Photoinduced toxicity of CdSe/ZnS quantum dots with ...

Int. J. Nanotechnol., Vol. 10, No. 12, 2013

Photoinduced toxicity of CdSe/ZnS quantum dots with different surface coatings to Escherichia coli Kavitha Pathakoti, Huey-Min Hwang*, Xiaojun Wang and Winfred G. Aker Department of Biology, Jackson State University, Jackson, MS 39217, USA Fax: +1-601-9796856 E-mail: [email protected] E-mail: [email protected] E-mail: [email protected] E-mail: [email protected] *Corresponding author Abstract: Quantum dot (QD) nanoparticles are increasingly used for various biomedical applications. To increase their biocompatibility, QDs are frequently coated with different surface coatings. Since QDs are photoactive under UV irradiation and may lead to oxidative stress, in this study the photoinduced toxicities of CdSe/ZnS QDs with three different coatings at three different wavelengths (530, 580 and 620 nm), along with a reference sample, a non-cadmium based QD CuInS2/ZnS were tested with E. coli under solar irradiation and dark conditions. QEI QDs coated with polyethylenimine (PEI) were found to be highly phototoxic to E. coli, conversely, QSA QDs with polyethyleneglycol coating and QSH QDs with amphiphilic polymer coating did not show toxicity. LC50 values for QEI QDs are smaller in the light exposure groups than those in the dark exposure groups. CuInS2/ZnS was less toxic to E. coli both under the light and dark conditions. The release of cadmium ions from the QD core was found to be negligible. Instead, the primary mechanism of the phototoxicity of QDs is the oxidative stress via formation of hydroxyl radicals that leads to lipid peroxidation and a concomitant decrease in reduced glutathione. The results also indicated that CdSe/ZnS QDs with PEI coating are highly toxic to E. coli and the non-cadmium based QD CuInS2/ZnS have better biocompatibility. Keywords: quantum dots; toxicity; E. coli; lipid peroxidation; reduced glutathione. Reference to this paper should be made as follows: Pathakoti, K., Hwang, H.M., Wang, X. and Aker, W.G. (2013) ‘Photoinduced toxicity of CdSe/ZnS quantum dots with different surface coatings to Escherichia coli’, Int. J. Nanotechnol., Vol. 10, No. 12, pp.1093–1108. Biographical notes: Kavitha Pathakoti is a Postdoctoral Research Associate at Jackson State University, Jackson, MS, USA. She received her Bachelor of Sciences from Osmania University in 1996 and MS in Environmental Sciences from Kakatiya University, India in 2003. She earned her PhD in Environmental Sciences in 2010 from Indian Institute of Chemical Technology, Hyderabad, India. Her research interests include environmental toxicology, biochemical toxicology, enzyme kinetics, oxidative stress biology and nanotoxicology.

Copyright © 2013 Inderscience Enterprises Ltd.

1093

1094

K. Pathakoti et al. She has authored or co-authored 17 papers in peer-reviewed journals. Currently, she is a reviewer for journals such as Science of the Total Environment, Toxicology and Environmental Chemistry and Nanoscale Research Letters. Huey-Min Hwang serves as Professor of Biology and Director of Environmental Science Master Program of Jackson State University (JSU). He received his PhD in Microbiology from the University of Georgia in 1985 and conducted his postdoctoral studies there from 1986 to 1989. He obtained his Bachelor of Science in Biology from National Taiwan Normal University in 1975. He was awarded the Outstanding Faculty Honoree of JSU and recognised by the HEADWAE Program of the State of Mississippi in 2002. His research interests include nanotoxicity, environmental toxicology, bioremediation, aquatic photochemistry and renewable energy. Currently, he serves as a guest editor or reviewer for journals such as Chemosphere, Environmental Toxicology and Chemistry, Environmental Toxicology, Journal of Photochemistry and Photobiology B: Biology and Environmental Science & Technology. He has authored or co-authored of over 90 publications in peer reviewed journals/books. Xiaojun Wang received her BS from China Agricultural University in 2007. She received her MS from Mississippi State University in 2010. Her research interests include cell culture and microbiological techniques. She was working as a Research Associate at Jackson State University. Winfred G. Aker received his BS in Meteorology from Purdue University, West Lafayette, Indiana, USA, in 1975 while on active duty in the military. Upon completion of his naval career, he earned and AS degree in Computer Technology in 1993 from Southwestern Community College, Chula Vista, California, USA. In 2007, he received his MS in Environmental Science from Jackson State University, Jackson, Mississippi, USA. Currently, he is pursuing a doctorate in Environmental Science, also from Jackson State University, concentrating on the mechanisms of nanotoxicity of metal oxide nanoparticles in aquatic environments.

1

Introduction

Quantum dots (QDs) are fluorescent semiconductor nanocrystals with unique optical and electrical properties [1,2]. Due to their high photo-stability, broad excitation and size tunable narrow emission spectrum, QDs have been used in various medical applications such as cellular imaging, cell tracking, targeted gene and drug delivery, high sensitivity biological assays and cancer detection [3–8]. To increase the biocompatibility of QDs, during the synthesis process a variety of surface coatings or functional groups are added to QDs with a desired bioactivity and additional secondary coatings to improve the water solubility. The most common coatings include amphiphilic polymers because they simultaneously exploit the hydrophobic nature of the quantum dot surface for cellular binding and the hydrophilic coating for solubilisation in aqueous solutions [9,10]. The carboxyl and amine residues can be used for attachment of proteins, nucleic acids, and other macromolecules which may be useful for the biomedical applications of the QDs. In the earlier studies, application of a surface coating material was found to be a major determining factor in

Photoinduced toxicity of CdSe/ZnS quantum dots

1095

causing the cytotoxicity of CdSe QDs [11]. Release of the core metal ions could induce generation of reactive oxygen species (ROS), leading to oxidative stress to the impacted living tissues or organisms [12–14]. In addition, free radicals were suggested as an important mechanism of QD toxicity [15]. QDs are also highly reactive, and are known to be liable to photo- and air-oxidation. Therefore, measurement of photoinduced toxicity should be included in the assessment of QDs impact on environmental health. Microorganisms act as primary decomposers and have an important role in element cycling, degradation of pollutants, and maintenance of ecological balance [16,17]. Bacteria serve as the foundation of many food chains in natural environments and are the most important agents that mediate biogeochemical cycling of naturally occurring and engineered materials; therefore, it is ecologically important to use a bacterial model to elucidate nanotoxicity mechanisms, severity of toxicity and persistence of nanomaterials. In the present study, E. coli was used as a bacterial model to study the effect of QDs coated with polyethylenimine (PEI), polyethyleneglycol (PEG) and only an amphiphilic coating at three different wavelengths. The aim of this paper is to investigate the photoinduced cytotoxicity of CdSe/Zns QDs with different coatings at three different wave lengths (530, 580 and 620 nm) to E. coli. In addition, the release of cadmium ions, ROS, lipid peroxidation and reduced glutathione were studied to understand the possible mechanisms of QDs.

2

Materials and methods

2.1 Quantum dot (QD) nanoparticles All the QD nanoparticles were procured from Ocean NanoTech Inc. (Springdale, AR). In our study, we used three different groups of QD nanoparticles with different coatings, with Ocean’s catalogue #QEI, QSA, and QSH. Each group consisted of QDs with emission wavelengths at 530, 580 and 620 nm. The QDs were coated with three different surface coatings, polyethylenimine (PEI) for the QEI, amine-polyethyleneglycol (PEG) for the QSA and only an amphiphilic polymer coating with a monolayer of octadecylamine for the QSH group. In addition, non cadmium QD CuInS2/ZnS (CuInS225) with amphiphilic polymer coating with carboxylic acid groups was used as the reference compound for comparison to all the above cadmium based QDs. The zeta potential of the QEI was about +50 mV and the total thickness of the organic layers was about 10 nm. The zeta potential of QSA ranged from –10 mV to +10 mV. The thickness of the total organic layers was about 6 nm. The zeta potential of QSH ranged from –30 mV to –50 mV and the thickness of the total organic layers was about 4 nm. The other physicochemical parameters of the study QDs are given in Table 1.

2.2 Transmission electron microscopy (TEM) For TEM the samples were prepared by drop-coating the nanoparticle suspension onto a carbon-coated copper grid (Ted Pella, CA) and then the samples were dried overnight at room temperature. The samples were observed using a TEM (JEOL JEM-1011).

1096

K. Pathakoti et al.

Table 1

Physicochemical properties of quantum dots

QD

Core/shell size (nm) Hydrodynamic size (nm) Composition

QSH 580

6.50

9.36 ± 2.75

QSH 530

3.40

7.34 ± 2.31

QSH 620

8.30

11.74 ± 2.31

QSA 580

6.50

10.86 ± 2.47

QSA 530

3.40

8.56 ± 2.53

QSA 620

8.30

12.56 ± 2.32

QEI 580

6.50

10.7 ± 3.69

QEI 530

3.40

8.19 ± 2.54

QEI 620

8.30

12.78 ± 3.24

CuInS2-25

7.5

11.14±2.31

CdSe/ZnS plus amphiphilic polymer coating with COOH on the surface

CdSe/ZnS plus amphiphilic polymer coating and PEG coating with –NH2 on the surface CdSe/ZnS plus amphiphilic polymer and PEI coating with primary, secondary and tertiary amine on the surface CuInS2-25 plus polymer coating with COOH on the surface

2.3 Cell viability assay E. coli (Migula) Castellani and Chalmers strain was prepared at 37°C overnight using Luria-Bertani (LB) broth. Then, the cultures were centrifuged at 3220 g for 10 min and resuspended in sterilised physiological saline. Bacterial density was adjusted to 2.2 × 109 – 3.0 × 109 bacteria/mL as determined by colony forming units (CFU) counting on LB Petri dishes. The stock solutions of the QDs were diluted to the various concentrations in 2 mL of distilled water. Hundred µL aliquots of freshly washed bacterial suspensions were added to the diluted solutions. The samples were exposed to midday sunlight for 30 min with agitation in a waterbath. Another group of the same samples were exposed under dark conditions wrapped with aluminium foil. (Solar irradiation outdoors: Irradiance: UVA range = 3.979 – 4.652 mW/cm2; UVB range = 3.1 – 3.7 MED/h). After 30 min of exposure time, the bacterial viability was determined using CFU counting on LB Petri dishes [18]. The cytotoxicity of the test QDs in terms of LC50 (the concentration of the nanoparticles proved to be fatal to 50% of the bacterium E. coli) was determined by the probit analysis method [19]. For all the other experiments i.e., cadmium ion measurement, ROS, lipid peroxidation (LPO), reduced glutathione (GSH), cell membrane integrity assay etc., the concentrations were chosen based on the LC50 values obtained from the cell viability assay.

2.4 Measurement of cadmium ions E. coli were exposed to different concentrations of QD samples for 30 min both under light and dark conditions (concentrations were chosen based on the LC50 values). After exposure, the solutions were filtered through 0.2 µm membrane filters (Corning Incorporated, Germany) and then solutions were digested with nitric acid. The final concentration of the nitric acid was less than 5% and the samples were further analysed by Inductively Coupled Plasma Mass Spectroscopy (ICP-MS, Varian Model No. 820-MS).


1097

2.5 Determination of reactive oxygen species (ROS) Intracellular ROS in the bacterial cells were measured using 3′-(p aminophenyl) fluorescein (APF) as described by Kim et al. [20]. APF has been recently used for detecting intracellular hydroxyl radicals (•OH), because they are resistant to auto-oxidation and do not respond to other oxidants such as singlet oxygen (1O2), superoxide radical (•O2–), and hydrogen peroxide (H2O2), in contrast with other fluorescent probes. The compound APF is not very fluorescent. However, when reacted with •OH it is converted into a strongly fluorescent compound [21,22]. The stock E. coli suspension was incubated with 10 µM APF for one hour in dark and then washed with PBS to remove excess dye after centrifugation. Then the E. coli suspension was adjusted to 2 × 107 CFU/mL and 50 µL aliquots of E. coli solutions loaded with fluorescent probes were added to 1 mL of various concentrations of QD nanoparticles respectively. Then 100 µL of the samples were added to 96-well plates and exposed to light for 30 min. The fluorescence intensity was measured before and after exposure with a micro plate reader (Triad series; Dynex Technologies, Chantilly, VA, USA) with 485 nm excitation and 535 nm emission filter. The formula [F(Final) – F(initial)]/F(Initial) × 100 was applied to calculate the percent increase in fluorescence. The data were normalised and expressed as fold increase in fluorescence intensity with respect to the controls (cells not treated). Similarly, the abiotic stress was measured without the E. coli.

2.6 Lipid peroxidation (LPO) LPO was determined based on the reaction of thiobarbituric acid (TBA) with malondialdehyde (MDA) to form a MDA-TBA adduct, referred as the peroxidation of lipids by the method of Esterbauer and Cheeseman [23]. One millilitre of treated bacterial culture was mixed with 2 mL of 10% (wt/vol) trichloroacetic acid and kept at room temperature for 10 min. The samples were centrifuged at 5000 g for 40 min to pellet the precipitated proteins and nanoparticles. Then 3 mL 0.67% TBA was added to the supernatants and incubated in a boiling water bath for 10 min. The samples were cooled and the absorbance was read at 532 nm in a micro plate reader (Triad series; Dynex Technologies, Chantilly, VA, USA). A standard curve was constructed based on the known amount of commercially bought malondialdehyde to the measured absorbance. The values were calculated as nanomoles of MDA formed per milligram (wet weight) of cells and the data was normalised as percent increase in MDA formed with respect to controls.

2.7 Reduced glutathione (GSH) GSH was estimated by the method previously described by Ellman [24]. Treated and untreated bacterial cells were disrupted by sonication in ice-cooled water and the supernatants were collected after centrifugation. Briefly, 50 µL of the supernatant was added to 0.2 mL of Tichloroacetic acid (5%) and then 0.25 mL of Ellman’s reagent (0.0198% DTNB in 1% sodium citrate) and the final assay volume was made up to 1 mL with phosphate buffer. The colour developed was read at 412 nm in a micro plate reader (Triad series; Dynex Technologies, Chantilly, VA, USA). The amount of glutathione was calculated using a GSH standard curve and calculated as nanomoles of GSH formed/mg protein. The data was normalised with respect to controls and expressed as percent

1098

K. Pathakoti et al.

decrease in GSH formed. Protein was estimated by the method of Bradford [25], using BSA as standard.

2.8 Membrane permeabilisation by propidium iodide (PI) uptake of cells Briefly, the treated bacterial cells were washed with PBS and incubated with 15 µM PI dye for 15 min at 37°C in the dark. Untreated samples and phosphate buffer were taken as control. The fluorescence was measured in a spectrofluorometer (Triad series; Dynex Technologies, Chantilly, VA, USA) with 485 nm excitation and 635 nm emission filters. Isopropanol treated cells were used as positive control and they gave the highest fluorescence intensity reading, fluorescence of all treatments was reported as a percentage of the fluorescence intensity units of this control. To prepare isopropanol-treated control, 1 mL of the cell suspension was dispensed into a 40-mL centrifuge tube containing 20 mL of isopropanol (70%). The mixture was incubated at 25°C for 1 h, with mixing every 15 min. Treated cells were washed twice by centrifugation for 15 min at 3200 g and 20°C, then resuspended in filter-sterilised 0.1% NaCl. Isopropanol-treated cells were adjusted to OD600 values similar to those of live suspensions.

2.9 Statistical analysis Data are expressed as mean ± standard deviation (SD) of three independent experiments, except the LC50 values in Table 2, which are expressed as LC50 ± Standard Error (SE). Differences between the control and treated samples were tested for statistical significance using unpaired Student’s t-test and values were considered to be significantly different when p < 0.05. Table 2

Cell viability of quantum dots against E. coli after exposure to sunlight and dark treatment

QDs QEI 580

Light LC50 ± SE

95% confidence limits (LL–UL)

Dark LC50±SE

95% confidence limits (LL–UL)

0.01 ± 0.0006

0.0095–0.012

0.063 ± 0.005

0.053–0.073

QEI 530

0.066 ± 0.006

0.0537–0.079

0.111 ± 0.022

0.066–0.156

QEI 620

0.015 ± 0.0008

0.066–0.156

0.029 ± 0.002

0.024–0.0358

32.46 ± 1.42

29.66–35.26

424 ± 3.17

418.13–430.59

CuInS2-25

*LL: lower limit; UL: upper limit.

3

Results

3.1 Cell viability The photoinduced toxicity of cadmium based QDs with three different coatings at three different wave lengths (530, 580 and 620 nm) were studied in E. coli both under the light and dark conditions, in comparison to the reference sample, a non-cadmium based QD CuInS2-25. The QSA and QSH QDs did not show any cytotoxicity to E. coli cells at the test concentrations up to 1 µM. Conversely, QEI QDs were highly toxic to E. coli both


1099

under the light and dark conditions. Figure 1 shows the representative TEM pictures of QEI QDs and all the QDs are highly stable and are in dispersed state. LC50 values are smaller in the light groups than those in the dark groups (Table 2). As for CuInS2-25, the non-cadmium based QD, extent of its cytotoxicity was much lower than the cadmium-based QDs under sunlight irradiation or in dark condition. Based on the LC50 mean values, rank of the cytotoxicity in light and dark exposure groups is QEI 580 > QEI 620 > QEI 530 > CuInS2-25 and QEI 620 > QEI 580 > QEI 530 > CuInS2-25, respectively. The viability count was 16% lower in the light control group compared to the dark control group (data not shown), indicating that the light inhibition occurs under outdoor irradiation. Based on these results, the cadmium based QEI QDs were chosen to further investigate the possible mechanisms of toxicity. Figure 1

TEM images of quantum dots (A) QEI 580 (250,000 X); (B) QEI 530 (300,000 X) and (C) QEI 620 (300,000 X)

3.2 Measurement of cadmium ions by ICPMS The release of cadmium ions in the test solution was measured by ICPMS and the results are presented in Figure 2. The results show that the cadmium concentrations are higher in the light groups, when compared to the dark counterparts. Figure 2

Cadmium (Cd) levels in the extracellular medium after E. coli cells were exposed to different concentrations of QDs for 30 min under light and dark conditions (see online version for colours)

1100

K. Pathakoti et al.

3.3 Role of ROS in exerting the toxicity of QDs ROS levels were measured by using the APF dye, mainly focusing on the •OH radicals and the data are presented in Figure 3. A significant concentration-dependent increase in intracellular ROS levels was observed in E. coli cells exposed to all the three QDs after light exposure. An increase of 99%, 62% and 97% increase in intracellular ROS was observed in QEI 530, QEI 580 and QEI 620 respectively at the highest concentrations tested. Similarly, increase in abiotic ROS was also observed at the same concentrations in all the three QDs (Figure 4). Conversely, after the dark exposure no significant increase in ROS levels was observed under abiotic condition nor in E. coli cells.

3.4 Role of LPO in exerting the toxicity of QDs A significant concentration-dependent increase (P < 0.05) in LPO, as indicated by MDA formation was observed in E. coli cells exposed to all the three QDs. However, the maximum percentage of lipid peroxidation induction was observed in the light groups at the highest test concentrations with increase of 53%, 57%, and 42% in QEI 580, QEI 530 and QEI 620, respectively, when compared to control (Figure 5). At the same concentrations, lipid peroxidation levels were significantly lower in E. coli in dark conditions. It is noteworthy that LPO activity was 13% higher in the light control group (80.08 ± 1.88 nM MDA/mg wet weight) compared to the dark control group (69.65 ± 0.49 nM MDA/mg wet weight. Figure 3

Level of intracellular ROS (hydroxyl radicals) after E. coli cells were exposed to various concentrations of QDs under solar irradiation (see online version for colours)

*Statistically significant from the control at p < 0.05.


1101

Figure 4

Abiotic production of ROS (hydroxyl radicals) after exposure to various concentrations of QDs under solar irradiation (see online version for colours)

Figure 5

Lipid peroxidation (% of control) in E. coli cells after exposure to QD nanoparticles under light (solar irradiation) and dark conditions (see online version for colours)

*Statistically significant from the relevant control at p < 0.05.

1102

K. Pathakoti et al.

3.5 Intracellular GSH levels after QD exposure E. coli cells exposed to all the three QDs showed reduction in GSH levels in a concentration-dependent manner. A reduction of 50%, 45% and 34% was observed in QEI 580, QEI 530 and QEI 620, respectively, when compared to control in the highest tested concentrations in the light group (Figure 6). Similarly, the GSH reduction levels were generally less in the dark groups compared to those of the light groups. Figure 6

Reduced glutathione (% of control) in E. coli cells after exposure to QD nanoparticles under light (solar irradiation) and dark conditions (see online version for colours)

*Statistically significant from the relevant control at p < 0.05.

3.6 Cell membrane integrity Cell membrane integrity was assessed using PI, the positive control isopropyl alcohol– treated cells showed highest fluorescence compared to the negative controls in E. coli cells (Figure 7). The cells treated with various concentrations of QDs for 30 min did not exhibit statistically significant increase in fluorescence, indicating that cell membranes were not damaged. As there are no significant differences between the control and treated cells when compared to the positive control, only the data of the highest tested concentrations are presented in Figure 7.

Photoinduced toxicity of CdSe/ZnS quantum dots Figure 7

4

1103

Measurement of cell membrane integrity by using propidium iodide in E. coli cells after treatment with QDs. Isopropanol treated cells were used as positive controls (+ve control) (see online version for colours)

Discussion

The unique fluorescent properties of QDs enable them to be used in cell imaging and cancer targeting. Recently, there has been an emergence of the use of QDs for the detection of bacteria, Escherichia coli [26]. The coating of QDs plays a key role in their biocompatibility, stability, weathering, and mobility [27]. A series of QDs coated with three different coatings (PEG, PEI and an amphiphilic coating) at different wavelengths were used in this study to investigate the mechanisms of their cytotoxicity by using the well known bacterial model, E. coli. Bacteria are one of the most susceptible living organisms to photo-induced damage [28]. Among them, E. coli has been used as an alternate test system for phototoxicity studies [29]. QSA QDs with amine-PEG coating and QSH QDs with only the amphiphilic polymer coating (carboxyl group on the surface) did not show any cytotoxicity to the E. coli cells at the test concentrations up to 1 µM. It was also observed that the CdSe/ZnS QEI QDs with PEI coating are highly toxic to E. coli after exposure to solar irradiation, when compared to those in dark conditions. Earlier studies also reported that UV exposure could result in increasing the cytotoxicity of CdSe QDs [20,30,31]. Adams et al. [32] reported that the bacterial growth inhibition was observed under illumination and the killing in darkness was also observed but at a reduced level. Among all the QDs QEI 580 was the most highly toxic to E. coli after light exposure while QEI 620 was the least toxic QD in this study. In contrast, QEI 620 was most highly toxic to E. coli under dark conditions, whereas CuInS2-25 was the least toxic. This finding is in agreement with the report from an in vivo study with mice that non-cadmium based CuInS2/Zn QD is less toxic to mice in comparison to CdTeSe/CdZnS QDs [33]. The higher toxicity of cationic PEI coated QDs can be attributed to their higher affinity for negatively charged bacteria [34]. The strong cationic property is responsible for inducing defects into lipid bilayers [35]. Earlier studies showed that CdTe QDs could effectively kill E. coli in a concentration-dependent manner [36]. It was also reported that the cytotoxicity of

1104

K. Pathakoti et al.

CdSe/ZnS QDs depended mainly on the surface coating material rather than the core materials [11,37]. In order to investigate the responsible mechanisms of QEI QD cytotoxicity, we measured the concentration of cadmium release in the medium, lipid peroxidation, reduced glutathione, cell membrane integrity and also the generation of ROS. The QD core metals could be released by oxidative and photolytic conditions and are known to be toxic at relatively low concentrations [38]. Cadmium ions are well known to cause cytotoxicity via several pathways; therefore, the cadmium ion released from the QD core material may be an important factor in influencing QD toxicity. QDs become unstable when exposed to UV light, eventually releasing Cd into the medium [31,38]. In the present study, the cadmium ion concentrations are higher in the light groups, when compared to the dark groups; however, the concentration levels appear too low to cause the toxicity. Coincidentally, it has been recently reported that the released metal ions are negligible and QD toxicity is mainly due to the specific function of the nanoparticles [39–41]. Our results are also in good agreement with the study by Kim et al. [20]. In that report, they indicated that the cadmium ion toxicity was negligible. ROS play an important role in causing the toxicity of nanoparticles by leading to secondary processes that ultimately cause cell damage (eg., lipid peroxidation, DNA damage and protein oxidation) and even cell death [12,42]. UV irradiation has the potential to generate a relatively high level of ROS during irradiation [43]. Lovric et al. [44] reported that ROS played the lead role in QD-induced cellular damage and the ROS generated from QDs under UV are known to be responsible for oxidative damage to sensitive biomolecules like DNA [45]. In the present study we observed a significant increase in intracellular ROS in E. coli cells in the form of •OH after light exposure in all of the three QDs. Formation of ROS under abiotic conditions was observed in our study, and the previous reports also showed that nanoparticles can exert oxidative effects under abiotic conditions [46–48]. After dark exposure there was no ROS formation in all the three QDs, suggesting some other mechanisms should account for the toxicity. Our results corroborate with the previous study by Dimpka et al. [49] in which they observed that the nanoparticles treated with APF dye do not produce any ROS. Toxicity has also been observed with nanoparticles in the absence of light, which suggests that toxicity may not be solely attributed to photoinduced ROS formation [32,50]. Lipid peroxidation is the first step of cellular damage and it is considered most deleterious, leading to alterations in cell membrane properties which in turn disrupt vital cellular functions [51,52]. Elevated lipid peroxidation levels were observed after exposure to QDs in the light groups, confirming that the phototoxicity is related to ROS generation by the QD nanoparticles. Glutathione is the major low-molecular-weight, non-enzymatic antioxidant which quenches oxyradicals through its sulfhydryl group. It also constitutes the first line of the cellular defense mechanism against oxidative injury and is the major intracellular redox buffer in ubiquitous cell types [53]. GSH is also a vulnerable target in cell defense, as different agents (environmental and other) can directly deplete or inactivate GSH. The depletion of GSH causes an imbalance of pro-oxidants (e.g. ROS) and antioxidants (e.g., GSH) leading to increased oxidative stress [54]. Hence, the depletion of GSH levels leading to lipid peroxidation is a major mechanism involved in QDs toxicity. To further understand the mechanisms of QD toxicity, cell membrane integrity was assessed by using the PI dye, which binds only to the damaged or dead cells [55]. The present results show that the cell membrane integrity is not affected by the QDs


1105

treatment on the E. coli cells. Earlier studies also reported that the cell membrane integrity is not affected by the nanoparticles [56,57]. The cytotoxicity mechanisms involved in the light exposure are the increase in ROS production and LPO with a concomitant decrease in GSH causing oxidative stress in E. coli cells. Under dark conditions, LPO and depletion of GSH are the mechanisms of toxicity. The release of cadmium ion and cell membrane damage do not account for the toxicity of these PEI coated QDs. Our results are in line with a recent study by Yang et al. [58], in which they showed that the QD toxicity is due to the potentially inhibitory binding material like PEI coating, rather than the released metals.

5

Conclusions

Overall, our results suggest that CdSe/ZnS QDs with PEI coating are more highly phototoxic to E. coli than the non-cadmium based CuInS2/ZnS. Intracellular ROS, LPO, GSH depletion are the mechanisms of for bacterial toxicity under light conditions. Nevertheless, the cell membrane integrity of the E. coli cells is not affected by the QDs. The inhibitory effects observed under the dark conditions suggest that lipid peroxidation with concomitant decrease in GSH are the mechanisms of toxicity. The toxicity caused by the QDs can be attributed to PEI surface coating which is highly toxic to the bacteria in comparison to the PEG coating and the amphiphilic coating. To overcome these problems, modification of PEI backbone with PEG should be used to lower the toxicity with increased stability. Additionally, use of non-cadmium based QD CuInS2/ZnS will result in better biocompatibility with less cytotoxicity. These findings are not only useful in promoting biomedical application of QDs, but also beneficial in the future design for biologically safe QDs.

Acknowledgements This study was supported by (1) NSF-SBIR grant # IIP-0823040 and (2) NSF-CREST program with grant #HRD-0833178. We wish to thank Dr. Andrew Wang of Ocean NanoTech Inc. (Springdale, AR) for the characterisation of QDs. We wish to thank Dr. Zikri Arslan for providing assistance in cadmium ion analysis with ICP-MS system. We thank Dr. Ming-Ju Huang for the helpful discussion in data analysis. In this study we used the JSU Molecular and Cellular Biology Core Lab which is supported by NIH-RCMI grant (#G12RR013459-13).

References 1 2

3

Bruchez, M., Moronne, M., Gin, P., Weiss, S. and Alivisatos, A.P. (1998) ‘Semiconductor nanocrystals as fluorescent biological labels’, Science, Vol. 281, pp.2013–2016. Dabbousi, B.O., Rodriguez-Viejo, J., Mikulec, F.V., Heine, J.R., Mattoussi, H., Ober, R., Jensen, K.F. and Bawendi, M.G. (1997) ‘(CdSe)ZnS Core-shell quantum dots: synthesis and characterization of a size series of highly luminescent nanocrystallites’, J. Phys. Chem. B, Vol. 101, pp.9463–9475. Day R.N., Periasamy, A. and Schaufele, F. (2001) ‘Fluorescence resonance energy transfer microscopy of localized protein interactions in the living cell nucleus’, Methods, Vol. 25, pp.4–18.

1106 4

5

6

7

8

9

10

11 12 13

14

15 16 17 18

19 20

21

22

K. Pathakoti et al.

Kaul, Z, Yaguchi, T, Kaul, S.C., Hirano, T., Wadhwa, R. and Taira, K. (2003) ‘Mortalin imaging in normal and cancer cells with quantum dot immuno-conjugates’, Cell Res., Vol. 13, pp.503–507. Dubertret, B., Skourides, P., Norris, D.J., Noireaux, V., Brivanlou, A.H. and Libchaber, A. (2002) ‘In vivo imaging of quantum dots encapsulated in phospholipid micelles’, Science, Vol. 298, pp.1759–1762. Pathak, S., Choi, S.K., Arnheim, N. and Thompson, M.E. (2001) ‘Hydroxylated quantum dots as luminescent probes for in situ hybridization’, J. Amer. Chem. Soc., Vol. 123, pp.4103–4104. Lee, L.Y., Ong, S.L., Hu, J.Y., Ng, W.J., Feng, Y., Tan, X. and Wong, S.W. (2004) ‘Use of semiconductor quantum dots for photostable immunofluorescence labeling of Cryptosporidium parvum’ App. Environ. Microbio., Vol. 70, pp.5732–5736. Brown, E., Mckee, T., diTomaso, E., Pluen A, Seed, B., Boucher, Y. and Jain, R.K. (2003) ‘Dynamic imaging of collagen and its modulation in tumors in vivo using second-harmonic generation’, Nat. Med., Vol. 9, pp.796–800. Wu, X., Liu, H., Liu, J., Haley, K.N., Treadway, J.A., Larson, J.P., Ge, N., Peale, F. and Bruchez, M.P. (2003) ‘Immunofluorescent labeling of cancer marker Her2 and other cellular targets with semiconductor quantum dots’, Nat. Biotechnol., Vol. 21, pp.41–46. Gussin, H.A., Tomlinson, I.D., Little, D.M., Warnement, M.R., Qian, H., Rosenthal, S.J. and Pepperberg, D.R. (2006) ‘Binding of muscimol-conjugated quantum dots to GABAc receptors’, J. Amer. Chem. Soc., Vol. 128, pp.15701–15713. Gao, X., Cui, Y., Levenson, R.M., Chung, L.W. and Nie, S. (2004) ‘In vivo cancer targeting and imaging with semiconductor quantum dots’, Nat. Biotechnol., Vol. 22, pp.969–976. Nel, A., Xia, T., Madler, L. and Li, N. (2006) ‘Toxic potential of materials at the nanolevel’, Science, Vol. 311, pp.622–627. Brayner, R., Ferrari-Iliou, R., Brivois, N., Djediat, S., Benedetti, M.F. and Fievet, F. (2006). ‘Toxicological impact studies based on Escherichia coli bacteria in ultrafine ZnO nanoparticles colloidal medium’, Nano Lett., Vol. 6, pp.866–870. Lin, W., Stayton, I., Huang, Y.W., Zhou, X.D. and Ma, Y. (2008) ‘Cytotoxicity and cell membrane depolarization induced by aluminum oxide nanoparticles in human lung epithelial cells A549’, Toxicol. Environ. Chem., Vol. 90, pp.983–996. Rzigalinski, B.A. and Strobl, J.S. (2009) ‘Cadmium-containing nanoparticles: perspectives on pharmacology and toxicology of quantum dots’, Toxicol. Appl. Pharm., Vol. 238, pp.280–288. Osler, G.H. and Sommerkorn, M. (2007) ‘Toward a complete soil C and N cycle: incorporating the soil fauna’, Ecology, Vol. 88, pp.1611–1621. Kertesz, M.A. and Mirleau, P. (2004) ‘The role of soil microbes in plant sulphur nutrition’, J. Exp. Bot., Vol. 55, pp.1939–1945. Thill, A., Zeyons, O., Spalla, O., Chauvat, F., Rose, J., Auffan, M. and Flank, A.M. (2006) ‘Cytotoxicity of CeO2 nanoparticles for Escherichia coli: physico-chemical insight of the cytotoxicity mechanism’, Environ. Sci. Technol., Vol. 40, pp.6151–6156. Finney, D.J. (1971) Probit Analysis, 3rd ed., Cambridge University Press, Cambridge, UK. Kim, J., Park, Y., Yoon, T.H., Yoon, C.S. and Choi, K. (2010) ‘Phototoxicity of CdSe/ZnSe quantum dots with surface coatings of 3-mercaptopropionic acid or tri-n-octylphosphine oxide/gum arabic in Daphnia magna under environmentally relevant UV-B light’, Aquat. Toxicol., Vol. 97, pp.116–124. Setsukinai, K., Urano, Y., Kakinuma, K., Majima, H.J. and Nagano T. (2003) ‘Development of novel fluorescence probes that can reliably detect reactive oxygen species and distinguish specific species’, J. Biol. Chem., Vol. 278, pp.3170–3175. Soh, N. (2006) ‘Recent advances in fluorescent probes for the detection of reactive oxygen species’, Anal. Bioanal. Chem., Vol. 386, pp.532–543.

Photoinduced toxicity of CdSe/ZnS quantum dots 23 24 25

26 27 28

29

30 31 32 33

34

35

36 37

38

39 40

41

42

1107

Esterbauer, H. and Cheeseman, K.H. (1990) ‘Determination of aldehydic lipid peroxidation products: malonaldehyde and 4-hydroxynonenal’, Methods Enzymol., Vol. 186, pp.407–421. Ellman, G.L. (1959) ‘Tissue sulfhydryl groups’, Arch. Biochem. Biophys., Vol. 82, pp.70–77. Bradford, M.M. (1976) ‘A rapid and sensitive method for the quantification of microgram quantities of protein utilizing the principle of protein-dye binding’, Anal. Biochem., Vol. 72, pp.248–254. Zhu, H., Sikora, U. and Ozcan, A. (2012) ‘Quantum dot enabled detection of Escherichia coli using a cell-phone’, Analyst, Vol. 137, pp.2541–2544. Mahendra, S., Zhu, H.G., Colvin, V.L. and Alvarez, P.J. (2008) ‘Quantum dot weathering results in microbial toxicity’, Environ. Sci. Technol., Vol. 42, pp.9424–9430. Laura, A.S., Josep, M.G., Thomas, L., Julia, H. and Ruben S. (2006) ‘Effect of natural sunlight on bacterial activity and differential sensitivity of natural bacterioplankton groups in northwestern Mediterranean coastal waters’, Appl. Environ. Microbiol., Vol. 72, pp.5806–5813. Verma, K., Agrawal, N., Misra, R.B., Farooq, M. and Hans, R.K. (2008) ‘Phototoxicity assessment of drugs and cosmetic products using E. coli’, Toxicol. In. Vitro., Vol. 22, pp.249–253. Bakalova, R., Ohba, H., Zhelev, Z., Ishikawa, M. and Baba, Y. (2004) ‘Quantum dots as photosensitizers?’, Nat. Biotechnol., Vol. 22, pp.1360–1361. Derfus, A.M., Chan, W.C.W. and Bhatia, S.N. (2004) ‘Probing the cytotoxicity of semiconductor quantum dots’, Nano Lett., Vol.4, pp.11–18. Adams, L.K., Lyon, D.Y. and Alvarez, P.J. (2006) ‘Comparative eco-toxicity of nanoscale TiO2, SiO2, and ZnO water suspensions’, Water Res., Vol. 40, pp.3527–3532. Pons, T., Pic, E., Lequeux, N., Cassette, E., Bezdetnaya, L., Guilleman, F., Marchal, F. and Dubertret, B. (2010) ‘Cadmium-free CuInS2/ZnS quantum dots for sentinel lymph node imaging with reduced toxicity’, ACS Nano, Vol. 4, pp.2531–2538. Yang, Y., Zhu, H., Colvin, V.L. and Alvarez, P.J. (2011) ‘Cellular and transcriptional response of pseudomonas stutzeri to quantum dots under aerobic and denitrifying conditions’, Environ. Sci. Technol., Vol. 45, pp.4988–4994. Leroueil, P.R., Berry, S.A., Duthie, K., Han, G., Rotello, V.M., McNerny, D.Q., Baker Jr., J.R., Orr, B.G. and Holl, M.M. (2008) ‘Wide varieties of cationic nanoparticles induced defects in supported lipid bilayers’, Nano Lett., Vol. 8, pp.420–424. Lu, Z., Li, C.M., Bao, H., Qiao, Y., Toh, Y. and Yang, X. (2008) ‘Mechanism of antimicrobial activity of CdTe quantum dots’, Langmuir, Vol. 24, pp.5445–5452. Hoshino, A., Hanaki, K., Suzuki, K. and Yamamoto, K. (2004) ‘Applications of T-lymphoma labeled with fluorescent quantum dots to cell tracing markers in mouse body’, Biochem. Biophys. Res. Commun., Vol. 314, pp.46–53. Hardman, R. (2006) ‘A toxicologic review of quantum dots: toxicity depends on physicochemical and environmental factors’, Environ. Health Perspect., Vol. 114, pp.165–172. Jiang, W., Mashayekhi, H. and Xing B. (2009) ‘Bacterial toxicity comparison between nanoand micro-scaled oxide particles’, Environ. Pollut., Vol. 157, pp.1619–1625. Priester, J.H., Stoimenov, P.K., Mielke, R.E., Webb, S.M., Ehrhardt, C., Zhang, J.P., Stucky, G.D. and Holden, P.A. (2009) ‘Effects of soluble cadmium salts versus CdSe quantum dots on the growth of planktonic Pseudomonas aeruginosa’, Environ. Sci. Technol., Vol. 43, pp.2589–2594. Baek, Y.W. and An, Y.J. (2011) ‘Microbial toxicity of metal oxide nanoparticles (CuO, NiO, ZnO, and Sb2O3) to Escherichia coli, Bacillus subtilis, and Streptococcus aureus’, Sci. Total Environ., Vol. 409, pp.1603–1608. Imlay, J.A. (2003) ‘Pathways of oxidative damage’, Annu. Rev. Microbiol., Vol. 57, pp.395–418.

1108 43

44

45 46

47

48

49

50

51

52

53

54

55 56 57

58

View publication stats

K. Pathakoti et al.

Clarke, S., Nadeau, J., Bahcheli, D., Zhang, Z. and Hollmann, C. (2005) ‘Quantum dots as phototoxic drugs and sensors of specific metabolic processes in living cells’, Conf. Proc. IEEE. Eng. Med. Biol. Soc., Vol. 1, pp.504–507. Lovric, J., Cho, S.J., Winnik, F.M. and Maysinger, D. (2005) ‘Unmodified cadmium telluride quantum dots induce reactive oxygen species formation leading to multiple organelle damage and cell death’, Chem. Biol., Vol. 12, pp.1227–1234. Green, M. and Howman, E. (2005) ‘Semiconductor quantum dots and free radical induced DNA nicking’, Chem. Commun., Vol. 1, pp.121–123. Applerot, G., Lipovsky, A., Dror, R., Perkas, N., Nitzan, Y., Lubart, R. and Gedanken, A. (2009) ‘Enhanced antibacterial activity of nanocrystalline ZnO due to increased ROS-mediated cell injury’, Adv. Funct. Mat., Vol. 19, pp.842–852. Xia, T., Kovochich, M., Brant J, Hotze, M., Sempf, J., Oberley, T., Sioustas, C., Yeh, J.I., Wiesner, M.R. and Nel, A.E. (2006) ‘Comparison of the abilities of ambient and manufactured nanoparticles to induce cellular toxicity according to an oxidative stress paradigm’, Nano Lett., Vol. 6, pp.1794–1807. Hussain, S., Boland, S., Baeza-Squiban, A., Hamel, R., Thomassen, L.C., Martens, J.A, Billon-Galland, M.A., Fleury-Feith, J., Moisan, F., Pairon, J.C. and Marano, F (2009) ‘Oxidative stress and proinflammatory effects of carbon black and titanium dioxide nanoparticles: role of particle surface area and internalized amount’, Toxicology, Vol. 260, pp.142–149. Dimkpa, C.O., Calder, A., Britt, D.W., McLean, J.E. and Anderson, A.J. (2011) ‘Responses of a soil bacterium, Pseudomonas chlororaphis O6 to commercial metal oxide nanoparticles compared with responses to metal ions’, Environ. Pollut., Vol. 159, pp.1749–1756. Daoud, W.A., Xin, J.H. and Zhang, Y.H. (2005) ‘Surface functionalization of cellulose fibers with titanium dioxide nanoparticles and their combined bactericidal activities’, Surf. Sci., Vol. 599, pp.69–75. Horie, M., Fukui, H., Nishio, K., Endoh, S., Kato, H., Fujita, K., Miyauchi, A., Nakamura, A., Shichiri, M., Ishida, N., Kinugasa, S., Morimoto, Y., Niki, E., Yoshida, Y. and Iwahashi, H. (2011) ‘Evaluation of acute oxidative stress induced by NiO nanoparticles in vivo and in vitro’, J. Occup. Health, Vol.53, pp.64–74. Rodea-Palomares, I., Gonzalo, S., Santiago-Morales, J., Leganés, F., García-Calvo, E., Rosal, R. and Fernández-Piñas, F. (2012) ‘An insight into the mechanisms of nanoceria toxicity in aquatic photosynthetic organisms’, Aquat. Toxicol., Vols. 122–123, pp.133–143. Meister, A. (1989) ‘Metabolism and function of glutathione’, in Dolphin, D., Poulson, R. and Auramovic, R. (Eds.): Glutathione: Chemical, Biochemical and Medical aspects. Part A, John Wiley and Sons, New York, pp.367–474. Brunet, L., Lyon, D.Y., Hotze, E.M., Alvarez, P.J. and Wiesner M.R. (2009) ‘Comparative photoactivity and antibacterial properties of C60 fullerenes and titanium dioxide nanoparticles’, Environ. Sci. Technol., Vol. 43, pp.4355–4360. Haugland, R.P. (1996) Handbook of Fluorescent Probes and Research Chemicals, Molecular Probes Inc., Eugene, Oregon. Spurlin, T.A. and Gewirth, A.A. (2007) ‘Effect of C60 on solid supported lipid bilayers’, Nano Lett., Vol. 7, pp.531–535. Lyon, D.Y. and Alvarex, P.J. (2008) ‘Fullerene water suspension (nC60) exerts antibacterial effects via ROS-independent protein oxidation’, Environ. Sci. Technol., Vol. 42, pp.8127–8132. Yang, Y., Wang, J., Zhu, H., Colvin, V.L. and Alvarez, P.J. (2012) ‘Relative susceptibility and transcriptional response of nitrogen cycling bacteria to quantum dots’, Environ. Sci. Technol., Vol. 46, pp.3433–3441.