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The use of charge flow and quenching (CFQ) to probe nucleic acid folds and folding
Methods 52 (2010) 141–149
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Review Article
The use of charge flow and quenching (CFQ) to probe nucleic acid folds and folding Edward K.Y. Leung a, Dipankar Sen b,* a b
Department of Biochemistry & Molecular Biology, The University of Chicago, Chicago, IL 60637, USA Department of Molecular Biology & Biochemistry, Simon Fraser University, Burnaby, British Columbia, Canada V5A 1S6
a r t i c l e
i n f o
Article history: Available online 8 June 2010
a b s t r a c t Charge flow and quenching (‘‘CFQ”) is a relatively new, versatile, and easily carried out methodology for probing a number of unique features of DNA and RNA folded structures, and of their folding pathways. An electrical charge (an electron hole, or radical cation) is injected site-specifically into the end of a predetermined reference helix within the larger DNA or RNA structure. The fate of the injected charge, as it percolates through the folded DNA or RNA is then monitored by mapping the oxidative consequences of the charge flow. Some of the kinds of structural and folding information that can be obtained from CFQ experiments include: a quantitative measure of helix–helix connectivity; the dynamics of specific bases; folding and unfolding pathways; the mapping of unusual, conformation-dependent, electronic properties of individual bases; extents of solvent exposure and susceptibility to quenching from the solvent. CFQ is a relatively new methodology, and is applicable to DNA and RNA structures and folds. In the near future it is expected that the range of applications of this methodology will increase dramatically. Ó 2010 Elsevier Inc. All rights reserved.
1. Introduction 1.1. Background Over the past two decades, intensive research has shown that DNA double helices, in aqueous solution, are able to conduct electrical charge (reviewed in [1–3]). Two classes of charge conduction have been studied: (1) a class in which the charge carrier is a nucleobase radical cation (an ‘‘electron hole”); and (2) a class in which it is a radical anion (an ‘‘excess electron”). On the whole, hole transfer has been studied in greater depth, and the term ‘‘charge conduction” will refer exclusively to hole transfer for the remainder of this paper. Charge flow in DNA can conveniently be initiated by photo-excitation of a sensitizer moiety, such as anthraquinone (AQ), covalently linked and p-stacked upon the end of a double helix. The base radical cation generated within the helix has been found to propagate along the helix over relatively long distances (at least 200 Å) [4–6]. Three possible mechanisms for photo-induced charge transfer through DNA were originally proposed: (1) molecular wire; (2) super-exchange; and (3) charge hopping (reviewed in [7]). In the molecular wire model, charge is injected from the donor to a bridge element, where it localizes prior to moving incoherently towards the acceptor (Fig. 1A). In the super-exchange mechanism, the charge does not localize on the bridge element, but tunnels from
* Corresponding author. Fax: +1 778 782 5583. E-mail addresses: [email protected] (E.K.Y. Leung), [email protected] (D. Sen). 1046-2023/$ - see front matter Ó 2010 Elsevier Inc. All rights reserved. doi:10.1016/j.ymeth.2010.06.006
the donor directly to the acceptor (Fig. 1B). The hopping model, by contrast, occurs via the hopping of charge in a thermally activated fashion, from guanine base to guanine base (Fig. 1C). Experimental data indicate that DNA charge transfer does not follow the molecular wire mechanism, because the strong distance dependence of charge migration through DNA does not behave like a molecular wire [8]. The super-exchange mechanism occurs only efficiently over short distances (<10 Å); however, experiments have found charge transfer in DNA up to 200 Å [4–6]. Therefore, the most plausible mechanism is that of charge hopping. Of the four DNA bases, guanine (G) is the most easily oxidized (Table 1) and the guanine radical cation, G+, is the most stable of the four possible radical cations [9]. G+ is the intermediate charge carrier in the hopping process, and adjacent Gs (for example 50 -GG-30 or 50 -GGG-30 ) provide a particularly good thermodynamic sink for the charge. Theoretical studies have shown the radical cation on the 50 guanines of sequences such as 50 -GG-30 or 50 -GGG-30 are particularly stabilized relative to guanines towards the 30 end (Table 2), but not vice versa [10]. A recent refinement of the charge hopping model, the ‘‘phononassisted polaron-like hopping” model, most successfully accounts for hole conduction [5,6,11]. In this model, the charge is delocalized over several DNA bases due to local structural distortions of the DNA helix. The base-pairing domain within the DNA base stack can act as a gate for DNA-mediated charge transfer. In other words, the charge transfer occurs as a hopping process between conformationally gated domains of well-stacked base pairs (reviewed in [7]). These delocalized domains may transiently form and break, and facilitate or inhibit charge transfer.
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Fig. 1. Mechanisms of charge transfer in DNA. (A) The induced charge is delocalized throughout the entire molecule in the molecular wire model. (B) The charge jumps directly from the charge donor to the charge acceptor in the super-exchange model. (C) The charge, in the form of a radical cation, migrates from guanine to guanine until it reaches the acceptor in the hole hopping model. The charge can tunnel through a stretch of consecutive adenines. (D) The charge is delocalized over several base pairs in the phonon-assisted polaron-like hopping model and propagates along the helix.
Table 1 Oxidation potential for nucleosides. The first five oxidation potentials were measured in (a) aprotic solvent (acetonitrile) conditions [9]. The remaining oxidative potentials were determined in protic (aqueous) solvents at (b) pH 7.0 [32] and (c) pH 8.0 [33]. The oxidation potentials of the bases are dependent on the solvent (aprotic versus protic) conditions, and therefore, hydrogen bonds will change the oxidation potentials of all nucleobases. Base
E0(V vs NHE)a
Guanine Adenine Cytosine Thymine Uracil rG rA dC dT r(8-oxoG) r(8-oxoA) r(5-OH-C) r(5-OH-U)
1.49 1.96 2.14 2.11 P2.39
E0(V vs NHE)b
E0(V vs NHE)c
pH
0.58 0.92 0.62 0.64
7 7 7 7 8 8 8 8
1.29 1.42 1.60 1.70
Table 2 Calculated oxidation potential for bases. (a) The oxidation potential of the 5’-most guanine is influenced by it adjacent bases [13]. (b) The oxidation potential of 8oxoguanine can also be influenced by it neighbouring bases [34]. Sequence
E0(V vs NHE)a
GGG GG GA GC GT G 8-oxoG (8-oxoG)G G(8-oxoG)
0.64 0.84 1.00 1.15 1.16 1.20
E0(V vs NHE)b
0.85 0.53 0.08 0.18
Any guanine that hosts the mobile radical cation transiently is susceptible to a side reaction with water (or, in some instances,
dissolved oxygen), leading to the formation of modest levels of such guanine oxidation products as 8-oxoguanine or diaminooxazalone (reviewed in [12]). The formation of these oxidation products (Fig. 2A, B, adapted from [12]) supplies a useful biochemical handle for tracing both the actual charge flow path though DNA, and its consequences. These are possible because treatment with hot piperidine (or other comparable aqueous base) breaks the DNA strand at sites containing the guanine oxidation products (Fig. 2C). The susceptibility of any specific guanine within a double helix to charge flow-related oxidation depends, in degrees, on the sequence context of the guanine [2–4]. Stretches of multiple guanines, such as 50 -GG-30 or 50 -GGG-30 , are particularly susceptible to oxidation, with the 50 -most guanine of the stretch the most extensively oxidized [2–4,12]. Densitometry trace of charge flowdependent DNA cleavage bands (‘‘CFDC”), separated by gel electrophoresis provides a convenient and quantitative means for mapping charge flow through double helices. The sequencing gel shown in Fig. 3 illustrates the patterns of CFDC in a typical DNA double helix. While the guanine ladder on the left shows more or less equivalent guanine band intensities for all the guanines within the 50 -32P-labeled strand, the guanine band intensities in the charge flow lane (‘‘Duplex reaction”) indicate how patterns of CFDC vary markedly (and predictably) at different guanines, depending on their location. Thus, isolated single guanines typically show low levels of CFDC, while the 50 -most guanine of a 50 -GG-30 stretch (for instance the ‘‘proximal” and ‘‘distal” guanine doublets) show guanine oxidation (and cleavage) heavily biased towards the 50 -most guanine. In a more extensive stretch of guanines (such as 50 -GGG-30 or the 50 -GGGGG-30 stretch shown in Fig. 3) the pattern of CFDC is more complex, although largely predictable [13]. Experimentally, it is not a surprise to find that any interruption to or deviation from the standard geometry and base p-stacking patterns found within a Watson–Crick base-paired double helix impacts on the efficiency of charge conduction. Where a p-stack is perturbed (such as with mismatches or bulges), charge migration efficiency diminishes notably [14,15]. Therefore, in principle, CFDC provides a simple measure of charge transfer through intact DNA helices as well as assemblages made of helices and other structural components, including but not restricted to DNA three-way and four-way helical junctions [16–19]. Such charge transfer patterns (and, in special cases, patterns of reductant-mediated quenching of charge flow) are able, in turn, to provide information on the structure and dynamics of the nucleic acid assemblage concerned. A detailed understanding of the secondary structure is required for the appropriate placement of the photoexcitation of a sensitizer moiety (AQ) and for interpreting the resulting CFDC patterns. A high-resolution structure, however, is not an absolute requirement, but may aid in the interpretation and help deconvolute any ambiguous CFDC patterns. 1.2. Some key applications of CFQ Key examples of the utility of charge flow and quenching patterns (‘‘CFQ”) in complexly folded DNA structures is in the study of ‘‘deoxyribosensors” (Fig. 4A), structural variants of classic three-way junctions, which changes conformation upon binding specified ‘‘analytes” or ‘‘ligands”, such as adenosine or argininamide [16,20]. Figure 4B shows the secondary structure of a typical deoxyribosensor construct (‘‘ArgA1.3”), originally designed to sense argininamide binding [20]. Charge flow was initiated by photo-excitation of an AQ tethered to the end of the ‘‘AQ stem”, and charge flow into the ‘‘signal stem” could then be monitored by measurement of oxidative damage, leading to DNA cleavage, at a 50 -GGG-30 motif located within that arm. In testing different deoxyribosensor constructs (which varied only in the sequences
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Fig. 2. (A) Guanine oxidation pathway. Oxazalone is known to be piperidine liable. (B) 8-Oxoguanine is only modestly piperidine liable, and therefore needs to be further oxidized in order to form products that are piperidine liable. (C) Hot piperidine treatment will result in the b- and d-elimination (figure adapted from [12]).
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Fig. 4. Deoxyribosensors. (A) In the absence of the ligand (‘‘off” state), the aptamer (shown as a bulge) is unstructured and the top helix does not stack very efficiently with the bottom helix and charge flow can be seen only in the top helix. The binding of the ligand, L (the ‘‘on” state) causes the aptamer to fold such that the top helix stacks very efficiently with the bottom helix and charge flow can been observed in the bases of the bottom helix. (B) The study of the ArgA1.3 argininamide deoxyribosensor led to the idea of studying DNA structures by charge flow. The arrow shows the preferred charge flow path and the guanine marked with an asterisk was particularly prone to charge flow-dependent oxidative damage (figure adapted from [20].
Fig. 3. Duplex guanine oxidative damage by charge transfer (figure adapted from [31]). The ‘‘G Ladder” lane shows the Maxam–Gilbert sequence ladder for guanines in the end-labeled DNA strand. The ‘‘Duplex Reaction” lane shows the DNA sample incorporating an AQ residue, irradiated for charge conduction, while the ‘‘Negative Control” shows a DNA, lacking the AQ, but also irradiated. The samples in both the ‘‘Duplex Reaction” and ‘‘Negative Control” lanes were worked up with hot aqueous piperidine.
immediately bordering the junction itself), it was found that in some constructs charge flowed from the AQ stem mainly or exclusively into the signal stem, while in other constructs it flowed to the aptamer stem (Fig. 4B). Charge flow patterns could therefore, in principle, be used to monitor pair-wise helix-stacking preferences in the different deoxyribosensor constructs [20]. In the ArgA1.3 construct, CFDC appeared to localize mainly in the aptamer stem rather than the signal stem [20]. A notable finding was that a guanine located at the apex of the aptamer loop (shown with an asterisk in Fig. 4B) showed strong oxidative damage at all argininamide concentrations tested. A high-resolution NMR structure of the argininamide aptamer bound to its ligand, however, has shown this apical guanine to be both unstacked and jutting out into the solvent [21]. The CFDC at any DNA site is the product of two distinct processes: (i) the efficiency of charge transfer to and from the guanine at that site (the charge transfer rate) versus (ii) the efficiency of the water reaction with the G+ species generated at that site (the water reaction rate). In a DNA double helix, the rate of charge transfer, step (i), is typically many orders of magnitude faster than the rate of the water oxidation reaction, step (ii) [3]. Owing to a general uniformity of the groove geometries of the B-type DNA
Fig. 5. Secondary structures of the DNA constructs used to study charge flowdependent oxidative damage. (A) The 8–17 deoxyribozyme and a double-strand control duplex (B) used to study the charge-flow dependent oxidative damage. The deoxyribozyme (E1) is bound to a DNA pseudosubstrate (PS) and the arrow indicates the site of cleavage on an active substrate. AQ is tethered to the 50 end of PS and the boxed nucleotides in E1 have been reported to be important for catalysis. The CFDC of the bases in red are used for subsequent analysis (figure adapted from [30]).
helix, it is reasonable to assume that the rate of water reaction does not vary enormously at the different guanines within the helix. Therefore, the overall CFDC pattern across the full set of guanines in a double helix provides a ‘‘snapshot” of the equilibrium distribution of charge along the length of the helix. In a more complex case, however, where the locus of a given guanine is not identical to the standard intra-helical location within a B-type helix, the water reaction rate can differ significantly from those of typical B-helix guanines. Such is likely to be the cause of the high level of CFDC seen at the apical guanine of Arg1A.3 (above); owing to its extrahelical location and enhanced exposure to solvent, it likely implies this base there is a suboptimal step (i) i.e., the rate of formation and persistence of its radical cation is less efficient than within a helix; nevertheless, step (ii) for this base should be relatively
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efficient and fast, i.e., once formed, this highly solvent-exposed G+ should react efficiently with water. Is it possible to distinguish between the processes (i) and (ii)? As discussed above, the level of observed damage at a particular base within a complexly folded nucleic acid reflects a balance between the rate of charge flow into, and out of, that base relative to the rate of water reaction of the resulting radical cation. Given that, it should be possible to distinguish between highly oxidized guanines located within helices from those that are extrahelical and more significantly exposed to the solvent. It was speculated that enriching the solvent with reducing agents might help to quench the G+ species and, in turn, prevent or lower the rate of the water reaction leading to less oxidative damage. Such a reductive quenching might be more efficient at solvent-exposed guanines when compared intra-helical guanines. This notion was explored with some success with the small, RNA-cleaving deoxyribozyme, the 8–17. As shown in Fig. 5A, the 8–17, smallest of the RNA-cleaving deoxyribozymes, has a 13 nucleotide catalytic core. The core is composed of a short (3 base pair) double-stranded stem terminating in an invariant AGC terminal loop and a key unpaired region (4–5 nucleotides), whose sequence conforms to the sequence consensus WCGR (where W = A or T and R = G or A) or to WCGAA [22]. Systematic mutagenesis of the 8–17 catalytic core, by using natural as well as non-natural base analogues, has provided a detailed picture of the involvement of individual residues and functional groups within the 8–17 catalytic core in hydrogen bonding and catalysis [23]. These data, as well as more recent in vitro selection experiments that have yielded new, 8–17-related deoxyribozymes [24] have identified certain nucleotides in the catalytic core (shown as boxed within Fig. 5A) as being indispensable for catalysis. However, only a limited number of structural analyses have been done on the 8–17. Presently, only a relatively few detailed structural studies on the 8–17 have been carried out. Liu and Lu conducted fluorescence resonance energy transfer (‘‘FRET”) analysis on a trifluorophore-labeled 8–17 and studied the spatial distance changes between the three helices (signal, AQ and catalytic stems) at different divalent metal ion concentrations [25]. Detailed contact photo-crosslinking analysis of the 8–17 complexed to its substrate have revealed the identity of a number of nucleobases in close proximity with the substrate’s scissile site [26,27]. These include bases from both the bulge loop (C23, G24) and terminal loop (G17) of the 8–17’s secondary structure; and, the invariant thymine base of the DNAzyme that participates in a wobble-base pair with a G adjacent to the scissile site of the substrate. Leung and Sen (2007) carried out CFQ experiments on the 8–17 deoxyribozyme bound to a DNA pseudosubstrate (shown in Fig. 5A) under different experimental conditions. The following results were obtained: (a) measurement of CFQ under conditions of varying magnesium ion concentration revealed distinct stages of 8–17 folding; (b) the folded 8–17 incorporates three distinct helical stems, as shown in Fig. 5A. CFQ revealed the helix–helix stacking preferences among the three stems, something that is ordinarily difficult to determine directly without resorting to NMR or X-ray crystallography. CFQ enabled a quantitative assessment of the
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stacking preferences of the DNAzyme’s three constituent helices; and (c) some completely unexpected results were found, such as an unprecedented CFDC of a catalytically indespensable cytosine, C23 (cytosines are not expected to be oxidized from charge flow) within the folded 8–17 complex, indicating unusual conformational and/or electronic properties for this key residue. 1.3. The use of ascorbic acid to deconvolute charge flow and water reaction rates The hyper-reactivity of C23, above, was investigated further. Both C23 and its neighbour, G24 (Fig. 5A), showed strikingly different CFDC values as a function of solution magnesium concentration (i.e. the extent of 8–17 folding). As discussed above, the observed CFDC levels at a specific base within a complexly folded nucleic acid arises from a balance between the rate of charge flow in and out of that base relative to the rate of water reaction with the resulting radical cation. Given that, it should be possible to distinguish between highly oxidized guanines that are located within helices from those that are extrahelical and more significantly exposed to the solvent. Leung and Sen (2007) tested whether dissolved reducing agents might quench different G+ species differentially, by essentially attenuating the water reaction. The initial expectation was that such reductants may more efficiently block CFDC at solvent-exposed guanines than at intra-helical guanines, for instance. It was found that ascorbic acid (Fig. 6A), at 50– 100 lM concentrations, reduced CFDC most notably at C23 and G24. By contrast, G17, located in an apical loop, responded poorly to ascorbic acid, suggesting that although apical, this guanine was not heavily exposed to the solvent (also confirming earlier mutagenesis data [23]). It was speculated whether a hindered form of ascorbic acid, ascorbic acid-6-palmitate ([28]) would show an even finer discrimination than ascorbic acid itself; indeed, for both C23 and G24, the smaller ascorbic acid was more effective at CFDC quenching. 1.4. A charge flow-linked end-effect in RNA and DNA There is another striking method that is applicable to the study of charge transfer in both DNA and RNA. As seen above, the workup of charge transfer through DNA involves treatment at high temperatures with a relatively strong base, such as 1 M aqueous piperidine. Piperidine, or bases of comparable strength, would normally hydrolyze RNA indiscriminately. For chemical sequencing and footprinting of RNAs, typically a much milder treatment with acidified aniline is used instead [28]. In order to directly compare charge transfer through DNA duplexes versus RNA duplexes, it is necessary to create a standard protocol that is applicable to both. Bergeron et al. (2008) showed that acidified aniline could in fact be used to break down both DNA and RNA strands at oxidized guanosine residues [29]. However, a particularly interesting end-effect was observed by these authors, from both RNA and DNA duplexes. The finding was that if there is a stretch of guanines at the extreme end of a DNA or RNA helix through which charge has flowed, then treatment of such helices with aniline results in a large conduction-
Fig. 6. L-Ascorbic Acid and L-ascorbic acid-6-palmitate. The chemical structures shown are L-ascorbic acid (A) and L-ascorbic acid-6-palmitate (B).
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Fig. 7. Charge transfer through DNA and DNA/RNA duplexes. (A) A schematic model duplex where AQ indicates the covalently appended anthraquinone to a short DNA oligonucleotide. The predicted charge transfer detector sequence, GGG, is shown as CT detector. The electron flow direction is indicated by the arrow. (B) A denaturing gel showing the charge transfer reactions of DNA and RNA duplexes mentioned above (figure adapted from [29]).
related end-effect (Fig. 7, band G1a for DNA, and band G1 for RNA; the mobility difference between these bands is likely due to higher molecular weight of RNA compared to DNA). Overall, two striking conduction-related end-effect differences were noted between DNA and RNA: (a) with regard to the end-effect itself, DNA generated three products, G1a, G1b, and G1c; however, RNA generated only one product (product G1); (b) More generally, the RNA and DNA double helices showed different intra-helical CFDC patterns. While, DNA showed the characteristic CFDC seen when piperidine is used, instead of aniline, to generate the strand breaks, RNA showed very low levels of intra-helix CFDC. Unpublished data on RNAs containing bulges and mismatches, however, suggest that (a) the introduction of mismatches within the RNA double helix do modulate or eliminate the end-effect; (b) a GG bulge out of the intra-helical RNA stem does show notable CFDC at the bulge guanines (Bergeron and Sen, unpublished results). These latter data testify to the high sensitivity of this newly developed tool, and its potential for application to both complexly folded DNAs and RNAs. 2. Methodology 2.1. Setting up a CFQ experiment CFQ works best over a relatively short distance scales (10– 50 bp). It can be used profitably with RNA or DNA folds that incorporate helices as well as more complexly structured or indeed unstructured elements. Described below are the steps that need to be taken in order to carry out a successful CFQ experiment.
(1) One or more reference helices must be designated. In a given construct there will be one reference helix. This is the helix into which charge is initially injected, and from which the trajectory and efficiency of charge flow are to be monitored. Typically, this helix should be at least 10–15 bp long. Whether it is an RNA or a DNA helix that serves as a reference helix, the most convenient way that a charge-injecting anthraquinone (AQ) moiety can be attached to is by constructing a short all-DNA oligonucleotide, 10–12 nt long, which has a 50 -appended AQ (see below). This AQ-DNA can then be incorporated into the reference helix by simple Watson–Crick hybridization. This overall arrangement is illustrated in Figure 7A. The sequence immediately adjacent to the appended and end-stacked AQ (the ‘‘injection sequence”) is crucial for efficient charge injection into the helix. Typically this initial sequence needs to contain 4–5 consecutive A-T or A-U base pairs. A convenient injection sequence can be AQ-50 -TATA.... [29] or AQ-50 -TTTA... [30]. (2) For a successful CFQ probing experiment, the injected charge needs to have a sufficiently conductive path (DNA and RNA sequences that incorporate at least one G-C base pair within every three base pair) from the injection sequence to the one or more detector sequences located at one or more location distal from the injection sequence (for instance, the GGG sequence that is labeled as ‘‘CT Detector” in Fig. 7A). It may be that in an RNA or DNA of interest, the helix chosen to be the reference helix (or indeed, other helices of interest) incorporate extended stretches of A-T or A-U base pairs (which are poor conductors of charge). In such cases, minor mutagenesis will be required to substitute an adequate number of G-C base pairs, appropriately spaced, into the desired conductive path. (3) Suitable detectors of charge flow need to be designed into the overall DNA or RNA complex being investigated. These, as described above, should be placed in helical elements, in one or more location distal to the reference helix and the injection sequence. Typically, such detectors can be short, intra-helical stretches of contiguous guanines, e.g. 50 -GGG30 (such as shown in Fig. 7A). Or, such stretches of contiguous guanines can be placed at the very termini of helices– these will generate the strong charge-flow dependent end-effect, described above, and are applicable for both RNA and DNA. Other than the above, there are very few requirements for carrying out of a successful CFQ experiment, which remains a uniquely low-tech and easily carried out method of probing structure, folding, and dynamics. The details of each steps required to set up a CFQ experiment are given below. These protocols are adaptable; however, the details given should work for most experiments. 2.2. DNA purification Chemically synthesized DNA oligonucleotides are ideally purified by denaturing polyacrylamide gel electrophoresis, with the full-sized DNA band visualized by UV shadowing. The oligonucleotides should be excised from the gel and eluted via a simple crushsoak, followed by concentration using one or more successive ethanol precipitations. The concentrations of the oligonucleotides can be determined by measurement of UV absorption at 260 nm. Purified oligonucleotides can be stored dry or in solution in TE buffer (10 mM Tris, pH 7.5, 0.1–1.0 mM EDTA), at 20 °C. 2.3. C6-amino DNA synthesis and purification All C6-amino 50 modified DNAs are ideally pre-treated by dissolving in 10 mM Tris HCl (pH 7.0), 0.5 mM EDTA, and extraction
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with two volumes of chloroform, repeated three times, to remove possible nitrogenous contaminants from the DNA synthesis procedure. The aqueous fractions can then be ethanol precipitated and the recovered DNA pellet washed with cold 70% (v/v) ethanol. The purified DNA pellet is then dissolved in deionized water, and its concentration can be determined by UV absorbance at 260 nm. The purified oligonucleotides can be stored, as above, at 20 °C. 2.4. Synthesis of AQ-NHS ester and AQ Coupling of C6-Amino DNA Anthraquinone-2-carboxylic acid and N-hydroxysuccinimide (NHS) are dissolved in anhydrous dimethylformamide (DMF) and pre-cooled in an ice bath. 1,3-dicyclohexylcarbodiimide (DCC), dissolved in anhydrous DMF, is then added drop-wise with constant stirring. The solution is removed from the ice bath, covered with aluminium foil, and stirred at room temperature under a nitrogen atmosphere overnight. The resulting solution is filtered under vacuum using Whatman filter-paper to remove insoluble dicyclohexylurea, a key by-product of the reaction. The success of NHS ester production can be monitored by spotting the sample on a TLC silica F254 plate, with anthraquinone-2-carboxylic acid and NHS (each dissolved in anhydrous DMF) as standards, and ethyl acetate as the resolving solvent. The products on the TLC plate can be visualized by illumination with UV at 254 nm wavelength. The AQ-NHS ester can now be dried in a rotary evaporator at 40 °C and then re-dissolved in chloroform. The sample is filtered under vacuum using Whatman filter-paper and the remaining solution dried using nitrogen gas. The final dried AQ-NHS ester is best stored covered from light in a dry area at room temperature. For coupling of the AQ to amino-labeled DNA, the AQ-NHS ester is first dissolved in anhydrous DMF to make a 20 lg/lL stock solu-
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tion. The standard AQ coupling reaction (typically 100 lL) consists of the approximately 10–12 nmoles C6-amino DNA, 75 mM Na2B4O7, and 7 lL of the AQ stock solution. The solution is covered with aluminium foil to keep it dark, and is shaken vigorously overnight at room temperature. The solution is briefly spun-down and the supernatant transferred to a new microcentrifuge tube. The AQ-DNA is then ethanol precipitated with NaCl and anhydrous ethanol and washed with cold 70% (v/v) ethanol. The pellet is fully resuspended in 100 lM triethylammonium acetate (TEAA) (pH 6.85) and an equal volume of chloroform. The two-phase solvent system is used to efficiently dissolve and separate the AQ-DNA from the un-coupled AQ. The aqueous layer is extracted two times with equal volumes of chloroform, lyophilized, and resuspended in 100 lM TEAA (pH 6.85). Detailed description of the synthesis and purification of the AQNHS ester can be found in Fahlman and Sen, 2002 [16], Sankar and Sen, 2004 [20], and Leung and Sen, 2007 [27]. 2.5. HPLC purification of AQ-coupled DNA The AQ-coupled DNA is purified by high-pressure liquid chromatography using a reverse-phase C-18 column. The solvent flow is set continuously at 1 ml/minute and the solvents are heated to 40 °C. The initial conditions are: 100% solvent A (20:1 100 lM TEAA (pH 6.85): acetonitrile), changing to 30% solvent B (100% acetonitrile) over 30 min with a linear gradient. Typically, un-coupled DNA has a retention time of approximately 10 min, while the AQcoupled DNA (‘‘AQ-DNA”) has a retention time of approximately 17 min (Fig. 8). Retention times will naturally vary for different DNA oligonucleotides, depending on the length of the DNA. To ensure that the collected fractions are indeed AQ-DNA the ratio is calculated of the area of the 260 nm absorbance peak relative to the
Fig. 8. (A) Anthraquinone coupled to DNA. (B) HPLC traces of anthraquinone coupled and un-coupled DNA. The exact retention times of the DNA oligonucleotides will vary depending on its length, but in all cases, the un-coupled DNA will elute from the reverse-phase HPLC column before the coupled DNA. DNA absorbs only at 260 nm (B) and AQ absorbs at both 260 and 335 nm (C), therefore the peak at approximately 12 min is the un-coupled DNA and the peak at approximately 22 min is the DNA coupled to AQ. The final broad peak at 32 min is the un-coupled AQ.
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335 nm absorbance peak. AQ-DNA oligonucleotides have a ratios ranging from 20–50 depending on the length of the DNA [16]. The fractions containing the AQ-DNA can then be lyophilized and re-dissolved in deinoized water. The AQ-DNA concentration can be determined by measurement of UV absorption at 260 nm. A typical yield of AQ-DNA conjugates ranges from 50–90% depending on the sequence and the synthesis batch. 2.6. RNA synthesis It is convenient to synthesize RNA strands using T7 RNA polymerase in vitro, with a double-stranded template that incorporates the T7 RNA polymerase promoter. The DNA template is typically amplified and rendered double-stranded using the polymerase chain reaction (PCR) using Vent or Taq DNA polymerase, under standard conditions. Transcription reactions can then be carried out using purified T7 RNA polymerase (30 lg) and the DNA template (2–5 lM) in 100 ll of transcription buffer (40 mM Tris–HCl, pH 7.9, 26 mM MgCl2, 2.5 mM spermidine, 10 mM DTT, 0.01% Triton X-100, 8 lM GTP, 4 lM ATP and CTP, 2 lM UTP). The reaction typically proceeds for 2–16 h at 37 °C. Upon completion, the transcription mixtures are treated with deoxyribonuclease I for 30 min at 37 °C, and the newly synthesized RNAs ethanol precipitated and size-purified by denaturing polyacrylamide gel electrophoresis. Dephosphorylation (for RNA) using calf intestinal alkaline phosphatase and 32P-end labeling (for DNA and RNA) using polynucleotide kinase can be carried out using standard protocols. 2.7. Preparation of assemblies DNA or RNA assemblies can be formed by annealing stoichiometric mixtures of all constituent oligonucleotides, including the AQDNA (1 lM each) and trace amounts of 50 -end labeled oligonucleotide, in 50 mM Tris–HCl (pH 7.5) and 0.1 mM EDTA. The solutions are heated at 90 °C for 30 s, cooled slowly to room temperature (60–90 min), then diluted 4-fold in irradiation buffer (final concentration: 50 mM Tris–HCl (pH 7.5), 50 mM NaCl, 1 mM MgCl2, and 0.1 mM EDTA; or any other desired solution conditions). The samples are incubated for approximately 15 min at room temperature prior to photo-irradiation to initiate charge flow. 2.8. Charge-flow inducing irradiation of the DNA or RNA complexes DNA/RNA solutions are placed in the wells of a 96-well polycarbonate or polyethylene plate pre-coated with glycogen (1 mg/ml) and placed under a Blak-Ray UVL-56 lamp (365 nm) for 60 min at a distance of 2 cm from the bulb. Temperature can be maintained by having the plate in contact with a shallow water bath. Following photo-irradiation, the samples are ethanol precipitated with the assistance of glycogen as carrier. 2.9. Piperidine treatment and denaturing gel electrophoresis The DNA pellets are dissolved in 10% (v/v) piperidine and incubated at 90 °C for 30 min. The treated DNA is lyophilized, dissolved in denaturing gel loading dye, heat denatured at 100 °C for 5 min, cooled to 22 °C, and loaded on denaturing polyacrylamide sequencing gels. The gels can be analyzed by phosphorimagery. 2.10. Aniline treatment Ethanol-precipitated DNA/RNA pellet is dissolved directly into an aniline acetate solution (20 lL aniline, 180 lL H2O, and 12 lL of glacial acetic acid) and strand scission is preformed at 60 °C for 15 min in the dark. Each sample is diluted with 80 lL of doubly
deionized water and is dried under vacuum for 1 h in a lyophilizer. The gels can be analyzed by phosphorimagery. 2.11. Quenching experiments with ascorbic acid DNA/RNA solutions are prepared as previously mentioned. Lascorbic acid or ascorbic acid-6-palmitate is added to a final concentration of 50 lM from stock solutions to the appropriate DNA solution just prior to irradiation. The L-ascorbic acid stock can be prepared with deinoized water and the ascorbic acid-6-palmitate stock solution can be prepared in dimethyl sulfoxide (DMSO) due to its low solubility in deinoized water. To equalize DMSO in all reactions to be irradiated, both the 0 and 50 lM L-ascorbic acid containing reactions are made up to a final DMSO concentration of 0.185% (v/v) to match the final concentration of DMSO that will be present in the ascorbic acid-6-palmitate-containing reactions. 2.12. Data analysis The densitometry analyses of the sequencing gels are carried out and the density of the band of interest was assessed as a percentage of the total signal contained within a particular lane. This procedure will compensate for any discrepancies of the total radioactive counts in different lanes. The normalized signal for a particular nucleotide obtained in the ‘‘dark” reaction (negative control) lane is subtracted from the signal of the same nucleotide in an irradiated sample lane. This corrected signal for a particular nucleotide is then divided by the corrected signal of the reference nucleotide to give the damage ratio. Any single intra-helical guanine can be used as the reference nucleotide as long as its CFDC is relative constant throughout all experimental conditions tested. 3. Concluding remarks CFQ is a new and versatile low-tech method for probing various structural parameters within complexly folded DNAs and RNAs. Future years will undoubtedly bring further innovations to the method; however, even in its present state, it is easily practised, and should provide practitioners with novel kinds of structural and folding information that is difficult to obtain without resorting to highresolution structural studies involving X-ray crystallography or NMR. One small innovation that might contribute to the ease of planning CFQ experiments is the synthesis and ready availability of an anthraquinone phosphoramidite for automated DNA/RNA synthesis. It is our hope that this challenge will be taken up by one or more of the larger commercial suppliers of phosphoramidites. References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] [16] [17] [18] [19]
E.M. Boon, J.K. Barton, Curr. Opin. Struct. Biol. 12 (2002) 320–329. B. Giese, Annu. Rev. Biochem. 71 (2002) 51–70. G.B. Schuster, Acc. Chem. Res. 33 (2000) 253–260. M.E. Nunez, D.B. Hall, J.K. Barton, Chem. Biol. 6 (1999) 85–97. P.T. Henderson, D. Jones, G. Hampikian, Y. Kan, G.B. Schuster, Proc. Natl Acad. Sci. USA 96 (1999) 8353–8358. A. Joy, G.B. Schuster, Chem. Commun. (Camb.) (2005) 2778–2784. H.A. Wagenknecht, Nat. Prod. Rep. 23 (2006) 973–1006. S. Priyadarshy, S.M. Risser, D.N. Beratan, J. Phys. Chem. 100 (1996) 17678– 17682. C.A.M. Seidel, A. Schulz, M.H.M. Sauer, J. Phys. Chem. 100 (1996) 5541–5553. H. Sugiyama, I. Saito, J. Am. Chem. Soc. 118 (1996) 7063–7068. M.A. O’Neill, J.K. Barton, J. Am. Chem. Soc. 126 (2004) 11471–11483. C.J. Burrows, J.G. Muller, Chem. Rev. 98 (1998) 1109–1152. I. Saito, M. Takayama, H. Sugiyama, K. Nakatani, A. Tsuchida, M. Yamamoto, J. Am. Chem. Soc. 117 (1995) 6406–6407. P.K. Bhattacharya, J.K. Barton, J. Am. Chem. Soc. 123 (2001) 8649–8656. S.O. Kelley, J.K. Barton, Science 283 (1999) 375–381. R.P. Fahlman, D. Sen, J. Am. Chem. Soc. 124 (2002) 4610–4616. R.P. Fahlman, R.D. Sharma, D. Sen, J. Am. Chem. Soc. 124 (2002) 12477–12485. D.T. Odom, E.A. Dill, J.K. Barton, Nucleic Acids Res. 29 (2001) 2026–2033. U. Santhosh, G.B. Schuster, Nucleic Acids Res. 31 (2003) 5692–5699.
E.K.Y. Leung, D. Sen / Methods 52 (2010) 141–149 [20] C.G. Sankar, D. Sen, J. Mol. Biol. 340 (2004) 459–467. [21] S.A. Robertson, K. Harada, A.D. Frankel, D.E. Wemmer, Biochemistry 39 (2000) 946–954. [22] S.W. Santoro, G.F. Joyce, Proc. Natl Acad. Sci. USA 94 (1997) 4262–4266. [23] A. Peracchi, M. Bonaccio, M. Clerici, J. Mol. Biol. 352 (2005) 783–794. [24] R.P. Cruz, J.B. Withers, Y. Li, Chem. Biol. 11 (2004) 57–67. [25] J. Liu, Y. Lu, J. Am. Chem. Soc. 124 (2002) 15208–15216. [26] Y. Liu, D. Sen, J. Mol. Biol. 381 (2008) 845–859.
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Methods journal homepage: www.elsevier.com/locate/ymeth
Review Article
The use of charge flow and quenching (CFQ) to probe nucleic acid folds and folding Edward K.Y. Leung a, Dipankar Sen b,* a b
Department of Biochemistry & Molecular Biology, The University of Chicago, Chicago, IL 60637, USA Department of Molecular Biology & Biochemistry, Simon Fraser University, Burnaby, British Columbia, Canada V5A 1S6
a r t i c l e
i n f o
Article history: Available online 8 June 2010
a b s t r a c t Charge flow and quenching (‘‘CFQ”) is a relatively new, versatile, and easily carried out methodology for probing a number of unique features of DNA and RNA folded structures, and of their folding pathways. An electrical charge (an electron hole, or radical cation) is injected site-specifically into the end of a predetermined reference helix within the larger DNA or RNA structure. The fate of the injected charge, as it percolates through the folded DNA or RNA is then monitored by mapping the oxidative consequences of the charge flow. Some of the kinds of structural and folding information that can be obtained from CFQ experiments include: a quantitative measure of helix–helix connectivity; the dynamics of specific bases; folding and unfolding pathways; the mapping of unusual, conformation-dependent, electronic properties of individual bases; extents of solvent exposure and susceptibility to quenching from the solvent. CFQ is a relatively new methodology, and is applicable to DNA and RNA structures and folds. In the near future it is expected that the range of applications of this methodology will increase dramatically. Ó 2010 Elsevier Inc. All rights reserved.
1. Introduction 1.1. Background Over the past two decades, intensive research has shown that DNA double helices, in aqueous solution, are able to conduct electrical charge (reviewed in [1–3]). Two classes of charge conduction have been studied: (1) a class in which the charge carrier is a nucleobase radical cation (an ‘‘electron hole”); and (2) a class in which it is a radical anion (an ‘‘excess electron”). On the whole, hole transfer has been studied in greater depth, and the term ‘‘charge conduction” will refer exclusively to hole transfer for the remainder of this paper. Charge flow in DNA can conveniently be initiated by photo-excitation of a sensitizer moiety, such as anthraquinone (AQ), covalently linked and p-stacked upon the end of a double helix. The base radical cation generated within the helix has been found to propagate along the helix over relatively long distances (at least 200 Å) [4–6]. Three possible mechanisms for photo-induced charge transfer through DNA were originally proposed: (1) molecular wire; (2) super-exchange; and (3) charge hopping (reviewed in [7]). In the molecular wire model, charge is injected from the donor to a bridge element, where it localizes prior to moving incoherently towards the acceptor (Fig. 1A). In the super-exchange mechanism, the charge does not localize on the bridge element, but tunnels from
* Corresponding author. Fax: +1 778 782 5583. E-mail addresses: [email protected] (E.K.Y. Leung), [email protected] (D. Sen). 1046-2023/$ - see front matter Ó 2010 Elsevier Inc. All rights reserved. doi:10.1016/j.ymeth.2010.06.006
the donor directly to the acceptor (Fig. 1B). The hopping model, by contrast, occurs via the hopping of charge in a thermally activated fashion, from guanine base to guanine base (Fig. 1C). Experimental data indicate that DNA charge transfer does not follow the molecular wire mechanism, because the strong distance dependence of charge migration through DNA does not behave like a molecular wire [8]. The super-exchange mechanism occurs only efficiently over short distances (<10 Å); however, experiments have found charge transfer in DNA up to 200 Å [4–6]. Therefore, the most plausible mechanism is that of charge hopping. Of the four DNA bases, guanine (G) is the most easily oxidized (Table 1) and the guanine radical cation, G+, is the most stable of the four possible radical cations [9]. G+ is the intermediate charge carrier in the hopping process, and adjacent Gs (for example 50 -GG-30 or 50 -GGG-30 ) provide a particularly good thermodynamic sink for the charge. Theoretical studies have shown the radical cation on the 50 guanines of sequences such as 50 -GG-30 or 50 -GGG-30 are particularly stabilized relative to guanines towards the 30 end (Table 2), but not vice versa [10]. A recent refinement of the charge hopping model, the ‘‘phononassisted polaron-like hopping” model, most successfully accounts for hole conduction [5,6,11]. In this model, the charge is delocalized over several DNA bases due to local structural distortions of the DNA helix. The base-pairing domain within the DNA base stack can act as a gate for DNA-mediated charge transfer. In other words, the charge transfer occurs as a hopping process between conformationally gated domains of well-stacked base pairs (reviewed in [7]). These delocalized domains may transiently form and break, and facilitate or inhibit charge transfer.
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Fig. 1. Mechanisms of charge transfer in DNA. (A) The induced charge is delocalized throughout the entire molecule in the molecular wire model. (B) The charge jumps directly from the charge donor to the charge acceptor in the super-exchange model. (C) The charge, in the form of a radical cation, migrates from guanine to guanine until it reaches the acceptor in the hole hopping model. The charge can tunnel through a stretch of consecutive adenines. (D) The charge is delocalized over several base pairs in the phonon-assisted polaron-like hopping model and propagates along the helix.
Table 1 Oxidation potential for nucleosides. The first five oxidation potentials were measured in (a) aprotic solvent (acetonitrile) conditions [9]. The remaining oxidative potentials were determined in protic (aqueous) solvents at (b) pH 7.0 [32] and (c) pH 8.0 [33]. The oxidation potentials of the bases are dependent on the solvent (aprotic versus protic) conditions, and therefore, hydrogen bonds will change the oxidation potentials of all nucleobases. Base
E0(V vs NHE)a
Guanine Adenine Cytosine Thymine Uracil rG rA dC dT r(8-oxoG) r(8-oxoA) r(5-OH-C) r(5-OH-U)
1.49 1.96 2.14 2.11 P2.39
E0(V vs NHE)b
E0(V vs NHE)c
pH
0.58 0.92 0.62 0.64
7 7 7 7 8 8 8 8
1.29 1.42 1.60 1.70
Table 2 Calculated oxidation potential for bases. (a) The oxidation potential of the 5’-most guanine is influenced by it adjacent bases [13]. (b) The oxidation potential of 8oxoguanine can also be influenced by it neighbouring bases [34]. Sequence
E0(V vs NHE)a
GGG GG GA GC GT G 8-oxoG (8-oxoG)G G(8-oxoG)
0.64 0.84 1.00 1.15 1.16 1.20
E0(V vs NHE)b
0.85 0.53 0.08 0.18
Any guanine that hosts the mobile radical cation transiently is susceptible to a side reaction with water (or, in some instances,
dissolved oxygen), leading to the formation of modest levels of such guanine oxidation products as 8-oxoguanine or diaminooxazalone (reviewed in [12]). The formation of these oxidation products (Fig. 2A, B, adapted from [12]) supplies a useful biochemical handle for tracing both the actual charge flow path though DNA, and its consequences. These are possible because treatment with hot piperidine (or other comparable aqueous base) breaks the DNA strand at sites containing the guanine oxidation products (Fig. 2C). The susceptibility of any specific guanine within a double helix to charge flow-related oxidation depends, in degrees, on the sequence context of the guanine [2–4]. Stretches of multiple guanines, such as 50 -GG-30 or 50 -GGG-30 , are particularly susceptible to oxidation, with the 50 -most guanine of the stretch the most extensively oxidized [2–4,12]. Densitometry trace of charge flowdependent DNA cleavage bands (‘‘CFDC”), separated by gel electrophoresis provides a convenient and quantitative means for mapping charge flow through double helices. The sequencing gel shown in Fig. 3 illustrates the patterns of CFDC in a typical DNA double helix. While the guanine ladder on the left shows more or less equivalent guanine band intensities for all the guanines within the 50 -32P-labeled strand, the guanine band intensities in the charge flow lane (‘‘Duplex reaction”) indicate how patterns of CFDC vary markedly (and predictably) at different guanines, depending on their location. Thus, isolated single guanines typically show low levels of CFDC, while the 50 -most guanine of a 50 -GG-30 stretch (for instance the ‘‘proximal” and ‘‘distal” guanine doublets) show guanine oxidation (and cleavage) heavily biased towards the 50 -most guanine. In a more extensive stretch of guanines (such as 50 -GGG-30 or the 50 -GGGGG-30 stretch shown in Fig. 3) the pattern of CFDC is more complex, although largely predictable [13]. Experimentally, it is not a surprise to find that any interruption to or deviation from the standard geometry and base p-stacking patterns found within a Watson–Crick base-paired double helix impacts on the efficiency of charge conduction. Where a p-stack is perturbed (such as with mismatches or bulges), charge migration efficiency diminishes notably [14,15]. Therefore, in principle, CFDC provides a simple measure of charge transfer through intact DNA helices as well as assemblages made of helices and other structural components, including but not restricted to DNA three-way and four-way helical junctions [16–19]. Such charge transfer patterns (and, in special cases, patterns of reductant-mediated quenching of charge flow) are able, in turn, to provide information on the structure and dynamics of the nucleic acid assemblage concerned. A detailed understanding of the secondary structure is required for the appropriate placement of the photoexcitation of a sensitizer moiety (AQ) and for interpreting the resulting CFDC patterns. A high-resolution structure, however, is not an absolute requirement, but may aid in the interpretation and help deconvolute any ambiguous CFDC patterns. 1.2. Some key applications of CFQ Key examples of the utility of charge flow and quenching patterns (‘‘CFQ”) in complexly folded DNA structures is in the study of ‘‘deoxyribosensors” (Fig. 4A), structural variants of classic three-way junctions, which changes conformation upon binding specified ‘‘analytes” or ‘‘ligands”, such as adenosine or argininamide [16,20]. Figure 4B shows the secondary structure of a typical deoxyribosensor construct (‘‘ArgA1.3”), originally designed to sense argininamide binding [20]. Charge flow was initiated by photo-excitation of an AQ tethered to the end of the ‘‘AQ stem”, and charge flow into the ‘‘signal stem” could then be monitored by measurement of oxidative damage, leading to DNA cleavage, at a 50 -GGG-30 motif located within that arm. In testing different deoxyribosensor constructs (which varied only in the sequences
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Fig. 2. (A) Guanine oxidation pathway. Oxazalone is known to be piperidine liable. (B) 8-Oxoguanine is only modestly piperidine liable, and therefore needs to be further oxidized in order to form products that are piperidine liable. (C) Hot piperidine treatment will result in the b- and d-elimination (figure adapted from [12]).
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Fig. 4. Deoxyribosensors. (A) In the absence of the ligand (‘‘off” state), the aptamer (shown as a bulge) is unstructured and the top helix does not stack very efficiently with the bottom helix and charge flow can be seen only in the top helix. The binding of the ligand, L (the ‘‘on” state) causes the aptamer to fold such that the top helix stacks very efficiently with the bottom helix and charge flow can been observed in the bases of the bottom helix. (B) The study of the ArgA1.3 argininamide deoxyribosensor led to the idea of studying DNA structures by charge flow. The arrow shows the preferred charge flow path and the guanine marked with an asterisk was particularly prone to charge flow-dependent oxidative damage (figure adapted from [20].
Fig. 3. Duplex guanine oxidative damage by charge transfer (figure adapted from [31]). The ‘‘G Ladder” lane shows the Maxam–Gilbert sequence ladder for guanines in the end-labeled DNA strand. The ‘‘Duplex Reaction” lane shows the DNA sample incorporating an AQ residue, irradiated for charge conduction, while the ‘‘Negative Control” shows a DNA, lacking the AQ, but also irradiated. The samples in both the ‘‘Duplex Reaction” and ‘‘Negative Control” lanes were worked up with hot aqueous piperidine.
immediately bordering the junction itself), it was found that in some constructs charge flowed from the AQ stem mainly or exclusively into the signal stem, while in other constructs it flowed to the aptamer stem (Fig. 4B). Charge flow patterns could therefore, in principle, be used to monitor pair-wise helix-stacking preferences in the different deoxyribosensor constructs [20]. In the ArgA1.3 construct, CFDC appeared to localize mainly in the aptamer stem rather than the signal stem [20]. A notable finding was that a guanine located at the apex of the aptamer loop (shown with an asterisk in Fig. 4B) showed strong oxidative damage at all argininamide concentrations tested. A high-resolution NMR structure of the argininamide aptamer bound to its ligand, however, has shown this apical guanine to be both unstacked and jutting out into the solvent [21]. The CFDC at any DNA site is the product of two distinct processes: (i) the efficiency of charge transfer to and from the guanine at that site (the charge transfer rate) versus (ii) the efficiency of the water reaction with the G+ species generated at that site (the water reaction rate). In a DNA double helix, the rate of charge transfer, step (i), is typically many orders of magnitude faster than the rate of the water oxidation reaction, step (ii) [3]. Owing to a general uniformity of the groove geometries of the B-type DNA
Fig. 5. Secondary structures of the DNA constructs used to study charge flowdependent oxidative damage. (A) The 8–17 deoxyribozyme and a double-strand control duplex (B) used to study the charge-flow dependent oxidative damage. The deoxyribozyme (E1) is bound to a DNA pseudosubstrate (PS) and the arrow indicates the site of cleavage on an active substrate. AQ is tethered to the 50 end of PS and the boxed nucleotides in E1 have been reported to be important for catalysis. The CFDC of the bases in red are used for subsequent analysis (figure adapted from [30]).
helix, it is reasonable to assume that the rate of water reaction does not vary enormously at the different guanines within the helix. Therefore, the overall CFDC pattern across the full set of guanines in a double helix provides a ‘‘snapshot” of the equilibrium distribution of charge along the length of the helix. In a more complex case, however, where the locus of a given guanine is not identical to the standard intra-helical location within a B-type helix, the water reaction rate can differ significantly from those of typical B-helix guanines. Such is likely to be the cause of the high level of CFDC seen at the apical guanine of Arg1A.3 (above); owing to its extrahelical location and enhanced exposure to solvent, it likely implies this base there is a suboptimal step (i) i.e., the rate of formation and persistence of its radical cation is less efficient than within a helix; nevertheless, step (ii) for this base should be relatively
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efficient and fast, i.e., once formed, this highly solvent-exposed G+ should react efficiently with water. Is it possible to distinguish between the processes (i) and (ii)? As discussed above, the level of observed damage at a particular base within a complexly folded nucleic acid reflects a balance between the rate of charge flow into, and out of, that base relative to the rate of water reaction of the resulting radical cation. Given that, it should be possible to distinguish between highly oxidized guanines located within helices from those that are extrahelical and more significantly exposed to the solvent. It was speculated that enriching the solvent with reducing agents might help to quench the G+ species and, in turn, prevent or lower the rate of the water reaction leading to less oxidative damage. Such a reductive quenching might be more efficient at solvent-exposed guanines when compared intra-helical guanines. This notion was explored with some success with the small, RNA-cleaving deoxyribozyme, the 8–17. As shown in Fig. 5A, the 8–17, smallest of the RNA-cleaving deoxyribozymes, has a 13 nucleotide catalytic core. The core is composed of a short (3 base pair) double-stranded stem terminating in an invariant AGC terminal loop and a key unpaired region (4–5 nucleotides), whose sequence conforms to the sequence consensus WCGR (where W = A or T and R = G or A) or to WCGAA [22]. Systematic mutagenesis of the 8–17 catalytic core, by using natural as well as non-natural base analogues, has provided a detailed picture of the involvement of individual residues and functional groups within the 8–17 catalytic core in hydrogen bonding and catalysis [23]. These data, as well as more recent in vitro selection experiments that have yielded new, 8–17-related deoxyribozymes [24] have identified certain nucleotides in the catalytic core (shown as boxed within Fig. 5A) as being indispensable for catalysis. However, only a limited number of structural analyses have been done on the 8–17. Presently, only a relatively few detailed structural studies on the 8–17 have been carried out. Liu and Lu conducted fluorescence resonance energy transfer (‘‘FRET”) analysis on a trifluorophore-labeled 8–17 and studied the spatial distance changes between the three helices (signal, AQ and catalytic stems) at different divalent metal ion concentrations [25]. Detailed contact photo-crosslinking analysis of the 8–17 complexed to its substrate have revealed the identity of a number of nucleobases in close proximity with the substrate’s scissile site [26,27]. These include bases from both the bulge loop (C23, G24) and terminal loop (G17) of the 8–17’s secondary structure; and, the invariant thymine base of the DNAzyme that participates in a wobble-base pair with a G adjacent to the scissile site of the substrate. Leung and Sen (2007) carried out CFQ experiments on the 8–17 deoxyribozyme bound to a DNA pseudosubstrate (shown in Fig. 5A) under different experimental conditions. The following results were obtained: (a) measurement of CFQ under conditions of varying magnesium ion concentration revealed distinct stages of 8–17 folding; (b) the folded 8–17 incorporates three distinct helical stems, as shown in Fig. 5A. CFQ revealed the helix–helix stacking preferences among the three stems, something that is ordinarily difficult to determine directly without resorting to NMR or X-ray crystallography. CFQ enabled a quantitative assessment of the
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stacking preferences of the DNAzyme’s three constituent helices; and (c) some completely unexpected results were found, such as an unprecedented CFDC of a catalytically indespensable cytosine, C23 (cytosines are not expected to be oxidized from charge flow) within the folded 8–17 complex, indicating unusual conformational and/or electronic properties for this key residue. 1.3. The use of ascorbic acid to deconvolute charge flow and water reaction rates The hyper-reactivity of C23, above, was investigated further. Both C23 and its neighbour, G24 (Fig. 5A), showed strikingly different CFDC values as a function of solution magnesium concentration (i.e. the extent of 8–17 folding). As discussed above, the observed CFDC levels at a specific base within a complexly folded nucleic acid arises from a balance between the rate of charge flow in and out of that base relative to the rate of water reaction with the resulting radical cation. Given that, it should be possible to distinguish between highly oxidized guanines that are located within helices from those that are extrahelical and more significantly exposed to the solvent. Leung and Sen (2007) tested whether dissolved reducing agents might quench different G+ species differentially, by essentially attenuating the water reaction. The initial expectation was that such reductants may more efficiently block CFDC at solvent-exposed guanines than at intra-helical guanines, for instance. It was found that ascorbic acid (Fig. 6A), at 50– 100 lM concentrations, reduced CFDC most notably at C23 and G24. By contrast, G17, located in an apical loop, responded poorly to ascorbic acid, suggesting that although apical, this guanine was not heavily exposed to the solvent (also confirming earlier mutagenesis data [23]). It was speculated whether a hindered form of ascorbic acid, ascorbic acid-6-palmitate ([28]) would show an even finer discrimination than ascorbic acid itself; indeed, for both C23 and G24, the smaller ascorbic acid was more effective at CFDC quenching. 1.4. A charge flow-linked end-effect in RNA and DNA There is another striking method that is applicable to the study of charge transfer in both DNA and RNA. As seen above, the workup of charge transfer through DNA involves treatment at high temperatures with a relatively strong base, such as 1 M aqueous piperidine. Piperidine, or bases of comparable strength, would normally hydrolyze RNA indiscriminately. For chemical sequencing and footprinting of RNAs, typically a much milder treatment with acidified aniline is used instead [28]. In order to directly compare charge transfer through DNA duplexes versus RNA duplexes, it is necessary to create a standard protocol that is applicable to both. Bergeron et al. (2008) showed that acidified aniline could in fact be used to break down both DNA and RNA strands at oxidized guanosine residues [29]. However, a particularly interesting end-effect was observed by these authors, from both RNA and DNA duplexes. The finding was that if there is a stretch of guanines at the extreme end of a DNA or RNA helix through which charge has flowed, then treatment of such helices with aniline results in a large conduction-
Fig. 6. L-Ascorbic Acid and L-ascorbic acid-6-palmitate. The chemical structures shown are L-ascorbic acid (A) and L-ascorbic acid-6-palmitate (B).
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Fig. 7. Charge transfer through DNA and DNA/RNA duplexes. (A) A schematic model duplex where AQ indicates the covalently appended anthraquinone to a short DNA oligonucleotide. The predicted charge transfer detector sequence, GGG, is shown as CT detector. The electron flow direction is indicated by the arrow. (B) A denaturing gel showing the charge transfer reactions of DNA and RNA duplexes mentioned above (figure adapted from [29]).
related end-effect (Fig. 7, band G1a for DNA, and band G1 for RNA; the mobility difference between these bands is likely due to higher molecular weight of RNA compared to DNA). Overall, two striking conduction-related end-effect differences were noted between DNA and RNA: (a) with regard to the end-effect itself, DNA generated three products, G1a, G1b, and G1c; however, RNA generated only one product (product G1); (b) More generally, the RNA and DNA double helices showed different intra-helical CFDC patterns. While, DNA showed the characteristic CFDC seen when piperidine is used, instead of aniline, to generate the strand breaks, RNA showed very low levels of intra-helix CFDC. Unpublished data on RNAs containing bulges and mismatches, however, suggest that (a) the introduction of mismatches within the RNA double helix do modulate or eliminate the end-effect; (b) a GG bulge out of the intra-helical RNA stem does show notable CFDC at the bulge guanines (Bergeron and Sen, unpublished results). These latter data testify to the high sensitivity of this newly developed tool, and its potential for application to both complexly folded DNAs and RNAs. 2. Methodology 2.1. Setting up a CFQ experiment CFQ works best over a relatively short distance scales (10– 50 bp). It can be used profitably with RNA or DNA folds that incorporate helices as well as more complexly structured or indeed unstructured elements. Described below are the steps that need to be taken in order to carry out a successful CFQ experiment.
(1) One or more reference helices must be designated. In a given construct there will be one reference helix. This is the helix into which charge is initially injected, and from which the trajectory and efficiency of charge flow are to be monitored. Typically, this helix should be at least 10–15 bp long. Whether it is an RNA or a DNA helix that serves as a reference helix, the most convenient way that a charge-injecting anthraquinone (AQ) moiety can be attached to is by constructing a short all-DNA oligonucleotide, 10–12 nt long, which has a 50 -appended AQ (see below). This AQ-DNA can then be incorporated into the reference helix by simple Watson–Crick hybridization. This overall arrangement is illustrated in Figure 7A. The sequence immediately adjacent to the appended and end-stacked AQ (the ‘‘injection sequence”) is crucial for efficient charge injection into the helix. Typically this initial sequence needs to contain 4–5 consecutive A-T or A-U base pairs. A convenient injection sequence can be AQ-50 -TATA.... [29] or AQ-50 -TTTA... [30]. (2) For a successful CFQ probing experiment, the injected charge needs to have a sufficiently conductive path (DNA and RNA sequences that incorporate at least one G-C base pair within every three base pair) from the injection sequence to the one or more detector sequences located at one or more location distal from the injection sequence (for instance, the GGG sequence that is labeled as ‘‘CT Detector” in Fig. 7A). It may be that in an RNA or DNA of interest, the helix chosen to be the reference helix (or indeed, other helices of interest) incorporate extended stretches of A-T or A-U base pairs (which are poor conductors of charge). In such cases, minor mutagenesis will be required to substitute an adequate number of G-C base pairs, appropriately spaced, into the desired conductive path. (3) Suitable detectors of charge flow need to be designed into the overall DNA or RNA complex being investigated. These, as described above, should be placed in helical elements, in one or more location distal to the reference helix and the injection sequence. Typically, such detectors can be short, intra-helical stretches of contiguous guanines, e.g. 50 -GGG30 (such as shown in Fig. 7A). Or, such stretches of contiguous guanines can be placed at the very termini of helices– these will generate the strong charge-flow dependent end-effect, described above, and are applicable for both RNA and DNA. Other than the above, there are very few requirements for carrying out of a successful CFQ experiment, which remains a uniquely low-tech and easily carried out method of probing structure, folding, and dynamics. The details of each steps required to set up a CFQ experiment are given below. These protocols are adaptable; however, the details given should work for most experiments. 2.2. DNA purification Chemically synthesized DNA oligonucleotides are ideally purified by denaturing polyacrylamide gel electrophoresis, with the full-sized DNA band visualized by UV shadowing. The oligonucleotides should be excised from the gel and eluted via a simple crushsoak, followed by concentration using one or more successive ethanol precipitations. The concentrations of the oligonucleotides can be determined by measurement of UV absorption at 260 nm. Purified oligonucleotides can be stored dry or in solution in TE buffer (10 mM Tris, pH 7.5, 0.1–1.0 mM EDTA), at 20 °C. 2.3. C6-amino DNA synthesis and purification All C6-amino 50 modified DNAs are ideally pre-treated by dissolving in 10 mM Tris HCl (pH 7.0), 0.5 mM EDTA, and extraction
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with two volumes of chloroform, repeated three times, to remove possible nitrogenous contaminants from the DNA synthesis procedure. The aqueous fractions can then be ethanol precipitated and the recovered DNA pellet washed with cold 70% (v/v) ethanol. The purified DNA pellet is then dissolved in deionized water, and its concentration can be determined by UV absorbance at 260 nm. The purified oligonucleotides can be stored, as above, at 20 °C. 2.4. Synthesis of AQ-NHS ester and AQ Coupling of C6-Amino DNA Anthraquinone-2-carboxylic acid and N-hydroxysuccinimide (NHS) are dissolved in anhydrous dimethylformamide (DMF) and pre-cooled in an ice bath. 1,3-dicyclohexylcarbodiimide (DCC), dissolved in anhydrous DMF, is then added drop-wise with constant stirring. The solution is removed from the ice bath, covered with aluminium foil, and stirred at room temperature under a nitrogen atmosphere overnight. The resulting solution is filtered under vacuum using Whatman filter-paper to remove insoluble dicyclohexylurea, a key by-product of the reaction. The success of NHS ester production can be monitored by spotting the sample on a TLC silica F254 plate, with anthraquinone-2-carboxylic acid and NHS (each dissolved in anhydrous DMF) as standards, and ethyl acetate as the resolving solvent. The products on the TLC plate can be visualized by illumination with UV at 254 nm wavelength. The AQ-NHS ester can now be dried in a rotary evaporator at 40 °C and then re-dissolved in chloroform. The sample is filtered under vacuum using Whatman filter-paper and the remaining solution dried using nitrogen gas. The final dried AQ-NHS ester is best stored covered from light in a dry area at room temperature. For coupling of the AQ to amino-labeled DNA, the AQ-NHS ester is first dissolved in anhydrous DMF to make a 20 lg/lL stock solu-
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tion. The standard AQ coupling reaction (typically 100 lL) consists of the approximately 10–12 nmoles C6-amino DNA, 75 mM Na2B4O7, and 7 lL of the AQ stock solution. The solution is covered with aluminium foil to keep it dark, and is shaken vigorously overnight at room temperature. The solution is briefly spun-down and the supernatant transferred to a new microcentrifuge tube. The AQ-DNA is then ethanol precipitated with NaCl and anhydrous ethanol and washed with cold 70% (v/v) ethanol. The pellet is fully resuspended in 100 lM triethylammonium acetate (TEAA) (pH 6.85) and an equal volume of chloroform. The two-phase solvent system is used to efficiently dissolve and separate the AQ-DNA from the un-coupled AQ. The aqueous layer is extracted two times with equal volumes of chloroform, lyophilized, and resuspended in 100 lM TEAA (pH 6.85). Detailed description of the synthesis and purification of the AQNHS ester can be found in Fahlman and Sen, 2002 [16], Sankar and Sen, 2004 [20], and Leung and Sen, 2007 [27]. 2.5. HPLC purification of AQ-coupled DNA The AQ-coupled DNA is purified by high-pressure liquid chromatography using a reverse-phase C-18 column. The solvent flow is set continuously at 1 ml/minute and the solvents are heated to 40 °C. The initial conditions are: 100% solvent A (20:1 100 lM TEAA (pH 6.85): acetonitrile), changing to 30% solvent B (100% acetonitrile) over 30 min with a linear gradient. Typically, un-coupled DNA has a retention time of approximately 10 min, while the AQcoupled DNA (‘‘AQ-DNA”) has a retention time of approximately 17 min (Fig. 8). Retention times will naturally vary for different DNA oligonucleotides, depending on the length of the DNA. To ensure that the collected fractions are indeed AQ-DNA the ratio is calculated of the area of the 260 nm absorbance peak relative to the
Fig. 8. (A) Anthraquinone coupled to DNA. (B) HPLC traces of anthraquinone coupled and un-coupled DNA. The exact retention times of the DNA oligonucleotides will vary depending on its length, but in all cases, the un-coupled DNA will elute from the reverse-phase HPLC column before the coupled DNA. DNA absorbs only at 260 nm (B) and AQ absorbs at both 260 and 335 nm (C), therefore the peak at approximately 12 min is the un-coupled DNA and the peak at approximately 22 min is the DNA coupled to AQ. The final broad peak at 32 min is the un-coupled AQ.
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335 nm absorbance peak. AQ-DNA oligonucleotides have a ratios ranging from 20–50 depending on the length of the DNA [16]. The fractions containing the AQ-DNA can then be lyophilized and re-dissolved in deinoized water. The AQ-DNA concentration can be determined by measurement of UV absorption at 260 nm. A typical yield of AQ-DNA conjugates ranges from 50–90% depending on the sequence and the synthesis batch. 2.6. RNA synthesis It is convenient to synthesize RNA strands using T7 RNA polymerase in vitro, with a double-stranded template that incorporates the T7 RNA polymerase promoter. The DNA template is typically amplified and rendered double-stranded using the polymerase chain reaction (PCR) using Vent or Taq DNA polymerase, under standard conditions. Transcription reactions can then be carried out using purified T7 RNA polymerase (30 lg) and the DNA template (2–5 lM) in 100 ll of transcription buffer (40 mM Tris–HCl, pH 7.9, 26 mM MgCl2, 2.5 mM spermidine, 10 mM DTT, 0.01% Triton X-100, 8 lM GTP, 4 lM ATP and CTP, 2 lM UTP). The reaction typically proceeds for 2–16 h at 37 °C. Upon completion, the transcription mixtures are treated with deoxyribonuclease I for 30 min at 37 °C, and the newly synthesized RNAs ethanol precipitated and size-purified by denaturing polyacrylamide gel electrophoresis. Dephosphorylation (for RNA) using calf intestinal alkaline phosphatase and 32P-end labeling (for DNA and RNA) using polynucleotide kinase can be carried out using standard protocols. 2.7. Preparation of assemblies DNA or RNA assemblies can be formed by annealing stoichiometric mixtures of all constituent oligonucleotides, including the AQDNA (1 lM each) and trace amounts of 50 -end labeled oligonucleotide, in 50 mM Tris–HCl (pH 7.5) and 0.1 mM EDTA. The solutions are heated at 90 °C for 30 s, cooled slowly to room temperature (60–90 min), then diluted 4-fold in irradiation buffer (final concentration: 50 mM Tris–HCl (pH 7.5), 50 mM NaCl, 1 mM MgCl2, and 0.1 mM EDTA; or any other desired solution conditions). The samples are incubated for approximately 15 min at room temperature prior to photo-irradiation to initiate charge flow. 2.8. Charge-flow inducing irradiation of the DNA or RNA complexes DNA/RNA solutions are placed in the wells of a 96-well polycarbonate or polyethylene plate pre-coated with glycogen (1 mg/ml) and placed under a Blak-Ray UVL-56 lamp (365 nm) for 60 min at a distance of 2 cm from the bulb. Temperature can be maintained by having the plate in contact with a shallow water bath. Following photo-irradiation, the samples are ethanol precipitated with the assistance of glycogen as carrier. 2.9. Piperidine treatment and denaturing gel electrophoresis The DNA pellets are dissolved in 10% (v/v) piperidine and incubated at 90 °C for 30 min. The treated DNA is lyophilized, dissolved in denaturing gel loading dye, heat denatured at 100 °C for 5 min, cooled to 22 °C, and loaded on denaturing polyacrylamide sequencing gels. The gels can be analyzed by phosphorimagery. 2.10. Aniline treatment Ethanol-precipitated DNA/RNA pellet is dissolved directly into an aniline acetate solution (20 lL aniline, 180 lL H2O, and 12 lL of glacial acetic acid) and strand scission is preformed at 60 °C for 15 min in the dark. Each sample is diluted with 80 lL of doubly
deionized water and is dried under vacuum for 1 h in a lyophilizer. The gels can be analyzed by phosphorimagery. 2.11. Quenching experiments with ascorbic acid DNA/RNA solutions are prepared as previously mentioned. Lascorbic acid or ascorbic acid-6-palmitate is added to a final concentration of 50 lM from stock solutions to the appropriate DNA solution just prior to irradiation. The L-ascorbic acid stock can be prepared with deinoized water and the ascorbic acid-6-palmitate stock solution can be prepared in dimethyl sulfoxide (DMSO) due to its low solubility in deinoized water. To equalize DMSO in all reactions to be irradiated, both the 0 and 50 lM L-ascorbic acid containing reactions are made up to a final DMSO concentration of 0.185% (v/v) to match the final concentration of DMSO that will be present in the ascorbic acid-6-palmitate-containing reactions. 2.12. Data analysis The densitometry analyses of the sequencing gels are carried out and the density of the band of interest was assessed as a percentage of the total signal contained within a particular lane. This procedure will compensate for any discrepancies of the total radioactive counts in different lanes. The normalized signal for a particular nucleotide obtained in the ‘‘dark” reaction (negative control) lane is subtracted from the signal of the same nucleotide in an irradiated sample lane. This corrected signal for a particular nucleotide is then divided by the corrected signal of the reference nucleotide to give the damage ratio. Any single intra-helical guanine can be used as the reference nucleotide as long as its CFDC is relative constant throughout all experimental conditions tested. 3. Concluding remarks CFQ is a new and versatile low-tech method for probing various structural parameters within complexly folded DNAs and RNAs. Future years will undoubtedly bring further innovations to the method; however, even in its present state, it is easily practised, and should provide practitioners with novel kinds of structural and folding information that is difficult to obtain without resorting to highresolution structural studies involving X-ray crystallography or NMR. One small innovation that might contribute to the ease of planning CFQ experiments is the synthesis and ready availability of an anthraquinone phosphoramidite for automated DNA/RNA synthesis. It is our hope that this challenge will be taken up by one or more of the larger commercial suppliers of phosphoramidites. References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] [16] [17] [18] [19]
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