Escherichia coli Inactivation by UVC-Irradiated C60: Kinetics and

Oct 15, 2011 - Su-Gyeong Kim , Love Kumar Dhandole , Young-Seok Seo , Hee-Suk Chung , Weon-Sik Chae , Min Cho , Jum Suk Jang. Applied Catalysis A: ...
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Escherichia coli Inactivation by UVC-Irradiated C60: Kinetics and Mechanisms Min Cho,†,‡ Samuel D. Snow,† Joseph B. Hughes,† and Jae-Hong Kim*,† †

School of Civil and Environmental Engineering, Georgia Institute of Technology, 200 Bobby Dodd Way, Atlanta, Georgia 30332, United States ‡ Division of Biotechnology, Advanced Institute of Environmental and Bioscience, College of Environmental and Bioresource Sciences, Chonbuk National University, Iksan, Jeonbuk 570-752, South Korea ABSTRACT: Motivated by recent studies that documented changes in fullerene toxicity after chemical transformation, C60 aggregates (nC60) were subject to UVC irradiation at monochromatic 254 nm and subsequently evaluated for antibacterial and bactericidal properties against Escherichia coli. The nC60 treated with UVC irradiation, referred to herein as UVC-irradiated C60, did not directly inhibit bacterial growth at concentrations up to 20 mg/L. In the presence of UVA and visible light, however, UVC-irradiated C60 rapidly inactivated E. coli, suggesting that photochemical production of reactive oxygen species (ROS) was involved. The use of ROS scavengers and probes determined that hydroxyl radicals were the primary ROS responsible for the E. coli inactivation. Results from protein release, lipid peroxidation, cell permeability, and intracellular enzyme assays suggest that the inactivation mechanism involves UVC-irradiated C60 diffusing through E. coli cell membrane and producing hydroxyl radicals within the cell. Further study on water-soluble C60 derivatives and possible transformative processes is, therefore, recommended based on the environmental implications of results presented herein that nC60 exposed to UVC irradiation is more toxic than parent nC60.

’ INTRODUCTION The toxicity, or lack thereof, of nC60 (aggregate of pristine C60 in the aqueous phase) has been a topic of many recent studies. Early studies reported toxic effects in various organisms, including mammalian cells, aquatic organisms, and bacteria.1 3 Contrary to these initial findings, some researchers reported much lower toxicity levels for nC60.4,5 Later, it was suggested that the method of nC60 formation could explain some of the initial toxicity reported; when tetrahydrofuran (THF) was used as an intermediate solvent, trace amounts of THF and its byproducts could have remained and exerted toxic effects.6 8 In the absence of solvent effect, researchers found nC60 to be only mildly toxic to Escherichia coli.8 This result was further supported by additional studies which showed that nC60’s crystalline structure caused self-quenching of photoexcitation,9 11 greatly inhibiting its ability to photochemically produce reactive oxygen species (ROS), which have been suggested as one of the C60’s primary toxicity mechanisms.3,12 Although nC60 themselves may not be of major environmental concern, few studies have examined the potential impacts of derivatized C60 in the aqueous environment. We have recently reported that nC60 is readily oxidized by ozone, forming watersoluble products containing as many as 29 oxygen atoms per C60,13 similar to fullerenol, a polyhydroxylated C60. Ozonated C60 was found to have low intrinsic antimicrobial properties yet a strong and unique photochemical toxicity pathway.14 Similarly, we also reported that nC60 could be readily oxidized by UVC irradiation, which is commonly used in water and wastewater r 2011 American Chemical Society

disinfection, resulting in the formation of highly oxidized, soluble C60 products (referred to herein as UVC-irradiated C60).15 A series of papers by Hou et al.16 18 also reported transformation of nC60 into water-soluble, multiple oxygenated products via sunlight. Yet, toxicological effects of these transformation products are unknown. Functionalization of C60 is known to affect its photochemical properties in a variety of ways,19,20 with a general decrease in photochemical reactivity with consecutive additions of functional groups13 caused by the disruption of C60’s pi-conjugated system.21 Fullerene derivatives, however, typically retain their ability to produce some ROS.16,22,23 Solubilizing C60 in the aqueous phase via chemical transformations with ozonation and UVC irradiation could in fact result in increased photochemical reactivity compared to nC60, since self-quenching is prevented. Additionally, functionalization may allow facile diffusion of individual C60 molecules (i.e., molecularly dispersed in the aqueous phase) through cell membranes, resulting in direct contact with intracellular materials. The objective of this study is to examine how UVC-irradiated C60 interacts with E. coli, a representative gram-negative bacterium, in order to gain a better understanding of environmental consequences of the phototransformation of nC60. Results suggest that UVC-irradiated C60 Received: July 1, 2011 Accepted: October 15, 2011 Revised: October 11, 2011 Published: October 15, 2011 9627

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Environmental Science & Technology effectively inactivates E. coli under UVA and visible light irradiation conditions through mechanisms that potentially involve transport of derivatized C60 through the cell membrane and subsequent photocatalytic production of 3 OH within the cell.

’ EXPERIMENTAL SECTION Materials. Water (>18 MΩ) produced from a Milli-Q Water Purification System (Millipore Co.) was used in preparation of all solutions and reagents. All chemicals used were analytical, reagent-grade. All glassware used in the experiments was washed with DI water, then autoclaved at 121 °C for 15 min. UVC-Irradiated C60 Preparation. An organic solvent-free preparation technique was used to avoid potential solvent interferences between prepared nC60 and microorganisms.8 Ultrasound (20 W) was continuously applied to 500 mL of ultrapure water containing 100 mg of C60 (99.9%, Materials Electronics Research Corporation, Tucson, AZ) in a sealed Pyrex bottle for 24 h at ambient temperature using an ultrasonicator (S-4000, Misonix Co.). The color of the solution became light orange, indicating that C60 had formed water-stable clusters. The resultant solution was filtered through a 0.45-μm cellulose filter (Millipore Co.) and stored in the dark. The filtrate containing 2 3 mg/L of nC60 was further concentrated to a stock solution of 30 mg/L using a rotary evaporator. The concentration of nC60 was measured using a UV/vis spectrophotometer at 350 nm (Agilent 8453, Agilent Co.)13 and a Shimadzu TOC-VWS analyzer. The nC60 was diluted in ultrapure water and subject to UVC-mediated transformation for 7 days in a magnetically stirred quartz reactor under water cooling using four UVC lamps emitting monochromatic radiation at 254 nm (Philips Co.). The incident light intensity of each lamp at the location of the reactor was measured at 11 mW/cm2 using a UVX Radiometer with 254nm sensor (UVP Co.). Preparation and Analysis of E. coli. E. coli was selected as a representative gram negative bacterium and was obtained from the American Type Culture Collection (ATCC). The bacteria were cultured and counted according to the method described by Cho et al.24 Briefly, E. coli (ATCC 8739) was inoculated in 50 mL of Tryptic Soy Broth in a 200-mL flask and grown for 18 h at 37 °C in a shaking incubator. The bacteria were harvested by centrifugation in a 50-mL conical tube at 1000g for 10 min and washed two times with 50 mL of phosphate buffered saline solution at pH 7.2 (PBS). An E. coli stock solution was prepared by resuspending the final pellets in 50 mL of PBS. The initial population of E. coli was approximately 105 cfu/mL and obtained by diluting the stock solution. The cell concentration was determined by a spreading plate method on nutrient agar, incubating at 37 °C for 24 h, with three replicate plates at each dilution. Inactivation Experiments. A suspension (20 mL) containing UVC-irradiated C60 and E. coli was placed in a 40-mL quartz reactor and vigorously stirred using a magnetic stirrer at 1100 rpm. Three blacklight blue (BLB) lamps (300 420 nm; Philips Co.) or six fluorescent lamps (FL, visible range with a small amount of UVA irradiation; Philips Co.) were placed 3 cm from the reactor surface. The emission spectra of these lamps were verified with an Acton Research Detection system (Spectrapro-500, USA). The incident light intensity by three BLB lamps was verified by ferrioxalate actinometry25 as 1.2  10 6 Einstein/L-s. The light intensity of six FLs at a representative wavelength of 365 nm on reactor surface was measured as 330 μW/cm2 using a UVX radiometer with a long-wave sensor (UVP Co.) at bandwidth of

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10 nm. The role of visible light was investigated after placing UV cutoff filter (Optivex Co.). Temperature and pH were adjusted to 19 ( 1 °C (air-cooling) and 7.1 (10 mM of phosphate buffer), respectively. Antibacterial Activity Assay. Antibacterial activity was assayed by determining the minimum inhibitory concentration (MIC).1 For assaying the MIC, modified minimal Davis (MD) media tubes containing 0 20 mg/L of UVC-irradiated C60 were inoculated overnight with a culture of E. coli (DH5α) and grown on Luria Bertani nutrient broth. Initial OD600 was controlled to 0.002 and the lack of growth of E. coli after overnight incubation indicated the MIC. Detection of Produced ROS. Furfuryl alcohol (FFA, 0.85 mM; Aldrich Co.) and p-chlorobenzoic acid (pCBA, 2 μM; Aldrich Co.) were used as indicators for detecting 1O2 and 3 OH, respectively.9 The concentrations were analyzed by means of HPLC (Agilent 1100; Agilent Co.). A C18 reverse-phase column (Agilent Zorbax RX-C18; Agilent Co.) was used with a diode-array UV detector at a wavelength of 230 and 237 nm for measuring the concentrations of FFA and pCBA, respectively. The production of O2 3 was measured by using XTT sodium salt (2,3-bis(2-methoxy-4-nitro5-sulfophehyl)-2H-tetrazolium-5-carboxanilide inner salt, 0.15 mM; Sigma Co.) as a probe which forms a purple product (λmax = 480 nm) from the reaction with O2 3 .9 Microbial Inactivation Mechanisms. The possible mechanisms of microbial inactivation were comparatively investigated by quantifying the degree of lipid peroxidation, the amount of protein oxidation or disruption, the change in cell wall permeability, and the degradation of intracellular enzymes during cell death. The time scale for all these experiments was adjusted in order to achieve 90% (1 log) inactivation of E. coli. All the experiments were conducted in triplicate and all the assay results were within a 90% confidence interval. Note that samples from photochemical experiments were collected and instantly transferred to amber bottles after centrifugation for all these analyses. Lipid peroxidation was analyzed by observing the quantity of malondialdehyde (MDA), a product of the lipid peroxidation, based on a reaction with thiobarbituric acid (TBA) which forms a pink MDA TBA product, following a method by Maness et al.26 The amount of proteins released from E. coli surface components was determined using a modified Bradford assay,27 while the amount of oxidized proteins (i.e., protein carbonyls) was measured using an OxyELISA oxidized protein quantification kit (Millipore Co.).28 To verify the possible change of cell permeability, the reaction between an intracellular enzyme, β-D-galactosidase, and its substrate, o-nitrophenyl-β-D-galactopyranoside (ONPG),29 was monitored following the method described by Zheng et al.29 The change of enzymatic activity after photocatalytic inactivation was investigated by means of the API-ZYM system (BioMerieux Co.).30 Additional experiments were also performed using a spheroplast of E. coli which was prepared by incubating fresh E. coli in a Tris-HCl buffered suspension containing sucrose, lysozyme, and ethylenediaminetetraacetic acid (EDTA) as described by Zheng et al.29

’ RESULTS AND DISCUSSION Inactivation Kinetics. Results from MIC tests suggested that addition of as high as 20 mg/L of UVC-irradiated C60 did not inhibit E. coli growth. These findings are consistent with previous results for ozonated C60 (no effect on the growth of E. coli up to 10 mg/L 14) as well as nC60 with varying degree of UVC 9628

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Figure 1. Kinetics of E. coli inactivation by UVC-irradiated C60 ([UVCirradiated C60]0 = 15 mg/L; [ozonated C60]0 = 10 mg/L14).

irradiation (increased MIC with increasing UVC irradiation time from 0 to 110 h with the same UVC intensity15). This contrasts with the previous findings that nC60 exhibits a MIC of approximately 2.0 mg/L.1,3 The difference resulted presumably due to different methods employed for nC60 synthesis (i.e., nC60 samples used in this study were thoroughly washed to remove THF and THF derivatives8). Collectively, this oxidative transformation of nC60, either by UVC or ozone, results in decreased intrinsic (i.e., in the absence of light) bacteriostatic activity of nC60. Figure 1 shows the effect of light sources on the inactivation of E. coli by the photocatalytic activity of UVC-irradiated C60. In control tests, E. coli was not inactivated by 15 mg/L nC60 under BLB or FL light irradiation within experimental time scale (results not shown). Consistent with the results of the MIC tests, E. coli was not inactivated when as much as 15 mg/L of UVC-irradiated C60 was applied for 150 min in the dark (results not shown). A minor level (0.1 log) of inactivation was observed under BLB light in the absence of C60, which may be attributed to action of UVA and UVB light. Note that 15 mg/L (and other concentrations in this study) refers to the initial mass concentration of nC60 before UVC irradiation, which would change due to addition of functional groups during UVC treatment. No E. coli inactivation was observed under fluorescent light alone within the experimental time scale. In contrast, a significant level of E. coli inactivation (2.4 log inactivation for 120 min) was observed when BLB light was irradiated. Figure 1 also shows a comparison with a previous study that reported the kinetics of E. coli inactivation by ozonated C60 (10 mg/L) which achieved only a 1.3 log inactivation for 150 min under the same light condition.14 Also note that nC60 under the same conditions would result in no measurable E. coli inactivation.14 Under FL light irradiation, 15 mg/L of UVC-irradiated C60 resulted in approximately 2 log inactivation (99%) in 135 min. When a UV cutoff filter was used to remove the UVA below 400 nm, typically comprising about 4% of the total light energy emitted from the fluorescence lamps, the inactivation kinetics

Figure 2. Effect of (a) UVC exposure time for the preparation of UVCirradiated C60, (b) UVC-irradiated C60 concentration, and (c) FL light intensity on the kinetics of E. coli inactivation. Experimental condition: (a) UVC irradiation time = 3, 5, 7 days; FL irradiation intensity at 365 nm = 330 μW/cm2; [UVC-irradiated C60]0 = 15 mg/L; (b) UVC irradiation time = 7 days; FL irradiation (six lamps) intensity at 365 nm = 330 μW/cm2; [UVC-irradiated C60]0 = 5, 10, 15 mg/L; (c) UVC irradiation time = 7 days; FL irradiation (six lamps) intensity at 365 nm = 110, 220, 330 μW/cm2; [UVC-irradiated C60]0 = 15 mg/L.

was slightly decreased. The observed inactivation levels for 150 min FL exposure with and without UV cutoff filters were 1.95 and 2.4 log, respectively. It is noteworthy that UVCirradiated C60 inactivates bacteria in response to visible light 9629

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Figure 3. Effect of N2 bubbling and addition of ROS scavenger on the kinetics of E. coli inactivation. Inset: degradation of pCBA as a function of time. ([UVC-irradiated C60]0 = 15 mg/L, light intensity at 365 nm by six FLs = 330 μW/cm2).

and the observed inactivation primarily resulted from the visible light irradiation. Most experiments and discussions presented below, therefore, focus on visible light irradiation condition which is more environmentally relevant. At a constant concentration of UVC-irradiated C60 (15 mg/L), the E. coli inactivation rate increased as nC60 was treated for longer UVC exposure time (from 3 to 7 days) (Figure 2a). We have reported that as UVC exposure time increased, nC60 disaggregated into more hydrophilic products with concurrent decrease in aggregate size.15 The increased hydrophilicity and available surface area may have contributed to the enhanced E. coli inactivation rate. As the UVC-irradiated C60 concentration was increased from 0 to 15 mg/L, the inactivation at 150 min of exposure increased from 0 to 2.4 log (Figure 2b), exhibiting a linear relationship between the concentration and log inactivation achieved (R2 = 0.99 when log inactivation was plotted versus concentration; inset in Figure 2b). The E. coli inactivation rate also increased as the light intensity of the FLs was increased from 110 (2 lamps) to 330 μW/cm2 (6 lamps) (Figure 2c). A squareroot dependence was found for the observed E. coli inactivation for 150 min and the light intensity (R2 = 0.98 when log inactivation was plotted versus square root of intensity). This observation is in agreement with results of previous studies on photocatalytic microbial inactivation.31,32 The E. coli inactivation observed above was caused by a photocatalytic production of ROS by UVC-irradiated C60. When N2 gas was bubbled in order to remove the residual oxygen, only a slight level of E. coli inactivation was observed (Figure 3; 0.15 log for 150 min). The low level of inactivation observed in the N2 purged system might have been caused by residual oxygen which was not completely removed during N2 bubbling. Among ROS, 1 O2 and O2 3 were not responsible for E. coli inactivation. First, when excess (30 mM) 1O2 and O2 3 scavengers (L-histidine and superoxide dismutase (SOD)) were added, there was little change in the inactivation kinetics (Figure 3), suggesting that both were not a primary disinfection agent. Second, no measurable

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degradation of FFA, a probe for 1O2, was observed over the experimental time scale, while degradation of XTT (0.15 mM), a probe for O2 3 , as indicated by increase of absorption at 480 nm due to formation of purple-colored product, was observed. It should be noted that, although O2 3 is known to have a potential for inactivating E. coli,24,33 1O2 would not inactivate E. coli even if a detectable amount had been produced. Control tests were performed with Rose Bengal (10 μM) under FL irradiation for 60 min to independently produce 1O2 (e.g., Φ(1O2) of RB (550 nm) = 0.75 in water34). The results of control test suggested that as much as 6.4  10 5 mg-min/L28 of 1O2 exposure did not induce E. coli inactivation (data not shown), although the same level of exposure under identical condition would lead to 99% of MS2 bacteriophage inactivation.28 Detection of O2 3 indicates that O2 3 might have been produced as a precursor for 3 OH production. The fact that E. coli inactivation was not inhibited by SOD, therefore, may suggest that either 3 OH production is kinetically favored over quenching by SOD, or SOD did not effectively quench O2 3 that was produced inside the cell. Regardless, the observed E. coli inactivation was directly caused by photochemically produced 3 OH, since E. coli inactivation was virtually prohibited in the presence of excess (30 mM) 3 OH scavenger, t-butanol (t-BuOH). Results obtained with pCBA as an 3 OH probe is shown in the inset of Figure 3 and suggests steady-state concentration of 3 OH is approximately 5.62  10 15 M.38 The Mechanism of Inactivation. Mechanism of E. coli inactivation by 3 OH has been delineated as a sequence of nonselective reactions of 3 OH with major cell wall components such as proteins and lipids35 before it reaches inner cell components, followed by increase in cell wall permeability and subsequent 3 OH penetration into cytoplasm, and ultimately cell death.36,37 Surface protein release during cell death, therefore, is indirect evidence for cell surface destruction by 3 OH. For example, a control experiment was performed to inactivate E. coli to achieve 0.5 log inactivation by 3 OH produced via a different pathway (i.e., mixing 0.1 mg/L of O3 and 1 mg/L of H2O2). It was found that approximately 0.03 ( 0.0015 mg/L of protein, measured by Bradford assay, were released from cell surface components of 2  107 cfu/mL of E. coli. Similar phenomena were observed when E. coli was inactivated using even weaker disinfectants (oxidants) such as ozone and chlorine dioxide38 compared to 3 OH. For the same level of inactivation, approximately 0.02 and 0.01 mg/L of protein was released from the same concentration of E. coli. Note that the initial lag phase in the kinetics (Figures 1 and 2) is commonly attributed to the time required for the chemical disinfectant to disrupt the cell surface components before penetrating into the cytoplasm and reacting with vital cell components.39 However, the following lines of evidence collectively suggest that the mechanism of E. coli inactivation by 3 OH, photochemically produced from UVC-irradiated C60, may be different from the aforementioned pathway which involves cell surface disruption as a critical, initial step. First, E. coli inactivated up to 1 log showed no measurable protein release (i.e., below the detection limit of 0.001 mg/L) under the same condition in which protein release was observed by 3 OH produced in different methods. Second, the concentration of protein carbonyls, a commonly used marker of protein oxidation,40 also did not increase as E. coli was inactivated up to 1 log. Third, MDA release was also not observed (below the detection limit of 0.01 nmol/(mg cell dry wt)) during E. coli inactivation up to 1 log. MDA is one of the 9630

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Figure 4. Degradation of intracellular β-D-galactosidase as a function of reaction time ([UVC- irradiated C60]0 = 15 mg/L, light intensity at 365 nm by six FLs = 330 μW/cm2).

most abundant aldehydes formed from the peroxidation of the lipid membranes,41 primarily consisting of polyunsaturated phospholipids,26 and MDA release provides direct evidence of cell membrane denaturation. Fourth, change in cell permeability, which would result from damage or alteration of cell surface components, was examined using ONPG as a probe.29,36 The ONPG hydrolysis, as indicated by the chromogenic product formation (420 nm), results only when ONPG penetrates into the cell and reacts with intracellular E. coli enzymes such as β-Dgalactosidase.29,42 No measurable ONPG hydrolysis was observed for E. coli inactivated up to 2 log by UVC-irradiated C60, similarly to intact E. coli. In contrast, ONPG hydrolysis was as high as approximately 1500 μmol/(min-mg cell dry wt) at 0.6 log inactivation using ozone, equivalent to 69% of that obtained with lysed cells.43 While no evidence points toward cell surface damage as a preceding step for cell death, E. coli inactivation was accompanied by degradation of intercellular enzyme, β-D-galactosidase (approximately 80% over 2 log inactivation) (Figure 4). The reaction between 3 OH and β-D-galactosidase was nearly instantaneous in an independent in vitro control experiment. When t-BuOH was added to the suspension, β-D-galactosidase degradation was prohibited, suggesting that enzyme was degraded by 3 OH. Similarly, other intracellular enzymes were found to be degraded during E. coli inactivation by UVC-irradiated C60. The API-ZYM enzyme assay performed with adding E. coli after 1 log inactivation by UVC-irradiated C60 also suggested that five intercellular enzymes, alkaline phosphatase, leucine arylamidase, acid phosphatase, β-galactosidase, and β-glucuronidase, showed decreased enzymatic activity. The above results collectively suggest that 3 OH was the primary agent for E. coli inactivation but the mechanism does not involve cell surface damage that would be expected when 3 OH were produced in the bulk phase. Consequently, penetration of UVC-irradiated C60 into E. coli and generation of 3 OH within the cytoplasm, causing internal damage and consequently cell death, is suspected. In such a case, the initial lag in the

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Figure 5. Inactivation kinetics of intact E. coli after initially exposing to UV irradiated C60 for 60 min and spheroplast E. coli (19 ( 1 °C, pH 7.1, UVC exposure: 7 days, [UV irradiated C60]0 = 15 mg/L, light intensity at 365 nm by FL (six lamps): 330 μW/cm2).

kinetics discussed above could be related to the time required for UVC-irradiated C60 to penetrate into the cytoplasm, not the time required for 3 OH to cause sufficient cell surface damage. Consistently, when E. coli was pre-exposed to UVC-irradiated C60 under dark for 60 min (during which no inactivation was observed) and then exposed to FL irradiation, the lag phase was significantly reduced from approximately 40 to 15 min with little change in the postshoulder kinetics (Figure 5). Another experimental result obtained with E. coli spheroplast (i.e., E. coli with most outer membrane and peptidoglycan layer removed)44 also supports this hypothesis. The reduction in the lag phase in E. coli spheroplast was approximately the same as E. coli that was pre-exposed to UVC-irradiated C60. This mechanism partly explains why E. coli was inactivated by a much lower exposure to 3 OH compared to the reported 3 OH CT value. The CT stands for the product of disinfectant concentration and contact time and is widely used as a parameter to gauge the efficiency of disinfectant. The steady-state 3 OH concentration measured using pCBA was approximately 5.62  10 15 M45 (inset of Figure 3), which translates into an 3 OH CT (the product of radical concentration and contact time) to induce 1 log inactivation of E. coli of 0.86  10 8 mg/L-min.46 This value is 698 times less than the CT values obtained from 3 OH CT of 6.0  10 6 mg/ L-min measured via different photocatalyst (TiO2).45 Alternatively, 0.86  10 8 mg/L-min of CT would not be sufficient to disrupt the surface structure of E. coli if produced in the bulk phase. Environmental Significance. This study suggests that nC60 in the aqueous phase, upon hydrophilic transformation by UVC irradiation, exhibits a markedly different biocidal activity compared to nC60. The results are similar to C60 transformed by ozone treatment. Ozonation of nC60 resulted in the formation of soluble, multiple-oxygenated products which readily inactivated E. coli in the presence of UVA light (by BLB lamps) and oxygen.14 Similar to the mechanism presented herein, penetration of ozonated C60 into E. coli cytoplasm and photochemical production of 3 OH was suspected as the primary reason for the 9631

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Environmental Science & Technology cell death. The inactivation kinetics had similar characteristics (i.e., presence of lag phase), although a direct quantitative comparison is not possible, and the values of 3 OH CT were also very similar: 0.86  10 8 mg/L-min for UVC-irradiated C60 and 1.1  10 8 mg/L-min for ozonated C60. This study highlights that understanding the fate of nC60 with potential transformation by UVC as well as ozone, either in nature or in engineered systems such as water and wastewater treatment, and the ecological impact of the transformation products is critical to assess the overall environmental impact of C60. Whereas this study focused on the interaction of UVC transformation product with a representative microorganism, further studies are required to assess its interaction with other microorganisms and higher organisms as well as transport behaviors in the environment. Other oxidative transformation routes, particularly in engineered systems, are possible and have potential to transform C60 into potentially more toxic forms. Many C60 derivatives are intentionally being manufactured for particular applications and the concerns and potential impacts of these materials have yet to be assessed. Finally, we acknowledge that more direct evidence is required to firmly prove the hypothesis of E. coli inactivation by penetration of UVCirradiated C60, which is the focus of our currently ongoing study.

’ AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected]; phone: 404-894-2216; fax: 404- 385-7087.

’ ACKNOWLEDGMENT This work was partly supported by the National Science Foundation (Award CBET-0932872) and Korean National Research Foundation (Korean Ministry of Education, Science and Technology, Award NRF-2011-35B-D00020). ’ REFERENCES (1) Lyon, D. Y.; Fortner, J. D.; Sayes, C. M.; Colvin, V. L.; Hughes, J. B. Bacterial cell association and antimicrobial activity of a C60 water suspension. Environ. Toxicol. Chem. 2005, 24 (11), 2757–2762. (2) Lovern, S. B.; Klaper, R. Daphnia magna mortality when exposed to titanium dioxide and fullerene (C60) nanoparticles. Environ. Toxicol. Chem. 2006, 25 (4), 1132–1137. (3) Sayes, C. M.; Fortner, J. D.; Guo, W.; Lyon, D.; Boyd, A. M.; Ausman, K. D.; Tao, Y. J.; Sitharaman, B.; Wilson, L. J.; Hughes, J. B.; West, J. L.; Colvin, V. L. The differential cytotoxicity of water-soluble fullerenes. Nano Lett. 2004, 4 (10), 1881–1887. (4) Oberdorster, E.; Zhu, S. Q.; Blickley, T. M.; McClellan-Green, P.; Haasch, M. L. Ecotoxicology of carbon-based engineered nanoparticles: Effects of fullerene (C60) on aquatic organisms. Carbon 2006 44 (6), 1112–1120. (5) Lyon, D. Y.; Alvarez, P. J. J. Fullerene water suspension (nC60) exerts antibacterial effects via ROS independent protein oxidation. Environ. Sci. Technol. 2008, 42 (21), 8127–8132. (6) Andrievsky, G.; Klochkov, V.; Derevyanchenko, L. Is the C60 fullerene molecule toxic?! Fullerenes, Nanotubes, Carbon Nanostruct. 2005, 13 (4), 363–376. (7) Henry, T. B.; Menn, F. M.; Fleming, J. T.; Wilgus, J.; Compton, R. N.; Sayler, G. S. Attributing effects of aqueous C60 nano-aggregates to tetrahydrofuran decomposition products in larval zebrafish by assessment of gene expression. Environ. Health Perspect. 2007, 115 (7), 1059–1065. (8) Zhang, B.; Cho, M.; Fortner, J. D.; Lee, J.; Huang, C. H.; Hughes, J. B.; Kim, J. H. Delineating oxidative processes of aqueous C60

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preparations: Role of THF peroxide. Environ. Sci. Technol. 2009, 43 (1), 108–113. (9) Lee, J.; Fortner, J. D.; Hughes, J. B.; Kim, J. H. Photochemical production of reactive oxygen species by C60 in the aqueous phase during UV irradiation. Environ. Sci. Technol. 2007, 41 (7), 2529–2535. (10) Lee, J.; Kim, J. H. Effect of encapsulating agents on dispersion status and photochemical reactivity of C60 in the aqueous phase. Environ. Sci. Technol. 2008, 42 (5), 1552–1557. (11) Lee, J.; Yamakoshi, Y.; Hughes, J. B.; Kim, J. H. Mechanism of C60 photoreactivity in water: Fate of triplet state and radical anion and production of reactive oxygen species. Environ. Sci. Technol. 2008 42 (9), 3459–3464. (12) Isakovic, A.; Markovic, Z.; Nikolic, N.; Todorovic-Markovic, B.; Vranjes-Djuric, S.; Harhaji, L.; Raicevi, N.; Romcevic, N.; VasiljevicRadovic, D.; Dramicanin, M.; Trajkovic, V. Inactivation of nanocrystalline C60 cytotoxicity by gamma-irradiation. Biomaterials 2006, 27 (29), 5049–5058. (13) Fortner, J. D.; Kim, D. I.; Boyd, A. M.; Falkner, J. C.; Moran, S.; Colvin, V. L.; Hughes, J. B.; Kim, J. H. Reaction of water-stable C60 aggregates with ozone. Environ. Sci. Technol. 2007, 41 (21), 7497–7502. (14) Cho, M.; Fortner, J. D.; Hughes, J. B.; Kim, J. H. Escherichia coli inactivation by water-soluble, ozonated C60 derivative: Kinetics and mechanisms. Environ. Sci. Technol. 2009, 43 (19), 7410–7415. (15) Lee, J.; Cho, M.; Fortner, J. D.; Hughes, J. B.; Kim, J. H. Transformation of aggregate C60 in the aqueous phase by UV irradiation. Environ. Sci. Technol. 2009, 43 (13), 4878–4883. (16) Hou, W. C.; Jafvert, C. T. Photochemistry of aqueous C60 clusters: Evidence of 1O2 formation and its role in mediating C60 phototransformation. Environ. Sci. Technol. 2009, 43 (14), 5257–5262. (17) Hou, W. C.; Jafvert, C. T. Photochemical transformation of aqueous C60 clusters in sunlight. Environ. Sci. Technol. 2009, 43 (2), 362–367. (18) Hou, W. C.; Kong, L. J.; Wepasnick, K. A.; Zepp, R. G.; Fairbrother, D. H.; Jafvert, C. T. Photochemistry of aqueous C60 clusters: Wavelength dependency and product characterization. Environ. Sci. Technol. 2010, 44 (21), 8121–8127. (19) Guldi, D. M.; Prato, M. Excited-state properties of C60 fullerene derivatives. Acc. Chem. Res. 2000, 33 (10), 695–703. (20) Guldi, D. M.; Asmus, K. D. Photophysical properties of monoand multiply-functionalized fullerene derivatives. J. Phys. Chem. A 1997, 101 (8), 1472–1481. (21) Krusic, P. J.; Wasserman, E.; Keizer, P. N.; Morton, J. R.; Preston, K. F. Radical reactions of C60. Science 1991, 254 (5035), 1183–1185. (22) Isakovic, A.; Markovic, Z.; Todorovic-Markovic, B.; Nikolic, N.; Vranjes-Djuric, S.; Mirkovic, M.; Dramicanin, M.; Harhaji, L.; Raicevic, N.; Nikolic, Z.; Trajkovic, V. Distinct cytotoxic mechanisms of pristine versus hydroxylated fullerene. Toxicol. Sci. 2006, 91 (1), 173–183. (23) Bogdanovic, V.; Stankov, K.; Icevic, I.; Zikic, D.; Nikolic, A.; Solajic, S.; Djordjevic, A.; Bogdanovic, G. Fullerenol C60(OH)24 effects on antioxidative enzymes activity in irradiated human erythroleukemia cell line. J. Radiat. Res. 2008, 49 (3), 321–327. (24) Cho, M.; Chung, H. M.; Choi, W. Y.; Yoon, J. Y. Different inactivation behaviors of MS-2 phage and Escherichia coli in TiO2 photocatalytic disinfection. Appl. Environ. Microbiol. 2005, 71 (1), 270–275. (25) Hatchard, C. G.; Parker, C. A. A new sensitive chemical actinometer 0.2. potassium ferrioxalate as a standard chemical actinometer. Proc. R. Soc. London Ser. A 1956, 235 (1203), 518–536. (26) Maness, P. C.; Smolinski, S.; Blake, D. M.; Huang, Z.; Wolfrum, E. J.; Jacoby, W. A. Bactericidal activity of photocatalytic TiO2 reaction: Toward an understanding of its killing mechanism. Appl. Environ. Microbiol. 1999, 65 (9), 4094–4098. (27) Zor, T.; Seliger, Z. Linearization of the Bradford protein assay increases its sensitivity: Theoretical and experimental studies. Anal. Biochem. 1996, 236 (2), 302–308. (28) Cho, M.; Lee, J.; Mackeyev, Y.; Wilson, L. J.; Alvarez, P. J. J.; Hughes, J. B.; Kim, J. H. Visible light sensitized inactivation of MS-2 9632

dx.doi.org/10.1021/es202269r |Environ. Sci. Technol. 2011, 45, 9627–9633

Environmental Science & Technology

ARTICLE

bacteriophage by a cationic amine-functionalized C60 derivative. Environ. Sci. Technol. 2010, 44 (17), 6685–6691. (29) Zheng, H.; Maness, P. C.; Blake, D. M.; Wolfrum, E. J.; Smolinski, S. L.; Jacoby, W. A. Bactericidal mode of titanium dioxide photocatalysis. J. Photochem. Photobiol. A 2000, 130 (2 3), 163–170. (30) Chudnicka, A.; Matysik, G. Research of enzymatic activities of fresh juice and water infusions from dry herbs. J. Ethnopharmacol. 2005, 99 (2), 281–286. (31) Lee, S.; Nishida, K.; Otaki, M.; Ohgaki, S. Photocatalytic inactivation of phage Qβ by immobilized titanium dioxide mediated photocatalyst. Water Sci. Technol. 1997, 35 (11 12), 101–106. (32) Bekbolet, M. Photocatalytic bactericidal activity of TiO2 in aqueous suspensions of E. coli. Water Sci. Technol. 1997, 35 (11 12), 95–100. (33) van Hemmen, J. J.; Meuling, W. J. Inactivation of Escherichia coli by superoxide radicals and their dismutation products. Arch. Biochem. Biophys. 1977, 182 (2), 743–8. (34) Muraseccosuardi, P.; Gassmann, E.; Braun, A. M.; Oliveros, E. Determination of the quantum yield of intersystem crossing of rosebengal. Helv. Chim. Acta 1987, 70 (7), 1760–1773. (35) Berg, J. M.; Tymoczko, J. L.; Stryer, L. Biochemistry, 6th ed.; Freeman: New York, 2006. (36) Tsuchido, T.; Katsui, N.; Takeuchi, A.; Takano, M.; Shibasaki, I. Destruction of the outer-membrane permeability barrier of Escherichia coli by heat treatment. Appl. Environ. Microbiol. 1985, 50 (2), 298–303. (37) Arana, I.; Santorum, P.; Muela, A.; Barcina, I. Chlorination and ozonation of waste-water: Comparative analysis of efficacy through the effect on Escherichia coli membranes. J. Appl. Microbiol. 1999, 86 (5), 883–888. (38) Cho, M.; Yoon, J. Measurement of OH radical CT for inactivating Cryptosporidium parvum using photo/ferrioxalate and photo/TiO2 systems. J. Appl. Microbiol. 2008, 104 (3), 759–766. (39) Driks, A. Bacillus subtilis spore coat. Microbiol. Mol. Biol. R. 1999, 63 (1), 1–20. (40) Dean, R. T.; Fu, S. L.; Stocker, R.; Davies, M. J. Biochemistry and pathology of radical-mediated protein oxidation. Biochem. J. 1997, 324, 1–18. (41) Wheatley, R. A. Some recent trends in the analytical chemistry of lipid peroxidation. TrAC, Trends Anal. Chem. 2000, 19 (10), 617–628. (42) Matthews, B. W. The structure of E. coli beta-galactosidase. Cr. Biol. 2005, 328 (6), 549–556. (43) Cho, M.; Kim, J.; Kim, J. Y.; Yoon, J.; Kim, J. H. Mechanisms of Escherichia coli inactivation by several disinfectants. Water Res. 2010 44 (11), 3410–3418. (44) Sullivan, C. J.; Morrell, J. L.; Allison, D. P.; Doktycz, M. J. Mounting of Escherichia coli spheroplasts for AFM imaging. Ultramicroscopy 2005, 105 (1 4), 96–102. (45) Cho, M.; Lee, Y.; Chung, H.; Yoon, J. Inactivation of Escherichia coli by photochemical reaction of ferrioxalate at slightly acidic and nearneutral pHs. Appl. Environ. Microbiol. 2004, 70 (2), 1129–1134. (46) Elovitz, M. S.; von Gunten, U.; Kaiser, H. P. Hydroxyl radical/ ozone ratios during ozonation processes. II. The effect of temperature, pH, alkalinity, and DOM properties. Ozone-Sci. Eng. 2000, 22 (2), 123–150.

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dx.doi.org/10.1021/es202269r |Environ. Sci. Technol. 2011, 45, 9627–9633