Introduction

The ability to quantify the concentrations of drugs, second messengers, hormones, and proteins is of fundamental biomedical importance. Although DNA microarray chips are revolutionizing biology by expanding our analyses from single-gene to genome-wide gene expression, analogous methods for the simultaneous study of the metabolome and proteome are not yet available. In addition, rapid monitoring of cellular events such as second messenger synthesis and hormone secretion in single cells is key to understand cellular organization in higher organisms, yet is still not fully accomplished. Finally, early

From: Methods in Molecular Biology, vol. 335: Fluorescent Energy Transfer Nucleic Acid Probes: Designs and Protocols Edited by: V. V. Didenko © Humana Press Inc., Totowa, NJ

pathogen detection is of increasing urgency in clinical diagnosis and biodefense in the face of newly emerging infectious diseases. For these applications, new biosensor technologies are needed.

Biosensors are hybrid analytical devices that amplify signals generated from the specific interaction between a receptor and an analyte of interest, through a biophysical mechanism. Biosensors use tissues, whole cells, artificial membranes, or cell components like proteins or nucleic acids as receptors, coupled to a physicochemical signal transducer. The ideal biosensor is extremely sensitive, specific for the target of interest, adaptable to a wide range of target analytes, compact, rugged, and consumes very little energy or other resources.

Particularly notorious targets for biosensors are small organic compounds such as theophylline (Fig. 1B). Theophylline is a bronchodilating drug widely used in the treatment of asthma, bronchitis, and emphysema, with a narrow therapeutic range (1). Its serum levels must, therefore, be monitored carefully to avoid toxicity. Detection of theophylline is particularly challenging because of its resemblance to the ubiquitous caffeine, which carries only one additional methyl group on the N7 of the purine ring (Fig. 1B).

RNA is a unique biopolymer in that it can carry genetic information, encoded in its linear sequence, and can bind ligands or substrates specifically, even catalyze chemical reactions, based on its ability to fold into complex three-dimensional structures. The high thermodynamic stability of the secondary structure (Watson-Crick basepairing) of RNA provides for a stable scaffold to acquire diverse tertiary structures that can recognize ligands with extremely high specificity and sensitivity, in some cases even surpassing those of antibodies (2). As a consequence, RNA has recently been proposed as ligand-spe-cific receptor component for biosensors (3-5), and even nature herself appears to resort to such RNA-based sensing (6).

RNA structural motifs called aptamers can be evolved in vitro to bind a desired ligand with high selectivity and tight binding affinity (2). The modularity of RNA structure allows one to incorporate aptamers into larger RNAs without loss of their binding properties. Breaker and co-workers have exploited this property to develop an allosteric RNA comprised of three elements: an aptamer as ligand receptor, a catalytic RNA called the hammerhead ribozyme as amplifier, and a communication module to couple their functions (4,7-12). Specifically, the aptamer and the communication module are incorporated into stem II of the hammerhead ribozyme in a way that catalytic activity is enhanced when the ligand binds. Such allosterically activated ribozymes are called aptazymes. Figure 1A shows the signal transduction mechanism for this aptazyme. In the absence of theophylline, the communication module is in a misaligned basepairing pattern, and the aptazyme-substrate complex is cata-lytically inactive. Upon addition of theophylline, the aptamer undergoes a con-

Fig. 1. Reaction pathway of the theophylline dependent aptazyme. (A) In the absence of theophylline (Left), the aptazyme-substrate complex is in a catalytically inactive conformation because the communication module (boxed) is misaligned. The characteristic high fluorescence resonance energy transfer (FRET) efficiency results in strong acceptor emission at 665 nm. Upon binding of theophylline (Middle), the catalytically active conformation of the ribozyme is formed by the realignment of the communication module (box). This event induces formation of domain 2, which is required for activity of the hammerhead ribozyme (14). Substrate cleavage occurs at the arrow, and the rapid dissociation of the 5' and 3' products follows (Right). The resulting separation of the fluorophores decreases FRET, and the donor emission at 565 nm increases at the expense of acceptor emission at 665 nm. (B) Chemical structures of theophylline and caffeine. The circles highlight their only difference: a methyl group on N7, which is distinguished by our aptazyme.

Fig. 1. Reaction pathway of the theophylline dependent aptazyme. (A) In the absence of theophylline (Left), the aptazyme-substrate complex is in a catalytically inactive conformation because the communication module (boxed) is misaligned. The characteristic high fluorescence resonance energy transfer (FRET) efficiency results in strong acceptor emission at 665 nm. Upon binding of theophylline (Middle), the catalytically active conformation of the ribozyme is formed by the realignment of the communication module (box). This event induces formation of domain 2, which is required for activity of the hammerhead ribozyme (14). Substrate cleavage occurs at the arrow, and the rapid dissociation of the 5' and 3' products follows (Right). The resulting separation of the fluorophores decreases FRET, and the donor emission at 565 nm increases at the expense of acceptor emission at 665 nm. (B) Chemical structures of theophylline and caffeine. The circles highlight their only difference: a methyl group on N7, which is distinguished by our aptazyme.

formational change (13), which aligns the communication module so that two adjacent G-A basepairs and a noncanonical A-U basepair in domain 2 of the hammerhead ribozyme can coaxially stack (14). Consequently, the catalyti-cally active conformation is accessed, the substrate backbone is cleaved, and the reaction products are released.

Although classically ribozyme activity induced by allosteric binding has been detected by radioisotope labeling (4), the use of fluorophores provides an attractive alternative for the following reasons: (1) Fluorophores do not carry the inherent risks in handling and disposal of radiolabeled nucleotides. (2) Fluorescence can be easily monitored and quantified directly in solution, whereas radioisotope assays require discontinuous analysis. Therefore, fluorescence detection accelerates the analysis process. (3) Large numbers of samples can be measured in short periods of time because microplate fluorescence readers and further miniaturized microarrays enable automation of the detection process. (4) The fluorescence probe shelf life is virtually unlimited compared to radioisotopes, which decay over time. In general, we have found fluorescence spectroscopy to be a versatile probe for studying cleavage kinetics and confor-mational changes of catalytic RNAs (15-23).

With this in mind, we have developed a biosensor component for theophylline based on the aptazyme described by Breaker et al. (15). Our aptazyme reports theophylline-induced cleavage in trans (i.e., with an external, replaceable substrate) by fluorescence resonance energy transfer (FRET) between a donor (Cy3) and an acceptor fluorophore (Cy5) covalently linked to the substrate termini (Fig. 1A). Before cleavage, the donor is close to the acceptor fluorophore and FRET occurs, which is characterized by a strong emission at the acceptor wavelength of 665 nm (Figs. 1 and 2A). Upon substrate cleavage and product dissociation, FRET breaks down and the donor specific emission at 565 nm increases (Figs. 1 and 2A). This breakdown of FRET provides an amplified signal for the presence of theophylline. The chosen pair of cyanine fluorophores for FRET is compatible with our goal to apply single-molecule fluorescence microscopy (20,21,23) to the detection of analyte binding.

This chapter reviews in detail the methods and protocols to prepare our theo-phylline specific aptazyme and to label its substrate with fluorophores. We also include detailed protocols to characterize by FRET the binding affinity of the target, theophylline, as well as the external substrate to the aptazyme (see Note 1).

Fig. 2. Fluorescence resonance energy transfer (FRET) readout for the theophylline dependent aptazyme as basis for a biosensor. (A) Emission spectrum of Cy3-Cy5 doubly labeled substrate. Upon addition of the aptazyme and theophylline (dotted line), donor emission increases while acceptor emission decreases, characteristic of a FRET decrease upon substrate cleavage and product release. (B) Excitation spectrum of the Cy3-Cy5 doubly labeled substrate. Upon addition of the aptazyme and theophylline (dotted line), donor excitation results in a lower acceptor signal than before, characteristic of a FRET decrease upon substrate cleavage and product release. (C) Lower panel: Fluorescence emission time traces of the donor (dark gray measured at 565 nm) and acceptor (light gray, measured at 665 nm) fluorophores upon excitation at 540 nm. The upper panel shows the resulting FRET time trace (black). Initially the relative FRET efficiency is constant at 0.8. After 60 s the aptazyme is added and the FRET ratio decreases as a result of cleavage and product release in the absence of theophyl-line. After approx 600 s theophylline is added, and the FRET decrease is enhanced as a result of accelerated cleavage induced by theophylline. This decrease is fit to a single-exponential decay (white line), whose rate is kobs. Addition of caffeine instead of theo-phylline does not cause a similar acceleration in cleavage (top panel, light gray). (D) Concentration dependence of the rate of FRET decrease, kobs, as a function of theo-phylline concentration. Theophylline concentrations in the low |jM range are sufficient to induce a measurable rate enhancement (inset). See text for experimental details.

2. Materials

2.1. Aptazyme Transcription and Purification

1. Polymerase chain reaction (PCR) buffer: 10 mM Tris-HCl, pH 8.3, 1.5 mM MgCl2, and 50 mM KCl.

2. Tris-EDTA (TE) buffer: 10 mM Tris-HCl, pH 7.0, 1 mM EDTA.

3. Tris-boric acid-EDTA (TBE) buffer: 100 mM Tris base, 70 mM boric acid, 2 mM EDTA.

4. Formamide gel loading buffer: 90% (v/v) formamide, 0.025% (w/v) xylene cyanol, and 0.025% (w/v) bromophenol blue in TBE buffer.

5. Buffered chloroform/isoamyl alcohol: 96% (v/v) chloroform, 4% (v/v) isoamyl alcohol. Add 1/3 vol TE buffer. Mix and allow the aqueous and organic phases to separate. Use only the chloroform/isoamyl alcohol layer (bottom).

6. Buffered phenol/chloroform: 50% (v/v) phenol, 50% (v/v) buffered chloroform/ isoamyl alcohol. Add one-third volume TE buffer. Mix and allow the aqueous and organic phases to separate. Use only the phenol/chloroform/isoamyl alcohol layer (bottom).

7. Transcription buffer: 120 mM HEPES-KOH, pH 7.6, 30 mM MgCl2, 40 mM dithiothreitol (DTT), 5 U/mL inorganic pyrophosphatase, 2 mM spermidine, and 0.01 % (w/v) Triton X-100.

8. Elution buffer: 0.1 mM EDTA, pH 8.0, 0.1% (w/v) sodium dodecyl sulfate (SDS), and 500 mM ammonium acetate.

9. DNA primers (Invitrogen, Carlsbad, CA).

10. Taq DNA polymerase (TaKaRa Biochemical, Berkeley, CA).

13. Commercial T7 RNA polymerase (e.g., TaKaRa Biochemical), or purified in native form from overexpressing Escherichia coli strain BL31/pAR1219 (24), or purified in histidine-tagged form from overexpressing E. coli strain BL21/pRC9 (25).

15. Spermidine (e.g., Fisher Scientific).

17. Inorganic pyrophosphatase (Sigma-Aldrich, St. Louis, MO).

18. Vertical slab gel electrophoresis apparatus (20 x 16 cm2), including glass plates, 1-mm spacers, 1-mm fitting seal, 1-mm one- or two-well comb, clamps, and aluminum plate (e.g., CBS Scientific, Del Mar, CA).

19. Centricon Plus-20 centrifugal filters (Millipore, Bedford, MA).

20. Hand-held ultraviolet (UV) lamp, wavelength 312 or 254 nm (e.g., Fisher Scientific).

21. Thin-layer chromatography (TLC) plate (20 x 20 cm2) with fluorescent indicator (e.g., Fisher Scientific).

22. Empty Poly-Prep chromatography column (Bio-Rad Laboratories, Hercules, CA).

2.2. Synthetic RNA Substrate Purification and Labeling

1. High-performance liquid chromatography (HPLC) system with C8-reversed phase column (e.g., ProStar system from Varian, Palo Alto, CA).

2. HPLC mobile phase B: 100% acetonitrile.

3. HPLC stationary phase A: 100 mM triethylammonium acetate, pH 7.0.

4. Synthetic RNA oligonucleotides (e.g., HHMI Biopolymer/Keck Foundation Biotechnology resource laboratory at the Yale University, School of Medicine, New Haven, CT).

5. Triethylamine trihydrofluoride (e.g., Sigma-Aldrich or Fisher Scientific).

6. N,N-dimethylformamide (e.g., Fisher Scientific; optional).

9. 14-mL Falcon centrifuge tube (e.g., Fisher Scientific).

10. Dimethyl sulfoxide (e.g., Fisher Scientific).

11. Cy5-succinimidyl ester (Amersham Biosciences, Piscataway, NJ).

2.3. Steady-State FRET Assays

1. Standard buffer: 50 mM Tris-HCl, pH 7.5, 10 mM MgCl2, and 25 mM DTT as oxygen scavenger.

2. AMINCO-Bowman 2 spectrofluorometer (Thermo Electron Corporation, Rochester, NY), or similar.

3. Quartz cuvet, 3-mm pathlength, 200 pL vol (Starna Cells, Atascadero, CA).

4. Theophylline (e.g., Sigma-Aldrich).

2.4. Time-Resolved FRET Assays

1. Frequency doubled Nd:VO4 pump laser, e.g., Millenia Xs-P (Spectra-Physics, Mountain View, CA).

2. Tunable Ti:Sapphire picosecond laser operating at 980 nm, e.g., Tsunami (Spectra-Physics, Mountain View, CA).

3. Pulse picker/frequency doubler module, e.g., model 3980-2 (Spectra-Physics).

4. Sample compartment Koala (ISS, Champaign, IL).

5. Microchannel plate photomultiplier tube, model R3809U-50 (Hamamatsu Corporation, Bridgewater, NJ).

6. Time-correlated single-photon counting card, SPC-630 (Becker and Hickl, Berlin, Germany).

7. Quartz cuvet, 10-mm sides, 80 pL volume (Starna Cells).

8. Emission band-pass filter, 25 mm diameter and 4-mm thick, HQ570/10m (Chroma, Rockingham, VT).

9. Fused silica filter-mimic, 25 mm diameter and 4 mm thick (Edmunds Industrial Optics, Barrington, NJ).

10. Circular neutral density variable filter (Edmund Industrial Optics).

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