An RNA-Aptamer-Based Assay for the Detection ... - ACS Publications

Mar 4, 2010 - Food and Environment Research Agency, Sand Hutton, York YO41 1LZ North Yorkshire, United Kingdom,. Somagenics, Inc., 2161 Delaware ...
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Anal. Chem. 2010, 82, 2652–2660

An RNA-Aptamer-Based Assay for the Detection and Analysis of Malachite Green and Leucomalachite Green Residues in Fish Tissue Sara L. Stead,*,† Helen Ashwin,† Brian H. Johnston,‡,§ Anne Dallas,‡ Sergei A. Kazakov,‡ Jonathan A. Tarbin,† Matthew Sharman,† Jack Kay,∇ and Brendan J. Keely⊥ Food and Environment Research Agency, Sand Hutton, York YO41 1LZ North Yorkshire, United Kingdom, Somagenics, Inc., 2161 Delaware Avenue, Santa Cruz, California 95060, Department of Pediatrics, Stanford University School of Medicine, Stanford, California 94305, Veterinary Medicines Directorate, Woodham Lane, New Haw, Addlestone, Surrey KT15 3LS, United Kingdom, and Department of Chemistry, University of York, Heslington Lane, York YO10 5DD, United Kingdom A robust screening assay employing solid phase extraction (SPE) followed by a novel aptamer-based procedure is presented for the rapid detection and semiquantitation of the triphenylmethane dye, Malachite Green (MG) and its primary metabolite Leucomalachite Green (LMG) in fish tissue. To the authors’ knowledge, this is the first reported use of an RNA aptamer for the development of a diagnostic assay for the detection of chemical residues in food. The aptamer based screening assay is found to be highly specific for MG; but has negligible affinity for the LMG metabolite. However, because the LMG metabolite is lipophilic and known to be highly persistent in tissues, an oxidation step has been incorporated within the sample cleanup procedure to ensure that all LMG residues are converted to MG prior to measurement. This article provides evidence that an oligonucleotide aptamer can be used as an alternative recognition element to conventional antibodies with application to the detection of residues in food. Furthermore, this finding has the future potential to reduce the number of animals currently being used in the production of antibodies for immunodiagnostic kits. Malachite Green (MG) (depicted in Figure 1) is a cationic triphenylmethane dye that has been used in commercial aquaculture since 1936.1 MG is especially active against the Saprolegnia fungus, which infects fish and fish eggs. It is also a very effective topical antiparasitic and antiprotozoan agent used in the treatment of farmed freshwater fish. However, there has been much concern about the use of MG in aquaculture, because of its potential * Author to whom correspondence should be addressed. Fax: +44 (0)1904 462111. E-mail: [email protected]. † Food and Environment Research Agency. ‡ Somagenics, Inc. § Department of Pediatrics, Stanford University School of Medicine. ∇ Veterinary Medicines Directorate. ⊥ Department of Chemistry, University of York. (1) Foster, F. J.; Woodbury, L. Prog. Fish-Cult. 1936, 3, 7–9. (2) Joint Committee on Mutagenicity and Committee on Carcinogenicity Statement on Mutagenicity and Carcinogenicity of Malachite Green (MG) and Leucomalachite Green (LMG). Available via the Internet at http:// www.iacom.org.uk/statements/COM04S4.htm, accessed August 3,2009.

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Figure 1. Chemical structures of Malachite Green (left) and Leucomalachite Green (right).

toxicity.2 Despite this, surveillance programs have identified a continuing incidence of MG and its main metabolitesLeucomalachite Green (LMG) (also depicted in Figure 1)sresidues in farmed fish samples. The leuco-form is more nonpolar (lipophilic) than the parent compound. Studies have shown that the LMG metabolite is predominant to MG in vivo, and highly persistent in edible fish tissue.3-5 Between 2003 and 2007, there were ca. 700 notifications under the EU Rapid Alert System for Food and Feed (RASFF) system, which involves veterinary medicines in foods of animal origin, of which 104 are related to the illegal use of MG.6 The EU has established a minimum required performance limit (MRPL) of 2 µg kg-1 for the sum of MG and LMG.7 Screening and confirmatory methods must be capable of reliably detecting LMG/MG at concentrations at or below 2 µg kg-1. Current laboratory procedures for the detection and quantification of MG/LMG are relatively complex, time-consuming, and tend to be reliant upon the use of instrumental analysis. Because of (3) Azmi, W.; Saini, R. K.; Banerjee, U. C. Enzyme Microb. Technol. 1998, 22, 185–191. (4) Plakas S. M., Doerge D. R. Turnipseed S. B. Disposition and metabolism of malachite green and other therapeutic dyes in fish. In Xenobiotics in Fish; Smith, D. J., Gingerich, W. H., Beconi-Barker, M. G., Eds.; Kluwer Academic Publishers/Plenum Publishers: New York, 1999. (5) Pointing, S. B.; Vrijmoed, L. L. P. World J. Microbiol. Biotechnol. 2000, 16, 317–318. (6) EC, Rapid Alert System of Food and Feed. Available via the Internet at http://ec.europa.eu/food/food/rapidalert/archive_en.htm, accessed August 18, 2009. (7) Commission Decision of 22 December 2003 amending Decision 2002/657/ EC as regards the setting of minimum required performance limits (MRPLs) for certain residues in food of animal origin (notified under document number C(2003) 4961), 2004/26/EC. Off. J. Eur. Union, L: Legis. (Engl. Ed.) 2004, L6, 38-39. 10.1021/ac902226v  2010 American Chemical Society Published on Web 03/04/2010

the strong absorption of MG in the visible range of the UV spectrum, HPLC with visible (VIS) detection has traditionally been used.8 More recently, LC-MS applications have been developed for the quantitative analysis of MG/LMG with detection limits below the EU MRPL of 2 µg kg-1.9-12 In the case of rapid screening methods, there has been limited progress, although enzyme-linked immunosorbent assay (ELISA)-based methods have also been reported.13 This paper describes the production and characterization of an RNA aptamer for application as an alternative approach to the rapid detection and screening for MG/ LMG residues in fish tissue. Aptamers are short, single-stranded oligonucleotide sequences derived from either DNA or RNA, generated against specific targets. These three-dimensional (3D) shapes possess specific structural, ligand-binding, and catalytic properties.14 Aptamers are rapidly generated in vitro using a technique called the systematic evolution of ligands by exponential enrichment (SELEX), thus removing the need for the use of animals in the production of recognition elements. The potential for the application of aptamers in residue diagnostics is considerable. A key challenge to their successful implementation is the transformation of the binding event into physically detectable signals.15 An RNA aptamer (38-mer consensus sequence) for MG has been reported previously.16 The MG aptamer was originally developed as an alternative to the chromophore-assisted laser inactivation (CALI) technique used in molecular biology to study the functions of specific genes.17 Binding of the aptamer with the cognate ligand was determined to result in an increase in the fluorescence of MG (by a factor of >2000), and, hence, an intrinsic detection mechanism. MG has a low quantum yield (Φ) for fluorescence, because of easy vibrational de-excitation. Binding of the aptamer affects the electronic structure and conformation of MG molecules, as a result of two complementary interactions, electrostatic forces, and base stacking. Once bound, the MG aromatic rings adopt a more coplanar, rotational stabilized structure, thus leading to an extended π-system, which is the likely origin of the observed red shift in its maximum absorption frequency. The three-dimensional (3D) structure of the MG aptamer (RNA) has been elucidated using one-dimensional (1D) 1H NMR for both the free and bound forms18 in aqueous solution. The ligand-binding site is defined by an asymmetric internal stemloop, which is flanked by a pair of helices. The RNA aptamer utilizes several tiers of stacked nucleotides arranged in pairs and triples, and a base quadruple to encapsulate the ligand.16 Because of the highly structured nature of the ligand-binding pocket, the aptamer discriminates between closely related (8) Tarbin, J. A.; Barnes, K. A.; Bygrave, J.; Farrington, W. H. H. Analyst 1998, 123, 2567–2571. (9) Allen, J. L.; Gofus, J. E.; Meinertz, J. R. J. AOAC Int. 1994, 77, 553–557. (10) Halme, K.; Lindfors, E.; Peltonen, K. J. Chromatogr. B 2006, 845, 74–79. (11) Dowling, G.; Mulder, P. P. J.; Duffy, C.; Regan, L.; Smyth, M. R. Anal. Chim. Acta 2006, 586, 411–419. (12) Lee, K. C.; Wu, J. L.; Cai, Z. J. Chromatogr. B 2006, 843, 247–251. (13) Yang, M. C.; Fang, J. M.; Kuo, T. F.; Wang, D. M.; Huang, Y. L.; Liu, L. Y.; Chen, P. H.; Chang, T. H. J. Agric. Food Chem. 2007, 55 (22), 8851–8856. (14) Huang, Z.; Szostak, J. W. RNA 2003, 9, 1456–1463. (15) Liu, J.; Lu, Y. Angew. Chem., Int. Ed. 2006, 118, 96–100. (16) Grate, D.; Wilson, C. Proc. Natl. Acad. Sci., U.S.A. 1999, 96, 6131–6136. (17) Jay, D. G.; Keshishian, H. Nature 1990, 348, 548–550. (18) Flinders, J.; DeFina, S. C.; Brackett, D. M.; Baugh, C.; Wilson, C.; Dieckmann, T. ChemBioChem 2004, 5, 62–72.

structural homologues of MG. Access to the binding pocket is restricted based on the conformation, charge, and size of the ligand. The characterization of an RNA MG aptamer and the subsequent development of a novel assay applicable for the rapid detection of MG/LMG residues in fish tissue is presented in this paper. EXPERIMENTAL SECTION Equipment. The fluorescence spectrophotometry was performed using a Model FP650 fluorescence spectrophotometer (Jasco, Essex, UK). Materials. The solventssacetonitrile, acetone, cyclohexane, dichloromethane, hexane, issoctane, methanol, pentane, and water (all HPLC-grade), and sodium acetateswere obtained from Fisher Scientific (Loughborough, U.K.). Glacial acetic acid, 2,3-dichloro5,6-dicyano-1,4-benzoquinone, ethylenediaminetetraacetic acid, 4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid, hydrochloric acid, magnesium chloride, potassium chloride, sodium chloride, sodium hydroxide, and sodium sulfate (all reagent-grade) were obtained from Sigma Aldrich (Dorset, U.K.). Phosphate buffered saline tablets were purchased from Calbiochem (San Diego, CA). Leucomalachite Green, Malachite Green oxalate, d5-Malachite Green picrate of the highest purity grade available were all obtained from Sigma Aldrich (Dorset, U.K.). Sodium dodecyl sulfate (10% solution) was obtained from Severn Biotech Ltd. (Worcestershire, U.K.). Tween 20 (polysorbate 20) was obtained from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). Molecular-grade water was obtained from HyPure (Fisher Scientific, Loughborough, U.K.). C8, C18, SCX (500 mg/6 mL) SPE cartridges were purchased from Varian (Palo Alto, CA). OASIS MCX and HLB (500 mg/6 mL) SPE cartridges were obtained from Waters (Milford, MA). Strata X SPE cartridges (60 mg/3 cc and 200 mg/6 cc) were obtained from Phenomenex (Macclesfield, U.K.). Preparation of the MG Aptamer. The RNA malachite green aptamer was synthesized according to the procedure described by Babendure et al.19 The 38-oligomer sequence (primary structure) of the RNA is given below, in the 5′ to 3′ direction: 5'-GGAUCCCGACUGGCGAGAGCCAGGUAACGAAUGGAUCC-3' The analysis of sodium dodecyl sulfate polyacrylamide gelelectrophoresis (SDS-PAGE) (Figure 2) shows that the transcription yielded RNAs with a length of 38-39 nucleotides, largely free of other species other than the N and N + 1 products. The two RNA products were pooled and used for the subsequent assay development. Preparation of HEPES-Based Assay Buffer. Potassium chloride (7.456 g), magnesium chloride (0.476 g), EDTA 0.1 mM (0.0292 g), 4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES) (2.381 g), and sodium dodecyl sulfate (SDS, 0.01%) were dissolved in 700 mL of molecular-grade water. The pH of the solution was adjusted to 7.3, using 1 M sodium hydroxide, and the final volume was adjusted to 1 L using molecular-grade water. (19) Babendure, J. R.; Adams, S. R.; Tsien, R. Y. J. Am. Chem. Soc. 2003, 125, 14716–14717.

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under gravity flow rate. The SPE cartridge was washed sequentially using the following wash solvents; 0.1 M HCl (2 mL), ethanol: water (70:30 v/v) (5 mL), dichloromethane (5 mL), and methanol (3 mL). The SPE cartridge was dried under vacuum for ∼5 min. Elution was performed using 95%:5% methanol:ammonia solution (2 × 3 mL increments) and the eluate (6 mL) was evaporated to dryness under a stream of nitrogen (+40-45 °C). Samples for aptamer analysis were reconstituted in HEPES buffer (2 mL). Samples for LC-MS/MS analysis were reconstituted in 2 mL of 1:3 (v/v) 5 M ammonium acetate (pH 4.5):methanol. Aptamer Assay Conditions. The conditions for the optimized fluorescence-based aptamer assay are detailed below. A fixed mass of RNA (2 µg) was preincubated with the test sample extract (2 mL) for 20 min (±5 min). The fluorescence signal associated with the RNA-MG complex was measured using fixed excitation (Ex) and emission (Em) wavelengths (Ex ) 618 nm/Em ) 643 nm). To correct for any matrix-related background fluorescence at these wavelengths, the fluorescence signal of the test sample extract was measured prior to incubation with the aptamer and a background subtraction correction made. Figure 2. Electropherogram showing the results of the SDS-PAGE analysis of Malachite Green RNA aptamers (15% denaturing). Lane 1 contains a decamer single-stranded DNA ladder (including marker sizes of 10, 20, 30, 40, 50, 60, 70, and 80 nucleotides). Lanes 2 and 3 contain the RNA products of two different transcription reactions yielding pure RNAs of length 38-39 nucleotides containing N and N + 1 products. N.B. RNA gel-mobility is slower than the equivalent single-stranded DNA products.

Preparation of 2,3-Dichloro-5,6-dicyano-1,4-benzoquinone (DDQ) Solution. 2,3-Dichloro-5,6-dicyano-1,4-benzoquinone (0.1136 g) was dissolved in acetonitrile (50 mL) to give a 0.01 M solution and placed in an amber glass bottle. The solution was prepared immediately prior to use. Fish Samples. Known blank trout (Salmo trutta and Oncorhynchus mykiss spp. and salmon Salmo salar) tissue were obtained from either controlled fish dosing studies or organic retail samples previously analyzed and confirmed negative for MG/LMG using confirmatory liquid chromatography tandem mass spectrometry (LC-MS/MS). The MG/LMG incurred salmon test material was obtained from the incurred tissue collection (http://incurredtissues.csl.gov.uk/, Veterinary Medicine Directorate, Addlestone, U.K.). Preparation of Fortified Fish Tissue. Analyte fortification was performed by applying a fixed volume of MG/LMG (as required) standard to finely sliced known blank tissue to achieve the appropriate concentration prior to extraction. Target tissue concentrations were in the range of 1-10 µg kg-1 (bracketing the current EU MRPL of 2 µg kg-1). The fortified samples were prepared in such a way that the final aqueous content changed by e2% via the addition of the spiking solution. Following fortification, the tissue was allowed to equilibrate for a minimum period of 15 min prior to extraction. Mixed-Mode Cation Solid-Phase Extraction Procedure for LMG/MG. The extraction solvent (0.5% acetic acid in acetonitrile (v/v)) (12 mL) was added to fish tissue (8 g) and homogenized for ca. 45 s. The homogenate was centrifuged for 10 min (3900 g, +4 °C). The supernatant (10 mL) was removed and oxidized using 0.1 M DDQ solution (800 µL) incubated at laboratory temperature in darkness for 15 min (±5 min). The oxidized extract was loaded onto a preconditioned OASIS MCX (500 mg/6 mL) SPE cartridge 2654

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RESULTS AND DISCUSSION Prior to the assay development phase, the performance of the RNA aptamer was evaluated and characterized using a buffered solution. RNA Stabilization. To stabilize and protect the RNA aptamer from ribonuclease (RNase) deactivation, a 4-(2-hydroxyethyl)-1piperazineethanesulfonic acid (HEPES)-based buffer, slightly below physiological pH (6.8-7.3), was developed for use as the binding buffer. The composition of this buffer was based on the binding buffers employed in previous intracellular-based studies using the MG RNA aptamer.20,21 The chaotrophic reagent (sodium dodecyl sulfate (SDS)) and scavenging reagent (ethylenediaminetetraacetic acid (EDTA)) were added to the buffer to deactivate RNase activity and chelate trace amounts of the divalent metal cations, e.g., Mg2+, Zn2+, Co2+, Mn2+, and Ca2+. As an additional measure, all manipulations involving the RNA aptamer were performed in an RNase-free environment (UV sterilization cabinet), using RNase free reagents and disposable glassware wherever possible. RNA Binding Affinity for MG. Apparent dissociation constants (Kd) for the aptamer have been reported for MG, ranging from 118 nM to 800 nM.19 The Kd value for the aptamer produced for use in this study was estimated to be ca. 600 nM, assuming a 1:1 binding model, using the following equation:

(

Y ) Bmax

free RNA Kd + free RNA

)

where Y is the response signal and Bmax is the maximum binding response observed. Spectrophotometric Characterization of the RNA-MG Complex in Solution. MG has an absorption maximum at 618 nm and a negligible fluorescence signal in aqueous solution. A spectrophotometric study was performed to investigate the (20) Baugh, C.; Grate, D.; Wilson, C. J. Mol. Biol. 2000, 301, 117–128. (21) Stojanovic, M. N.; Kolpashnikov, D. M. J. Am. Chem. Soc. 2004, 126, 9266– 9270.

Figure 3. Spectrofluorometric trace showing the fluorescence signal intensity (y-axis) associated with the RNA-MG complex before (black line) and after (dashed line) the addition of 50 µL of 1 M sodium hydroxide and following pH adjustment to the optimum pH ca. 7.0 (dotted line).

fluorescence signal enhancement associated with the constriction of MG when encapsulated by the aptamer. Three-dimensional (3D) measurements were performed whereby both the excitation (Ex) and emission (Em) wavelengths were scanned simultaneously over fixed ranges (Ex ) 550-650 nm) and (Em ) 550-750 nm). The following samples were included: (control 1) free MG, 0.5 µg mL-1; (control 2) free RNA, 0.5 µg mL-1; and (test 1) RNA-MG binding complex, each component present at 0.5 µg mL-1. The results (data not shown) reveal no significant emission peaks (>200 units) in the range of 640-650 nm in either of the two control samples. Emission peaks with a fluorescence intensity of 600-900 units at 640-650 nm were observed in the test sample, arising from the RNA-MG complex. The intensity of the emission peak for the RNA-MG complex was observed to be maximized between excitation wavelengths of 618-630 nm. The excitation wavelength was fixed at 618 nm and emission was monitored between 640 nm and 650 nm for the subsequent assay development experiments, unless otherwise stated. A fixed mass of the RNA (3 µg, equivalent to 239.6 nmoL-1) was incubated with increasing amounts of MG in assay buffer and the fluorescence signal was measured to evaluate the linearity of the binding interaction. To preserve the limited stock of the RNA, replication was not performed. The binding interaction between MG and the RNA produces a clear linear increase in fluorescence intensity over the range of 0-40 ng of MG in the assay buffer. In this range, a squared correlation coefficient (R2) of 0.997 was achieved (data not shown). The linearity of the fluorescence response plateaus between 40 ng and 60 ng of MG (equivalent to 121.4 nmoL-1 to 182.1 nmoL-1). Given that the reaction was performed with the aptamer in molar excess of the ligand, it is likely that the lack of increase in fluorescence intensity above 40 ng MG reflects fluorescence quenching rather than binding domain saturation.

The binding interaction between the aptamer and the MG major metabolite (LMG) was also investigated. Excitation and emission wavelengths between 250 nm and 750 nm were scanned synchronously. No significant change in fluorescence intensity associated with LMG (0.5 µg mL-1) was observed in the presence of the aptamer (data not shown). A further binding site competition assay was conducted, whereby the fluorescence signal associated with the RNA-MG complex was monitored in the presence of an ∼5-fold molar excess of LMG. The fluorescence signal of the RNA-MG complex was unaffected by the presence of the LMG competitor (data not shown). Based on these findings, it was concluded that LMG must be oxidized to the chromogenic (MG) form prior to analysis in the final assay, because of the apparent lack of binding affinity. Effect of pH on the RNA-MG Complex. The fluorescence spectrum of the RNA-MG complex with each component present at 0.5 µg mL-1 was measured prior to (and 1 min after) the addition of a strong alkali (1 M sodium hydroxide). The results (Figure 3) reveal that the RNA-MG fluorescence signal was lost in the presence of the alkali, indicating that the binding interaction was disrupted. At high pH (>9), RNA becomes denatured; the secondary and tertiary structures forming the ligand-binding domain are disrupted resulting in the dissociation of the ligand and loss of the fluorescence signal. The RNA aptamer was observed to be permanently degraded following exposure to alkali (alkali hydrolysis), as the fluorescence signal was not recoverable when the pH was returned to the optimum (ca. pH 7.0). The pH tolerance for the RNA-MG complex was investigated to establish the operational range. The assay buffer was adjusted to generate pH values between 1 and 14 units and the fluorescence signal of the complex (0.5 µg mL-1 RNA and 0.1 µg mL-1 MG) was recorded. The results (Figure 4) show that the operational pH range for the assay buffer was determined to be between 5 Analytical Chemistry, Vol. 82, No. 13, April 1, 2010

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Figure 4. Effect of assay buffer pH on the RNA-MG complex formation as measured by the fluorescence signal intensity (Ex ) 618 nm, Em ) 643 nm). The operational pH range for the assay buffer was found to be 5 and 9 (dotted-line box).

and 9 units (dotted line box), with the optimum signal being observed between pH 6 and 7.5. Effect of Metal Cations in the Assay Buffer. The composition of the HEPES-based assay buffer was examined to determine the critical constituents. The effect of varying the divalent metal cation (Mg2+) content and the requirement for the presence of monovalent cations, (K+) and (Na+) were considered. The most commonly used ions in RNA folding studies are group I (alkali) and group II (alkaline-earth) metals. Of these, K+ and Mg2+ are the most relevant to in vivo conditions. These ions have closed-shell electronic structures and interact with RNA predominantly via electrostatic forces.22 The ionic radius is relevant to ion-RNA interactions for two reasons. First, the steric problem of fitting an ion into a given chelation site, and second, it determines the hydration energy. The smaller the ionic radius, the greater the charge density and the stronger its interactions with the dipole moment of water. Five concentrations of Mg2+ were examined (0, 1, 5, 10, and 100 mM) and the rate of formation and stability of the RNAMG complex was monitored for a 30-min period. The RNA (1 µg) was added directly to a cuvette positioned in the spectrophotometer containing a solution of MG (10 ng mL-1). The instrument was programmed to perform a series of 60-s interval measurements (Ex ) 618 nm, Em. scanned between 640 nm to 660 nm) over a 30-min period. The results (Figure 5) reveal that the presence of Mg2+ is required for the formation of the RNA-MG complex as measured by the highest fluorescence intensity. The presence of Mg2+ ions was found to exert a complex stabilizing effect. The strongest fluorescence signal (1 mM Mg2+) was seen to plateau at ca. 450 s, followed by a rapid decline after ca. 1300 s, which is indicative of complex dissociation. Mg2+ ions are an essential component of the assay buffer and a concentration of 5-10 mM was deemed optimal for complex formation and stability, giving the next strongest fluorescence signal that did not decline over a 30-min period. Titration with Mg2+ allows large RNAs to adopt more-compact conformations termed electrostatic collapse, via shielding of the phosphate charges and thus, reducing the radius of gyration (22) Draper, D. E.; Grilley, D.; Soto, A. M. Annu. Rev. Biophys. Biomol. Struct. 2005, 34, 221–243.

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Figure 5. Plot showing the effect of varying the concentration of Mg2+ ions in the assay buffer (0, 1, 5, 10, and 100 mM) on the RNAMG complex formation and stability. A series of 60 s interval fluorescent measurements were taken (excitation fixed at 618 nm, emission monitored between 640 to 660 nm) in the presence of RNA (1 µg) and MG (10 ng mL-1) for a 30 min period.

Figure 6. Plot showing the effect of the monovalent metal cations, Na+ and K+ on the RNA-MG complex formation characteristics and stability. A series of 60 s interval fluorescent measurements were taken (Ex ) 618 nm, Em monitored between 640 nm to 660 nm) in the presence of RNA (1 µg) and MG (10 ng mL-1) for a 30-min period.

(RG).22 Divalent cations, in particular Mg2+ are more effective at inducing partial electrostatic collapse of RNA than monovalent ions.23 The requirement for K+ was determined in a similar manner as described for Mg2+ using a single concentration. Assay buffer was prepared in the absence of KCl (buffer base minus KCl) and also with KCl replaced by NaCl (buffer base plus NaCl). The degree of formation of the fluorescent RNA-MG complex was expressed as the measured change in fluorescence signal as a function of time, according the equation below: Formation of RNA-MG complex ) FL units at 600 s - FL units at 0 s 600 s The results (Figure 6) show that, in the absence of KCl or NaCl, the most intense fluorescence signal was obtained. The (23) Takamoto, K.; He, Q.; Morris, S.; Chance, M. R.; Brenowitz, M. Nat. Struct. Biol. 2002, 9, 928–933.

Figure 7. Plot showing the results of the time interval experiment to determine the time scale for the RNA-MG complex formation and the stability of the associated fluorescence signal at +16-20 °C. A fixed mass of MGA (2 µg) was added to a standard solution of MG (25 ng mL-1) and a series of 1-min-interval fluorescence measurements (Ex ) 618 nm, and Em ) 643 nm) were made over a 90-min period.

formation of the RNA-MG complex in the absence of KCl occurred at 1.1 FL units per second. After 1260 s, the fluorescence intensity declined rapidly indicative of complex dissociation. Both the complex formation and subsequent stability were found to be comparable in the presence of either NaCl or KCl (standard composition), although NaCl promoted a stronger fluorescence signal. Complex formation occurred at 0.7 FL units per s in the presence of either cation. The presence of a monovalent metal cation (either Na+ or K+) was deemed a requirement for the RNA-MG complex stability. Characteristics of RNA-MG Complex Formation. The time required for RNA-MG complex formation in the optimized assay buffer was monitored at ambient temperature (+16-20 °C). RNA (2 µg) was added to a quartz cuvette containing a MG solution (25 ng mL-1) prepositioned in the spectrophotometer. The instrument was programmed to perform a series of scans at 1-min intervals for a total of 90 min. Maximum binding (defined as the most intense fluorescence signal recorded) was achieved after ca. 16 min (see Figure 7). Thus, the formation of the binding complex occurs in a time-dependent manner. Once formed, the complex-associated fluorescence remains stable (not declining below 90% of the maximum signal) for up to 60 min. The RNAMG complex dissociation was determined to be a gradual process, with the fluorescence remaining reasonably stable over the entirety of the monitoring period. Effect of Temperature on the Formation and Stability of the RNA-MG Complex. The effect of four different temperaturess0 (ice), +4-8 °C (fridge), +16-20 °C (laboratory, light) and +35 °Cson the rate of the RNA-MG complex formation and longer-term stability were investigated. MG solution (10 ng mL-1) was dispensed into cuvettes and preincubated at the designated temperature prior to introduction of the aptamer. Immediately after the addition of RNA (1 µg), a measurement was taken. Further measurements were taken at intervals of 5, 10, 15, 20, 30, 60, 120, and 180 min on day 1 and daily for the 10 subsequent days (excluding weekend days). After the initial 10-day period, measurements were taken at regular weekly intervals until the fluorescence signal declined to e50% of the starting signal.

Figure 8. Plot of the effect of different storage temperatures on the formation of the MGA-MG complex and stability over a 48-day period monitored by the fluorescence response at Ex ) 618 nm, Em ) 643 nm. MG was present, at a concentration of 10 ng mL-1, in the assay buffer, RNA at 1 µg, four temperatures were investigated: (2) 0 °C, (*) +4-8 °C, (9) room temperature (+16-20 °C), and (b) +35 °C.

The results (Figure 9, shown later in this work) reveal that both the time scale required for the RNA-MG complex formation and subsequent stability are affected by temperature. The aliquots stored at 0 °C and +35 °C produced the weakest signals and the fluorescence declined to ca. 50% after one week of storage. The reaction performed at 0 °C achieved ca. half the maximum fluorescence, compared to that performed at laboratory temperature in 180 min. The formation of the complex was significantly inhibited at +35 °C. The fluorescence signals for both the reactions performed at 0 °C and 35 °C were not fully recoverable when the samples were returned to laboratory temperature, suggesting that the aptamer was denatured under these storage conditions. The reaction performed at fridge temperature required the longest time to achieve the maximum signal. Once achieved, however, the complex had the highest intensity and was the most stable, surviving for ca. 4 weeks. It was postulated that the enhanced fluorescence signal and stability observed under fridge conditions could be a cumulative vibrational stabilization effect resulting from the molecular constriction of MG within the RNA binding pocket and the cold temperature. The reaction performed at laboratory temperature in the light gave its signal maximum in the first 60 min. The signal remained stable (within a 10% reduction of the maximum) for eight days. At temperatures outside of the range (+16-20 °C) used for the original aptamer selection, the formation of the secondary structure and ligand binding domain formation may be disrupted. Incubation temperature is a critical factor, with the range of +16-20 °C being determined to produce optimal results. The results (Figures 7 and 8) are consistent with the suggestion that RNA folding/unfolding occurs in discrete steps and transient intermediate structures are formed during the process.24 Organic Solvent Tolerance of the RNA. The performance of the RNA was investigated in the presence of two polar organic solvents: methanol and acetonitrile. MG solutions (10 ng mL-1) were prepared in varying volumes of assay buffer and either methanol or acetonitrile to produce solutions containing 2%, (24) Onoa, B.; Tinoco, I. Curr. Opin. Struct. Biol. 2004, 14, 374–379.

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Figure 9. Effect of percentages of the two organic solvents (acetonitrile (light gray bars) and methanol (black bars)) in assay buffer on the RNA-MG complex activity, in terms of fluorescence intensity, compared to that of the complex in 100% aqueous buffer. The mass of MGA was fixed at (2 µg) and MG at 10 ng mL-1.

5%, 10%, 15%, 20%, 30%, or 50% organic solvent (v/v). A fixed mass of the RNA (2 µg) was added to each aliquot (2 mL) of the MG solution, incubated for 15-20 min at +16-20 °C, and the fluorescence was measured. The fluorescence signal was calculated as a percentage of the mean intensity of the RNAMG complex response in the presence of 100% assay buffer. The results (Figure 9) reveal that, even in the presence of 2% organic solvent, the activity of the RNA is significantly reduced (ca. 30% decrease). Acetonitrile has a greater inhibitory affect than methanol. Even in the presence of 50% methanol, a measurable fluorescence signal could still be observed. Hydrogen bonding interactions are essential for the formation of the rigid RNA-MG complex. Methanol, which is a protic solvent, displays some hydrogen bonding, which is not achieved with acetonitrile. Thus, whenever possible the binding reaction should be performed in a 100% aqueous solution to generate the optimum fluorescence signal. Dose Response Curve. To characterize the performance of the RNA over an extended concentration range, a concentration response curve was prepared. A total of 12 MG solutions were prepared in assay buffer, spanning concentrations of 0.3-1040 ng mL-1 (1.0-3160 nmol L-1). A fixed mass of RNA (2 µg) was used for each reaction. After a 15-20 min incubation period at +16-20 °C, the intensity of the fluorescence signal was measured. The results (Figure 10) were plotted on a semilogarithmic scale and a regression line fitted using the four-parameter logistic equation (4-PL). A sigmoidal curve with the steepest region falling between 10 nmol L-1 and 100 nmol L-1 (3.3-33 ng mL-1) of MG was obtained. The half maximal effective concentration (EC50) for the RNA-MG complex was calculated as 46.6 nmol L-1 (15.4 ng mL-1), representing the concentration of MG at which 50% of the maximal fluorescence is observed at a fixed RNA mass of 2 µg. The threshold concentration of MG at which the signal was detectable above the background noise was 5 nmol L-1 (2 ng mL-1). Maximal binding (Bmax) occurred in the range of 158-316 nmol L-1 (52-104 ng mL-1), observed as the maximal fluorescence signal indicating that all available binding domains are occupied. Development of the Aptamer-Based MG Diagnostic Assay. Initially, two different assay designs were evaluated to select the optimum format for the MG diagnostic assay using the RNA. The fluorescence-based direct assay and a surface plasmon resonance (SPR) biosensor assay were compared. The fluorescence-based 2658

Analytical Chemistry, Vol. 82, No. 13, April 1, 2010

Figure 10. Concentration response curve in assay buffer showing the response of the RNA-MG complex (Ex ) 618 nm, Em ) 643 nm) plotted against the dose of free MG in solution (concentration expressed in terms of log10 nmol L-1). The mass of the RNA was fixed at 2 µg per reaction. The threshold level (equivalent to 5 nmol L-1) and the EC50 value (equivalent to 46.6 nmol L-1) are indicated.

direct assay was deemed the most appropriate format, in terms of the requirements for RNA-MG complex formation, analytical performance, and other considerations such as technology transferability, cost per analysis, and ease of operation. The procedure adopted uses a fixed mass of RNA (2 µg), which is preincubated with the test sample (2 mL) for 20 min (±5 min). The fluorescence signal associated with the RNA-MG complex is measured using fixed excitation and emission wavelengths (Ex ) 618 nm/Em ) 643 nm). To correct for any matrix-related background fluorescence at these wavelengths, the fluorescence signal of the test sample was measured prior to incubation with the aptamer and a background subtraction correction made. Development of the Extraction Procedure Compatible with the Aptamer-Based Assay Format. Initially, an approach based on the QuEChERS method for the rapid preparation of samples prior to multiresidue pesticide analysis25 was investigated for both extraction and cleanup. To assess the suitability of the QuEChERS style method for this application, salmon tissue samplessuntreated and fortified with LMG (20 µg kg-1)swere extracted using acetonitrile, along with a process blank, according to the basic QuEChERS procedure. The procedure was modified to evaluate the performance of different sorbent phases. Thus, five replicate aliquots (equivalent to 1 g of tissue) of the supernatant were collected. All aliquots were treated with the oxidizing agent, 2,3-dichloro-5,6-dicyano-1,4-benzoquinone (DDQ) to convert residues of LMG to MG. The oxidation reaction was conducted using 0.01 M DDQ for 15 min at laboratory temperature in darkness. One aliquot of the supernatant was evaporated to dryness, to assess the assay tolerance to crude extract (i.e., without further cleanup). The remaining four aliquots were subject to purification via dispersive solidphase extraction (dSPE), using bulk sorbent material. Four different reversed-phase sorbents were evaluated, including silica-based C18 and C8 and polymeric-based OASIS HLB and Strata X. Following the evaporation step, all of the extracts were reconstituted in aqueous assay buffer and analyzed via the (25) Anastassiades, M.; Lehotay, S. J.; Stanjnbaher, D.; Schenck, F. J. J. AOAC Int. 2003, 86, 412–431.

Table 1. Comparison between Crude Extract and a Variety of Dispersive “QuEChERS”-Based Solid-Phase Extractions Using the Aptamer-Based Fluorescence Assay (RNA 2 µg, Ex ) 618 nm, Em ) 643 nm)a Measured Fluorescence Intensity, Based on the Crude Solvent Extract Plus Type of dSPE Cleanup Silica

Polymeric

extract type

none

C18

C8

OASIS

Strata-X

blank salmon spiked salmon (20 µg kg-1) positive control 1 solvent standard, taken through full method (20 µg kg-1) positive control 2 matrix-matched standard, blank salmon extract spiked immediately before measurement (20 µg kg-1)

80.4 89.3 689.3

64.5 59.1 789.2

75.6 68.5 795.6

60.7 67.5 643.5

58.8 64.3 699.8

850.6

865.8

895.4

902.5

911.7

a

LMG fortified into salmon tissue at 10× MRPL concentration (20 µg kg-1).

aptamer-based test. After the addition of RNA, a fluorescence enhancement above the level recorded in the untreated controls was not observed in any of the five fortified extracts (see Table 1). Strong fluorescence signals were observed in both positive control samples: a solvent standard, taken through the full method, and a matrix-matched standard, fortified post-extraction. This finding showed that solvent solutions of LMG/MG can be recovered via a QuEChERS style procedure and that RNA-MG binding remains active in the presence of both the crude and purified fish tissue extracts. Because of the lack of apparent recovery in the presence of fish tissue, it was hypothesized that the MG/LMG was associated with specific matrix component(s) and, in the bound conformation, is not recognized by the aptamer. A series of further experiments were performed to help overcome this technical issue including the addition of a defatting solvent to remove more of the nonpolar lipid component. For example, prior to the dSPE step, the acetonitrile extract was partitioned using an immiscible nonpolar organic solvent. The following solvents were evaluated in ascending order, relative to polarity: hexane < isooctane (2,2,4-trimethylpentane) < pentane < cyclohexane. The defatting wash step using nonpolar solvents was not observed to improve the assay performance (data not shown). Based on the lack of improvement in recovery after defatting, it was concluded that MG/LMG residues may be strongly associated with, or encapsulated by, the fish lipid component or the fatty acid fraction at the point of the aqueous reconstitution step immediately prior to measurement. As an alternative approach, MG residues may be recovered via solid-phase extraction, using cartridges prepacked with a cation exchange sorbent.8,26 A procedure based on the use of acidified acetonitrile extraction solvent and cation exchange purification was evaluated for performance with the aptamer assay. A comparison between a bonded silica (SCX Bond Elut, Varian 500 mg/6 mL) and a modified cross-linked polymeric based sorbent (OASIS MCX, Waters 500 mg/6 mL) was made. The background fluorescence signal in both the tissue and the method blank control samples was determined to be higher using the silicabonded phase. This background fluorescence signal was attributed to the presence of silica-derived “fines” in the final extract and was determined to mask the response of low concentrations of (26) Stubbings, G.; Tarbin, J.; Cooper, A.; Sharman, M.; Bigwood, T.; Robb, P. Anal. Chim. Acta 2005, 547 (2), 262–268.

Table 2. Calculated MG Recovery Obtained from Salmon Tissue Fortified with LMG at Three Concentrations 1, 2, and 5 µg kg-1 (n ) 5) Recovery, as Determined Using LMG Spikes (Measured by LC-MS/MS as MG) (%) replicate 1 2 3 4 5 mean standard deviation, SD coefficient of variation, %CV a

[LMG] ) 1 µg [LMG] ) 2 µg [LMG] ) 5 µg kg-1 kg-1 kg-1 79 76 76 92 97 84 10.0

87 75 98 101 83 89 10.9

92 84 90 75 73 83 8.5

11.9

12.3

10.3

These extracts were analyzed quantitatively using LC-MS/MS.

MG residues. In contrast, the polymeric MCX sorbent was able to withstand a greater pH range (0-14) than the silica-based sorbent. For these reasons, the polymeric sorbent was selected for further evaluation and use in the final procedure. To assess the performance of (i) the developed SPE procedure and (ii) the efficiency of the oxidation reaction, a recovery experiment was conducted and sample extracts were analyzed quantitatively via a LC-MS/MS based procedure reported by Scherpenisse and Bergwerff.27 Aliquots of known blank salmon tissue were fortified with LMG at three concentrations (1, 2, and 5 µg kg-1) and were extracted according to the developed procedure. Following the final sample evaporation step, the sample residue was reconstituted in a solvent compatible with LC-MS/MS analysis. The results from this experiment (Table 2) revealed that the individual sample MG recovery exceeded 70% at each of the concentrations tested. The mean recoveries were determined to be 80-90% (n ) 5), and the intrabatch repeatability (as a coefficient of variation) was