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Development of an Advanced Electrochemical DNA Biosensor for Bacterial Pathogen Detection

2007; Elsevier BV; Volume: 9; Issue: 2 Linguagem: Inglês

10.2353/jmoldx.2007.060052

1943-7811

Joseph C. Liao, Mitra Mastali, Yang Li, Vincent Gau, Marc A. Suchard, Jane T. Babbitt, Jeffrey Gornbein, Elliot M. Landaw, Edward R.B. McCabe, Bernard M. Churchill, David A. Haake,

Electrochemical Analysis and Applications

Electrochemical sensors have the capacity for rapid and accurate detection of a wide variety of target molecules in biological fluids. We have developed an electrochemical sensor assay involving hybridization of bacterial 16S rRNA to fluorescein-modified detector probes and to biotin-modified capture probes anchored to the sensor surface. Signal is generated by an oxidation-reduction current produced by the action of horseradish peroxidase conjugated to an anti-fluorescein monoclonal Fab. A previous study found that this electrochemical sensor strategy could identify uropathogens in clinical urine specimens. To improve assay sensitivity, we examined the key steps that affect the current amplitude of the electrochemical signal. Efficient lysis and release of 16S rRNA from both gram-negative and -positive bacteria was achieved with an initial treatment with Triton X-100 and lysozyme followed by alkaline lysis, resulting in a 12-fold increase in electrochemical signal compared with alkaline lysis alone. The distance in nucleotides between the target hybridization sites of the detector and capture probes and the location of fluorescein modification on the detector probe contributed to a 23-fold change in signal intensity. These results demonstrate the importance of target-probe and probe-probe interactions in the detection of bacterial 16S rRNA using an electrochemical DNA sensor approach. Electrochemical sensors have the capacity for rapid and accurate detection of a wide variety of target molecules in biological fluids. We have developed an electrochemical sensor assay involving hybridization of bacterial 16S rRNA to fluorescein-modified detector probes and to biotin-modified capture probes anchored to the sensor surface. Signal is generated by an oxidation-reduction current produced by the action of horseradish peroxidase conjugated to an anti-fluorescein monoclonal Fab. A previous study found that this electrochemical sensor strategy could identify uropathogens in clinical urine specimens. To improve assay sensitivity, we examined the key steps that affect the current amplitude of the electrochemical signal. Efficient lysis and release of 16S rRNA from both gram-negative and -positive bacteria was achieved with an initial treatment with Triton X-100 and lysozyme followed by alkaline lysis, resulting in a 12-fold increase in electrochemical signal compared with alkaline lysis alone. The distance in nucleotides between the target hybridization sites of the detector and capture probes and the location of fluorescein modification on the detector probe contributed to a 23-fold change in signal intensity. These results demonstrate the importance of target-probe and probe-probe interactions in the detection of bacterial 16S rRNA using an electrochemical DNA sensor approach. Microfabrication technology has enabled the development of electrochemical DNA biosensors with the capacity for sensitive and sequence-specific detection of nucleic acids.1Drummond TG Hill MG Barton JK Electrochemical DNA sensors.Nat Biotechnol. 2003; 21: 1192-1199Crossref PubMed Scopus (1957) Google Scholar2Gau JJ Lan EH Dunn B Ho CM Woo JC A MEMS based amperometric detector for E. coli bacteria using self-assembled monolayers.Biosens Bioelectron. 2001; 16: 745-755Crossref PubMed Scopus (188) Google Scholar3Gooding JJ Electrochemical DNA hybridization biosensor.Electroanalysis. 2002; 14: 1149-1156Crossref Scopus (410) Google Scholar4Palecek E Jelen F Electrochemistry of nucleic acids and development of DNA sensors.Crit Rev Anal Chem. 2002; 3: 261-270Crossref Scopus (165) Google Scholar5Wang J Electrochemical nucleic acid biosensors.Anal Chim Acta. 2002; 469: 63-71Crossref Scopus (592) Google Scholar The ability of electrochemical sensors to directly identify nucleic acids in complex mixtures is a significant advantage over approaches such as polymerase chain reaction (PCR) that require target purification and amplification. Application of DNA sensor technology to infectious diseases has the potential for recognition of pathogen-specific signature sequences in biological fluids. The most frequent body fluid submitted to clinical microbiology laboratories is urine. Culture-based methods for isolation and identification of uropathogens are time consuming and labor intensive, contributing significantly to the estimated $3.5 billion annual cost of treating urinary tract infections in the United States.6Griebling TL Urologic diseases in America project: trends in resource use for urinary tract infections in men.J Urol. 2005; 173: 1288-1294Abstract Full Text Full Text PDF PubMed Scopus (138) Google Scholar,7Griebling TL Urologic diseases in America project: trends in resource use for urinary tract infections in women.J Urol. 2005; 173: 1281-1287Abstract Full Text Full Text PDF PubMed Scopus (134) Google Scholar A rapid, automated point-of-care system for identification of bacterial pathogens would have a significant impact on the clinical management of urinary tract infection (UTI) and infectious diseases in general. A general approach for species-specific identification of bacterial pathogens using an electrochemical sensor involves hybridization of single-stranded oligonucleotide capture and detector probes to target 16S rRNA.2Gau JJ Lan EH Dunn B Ho CM Woo JC A MEMS based amperometric detector for E. coli bacteria using self-assembled monolayers.Biosens Bioelectron. 2001; 16: 745-755Crossref PubMed Scopus (188) Google Scholar,8Gau V Ma SC Wang H Tsukuda J Kibler J Haake DA Electrochemical molecular analysis without nucleic acid amplification.Methods. 2005; 37: 73-83Crossref PubMed Scopus (105) Google Scholar The capture probe anchors the target to the sensor, whereas the detector probe signals the presence of the target through a reporter molecule (Figure 1). Binding of the capture and detector probes to the nucleic acid target creates a three-component "sandwich" complex on the sensor surface.9Campbell CN Gal D Cristler N Banditrat C Heller A Enzyme-amplified amperometric sandwich test for RNA and DNA.Anal Chem. 2002; 74: 158-162Crossref PubMed Scopus (129) Google Scholar10Dequaire M Heller A Screen printing of nucleic acid detecting carbon electrodes.Anal Chem. 2002; 74: 4370-4377Crossref PubMed Scopus (112) Google Scholar11Umek RM Lin SW Vielmetter J Terbrueggen RH Irvine B Yu CJ Kayyem JF Yowanto H Blackburn GF Farkas DH Chen YP Electronic detection of nucleic acids: a versatile platform for molecular diagnostics.J Mol Diagn. 2001; 3: 74-84Abstract Full Text Full Text PDF PubMed Scopus (267) Google Scholar12Williams E Pividori MI Merkoci A Forster RJ Alegret S Rapid electrochemical genosensor assay using a streptavidin carbon-polymer biocomposite electrode.Biosens Bioelectron. 2003; 19: 165-175Crossref PubMed Scopus (52) Google Scholar The fluorescein-modified detector probe enables binding of an antifluorescein-conjugated horseradish peroxidase reporter enzyme to the target-probe complex.8Gau V Ma SC Wang H Tsukuda J Kibler J Haake DA Electrochemical molecular analysis without nucleic acid amplification.Methods. 2005; 37: 73-83Crossref PubMed Scopus (105) Google Scholar The addition of a redox substrate and application of a fixed potential between the working and reference sensor electrodes creates a horseradish peroxidase-mediated redox cycle that is detected by the electrochemical sensor as current.13Fanjul-Bolado P Gonzalez-Garcia MB Costa-Garcia A Amperometric detection in TMB/HRP-based assays.Anal Bioanal Chem. 2005; 382: 297-302Crossref PubMed Scopus (104) Google Scholar,14Mecheri B Piras L Ciotti L Caminati G Electrode coating with ultrathin films containing electroactive molecules for biosensor applications.IEEE Sens J. 2004; 4: 171-179Crossref Scopus (11) Google Scholar In this way, the amplitude of the electroreduction current reflects the concentration of the target-probe complexes on the sensor surface. Initial proof-of-concept research has indicated that this electrochemical sensor approach has the potential for species-specific detection of uropathogens in clinical urine specimens.15Sun CP Liao JC Zhang YH Gau V Mastali M Babbitt JT Grundfest WS Churchill BM McCabe ER Haake DA Rapid, species-specific detection of uropathogen 16S rDNA and rRNA at ambient temperature by dot-blot hybridization and an electrochemical sensor array.Mol Genet Metab. 2005; 84: 90-99Crossref PubMed Scopus (42) Google Scholar,16Liao JC Mastali M Gau V Suchard MA Moller AK Bruckner DA Babbitt JT Li Y Gornbein J Landaw EM McCabe ER Churchill BM Haake DA Use of electrochemical DNA biosensors for rapid molecular identification of uropathogens in clinical urine specimens.J Clin Microbiol. 2006; 44: 561-570Crossref PubMed Scopus (188) Google Scholar However, the sensitivity of the detection system seems to be limited by inefficiencies in bacterial lysis and probe-target hybridization. In this study, we have conducted a detailed examination of the determinants of electrochemical signal intensity, namely 1) bacterial lysis and release of the 16S rRNA target molecules, 2) the length of the capture-detector probe complex, 3) the distance between the capture and detector probe hybridization regions on the 16S rRNA target, 4) the location of fluorescein on the detector probe, and 5) probe-probe and probe-target interactions during hybridization with mixtures of detector probes. Major improvements in signal intensity were achieved, contributing significantly toward our goal of developing a micro-fluidics-based "lab-on-a-chip" electrochemical sensor assay for bacterial pathogen detection. The following American Type Culture Collection (ATCC) strains were obtained from the University of California-Los Angeles (UCLA) Clinical Microbiology Laboratory: Escherichia coli strain 35218, Klebsiella pneumoniae strain 13883, Klebsiella oxytoca strain 49131, Enterobacter aerogenes strain 13048, Enterobacter cloacae strain 13047, Proteus mirabilis strain 12453, Pseudomonas aeruginosa strain 10145, Citrobacter freundii strain 8090, and Enterococcus faecalis strain 49532. Additional strains of uropathogenic bacteria were obtained from the UCLA Uropathogen Specimen Bank: E. coli strain Ec103, K. pneumoniae strain Kp295, P. mirabilis strain Pm278, P. aeruginosa strain Pa3, and E. faecalis strain Eo111. Isolation of uropathogens from clinical urine specimens was approved by the UCLA and VA Greater Los Angeles Healthcare System Institutional Review Boards. The identity of all clinical strains was determined by standard biochemical assays and verified by 16S rRNA gene sequencing. The 16S rRNA genes were PCR amplified with universal primers 8UA and 1485B.17Brosius J Palmer ML Kennedy PJ Noller HF Complete nucleotide sequence of a 16S ribosomal RNA gene from Escherichia coli.Proc Natl Acad Sci USA. 1978; 75: 4801-4805Crossref PubMed Scopus (2004) Google Scholar The amplified product was purified by using the QIAquick PCR purification kit (Qiagen, Inc., Chatsworth, CA) and directly sequenced using primer pairs 8UA/907B and 774A/1485B as described previously.18Summanen PH Durmaz B Vaisanen ML Liu C Molitoris D Eerola E Helander IM Finegold SM Porphyromonas somerae sp. nov., a pathogen isolated from humans and distinct from porphyromonas levii.J Clin Microbiol. 2005; 43: 4455-4459Crossref PubMed Scopus (40) Google Scholar DNA sequencing was performed at the W.M. Keck Foundation Biotechnology Resource Laboratory (New Haven, CT). Isolates were inoculated into Brucella broth with 15% glycerol (BBL, Annapolis, MD) and were stored at −70°C. All experiments reported here involved bacteria grown overnight in Luria broth, inoculated into Luria broth, and grown to logarithmic phase as measured by OD600. Concentration of the logarithmic phase specimens was determined by serial plating, typically yielding 107 to 108 bacteria/ml. Oligonucleotide probes were synthesized by MWG Biotech (High Point, NC). Capture probes were synthesized with a 5′ biotin modification. Detector probes were synthesized with 5′- and/or 3′-fluorescein modifications. Oligonucleotide probe pairs were designed to hybridize with species-specific regions of the 16S rRNA molecules of E. coli, E. faecalis, P. mirabilis, and P. aeruginosa. Oligonucleotides were also designed as capture and detector probes for the Klebsiella-Enterobacter group, the family Enterobacteriaceae, and as universal bacterial probes. Probe pairs were studied with and without a gap between the hybridization regions on the 16S rRNA target. The sequences of all oligonucleotide probes used in this study are shown in Table 1.Table 1Sequences of Oligonucleotide Probes Used in This WorkProbe designation*Capture probes were 5′-modified with biotin. Detector probes were 5′- and/or 3′-modified with fluorescein. Probe sequence numbering based on E. coli 16S rDNA. EB, Enterobacteriaceae family; EC, Escherichia coli; EF, Enterococcus species; KE, Klebsiella-Enterobacter group; PA, Pseudomonas aeruginosa; PM, Proteus mirabilis; UNI, universal bacterial.Sequence (5′-3′)Capture probes EB1172C (35-mer)5′-CGGACTACGACRYACTTTATGAGGTCCGCTTGCTC-3′ EC434C (35-mer)5′-GTCAATGAGCAAAGGTATTAACTTTACTCCCTTCC-3′ EC430C (35-mer)5′-GAGCAAAGGTATTAACTTTACTCCCTTCCTCCCCG-3′ EF207C (35-mer)5′-TTGGTGAGCCGTTACCTCACCAACTAGCTAATGCA-3′ EF165C (35-mer)5′-GTCCATCCATCAGCGACACCCGAAAGCGCCTTTCA-3′ KE434C (35-mer)5′-GTCAATCGMCRAGGTTATTAACCTYAHCGCCTTCC-3′ PA111C (35-mer)5′-CCCACTTTCTCCCTCAGGACGTATGCGGTATTAGC-3′ PA972C (35-mer)5′-TGAGTTCCCGAAGGCACCAATCCATCTCTGGAAAG-3′ PM188C (35-mer)5′-GGGTTCATCCGATAGTGCAAGGTCCGAAGAGCCCC-3′ UNI782C (27-mer)5′-CATCGTTTACGGCGTGGACTACCAGGG-3′Detector probes EB1137D (35-mer)5′-GAGGTCGCTTCTCTTTGTATRYGCCATTGTAGCAC-3′ EB1141D (35-mer)5′-TCGCGAGGTCGCTTCTCTTTGTATRYGCCATTGTA-3′ EC393D (35-mer)5′-CTGAAAGTACTTTACAACCCGAAGGCCTTCTTCAT-3′ EC394D (35-mer)5′-GCTGAAAGTACTTTACAACCCGAAGGCCTTCTTCA-3′ EC395D (35-mer)5′-CGCTGAAAGTACTTTACAACCCGAAGGCCTTCTTC-3′ EC396D (35-mer)5′-CCGCTGAAAGTACTTTACAACCCGAAGGCCTTCTT-3′ EC397D (35-mer)5′-CCCGCTGAAAGTACTTTACAACCCGAAGGCCTTCT-3′ EC398D (35-mer)5′-CCCCGCTGAAAGTACTTTACAACCCGAAGGCCTTC-3′ EC399D (35-mer)5′-TCCCCGCTGAAAGTACTTTACAACCCGAAGGCCTT-3′ EF165D (35-mer)5′-GTCCATCCATCAGCGACACCCGAAAGCGCCTTTCA-3′ EF171D (35-mer)5′-CCGCGGGTCCATCCATCAGCGACACCCGAAAGCGC-3′ EF207D (35-mer)5′-TTGGTGAGCCGTTACCTCACCAACTAGCTAATGCA-3′ KE393D (35-mer)5′-CTGAAAGTGCTTTACAACCCGAAGGCCTTCTTCAT-3′ KE399D (35-mer)5′-TCCCCGCTGAAAGTGCTTTACAACCCGAAGGCCTT-3′ PA68D (35-mer)5′-TTCCGGACGTTATCCCCCACTACCAGGCAGATTCC-3′ PA74D (35-mer)5′-GCCCGTTTCCGGACGTTATCCCCCACTACCAGGCA-3′ PA932D (35-mer)5′-CAGCATGTCAAGGCCAGGTAAGGTTCTTCGCGTTG-3′ PM147D (35-mer)5′-GGTCCGTAGACATTATGCGGTATTAGCCACCGTTT-3′ PM153D (35-mer)5′-TGCTTTGGTCCGTAGACATTATGCGGTATTAGCCA-3′UNI751D (31-mer)5′-TATCTAATCCTGTTTGCTCCCCACGCTTTCG-3′* Capture probes were 5′-modified with biotin. Detector probes were 5′- and/or 3′-modified with fluorescein. Probe sequence numbering based on E. coli 16S rDNA. EB, Enterobacteriaceae family; EC, Escherichia coli; EF, Enterococcus species; KE, Klebsiella-Enterobacter group; PA, Pseudomonas aeruginosa; PM, Proteus mirabilis; UNI, universal bacterial. Open table in a new tab Microfabricated electrochemical sensor arrays with an alkanethiolate self-assembled monolayer were obtained from GeneFluidics (Monterey Park, CA). Self-assembled monolayer integrity was confirmed by cyclic voltammetry19Bard AJ Faulkner LR Potential Sweep Methods. John Wiley & Sons, Hoboken NJ2001: 226-260Google Scholar using a 16-channel potentiostat (GeneFluidics). After cyclic voltammetry characterization, sensor arrays were washed and dried. Washing steps were performed by applying a stream of deionized H2O to the sensor surface for approximately 2 to 3 seconds followed by 5 seconds of drying under a stream of nitrogen. To functionalize the sensor surface, 4 μl of 0.5 mg/ml streptavidin (Calbiochem, San Diego, CA) in H2O was added to the alkanethiol-activated sensors, incubated for 10 minutes at room temperature, and washed off. Biotinylated capture probes (4 μl, 1 μmol/L in 1 mol/L phosphate buffer, pH 7.4) were added to the streptavidin-coated sensors. Phosphate buffer (1 mol/L), pH 7.4, was prepared by mixing 1 mol/L NaH2PO4 and 1 mol/L K2HPO4 in a 19:81 (v/v) ratio, respectively, and adjusting the pH to 7.4. After 30 minutes of incubation at room temperature, the sensor array was washed and dried, completing the surface preparation. Logarithmic phase bacterial cells were concentrated by centrifugation at 10,000 rpm for 5 minutes. Lysis of bacterial cells was performed by the addition of 10 μl of one or more of the following: 1 mol/L NaOH, 0.1% Triton X-100 in 20 mmol/L Tris-HCl, pH 8.0, 2 mmol/L ethylenediamine tetraacetic acid, and 1 mg/ml lysozyme (Sigma, St. Louis, MO). After incubation at room temperature, 50 μl of the detector probe (0.25 μmol/L) in 2.5% bovine serum albumin (Sigma) and 1 mol/L phosphate buffer, pH 7.4, was added to the bacterial lysate. The detector probe/bacterial lysate mixture was incubated for 10 minutes at 65°C to allow hybridization of the detector probe to target rRNA. Four microliters of the bacterial lysate/detector probe mixture was deposited on each of the working electrodes in the sensor array. The sensor array was incubated for 15 minutes at 65°C in a humidified chamber. After washing and drying, 4 μl of 0.5 U/ml anti-fluorescein horseradish peroxidase (HRP) Fab conjugate (diluted in 0.5% casein in 1 mol/L phosphate buffer, pH 7.4; Roche Diagnostics, Mannheim, Germany) were deposited on each of the working electrodes for 10 to 15 minutes. After washing and drying, a prefabricated plastic well manifold (GeneFluidics) was bonded to the sensor array. HRP substrate solution (50 μl of K-Blue Aqueous TMB; Neogen, Lexington, KY) was placed on each of the sensors in the array to cover all three of the electrodes. Measurements were immediately and simultaneously taken for all 16 sensors. The entire assay protocol was completed within 45 minutes from the initiation of bacterial lysis. Amperometric current versus time was measured using a multichannel potentiostat (GeneFluidics). The voltage was fixed at −200 mV (versus reference), and the electroreduction current was measured at 60 seconds after the HRP redox reaction reached steady state. Negative controls were included in each experiment in which 1 mol/L phosphate buffer, pH 7.4, was used as the target instead of bacterial lysate. All samples were analyzed in duplicate. Experiments were performed on ATCC strains to verify probe specificity using a 16-sensor array "UTI chip" in which the UNI782C, EB1176C, EC434C, KE434C, PM187C, PA102C, and EF207C 5′-biotinylated capture probes (defined in Table 1) were tested in duplicate. The two remaining sensors in the array served as negative controls (using capture probe UNI782C in 1 mol/L phosphate buffer, pH 7.4, instead of bacterial lysate). Bacterial lysates were combined with a mixture of the following 3′-fluorescein-labeled detector probes: UNI751D, EB1141D, EC399D, KE399D, PM153D, PA74D, and EF171D (defined in Table 1). The degree of variance in the electrochemical sensor measurements was determined by comparing duplicate measurements for all experiments. The background signal level was determined by averaging the log10 results of the two negative control sensors. Positive signals were defined as signals greater than 5 SD (in log10 units) over background. An effective 16S rRNA detection system for uropathogens requires efficient lysis of gram-negative and gram-positive organisms. Previous work had shown that alkaline lysis efficiently released 16S rRNA from gram-negative but not gram-positive bacteria.16Liao JC Mastali M Gau V Suchard MA Moller AK Bruckner DA Babbitt JT Li Y Gornbein J Landaw EM McCabe ER Churchill BM Haake DA Use of electrochemical DNA biosensors for rapid molecular identification of uropathogens in clinical urine specimens.J Clin Microbiol. 2006; 44: 561-570Crossref PubMed Scopus (188) Google Scholar For this reason, we examined a variety of methods for rapid release of 16S rRNA from the gram-positive uropathogen, E. faecalis. Given the thicker cell wall of gram-positive organisms, we considered whether membrane-active detergents and/or peptidoglycan-specific enzymes would be useful components of an effective lysis strategy. As shown in Figure 2, NaOH with or without the detergent Triton X-100 did not lyse Enterococcus cells sufficiently for electrochemical detection of 16S rRNA above negative control signal levels. However, the combination of 0.1% Triton X-100 plus 1 mg/ml lysozyme resulted in a threefold increase in current output over negative control. The approach that yielded the highest electrochemical signal intensity was a strategy in which Enterococcus cells were initially treated with the combination of Triton X-100 and lysozyme for 5 minutes followed by treatment with NaOH for an additional 5 minutes. This two-step lysis method resulted in a 12-fold increase in electrochemical signal compared with alkaline lysis alone. The sequence in which these treatments were applied was important. Alkaline lysis before application of Triton X-100 and lysozyme was not as effective as treatment with NaOH after the detergent-enzyme combination. Longer periods of lysis did not further enhance signal intensity. Lysis of gram-negative uropathogens (eg, E. coli, P. mirabilis, K. pneumoniae, and P. aeruginosa) with Triton X-100 and lysozyme or Triton X-100 and lysozyme followed by NaOH resulted in successful electrochemical detection of 16S rRNA, although the results were not significantly better than lysis with NaOH alone (data not shown). Therefore, this two-step process can be considered a universal lysis strategy for release of bacterial 16S rRNA and was used for the remainder of the experiments described here involving E. faecalis. For the sake of simplicity, subsequent experiments involving gram-negative organisms involved lysis with NaOH alone. Use of various concentrations of the denaturing detergent sodium dodecyl sulfate coupled with nonspecific proteases (eg, proteinase K or Pronase) did not further improve electrochemical signal strength (data not shown). The effect of distance between the capture and detector probe hybridization sites on the 16S rRNA target was examined. Experiments involving a variety of 16S rRNA targets using capture and detector probes with hybridization sites separated by a gap of >300 nucleotides produced no significant electrochemical current output, even though these probes were known to function well as members of juxtaposed probe pairs (data not shown). In contrast, capture and detector probes with hybridization sites separated by relatively short interprobe gaps of up to six nucleotides yielded positive signals. As shown in Figure 3, there was a negative correlation (Pearson product-moment correlation coefficient r = −0.84) between signal intensity and the size of the gap between the capture and detector probe hybridization sites, even for very short interprobe gaps. Maximal signal intensity required eliminating the interprobe hybridization site gap between probe pairs specific for the Klebsiella-Enterobacter group, P. aeruginosa, and P. mirabilis (Table 2), which bind to various regions of the 16S rRNA target. These results indicate that the effect of an interprobe gap on signal intensity is independent of the 16S rRNA target hybridization site and species of origin.Table 2Effect of a Gap between the Capture and Detector Probe Hybridization SitesTarget*Ec, E. coli; Ef, E. faecalis; Kp, K. pneumoniae; Pa, P. aeruginosa; Pm, P. mirabilis.Probe pair (6-nucleotide gap)nA ± SDProbe pair (0-nucleotide gap)nA ± SDFold change (0:6 nucleotide)EcEC 434C/393D†3′-Fluorescein-modified detector probe.590 ± 65EC 434C/399D†3′-Fluorescein-modified detector probe.3242 ± 855.49‡P < 0.001.EfEF 207C/165D†3′-Fluorescein-modified detector probe.1202 ± 85EF 207C/171D†3′-Fluorescein-modified detector probe.1777 ± 571.48KpKP 434C/393D†3′-Fluorescein-modified detector probe.2106 ± 186KE 434C/399D†3′-Fluorescein-modified detector probe.7789 ± 933.70‡P < 0.001.KpKP 972C/932D§5′-Fluorescein-modified detector probe.1438 ± 151KE 972C/938D§5′-Fluorescein-modified detector probe.2975 ± 6102.07¶P < 0.05.PaPA 102C/68D†3′-Fluorescein-modified detector probe.2483 ± 85PA 102C/74D†3′-Fluorescein-modified detector probe.5455 ± 352.20|P < 0.01.PmPM 187C/147D†3′-Fluorescein-modified detector probe.908 ± 19PM 187C/153D†3′-Fluorescein-modified detector probe.3361 ± 2483.70‡P < 0.001.* Ec, E. coli; Ef, E. faecalis; Kp, K. pneumoniae; Pa, P. aeruginosa; Pm, P. mirabilis.† 3′-Fluorescein-modified detector probe.‡ P < 0.001.§ 5′-Fluorescein-modified detector probe.¶ P < 0.05.| P < 0.01. Open table in a new tab We considered whether the location of the fluorescein (the binding site for the anti-fluorescein Fab-HRP conjugate) on the detector probe affected signal intensity. To examine this question, we compared the signal intensity produced using 3′- versus 5′-fluorescein-modified detector probes. Use of 3′-fluorescein-modified detector probes resulted in greater signal intensity than 5′-fluorescein-modified detector probes for detection of both E. faecalis (Figure 4A) and E. coli (Figure 4B) 16S rRNA. As shown in Figure 4, there was an additive effect of removing the interprobe gap between the capture and detector probe hybridization sites and moving the fluorescein modification from the 5′ to the 3′ end of the detector probe. Experiments using fivefold dilutions of enterococcal and E. coli cells showed that the probe pair with a 0-nucleotide gap and a 3′-fluorescein-modified detector probe had a 23- to 29-fold lower limit of detection sensitivity compared with the probe pair with a 6-nucleotide gap and a 5′-fluorescein-modified detector probe (Figure 5).Figure 5Sensitivity of the electrochemical sensor assay as a function of locations of probe hybridization and fluorescein modification. A: Electrochemical sensor results from fivefold serial dilutions of enterococcal cells using capture probe EF207C paired with detector probes EF165D or EF171D modified by fluorescein at the 5′ and 3′ positions, respectively. B: Electrochemical sensor results from fivefold serial dilutions of E. coli cells using capture probe EC434C paired with detector probes EC393C or EC399C modified by fluorescein at the 5′ and 3′ positions, respectively. The dashed horizontal lines indicate current output thresholds for duplicate results significantly greater than negative control (P < 0.01). 3′-Fluorescein modification of the detector probe combined with continuity between the detector and capture probe hybridization sites resulted in a 23-fold improvement in sensitivity of electrochemical sensor assay for detection of enterococci (from 232,000 down to 9900 cells) and E. coli (from 6500 down to 280 cells).View Large Image Figure ViewerDownload Hi-res image Download (PPT) As shown in Table 3, we examined whether the effect of the location of fluorescein modification could be generalized for a variety of detector probes and targets. Some of the detector probes (UNI751D, EB1137D, and EC399D) were also modified with fluorescein at both the 5′ and 3′ positions. Significant increases in electrochemical current output were achieved using fluorescein modification at the 3′ position compared with the 5′ position for all of the species-specific detector probes. Interestingly, the location of fluorescein modification had no effect on signal intensity in the case of the universal (UNI751D) bacterial detector probe. In addition, fluorescein labeling of the detector probe at both the 3′ and 5′ positions did not enhance signal strength beyond that achieved with 3′ modification alone.Table 3Effect of Location of Detector Probe Fluorescein ModificationTarget*Ec, Escherichia coli; Ef, Enterococcus faecalis; Pa, Pseudomonas aeruginosa; Pm, Proteus mirabilis.Probe pair5′ Fluorescein (nA ± SD)3′ Fluorescein (nA ± SD)Fold change (3′:5′)EfEF 207C/165D†Six-nucleotide gap between the capture and detector probe hybridization sites.236 ± 591202 ± 855.10‡P < 0.001.EfEF 207C/171D§Zero-nucleotide gap between the capture and detector probe hybridization sites.1093 ± 321777 ± 571.63EcEC 434C/399D§Zero-nucleotide gap between the capture and detector probe hybridization sites.2781 ± 1447313 ± 3362.63¶P < 0.01.EcEC 434C/393D†Six-nucleotide gap between the capture and detector probe hybridization sites.2993 ± 6744288 ± 6031.43EcEC 430C/393D§Zero-nucleotide gap between the capture and detector probe hybridization sites.5650 ± 2706816 ± 8581.21EcEB 1176C/1137D|Four-nucleotide gap between the capture and detector probe hybridization sites.1122 ± 352528 ± 1442.25¶P < 0.01.EcUNI 782C/751D§Zero-nucleotide gap between the capture and detector probe hybridization sites.1890 ± 4912053 ± 1891.09PmPM 188C/147D†Six-nucleotide gap between the capture and detector probe hybridization sites.420 ± 58806 ± 451.92**P < 0.05.PmUNI 782C/751D§Zero-nucleotide gap between the capture and detector probe hybridization sites.1388 ± 101502 ± 331.08PaPA 111C/68D†Six-nucleotide gap between the capture and detector probe hybridization sites.668 ± 731295 ± 2381.94**P < 0.05.PaPA 972C/932D†Six-nucleotide gap between the capture and detector probe hybridization sites.393 ± 351314 ± 1153.34‡P < 0.001.* Ec, Escherichia coli; Ef, Enterococcus faecalis; Pa, Pseudomonas aeruginosa; Pm, Proteus mirabilis.† Six-nucleotide gap between the capture and detector probe hybridization sites.‡ P < 0.001.§ Zero-nucleotide gap between the capture and detector probe hybridization sites.¶ P < 0.01.| Four-nucleotide gap between the capture and detector probe hybridizati