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Rapid, single-cell electrochemical detection of Mycobacterium tuberculosis using colloidal gold nanoparticles Benjamin Y.C. Ng, Wei Xiao, Nicholas P. West, Eugene J.H. Wee, Yuling Wang, and Matt Trau Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.5b03121 • Publication Date (Web): 18 Sep 2015 Downloaded from http://pubs.acs.org on September 21, 2015

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Analytical Chemistry

Rapid, single-cell electrochemical detection of Mycobacterium tuberculosis using colloidal gold nanoparticles Benjamin Y.C. Ng,1,2 Wei Xiao,1 Nicholas P. West,2 Eugene J.H. Wee,1* Yuling Wang,1* and Matt Trau1,2* 1

Centre for Personalized NanoMedicine, Australian Institute for Bioengineering and

Nanotechnology, The University of Queensland, QLD 4072, Australia. 2

School of Chemistry and Molecular Biosciences, The University of Queensland, QLD

4072, Australia. *[email protected], [email protected], [email protected]

ACS Paragon Plus Environment


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ABSTRACT: Tuberculosis (TB) remains a global health threat, with over a third of the world population suffering from the disease, and 1.5 million deaths due to the disease in 2013 alone. Despite significant advances in TB detection strategies in recent years, a bigger push towards detecting TB in the shortest and easiest way possible at the point-ofcare (POC) is still in demand. To this end, we have designed a simple yet rapid and sensitive bioassay that detects Mtb DNA electrochemically using colloidal gold nanoparticles. This assay couples rapid isothermal amplification of target DNA that is specific to Mtb with gold nanoparticle electrochemistry on disposable screen printed carbon electrodes. The assay is capable of detecting a positive differential pulse voltammetry (DPV) response from as low as 1 CFU of Mtb bacilli DNA input material, and having shown its exquisite sensitivity over a conventional gel based readout. The translation of our assay onto a portable potentiostat was also demonstrated, with promising results. We believe that our assay has significant potential for translation into broader bioassay applications or development as a POC diagnostic tool.


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INTRODUCTION Tuberculosis (TB), which is caused by the acid-fast bacillus Mycobacterium tuberculosis (Mtb), has one of the highest infection rates in the world, with over a third of the world population suffering from the disease, and 1.5 million deaths due to the disease in 2013 alone.1 As such, it remains a vital health threat to people from many parts of the world, thus prompting the World Health Organization (WHO) to initiate a global plan to stop TB. The WHO has identified several key factors to eradicate the disease; among these factors is the urgent need to rapidly identify cases of TB in order to reduce time to treatment and subsequently prevent disease transmission.1 Conventional TB diagnostics include the tuberculin skin test and sputum smear microscopy, methods that are acknowledged to have several issues ranging from poor sensitivity and/or specificity and false negative results.2-4 A relatively new test, the interferonγ release assay, was shown to be highly sensitive but does not differentiate between active TB disease and latent TB infection.4,5 Moreover, it requires storage of whole blood or viable white blood cells, which can be challenging in resource-poor settings.4,5 While TB culture remains the gold standard for definitive diagnosis, it is a lengthy process that significantly delays time to treatment for TB patients, and in turn, increases the risk of continued transmission of the disease and thus mortality rates.6 In resource-poor settings, a large hurdle that remains is the challenge in bringing the capability of detecting TB out of the laboratory and into point-of-care (POC) – especially in areas where rapid turnaround diagnosis is most needed, and where sophisticated diagnostic equipment are unavailable.6 In 2010, the Xpert MTB/RIF real-time PCR platform7 was endorsed by the WHO for the simultaneous diagnosis of TB and Rifampicin (RIF) resistance.1 Most significantly, it is seen as an advance in the right direction of decreasing time to an accurate diagnosis.8 However, the Xpert MTB/RIF device does present some limitations related to potential POC use. The high cost instrumentation and single-use cartridges increases the economic burden in resource-poor settings, even after heavy subsidization through sponsorships.9 Its reliance on a stable power supply, regular maintenance and calibrations all contribute to a challenge in many remote settings.9,10 Therefore, we believe that there still is an urgent need for new diagnostic tools that can be easily used as a screening tool in remote areas of high TB burden, and at a much lower cost than conventional diagnostic methods.3,6 In recent years, methods to isothermally amplify DNA have become increasingly popular due to their potential for on-field DNA detection strategies. Unlike traditional PCR based



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methods such as that used in the Xpert MTB/RIF device, isothermal amplification methods circumvent the need for thermal cycling.9,11 This is a major advantage when considering that a consistent supply of power for thermal cycling cannot be taken for granted at POC, especially in remote settings. Of the various isothermal amplification methods, Recombinase Polymerase Amplification (RPA)12 was recently demonstrated for Mtb detection.13 The study uses a fluorescence-based detection platform which, while effective, might present a potential cost barrier in low resource applications. Hence there is still a need for alternative low-cost readout methods of RPA that can be readily deployed in the field. Electrochemical sensors have inherent advantages such as improved sensitivity to minute amounts of sample input at relatively low cost.14,15 In particular, gold nanoparticles (AuNPs) have been used extensively as labels for electrochemical detection due to their simple synthesis, ease of bio-functionalization and the monodispersity of particle sizes.14,16,17 Moreover, AuNPs can be chemically oxidised in Br2 or HCl to form electrochemically active species AuCl4-, which can then be detected with electrochemical analytical tools such as differential pulse voltammetry (DPV).14,15 Recently, the direct detection of Mtb using surface plasmon resonance with the hybridization of AuNP labelled probes was demonstrated;18 however, the method required the hybridization of gold labelled probes to complementary DNA targets – a process that requires laboratory equipment for thermal denaturation of DNA and the detection of spectral shifts.18 Such requirements could hinder a translation of these methods to POC detection in the field. Herein, we describe a method to cheaply and rapidly interpret RPA amplified Mtb DNA using electrochemical detection of colloidal gold nanoparticles after DNA extraction from Mtb. This bioassay is highly specific to Mtb, has a sensitivity of a single Mtb cell and can be analysed with portable electrochemical readout devices for POC applications. In addition, this bioassay makes use of disposable screen printed carbon electrodes (SPCEs) surface functionalized with streptavidin (SA) molecules that have desirable characteristic such as good reproducibility19 at a relatively cheap cost (~USD $5 per SA modified SPCE). An added advantage of being disposable, the use of SPCEs potentially avoids issues with contamination or time consuming cleaning process typically associated with conventional carbon or gold electrodes.20


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EXPERIMENTAL SECTION Materials. Carboxyl coated magnetic beds were purchased from Thermo Fisher (Cat# 45152105-050250). TwistAmp Basic RPA Kit was purchased from TwistDX (Cat# TABAS01KIT). Biotin-11 DUTP was purchased from Biotium (Cat# 40029). Primers were purchased from Integrated DNA Technologies. All other chemicals were purchased from Sigma-Aldrich. DNA sample acquisition. M. tuberculosis H37Ra was cultured in Middlebrook 7H9 complete broth with supplementation (10% ADC, 0.2% glycerol and 0.02% tyloxapol). Samples for assay testing were taken at mid-exponential phase (OD600 = 0.8). These samples were serially diluted and plated on Middlebrook 7H11 agar (0.5% glycerol and 10% OADC) and the number of colony forming units (CFU) was visually determined. Nucleic acid extraction and purification. We have employed a modified Solid Phase Reversible Immobilization (SPRI) protocol21,22 with Guanidium-HCl lysis buffer to sample for genomic DNA (gDNA) from Mtb cells. Mtb gDNA was extracted directly from culture by adding 2 volumes of an optimized lysis buffer (100 mM Tris-HCl pH 8.0, 3 M guanidiumHCl. 400 ng/µL RNase A and 2% v/v Triton-X) to 1 volume of culture. After 15 min incubation at room temperature, nucleic acids were purified with SPRI. 1 volume of lysate was incubated with 2 volumes of 1 micron carboxyl coated magnetic beads in a binding buffer (10 mM Tris-HCl pH 8.0, 20% PEG8000, 2.5 M NaCl) for 10 minutes. These DNA bound beads were then separated from the lysate with a magnet and washed twice with 100% isopropanol, twice with 80% ethanol and eluted in one volume of water. Nucleic acid amplification. Nucleic acid amplification was performed with the TwistAmp Basic RPA Kit as recommended by the manufacturer with some modifications. 0.5 µL of 1mM biotin-11 dUTP was added into 12.5 µL reactions. These reactions were performed at 38°C for 20 min using 1 µL of the nucleic acid extraction and 800 nM of each primer. We designed a set of primers that amplified a region within the RNA polymerase beta subunit (RpoB) gene of Mtb (Table 1). Mutations occurring in a defined 81 base pair region within the RpoB, commonly termed the Rifampicin Resistance Determining Region (RRDR), limit the effectiveness of the drug Rifampicin.23,24 Following amplification, 3 µL of the RPA reaction was visualized by gel electrophoresis.



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Table 1. List of RPA primers used. GenBank Accession numbers are as given.

Target / GenBank

M. tuberculosis RRDR

Accession

CP003248.2

5'-Forward-3'

AACCGACGACATCGACCACTTCGGCAACCG

5'-Reverse-3'

CCAGCGCCGACAGTCGGCGCTTGTGGGTCAA

Synthesis of AuNPs. AuNPs were prepared according to the procedure reported.25 Briefly, 100 mL of 1 mM chloroauric acid (HAuCl4) was heated until boiling, 10 mL of 1% 38.8 mM sodium citrate (Na3C6H5O7) was then added to the flask and the mixture was heated for another 10 minutes. After 10 minutes, the flask was brought to cool and the contents stirred for 30 minutes. Surface modification of AuNPs with streptavidin. The as-prepared AuNPs were then coated by SA according to the reported protocol with slight modification.26 Briefly, 1 mL of AuNPs was mixed with 10 µL of 1 M NaHCO3 for pH adjustment (9-10). 25 µL of 1 mg/mL streptavidin was then incubated with AuNPs at RT for 30 min, followed by the addition of 20 µL of 2% PEG. AuNP-SA was then centrifuged at 12500 rpm twice at 4 °C for 25 minutes to remove the excess SA and PEG. The pellet was then dissolved in 200 µL PBS. Electrochemical assay and readout with AuNP labels. Amplified DNA from RPA was first purified using the protocol for DNA purification from above, but with 0.8 volumes of magnetic bead solution to 1 volume of RPA reaction. 3 µL of each sample was incubated on the working electrode of the SPCEs for 15 minutes at room temperature. Following which, the samples were washed out with 60 µL 1 M phosphate buffered saline (PBS) 3 times to remove all unbound DNA. An optimized blocking buffer (5 µL of 0.1% BSA and 5 µL of 0.5% PEG) was then added to the working electrode and left for 15 minutes to block the surfaces of the working electrode. The blocking buffer was then removed and the electrode surface washed with 60 µL of 1 M PBS 3 times. 3 µL of AuNP-SA was added and left to incubate on the working electrode surface for 10 minutes at room temperature. The unbound AuNP-SA particles were then washed off using 1 M PBS with 0.1% Tween-20 3 times. The AuNPs were subsequently electrochemically activated with 60 µL of 0.1 M HCl at a preoxidation potential of 1.3V for 30 seconds to oxidise all Au0 to Au3+. The subsequent reduction of this species was then detected with Differential Pulse Voltometry (DPV)


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response, which was taken from a starting potential of 0.15 V to an ending potential of 0.45 V, with a step potential of 0.004 V, pulse period of 0.2 s, and pulse amplitude of 0.05 V. Instruments. AuNPs were characterized with transmission electron microscopy (TEM) (JEOL-2100). UV-Vis absorption spectra of AuNPs before and after SA coating were determined with the Nanodrop spectrophotometer ND-1000. DPV responses were measured on a workstation potentiostat (CH Instruments) and hand-held potentiostat (Metrohm). RESULTS AND DISCUSSION Scheme design and characterization of gold nanoparticles: In our work, we have combined the versatility of AuNPs as labels for electrochemical response with the benefits of isothermal DNA amplification for highly sensitive and rapid POC detection methodology for Mtb DNA (Fig. 1). We first sample for Mtb DNA using SPRI, which avoids the reliance on multiple centrifugation steps often used in conventional DNA extraction kits and thus more suitable for POC applications.

Figure 1. Schematic representation of the electrochemical detection strategy from DNA extraction to readout. A positive DPV signal is generated when there are sufficient AuNPs immobilized on the working electrode surface. After gDNA purification, we used RPA as it has been shown to be a robust and sensitive isothermal method for DNA amplification12,13 (other isothermal methods may also be applicable for amplifying our target sequence). The RPA was supplemented with biotindUTPs that randomly substitute dTTPs to create biotinylated amplicons. The SPRI method used earlier for sampling is then used again to select for specifically amplified high molecular weight products while removing excess biotin dUTPs and primers/dimers.



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We then utilize the highly specific and strong binding interaction between biotin and SA in a sandwich format labelling assay. Biotinylated amplicons were then allowed to spontaneously react with SA-coated surface of the working electrode on SPCEs. As the amplicons had multiple biotins, the free biotins could function as a substrate for subsequent AuNP-SA binding. Following a wash to remove excess DNA, a BSA/PEG blocking buffer was used prior to the addition of AuNP-SA to minimize non-specific adsorption. Finally, the amount of captured AuNPs, representing the presence of amplified Mtb DNA, was detected electrochemically. To the best of our knowledge, there is currently no Mtb DNA detection assay combining the use of RPA with AuNP-based electrochemical detection on SPCEs. The as-prepared AuNPs were characterized by TEM (Fig. 2A) and UV-Vis absorption spectroscopy (Fig. 2B) to investigate their size and monodispersity as well as their optical properties, respectively. The TEM image showed a monodispersed AuNP with the size of ~13 nm. The UV-Vis absorbance spectrum of AuNPs before SA coating was observed at 520 nm (Fig. 2B). Upon SA coating, the absorption of AuNPs red-shifted to 526 nm, which is due to the change of the surrounding refractive index.

A

B

Figure 2. (A) TEM image of as-prepared AuNPs. (B) Absorption spectra of AuNPs before and after streptavidin (SA) coating. A red shift indicates after the coating of SA on AuNPs. Sensitivity of assay for TB detection: In many nucleic acid detection assays,13 insertion elements IS6110 and IS1081 are popular targets due to multiple copies of these sequences per Mtb genome that could result in greater assay sensitivity.27 However, IS6110 has also been found in other mycobacterial species,27 and thus may limit its use as a specific target for TB diagnostic purposes. Unlike IS1081 and IS6110, the RRDR is present only as a single copy in the genome, is highly specific to the Mtb genome and is currently used as a target in the Xpert MTB/RIF platform due to minimal cross reactivity with other innocuous mycobacterial


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species.7,28 Using our primer set for RRDR, we determined the limit of detection of this assay by titrating aliquots of Mtb cells and assaying them with our method. The same aliquots were also grown on culture plates to estimate bacterial titre which is measured by the number of colony forming units (CFU). As controls we used a no DNA template (NTC) to verify the fidelity of the amplification reaction, and a PBS buffer only to discern any effect of nonspecific binding of AuNPs on the working electrode. Gel electrophoresis was used to visualize and validate RPA amplification, and also as a comparison to determine the sensitivity of the electrochemical readout (Fig. 3).

Figure 3. (A) DPV responses for AuNP-SA labelled DNA duplexes immobilized on SPCE from 1000, 100, 10, 1 CFU of Mtb, NTC and PBS. (B) Mean DPV peak current values of products with input samples of (L-R) 1000, 100, 10, 1 CFU of Mtb, NTC and PBS. The error bars represent the standard deviation of a minimum of 3 separate experimental replicates. (C) Gel electrophoresis of products of amplification with input Mtb samples serial diluted 10-fold (L-R) 1000, 100, 10, 1 CFU and NTC.

Our results showed that as low as 1 CFU could be detected via the electrochemical assay with DPV responses 5 times higher than that of controls (Fig. 3A and 3B), however, only a faint band corresponding to a dilution of 10 CFU could be observed under gel electrophoresis (Fig. 3C), thus underscoring the superior sensitivity of electrochemical-based readouts. Furthermore, the minimal responses of both the NTC and PBS only controls are evidence of sufficient blocking of the working electrode and AuNP surfaces to prevent non-specific binding and consequentially, negligible background noise. The relative standard deviation of our assay is 8.3% (n = 18), thus showing good inter-assay reproducibility. While there have been reports of isothermal Mtb DNA detection methods that claim detection limits approaching a single Mtb cell, however, it should also be noted that these



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methods commonly target insertion elements IS6110 and IS1081 which appear in multiple copies per Mtb cell and may have specificity issues.13,29 As a comparison, the Xpert MTB/RIF real time PCR platform, which uses similar molecular targets as our assay, has reported a sensitivity of 131 CFU.8 We thus believe that the high sensitivity of our method could find applications in detecting trace amounts of Mtb present in samples, such as in paucibacillary cases of childhood TB,30 or bacteria present in the oral mucosa of TB patients.31 To further demonstrate the stringency of the BSA/PEG blocking step and that any response was a result of biotinylated DNA, a control was performed with the various dilutions of Mtb cells for which no biotin dUTPs were added to the RPA reaction. The control experiments showed that the DPV responses for non-biotinylated DNA products were minimal thus proving the efficacy of the BSA/PEG block step and that any observed response was indeed due the presence of biotinylated DNA. (Fig. 4).

Figure 4. Peak current values of 1000 CFU, 10 CFU, 1 CFU, PBS and NTC samples without any biotin incorporation during RPA but with AuNPs added. Specificity of the assay: In the interest of demonstrating assay specificity towards Mtb, we challenged the assay with non-Mtb targets: Escherichia coli and HeLa DNA. 1 ng of gDNA for each bacterial species was used as template for RPA. As expected, only the Mtb sample gave a significant positive DPV signal as compared to the other non-Mtb samples (Fig. 5). The gel electrophoresis results also validated the DPV responses, where only an amplification band was seen in the Mtb lane.


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Figure 5. (A) Mean DPV peak current values of post-amplification products with input DNA templates from (L-R) Mtb, HeLa, E.coli and NTC. The error bars represent the standard deviation of a minimum of 3 separate experimental replicates. (B) Gel electrophoresis of runs in Fig. 5 (A). Point-of-care detection with handheld potentiostat: Towards the eventual goal of realizing a POC diagnostic assay, we have also translated our assay onto a portable potentiostat (Fig. S1A), which can be powered by a laptop computer and therefore better suited for field applications. The results were agreeable with earlier results obtained with the workstation electrochemical sensor, with Mtb positive samples exhibiting much higher DPV responses as compared to no template controls (Fig. S1B). The entire assay, which costs under USD $10 per run, can be completed in under 90 minutes. This assay can potentially provide a patient with a definitive test within the span of a visit, and thus allow for the timely treatment of TB positive cases. All steps in the assay can be performed and interpreted easily with minimal training, relative to other laboratory based DNA detection methods such as line probe assays,32 fluorescence imaging13 and high performance liquid chromatography.33 CONCLUSIONS In conclusion, we have developed a low cost yet highly sensitive assay that is able to detect as low as 1 CFU of Mtb DNA in less than 90 minutes with high specificity, by taking advantage of the benefits of electrochemical detection of RPA products with colloidal gold nanoparticles. The use of RPA and disposable SPCEs potentially adds convenience and portability at low cost while avoiding contamination issues on the field. By making use of portable electrochemical sensors, the assay has very high potential in low resource POC settings. We believe that our assay can be used as a complement to current diagnostic



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methods and address the growing need for rapid and sensitive detection of TB in remote regions that currently have limited means to do so. ACKNOWLEDGMENTS B.N. acknowledges the TB research scholarship awarded by the Centre for Australian Military and Veterans’ Health and administered through the School of Public Health, University of Queensland. This project is partly supported by the Australian Research Council under the Discovery Early Career Researcher Award scheme (DE 140101056) awarded to Y.W. The authors would also like to thank MEP Australia for generously loaning the portable potentiostat used in DPV experiments. REFERENCES (1) World Health Organization 2014. Global tuberculosis report 2014. http://www.who.int/tb/publications/global_report/en/. Accessed May 16, 2015. (2) Andersen, P.; Munk, M. E.; Pollock, J. M.; Doherty, T. M. Lancet 2000, 356, 1099-1104. (3) Lawn, S. D.; Zumla, A. I. Lancet 2011, 378, 57-72. (4) Maartens, G.; Wilkinson, R. J. Lancet 2007, 370, 2030-2043. (5) Pai, M.; Minion, J.; Sohn, H.; Zwerling, A.; Perkins, M. D. Clin. Chest Med. 2009, 30, 701716. (6) Wallis, R. S.; Pai, M.; Menzies, D.; Doherty, T. M.; Walzl, G.; Perkins, M. D.; Zumla, A. Lancet 2010, 375, 1920-1937. (7) Lawn, S. D.; Nicol, M. P. Future Microbiol. 2011, 6, 1067-1082. (8) Helb, D.; Jones, M.; Story, E.; Boehme, C.; Wallace, E.; Ho, K.; Kop, J.; Owens, M. R.; Rodgers, R.; Banada, P.; Safi, H.; Blakemore, R.; Lan, N. T. N.; Jones-Lopez, E. C.; Levi, M.; Burday, M.; Ayakaka, I.; Mugerwa, R. D.; McMillan, B.; Winn-Deen, E.; Christel, L.; Dailey, P.; Perkins, M. D.; Persing, D. H.; Alland, D. J. Clin. Microbiol. 2010, 48, 229-237. (9) McNerney, R.; Daley, P. Nature Reviews Microbiology 2011, 9, 204-213. (10) Boehme, C. C.; Nicol, M. P.; Nabeta, P.; Michael, J. S.; Gotuzzo, E.; Tahirli, R.; Gler, M. T.; Blakemore, R.; Worodria, W.; Gray, C.; Huang, L.; Caceres, T.; Mehdiyev, R.; Raymond, L.; Whitelaw, A.; Sagadevan, K.; Alexander, H.; Albert, H.; Cobelens, F.; Cox, H.; Alland, D.; Perkins, M. D. Lancet 2011, 377, 1495-1505. (11) McNerney, R.; Maeurer, M.; Abubakar, I.; Marais, B.; McHugh, T. D.; Ford, N.; Weyer, K.; Lawn, S.; Grobusch, M. P.; Memish, Z.; Squire, S. B.; Pantaleo, G.; Chakaya, J.; Casenghi, M.; Migliori, G. B.; Mwaba, P.; Zijenah, L.; Hoelscher, M.; Cox, H.; Swaminathan, S.; Kim, P. S.; Schito, M.; Harari, A.; Bates, M.; Schwank, S.; O'Grady, J.; Pletschette, M.; Ditui, L.; Atun, R.; Zumla, A. J. Infect. Dis. 2012, 205, S147-S158. (12) Piepenburg, O.; Williams, C. H.; Stemple, D. L.; Armes, N. A. PLoS Biol. 2006, 4, 11151121. (13) Boyle, D. S.; McNerney, R.; Low, H. T.; Leader, B. T.; Perez-Osorio, A. C.; Meyer, J. C.; O'Sullivan, D. M.; Brooks, D. G.; Piepenburg, O.; Forrest, M. S. PLoS One 2014, 9, e103091. (14) Leng, C. A.; Lai, G. S.; Yan, F.; Ju, H. X. Anal. Chim. Acta 2010, 666, 97-101. (15) Xu, Q.; Yan, F.; Lei, J.; Leng, C.; Ju, H. Chemistry-a European Journal 2012, 18, 49944998. (16) Low, K. F.; Rijiravanich, P.; Singh, K. K. B.; Surareungchai, W.; Yean, C. Y. Journal of Biomedical Nanotechnology 2015, 11, 702-710. (17) Pinijsuwan, S.; Rijiravanich, P.; Somasundrum, M.; Surareungchai, W. Anal. Chem. 2008, 80, 6779-6784.


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(18) Baptista, P. V.; Koziol-Montewka, M.; Paluch-Oles, J.; Doria, G.; Franco, R. Clin. Chem. 2006, 52, 1433-1434. (19) Yu, H.; Yan, F.; Dai, Z.; Ju, H. X. Anal. Biochem. 2004, 331, 98-105. (20) Fischer, L. M.; Tenje, M.; Heiskanen, A. R.; Masuda, N.; Castillo, J.; Bentien, A.; Emneus, J.; Jakobsen, M. H.; Boisen, A. Microelectronic Engineering 2009, 86, 1282-1285. (21) Deangelis, M. M.; Wang, D. G.; Hawkins, T. L. Nucleic Acids Res. 1995, 23, 4742-4743. (22) Hawkins, T. L.; Oconnormorin, T.; Roy, A.; Santillan, C. Nucleic Acids Res. 1994, 22, 4543-4544. (23) Miller, L. P.; Crawford, J. T.; Shinnick, T. M. Antimicrob. Agents Chemother. 1994, 38, 805-811. (24) Ramaswamy, S.; Musser, J. M. Tuber Lung Dis 1998, 79. (25) Daniel, M. C.; Astruc, D. Chem. Rev. 2004, 104, 293-346. (26) Yokota, S. In Immunoelectron Microscopy: Methods and Protocols, Schwartzbach, S. D.; Osafune, T., Eds.; Humana Press Inc: Totowa, 2010, pp 109-119. (27) Coros, A.; DeConno, E.; Derbyshire, K. M. J. Bacteriol. 2008, 190, 3408-3410. (28) Piatek, A. S.; Tyagi, S.; Pol, A. C.; Telenti, A.; Miller, L. P.; Kramer, F. R.; Alland, D. Nat. Biotechnol. 1998, 16, 359-363. (29) Aryan, E.; Makvandi, M.; Farajzadeh, A.; Huygen, K.; Bifani, P.; Mousavi, S. L.; Fateh, A.; Jelodar, A.; Gouya, M. M.; Romano, M. Microbiol. Res. 2010, 165, 211-220. (30) Newton, S. M.; Brent, A. J.; Anderson, S.; Whittaker, E.; Kampmann, B. Lancet Infect. Dis. 2008, 8, 498-510. (31) Wood, R. C.; Luabeya, A. K.; Weigel, K. M.; Wilbur, A. K.; Jones-Engel, L.; Hatherill, M.; Cangelosi, G. A. Sci. Rep. 2015, 5, 8668. (32) Luetkemeyer, A. F.; Kendall, M. A.; Wu, X. Y.; Lourenco, M. C.; Jentsch, U.; Swindells, S.; Qasba, S. S.; Sanchez, J.; Havlir, D. V.; Grinsztejn, B.; Sanne, I. M.; Firnhaber, C.; Adult, A. C. T. G. A. J. Clin. Microbiol. 2014, 52, 1052-1059. (33) Butler, W. R.; Jost, K. C.; Kilburn, J. O. J. Clin. Microbiol. 1991, 29, 2468-2472.



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