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Mechanism of Antimicrobial Activity of CdTe Quantum Dots Zhisong Lu,†,‡ Chang Ming Li,*,†,‡ Haifeng Bao,†,‡ Yan Qiao,†,‡ Yinghui Toh,† and Xu Yang§ School of Chemical and Biomedical Engineering, Nanyang Technological UniVersity, 70 Nanyang DriVe, Singapore 637457, Singapore, Center for AdVanced Bionanosystems, Nanyang Technological UniVersity, 70 Nanyang DriVe, Singapore 637457, Singapore, and College of Life Science, Central China Normal UniVersity, Wuhan 430079, PR China ReceiVed December 31, 2007. ReVised Manuscript ReceiVed February 28, 2008 The antimicrobial activity and mechanism of CdTe quantum dots (QDs) against Escherichia coli were investigated in this report. Colony-forming capability assay and atomic force microscopy (AFM) images show that the QDs can effectively kill the bacteria in a concentration-dependent manner. Results of photoluminescence spectrophotometry, confocal microscopy, and antioxidative response tests indicate that the QDs bind with bacteria and impair the functions of a cell’s antioxidative system, including down-regulations of antioxidative genes and decreases of antioxidative enzymes activities. The oxidative damage of protein and lipid is also observed with thiobarbituric reacting substances and protein carbonyl assays, respectively. On the basis of these results, it is proposed that the mechanism of the antimicrobial activity of CdTe QDs involves QDs-bacteria association and a reactive oxygen species-mediated pathway. Thus, CdTe QDs could have the potential to be formulated as a novel antimicrobial material with excellent optical properties.
Introduction With the remarkable advances of nanotechnology in recent years, nanoparticles with unique chemical and physical properties have demonstrated an increasing importance in biological, biomedical, and pharmaceutical applications. Owing to their large specific surface area and high bioactivity, inorganic nanomaterials are regarded as good candidates to replace traditional organic antimicrobial agents that are extremely irritant and toxic. A number of nanoparticles with antimicrobial activities have been reported recently,1–9 of which silver nanoparticles have been well studied and reported to accumulate in the Escherichia coli (E. coli) membrane to possess effectively antibacterial effects.10 Metal oxide nanoparticles, another type of nanomaterials including titanium dioxide (TiO2), silicon dioxide (SiO2), and zinc oxide (ZnO) nanoparticles, also display excellent biocidal activities against both Gram-positive and Gram-negative bacteria.7 Although all these compounds are photosensitive and can generate reactive oxygen species (ROS) * Corresponding author. Tel.:+65 67904485; fax: +65 67911761. E-mail:
[email protected]. † School of Chemical and Biomedical Engineering, Nanyang Technological University. ‡ Center for Advanced Bionanosystems, Nanyang Technological University. § Central China Normal University. (1) Sondi, I.; Salopek-Sondi, B. J. Colloid Interface Sci. 2004, 275, 177–182. (2) Gogoi, S. K.; Gopinath, P.; Paul, A.; Ramesh, A.; Ghosh, S. S.; Chattopadhyay, A. Langmuir 2006, 22, 9322–9328. (3) Panacek, A.; Kvitek, L.; Prucek, R.; Kolar, M.; Vecerova, R.; Pizurova, N.; Sharma, V. K.; Nevecna, T.; Zboril, R. J. Phys. Chem. B 2006, 110, 16248– 16253. (4) Lyon, D. Y.; Adams, L. K.; Falkner, J. C.; Alvarezt, P. J. EnViron. Sci. Technol. 2006, 40, 4360–4366. (5) Kim, Y. H.; Lee, D. K.; Cha, H. G.; Kim, C. W.; Kang, Y. C.; Kang, Y. S. J. Phys. Chem. B 2006, 110, 24923–24928. (6) Rosemary, M. J.; MacLaren, I.; Pradeep, T. Langmuir 2006, 22, 10125– 10129. (7) Adams, L. K.; Lyon, D. Y.; Alvarez, P. J. Water Res. 2006, 40, 3527–3532. (8) Li, P.; Li, J.; Wu, C. Z.; Wu, Q. S.; Li, J. Nanotechnology 2005, 16, 1912–1917. (9) Lee, D.; Cohen, R. E.; Rubner, M. F. Langmuir 2005, 21, 9651–9659. (10) Lok, C. N.; Ho, C. M.; Chen, R.; He, Q. Y.; Yu, W. Y.; Sun, H.; Tam, P. K. H.; Chiu, J. F.; Che, C. M. J. Biol. Inorg. Chem. 2007, 12, 527–534.
under high-intensity light at a specific wavelength, only TiO2 exhibits a significant antibacterial activity when sunlight is applied as the excitation source.11,12 In comparison to TiO2 nanoparticles, quantum dots (QDs) have superior size-dependent optical properties and could be another type of attractive nanomaterials. QDs are nanoscale crystalline clusters synthesized from semiconducting materials.13–16 Currently, they are widely used as effectively alternatives or complementary tools to conventional fluorescent dyes in advanced biosensors,17 cell imaging18 and in ViVo animal tracking19 because of their great photostability, bright photoluminescence, narrow emission, and broad UV excitation. Since fluorescent detection plays an important role in both studies of complex microbial populations and the identification of bacteria, the construction of probe-conjugated QDs for single bacterium imaging is one of the most potential research areas in biological applications of QDs.20–22 Because of their broad potential (11) Wei, C.; Lin, W. Y.; Zainal, Z.; Williams, N. E.; Zhu, K.; Kruzic, A. P.; Smith, R. L.; Rajeshwar, K. EnViron. Sci. Technol. 1994, 28, 934–938. (12) Hajkova, P.; Patenka, P. S.; Horsky, J.; Horska, I.; Kolouch, A. Tissue Eng. 2007, 13, 908. (13) Bao, H. F.; Cui, X. Q.; Li, C. M.; Zang, J. F. Nanotechnology 2007, 18, 455701. (14) Bao, H. F.; Wang, E. K.; Dong, S. J. Small 2006, 2, 476–480. (15) Cao, X. D.; Li, C. M.; Bao, H. F.; Bao, Q. L.; Dong, H. Chem. Mater. 2007, 19, 3773–3779. (16) Li, R.; Li, C. M.; Bao, H. F.; Bao, Q. L.; Lee, V. S. Appl. Phys. Lett. 2007, 91, 223901. (17) Constantine, C. A.; Gattas-Asfura, K. M.; Mello, S. V.; Crespo, G.; Rastogi, V.; Cheng, T.-C.; DeFrank, J. J.; Leblanc, R. M. Langmuir 2003, 19, 9863–9867. (18) Biju, V.; Muraleedharan, D.; Nakayama, K.-i.; Shinohara, Y.; Itoh, T.; Baba, Y.; Ishikawa, M. Langmuir 2007, 23, 10254–10267. (19) Voura, E. B.; Jaiswal, J. K.; Mattoussi, H.; Simon, S. M. Nat. Med. 2004, 10, 993–998. (20) Chalmers, N. I.; Palmer, R. J.; Du-Thumm, L.; Sullivan, R.; Shi, W. Y.; Kolenbrander, P. E. Appl. EnViron. Microbiol. 2007, 73, 630–636. (21) Hirschey, M. D.; Han, Y. J.; Stucky, G. D.; Butler, A. J. Biol. Inorg. Chem. 2006, 11, 663–669. (22) Edgar, R.; McKinstry, M.; Hwang, J.; Oppenheim, A. B.; Fekete, R. A.; Giulian, G.; Merril, C.; Nagashima, K.; Adhya, S. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 4841–4845.
10.1021/la704075r CCC: $40.75 2008 American Chemical Society Published on Web 04/18/2008
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gene name sod A zwf nfo 16S
upper lower upper lower upper lower upper lower
sequence (5′ > 3′)
bases (bp)
products (bp)
AGCTGATCACCAAACTGGACC CGCTTTTTCAAATTCTGC AATGTTCTGAAGTCGTGGTCT ATGCGTCTGATTAAAGGTT ATGCCTTTATAGATGAAATGC AGATGTTCGAATTTAAACCC CGCAAGGTTAAAACTCA TACTTCTTTTGCAACCCACTC
21 18 21 19 21 20 17 21
194
applications, study of antimicrobial activity of QDs has been logically fueled up. Under irradiation, QDs can produce free radicals, of which the quality and the type are determined by their core materials.23 It is known that excess free radicals are harmful to microbes. The liberation of free heavy metal ions from QDs could also be toxic to bacteria. There are a few of reports on antimicrobial activities of QDs.24,25 Kloepfer et al. discovered that cadmium selenium (CdSe) QDs could inhibit the growth of bacteria.24 Recently, Dwarakanath et al. demonstrated that antibody-QD conjugates exhibited stronger antibacterial effects versus bare QDs.25 However, all studies are focused on the antimicrobial effects of CdSe-core QDs. Because their chemical and physical properties play vital roles in bioactivities, QDs with different core materials may have a distinct interaction with microbes. Thus, it is of great importance to investigate the antimicrobial activities of QDs that are composed of other core materials. The mechanism of the interaction between nanoparticles and bacteria needs to be better understood. Although ROS has been proposed to be a key player in mediating the antimicrobial activity of QDs,24,25 the studies so far are still limited to determine the survival rate of bacteria exposed to QDs. The effects of QDs on bacterial antioxidative response, biomolecule oxidative damage, and the biological molecular mechanism of QDs’ antibacterial properties have not been studied yet. In this study, the investigation of the antimicrobial activity and mechanism of CdTe QDs, one of the most widely used QDs, is reported. The binding capability of the QDs on the surface of E. coli is explored to verify the diffusion pathway of ROS. Further, we systematically studied the toxic effects of the QDs on the antioxidative system and biomolecules by measuring the oxidative damage of E. coli and determining the expression levels of related genes and activities of antioxidative enzymes. On the basis of these results, a distinct biomolecular mechanism of CdTe QDs’ antimicrobial activity is proposed.
Experimental Section Unless stated otherwise, all chemicals were purchased from SigmaAldrich. Bacteria Growth and CdTe QDs Treatment. High fluorescent mercaptosuccinic acid (MSA)-capped CdTe nanocrystals were synthesized according to Bao’s route.14 The synthesized CdTe QDs with orange emission and good physical and optical properties were applied in the following antimicrobial tests and toxicological assays (Supporting Information Figure S1). Before conducting antimicrobial assays, freshly synthesized QDs were washed three times in 50% ethanol by centrifugation (4000g, 5 min) to remove residual chemicals. The pigmented layer was suspended into phosphatebuffered solution (PBS) (pH 7.4) and diluted to desired concentrations. E. coli K12 (ATCC 29181) was grown with aeration at 37 °C in an (23) Ipe, B. I.; Lehnig, M.; Niemeyer, C. M. Small 2005, 1, 706–709. (24) Kloepfer, J. A.; Mielke, R. E.; Nadeau, J. L. Appl. EnViron. Microbiol. 2005, 71, 2548–2557. (25) Dwarakanatha, S.; Bruno, J. G.; Athmaram, T. N.; Bali, G.; Vattem, D.; Rao, P. Folia Microbiol. (Prague) 2007, 52, 31–34.
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LB medium, a mixture containing 10 g of peptone, 5 g of yeast extract, and 10 g of NaCl per liter (pH 7.0). When the absorbencies of bacteria cultures rose to 0.6 at 600 nm (A600), bacteria were collected by centrifugation at 4000g for 10 min. After being washed, the pellet was suspended in PBS to obtain the original absorbency again. Different concentrations of QDs were dropped into suspensions, respectively, and the mixtures were incubated at 37 °C for 2 h. Colony-Forming Capability Test. After being exposed to different concentrations of QDs at 37 °C for 2 h, E. coli suspensions were diluted to 2 × 103 cells per milliliter. 100 µL of the cell suspension was spread onto LB agar plates. The number of the colonies was counted after agar plates were incubated at 37 °C in the dark overnight. The survival percentage was used to evaluate the antimicrobial effect of QDs. Survival percentage was defined as the following formula:
Survival% )
Colony numbers of treated bacteria × 100% Colony numbers of control bacteria
Atomic Force Microscopy (AFM). A 100 µL portion of bacteria suspension was dropped onto the mica surface. After a 10 min incubation at room temperature, the substrate was rinsed with distilled water three times, followed by a N2 blow-dry. The morphologies and cell wall structures of the bacteria were captured by using AFM (SPM 3100, Veeco Instruments, Inc., U.S.A.). All AFM measurements were carried out in contact mode at ambient temperature in air. Spectrophotometry. Photoluminescence (PL) spectra were recorded by using an Aminco Bowman II luminescence spectrometer (Thermo Electron, U.S.A.) with an illumination source at 400 nm. After being treated with 200 nM QDs for 2 h, E. coli suspensions were centrifuged at 4000g for 10 min. The supernatants were directly applied in the measurement of emission spectra. PL spectra of pellets were monitored after resuspending them into 3 mL of PBS. Laser Scanning Confocal Microscopy. Fluorescent images were acquired with a Zeiss LSM 510 Meta confocal microscope. Bacteria were washed with PBS three times before conducting the test to remove free QDs. In order to exclude the interference of bacteria autofluorescence, the excitation laser and filter used in the assay were argon 488 nm and BP530-600, respectively. Semiquantitative Reverse Transcription Polymerase Chain Reaction (RT-PCR) Assay. Total RNA samples were extracted from nontreated and QDs-treated bacteria, respectively, using TRIZOL reagent (Invitrogen, U.S.A.) according to the manufacturer’s protocol. The reverse transcription was conducted by using a firststrand synthesis system (Invitrogen, U.S.A.) to produce cDNA. The primers for PCR were synthesized as shown in Table 1. Incubation at 94 °C for 5 min to prevent the formation of an RNA secondary structure was followed by performing PCR amplification for 30 cycles. Amplified products were separated by agarose gel electrophoresis (1.2%) and visualized under UV light after being stained with ethidium bromide. The LabImage software (Kapelan GmbH, Bio-Imaging Solutions, Germany) was used to determine the intensities of the amplified products appearing as bands with the predicted sizes in the images. The net intensity of the 16S rRNA gene was used to normalize the data to correct any error in the amount of input RNA. Enzyme Activity Assay. The peroxidase activity was measured in reaction mixtures containing 10 mM H2O2, 1.6 mM o-diarising and 40 mM 3-amino-1,2,4-triazole in 50 mM potassium phosphate
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Figure 1. Antimicrobial activity of CdTe QDs. After being exposed to different concentrations of QDs at 37 °C for 2 h, E. coli cells were spread onto LB medium agar plates and incubated at 37 °C for 12 h (n ) 3). *: p < 0.05, **: p < 0.01. Inset shows the formation of bacterial colonies on agar plates.
+ 0.5 mM EDTA with pH 7.0 at 25 °C. After 20 µL of protein supernatant was added into the mixture, the change of the absorbency at 436 nm was measured for 60 s. One unit of the peroxidase activity was defined as one optical change at 436 nm/min. The activity of superoxide dismutase (SOD) was assayed by the xanthine oxidase/cychrome c method.26 The reaction mixture contained 10 mM cytochrome c and 50 mM xanthine in 50 mM potassium phosphate + 0.1 mM EDTA with pH 7.8 at 25 °C. The reduction of cytochrome c by superoxide anion, which is generated from the molecular oxygen by the reduction of xanthine, was measured by UV–vis spectrometry at 550 nm. Reactions were started by adding xanthine oxidase in an amount sufficient to cause an absorbance change of 0.025/min in the cytochrome c assay at pH 7.8. One unit of the SOD activity was defined as a 50% decrease in the rate of cytochrome c reduction. Measurement of Protein Carbonyl and Lipid Peroxides. Contents of protein carbonyl were detected by the reaction with 2,4-dinitrophenylhydrazine (DNPH).27 The product, 2,4-dinitrophenylhydrazone, was spectrophotometrically quantified at 370 nm. 200 µL of supernatant was mixed with 1 mL of 10 mM DNPH in 2 M HCl. The control sample contained 1 mL of 2 M HCl instead of the DNPH solution. Samples were incubated in the dark for 1 h, vortexing every 10 min. After adding 500 µL of 10% trichloroacetic acid (TCA), mixtures were centrifuged for 15 min at 12 000g. Supernatants were discarded, and pellets were washed three times with 1 mL of ethanol-butylacetate (1:1 v/v) to remove free DNPH. Then, pellets were dissolved in 1.5 mL of 6 M guanidine-HCl and centrifuged as above. The supernatant was collected, and its absorbency was measured at 370 nm. Carbonyl content was expressed as micromoles of carbonyl per milligram of protein with a molar extinction coefficient of 22.0 mM-1 cm-1. Products of lipid peroxidation were measured by the thiobarbituric reacting substances (TBARS) assay.28 The TCA-treated cell suspension was centrifuged for 10 min at 4 000g. The supernatant was mixed with 2 mL of 20 mM butylated hydroxytoluence and saturated thiobarbituric in 0.1 M HCl. After being heated at 100 °C for 60 min, a 1.5 mL aliquot was chilled with ice followed by mixing with 1.5 (26) Attar, F.; Keyhani, E.; Keyhani, J. Appl. Biochem. Microbiol. 2006, 42, 101–106. (27) Oliver, C. N.; Starke-Reed, P. E.; Stadtman, E. R.; Liu, G. J.; Carney, J. M.; Floyd, R. A. Proc. Natl. Acad. Sci. U.S.A. 1990, 87, 5144–5147. (28) Yousef, M. I.; Awad, T. I.; Elhag, F. A.; Khaled, F. A. Toxicology 2007, 235, 194–202.
mL of butanol. The mixture was centrifuged for 10 min at 4 000g to collect the supernatant for spectrophotometrically measurement at 535 nm. Contents of TBARS were presented as picomoles per milligram of protein by applying a molar extinction coefficient of 156 mM-1cm-1. Data Analysis. Data of enzyme activities were expressed as units per milligram of protein. The protein content of each sample was measured with a BCA protein assay kit. All tests were repeated in independent experiments. Differences between different groups were tested for significance using Student’s t test (Origin 6.0).
Results and Discussion The Gram-negative bacterium E. coli is selected as a testing microbe since it is a well-studied model organism in antimicrobial experiments. The antimicrobial activity of CdTe QDs was examined by comparing the colony-forming capability of nontreated and QD-treated bacteria. As shown in Figure 1, even at a concentration as low as 80 nM, the QDs are observed to have a significant toxic effect on E. coli. When the concentration of QDs reaches 200 nM, a 92% decrease in the viability of bacteria is found. The results demonstrate that CdTe QDs can effectively kill bacteria in a concentration-dependent manner. It is quite similar to the reported antibacterial effect of CdSe-core QDs where the number of colony-forming units (CFUs) reduced when the exposure concentration of CdSe QDs increased.25 The antibacterial concentration range determined in our study is also consistent with that for CdSe QDs reported by Kloepfer et al.24 These results indicate that both CdSe QDs and CdTe QDs may share the same antimicrobial mechanism. Antimicrobial effects are usually accompanied by a change in bacterial surface morphology. To further verify the antibacterial activity, contact-mode AFM microscopy was applied to evaluate the effect of CdTe QDs on E. coli surface morphology. The rod shape, the surface structure, and flagella of individual E. coli cells can be clearly seen in Figure 2. As AFM images show, CdTe QD treatment significantly changes the morphology of the bacterium. Small dots on the native cell surface are all replaced by big clusters after CdTe QD incubation (Figure 2B,F). From section curves, it can be observed that pores on the surface of
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Figure 2. CdTe QD-induced surface morphological change of E. coli. (A) Friction image of nontreated E. coli. (B) Friction image of QD-treated E. coli. (C) Height image of nontreated E. coli. (D) Height image of QD-treated E. coli. The image size in panels A-D is 4 µm. (E) 3-D height magnified image of nontreated E. coli surface. (F) 3-D height magnified image of QD-treated E. coli surface. The image size of panels E and F is 500 nm. (G) Section curve for image E. (H) Section curve for image F.
the treated bacterium are much deeper than those on the native E. coli cell wall. Oxidative damage to the cell wall of Saccharomyces cereVisiae have been reported to cause similar membrane damage.29 The results of colony-forming assay and AFM microscopy indicate that CdTe-core QDs can also act as an antibacterial nanomaterial and could have the potential to be formulated as novel antimicrobial materials with superior optical properties. (29) Pereira, R. D.; Geibel, J. Mol. Cell. Biochem. 1999, 201, 17–24.
It is unknown how CdTe QDs react with E. coli to cause the antibacterial effects. Heavy metal ions and oxides from the CdTe core of QDs are suspected to be the possible cause. As reported, the release of heavy metal ions and oxides is always accompanied by the decomposition of QDs, leading to the blue shift of the emission peak.30,31 To prove this assumption, PL spectra of both QDs-bacterial mixture and bacteria suspension were monitored (30) Boldt, K.; Bruns, O. T.; Gaponik, N.; Eychmuller, A. J. Phys. Chem. B 2006, 110, 1959–1963. (31) Derfus, A. M.; Chan, W. C. W.; Bhatia, S. N. Nano Lett. 2004, 4, 11–18.
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Figure 3. PL spectra of QDs-bacteria mixtures and bacteria pellet suspensions. (A) PL spectra of QDs-bacteria mixtures that were measured at an interval of 20 min. (B) PL spectra of bacteria before (dash line) and after (solid line) 2 h QD incubation. The inset shows the fluorescence of QD-treated and nontreated bacterial pellets with the illumination at 330 nm.
respectively before and after the incubation. It can be seen from Figure 3A that peak intensities of QD emission gradually decline in the QDs-bacteria mixture as the incubation time increases, while the emission peak wavelength slightly shifts from 602 to 605 nm. The red shift caused by the aggregation of QDs on the surface of E.coli is observed instead of a blue shift, possibly indicating that CdTe QDs do not decompose during the incubation process to release heavy metal ions for the antimicrobial activity. QDs can transfer energy to nearby oxygen molecules to induce the formation of singlet molecular oxygen (1O2) and hydroxyl free radicals (OH•).23,32–34 Our study shows that CdTe QDs can increase the level of ROS in bacterial culture medium (Support(32) Bakalova, R.; Ohba, H.; Zhelev, Z.; Ishikawa, M.; Baba, Y. Nat. Biotechnol. 2004, 22, 1360–1361. (33) Green, M.; Howman, E. Chem. Commun. 2005, 121–123. (34) Bakalova, R.; Ohba, H.; Zhelev, Z.; Nagase, T.; Jose, R.; Ishikawa, M.; Baba, Y. Nano Lett. 2004, 4, 1567–1573.
ing Information Figure S2). ROS could be the key players in the biocidal effects of CdTe QDs. However, the lifetime of 1O2 is about 10-6 to 10-5 seconds,35 and OH• can only react with the surrounding molecules.36 CdTe QDs can self-assembled on bacterial flagella nanotubes.37 If ROS play a key role in the antimicrobial effect, CdTe QDs must be close enough to bacterial cells. In this study, PL spectrophotometry and confocal microscopy were conducted to investigate whether QDs can bind with E. coli. As shown in figure 3B, no emission peak is observed in bacteria pellet suspension before QDs treatment. After a 2 h exposure to 200 nM QDs, a significant emission peak corre(35) Kukreja, R. C.; Jesse, R. L.; Hess, M. L. Mol. Cell. Biochem. 1992, 111, 17–24. (36) Oren, D. A.; Charney, D. S.; Lavie, R.; Sinyakov, M.; Lubart, R. Biol. Psychiatry 2001, 49, 464–467. (37) Kumara, M. T.; Tripp, B. C.; Muralidharan, S. J. Phys. Chem. C 2007, 111, 5276–5280.
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Figure 4. Confocal micrographs of nontreated and QD-treated E. coli. (A) Fluorescent micrograph of nontreated E. coli. (B) Bright field micrograph of nontreated E. coli. (C) Merged micrograph of A and B. (D) Fluorescent micrograph of nontreated E. coli. (E) Bright field micrograph of nontreated E. coli. (F) Merged micrograph of D and E.
sponding to the QDs appears in the pellet suspension. The inset of figure 3B also illustrates that the pellet of QD-treated bacteria under illumination at 330 nm emits orange fluorescence, whereas the E. coli pellet without QD exposure has no any emission. It is known that the diffusion coefficient of oxygen is about 10-5 cm2 s-1, and ROS should have an identical value.38 The time for ROS diffusion into the bacterium should be around 10-7 s, considering the nanoscale thickness of the E.coli cell wall and membrane.39 Therefore, in comparison to the lifetime of ROS (∼10-5 to 10-6s), QD-induced ROS could have enough time to diffuse into the cells. Confocal microscopy further demonstrates the association of QDs and bacteria (Figure 4). Most of the QD-exposed E. coli cells present fluorescence under the irradiation, but no fluorescence can be observed in nontreated bacteria. As reported, QDs without surface conjugations cannot be transported into the bacteria owing to the lack of endocytosis mechanism.40 In this work, QDs should only be adsorbed on the surface and flagella of E. coli. The AFM images in Supporting Information Figure S3 show the binding of QDs on the cell wall and flagella of E.coli cells. Results of both PL and confocal micrographs indicate that the MSA-capped CdTe QDs can bind on the surface of bacteria, providing an immediate contact for the easy diffusion of ROS into the bacterium to play a vital role in the antibacterial activity of QDs. In biological aerobic organisms, an antioxidative system can protect the cells from the oxidative damage caused by ROS. (38) Gubbins, K. E.; Walker, R. D. J. Electrochem. Soc. 1965, 112, 469–471. (39) Anuntalabhochai, S.; Chandej, R.; Phanchaisri, B.; Yu, L. D.; Vilaithong, T.; Brown, I. G. Appl. Phys. Lett. 2001, 78, 2393–2395. (40) Sweeney, R. Y.; Mao, C. B.; Gao, X. X.; Burt, J. L.; Belcher, A. M.; Georgiou, G.; Iverson, B. L. Chem. Biol. 2004, 11, 1553–1559.
Thus, although the adsorption of QDs on surfaces of E. coli could possibly make ROS enter into a bacterial cell, their antibacterial effect still depends on the antioxidative response of bacteria. The response of the E. coli antioxidative system to CdTe QDs should be also considered in the mechanism of the QDs antimicrobial activity. In this work, with a semiquantitative RT-PCR assay, a number of ROS-related genes were chosen as target genes to explore whether QDs could affect the cellular antioxidative system by regulating gene expressions. nfo, sod A, and zwf genes encode endonuclease IV, manganese-superoxide dismutase (Mn-SOD) and glucose-6-phosphate dehydrogenase (G6PDH), respectively. 16 S rRNA, a noncoding RNA gene, was used as the internal control in this experiment. Figure 5 shows that expressions of three target genes are significantly down-regulated in 200 nM QD-treated samples. The expression level of the nfo gene is inhibited up to 50% by the QDs treatment. The down-regulation of the nfo gene can inhibit the activity of endonuclease IV, and further affect the DNA repair system. About 10% downregulations of sod A and zwf are observed. Since Mn-SOD and G6PDH are vital enzymes that can eliminate ROS in E. coli cells under the stress environment, down-regulations of sod A and zwf genes could significantly reduce activities of antioxidative enzymes to suppress the cell antioxidation capability. To further confirm whether gene expression down-regulation could affect the antioxidative system, activities of antioxidative enzymes in QD-treated E. coli were investigated. Two of the most crucial enzymes in the ROS elimination system, SOD and peroxidase, were monitored. As illustrated in Figure 6, QDs could cause the decline of activities of both peroxidase and SOD in a dose-dependent manner. Even if the concentration is only
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Figure 5. QD-induced down-regulations of ROS-related genes in E. coli. (n ) 3). *: p < 0.05, **: p < 0.01.
Figure 7. QD-induced oxidative damage to proteins and lipids in E. coli. (A) Effects of QDs on the level of TBARS. (B) Effects of QDs on the level of protein carbonyl. (n ) 3). *: p < 0.05, **: p < 0.01. Figure 6. Effects of the QDs on activities of antioxidative enzymes in E. coli: (A) peroxidase; (B) SOD. (n ) 3). *: p < 0.05, **: p < 0.01.
40 nM, QDs could still inhibit the activity of peroxidase significantly. For the SOD activity, it was negatively affected by an exposure to 120 or higher nM QDs (p < 0.05, compared to control samples). It can be found in the figure that 200 nM QDs could lead to decreases of activities for both peroxidase and SOD. These results are greatly in agreement with the findings in the RT-PCR experiment. Thus, treatments with QDs can not only down-regulate expressions of ROS-related genes, but also reduce activities of antioxidative enzymes in E. coli. Apparently, both effects of QDs on E. coli revealed from RT-PCR and enzyme activity assays could result in the accumulation of ROS in bacteria. Once ROS begins to accumulate, the oxidative stress in cells can first damage biomolecules such as protein and lipid. Thus, oxidative damage to biomolecules is regarded as indicators of
the formation of oxidative stress in cells. It is essential to study whether ROS accumulation in bacteria causes the oxidative stress and damage of biomolecules for fully understanding the antibacterial mechanism. In this work, TBARS and protein carbonyl contents, biomarkers of lipoperoxidation, and protein oxidative modifications were used to quantitatively determine levels of oxidative damage to lipids and proteins in E. coli, respectively. Results of the lipoperoxidation damage demonstrate a dose–effect relationship between TBARS and QD concentrations (Figure 7A). At concentrations higher than 120 nM, the level of TBARS in cells increases significantly. Protein carbonyl contents in E. coli are also related to QD exposure levels. It seems that proteins are more sensitive to oxidative stress induced by QDs than are lipids. 40 nM QDs were enough for the incurrence of significant oxidative damage of the proteins (Figure 7B). This may be one of the causes of lower enzyme activities in the antioxidative system after exposure to QDs. The result here proves
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Figure 8. Schematic diagram of the mechanism of CdTe QD antimicrobial activity.
the formation of oxidative stress and oxidative damage to biomolecules in E. coli. Lovric´ et al. recently reported that N-acetylcysteine, an antioxidant, could effectively prevent the deleterious effects of QDs-induced ROS in cultured cells,41,42 also suggesting that QD-induced ROS plays an essential role in the cytotoxicity of QDs. This is greatly consistent with ROSrelated bactericidal effects observed in this work. On the basis of these discoveries, we propose the mechanism of the antimicrobial activity of CdTe QDs that involves both the surface binding and a ROS-dependent pathway, as shown in Figure 8, in which QDs bound on the bacteria surface make the reaction of ROS with cells easily. CdTe QDs can also downregulate expressions of antioxidative genes, resulting in lowlevel translations of antioxidative enzymes to suppress the capability of removing ROS. Obviously, both effects of CdTe QDs can break the oxidant/antioxidant balance and then cause accumulation of ROS in bacteria. The superfluous ROS in bacteria oxidizes proteins and lipids, and triggers the necrosis cellular signals transduction, leading to the biocidal effects. Although TiO2, SiO2, ZnO, and QD nanoparticles possess different energy gaps, they are all reported to produce ROS.7,23 They may have an antibacterial mechanism similar to that of CdTe QDs. The proposed mechanism might provide scientific insight to other nanoparticle-induced biocidal activity. However, the effect of different energy gaps possessed by different nanoparticles and QDs with different sizes on the amount of ROS produced and antibacterial mechanism is not clear, and is currently being investigated in the author’s laboratory. Many of the reports on the antibacterial effects and cytotoxicity of nanomaterials focus on the toxins generated by nanomaterials. However, the mechanism of CdTe QD antimicrobial activity proposed in this study suggests that, besides nanoparticle-induced ROS, the response of the antioxidative system should also be considered as a key (41) Lovric´, J.; Bazzi, H. S.; Cuie, Y.; Fortin, G. R. A.; Winnik, F. M.; Maysinger, D. J. Mol. Med. 2005, 83, 377–385. (42) Lovric´, J.; Cho, S. J.; Winnik, F. M.; Maysinger, D. Chem. Biol. 2005, 12, 1227–1234.
event in the antimicrobial activity and cytotoxicity of nanoparticles.
Conclusions In conclusion, CdTe QDs are demonstrated to be antimicrobial nanoparticles for the first time in this report. Those QDs can bind on the bacterial surface and negatively affect the function of cellular antioxidative systems, including down-regulations of antioxidative genes and decreases of antioxidative enzyme activities. In addition, the bacterial survivability and the level of oxidative damage to the biomolecules are also dependent on the exposure concentration of QDs. On the basis of these results, the mechanism of the antimicrobial activity of MSA-capped CdTe QDs, which involves the formation of the QDs-bacteria complex and a QD-related ROS-mediated pathway, is proposed. Although it has been reported that QDs could induce cytotoxicity to animal cells, as summarized by Hardman,43 the toxic concentration level is in the range of several micromoles per liter, which is much higher than the antimicrobial concentration observed in this report. Therefore, QDs have the potential to be formulated as novel antimicrobial materials with excellent optical properties. However, the use of QDs as antimicrobial materials should be cautious and limited until their long-term toxic effects are completely clear and biocompatibility is significantly improved. Acknowledgment. This work was financially supported by the Center for Advanced Bionanosystems, Nanyang Technological University, Singapore, under the Research Programs for Electronic Biochip. Supporting Information Available: Details about characterization of MSA-capped CdTe QDs, cytochrome c reduction assay, and AFM micrographs of QD-bacterium association. This material is available free of charge via the Internet at http://pubs.acs.org. LA704075R (43) Hardman, R. EnViron. Health Perspect. 2006, 114, 165–172.
