Response of Soil Microorganisms to As-Produced and Functionalized

Nov 19, 2012 - Results show that repeated applications of AP-SWNTs can affect microbial community structures and induce minor changes in soil metaboli...
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Response of Soil Microorganisms to As-Produced and Functionalized Single-Wall Carbon Nanotubes (SWNTs) Zhonghua Tong,†,⊥ Marianne Bischoff,† Loring F. Nies,‡ Phillip Myer,∥ Bruce Applegate,§,∥ and Ronald F. Turco†,* †

College of Agriculture − Laboratory for Soil Microbiology, School of Civil Engineering, § Department of Food Science, ∥ Department of Biological Sciences, Purdue University, West Lafayette, Indiana 47907, United States ⊥ Department of Chemistry, University of Science & Technology of China, Hefei, Anhui 230026, China ‡

S Supporting Information *

ABSTRACT: The use of single-wall carbon nanotubes (SWNTs) in manufacturing and biomedical applications is increasing at a rapid rate; however data on the effects of a potential environmental release of the materials remain sparse. In this study, soils with either low or high organic matter contents as well as pure cultures of E. coli are challenged with either raw as-produced SWNTs (AP-SWNTs) or SWNTs functionalized with either polyethyleneglycol (PEG-SWNTs) or m-polyaminobenzene sulfonic acid (PABS-SWNTs). To mimic chronic exposure, the soil systems were challenged weekly for six weeks; microbial activities and community structures for both the prokaryote and eukaryote community were evaluated. Results show that repeated applications of AP-SWNTs can affect microbial community structures and induce minor changes in soil metabolic activity in the low organic matter systems. Toxicity of the three types of SWNTs was also assessed in liquid cultures using a bioluminescent E. coli-O157:H7 strain. Although decreases in light were detected in all treated samples, low light recovery following glucose addition in AP-SWNTs treatment and light absorption property of SWNTs particles suggest that AP-SWNTs suppressed metabolic activity of the E. coli, whereas the two functionalized SWNTs are less toxic. The metals released from the raw forms of SWNTs would not play a role in the effects seen in soil or the pure culture. We suggest that sorption to soil organic matter plays a controlling role in the soil microbiological responses to these nanomaterials.

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INTRODUCTION Since their isolation and characterization in 1991,1 single-wall carbon nanotubes (SWNTs) have been subjected to hundreds of research studies addressing their intrinsic behavior and potential use.2 SWNTs are composed of graphite sheets rolled into hollow cylinders with nanosized diameters and can be uncapped or capped using fullerenes. They are now used for purposes ranging from electronics to building materials. With their expanding role in manufacturing, some environmental exposure is likely raising concerns about their potential environmental risk.3 Understanding the potential environmental consequences of environmental exposure of SWNTs is warranted as their toxicity has been reported. For example, SWNTs have been shown to induce dose-dependent lesions in the lungs of mice4 and human cervical carcinoma HeLa cell apoptosis.5 Roberts et al. showed that SWNTs solubilized with lysophophatidylcholine could induce mortality in Daphnia magna.6 SWNTs have also been shown to cause respiratory toxicity and some physiological changes in rainbow trout.7 Antimicrobial activity © XXXX American Chemical Society

of SWNTs has also been demonstrated and attributed to cell membrane damage following contact with the SWNTs.8 Pristine SWNTs are minimally soluble in water and most organic solvents; the solubility of small diameter SWNTs is only 95 and 1 mg L−1 in 1,2-dichlorobenzene and toluene, respectfully.9,10 Bundled SWNTs strongly resist dispersion in water and, if dispersed, tend to reaggregate or aggregate with macromolecules or deposit on surfaces.10 To overcome this issue, most technical applications of SWNTs take advantage of the use of end-group and sidewall functionalization to improve dispersion.11 Noncovalent solubilization can also be achieved by wrapping the SWNTs in an organic polymer.12 Functionalized SWNTs (f-SWNTs) have been shown to be less cytotoxic, which improves their biocompatibility for potential pharmaceutical applications.13,14 To date, few studies have Received: August 10, 2012 Revised: November 16, 2012 Accepted: November 19, 2012

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aqueous suspension of f-SWNTs each at 10 and 50 μg g−1, and then mixed with a spatula. Accounting for impurities (product mean purity: AP-SWNTs 50%, f-SWNTs 85%) and functional group content (PEG:SWNTs, 30:70 w/w; PABS:SWNTs, 65:35 w/w), the actual concentrations of SWNTs applied were 500 μg g−1 soil for AP-SWNTs, 6 and 30 μg g−1 soil for PEG-SWNTs, 3 and 15 μg g−1 soil for PABS-SWNTs. Soil moisture content was adjusted to an equivalent of −0.33 bar, which corresponds to water content of 29.2% for Drummer soil, and 15.2% for Tracy soil. Water only controls were also included, and all treatments were done in triplicate, and the experiment was repeated. The CO2 released from the incubated soils was trapped in 10 mL of 1 M KOH and determined by back-titration with HCl. Soil Microcosm Study. To follow the effects of repeated applications, soil samples (100 g dry weight equivalent, in triplicate) were weighed into 250 mL screw-top jars and preincubated at 23 °C for 4 days. After the preincubation, nanomaterials were added to the microcosms as dry APSWNTs applied at 1000 μg g−1 soil or an aqueous suspension of functionalized SWNTs (f-SWNTs) each at 10 and 50 μg g−1 followed by thoroughly mixing with soils. The application was repeated weekly for six weeks and at the end of the experiments a total of 6000 μg g−1 of AP-SWNTs, 60 and 300 μg g−1 of each f-SWNTs were applied. A water only control was used. Soil moisture was maintained at −0.33 bar equivalent by gravimetrically adjusting the moisture content using sterile distilled H2O as needed. Microbial Activity Assessment. Weekly for the 6 week study, the ability of the standing biomass to mineralize glucose was determined by treating subsamples from each microcosm with radiolabeled glucose (D-glucose-UL-14C, specific activity 264 mCi mmol−1, Sigma-Aldrich, St. Louis, MO) to create a substrate induced respiratory response. A subsample (10 g dry weight equivalent) was mixed thoroughly with a mixture of 2.5 mg nonlabeled glucose and 47.5 mg of talcum, and supplemented with 0.15 μCi 14C-labeled glucose. The samples were incubated at 23 °C and the evolved 14C−CO2 was trapped in 10 mL of 1 M KOH. The alkali trap was recovered after 3 and 24 h of incubation. Treatments were done in triplicate. A 1 mL aliquot of each KOH trap solution was removed to a 22 mL scintillation vial, which was mixed with 15 mL of scintillation cocktail (Econosafe, Research Products International, Mt. Prospect, IL). Vials were stored in the dark at room temperature overnight prior to liquid scintillation counting using a Packard 1600 TR liquid scintillation analyzer (PerkinElmer, Shelton, CT) with quench correction. Microbial Biomass Analysis. After SWNTs were applied weekly for 6 times and incubated for a week, subsamples were taken from the microcosms, lyophilized, and stored frozen until analysis. Dried soil samples (2−5 g) were used to estimate microbial biomass size using phospholipid-derived phosphate (PL-PO4).25 The total lipid was extracted from soil samples using a 1:2:0.8 chloroform/methanol/phosphate buffer (50 mM, pH 7.4) solution and fractionated on a silicic acid column into neutral, glycol-, and phospholipid by sequential elution with chloroform, acetone, and methanol. The phospholipid fraction was collected and a 100 μL portion was used to determine the PL-PO4 concentration. Phosphate derived from the phospholipids was colorimetrically determined by measuring O.D. at 610 nm after potassium persulfate digestion.25 DNA Extraction. SWNTs were applied weekly for 6 weeks. After the seventh week, subsamples were taken from each

investigated the effects of raw or as produced SWNTs compared with functionalized forms on soil microorganisms. As microorganisms play a key role in all biogeochemical cycling, how they respond to possible stress conditions is suggested as an early indicator of potential long-term soil system responses.15 It has been shown that under certain culture conditions, fullerene C60 can inhibit bacterial activity.16 When some known confounding factors were avoided, a lack of antibacterial activity by fullerene were observed.17 Natural organic matter have been shown to mitigate the toxic effects of nanomaterials,18,19 which supported our findings that few adverse effects of C60 are expressed in soil system studies20 suggesting that the soil matrix is playing a critical role in modulating the interaction between microorganisms and introduced nanomaterials. We suggest that soil may play a similar role with SWNTs. In this study, we examined the response of microorganisms to SWNTs in two soils. We evaluated both the raw form or as produced form of SWNTs (including production catalyst contaminants) and the more purified and dispersed forms because both types could be utilized or transported in large volumes. The unpurified or as produced forms with production contaminants would be transported to manufacturing facilities and are suggested as having the greatest potential for occurring in large spills. As produced forms of SWNTs made using catalytic methods can contain a significant amount of metal residues21 including Ni, Y, or Pb. The mobilized metallic impurities have been shown to be at toxicologically significant levels.22,23 We use a bioluminescent E. coli O157:H7 (denoted as E. coli-lux), which contains the luxCDABE cassette expressed from a lac promoter on a multi copy plasmid containing a kanamycin resistance marker,24 to develop an understanding of the direct response of bacteria to the different forms of SWNTs. It is imperative to understand the interaction between SWNTs and soil microbes/soil function as their introduction into the environment is increasing with the increasing utilization of these materials.

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MATERIALS AND METHODS Nanomaterials and Soils. The following SWNTs were purchased as detailed in the Supporting Information: raw asproduced SWNTs (AP-SWNTs), SWNTs functionalized with polyethyleneglycol (PEG-SWNTs) and SWNTs functionalized with m-polyaminobenzene sulfonic acid (PABS-SWNTs). AP-, PEG-, and PABS-SWNTs were suspended in waters using sonication (1 h) at 0.5, 2.5, and 2.5 mg mL−1, respectively. The Drummer soil (fine-silty, mixed, superactive, mesic Typic Endoaquoll, 3.6% organic matter, pH 6) was collected from continuous corn no-till plots at Purdue Agriculture Research and Education Center located northwest of the Purdue University campus and a Tracy soil (coarse-loamy, mixed, active, mesic Ultic Hapludalfs, 1.5% organic matter, pH 5.5) was collected from the Pinney - Purdue Agriculture Center located on the county line between Porter and LaPorte counties, in Northwest Indiana. All nanotube suspensions were agitated before application to soils or media to ensure a suspension. Details on soil preparation and characteristics are provided in the Supporting Information. Soil Basal Respiration. A 30 day single application basal respiration study was conducted using biometer flask microcosms as previously described.20 Briefly, following preincubation, soil (50 g dry weight equivalent) was treated with SWNTs. Dry AP-SWNTs were applied at 1000 μg g−1 soil and B

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microcosm to isolate genomic DNA using the FastDNA SPIN kit for soil (MP Biomedical, Solon, OH) according to the manufacturer’s instructions. The amount and quality of DNA extracts were determined using a NanodropTM ND-1000 Spectrophotometer (NanoDrop Technologies, Wilmington, DE). PCR was performed using bacterial and eukaryotic specific primers and the community profiles were analyzed using denaturing gradient gel electrophoresis (DGGE). PCR-DGGE Fingerprinting. 16S rRNA gene fragments for DGGE analysis were obtained using primer set F338 with a 40 bp GC clamp attached at 5′ end and R534 as described by Muyzer et al.26 For eukaryote 18S rRNA gene amplification, primers F1427-GC and R1616,27 which amplify a fragment of approximately 210 bp, were used with a modified cycling program. DGGE was performed on 8% polyacrylamide gels with a denaturant gradient 35−70% for bacteria and 25−65% for eukaryote, where 100% denaturant is defined as 7 M urea plus 40% v/v formamide. The sequences of the primer sets and the GC-clamp, PCR cycling program, and information on DGGE analysis are provided in the Supporting Information. The DGGE profiles were analyzed using Quantity One, version 4.6.7 (Bio-Rad Laboratories, Hercules, CA, USA). Cluster analysis of profile similarity was performed using the unweighted pair group method with arithmetic mean (UPGMA). Recovery and Sequencing of Selected DGGE Bands. The DGGE bands which appeared to be either enhanced or reduced following SWNTs treatment were excised and suspended in 100 μL Rnase/Dnase-free water overnight to elute DNA. Eluted DNA fragments were reamplified with the same primers as described above and examined by DGGE to ensure purity and correct mobility. A second round of excision and elution was performed if necessary. To sequence the bands, PCR amplification was performed with primers used for DGGE analysis without the GC clamp. The PCR product was purified by using QIAquick gel extraction kit (QIAGEN, Valencia, CA) according to the manufacturer’s instruction and directly sequenced28 at Purdue University’s Genomics Facility. Nucleotide sequences were compared to sequences in the National Center for Biotechnology Information (NCBI) GenBank database using the BLASTn search program. Sequence data of DGGE bands obtained in this study have been deposited in the GenBank database (Table 1) under accession numbers GQ470417- GQ470426. Metal Impurities in AP-SWNTs and Release into Aqueous Solution. AP-SWNTs were assayed for metal content by ashing followed with acid digestion, and metal impurities quantification by ICP-MS.29 The release of soluble metals into aqueous solution was measured by shaking the APSWNTs in either distilled water or soil solution extract, followed by ICP-MS quantification. Detailed information on sample preparation and ICP-MS analysis was provided in the Supporting Information. Bioluminescence Assay. A bioluminescent E. coli O157:H7 was constructed by introducing a plasmid-based luxCDABE gene cassette under the control of the lac promoter and the plasmid also carries a kanamycin resistance marker,24 denoted as E. coli-lux. It is important to note that due to the high copy number of the plasmid the strain is constitutively luminescent.24 The strain was maintained on LB agar supplemented with 100 μg mL−1 kanamycin (Sigma-Aldrich, St. Louis, MO) as a source of selective pressure to maintain the bioluminescent phenotype. The inoculum was prepared by

Table 1. Sequences Similarities of Excised DGGE Bands band (Genbank accession number) F338-GC/R534 Primer set A (GQ470417)) B (GQ470418) C (GQ470419) F1427-GC/ R1616 primer set H (GQ470421) I (GQ470422) J (GQ470423) K (GQ470424) L (GQ470425) M (GQ470426)

most closely related database entry (Genbank accession number)

% sequence similarity (no. of bases)a

band intensityb

Uncultured bacterium clone (EU134745.1) Uncultured gamma proteobacterium (DQ676408.1) Uncultured Lysobacter sp. (AM935900.1)

95 (160)

−

96 (160)

+

97 (160)

+

Uncultured syndiniales clone (EU793698.1) Stylonychia bifaria (FM209296.1) Cercozoa sp. DDB-2008d (EU567254.1) Coniophora marmorata (AM946632.1) Parasitorhabditis obtuse (EU003189.1) Uncultured alveolate clone (AY179988.1)

88 (165)

−

100 (162)

+

97 (165)

−

100 (165)

−

94 (160)

+

94 (162)

+

a

The numbers in parentheses are the numbers of bases used to calculate the levels of sequence similarity. bStimulated band marked +, inhibited band marked −, compared to the soil control.

growing overnight to an OD at 600 nm of 0.9 to 1.0 in LB media (+ kanamycin) at 37 °C with shaking. Cells were harvested by centrifugation (7000 rpm, 10 min), washed, and resuspended with 50 mM phosphate buffer (pH 7.0). The cell number was adjusted to approximately 5 × 108 colony forming units (CFU) per ml and 10 μL added to test tubes containing 990 μL of phosphate buffer with different concentration of SWNTs. AP-SWNTs (10, 15, 25, 50 μg mL−1) and two fSWNTs (each at 25, 63, 100, 150 μg mL−1) were included along with a no-treatment control. Again considering the impurity and functional group content, the actual concentrations of SWNTs were 5, 7.5, 12.5, and 25 μg mL−1 for APSWNTs, 14.9, 37.2, 59.5, and 89.3 μg mL−1 for PEG-SWNTs, 7.4, 18.6, 29.8, and 44.6 μg mL−1 for PABS-SWNTs. All treatments were done in triplicate. Bioluminescence (relative light units; RLU) of E. coli-lux was measured for 2 s with a luminometer (Zylux corporation, Huntsville. AL) at 10 and 30 min after the mixing. The bioluminescence response of bacteria is directly dependent on levels of ATP, FMNH2, and NADPH.30 When bacteria cells are impaired, they will show lower metabolic activities and light production. Therefore, a stock glucose solution was added at 1% in each tube and the bioluminescence was measured at 30, 60, and 105 min after glucose amendment. Data Analysis. Mean values and standard deviation of CO2 production in soil respirations, total microbial biomass, and relative bioluminescence (%) were calculated. Analysis of variance (ANOVA) and Tukey’s test were performed to compare treatment effects and significant differences with SAS, version 9.1 (Cary, NC) at α = 0.05. A P value of 0.05) differences between treatments within a soil (parts a and b of Figure 2), respectfully. However, in the Tracy soil, but not the Drummer, 24 h cumulative 14C−CO2 production in samples treated with APSWNTs was significantly lower than the control (part d of Figure 2) after four weeks of AP-SWNTs application. It should be noted that for the Tracy soil 24 h production of CO2 (related to biomass growth) continued to decline, as more APSWNTs were added past week four (part d of Figure 2). With substrate induced response, CO2 evolution during the first three hours post glucose addition reflects the activity of the standing microbial biomass, while CO2 formed between 3 and 24 h reflects the CO2 released from a growing population.39 At 24 h, 25 to 27% and 30 to 33% of the applied glucose in the control Drummer and Tracy soils respectively were typically mineralized to CO2 (parts c and d of Figure 2). More CO2 was released from the Tracy soil than the Drummer soil in 24 h, although less CO2 was produced during the first 3 h in Tracy

control soil was two times greater than the Tracy control soil (Figure S1 of the Supporting Information). It should be noted that basal respiration in the Drummer soil is typically much higher than in Tracy soil.37 For either soil, no significant difference (P > 0.05) in cumulative CO2 as related to the application of any nanomaterial was observed. Noncumulative respiration data collected at each of the 12 time points were subjected to Tukey’s test and showed no significant differences (P > 0.05) between any treatments and the controls within each soil type. These findings are similar to the respiration result when we examined CO2 production for soils treated with either 1000 μg g−1 of solid C60 or 1 μg C60 g−1 soil in aqueous suspension (nC60).20 In the long term repeat application study, substrate induced respiratory response was used to estimate the impact of the nanomaterials on the ability of the standing biomass to mineralize glucose or grow in the presence of increasing levels of SWNTs. 14C-labeled glucose was used and the evolved 14C− CO2 was monitored to compare the impacts of raw and fSWNTs on the activity of the soil microbial community. Whereas the level of 14C−CO2 production at 3 h post glucose E

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soil. These results suggest that glucose addition induced a strong biomass growth response in the Tracy soil, the overall biomass in Tracy soil is smaller, and the microorganisms appear to be dominated by r-strategists, which are characterized by greater growth rates when substrates are abundant.40 For the Tracy soil, repeated introduction of AP-SWNTs suppressed the growth response and suggests high levels of AP-SWNTs are toxic to the growing biomass. This effect would have been missed in static systems where growth is not induced. The source of the suppression with AP-SWNTs is confounded by the fact the SWNTs were synthesized using Ni/Y catalyst according to the supplier and high levels of Ni have been shown to suppress activity in the soil microbial community.41,42 In our study, AP-SWNTs were applied at 1000 μg g−1 soil weekly, and have accumulated to 6000 μg g−1 in 6 weeks. On the basis of our metal release data, the theoretical level that soluble metals could reach would be 37.0 and 50.4 μg g−1 in the Drummer and Tracy soil, respectively. As stated previously, other work has shown metals applied at similar levels do not exert inhibitory effect on soil process.38 The low level of water recoverable metal from the AP-SWNTs had no effect on E. coli-lux suggesting free metal was not influencing soil processes. Therefore, the growth suppression caused by the repeated applications of SWNTs to the Tracy soil cannot be attributed to the free metals alone and reflect a toxic effect of SWNTs or SWNT carrying metal into a cell. Microbial Biomass. At the end of the 6 week incubation, subsamples removed from the microcosms were lyophilized and stored frozen. The total biomass level was estimated by phospholipid phosphate (PL-PO4), which ranged from 12 to 14 nmol PL-PO4 g−1 dry soil for Drummer, and 6−8 nmol g−1 for Tracy (Figure S2 of the Supporting Information). However, no significant difference (P > 0.05) in the size of microbial biomass was observed between the treated samples and the controls on either soil. This result implies no effects of the repeated applications of SWNTs on the total biomass and also suggests that the applied concentration of catalyst metals did not affect the biomass levels in either soil. A similar response had been observed when other soil microbial communities are treated with high level of heavy metals.37 DGGE Analysis. For both Drummer and Tracy soils, the 16S rRNA gene DGGE profiles (part a of Figure 3, Figure S3 of the Supporting Information) revealed a banding pattern that reflected the high bacterial diversity typical in soil.43 In Drummer soil, band A in the control soil was inhibited in samples treated with AP-SWNTs or f-SWNTs, whereas bands B, C, and D in the control soil were enhanced by f-SWNTs. In Tracy soil, the profiles of all treatments are similar with only one band missing in AP-SWNTs treated samples. Following the application of the three types of SWNTs, eukaryotic DGGE fingerprints yielded many visible bands (part b of Figure 3). In Drummer soil, band H from the control soil was inhibited following AP-SWNTs application and band I was enhanced by PABS-SWNTs. In Tracy soil, soil control bands J and K were inhibited by all the three types of SWNTs, while the intensity of bands L and M was increased in samples treated with PEGSWNTs at a higher concentration. In general, AP-SWNTs inhibited some bands in both bacteria and eukaryotic communities, whereas some bands appeared or their intensity increased as a result of f-SWNTs exposure. Clustering of the profiles (Figure S4 of the Supporting Information) shows that both bacterial and eukaryotic DGGE profiles are of more than 70% similarity for both soils

Figure 3. DGGE profiles from 16S rRNA gene fragments (a) and 18S rRNA gene fragments (b) amplified from samples treated with SWNTs, and the control soil after repeated application for 6 weeks. Lane 1, control; Lane 2, AP-SWNTs; Lane 3, PEG-SWNTs 6; Lane 4, PEG-SWNTs 30; Lane 5, PABS-SWNTs 3; Lane 6, PABS-SWNTs 15. The number following the materials indicate the concentration (μg SWNTs g−1soil) of nanotubes for repeated application. Letters on the figure indicate the bands with greater intensity that were extracted for sequencing.

suggesting the SWNTs treatments did not lead to significantly altered microbial communities although some community shifts were observed. To better understand changes in biodiversity prompted by the incorporation of SWNTs, DNA bands which showed an increase or decrease in intensity, when compared to the controls, were extracted from the DGGE gels, subjected to reamplification and then direct sequence analysis. The resulting DNA sequences were compared to the GenBank database using a BLASTn search and the resulting sequence with 96−100% for bacterial DGGE bands and 92 to 100% for eukaryotic DGGE bands similarities with published data are shown in Table 1. Band A, which is lost from the Drummer soil, is closely related to an uncultured proteobacterium clone and is absent in all samples treated with either AP-SWNTs or f-SWNTs indicating it is sensitive to SWNTs application. Proteobacterim are a class of environmentally important bacteria that includes Enterobacteriaceae and Pseudomonadaceae, which are common in soil and the environment.44−46 Bands B and C appeared in samples treated with f-SWNTs and showed high identity to an uncultured gamma proteobacterium (96%) and Lysobacter pocheonensis (97%), respectively. Lysobacter pocheonensis is a proteobacteria but the exact function of this species is unclear. Lysobacter lack flagella but are highly mobile via a gliding F

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mechanism and are known to produce extracellular enzymes to cope with environmental stimuli.47 Six bands were excised from the DGGE gel of 18S rRNA gene fragments and found to show phylogenetic affiliations to three phyla, Dinoflagellata (band H), protozoa (band I, J, and M) and fungus (band K and L). Band H, which was inhibited in Drummer soil following APSWNTs treatment, is related to an uncultured syndiniales clone typically assumed to be parasitic.48 Band I shows the highest matching identity (100%) to the ciliate Stylonychia bifaria, which has been reported to occur in soil but has not been defined as to their exact function.49 While in the Tracy soil, bands J and M are related to Cercozoa sp. and uncultured alveolate clone, respectively. Cercozoa are important in soil for nutrient cycling, whereas the exact role of the alveolate are unclear but are suggested to be bacterial predators.50 Bands K and L show high identity to fungus Coniophora marmorata and Parasitorhabditis obtuse respectively both are encountered in soil and the Coniophora is associated with wood decomposition.51 Clearly our understanding of complex soil communities is nascent. Furthermore, beyond identification of changes occurring in the structure of the biomass, an advance in our understanding of how these changes are linked to functions is needed. In this study, we repeatedly applied raw-SWNTs accumulating them to high concentration. In the soil with low organic matter, some inhibition was observed in 24 h after glucose amendment. This was noted starting at week 4 or when approximately 2000 μg g−1 AP-SWNTs had been applied. Whereas the direct role of the catalyst metals as an inhibitor agent can be ruled out as the dissolved metals were at low concentration, the delivery of the metals to the cells may be enhanced near the tubes. The extremely high concentration of SWNTs may occur in soil following a spill, manufacturing error or with repeated application of a SWNT containing biosolid. Clearly, the best approach for the environmental management of manufactured nanotubes remains a significant question. Procedures such as land application with high rates of SWNTs should be approached with caution. Whereas work has been published on the toxicity of carbon nanotubes using a variety of biological test systems,4−7 we present a study on the impact of production grade or functionalized SWNTs on soil microbial activity. Moreover, we have evaluated the impact of nanotubes on E. coli in pure culture. Our results demonstrated a distinct effect of SWNTs on microorganisms in the different test systems. The raw SWNTs appeared to be toxic to the metabolic activity of bacteria cells, caused minor shifts in soil microbial community structure, and also produced a minor effect on community metabolic function. The concentrations of soluble catalyst metals released into soil solution extracts were determined and they are below the range which has been shown to be toxic. However, toxicity of metals in soil has never been tested in the presence of carbon nanotubes and other work has shown metal toxicity to be enhanced when associated with carbon nanotubes. We hypothesize that soil organic matter is also a major factor contributing to the differential effects, as previous studies have reported the high retention of hydrophobic contaminants by organic matter in soils52 and we report differences in a low and high organic matter soil. Building off of previous findings,20,53 this work points out the complex nature of materials in natural systems such as soil and the role the environmental matrix will have in modulating the effect of nanomaterials.

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ASSOCIATED CONTENT

S Supporting Information *

Property of nanomaterials and soil samples, soil respiration, and biomass analysis, level of metal impurity eluted into aqueous solution, a tube-in-tube method to analyze light decrease, bioluminescence-based toxicity assay, bacterial and eukaryotic PCR-DGGE analysis, dendrogram (UPGMA) of DGGE profiles are provided. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]; phone; tel: 1 765 494 8077; fax: +1 765 496 2926. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors acknowledge support from the National Science Foundation (NSF) under Award EEC-0404006 and EPA STAR Award RD-83172001-0, the Chinese Academy of Sciences (KZCX2-YW-QN505), the National Science Foundation of China (No. 21077099), and the Fundamental Research Funds for the Central Universities (WK2060190007). The authors would like to thank Mr. Steve Sassman for conducting the ICPMS analysis in this study.

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REFERENCES

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dx.doi.org/10.1021/es303251r | Environ. Sci. Technol. XXXX, XXX, XXX−XXX