Enzymatic Litmus Test for Selective Colorimetric Detection of C–C

Mar 14, 2019 - Addition of DNA containing C–C mismatches reactivates urease via binding of Ag(I), allowing restoration of urease activity, hydrolysi...
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Enzymatic Litmus Test for Selective Colorimetric Detection of C−C Single Nucleotide Polymorphisms Michael G. Wolfe, M. Monsur Ali, and John D. Brennan* Biointerfaces Institute, McMaster University, 1280 Main Street West, Hamilton, ON L8S 4O3, Canada

Anal. Chem. Downloaded from pubs.acs.org by EAST CAROLINA UNIV on 03/14/19. For personal use only.

S Supporting Information *

ABSTRACT: A paper based litmus test has been developed using modulation of urease enzyme activity for detection of C−C mismatch single nucleotide polymorphisms (SNPs) by the naked eye. Urease is first inactivated with silver ions and printed onto paper microzones. Addition of DNA containing C−C mismatches reactivates urease via binding of Ag(I), allowing restoration of urease activity, hydrolysis of urea to produce ammonia, and an increase in pH, which is monitored colorimetrically using a pH indicator with a limit of detection of 11 nM DNA in 40 min. The assay system is easy to use, portable, and stable for at least 30 days at ambient temperature. To assess the versatility and practical application of the paper sensor, we used it to identify a G > C transversion present in human genomic DNA from a ductal carcinoma cell line, a mutation commonly found in breast cancer. We believe this new assay system has the potential to be a low-cost method for rapidly identifying DNA with the C−C mismatch SNP as a means of cancer screening in resource-limited areas.

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INTRODUCTION With over 9 million single nucleotide polymorphisms (SNPs) identified in the human genome, these mutations account for approximately 90% of human genetic variation.1,2 SNPs are related to various worldwide health problems such as cancer, genetic diseases, and development of drug-resistant bacteria.3−7 The past decade has seen tremendous advancements in methods to identify these SNPs, including those based on DNA microarrays, quantitative polymerase chain reactions and next generation sequencing methods.8,9 However, these methods are still limited to laboratory settings which require expensive instrumentation and highly trained personnel. With the high cost of both specialized equipment and trained technicians, traditional laboratory diagnostics are increasingly being challenged by the advancement in pointof-care (POC) diagnostic technologies.10−13 Paper based POC diagnostics in particular provide many advantages over labbased assays, such as being low-cost, disposable, simple to manufacture, and portable for use in resource-limited areas.14−21 Paper based biosensors have been developed by our lab and others to identify a variety of targets including environmental toxins,22,23 proteins,24−26 bacteria,27,28 DNA, and RNA.29 Paper based biosensors have been designed using a multitude of detection platforms, including colorimetric, fluorimetric, chemiluminescent, and electrochemical outputs.14−17,19 Of these platforms, colorimetric detection is the simplest method to allow equipment-free detection by the naked eye and can be developed by linking enzymatic reactions to simple pH sensitive dyes.20,21,30−32 © XXXX American Chemical Society

It is well-known that urease, a Ni(II) dependent enzyme, is susceptible to trace metal inhibition,33−38 and based on this phenomena, methods for detection of several metal ions have been developed.39−43 Hg(II), Ag(I), and Cu(II) are particularly effective urease inhibitors due to their strong reactivity with thiol, amine, and oxygen functional groups within the enzyme active site.34,36,38 In parallel, DNA undergoes numerous interactions involving complexation with these same metals. One key interaction is the complexation between cytosine-cytosine (C−C) mismatched DNA base pairs and Ag(I) ions.44−46 We hypothesized that the C−C mismatch DNA could extract the Ag(I) ions from the urease/Ag(I) complex, resulting in reactivation of the urease to hydrolyze the urea substrate and produce ammonia. To generate a signal, the substrate solution is set to an initial pH which is slightly acidic (pH 5.8) and contains a pH indicator, phenol red, which is yellow below pH 7.0. Once the pH increases due to the formation of ammonia by the urease catalyzed reaction, the indicator produces a red color, which can be observed with the naked eye (Figure 1). In the case where the DNA in the sample is fully matched (no mutation), the urease enzyme should remain inactive and the solution color will remain yellow. We show that this assay works both in a solution-based microwell plate assay and as a printed paper sensor. We further demonstrate that the sensing method can be used to detect a Received: January 14, 2019 Accepted: March 7, 2019

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DOI: 10.1021/acs.analchem.9b00235 Anal. Chem. XXXX, XXX, XXX−XXX

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μL) by hand and then depositing 10 μL onto each microwell zone using a TECAN EVO 200 liquid handler. Samples were dried for a minimum of 2 h, and typically overnight at room temperature. Once dry, the bioactive paper was stored at room temperature until used. Analysis of DNA Activity on Paper. DNA samples were tested by adding samples (ranging from 0−50 μM of dsDNA in 150 mM NaCl, pH 5.8, 10 μL) to a paper microzone well followed by incubation for 20 min at room temperature. Substrate (500 mM urea, 14 mM phenol red, pH 5.8, 18 μL) was then added and color was allowed to develop for 20 min, after which images were obtained using a BlackBerry Q5 cellular phone or LG G5 cellular phone operated in automatic mode. Images were analyzed using ImageJ software as follows: images were split into red/blue/green color channels, with the blue and red channels being discarded. The green channel was inverted, and then the intensity of color in each zone was quantified using the “measure” function yielding the values on a scale of 0−255. Wells containing only pullulan (no enzyme) were used to subtract the background signal. Assay Validation Using p53 Model System. Using purified human genomic DNA containing a G > C transversion in the TP53 gene, the polymerase chain reaction (PCR) was performed to amplify a 50-nt region containing the mutation of interest. Using a fully complementary forward and reverse primer (bt-549-wtRP and bt-549-FP, respectively), a dsDNA oligonucleotide was produced to mimic a healthy control (termed “p53wt”). To keep the G > C mutation present, a separate PCR was performed using a different reverse primer that contained the C nucleotide mismatch (bt-549-mRP). In DNA containing the G > C transversion, amplification using the bt-549-mRP primer resulted in a percentage of dsDNA containing a C−C mismatch (denoted as “p53m”). The PCR amplification was performed in two rounds. The first round PCR mixture (50 μL) contained 100 ng of the genomic DNA along with 1 μM forward primer, 1 μM reverse primer (wildtype or mutant), 200 mM dNTPs, 1× PCR reaction buffer (75 mM Tris-HCl, 2 mM MgCl2, 50 mM KCl, 20 mM (NH4)2SO4, pH 9), and 2.5 U DNA polymerase (BTL-10043, Biotools, Spain). Thermal cycles were performed as follows: 94 °C for 60 s, followed by 25 cycles of 90 °C for 45 s, 55 °C for 30 s, and 70 °C for 30 s. After this, the PCR product was diluted 1:50 in ddH2O. Then 1 μL of this diluted product was further amplified an additional 10 rounds using the same thermal cycles as above. PCR products were confirmed by agarose gel electrophoresis (2% w/v containing 1× SYBR GOLD) to ensure successful amplification. The PCR products were purified using a 30 kDa spin column (#UFC5000396, Millipore), and the concentration was determined using a NanoQuant plate on a TECAN M200 plate reader. Following the paper assay format previously described, the two p53 dsDNA constructs were compared. Samples containing ∼150 ng of amplified DNA, corresponding to a concentration of 600 nM, were added to the paper test zone (in 150 mM NaCl, 10 μL) for 20 min. After incubation, substrate (500 mM urea, 14 mM phenol red, pH 5.8, 18 μL) was added and images were taken after 20 min and analyzed as described above. Stability of Paper Biosensor and Microwell Plate Assay. The paper biosensor and microwell plate assay system were prepared and left at room temperature on a lab bench in ambient lighting without any special storage conditions. DNA samples were prepared and aliquoted out for each day and then

Figure 1. General schematic of the proposed assay. Urease (blue circle) is mixed with Ag(I) (black circle), inactivating the enzyme. Addition of dsDNA containing a C−C mismatch sequesters the Ag(I) ions from urease and reactivates the enzyme. Active urease then hydrolyzes urea into carbon dioxide and ammonia, which increases the pH of the system. This increase in pH causes the phenol red indicator to change from yellow to red (top image), which can be monitored by the naked eye. In the presence of fully matched DNA, urease remains inactive and no color change occurs (bottom image).

C−C mismatch in the TP53 gene from a mutant human breast ductal carcinoma cell line, a key biomarker for breast cancer detection.

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EXPERIMENTAL SECTION Oligonucleotides and Chemicals. All DNA oligonucleotides (Table S1) were purchased from Integrated DNA Technologies (IDT). Pullulan (molecular weight of ∼200 000 Da) was purchased from Polysciences Inc. Nitrocellulose membranes (HF180) were purchased from Millipore. Urease from Canavalia ensiformis (U1500, 40 318 U/g) was from Sigma-Aldrich (Oakville, Canada). Human genomic DNA containing a G > C transversion (HTB-122DQ) was purchased from Cedarlane Laboratories (Burlington, Canada). All other chemicals were purchased from Sigma-Aldrich and used without further purification. Solution-Based Urease/Ag(I) Assay. DNA oligonucleotides were mixed (in 150 mM NaCl) and heated at 90 °C for 5 min and then cooled to room temperature for 20 min. AgNO3 (62.5 nM in ddH2O, 16 μL) was mixed with urease (227 nM in ddH2O, 2 μL) for 20 min to inactivate the enzyme. DNA (ranging from 0−50 μM, 18 μL) was then added and mixed for 20 min. Addition of substrate (500 mM urea, 14 mM phenol red, pH 5.8, 82 μL) initiated the color change. End point absorbance was measured at 570 nm after 20 min using a Tecan M1000 platereader. “Percent activity” was determined by the following formula: percent activity = (A T − A 0)/(AP − A 0) × 100%

(1)

where AT is the absorbance of the test sample, A0 is the absorbance of the blank, and AP is the absorbance of the positive control (urease without any Ag(I) present). Preparation of Bioactive Paper. A wax microzone plate consisting of 96 × 4 mm diameter wells was printed in an 8 × 12 pattern onto a HF180 nitrocellulose membrane using a Xerox ColorQube 8570N wax printer and then heated at 125 °C for 3 min in a lab oven to melt the wax into the paper pores. The paper was cooled at room temperature for 30 min before use. A master-mix was made by mixing urease (2.3 μM, 100 μL), AgNO3 (6.25 μM, 100 μL), and pullulan (8% w/v, 800 B

DOI: 10.1021/acs.analchem.9b00235 Anal. Chem. XXXX, XXX, XXX−XXX

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Analytical Chemistry stored frozen at −20 °C until testing. We chose to run the assays at DNA concentrations that had been shown to produce high enzyme activity. Therefore, both the paper assay and the microwell plate assay were performed with 2.5 μM DNA using conditions as noted above. The end point color intensity was normalized to 100% at day zero and relative activity was determined over a period of 30 days.

of 68.9 ± 1.1 nM, compared to 20.6 ± 1.1 nM for TC2. The limit of detection (LOD) was 11 nM for a single dsDNA mismatch (TCAs ± 3 σ) and 4.5 nM for DNA with two C−C mismatches. The above results reveal that the detection system performs well in solution and suggests that optimal differentiation between 0, 1, and 2 mismatches occurs at a DNA concentration of approximately 50 nM. However, this assay platform requires multiple steps and uses an expensive microplate reader for quantification. To alleviate these limitations, the assay was converted into a paper based sensor platform (Figure 3). The urease/Ag(I) mixture was mixed with

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RESULTS AND DISCUSSION The minimum concentration of the urease enzyme required to alter the pH and elicit the change of color from yellow to pink, as well as the ratio between urease and Ag(I) to fully inactivate the enzyme, were first determined. The experimental results showed that 4.5 nM of urease was required to produce a full color change within 20 min (Figures S1 and S2a). The results also showed that 10 nM of Ag(I) was able to completely inactivate 4.5 nM of the enzyme (Figure S2b). This ratio of ∼2:1 Ag(I):urease falls within the range of 2:1 to 12:1 that has previously been reported in the literature.33,47,48 Using these optimized enzyme and inhibitor concentrations, it was possible to reduce the total assay time to 40 min (20 min incubation +20 min color development), making the assay compatible with rapid testing. The ability of the urease/Ag(I) system to detect DNA containing C−C mismatches in solution was then examined using a microwell plate assay format. Three DNA duplexes were used to evaluate the assay system: (1) a fully matched duplex of 25 nucleotides as a control (termed “TCAs” in Table S1 in the Supporting Information), (2) a duplex DNA of 25 nucleotides with a single C−C mismatch (“TC1”), and (3) a duplex DNA of 25 nucleotide with a double C−C mismatch (“TC2”). The DNA duplexes were incubated with the inactivated urease/Ag(I) complex for 30 min and the recovery of enzyme activity was observed based on the color development 30 min after addition of the urease/phenol red substrate solution. The results shown in Figure 2 indicated that

Figure 3. Schematic of the paper assay. Deactivated urease in a pullulan solution is printed on microzones and dried to form pullulan films. Addition of the aqueous solution of dsDNA dissolves the pullulan, liberating the deactivated urease. DNA that is fully complementary has no affinity for the Ag(I) ions resulting in no urease reaction, as observed by the appearance of a yellow color (left side of image). However, DNA containing a C−C mismatch (right side of image) allows the Ag(I) ions to be sequestered from the enzyme, activating urease which produces urea and generates a red color. Inclusion of a pH sensitive dye in the urea solution allows the reaction to be monitored colorimetrically.

a solution of pullulan and transferred to paper using an automated liquid handler to create a system with urease/Ag(I) encapsulated in pullulan. The pullulan provides an oxygenimpermeable thin film and has been previously shown to provide long-term stability to entrapped biomolecules.49−53 To test urease activity on paper, a volume of 18 μL of substrate solution (urea with phenol red) was added and the paper was imaged after 20 min. In the case of the paper sensor, a substantially larger concentration of urease (2.2 μM) was required to obtain full activity (Figure S3a). The need for an almost 500-fold increase in enzyme concentration may be due to denaturation of a fraction of the enzyme upon printing and to the higher viscosity of the solution containing the dissolved pullulan, which would be expected to slow the diffusion-limited rate of the enzyme reaction.54 Given the higher urease concentration, it was necessary to increase the concentration of Ag(I) to 6.3 μM to completely inactivate the enzyme (Figure S3b). This 3:1 Ag(I):urease ratio again falls within the

Figure 2. Effect of dsDNA on the reactivation of urease in solution. TCAs contains no mismatches, TC1 contains a single C−C mismatch, and TC2 contains two C−C mismatches. Error bars represent the standard deviation from triplicate measurements.

the color of the solution with control DNA (TCAs, fully matched) showed low urease activity (10% activity at 140 nM DNA), confirming the fully matched duplex was unable to effectively reactivate urease. Conversely, both the TC1 (single mismatch duplex) and TC2 (double mismatch duplex) increased the activity of urease up to as high as 80% of the maximum value in a manner that was found to be dependent on the number of C−C mismatches, with TC1 having an EC50 C

DOI: 10.1021/acs.analchem.9b00235 Anal. Chem. XXXX, XXX, XXX−XXX

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Figure 4. Reactivation of urease on paper using the various dsDNA constructs. (A) Comparing sensitivity for homogeneous solutions of each dsDNA construct. Image above the graph shows the color of each well. (B) Sensitivity of a heterogeneous combination of TC1 and TCAs kept at a combined concentration of 1.7 μM. Error bars represent the standard deviation from triplicate measurements.

complete inhibition of urease was first determined using the same method as shown for the silver ions. The results showed that both copper and mercury had similar inhibitory effects on urease as silver, however zinc was much less effective at inhibiting urease (Figure S4a). Addition of C−C mismatched duplex DNA (TC1 at 1.7 μM) to each of the inactivated urease samples revealed that only the urease which was inactivated with silver ions could restore its activity, indicating that interaction between the C−C mismatch and silver ions is specific and thus that restoration of urease activity is proof of the presence of a C−C mismatch (Figure S4b). These results are consistent with previous reports.44,58 To investigate if the sequence or location of the C−C mismatch had an effect on the ability to reactivate urease, the parent strand was scrambled (termed “scrambled TCS” or “scTCS”) and hybridized on complementary scTC1 oligonucleotides that contained a single C−C mismatch at 4 separate sites (termed scTC1.A to scTC1.D). A scrambled TCAs was used as a control, with all sequences listed in Table S1 of the SI. Reactivation of urease by each of the five scrambled constructs was evaluated and demonstrated that the location of the mismatch had a significant effect on the ability to reactivate urease (Figure S5). As expected, all four scrambled sequences were able to reactive the inhibited urease to a much higher degree than the complementary scTCAs; however, the degree of reactivation varied by up to a factor of 2. The low degree of reactivation by scTC1.A is possibly due to the C−C mismatch being located on the terminus of the dsDNA construct, which may result in reduced hybridization in this region, thereby decreasing the ability for this sequence to effectively bind the Ag(I) ions. We also evaluated the effect of having multiple C−C mismatches by examining the reactivation of urease by a 20 nt polyC sequence. As shown in Figure S6, the polyC caused reactivation of urease at a level similar to the TC1 construct. It is expected that the polyC sequence can randomly form intraand/or intermolecular C−C mismatches that could sequester Ag(I) to increase the stability of these interactions and thereby restore the activity of urease. To demonstrate the analytical utility of the paper based urease/Ag(I) assay, the sensor was used to detect mutations in the tumor protein p53 gene TP53; the gene responsible for

range of 2:1 to 12:1 Ag(I):urease previously reported in the literature.33,47,48 Figure 4A shows colorimetric outputs from the paper sensor upon introduction of the control DNA and the sequences with one or two C−C mismatches. In this case, a volume of 10 μL of DNA sample was first added, incubated for 20 min, followed by addition of substrate solution and imaging after 20 min of color development. Once again, the mismatched duplexes reactivated the enzyme to produce color while the fully matched control duplex required almost 100 times more DNA to produce a comparable signal. TC1 had an EC50 of 602 ± 207 nM, compared to 195 ± 34 nM for TC2. At very high DNA concentrations, one can see that the TCAs eventually activates the enzyme, likely due to DNA nonspecifically abstracting the Ag(I) ions. Even so, it was possible to easily differentiate DNA samples with 0, 1, or 2 mismatches when using a DNA concentration in the range of 300−600 nM, and a single C−C mismatched duplex could be detected at a concentration of 34 nM, while the LOD of a double C−C mismatch was 17 nM of DNA, or 34 nM of total C−C mismatches. While the limit of detection is poorer than competing SNP detection platforms (typically in the fM to pM range),1 the paper assay does have the ability to easily differentiate between normal DNA and that with a C−C mismatch so long as an appropriate concentration of DNA is applied to the test and has the advantage of being inexpensive, simple to use, and amenable to reading by eye.55−57 To investigate the ability of the paper sensor to detect heterogeneous mixtures comprised of fully matched dsDNA and DNA containing a single C−C mismatch, samples containing varying ratios of TC1 and TCAs were tested. The total amount of DNA was kept at a concentration of 1.7 μM, as this value was shown to have high activity with TC1 compared to TCAs on the paper sensor (61 ± 4% activity and 14 ± 5% activity, respectively). The results (Figure 4B) show that the activity of urease was increased in samples containing as little as 10% TC1 (based on urease activity with 100% TCAs + 3SD). To show that the interaction of the C−C mismatch with Ag(I) ions is specific, three other metals were tested, including zinc-, copper-, and mercury-inactivated urease. For this purpose, the required concentration of each metal ion for D

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prior to performing the urease assay, and thus alleviating the need for a thermal cycler. Since paper-based POC diagnostic tests are designed to be used in the field or at the patient’s bedside, it is vital that paper based sensors remain stable under ambient storage conditions. The stability of the both the solution assay components and paper-based sensors was investigated over a period of 30 days with the devices stored at room temperature in ambient lighting conditions. In solution, the urease lost 99% activity after 3 days. However, the printed assay components lost only 2% activity after 3 days and retained greater than 60% activity after a month at room temperature and in ambient light (Figure S8). This data agrees with our previous results, which demonstrates the ability of enzymes to retain high activity after entrapment in pullulan films.49−51

encoding the p53 protein. Tumour protein p53 is a transcription factor which plays a significant role in suppressing tumor growth and development.59,60 Mutations in p53 are associated with approximately 50% of all human cancers, making it an important biomarker for biosensor development.61,62 Using a sample of purified human genomic DNA from a ductal carcinoma cell line, a 50-nt DNA region containing a G > C transversion mutation in the TP53 gene was PCR amplified. A reverse primer (mRP) was designed to be complementary to the mutation with the exception of a deoxycytidine at the transversion site. PCR amplification using this primer results in a portion of the PCR products containing a C−C mismatch. To mimic healthy genomic DNA, a separate PCR amplification was performed using the same carcinomic DNA, but with a separate reverse PCR primer (wtRP) that was fully complementary to the mutation site. Amplification in this case results in a fully complementary dsDNA with no mismatches, mimicking what a healthy cell line would produce (i.e., no C−C mismatch). We chose to use the same carcinomic DNA to produce both the p53m and p53wt data to ensure that the results were directly due to the PCR manipulations and not as the result of any background components/contaminants found in the samples. Figure S7 shows a schematic to illustrate the amplification strategy. Following amplification, a concentration of 600 nM of either wild-type or mutant p53 DNA was applied to the paper sensor, and the output of the paper sensor was determined, as shown in Figure 5. As expected, the system was able to detect a single

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CONCLUSIONS We have described a simple paper biosensor capable of detecting C−C mismatches by the naked eye in under 1 h. The paper assay format was not only faster than the microwell plate assay, but it required fewer pipetting steps and reduced reagent volumes. While we note that the limit of detection is poorer than some competing SNP detection platforms, the paperbased sensor can distinguish between DNA species with 0, 1, or 2 C−C mismatches so long as an appropriate concentration of DNA is used for the test. In addition, the paper-based assay provides relatively rapid turnaround times and is easy to use and inexpensive to produce. By designing gene-specific primers, the paper based system was capable of identifying a TP53 G > C transversion in genomic DNA from a human ductal carcinoma cell line. Hence, this simple paper-based sensor may find utility as a rapid means of assessing disease states that arise from C−C mismatch SNPs.

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ASSOCIATED CONTENT

* Supporting Information S

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.analchem.9b00235.

Figure 5. Detection of a C−C mutation in a p53 mutant gene sequence compared to the wild-type gene using the paper platform. Inset images show the colorimetric results of the paper assay. Error bars represent the standard deviation from triplicate measurements. *** = p < 0.001.

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List of the oligonucleotides used in this study; optimization of the urease/Ag(I) assay in solution and on paper; metal ion selectivity testing; PCR amplification strategy for genomic DNA from a ductal carcinoma cell line; biosensor stability data (PDF)

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. C−C mismatch (ρ = 0.001) in DNA from the cancerous cell line, with the results observable by the naked eye. In this case, the color change was smaller than that produced for the synthetic DNA (Figure 4), likely because only a small portion of the mutant PCR product contained the C−C mismatch. Even so, these results show that the system can detect heterogeneous mixtures containing C−C mismatches if appropriate PCR primers are used to amplify the gene of interest. Although our current version of the test required an offline DNA amplification step, it should be noted that ondevice isothermal amplification strategies such as strand displacement amplification or rolling circle amplification are possible,29,63 potentially allowing for on-device amplification

ORCID

John D. Brennan: 0000-0003-3461-9824 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors thank the Natural Sciences and Engineering Research Council of Canada for funding this work. We also thank the Canadian Foundation for Innovation and the Ontario Ministry of Research and Innovation for Infrastructure funding to the Biointerfaces Institute. J.D.B. holds the Canada Research Chair in Point of Care Diagnostics. E

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(36) Zaborska, W.; Krajewska, B.; Olech, Z. J. Enzyme Inhib. Med. Chem. 2004, 19, 65−69. (37) Du, N.; Chen, M.; Liu, Z.; Sheng, L.; Xu, H.; Chen, S. Chem. Cent. J. 2012, 6, 154. (38) Krajewska, B. J. Enzyme Inhib. Med. Chem. 2008, 23, 535−542. (39) Syshchyk, O.; Skryshevsky, V. A.; Soldatkin, O. O.; Soldatkin, A. P. Biosens. Bioelectron. 2015, 66, 89−94. (40) Do, J. S.; Lin, K. H. J. Taiwan Inst. Chem. Eng. 2016, 63, 25−32. (41) Kuswandi, B. Anal. Bioanal. Chem. 2003, 376, 1104−1110. (42) Tsai, H.-C.; Doong, R.-A.; Chiang, H.-C.; Chen, K.-T. Anal. Chim. Acta 2003, 481, 75−84. (43) Turdean, G. L. Int. J. Electrochem. 2011, 2011, 1−15. (44) Ono, A.; Cao, S.; Togashi, H.; Tashiro, M.; Fujimoto, T. Chem. Commun. 2008, 39, 4825−4827. (45) Tanaka, Y.; Kondo, J.; Sychrovský, V.; Š ebera, J.; Dairaku, T.; Saneyoshi, H.; Urata, H.; Torigoe, H.; Ono, A. Chem. Commun. 2015, 51, 17343−17360. (46) Dairaku, T.; Furuita, K.; Sato, H.; Š ebera, J.; Nakashima, K.; Ono, A.; Sychrovský, V.; Kojima, C.; Tanaka, Y. Inorg. Chim. Acta 2016, 452, 34−42. (47) Rezaei Behbehani, G.; Saboury, A. A.; Taherkhani, A.; Barzegar, L.; Mollaagazade, A. J. Therm. Anal. Calorim. 2011, 105, 1081−1086. (48) Mazzei, L.; Cianci, M.; Gonzalez Vara, A.; Ciurli, S. Dalt. Trans. 2018, 47, 8240−8247. (49) Jahanshahi-Anbuhi, S.; Pennings, K.; Leung, V.; Liu, M.; Carrasquilla, C.; Kannan, B.; Li, Y.; Pelton, R.; Brennan, J. D.; Filipe, C. D. M. Angew. Chem., Int. Ed. 2014, 53, 6155−6158. (50) Jahanshahi-Anbuhi, S.; Kannan, B.; Leung, V.; Pennings, K.; Liu, M.; Carrasquilla, C.; White, D.; Li, Y.; Pelton, R. H.; Brennan, J. D.; Filipe, C. D. M. Chem. Sci. 2016, 7, 2342−2347. (51) Kannan, B.; Jahanshahi-Anbuhi, S.; Pelton, R. H.; Li, Y.; Filipe, C. D. M.; Brennan, J. D. Anal. Chem. 2015, 87, 9288−9293. (52) Hsieh, P. Y.; Monsur Ali, M.; Tram, K.; Jahanshahi-Anbuhi, S.; Brown, C. L.; Brennan, J. D.; Filipe, C. D. M.; Li, Y. ChemBioChem 2017, 18, 502−505. (53) Ali, M. M.; Brown, C. L.; Jahanshahi-Anbuhi, S.; Kannan, B.; Li, Y.; Filipe, C. D. M.; Brennan, J. D. Sci. Rep. 2017, 7, 12335. (54) Berg, O. G.; von Hippel, P. Annu. Rev. Biophys. Biophys. Chem. 1985, 14, 131−160. (55) Hu, S.; Li, N.; Liu, F. Chem. Commun. 2018, 54, 3223−3226. (56) Wu, T.; Xiao, X.; Zhang, Z.; Zhao, M. Chem. Sci. 2015, 6, 1206−1211. (57) Deng, R.; Tang, L.; Tian, Q.; Wang, Y.; Lin, L.; Li, J. Angew. Chem., Int. Ed. 2014, 53, 2389−2393. (58) Torigoe, H.; Okamoto, I.; Dairaku, T.; Tanaka, Y.; Ono, A.; Kozasa, T. Biochimie 2012, 94, 2431−2440. (59) Vogelstein, B.; Lane, D.; Levine, A. J. Nature 2000, 408, 307− 310. (60) Vousden, K. H.; Lu, X. Nat. Rev. Cancer 2002, 2, 594−604. (61) Joerger, A.; Fersht, A. Oncogene 2007, 26, 2226−2242. (62) Olivier, M.; Hollstein, M.; Hainaut, P. Cold Spring Harbor Perspect. Biol. 2010, 2, 1−17. (63) Lafleur, L. K.; Bishop, J. D.; Heiniger, E. K.; Gallagher, R. P.; Wheeler, M. D. Lab Chip 2016, 16, 3777−3787.

REFERENCES

(1) Chang, K.; Deng, S.; Chen, M. Biosens. Bioelectron. 2015, 66, 297−307. (2) Auton, A.; Abecasis, G. R.; Schloss, J. A. Nature 2015, 526, 68− 74. (3) Bond, G. L.; Hu, W.; Bond, E. E.; Robins, H.; Lutzker, S. G. Cell 2004, 119, 591−602. (4) Zhao, X. Cancer Res. 2004, 64, 3060−3071. (5) Burton, P. R.; Clayton, D. G.; Cardon, L. R. Nature 2007, 447, 661−678. (6) Levin, B. R.; Perrot, V.; Walker, N. Genetics 2000, 154, 985−997. (7) Ramaswamy, S. V.; Reich, R.; Dou, S.-J.; Jasperse, L.; Pan, X.; Wanger, A.; Quitugua, T.; Graviss, E. A. Antimicrob. Agents Chemother. 2003, 47, 1241−1250. (8) Deulvot, C.; Charrel, H.; Marty, A. BMC Genomics 2010, 11, 468. (9) Knez, K.; Spasic, D.; Janssen, K. P. F.; Lammertyn, J. Analyst 2014, 139, 353−370. (10) Urdea, M.; Penny, L.a; Olmsted, S. S.; Giovanni, M. Y.; Kaspar, P. Nature 2006, 444, 73−79. (11) Turner, A. P. F. Chem. Soc. Rev. 2013, 42, 3184. (12) Yager, P.; Floriano, P. N.; Mcdevitt, J. IEEE Pulse 2011, 2, 40− 50. (13) Gubala, V.; Harris, L. F.; Ricco, A. J.; Tan, M. X.; Williams, D. E. Anal. Chem. 2012, 84, 487−515. (14) Pelton, R. TrAC, Trends Anal. Chem. 2009, 28, 925−942. (15) Yetisen, A. K.; Akram, M. S.; Lowe, C. R. Lab Chip 2013, 13, 2210−2251. (16) Martinez, A. W.; Phillips, S. T.; Whitesides, G. M.; Carrilho, E. Anal. Chem. 2010, 82, 3−10. (17) Li, X.; Ballerini, D. R.; Shen, W. Biomicrofluidics 2012, 6, 12− 14. (18) Posthuma-Trumpie, G. A.; Korf, J.; Van Amerongen, A. Anal. Bioanal. Chem. 2009, 393, 569−582. (19) Hu, J.; Wang, S.; Wang, L.; Li, F.; Pingguan-Murphy, B.; Lu, T. J.; Xu, F. Biosens. Bioelectron. 2014, 54, 585−597. (20) Parolo, C.; Merkoçi, A. Chem. Soc. Rev. 2013, 42, 450−457. (21) Nery, E. W.; Kubota, L. T. Anal. Bioanal. Chem. 2013, 405, 7573−7595. (22) Hossain, S. M. Z.; Luckham, R. E.; Smith, A. M.; Lebert, J. M.; Davies, L. M. Anal. Chem. 2009, 81, 5474−5483. (23) Hossain, S. M. Z.; Luckham, R. E.; McFadden, M. J.; Brennan, J. D. Anal. Chem. 2009, 81, 9055−9064. (24) Carrasquilla, C.; Little, J. R. L.; Li, Y.; Brennan, J. D. Chem. Eur. J. 2015, 21, 7369−7373. (25) Wolfe, M. G.; Zhang, Q.; Hui, C.; Radford, K.; Nair, P.; Brennan, J. D. Analyst 2016, 141, 6438−6443. (26) Hui, C. Y.; Liu, M.; Li, Y.; Brennan, J. D. Angew. Chem., Int. Ed. 2018, 57, 4549−4553. (27) Hossain, S. M. Z.; Ozimok, C.; Sicard, C.; Aguirre, S. D.; Ali, M. M.; Li, Y.; Brennan, J. D. Anal. Bioanal. Chem. 2012, 403, 1567− 1576. (28) Liu, M.; Zhang, Q.; Kannan, B.; Botton, G.; Yang, J.; Soleymani, L.; Brennan, J.; Li, Y. Angew. Chem. 2018, 130, 12620− 12623. (29) Liu, M.; Hui, C. Y.; Zhang, Q.; Gu, J.; Kannan, B.; JahanshahiAnbuhi, S.; Filipe, C. D. M.; Brennan, J. D.; Li, Y. Angew. Chem. 2016, 128, 2759−2763. (30) Zhu, X.; Li, G.; Chen, X.; Feng, C.; Chen, T.; Shi, H.; Mao, X.; Bo, B. Anal. Chem. 2017, 89, 6631−6636. (31) Yang, Y.; Li, C.; Shi, H.; Chen, T.; Wang, Z.; Li, G. Talanta 2019, 192, 325−330. (32) Tram, K.; Kanda, P.; Salena, B. J.; Huan, S.; Li, Y. Angew. Chem., Int. Ed. 2014, 53, 12799−12802. (33) Ambrose, E.; Kridl, A. G.; Kistiakowsky, G. B. J. Am. Chem. Soc. 1951, 73, 7−11. (34) Shaw, W. H. R. J. Am. Chem. Soc. 1954, 76, 2160−2163. (35) Shaw, W. H. R.; Raval, D. N. J. Am. Chem. Soc. 1961, 83, 3184− 3187. F

DOI: 10.1021/acs.analchem.9b00235 Anal. Chem. XXXX, XXX, XXX−XXX