values ranged from 256 to 9429 lg g 1 (dry weight) for Zn2+ treatment, implying that the uptake of Zn from ZnO NPs was substantially greater than that for Zn2+. The root uptake (of the initial mass of Zn in the solution) for ZnO NP treatment ranged from 8.6% to 43.5%, while for Zn2+ treatment they were 1.66% to 17.44%. The low values of the translocation factor for both ZnO NP (0.001–0.05) and Zn2+ (0.05–0.27) treatments implied that the potential for translocation of Zn NPs from roots to shoots was limited. ZnO NP distribution in the root tissues of S. tabernaemontani was confirmed by scanning electron microscopy (SEM). Transmission electron microscopy (TEM) demonstrated that ZnO NPs could pass through plant cell walls, and were present within the plant cells of S. tabernaemontani. Ó 2014 Elsevier Ltd. All rights reserved.

1. Introduction Nanotechnology, the design and manufacture of nanoscale (<100 nm) materials with unique properties, has opened up a burgeoning new field of applications in consumer products, chemical and medical equipment, information technology and the energy sector (Kümmerer et al., 2011; Lee et al., 2012). Zinc oxide (ZnO) ⇑ Corresponding author. Tel.: +65 6790 6619; fax: +65 6790 6620. E-mail address: [email protected] (D. Zhang). http://dx.doi.org/10.1016/j.chemosphere.2014.06.041 0045-6535/Ó 2014 Elsevier Ltd. All rights reserved.

nanoparticles (NPs) are wide band-gap semiconductors that exhibit near UV emission and transparent conductivity, which makes them particularly attractive in their application as electronic sensors and solar voltaics (Lin and Xing, 2008; Ma et al., 2013). In particular, ZnO NPs have been widely exploited for their photolytic properties and are finding extensive application in personal care products because of their ultraviolet-blocking ability (Hoffmann et al., 1995; Li et al., 2008). NPs can be released into the environment deliberately or accidentally (Stampoulis et al., 2009; Lee et al., 2010). Since NPs often

212

D. Zhang et al. / Chemosphere 120 (2015) 211–219

exhibit physical and chemical properties substantially different from those of their bulk counterparts, and these materials can bind to and transport other compounds within cells or tissues, concern has been growing with regard to their fate and toxicity in the aquatic environment (Peralta-Videa et al., 2011). Results from material flow analyses suggest that a major fraction of the NPs in commercial products may eventually be released into sewer systems, and thus reach wastewater treatment plants (WWTPs) (Kim et al., 2010). Despite the fact that the incorporation of NPs into sewage sludge (to levels of nearly 100 lg L 1) through aggregation and/ or adsorption may account for NP removal from WWTP (Kiser et al., 2009), ZnO concentrations have been estimated to range from 0.34 to 1.42 lg L 1 in WWTP effluents (Gottschalk et al., 2009). Efficient removal of NPs from wastewater is particularly important in view of their possible persistence, and increasing evidence for their ecotoxicity (Limbach et al., 2008). Constructed Wetlands (CWs) are engineered wastewater treatment systems that encompass a plurality of treatment modes including biological, chemical and physical processes, which are all akin to processes occurring in natural treatment wetlands (Kadlec and Knight, 1996). In the recent past, CWs have become a popular option for wastewater treatment because of their high pollutant removal efficiency, easy operation and maintenance, and low cost and energy requirements (Vymazal, 2011). Closely interacting with soil, water and the atmospheric compartments, higher plants may constitute one of the main routes of exposure to NPs in the aquatic environment (Navarro et al., 2008; Ma et al., 2010). Direct uptake and translocation of microcontaminants by plants has been considered as one of the most important mechanisms for phytoremediation (Burken and Schnoor, 1998; Collins et al., 2006). Notwithstanding increased knowledge that has been gained on the human toxicology of NPs, few studies have considered interaction of NPs with plants (Ma et al., 2010; Rico et al., 2011), and very little is known about the mechanisms of biological uptake and accumulation of NPs in plants, and transport between their biological compartments (Ju-Nam and Lead, 2008). In particular, very few plant species have been studied, and the majority of these were edible plants (Nair et al., 2010; Rico et al., 2011), such as ryegrass (Lin and Xing, 2008), zucchini (Stampoulis et al., 2009), lettuce (Lin and Xing, 2007), crop plants (Lee et al., 2012) and wheat (Larue et al., 2012). To date, there have been very few reports assessing uptake and translocation of NPs by higher aquatic plants that are used in treatment wetlands. Therefore, fundamental questions remain on the toxic effects of NPs on higher aquatic plants and the impact of plant species and NP physicochemical properties on the plant uptake potential (Nair et al., 2010; Rico et al., 2011; Ma et al., 2013). The primary objective of this study was to investigate the uptake and bioaccumulation of Zn by the wetland plant Schoenoplectus tabernaemontani exposed to ZnO NPs as well as Zn2+. The specific aims were: (i) to investigate the adverse effects on S. tabernaemontani growth induced by treatment with ZnO NPs and Zn2+; (ii) to assess the potential for bioaccumulation and translocation of Zn NPs by S. tabernaemontani; and (iii) to observe the localization and distribution of ZnO NPs within the plant roots of S. tabernaemontani.

2. Materials and methods 2.1. Preparation of metal salt solution and ZnO NP suspensions Zinc sulfate heptahydrate (99.8% purity) and ZnO NP with a nominal primary particle size of 35 ± 5 nm (99.8% purity) were purchased from Sigma–Aldrich (Singapore). ZnO NP suspensions at concentrations of 0 (control), 10, 100 and 1000 mg L 1 were

prepared by adding the appropriate amounts of ZnO NPs or Zn2+ to Milli-Q water. To avoid aggregation, the ZnO NP suspensions were sonicated for 1 h (VWR 75T Aquasonic sonicator, 30 °C, 100 W, 40 kHz) prior to the use (Lin and Xing, 2007). Small magnetic bars were placed in the suspensions for stirring to avoid aggregation of the particles. After sonication, the ZnO NP suspensions became homogeneously turbid and then were allowed to settle for 5 min. The top (90%) (approximately) of the suspension was drawn and placed in a clean beaker. The bottom portion of the suspension consisting of a few visible particle aggregates was discarded to reduce the presence of aggregates outside of the nanorange. The pH value of the suspension was 6.7 ± 0.1. NPs in the nutrient solution (see below) were extracted using a vacuum probe and filtered by nylon membrane filters (0.2 lm; Whatman). Primary particle size and morphology analysis is confirmed using transmission electron microscopy (TEM, JEOL 3010). Specific surface area was measured with the Brunauer–Emmett– Teller (BET) method. Electrophoretic mobility was converted to zeta potentials using a Zetasizer Nano Z90 (Malvern Instruments, UK). Dissolution kinetics of ZnO NPs were determined. ZnO NPs suspensions at the maximum exposure concentration (1000 mg L 1) were used. At times of 4, 8, 16, 24 and 72 h, the concentration of dissolved Zn in the suspensions, was determined by Zn elemental analysis via inductively coupled plasma optical emission spectroscopy (ICP-OES, Perkin–Elmer, Optima 2100, USA) after centrifugation (10 000 rpm for 30 min). 2.2. Hydroponic culture 0.2 m high S. tabernaemontani was purchased from Uvaria Tide (Singapore), and thoroughly washed to remove any soil particles attached to the plants. A 25% strength Hoagland nutrient solution was prepared with the following composition (in mmol L 1): 0.75 K2SO4, 0.65 MgSO4
7H2O, 2.0 Ca(NO3)2, 0.1 KCl, 0.25 KH2PO4, 1  10 3 H3BO3, 1  10 3 MnSO4
H2O, 1  10 4 CuSO4
5H2O, 5  10 6 (NH4)6Mo7O24, 0.1 Fe-EDTA. All of these chemicals were purchased from Sigma–Aldrich (Singapore) and were of > 98% purity. The plants were then acclimatized to Hoagland nutrient solutions for four weeks. Then the plants with a uniform size were transferred to 2 L mesocosms (4 plants per mesocosm corresponding to the four exposure periods studied, i.e., 3, 7, 14 and 21 d) which contained the modified Hoagland nutrient solution enriched with either ZnO NPs or Zn2+. Two sets of experiments (24 assays) were performed to evaluate the potential for Zn accumulation and translocation in S. tabernaemontani. Treatment 1 was set up to include 9 mesocosms (with three replicates each) spiked with ZnO NPs at Zn levels of 10, 100 and 1000 mg L 1, respectively. In treatment 2, ZnCl2 was spiked into another 9 vessels (with three replicates each) containing nutrient solutions at the same Zn levels of 10, 100 and 1000 mg L 1. Control tests (3 vessels) were also performed where neither ZnO NPs or Zn ions were spiked into the nutrient solutions Additionally, in order to investigate the contribution of Zn2+ released from ZnO NPs to plant phytotoxicity, a set of 3 mesocosms spiked with Zn2+ at 4.0 mg L 1 was included in the experimental design, as the equilibrium concentration of dissolved Zn2+ was measured in the ZnO NP suspension (1000 mg L 1) to be 3.75 mg L 1 (see Section 3.1). Each assay was covered with a plastic sheet with 4 holes through which the plant shoots could extend for exposure to light. The plants were harvested at each of the four exposure periods studied (i.e., 3, 7, 14 and 21 d) for biomass weight and Zn analysis. 2.3. Quantification of zinc in plant tissues At the end of each exposure period, the plants were washed with tap water followed by rinsing with Milli-Q water three times.

D. Zhang et al. / Chemosphere 120 (2015) 211–219

The plant tissues were weighed for wet weight and again after being dried at 70 °C for 24 h. Thereafter, the roots and shoots (0.5 g) were digested in an Anton Paar Microwave Reaction System (Multiwave 3000, Alpha Analytical USA) following the USEPA 3051 method. The concentrations of Zn in the plant tissues were determined using ICP-OES. The instrument was calibrated from 0.1 to 10 ppm in the axial configuration at 250 nm using a zinc reference standard solution. 2.4. Electron microscopy Fresh roots of S. tabernaemontani were thoroughly washed with Milli-Q water. The rootlets were cut and coated with gold for 60 s (ca. 1 nm thickness of gold) by using a sputter coater. ZnO NPs in both suspension and root tissues were observed under a scanning electron microscope (SEM, JEOL-6700). Root samples for TEM analyses were prepared following the procedures below. In brief, S. tabernaemontani root segments (diameter, 1 mm; height, 2 mm) were taken from roots above the apical part of the root tip, and prefixed in 4% glutaraldehyde diluted with 0.1 M cacodylate buffer for 4 h. Then the tissues were rinsed with 0.1 M phosphate buffer (pH 7.5) twice for 10 min, and post-fixed in 1% osmium tetroxide and 0.1 M cacodylate buffer for 1 h. The tissues were then rinsed with 0.1 M cacodylate buffer and dehydrated in a graded ethanol series (50%, 70%, 80%, 90%, and 99.9%). Resin infiltration and embedding were in a graded resin: 30%, 50% and 70% for 2 h and 100% overnight. The tissues were then embedded in beam capsules and polymerized at 60 °C for 72 h; once embedded approximately 150 nm thick (semi-thin) cross sections were cut perpendicularly using a diamond wafer blade. Ultra-thin sections (thickness < 70 nm) were also stained with uranyl acetate (45 min) and lead citrate (3 min). The sections were observed by TEM. 2.5. Statistical analysis The results are presented as mean ± SD (standard deviation) for each concentration. Tests to determine statistical differences between treatments were carried out by comparing the critical value through ANOVA one-way analysis of variance (SPSS Statistics V17.0). Comparisons were considered significantly different at p < 0.05. 3. Results and discussion 3.1. Characterization of ZnO NPs Very little is currently known about the actual level of Zn NPs in both municipal wastewaters and natural aquatic systems. Gottschalk et al. (2009) modeled the predicted release of ZnO NPs to aquatic systems to result in estimated environmental

213

concentrations on the order of approximate 100 lg kg 1 in sediments and soils, and 0.34–1.4 lg L 1 in treated waters. The concentrations used in our experiments were significantly higher than these environmentally relevant levels, because our aim was to investigate both the potential for phytotoxicity and the mechanisms of bioaccumulation and translocation of these NPs in the higher aquatic plant. Moreover, the release of these NPs may continue to increase over time due to their large scale production and use. Size distribution of the ZnO NPs was measured (Fig. S1 in the Supporting Information), and found to be 19–47 nm, with a mean size of 33 ± 8 nm (n = 100), concurring with information provided by the NP producer. Fig. 1 shows a TEM image of the ZnO NPs in nutrient solution; near-spherical individual NPs were observed in the solution (Fig. 1C), and their surface area was found to be 43 m2 g 1. Titration solutions of 0.01 M HCl, 0.01 M NaOH and 0.1 M NaOH were used to determine the zeta potential, which was shown to be 5.4 mV (pH = 6.8) and 2.6 mV (at pH = 6.4) at the beginning and end of the experiment. To determine the concentration of Zn2+ released by ZnO NP suspension, the dissolved Zn2+ level was measured in a ZnO NP suspension (1000 mg L 1) over a period of 3-d. An equilibrium concentration of 3.75 mg L 1 was reached after 24 h, and remained constant for 3 d (Fig. S2 in the SI). 3.2. Effect of ZnO NPs and Zn2+ treatment on plant growth Fig. 2 shows the effect of ZnO NPs and Zn2+ treatments on the growth of S. tabernaemontani. Phytotoxicity exerted by ZnO NPs and Zn2+ was dependent on the Zn concentration as well as exposure time. S. tabernaemontani showed significant (p < 0.05) inhibition compared to the controls when treated by Zn2+ and ZnO NPs at 1000 mg L 1. Zn2+ showed more pronounced effect (the shoots became yellow, withered and dry) of toxicity than did ZnO NPs. After 21-d, at Zn concentrations of 10, 100 and 1000 mg L 1, the S. tabernaemontani was inhibited by 9%, 28% and 54% by Zn2+ ions (as compared to controls), while values were 6%, 13% and 41% for ZnO NPs. Our results are concordant with a recent ZnO investigation by Lin and Xing (2008) who reported a significant decrease in root elongation when Lolium perenne (ryegrass) was exposed to high concentrations of ZnO NPs, with the biomass reduced by 50% after a 12-d exposure to ZnO NPs at a level of 1000 mg L 1. Determination of phytotoxicity of NPs is somewhat complicated, due to the potential dissolution of metallic ions from the NPs along with the potential toxicity of the NPs themselves (Ma et al., 2010). In order to investigate the contribution to the plant toxicity observed from the dissolved ions possibly released from the ZnO NPs, we included a treatment level of 4.0 mg L 1 Zn2+ (the measured level released by ZnO NPs at a concentration of 1000 mg L 1) into our experimental design. Fig. 2 shows that there

Fig. 1. TEM image of ZnO NPs in the nutrient solution. (A) 0.5 lm; (B) 100 nm and (C) 50 nm.

214

D. Zhang et al. / Chemosphere 120 (2015) 211–219

30

Biomass of S. tabernaemontani (g)

10 Zn ions

26

100 Zn ions

a a

1000 Zn ions

24

themselves was apparently not sufficient to cause significant toxicity. Similarly, Lee et al. (2010) investigated the effects of metal oxide NPs on the development of Arabidopsis thaliana, and found that exposure to 400 mg L 1 ZnO NPs (which released 14.6 mg L 1 of soluble Zn) prevented 94% of the seeds from germination, and completely halted root elongation. In contrast, exposure to the equivalent concentration of dissolved Zn (by adding ZnCl2 to a level of 14.6 mg L 1) resulted in significantly lower toxicity. Additionally, Lin and Xing (2008) examined the phytotoxicity of ZnO NPs to ryegrass, and reported that the dissolved Zn in the ZnO NP-treated nutrient solutions was 6 ± 2 mg L 1, which was lower than the toxic threshold of Zn2+ to the ryegrass. This strongly suggested that the phytotoxicity of ZnO NPs could not directly result from the Zn dissolution into the bulk nutrient solution.

A

Control 28

a

4 Zn ions 22 20

ac 18 16 14 12

bc 0

5

10

15

20

25

Days

Biomass of S. tabernaemontani (g)

30

3.3. Accumulation of Zn in plant tissues

B

Control 28

10 Zn NPs

26

100 Zn NPs

24

1000 Zn NPs

3.3.1. Accumulation of Zn in the roots Fig. 3A shows Zn levels (lg g 1 dry weight) in the roots of S. tabernaemontani after ZnO NP and Zn2+ treatment. Statistical analysis shows that there was a significant (p < 0.05) correlation between the Zn level in the roots with ZnO NP treatment, and the external Zn NP concentrations. These levels ranged from 402 to 36 513 lg g 1 (dry weight), while the values were 256–9429 lg g 1 for Zn2+-treated mesocosms. These values suggested that the uptake of Zn from ZnO NPs was substantially greater than for Zn2+. Furthermore, at the external concentrations of 10, 100 and 1000 mg L 1, respectively, the accumulated Zn concentrations in the roots of ZnO NP-treated plants were 2.5, 2.7 and 3.9 times higher than those for Zn2+ treated plants. This finding is in good agreement with that of Lin and Xing (2008) who reported that the concentrations of Zn in the roots under ZnO NP treatment were 3.6 times higher than those from Zn2+ treatment at external concentrations of 1000 mg L 1. A bioaccumulation factor (BAF) is defined as the ratio of the concentration of a chemical in the tissue of an aquatic species to its concentration in water (U.S. EPA, 2000). Table 1 shows that

a a ac ac

4 Zn ions 22 20 18

bc

16 14 12

0

5

10

15

20

25

Days Fig. 2. Effect of Zn2+ treatment (A) and ZnO NP treatment (B) at the concentrations of 10, 100 and 1000 mg L 1, and 4 mg L 1 Zn2+ (A and B) on the biomass of S. tabernaemontani (fresh weight). The values were given as mean ± SD (standard deviation) of three replicates.

was no significant (p > 0.05) difference in plant growth between this treatment of 4.0 mg L 1 Zn2+ and the control, implying that the concentration of dissolved Zn2+ released from ZnO NPs

Zn levels in the roots (µg g-1)

A

7,000

1,000

a

800 600 400

a

a

a

a

d-14

d-7

d-14

ZnO NP treatment

Zn ion treatment

a

25

a

a a

a

100

2,000 1,800 1,600 1,400 1,200 1,000 800 600 400 200 0

a

80

a

20

a

15

60

a

40

a

20

5 0

a a

a a

a a

0 d-3 d-7 ZnO NP treatment

d-14 d-21 Zn ion treatment 2+

b

d-3 d-7 ZnO NP treatment

Zn ion treatment

100 mg L-1

d-3

d-7

ZnO NP treatment

d-14

1

b b

b b a

a

d-7

ZnO NP treatment

Zn ion treatment

d-14 d-21 Zn ion treatment

1000 mg L-1

d-3

d-21

Fig. 3. The concentrations of Zn under ZnO NP and Zn treatment: in the roots; and B: in the shoots (unit: lg g p < 0.05, different letters indicate significant different values between ZnO NP treatment and Zn ion treatment.

b

b

0

d-21

120 10 mg L-1

30

b

10,000 5,000

d-3

d-21

35

10

15,000

0 d-7

ZnO NP treatment

Zn levels in the shoots (µg g-1)

b b

b

1,000 d-3

B

3,000

b

0

a

20,000

2,000

200

25,000

a

a

a

30,000

a

4,000

a

a

1000 mg L-1

35,000

a

5,000

a

a

40,000

a

100 mg L-1

6,000

10 mg L-1

a

a

d-14

d-21

Zn ion treatment

dry weight). For each concentration, ANOVA, significant at

215

D. Zhang et al. / Chemosphere 120 (2015) 211–219

the BAFs for the roots of S. tabernaemontani in the ZnO NP-treated mesocosms ranged from 20.96 to 84.13, while values for the Zn2+ -treated mesocosms ranged from 4.05 to 33.71. All of the BAFs for ZnO NP treatment were higher than those for Zn2+ treatment, implying that S. tabernaemontani had greater potential to take up Zn from ZnO NPs than from Zn2+ treatment. The uptake percentage for Zn (%) is defined as the ratio of the assimilated mass (mg) of Zn in the plant tissues to the initial mass (mg) of Zn in the nutrient solution. Table 2 shows that the uptake percentage (%) of Zn in the plant roots ranged from 8.61% to 43.51% for the ZnO NP-treated plants, while the values were 1.66% to 17.44% for the Zn2+-treated plants. The relatively high range of values for the NP-treated plants shows the potential use of these aquatic plants for phytoremediation of NPs in wastewaters. The variability of biological response to NPs, and the potential for plant uptake of NPs was found to depend largely upon the physicochemical properties of NPs as well as the plant species (Ma et al.,

2010; Nair et al., 2010; Rico et al., 2011). In this present study, the levels of total Zn in the roots of S. tabernaemontani treated by ZnO NPs ranged from 402 to 36,513 lg g 1. In a similar manner, Hernandez-Viezcas et al. (2011) using ICP-OES showed that Zn concentrations in plant (velvet mesquite) tissues treated with ZnO NPs were 2102 and 1135 lg g 1 dry weight in the roots and stems of the plant. On the other hand, Lin and Xing (2008) found that the Zn concentrations measured in the roots of ryegrass (L. perenne) being treated with ZnO NP were only 140 lg g 1. Such differential accumulation of NPs in the plants may also be substantial due to the differences among in plant species, since plants with different xylem structures may demonstrate different uptake kinetics for NPs (Ma et al., 2010). For example, López-Moreno et al. (2010) investigated the fate of CeO2 NPs in different crops and reported that at 4000 mg L 1 of CeO2, the concentrations of Ce varied significantly between plant species (approximately 300 lg g 1 for corn, 400 lg g 1 for soybean, 3000 lg g 1 for tomato, and 6000 lg g 1

Table 1 BAFs in the roots. Concentrations in the solution (mg L

1

BAFsa in the roots

)

Statistical analysisb

Day-3

Day-7

Day-14

Day-21

ZnO NP treatment

10 100 1000

40.24 28.32 20.96

45.84 33.84 27.12

52.72 49.12 31.20

84.13 59.62 36.51

a ac bc

Zn2+ treatment

10 100 1000

25.6 10.09 4.05

28.16 13.47 4.77

29.60 15.06 4.99

33.71 22.17 9.42

bc b b

Note: a Bioaccumulation factors (BAFs) = detected concentrations of Zn in the roots under ZnO NP treatment/initial concentrations of ZnO NPs in the nutrient solution. b Different letters indicate significant difference of BAF at different concentrations between ZnO NP and Zn2+ treatment, significant at p < 0.05 (one-way ANOVA).

Table 2 The assimilated mass (mg) and uptake percentage (%) of Zn in the roots. Concentrations in the solution (mg L 1)

Assimilated mass (mg)a

Statistical analysisc

Uptake percentage (%)b

Day-3

Day-7

Day-14

Day-21

ZnO NP treatment

10 100 1000

2.71 16.01 86.12

2.84 24.17 104.61

2.91 24.30 111.61

Day-3

Day-7

Day-14

Day-21

4.35 27.67 140.63

a a b

27.13 16.01 8.61

28.42 24.17 10.46

29.07 24.30 11.16

43.51 27.67 14.06

a b cd

Zn2+ treatment

10 100 1000

1.73 5.70 16.64

1.75 6.93 18.39

1.63 7.45 17.84

1.74 10.29 36.32

a a a

17.26 5.70 1.66

17.46 6.93 1.84

16.32 7.45 1.78

17.44 10.29 3.63

bd c c

Statistical analysisd

Note: a Assimilated mass of Zn (mg) = root weight (g)  detected concentrations of Zn in roots (lg g 1). b Uptake percentage (%) = assimilated mass of Zn (mg) in the roots/initial mass of ZnO NPs in the nutrient solution (mg)  100. c Different letters indicate significant difference of assimilated mass at different concentrations between ZnO NP and Zn2+ treatment, significant at p < 0.05 (one-way ANOVA). d Different letters indicate significant difference of uptake percentage at different concentrations between ZnO NP and Zn2+ treatment, significant at p < 0.05 (one-way ANOVA).

Table 3 BAFs in the shoots. Concentrations in the solution (mg L

1

)

BAFs in the shootsa

Statistical analysisb

Day-3

Day-7

Day-14

Day-21

ZnO NP treatment

10 100 1000

0.64 0.07 0.02

1.92 0.18 0.08

2.48 0.23 0.10

2.72 0.58 0.11

a b b

Zn2+ treatment

10 100 1000

1.28 0.11 0.30

1.32 0.15 0.79

2.56 0.23 1.43

2.81 0.95 1.71

a b ab

Note: a Bioaccumulation factors (BAFs) = detected concentrations of Zn in the shoots under ZnO NP treatment/initial concentrations of ZnO NPs in the nutrient solution. b Different letters indicate significant difference of BAF at different concentrations between ZnO NP and Zn2+ treatment, significant at p < 0.05 (One-way ANOVA).

216

D. Zhang et al. / Chemosphere 120 (2015) 211–219

Table 4 The assimilated mass (mg) and uptake percentage (%) of Zn in the shoots. Concentrations in the solution (mg L 1)

Assimilated mass (mg)a

Statistical analysisc

Uptake percentage (%)b

Statistical analysisd

Day-3

Day-7

Day-14

Day-21

Day-3

Day-7

Day-14

Day-21

ZnO NP treatment

10 100 1000

0.11 0.10 0.22

0.30 0.26 0.73

0.34 0.28 0.90

0.35 0.67 1.06

a a a

1.07 0.10 0.02

2.97 0.26 0.07

3.42 0.28 0.09

3.52 0.67 0.11

a b b

Zn2+ treatment

10 100 1000

0.22 0.16 3.08

0.20 0.26 7.63

0.35 0.29 12.86

0.36 1.10 16.47

a a b

2.16 0.16 0.31

2.05 0.26 0.76

3.53 0.29 1.29

3.63 1.10 1.65

a b b

Note: a Assimilated mass of Zn (mg) = shoot weight (g)  detected concentrations of Zn in shoots (lg g 1). b Uptake percentage (%) = assimilated mass of Zn (mg) in the shoots/initial mass of ZnO NPs in the nutrient solution (mg)  100. c Different letters indicate significant difference of assimilated mass at different concentrations between ZnO NP and Zn2+ treatment, significant at p < 0.05 (one-way ANOVA). d Different letters indicate significant difference of uptake percentage at different concentrations between ZnO NP and Zn2+ treatment, significant at p < 0.05 (one-way ANOVA).

for alfalfa). This variation might be due to the differences in xylem structure of plants, and has been considered as a critical factor in the speed of water transport (Guthrie, 1989). Another reason for the wide ranging results is the variability among the NPs used, as the tissue-penetration ability of NPs depends strongly on their physicochemical properties (Santos et al., 2010). Similarly, Slomberg and Schoenfisch (2012) evaluated the phytotoxicity of 14, 50 and 200 nm Si NPs at 250 mg L 1 exposed to A. thaliana over 3 and 6 weeks in a hydroponic growth medium. The authors reported Si levels in the roots of 43 992 lg g 1 (14 nm), 70 442 lg g 1 (50 nm) and 60 217 (200 nm) lg g 1.

Fig. 4. SEM image of S. tabernaemontani root surface under treatment of (A) control; (B) 1000 mg L 1 Zn2+ ions and (C) 1000 mg L 1 ZnO NPs. Nanoparticles are pointed by arrows.

3.3.2. Translocation of Zn from roots to shoots Total Zn levels in the shoots after both treatments of ZnO NPs and Zn2+ are shown in Fig. 3B. Zn concentrations in the shoots for the ZnO NP treatment ranged from 6.4 to 110.4 lg g 1, while for Zn2+ treatment the levels ranged from 11.2 to 1710 lg g 1 (dry weight). Zn levels remained much lower in the shoots than in the roots, implying that the Zn translocation potential from roots to shoots in S. tabernaemontani for both treatments was limited. This present finding is consistent with Lin and Xing (2008) who indicated that ZnO NPs tended to be attached so strongly to the surface of the roots that few could be translocated to the shoots of ryegrass. The translocation factor (TF) for Zn is defined as the concentration ratio of shoot to root (Cshoot/Croot). The low TF values of 0.02– 0.05 for Zn at a ZnO NP treatment at a concentration of 10 mg L 1, and the even lower TF value of 0.001–0.003 at a concentration of 1000 mg L 1 implied that the translocation potential was reduced with increasing external concentrations of ZnO NPs in the nutrient solution. Compared to the Zn2+ treatment (with TF values in the range of 0.05–0.27), the relatively low TF value for the ZnO NP treatment indicates that it was more difficult to translocate Zn (as NPs) to the shoot compared to Zn2+. The BAFs of Zn for the shoots of S. tabernaemontani under both treatments are shown in Table 3. The BAFs for the ZnO NP-treated plants ranged from 0.02 to 2.72, while for Zn2+ treatment the values ranged from 0.11 to 2.81. Table 4 shows the assimilated mass (mg) and uptake percentage (% of mass initially in solution) for Zn in the plant shoots under both treatments. The shoot uptake with ZnO NP treatment ranged from 0.02% to 3.52%, while for Zn2+ treatment the value was between 0.16% and 3.63%. Our findings are in good agreement with previous studies which indicated that the total Zn levels in the shoots subsequent to ZnO NP treatment were less than 10% (Navarro et al., 2012). The mechanisms of removal of nanoparticles from wetland systems include: (i) plant uptake (as well as adsorption to plant

217

D. Zhang et al. / Chemosphere 120 (2015) 211–219

vc.

A

B

nu.

vs. cw.

vs.

C

D

cm.

cw. Pl.

Pl. Fig. 5. Transmission electron microscopy. (A) and (B) TEM image of the cellular structure of S. tabernaemontani containing ZnO NPs on 5 lm section after 21 d exposure to a solution of ZnO NPs at the concentration of 1000 mg L 1; (B) ZnO NPs have penetrated both the cell wall and plasma membrane in the root of S. tabernaemontani, and aggregated in the interface between the cell wall and the plasma membrane; (C) ZnO NPs could be transported through plasmodesm within the cell and clearly be seen in the intracellular space and (D) higher magnification of the ZnO NPs in (C). vc: vascular cylinder; cw: cell wall; cm: cell membrane; pl: plasmodesm; vs: vacuole; nu: nucleus. Nanoparticles are identified by arrows.

Fig. 6. Transmission electron microscopy. (A) TEM image shows the presence of ZnO NPs around a single organelle in the cell of the S. tabernaemontani root with external concentration of 1000 mg L 1; ZnO NPs were aggregated on the membrane of the organelle and seen as beads on the string after 21 d; (B–C) ZnO NPs attached on the membrane of the same organelle and from enlarged imaging of A. Nanoparticles were pointed by arrows.

surfaces); and (ii) dissolution, aggregation, and sedimentation. This study focused only on uptake and translocation and bioaccumulation by the higher aquatic plant. The other mechanisms (mainly aggregation and sedimentation) were not investigated. Nevertheless, our results showed that the higher aquatic plant could account for the removal of up to 47% of the initial mass of ZnO NPs in solution (Tables 2 and 3). 3.4. SEM and TEM results To further clarify the nature of NP uptake by the higher aquatic plant S. tabernaemontani, SEM was used to observe the localization of ZnO NPs in the plant root tissues. Fig. 4 shows the SEM images of the roots of S. tabernaemontani in both the treatments and control.

No particles were observed on the root epidermis of the control (Fig. 4A) and the Zn2+ treatment (Fig. 4B), while the presence of monodispersed ZnO NPs and small clusters of aggregated ZnO NPs on the root epidermis was observed in the NP-treated plants (Fig. 4C). In order to determine whether NPs could enter root cells of S. tabernaemontani, the root tips exposed to ZnO NPs (1000 mg L 1) were further analysed by TEM (Fig. 5). After the 21-d exposure period, ZnO NPs had not only penetrated both the cell wall and plasma membrane (Fig. 5A and B), but could also be transported through the plasmodesm within the cell, and could clearly be seen in the intercellular space of the cell (Fig. 5C and D). Recent studies have already demonstrated that NPs can penetrate different biological barriers, from mammalian cells, the blood–brain barrier of humans to plant cells (Lee et al., 2008; Lin

218

D. Zhang et al. / Chemosphere 120 (2015) 211–219

and Xing, 2008). Cellular penetration is the most accepted mode of action by which NPs interact with plants (Lin et al., 2009; Chen et al., 2010). Transport of NPs into plant cells is complicated due to the high turgor pressure in plant cells combined with the presence of the rigid cell wall hindering internalization. It is assumed that the diameters of NPs (core: 1–100 nm) relative to the diameter of plant cell wall pores (3.5–5 nm) would already indicate a restriction in the ability of NPs to penetrate the plant cell via active/passive transport (Carpita and Gibeaut, 1993). However, results collected to date on whole plant cells suggest that an NP’s ability to penetrate a cell wall depends more on its other properties (Miralles et al., 2012). Several mechanisms have been identified which would allow NPs to penetrate plant cells, for instance, resembling endocytosis or nonendocytic penetration allows absorption of these materials/molecules (Etxeberria et al., 2006; Harris and Bali, 2008), inducing the formation of new and large size pores (Navarro et al., 2008; Lin and Xing, 2008), and binding to carrier proteins through aquaporin, and ion channels. (Nair et al., 2010; Rico et al., 2011). In particular, NP internalization studies using isolated plant cells have suggested that the endocytosis pathway may be involved in key cellular processes and active-transport mechanisms such as nutrient uptake and the regulation of plasma membrane receptors, as well as plasma membrane recycling and signalling (Miralles et al., 2012). The TEM image of the NP-treated root cells of S. tabernaemontani (Fig. 6) shows that the ZnO NPs aggregated around an organelles showing up as beads on a string. Once inside a plant cell, NPs can be transported apoplastically or symplastically through plasmodesmata (Lucas and Lee, 2004). In cell suspensions, internalization can occur via fluid-phase endocytosis, which is the incorporation of solutes from the apoplast to the vacuole via vesicles generated at the plasma membrane (Etxeberria et al., 2009). Plasmodesmata or intercellular bridges were reported to be cylindrical channels approximate 40 nm in diameter (Torney et al., 2007). Etxeberria et al. (2006) demonstrated that Platanus occidentalis (sycamore) cells accumulated quantum dots in large, spherical cytoplasmic organelles that are thought to be diverse storage compartments used in the indiscriminate trapping that is characteristic of fluid-phase endocytosis. Slomberg and Schoenfisch (2012) investigated the phytotoxicity of Si NPs exposed to Arabidopsis thaliana at concentrations of 250 and 1000 mg L 1. The authors reported that within the root cells, the Si NPs were localized in the cytoplasm of the cell surrounding organelles. Wang et al. (2012) provided evidence for the transport and bioaccumulation of CuO NPs (20–40 nm) in maize; the authors demonstrated that CuO NPs could exist both in the intercellular space, and the cytoplasm of cortical cells. In the present study, the existence of ZnO NPs in the intercellular space, and the presence of ZnO NPs around the organelles demonstrate that NPs may pass through the epidermis and cortex.

4. Conclusions This study reports on the fate and the bioaccumulation potential of ZnO NPs in hydroponic mesocosms planted with S. tabernaemontani. S. tabernaemontani in wetland mesocosms exhibited concentration-dependent growth inhibition by ZnO NPs. Such phytotoxicity exhibited by the ZnO NPs was not directly resulted from their dissolution in the bulk nutrient solution. S. tabernaemontani clearly had the capacity to accumulate Zn in their roots in both ZnO NP and Zn2+ treatment, the bioaccumulation potential of ZnO NPs was however substantially greater than Zn2+. However, their low translocation factor, implied that the translocation of Zn NPs from the roots to shoots of S. tabernaemontani was limited. SEM observations confirm the presence of ZnO NPs in the root

tissues, while TEM analysis demonstrated that NPs could penetrate the cell walls and reach the organelles of S. tabernaemontani. Despite the fact that concentrations used in our experiments were higher than levels that are currently environmentally relevant (based on modeled estimates of environmental release), our results nevertheless indicate both the potential for phytotoxicity of these NPs and the mechanisms by which they may be bioaccumulated and translocated in the higher aquatic plant. Such information may well be important because the release of these NPs will continue to increase over time due to their large scale production and use. Future in-depth research is needed on nanoparticle behaviour and the complex soil–nanoparticle–plant interactions in actual treatment wetlands before efficient phytoremediation of NP-contaminated wastewater may be realized. Appendix A. Supplementary material Supplementary material associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/ j.chemosphere.2014.06.041. References Burken, J.G., Schnoor, J.L., 1998. Predictive relationships for uptake of organic contaminants by hybrid poplar trees. Environ. Sci. Technol. 32, 3379–3385. Carpita, N.C., Gibeaut, D.M., 1993. Structural models of primary cell walls in flowering plants: consistency of molecular-structure with the physical properties of walls during growth. Plant J. 3, 1–30. Chen, R., Ratnikova, T.A., Stone, M.B., Lin, S., Lard, M., Huang, G., Hudson, J.S., Ke, P.C., 2010. Differential uptake of carbon nanoparticles by plant and mammalian cells. Small 6 (5), 612–617. Collins, C., Fryer, M., Grosso, A., 2006. Plant uptake of non-ionic organic chemicals. Environ. Sci. Technol. 40, 45–52. Etxeberria, E., Gonzalez, P., Baraja-Fernandez, E., Romero, J.P., 2006. Fluid phase endocytic uptake of artificial nano-spheres and fluorescent quantum dots by sycamore cultured cells: evidence for the distribution of solutes to different intracellular compartments. Plant Signal Behav. 1, 196–200. Etxeberria, E., Gonzalez, P., Pozueta, J., 2009. Evidence for two endocytic transport pathways in plant cells. Plant Sci. 177 (4), 341–348. Gottschalk, F., Sonderer, T., Scholz, R.W., Nowack, B., 2009. Modeled environmental concentrations of engineered nanomaterials (TiO2, ZnO, Ag, CNT, fullerences) for different regions. Environ. Sci. Technol. 43 (24), 9216–9222. Guthrie, R.L., 1989. Xylem structure and ecological dominance in a forest community. Am. J. Bot. 76, 1216–1228. Harris, A.T., Bali, R., 2008. On the formation and extent of uptake of silver nanoparticles by live plants. J. Nanopart. Res. 10, 691–695. Hernandez-Viezcas, J.A., Castillo-Michel, H., Servin, A.D., Peralta-Videa, J.R., GardeaTorresdey, J.L., 2011. Spectroscopic verification of zinc absorption and distribution in the desert plant Prosopis juliflora-velutina (velvet mesquite) treated with ZnO nanoparticles. Chem. Eng. J. 170, 346–352. Hoffmann, M.R., Martin, S.T., Choi, W., Bahnemann, D.W., 1995. Environmental application of semiconductor photocatalysis. Chem. Rev. 95, 69–96. Ju-Nam, J., Lead, J.R., 2008. Manufactured nanoparticles: an overview of their chemistry, interactions and potential environmental implications. Sci. Total Environ. 400, 396–414. Kadlec, R.H., Knight, R.L., 1996. Treatment Wetlands. CRC Press Inc., Boca Raton, FL. Kim, B., Park, C.S., Murayama, M., Hochella, M.F., 2010. Discovery and characterization of silver sulphide nanoparticles in final sewage sludge products. Environ. Sci. Technol. 44, 7509–7514. Kiser, M.A., Westerhoff, P., Benn, T., Wang, Y., Perez-Rivera, J., Hristovski, K., 2009. Titanium nanomaterial removal and release from wastewater treatment plants. Environ. Sci. Technol. 43 (17), 6757–6763. Kümmerer, K., Menz, J., Schubert, T., Thielemans, W., 2011. Biodegradability of organic nanoparticles in the aqueous environment. Chemosphere 82, 1387– 1392. Larue, C., Laurette, J., Herlin-boime, N., Khodja, H., Fayard, B., Flank, A., Brisset, F., Carriere, M., 2012. Accumulation, translocation and impact of TiO2 nanoparticles in wheat (Triticum aestivum spp.): influence of diameter and crystal phase. Sci. Total Environ. 431, 197–208. Lee, C.W., Mahendra, S., Zodrow, K., Li, D., Tsai, Y.C., Braam, J., 2010. Developmental phytotoxicity of metal oxide nanoparticles to Arabidopsis thaliana. Environ. Toxicol. Chem. 29 (3), 669–675. Lee, W.M., An, Y.J., Yoon, H., Kwbon, H.S., 2008. Toxicity and bioavailability of copper nanoparticles to the terrestrial plants mung bean (Phaseolus radiates) and wheat (Triticum aestrivum): plant agar test for water-insoluble nanoparticles. Environ. Toxicol. Chem. 27, 1915–1921. Lee, W.M., Jin, I.K., An, Y.J., 2012. Effect of silver nanoparticles in crop plant Phaseolus radiates and Sorghum bicolor: media effect on phytotoxicity. Chemosphere 86, 491–499.

D. Zhang et al. / Chemosphere 120 (2015) 211–219 Li, Q., Mahendra, S., Lyon, D.Y., Brunnet, L., Liga, M.V., Li, D., Alvarez, P.J., 2008. Antimicrobial nanomaterials for water disinfection and microbial control: potential applications and implications. Water Res. 42 (18), 4591–4602. Limbach, L.K., Bereiter, R., Müller, E., Krebs, R., Gälli, R., Stark, W.J., 2008. Removal of oxide nanoparticles in a model wastewater treatment plant: influence of agglomeration and surfactants on clearing efficiency. Environ. Sci. Technol. 42, 5828–5833. Lin, D.H., Xing, B.S., 2007. Phytotoxicity of nanoparticles: Inhibition of seed germination and root growth. Environ. Pollut. 150, 243–250. Lin, D.H., Xing, B.S., 2008. Root uptake and phytotoxicity of ZnO nanoparticles. Environ. Sci. Technol. 42, 5580–5585. Lin, S., Reppert, J., Hu, Q., Hudson, J.S., Reid, M.L., Ratnikova, T.A., Rao, A.M., Luo, H., Ke, P.C., 2009. Uptake, translocation, and transmission of carbon nanomaterials in rice plants. Small 5 (10), 1128–1132. López-Moreno, M.L., Rosa, G.D.L., Hernández-Viezcas, J.A., Peralta-Videa, J.R., Gardea-Torresdey, J.L., 2010. X-ray absorption spectroscopy (XAS) corroboration of the uptake and storage of CeO2 nanoparticles and assessment of their differential toxicity in four edible plant species. J. Agric. Food Chem. 58, 3689–3693. Lucas, W.J., Lee, J.Y., 2004. Plasmodesmata as a supracellular control network in plants. Nat. Rev. Mol. Cell Biol. 5, 712–726. Ma, H., Williams, P.L., Diamond, S.A., 2013. Ecotoxicity of manufactured ZnO nanoparticles – a review. Environ. Pollut. 172, 76–85. Ma, X., Geiser-Lee, J., Deng, Y., Kolmakv, A., 2010. Interaction between engineered nanoparticles (ENPs) and plants: phytotoxicity, uptake and accumulation. Sci. Total Environ. 408, 3053–3061. Miralles, P., Church, T.L., Harris, A.T., 2012. Toxicity, uptake, and translocation of engineered nanomaterials in vascular plants. Environ. Sci. Technol. 46, 9224–9239. Nair, R., Varghese, S.H., Nair, B.G., Maekawa, T., Yoshida, Y., Kumar, D.S., 2010. Nanoparticulate material delivery to plant. Plant Sci. 179, 154–163.

219

Navarro, E., Baun, A., Behra, R., Hartmann, N.B., Filser, J., Miao, A.J., Quigg, A., Santschi, P.H., Sigg, L., 2008. Environmental behaviour and ecotoxicity of engineered nanoparticles to algae, plants, and fungi. Ecotoxicology 17, 372–386. Navarro, D.A., Bisson, M.A., Aga, D.S., 2012. Investigating uptake of waterdispersible CdSe/ZnS quantum dots nanoparticles by Arabidopsis thaliana plants. J. Hazard. Mater. 211–212, 427–435. Peralta-Videa, J.R., Zhou, L., Lopez-Moreno, M.L., Rosa, G., Hong, J., GardeaTorresdey, J.L., 2011. Nanomaterials and the environment: A review for the Biennium 2008–2010. J. Hazard. Mater. 186, 1–15. Rico, C.M., Majumdar, S., Duarte-Gardea, M., Peralta-Videa, J.R., Gardea-Torresdey, J.L., 2011. Interaction of nanoparticles with edible plants and their possible implications in the food chain. J. Agric. Food Chem. 59, 3485–3498. Santos, A.R., Miguel, A.S., Tomaz, L., Malhó, R., Maycock, C., Vaz Patto, M.C., Fevereiro, P., Oliva, A., 2010. The impact of CdSe/ZnS quantum dots in cell of Medicago sativa in suspension culture. J. Nanobiotechnol. 8 (1), 24. Slomberg, D.L., Schoenfisch, M.H., 2012. Silica nanoparticle phytotoxicity to Arabidopsis thaliana. Environ. Sci. Technol. 46, 10247–10254. Stampoulis, D., Sinha, S.K., White, J.C., 2009. Assay-dependent phytotoxicity of nanoparticles to plants. Environ. Sci. Technol. 43, 9473–9479. Torney, F., Trewyn, B.G., Lin, V.S.-Y., Wang, K., 2007. Mesoporous silica nanoparticles deliver DNA and chemicals into plants. Nat. Nanotechnol. 2, 295–300. U.S. Environmental Protection Agency, 2000. Methodology for Deriving Ambient Water Quality Criteria for the Protection of Human Health. Technical Support Document Volume 2: Development of National Bioaccumulation Factors. Office of Water EPA-822-R-03-030. Wang, Z., Xie, X., Zhao, J., Liu, X., Feng, W., White, J.C., Xing, B., 2012. Xylem- and phloem-based transport of CuO nanoparticles in maize (Zea mays L.). Environ. Sci. Technol. 46, 4434–4441. Vymazal, J., 2011. Constructed wetlands for wastewater treatment: five decades of experience. Environ. Sci. Technol. 45 (1), 61–69.