Article pubs.acs.org/IECR
Direct Detection System for Escherichia coli Using Au−Ag Alloy Microchips Wan-Joong Kim,† Sanghee Kim,*,‡ Ae Rhan Kim,§ and Dong Jin Yoo*,∥ †
Biosensor Research Team, Electronics and Telecommunications Research Institute, Daejeon 305-700, Republic of Korea Department of Mechanical Systems Engineering, Hansung University, Seoul 136-792, Republic of Korea § Division of Chemical Engineering and ∥Department of Hydrogen and Fuel Cells Engineering, Specialized Graduate School, Chonbuk National University, Jeollabuk-do 561-756, Republic of Korea ‡
ABSTRACT: Au−Ag alloy nanoparticles (NPs) were prepared by the reduction of metal ion mixtures in aqueous sodium citrate solution using sodium borohydride (NaBH4). The resulting Au−Ag alloy NPs were analyzed by various techniques. Alloyattached chips for the detection of microorganisms were fabricated simply by the attachment of Au−Ag alloy nanoparticles onto glass slides after silanization through self-assembled monolayers (SAMs) for the formation of activated amine (−NH2) as a terminal function group. The alloy-attached chips were investigated for their ability to bind the target Escherichia coli (E. coli) in water. E. coli was detected in water as a function of time and concentration by UV−vis spectroscopic measurements based on the interaction between the alloy-attached chip and E. coli. Field-emission scanning electron microscopy (FE-SEM) was used to directly observe the E. coli captured on the alloy chips. These studies demonstrated that E. coli in drinking water can be directly detected with Au−Ag alloy microchips without requiring any interaction between an antibody and an antigen.
1. INTRODUCTION E. coli is found in large numbers in the intestinal tracts of humans and other warm-blooded animals, and most strains spread widely in any natural environment.1 E. coli can be deadly, especially for children or the elderly, and it is the major cause of infectious outbreaks with serious consequences, often caused by ingesting contaminated food or water.2−5 Therefore, the presence of E. coli in foodstuffs and drinking water is a chronic worldwide problem. Recently, several rapid assays for detecting E. coli based on different measuring methods, such as DNA-based assays,6−8 fluorescence,9−11 metal nanoparticles,12−15 electrochemical sandwich immunoassays,16−18 and optical assays,19,20 have been developed. DNA-based assays, which are the most specific and sensitive methods available, are routinely used as a confirmatory assay. Simpson and Lim demonstrated a method that reduced the time required for detection from 10 h to about 2 h by direct polymerase chain reaction (PCR) of bacteria from fiber-optic waveguides.21 Stender et al. published a new fluorescence in situ hybridization (FISH) method using peptide nucleic acid (PNA) probes and an array scanner for the rapid detection, identification, and enumeration of E. coli.22 Cheng et al. demonstrated a rapid, specific, and sensitive method for assaying E. coli using biofunctional magnetic nanoparticles (BMNPs) in combination with adenosine triphosphate (ATP) bioluminescence.12 Using a microfluidic device, Han et al. demonstrated that E. coli K-12 can be detected in a Y-channel polydimethylsiloxane (PDMS) microfluidic device through optical fiber monitoring of latex immunoagglutination.23 These reports describe E. coli assay methods that are rapid and have improved sensitivity and specificity; however, all are too complicated, laborious, and expensive to be implemented in portable biosensors. These previously reported sensing methods also usually require the detection of antibodies for © XXXX American Chemical Society
immunoassays. Therefore, better, more sensitive, and simpler methods are still needed to detect E. coli. Recently, silver nanoparticles (Ag NPs) have attracted a great deal of attention because of their unique optical and electronic properties and potential applications in biological, optoelectronic, and photonic technologies.24−28 Ag NPs have a strong affinity to E. coli, which bears numerous amine (−NH2) and thiol (−SH) groups.29 In addition, Ag NPs have strong biocidal effects on various species of bacteria, including E. coli, and are highly toxic to microorganisms.30,31 Despite various applications, Ag NPs have not yet been used as mediation materials in sensors to directly detect specific biomolecules such as microorganisms, proteins (antigens), and others because of the instability of Ag NPs in both air and water. Therefore, it would be a great benefit to have a more stable material with Ag properties for practical applications of Ag NPs, such as in sensor devices.32,33 It is known that Au−Ag alloy NPs, bimetallic nanoparticles with greater stability, can be easily prepared from HAuCl4·3H2O and AgNO3 reagents through the reduction of sodium citrate because Au and Ag have the same face-centered cubic (FCC) crystal structures and the lattice constants of Au (0.4078 nm) and Ag (0.4086 nm) are very similar.32 In this study, we synthesized stable Au−Ag alloy NPs and deposited these Au−Ag alloy NPs onto glass devices using the interactions between the metal ions and the reactivated amine group on the glass formed by self-assembled monolayers (SAMs) for sensor chips. The alloy-attached devices were investigated for their ability to bind the target E. coli in water. Received: August 25, 2012 Revised: April 26, 2013 Accepted: May 8, 2013
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Figure 1. Characterization of prepared Au−Ag alloy NPs: (a) UV−visible absorption spectrum (Au/Ag = 1:9) and (inset) fraction of each particle size, (b) TEM image of Au−Ag alloy nanoparticles, (c) EDXS analysis showing each component alloy NP as a 1:9 ratio, (d) wide-angle XRD pattern analysis showing that the peak position corresponds to the lattice structure of Au and Ag in alloy NPs. XPS spectra of sodium-citrate stabilized Au-Ag alloy nanoparticles (e) Au 4f and (f) Ag 3d.
We detected E. coli in water as a function of time and concentration by UV−visible absorption spectroscopy based on the interaction between the Au−Ag alloy-attached chip and E. coli without detection antibody. This simple method using Au− Ag alloy NPs might be very useful for the development of lowcost portable biosensors for the detection of microorganisms in water supplies or water purifiers.
Deionized water was distilled in a Milli-Q water purification system. E. coli-attached chips were analyzed by UV−visible absorption spectroscopy (U-3501, Hitachi, Tokyo, Japan). Oxygen plasma analysis was performed using an SPI PlasmaPrep II Plasma etcher. Field-emission scanning electron microscopy (FE-SEM) images were collected on an FEI Sirion-400 field-emission scanning electron microscope. Transmission electron microscopy (TEM) images were obtained using a JEM-ARM200F JEOL instrument equipped with an energy-dispersive X-ray spectroscopy (EDXS) analyzer at an accelerating voltage of 200 kV. X-ray photoelectron spectroscopy (XPS) analysis was conducted using an ESCA LAB 210D (VG Scientific Ltd., East Grinstead, U.K.) analyzer with a monochromatic Al Kα X-ray source at a power of 12.5 kV at 250 W and scanning range from 0 to 1400 eV. High-resolution spectra were obtained with a 50 eV pass energy, 0.05 eV energy
2. EXPERIMENTAL SECTION 2.1. Materials and Measurement. 3-Aminopropyltriethoxylsilane (APTES), gold(III) chloride trihydrate (HAuCl4·3H2O), silver nitrate (AgNO3), sodium borohydride (NaBH4), and sodium citrate tribasic dehydrate (Na3C6H5O7) were purchased from Sigma-Aldrich (St. Louis, MO). E. coli (KCCM41036) was purchased from the Korean Culture Center of Microorganisms (KCCM, Seoul, Korea). B
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Target monolayers of APTES were formed on the surface of the hydroxylated glass substrates. The amino-silanized surface was rinsed with ethanol and baked at 120 °C for 15 min. Reactive amine groups were then introduced onto the surface. For deposition with alloy nanoparticles onto the glass, the reactivated amine groups on the glass were incubated with a solution of Au−Ag alloy NPs (0.4 nM) at 22 °C overnight. After incubation, the deposited alloy glass was rinsed three times with DI water. 2.4. Bacterial Preparation. E. coli (KCCM41036) was resuspended in Luria−Bertani (LB) broth (1 g of tryptone, 0.5 g of yeast extract, and 0.5 g of NaCl in 1 L of DI water, adjusted to pH 7.0). E. coli was grown at 30 °C overnight with aeration in LB broth supplemented with 0.5 wt % glucose. The culture was then used to inoculate fresh LB broth containing 0.5 wt % glucose at 1:100 dilutions and grown at 30 °C with aeration. The optical density at 600-nm wavelength (OD600) was measured using a spectrophotometer (Thermo Spectronic, Cambridge, U.K.). The number of cells was determined using a counting chamber. The correlation between optical density and number of cells was determined and used to generate the growth curve of E. coli under batch conditions.
step size, and 200 ms dwell time. A nonlinear, Shirley-type baseline and an iterative fitting program (Thermo Advantage 3.31) were used to deconvolve the XPS peaks. The peak-fitting process was repeated until an acceptable fit was obtained. In addition, X-ray diffraction (XRD) patterns were collected on an RU-200BHD instrument (Rigaku, Tokyo, Japan) operating at a voltage of 40 kV and a current of 40 mA in the range of 2θ values between 10° and 90° in steps of 0.02° at a speed of 3°/ min. 2.2. Preparation of Au−Ag Alloy NPs. Au−Ag alloy NPs (Figure 1) with sizes of 13 nm were prepared by the known procedures34 as follows: Briefly, while a mixture of 0.8 mg of HAuCl4·3H2O, 3.1 mg of AgNO3, and 50 mg of sodium citrate dissolved in 95 mL of deionized (DI) water was being stirred at ∼4 °C, 0.25 mL of ice-cold 500 mM NaBH4 was added at 1min intervals. The colloidal solution was stirred for an additional 5 min at ∼4 °C. After being stirred for another 30 min at room temperature, the mixture was filtered through a 0.45-μm polymer membrane filter. 2.3. Preparation of the Alloy-Attached Chip. The glass substrate was cleaned by sonication in methanol for 15 min, rinsed with water, and blown dry with nitrogen before being exposed to oxygen plasma radiation. As shown in the schematic diagram of Figure 2a, the hydroxylation was also carried out within ∼1−5 min after oxygen plasma treatment to form hydroxyl groups (OH) on the glass substrates, which were exposed to laboratory atmospheric conditions of 22 °C temperature and 32% humidity. The hydroxylated surface was silanized in ethanol (0.5% DI water) containing 1 wt % APTES.
3. RESULTS AND DISCUSSION 3.1. Preparation of Au−Ag Alloy NPs. Colloidal dispersions of bimetallic Au−Ag alloy NPs of different molar
Figure 3. Schematic representation of the mechanism of E. coli capture by Au−Ag alloy NPs through SAMs (self-assembled monolayers) of thiol groups on the cysteine domain of E. coli on the surface of the alloy.
ratios (Au/Ag = 1:9) were synthesized with a slight modification of the conventional citrate reduction method, using reduction by NaBH4 without refluxing. As shown in Figure 1, the prepared Au−Ag alloy NPs were verified using UV−visible absorption spectroscopy, TEM, EDXS, XPS, and XRD. The chemically synthesized Au and Ag NPs showed surface plasmon absorption bands with maxima at around 400 and 520 nm, but the absorption bands of Au−Ag alloy NPs (Figure 1a) appeared at around 422 nm when the molar ratio of HAuCl4·3H2O to AgNO3 was 1:9. The Au−Ag alloy NPs absorption showed a sharper peak than Au NPs that was slightly red-shifted compared with that of Ag NPs. Figure 1b shows a TEM image of Au−Ag alloy NPs used to examine the
Figure 2. Schematic representation of the surface modification of a glass chip and the capture of E. coli by Au−Ag alloy NPs: (a) preparation of surface functionalization by O2 plasma and APTES treatment for further Au−Ag alloy NP-attached glass chips, (b) FESEM image of Au−Ag alloy NP-attached glass chip, and (c) capture sequence of target E. coli by Au−Ag alloy NPs deposited on a glass chip and detection of captured E. coli using UV−visible absorption spectra. C
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Figure 4. Characterization of E. coli by UV−visible spectrophotometry: (a) wavelength shift (Δλ) measured by UV−visible spectroscopy at different concentrations of E. coli for 48 h and (b) UV−visible absorption spectra and (c) degree of wavelength shift (Δλ) of 4 × 103 cfu/mL E. coli measured at different incubation times.
size distribution, and the inset in Figure 1a shows a size of 13 ± 2 nm. For the ingredient ratio of gold to silver, the alloy NPs were analyzed using EDXS (Figure 1c). The prepared alloy NPs were identified as having a 1:9 ratio of Au to Ag. 3.2. Characterizations of Au−Ag Alloy NPs. The prepared Au−Ag NPs were characterized by XRD to verify the presence of Au−Ag alloy in the mixture of monometallic particles. The XRD pattern was collected and characterized by four peaks corresponding to the (111), (200), (220), and (311) lattice planes, as shown in Figure 1d. XRD peaks corresponding to the four lattice planes were positioned at 37.86°, 44.31°, 64.56°, and 77.34°. The 2θ values of the peaks corresponding to the (111) and (200) facets were more prominent than those of the other planes, as was previously observed for Au−Ag alloy NPs.35−38 It has been reported that higher molar ratios of Ag induce broader peaks for the (220) and (311) lattice planes.39 Two additional broad peaks observed around 2θ = 64.56° and 77.34° indicate that both metallic Au and Ag were contained in the prepared NPs in composite form. Because the 2θ values corresponding to the (220) and (311) structures of metallic Au and Ag are known to be 64.48°, 77.71°, 64.55°, and 77.48°,36 the values for the diffraction peaks of the (220) and (311) lattice planes were found to be closer to those of metallic Ag. This would result from the higher content of metallic Ag than Au (Au/Ag = 1:9) used for the preparation of the Au−Ag alloy NPs. The prepared Au−Ag alloy NPs were further analyzed by XPS to obtain additional information in terms of composite structure; shifts were obtained in the spectra, as shown in Figure 1e,f. Both the Au and Ag spectra showed the characteristic two spin−orbit couplings of Au (4f5/2 and
4f 7/2) and Ag (3d3/2 and 3d5/2), with shifts of the peak positions compared with those of the pure forms of metallic gold and silver.40,41 In the Au 4f spectrum, the binding energies of Au 4f5/2 and Au 4f 7/2 were determined to be 87.1 and 83.5 eV, respectively (Figure 1e). These observed shifts in binding energy are indicative of an Au−Ag alloy structure that contains both Au and Ag atoms, as previously reported.35,42,43 The peak binding energies moved toward lower values than for bulk metallic gold determined at 84.0 and 87.7 eV.43 The XPS results suggest that metallic Au and Ag are present in nonzero valence states because the Au 4f and Ag 3d peaks appear at 84.0 and 368.1 eV when gold and silver are in their zerovalent states (Au0 and Ag0).44 In addition, upon oxidation of Au in the alloy composite structure, the binding energies are known to increase and were measured at 85.5 eV (Au 4f5/2) and 86.3 eV (Au 4f 7/2).45 Our XPS results provide evidence that there was a change in the surface orientation of metallic Au because the Au 4f binding energies exhibited a negative shift. On the other hand, the Ag 3d3/2 and Ag 3d5/2 binding energies were determined at 373.8 and 367.8 eV, respectively (Figure 1f). As for the peak positions of Au 4f, the binding energies of Ag 3d also shifted toward lower values than for bulk metallic silver.36 The oxidation of silver in the Au−Ag alloy composite structure due to electron transfer is known to shift the binding energy to lower values.43,46 It is expected that the metallic silver was present as Ag+ and was closely packed by sodium citrate ions of surfactant around the alloy composite. 3.3. Glass Chip Fabrication. Figure 2a presents a schematic of the process used for generating biochips prepared from alloy NPs on a glass surface. For the deposition of alloy NPs, the glass chip was treated with oxygen plasma radiation to D
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Figure 5. UV−visible spectrophotometric measurements displaying the capability of Au−Ag alloy NPs to target E. coli: (a) almost no signal in the absence of Au−Ag alloy NPs, (b) characteristic peak determined in the presence of Au−Ag alloy NPs, (c) Au−Ag alloy NPs in uncontaminated DI water displaying the plasmon peak of metallic NPs, and (d) shift of peak position of water contaminated with E. coli.
interaction. The most important reason to select alloy NPs for the detection of E. coli is that the alloy material has the benefits of both gold and silver materials. Generally, alloy NPs are more chemically stable than other materials when they are dispersed in aqueous solutions, similarly to gold NPs. In addition, alloy NPs show a narrow intense plasmon absorption band at around 420 nm, similarly to silver NPs. One possible mechanism for the direct detection of E. coli is schematically shown in Figure 3. This direct detection mechanism, which does not require capture antibody, can be explained as follows: (i) Partially positive surfaces of Au−Ag alloy NPs are formed by oxidation. (ii) The unshared electron pair of the nucleophile, which can be a thiol or amine functional group of a cysteine residue contained in E. coli, attacks the Au−Ag alloy NPs to form a new bond, and subsequently, a hydrogen ion (H+) departs, leaving a pair of electrons. 3.5. Detection Results of E. coli. The concentrations of E. coli tested ranged from 4 × 107 to 4 × 102 colony forming units (cfu)/mL. As shown in Figure 4a, the wavelength shifted by 4, 10, 22, 23, 24, and 25 nm for concentrations of 4 × 107, 4 × 106, 4 × 105, 4 × 104, 4 × 103, and 4 × 102 cfu/mL, respectively, providing a linear dose−response curve for the detection of E. coli in water. In other words, the detection limit of Au−Ag alloy NPs for E. coli was found to be 4 × 102 cfu/mL. This value is in good agreement with that of a microarray detection system, which showed a detection limit of 5 × 103 cfu/mL.47 After alloy chips had been soaked in various concentrations of E. coli for 48 h, they were analyzed by UV−visible absorption spectroscopy, as shown in Figure 4b.
form hydroxyl groups on the surface. APTES molecules were introduced onto the surface through SAMs for the deposition of alloy NPs because alloy NPs containing gold and silver adhere well to amine groups (or thiol groups) that have lone pairs of electrons. The reaction conditions for the successful deposition of Au−Ag alloy NPs on a glass surface were found to include a high concentration of Au−Ag alloy NPs (∼8 nM) and a reaction time of greater than 12 h at room temperature. Figure 2b is an FE-SEM image of an Au−Ag alloy-deposited glass chip shown at a concentration of 728 Au−Ag alloy NPs per square micrometer. After the Au−Ag alloy glass chips had been prepared, they were soaked in of E. coli solutions of various concentrations in water. Then, the target E. coli in water was captured by the alloy-deposited glass chips and detected by UV−visible absorption spectroscopy, as shown in Figure 2c. 3.4. Detection Strategy for E. coli with Au−Ag Alloy. Our results indicate a method for the direct detection of E. coli using Au−Ag alloy chips that are simple to prepare through the attachment of Au−Ag alloy NPs to a glass surface. We employed Au−Ag alloy NPs as a sensing material for the target microorganisms. Detection of the microorganisms without requiring capture antibody was achieved by utilizing the chemical interactions between the Au−Ag alloy NPs and the thiol (and amine) functional groups of cysteine residues contained on E. coli. Because gold and silver surfaces easily absorb functional groups such as thiol (−SH), amine (−NH2), and cyanide (−CN) through their lone pairs of electrons, Au− Ag alloy NPs can directly detect the microorganisms in a water supply or water purifier without any antibody−antigen E
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seen in the FE-SEM images of different sizes in Figure 6c−f. These results indicate that the wavelength shifted as the concentration of E. coli or the chip incubation time increased.
4. CONCLUSIONS A sensor using Au−Ag alloy NPs coated on a glass chip can directly detect the concentration of E. coli in DI water by UV− visible absorption spectroscopy, without requiring any capture antibody. To prepare these sensors, we synthesized bimetallic Au−Ag alloy nanoparticles from gold(III) chloride trihydrate, silver nitrate, and sodium citrate tribasic dehydrate by a reduction method employing sodium borohydride. The fabricated alloy NPs were characterized by UV spectroscopy, TEM, EDXS, XPS, and XRD. An Au−Ag alloy-deposited glass chip used for the simple detection of E. coli in water was fabricated by the attachment of the Au−Ag alloy NPs to glass slides through activated amine terminal functional groups using silanization through SAMs. Our UV−visible absorption spectroscopy analysis showed a linear dose−response curve for the wavelength shift when E. coli was in the concentration range from 4 × 107 to 4 × 102 cfu/mL. The peak wavelength of Au− Ag-alloy-NP-coated glass chips that had captured E. coli was found to be proportional to the incubation time at a fixed E. coli concentration. In addition, the E. coli captured by the alloy-NPcoated glass chip were examined by FE-SEM. The results demonstrate the ability of the chips to distinguish between safe drinking water and polluted water using only Au−Ag alloy NPs without capture antibody.
Figure 6. FE-SEM images of E. coli captured by Au−Ag alloy NPs on a glass chip at different concentrations: (a) 4 × 102, (b) 4 × 103, (c,e) 4 × 105, and (d,f) 4 × 107 cfu/mL. Arrows indicate captured E. coli by Au−Ag alloy NPs having a typical size ∼ 1 micron.
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AUTHOR INFORMATION
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The results show the ability of the alloy chips to distinguish between safe and polluted drinking water. The wavelength was checked by UV−visible absorption spectroscopy at incubation times of 1, 2, 4, 10, 12, and 48 h for an E. coli concentration of 4 × 103 cfu/mL, as shown in Figure 4c. The results show that the wavelength increased in proportion to the incubation time at a fixed concentration of E. coli. The ability of the Au−Ag alloy NPs to detect E. coli in water was demonstrated as shown in Figure 5. A glass chip whose surface had been modified with APTES showed no sensing capability to E. coli in the absence of Au−Ag alloy NPs, as no distinctive signals were observed when E. coli was introduced onto an APTES-treated glass chip (Figure 5a). On the other hand, the characteristic peak of the Au−Ag alloy NPs was detected around 424 nm when the alloy NPs were introduced onto the glass chip surface, as shown in Figure 5b. After 4 × 103 cfu/mL E. coli had been incubated for 15 h with Au−Ag alloy NPs on a glass chip surface, a peak shift was clearly observed (Figure 5b). The ability of the Au−Ag alloy NPs to differentiate between E. coli-contaminated water and clean water is also demonstrated in Figure 5. There was no peak shift when the clean water was allowed to incubate with the Au−Ag alloy NPs (Figure 5c). The distinctive peak shift (Figure 5d) reflects the presence of E. coli captured by Au−Ag alloy NPs on the glass chip surface after 4 × 103 cfu/mL E. coli had been incubated for 15 h. The E. coli captured by the alloy NPs on the glass chip was examined by FE-SEM, as shown in Figure 6. In low E. coli concentrations of 4 × 102 and 4 × 103 cfu/mL, the E. coli could not be easily captured, as shown in the FE-SEM images in Figure 6a,b. However, the E. coli captured at high E. coli concentrations of 4 × 105 and 4 × 107 cfu/mL can be clearly
*E-mail:
[email protected] (S.K.),
[email protected] (D.J.Y.). Tel.: +82-63-270-3608 (D.J.Y.). Fax: +82-63-2703909 (D.J.Y.). Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS This work was supported by the KOCI (11ZC1110, Basic Research for the Ubiquitous Life Care Module Development) and by the Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Education, Science and Technology (20110010538). Also, this research was supported by Hansung University for S.K.
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REFERENCES
(1) Villani, F.; Russo, F.; Blaiotta, G.; Moschetti, G.; Ercolini, D. Presence and characterisation of verotoxin producing E. coli in fresh Italian pork sausages, and preparation and use of an antibiotic-resistant strain for challenge studies. Meat Sci. 2005, 70, 181. (2) Farnleitner, A. H.; Hocke, L.; Beiwl, C.; Kavka, G. G.; Zechmeister, T.; Kirschner, A. K. T.; Mach, R. L. Rapid enzymatic detection of Escherichia coli contamination in polluted river water. Lett. Appl. Microbiol. 2001, 33, 246. (3) Phillips, C. A. The epidemiology, detection and control of Escherichia coli O 157. J. Sci. Food Agric. 1999, 79, 1367. (4) Varma, J. K.; Greene, K. D.; Reller, M. E.; DeLong, S. M.; Trottier, J.; Nowicki, S. F.; DiOrio, M.; Koch, E. M.; Bannerman, T. L.; York, S. T. Escherichia coli O157 infection following exposure to a contaminated building. J. Am. Med. Assoc. 2003, 290, 2709. (5) Mohle-Boetani, J. C.; Farrar, J. A.; Werner, S. B.; Minassian, D.; Bryant, R.; Abbott, S.; Slutsker, L.; Vugia, D. J. Escherichia coli O157
F
dx.doi.org/10.1021/ie3022797 | Ind. Eng. Chem. Res. XXXX, XXX, XXX−XXX
Industrial & Engineering Chemistry Research
Article
and Salmonella infections associated with sprouts in California, 1996− 1998. Ann. Intern. Med. 2001, 135, 239. (6) Mao, X.; Yang, L.; Su, X.-L.; Li, Y. Nanoparticles amplification based quartz crystal microbalance DNA sensor for detection of E. coli O157:H7. Biosens. Bioelectron. 2006, 21, 1185. (7) Fratamico, P. M.; DebRoy, C.; Strobaugh, T. P., Jr.; Chen, C.-Y. DNA sequence of the Escherichia coli O103 O antigen gene cluster and detection of enterohemorrhagic E. coli O103 by PCR amplification of the wzx and wzy genes. Can. J. Microbiol. 2005, 51, 515. (8) Shriver-Lake, L. C.; Turner, S.; Taitt, C. R. Rapid detection of Escherichia coli O157:H7 spiked into food matrices. Anal. Chim. Acta 2007, 584, 66. (9) Arcidiacono, S.; Pivarnik, P.; Mello, C. M.; Senecal, A. Cy5 labeled antimicrobial peptides for enhanced detection of Escherichia coli O157:H7. Biosens. Bioelectron. 2008, 23, 1721. (10) Stender, H.; Oliveira, K.; Rigby, S.; Bargoot, F.; Coull, J. Rapid detection, identification, and enumeration of Escherichia coli by fluorescence in situ hybridization using an array scanner. J. Microbiol. Methods 2001, 45, 31. (11) Tanji, Y.; Furukawa, C.; Na, S.-H.; Hijikata, T.; Miyanaga, K.; Unno, H. Escherichia coli detection by GFP-labeled lysozymeinactivated T4 bacteriophage. J. Biotechnol. 2004, 114, 11. (12) Cheng, Y.; Liu, Y.; Huang, J.; Li, K.; Zhang, W.; Xian, Y.; Jin, L. Combining biofunctional magnetic nanoparticles and ATP bioluminescence for rapid detection of Escherichia coli. Talanta 2009, 77, 1332. (13) Zhao, X.; Hilliard, L. R.; Mechery, S. J.; Wang, Y.; Bagwe, R. P.; Jin, S.; Tan, W. A rapid bioassay for single bacterial cell quantitation using bioconjugated nanoparticles. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 15027. (14) Kalele, S. A.; Kundu, A. A.; Gosavi, S. W.; Deobagkar, D. N.; Deobagkar, D. D.; Kulkarni, S. K. Rapid detection of Escherichia coli by using antibody-conjugated silver nanoshells. Small 2006, 2, 335. (15) Singh, A. K.; Senapati, D.; Wang, S.; Griffin, J.; Neely, A.; Candice, P.; Naylor, K. M.; Varisli, B.; Kalluri, J. R.; Ray, P. C. Gold nanorod based selective Identification of Escherichia coli bacteria using two-photon Rayleigh scattering spectroscopy. ACS Nano 2009, 3, 1906. (16) Muhammad-Tahir, Z.; Alocilja, E. C. Fabrication of a disposable biosensor for Escherichia coli O157:H7 detection. IEEE Sens. J. 2003, 3, 345. (17) Tokarskyya, O.; Marshall, D. L. Immunosensors for rapid detection of Escherichia coli O157:H7Perspectives for use in the meat processing industry. Food Microbiol. 2008, 25, 1. (18) Heo, J.; Hua, S. Z. An overview of recent strategies in pathogen sensing. Sensors 2009, 9, 4483. (19) Van Poucke, S. O.; Nelis, H. J. Development of a sensitive chemiluminometric assay for the detection of beta-galactosidase in permeabilized coliform bacteria and comparison with fluorometry and colorimetry. Appl. Environ. Microbiol. 1995, 61, 4505. (20) Geissler, K.; Manafi, M.; Amoros, I.; Alonso, J. L. Quantitative determination of total coliforms and Escherichia coli in marine waters with chromogenic and fluorogenic media. J. Appl. Microbiol. 2000, 88, 280. (21) Simpson, J. M.; Lim, D. V. Rapid PCR confirmation of E. coli O157:H7 after evanescent wave fiber optic biosensor detection. Biosens. Bioelectron. 2005, 21, 881. (22) Stender, H.; Oliveira, K.; Rigby, S.; Bargoot, F.; Coull, J. Rapid detection, identification, and enumeration of Escherichia coli by fluorescence in situ hybridization using an array scanner. J. Microbiol. Methods 2001, 45, 31. (23) Han, J.-H.; Heinze, B. C.; Yoon, J.-Y. Single cell level detection of Escherichia coli in microfluidic device. Biosens. Bioelectron. 2008, 23, 1303. (24) Nam, J. M.; Thaxton, C. S.; Mirkin, C. A. Nanoparticle-based bio-bar codes for the ultrasensitive detection of proteins. Science 2003, 301, 1884. (25) Seker, F.; Malenfant, P. R. L.; Larsen, M.; Alizadeh, A.; Conway, K.; Kulkarni, A. M.; Goddard, G.; Garaas, R. On-demand control of
optoelectronic coupling in gold nanoparticle arrays. Adv. Mater. 2005, 17, 1941. (26) Reece, P. J. Plasmonics: Finer optical tweezers. Nat. Photonics 2008, 2, 333. (27) Larguinho, M.; Baptista, P. V. Gold and silver nanoparticles for clinical diagnosticsFrom genomics to proteomics. J Proteomics 2012, 75, 2811. (28) Doria, G.; Larguinho, M.; Dias, J. T.; Pereira, E.; Franco, R.; Baptista, P. V. Gold−silver-alloy nanoprobes for one-pot multiplex DNA detection. Nanotechnology 2010, 21, 255101. (29) Ahirwal, G. K.; Mitra, C. K. Direct electrochemistry of horseradish peroxidase−gold nanoparticles conjugate. Sensors 2009, 9, 881. (30) Slawson, R. M.; Van Dyke, M. I.; Lee, H.; Trevors, J. T. Germanium and silver resistance, accumulation, and toxicity in microorganisms. Plasmid 1992, 27, 72. (31) Zhao, G.; Stevens, S. E. Multiple parameters for the comprehensive evaluation of the susceptibility of Escherichia coli to the silver ion. Biometals 1998, 11, 27. (32) Luis, M.; Marzán, L. Tailoring surface plasmons through the morphology and assembly of metal nanoparticles. Langmuir 2006, 22, 32. (33) Wilcoxon, J. Optical absorption properties of dispersed gold and silver alloy nanoparticles. J. Phys. Chem. B 2009, 113, 2647. (34) Mallin, M. P.; Murphy, C. J. Solution-phase synthesis of sub-10 nm Au−Ag alloy nanoparticles. Nano Lett. 2002, 11, 1235. (35) Wang, A. Q.; Lui, J. H.; Lin, S. D.; Mou, C. Y. J. A novel efficient Au−Ag alloy catalyst system: Preparation, activity, and characterization. Catal. 2005, 233, 186. (36) Philip, D. Molecular and biomolecular spectroscopy. Spectrochim. Acta A 2009, 73, 374. (37) Sawle, B. D.; Salimath, B.; Deshpande, R.; Berdre, M. D.; Prabhakar, B. K.; Venkataraman, A. Biosynthesis and stabilization of Au and Au−Ag alloy nanoparticles by fungus Fusarium semitectum. Sci. Technol. Adv. Mater. 2008, 9, 1. (38) Wang, A.; Hsieh, Y. P.; Chen, Y. F.; Mou, C. Y. Au−Ag alloy nanoparticle as catalyst for CO oxidation: Effect of Si/Al ratio of mesoporous support. J. Catal. 2006, 237, 197. (39) Chen, D. H.; Chen, C. J. Formation and characterization of Au− Ag bimetallic nanoparticles in water-in-oil microemulsions. J. Mater. Chem. 2002, 12, 1557. (40) Senapati, S.; Ahmad, A.; Khan, M. I.; Kumar, R. Extracellular biosynthesis of bimetallic Au−Ag alloy nanoparticles. Small 2005, 1, 517. (41) Watson, R. E.; Hudis, J.; Perlman, M. L. Charge flow and d compensation in gold alloys. Phys. Rev. B 1971, 4, 4139. (42) Kim, M. J.; Na, H. J.; Lee, K. C.; Yoo, E. A.; Lee, M. Preparation and characterization of Au−Ag and Au−Cu alloy nanoparticles in chloroform. J. Mater. Chem. 2003, 13, 1789. (43) Han, S. W.; Kim, Y.; Kim, K. J. Dodecanethiol-derivatized Au/ Ag bimetallic nanoparticles: TEM, UV/VIS, XPS, and FTIR Analysis. J. Colloid Interface Sci. 1998, 208, 272. (44) Devarajan, S.; Bera, P.; Sampath, S. Bimetallic nanoparticles: A single step synthesis, stabilization, and characterization of Au−Ag, Au−Pd, and Au−Pt in sol−gel derived silicates. J. Colloid Interface Sci. 2005, 290, 117. (45) Arrii, S.; Morfin, F.; Renouprez, J.; Rousset, J. L. Oxidation of CO on gold supported catalysts prepared by laser vaporization: Direct evidence of support contribution. J. Am. Chem. Soc. 2004, 126, 1199. (46) Liu, J. H.; Wang, A. Q.; Chi, Y. S.; Lin, H. P.; Mou, C. Y. Synergistic effect in an Au−Ag alloy nanocatalyst: CO oxidation. J. Phys. Chem. B 2005, 109, 40. (47) Wolter, A.; Niessner, R.; Seidel, M. Detection of Escherichia coli O157:H7, Salmonella typhimurium, and Legionella pneumophila in water using a flow-through chemiluminescence microarray readout system. Anal. Chem. 2008, 80, 5854.
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dx.doi.org/10.1021/ie3022797 | Ind. Eng. Chem. Res. XXXX, XXX, XXX−XXX
