ABSTRACT
DNA base flipping, which was first observed for the C5-cytosine DNA methyltransferase M.HhaI, results in a complete removal of the stacking interactions between the target base and its neighbouring bases. We have investigated whether duplex oligodeoxynucleotides containing the fluorescent base analogue 2-aminopurine can be used to sense DNA base flipping. Using M.HhaI as a paradigm for a base flipping enzyme, we find that the fluorescence intensity of duplex oligodeoxynucleotides containing 2-aminopurine at the target site is dramatically enhanced (54-fold) in the presence of M.HhaI. Duplex oligodeoxynucleotides containing 2-aminopurine adjacent to the target cytosine show little fluorescence increase upon addition of M.HhaI. These results clearly demonstrate that duplex oligodeoxynucleotides containing 2-aminopurine at the target site can serve as fluorescence probes for base flipping. Another enzyme hypothesized to use a base flipping mechanism is the N6-adenine DNA methyltransferase M.TaqI. Addition of M.TaqI to duplex oligodeoxynucleotides bearing 2-aminopurine at the target position, also results in a strongly enhanced fluorescence (13-fold), whereas addition to duplex oligodeoxynucleotides containing 2-aminopurine at the 3'- or 5'-neighbouring position leads only to small fluorescence increases. These results give the first experimental evidence that the adenine-specific DNA methyltransferase M.TaqI also flips its target base.
Specific binding of proteins to DNA often leads to drastic conformational changes in the DNA structure. Proteins can bend, kink, unwind and melt the DNA or can flip out a whole base (nucleotide) from the double helix. Base flipping was initially observed by X-ray crystallography for the C5-cytosine DNA methyltransferase from Haemophilus haemolyticus (M.HhaI) (1). Subsequently, a second C5-cytosine DNA methyltransferase, from Haemophilus aegypticus (M.HaeIII) (2), the thymidine dimer excision repair enzyme endonuclease V (3) and the uracil DNA glycosylase (4) were seen to use a base flipping mechanism. In all four protein-DNA complexes a base is completely rotated out of the DNA double helix. The C5-cytosine DNA methyltransferases and the uracil DNA glycosylase have their target bases flipped out of the DNA helix into a cleft within each enzyme, where catalysis takes place. In contrast, with endonuclease V one of the partner adenines of the thymidine dimer is flipped into a hydrophobic pocket of the enzyme permitting the catalytic residues to reach the target thymidine dimer. In all these cases base flipping allows the catalytic machinery to gain access to the target site within the bulky DNA substrate.
It has been postulated that base flipping occurred early in evolution and that it might be found in many DNA modifying enzymes (5). Possible candidates are other C5-cytosine DNA methyltransferases, N6-adenine and N4-cytosine DNA methyltransferases as well as DNA glycosylases and DNA repair enzymes. This hypothesis is supported by a number of X-ray structures of enzymes, which have been crystallized without their DNA substrates. The catalytic sites of O6-methylguanine DNA methyltransferase (6), [beta]-glucosyl transferase (7), exonuclease III (8), photolyase (9), endonuclease III (10), DNA 3-methyl adenine glycosylase II (11,12) and N4-cytosine DNA methyltransferase M.PvuII (13) were all found in the interior of the proteins. In each instance, base flipping appears to be necessary to bring the active site amino acid residues in close proximity to the target deoxynucleotide.
A similar example is the N6-adenine DNA methyltransferase from Thermus aquaticus (M.TaqI), which catalyses the methyl group transfer from S-adenosyl-l-methionine (AdoMet) to the exocyclic amino group of adenine within the double-stranded (ds) 5'-TCGA-3' DNA sequence (14). The structure of M.TaqI in complex with its cofactor consists of two domains which form a cleft wide enough to accommodate dsDNA (15). However, modelling of B-DNA into the M.TaqI structure showed that the distance between the target adenine and the activated methyl group of the cofactor is 15 Å, which is too large for direct methyl group transfer (16). By rotating the adenine out of the DNA helix towards the cofactor, the distance between the methyl group donor and acceptor can be significantly reduced, and hence a base flipping mechanism was proposed for M.TaqI. A structural comparison of M.TaqI with the base flipping C5-cytosine DNA methyltransferase M.HhaI showed that the catalytic domains of both enzymes have a very similar fold (17), although primary structural similarities between the two enzymes are limited to a few sequence motifs (18,19). In a superimposition of the catalytic domains of M.TaqI and M.HhaI both the cofactors and the catalytic motifs (N/D P P Y/F/W-motif for N6-adenine DNA methyltransferases and PC-motif for C5-cytosine DNA methyltransferases) overlay very well (17). Because of the similar architecture of their catalytic domains a shared base flipping mechanism seems likely.
In general, X-ray crystallography of protein-DNA complexes provides the absolute proof of DNA base flipping. However, cocrystallization of proteins with their DNA substrates is often cumbersome, and in some cases cocrystals might never be obtained. Therefore, other methods suitable to detect DNA base flipping would be of great value. Recently, tighter binding of the two C5-cytosine DNA methyltransferases M.HhaI and M.HpaII to duplex oligodeoxynucleotides containing mismatched bases at their target sites (20,21) and of the N6-adenine DNA methyltransferase M.EcoRV to duplexes containing unnatural bases with reduced hydrogen bonding capabilities at the target site (22) was reported. An alternative approach to detect base flipping might use the fluorescent base analogue 2-aminopurine. This base is a very sensitive probe to detect conformational changes in DNA structure (23), and it can be incorporated into oligodeoxynucleotides by solid phase DNA synthesis (24-28). The 2-aminopurine fluorescence is highly quenched in polynucleotides (29), due to stacking interactions with neighbouring bases (30,31) and would be expected to increase dramatically if the 2-aminopurine base is flipped out of the DNA helix. Furthermore, the 2-aminopurine fluorescence excitation and emission maxima are well separated from those of tryptophan and tyrosine residues in proteins, increasing its usefulness for studying protein-DNA interactions. Duplex oligodeoxynucleotides containing 2-aminopurine have been used to study the Klenow fragment of DNA polymerase I (32,33), T4 DNA polymerase (34-36), T7 RNA polymerase (37-39), Escherichia coli RNA polymerase (40) and helicases (41,42). Recently, duplex oligodeoxynucleotides containing 2-aminopurine have been used to study the N6-adenine DNA methyltransferase M.EcoRI, and a DNA base flipping mechanism was suggested for this enzyme (43).
Using the C5-cytosine DNA methyltransferase M.HhaI as a paradigm for a base flipping enzyme, we demonstrate that 2-aminopurine incorporated in DNA serves as an excellent spectroscopic probe for DNA base flipping. M.HhaI recognizes the specific tetranucleotide DNA sequence 5'-GCGC-3' (44) and catalyses the methyl group transfer from AdoMet to the 5-position of the first cytosine (45). Titration with M.HhaI of a duplex oligodeoxynucleotide, in which the first cytosine is replaced with 2-aminopurine, yields a large increase in the 2-aminopurine fluorescence intensity. In addition, the N6-adenine DNA methyltransferase M.TaqI, which was suggested to use a base flipping mechanism, also shows a strongly enhanced 2-aminopurine fluorescence after addition to a duplex oligodeoxynucleotide, in which the target adenine is replaced by 2-aminopurine. Thus, duplex oligodeoxynucleotides containing 2-aminopurine can serve as spectroscopic probes for DNA base flipping in protein-DNA complexes. In addition, these duplex oligodeoxynucleotides should be very useful to study the dynamics of DNA base flipping.
UV absorption spectra were measured in H2O using a Varian Cary 3E spectrometer. HPLC was performed using a Beckmann System Gold equipped with a programmable solvent module 125, a diode array detector module 168, an analog interface module 406 and a Shimadzu spectrofluorometric detector RF-551. Fluorescence data were collected with a SLM-Aminco AB2 spectrofluorometer. The excitation and emission bandwidths were adjusted to 2 and 8 nm, respectively. Displayed emission spectra were corrected for lamp intensity fluctuations. Fluorescence titrations were performed with an excitation wavelength of 320 nm and an emission wavelength of 381 nm to lower the influence of the natural DNA base absorption and the protein tryptophan fluorescence. Emission spectra were recorded with an excitation wavelength of 320 nm. Due to the optical arrangement of the SLM AB2 all 2-aminopurine emission spectra show an extra shoulder at around 380 nm.
Oligodeoxynucleotides were synthesized on an Applied Biosystems 392 DNA/RNA synthesizer, using standard [beta]-cyanoethyl phosphoramidite chemistry or were purchased from MWG Biotech. 2-Aminopurine, C5-methylcytosine and N6-methyladenine phosphoramidites were purchased from Glen Research. Phosphoramidites for the normal bases and the synthesizer chemicals were obtained from Applied Biosystems and MWG Biotech. Phosphoramidites were dissolved manually in dry acetonitrile at 0.1 M concentration. Sequences of synthesized oligodeoxynucleotides are given in Table 1.
Evaluation of the coupling yield using the dimethoxytrityl cation assay gave an average of 98.5% per step. Deblocking was achieved by heating in aqueous ammonia (33%, Baker) at 55°C for 8 h. Syntheses were performed `trityl on', and oligodeoxynucleotides containing the dimethoxytrityl group were purified by reversed phase HPLC (Hypersil ODS, 5 µm, 120 Å, 250 × 8 mm column, Bischoff) with a linear acetonitrile gradient (13-31% in 20 min, 2.5 ml/min) in triethylammonium acetate buffer (0.1 M, pH 7.0). After detritylation with acetic acid (80%) and ethanol precipitation oligodeoxynucleotides were desalted by gel filtration (NAP column, Pharmacia). Concentrations of oligodeoxynucleotides were determined by UV absorbance at 260 nm using the sum of the extinction coefficients of the individual bases.
Base composition of oligodeoxynucleotides was analysed by enzymatic digestion with snake venom phosphodiesterase and alkaline phosphatase followed by analytical reversed phase HPLC (26,46). Normalized peak areas of UV absorption at 254 nm correlated well with the expected base composition of the individual oligodeoxynucleotides. Oligodeoxynucleotides were 5'-labelled by treatment with T4 polynucleotide kinase (New England Biolabs) and [[gamma]-32P]ATP (0.4 MBq/µl, Hartmann Analytik) and analysed by denaturing PAGE. Radioactivity was quantified using a Phosphoimager (Molecular Imager System GS 525, Bio-Rad). All oligodeoxynucleotides showed a purity >95%.
M·HhaI was essentially isolated as described earlier (47); exhaustive dialysis prior to chromatography yielded AdoMet-free enzyme. The enzyme was purified to homogeneity by passing it through a pre-column of Q-Sepharose followed by column chromatography on S-Sepharose.
M.TaqI was expressed in ER 2267 Escherichia coli cells (New England Biolabs) harbouring pAGL15-M13, which carries the gene for M.TaqI under the inducible tac-promotor. pAGL15-M13 was kindly provided by Drs Jack Benner and Cheryl Baxa, New England Biolabs. Cells were grown at 37°C to an optical density of 0.6 at 600 nm and induced with IPTG (0.1 mM final concentration). After induction (4 h) cells were harvested by centrifugation (30 min at 3500 r.p.m.) and lysed by sonication in buffer A (Mes/HEPES/sodium acetate buffer mix, 6.7 mM each, pH 7.5, 1 mM EDTA, 10 mM [beta]-mercaptoethanol and 10% glycerol), containing sodium chloride (200 mM) and phenylmethylsulfonyl fluoride (25 mg/l). After centrifugation (1 h at 35 000 g) the supernatant was loaded onto a cation exchange column (Poros HS/M, 26 × 95 mm, Perseptive Biosystems) and M.TaqI was eluted with a linear gradient of sodium chloride (0.2-1.0 M) in buffer A. Fractions containing M.TaqI were pooled, 3-fold diluted with buffer A and loaded onto a heparin column (heparin Sepharose CL 6B, 26 × 95 mm, Pharmacia). M.TaqI was eluted with a linear gradient of sodium chloride (0.2-1.0 M) in buffer A. Fractions containing M.TaqI were pooled, and the protein solution was concentrated by ultrafiltration (Centriprep 10, Amicon). The concentrated protein solution (10 ml) was loaded onto a gel filtration column (Superdex 75, 26 × 600 mm, Pharmacia) and eluted with a Tris-HCl buffer (20 mM, pH 7.4) containing potassium chloride (600 mM), EDTA (0.2 mM), DTE (2 mM) and glycerol (10%). Fractions containing M.TaqI were pooled, 2-fold diluted with glycerol and stored at -20°C. M.TaqI was at least 95% pure as judged by polyacrylamide gel electrophoresis in the presence of sodium dodecyl sulfate (48) and staining with Coomassie Blue. However, it was found that some fraction of the protein still contained the cofactor AdoMet. AdoMet bound to M.TaqI was removed as previously described (49).
Concentrations of M.HhaI and M.TaqI were estimated by the method of Bradford (50) (Coomassie Protein Assay Reagent, Pierce) using bovine serum albumin as standard. Concentrations were calculated using molecular weights of 37 100 g/mol for M.HhaI and 47 900 g/mol for M.TaqI.
DNA binding studies with M.HhaI were performed by incubating M.HhaI (50 nM) with 5'-32P-labelled duplexes oligodeoxynucleotides (5 nM) in buffer B (10 mM Tris-HCl, pH 7.4, 50 mM sodium chloride, 0.5 mM EDTA and 2 mM [beta]-mercaptoethanol) supplied with glycerol (5%) and, where applicable, S-adenosyl-l-homocysteine (100 µM) for 30 min at 8°C. Aliquots (5 µl) were loaded onto a non-denaturing polyacrylamide gel (13%, acrylamide:bisacrylamide = 19:1) and electrophoresis was carried out in Tris-borate (45 mM, pH 8.3) and EDTA (1 mM) at 8°C for 2 h. Dried gels were autoradiographed to an X-ray film and the films were scanned with a ScanMaker E6 densitometer (Microtek). DNA binding of M.TaqI was analysed by incubation of M.TaqI (30 nM) with 5'-32P-labelled duplexes oligodeoxynucleotides (1 nM) in buffer C (10 mM Tris-acetate, pH 7.9, 50 mM potassium acetate, 10 mM magnesium acetate, 1 mM DTT and 0.01% Triton X-100) supplied with BSA (0.1 mg/ml) and glycerol (10%) at 0°C for 10 min. Aliquots (10 µl) were loaded onto a non-denaturing polyacrylamide gel (10%, acrylamide:bisacrylamide = 19:1) and electrophoresis was carried out in a buffer (pH 8.5) containing Tris-HCl (25 mM), glycine (192 mM) and EDTA (1 mM) at 0°C for 2 h. Radioactivity in the gel was visualized by autoradiography using a Bio-Rad Phosphoimager.
Fluorescence measurements were performed in 5 × 5 mm cuvettes at 25°C. Titrations of duplex oligodeoxynucleotides (20 and 250 nM) with M.HhaI were performed in buffer B and with M.TaqI in buffer C (see gel mobility shift assays). S-adenosyl-l-homocysteine (50 µM) was added to buffer B for titrations of 2CGC and GC2G with M.HhaI. Duplex oligodeoxynucleotides were made by mixing strands containing 2-aminopurine or non-fluorescent control strands with a 1.1-fold molecular excess of methylated complementary strands in the appropriate buffer and heating at 95°C for 2 min followed by cooling to room temperature over a period of 4 h. Enzymatic digestion of the duplex oligodeoxynucleotide G2GC was done by addition of DNase I (0.1 µg, Boehringer Mannheim) and snake venom phosphodiesterase (2 µg, Boehringer Mannheim) to a solution (400 µl) containing G2GC (250 nM) and buffer B supplemented with magnesium chloride (5 mM).
Table 1
All fluorescence emission spectra and fluorescence intensities from titrations were corrected for protein tryptophan fluorescence by subtraction of control spectra and control titrations, where the 2-aminopurine fluorophor was replaced by adenine in the duplex oligodeoxynucleotides. In addition, fluorescence data were corrected for variable background emission of the solutions. This was necessary, because the 2-aminopurine fluorescence is highly quenched in duplex oligodeoxynucleotides and its fluorescence intensity at low duplex concentration used in this study (20 and 250 nM) is small compared with the background. Therefore, the fluorescence intensities of the various duplex oligodeoxynucleotides were measured at higher concentrations (typically between 0.5 and 4 µM). Fluorescence intensities of all duplex oligodeoxynucleotides containing 2-aminopurine showed a linear dependence on their concentrations. Fluorescence intensities at 20 and 250 nM concentration free of background were calculated from the obtained slopes. Titration data (fluorescence intensities as a function of total enzyme concentrations) were fitted to the real solution of the quadratic binding equation for one binding site (51) using the data analysis program GraFit (52).
For fluorescence measurements with M.HhaI, which catalyses the methylation of the first cytosine within the ds 5'-GCGC-3' DNA sequence, we used three hemimethylated duplex oligodeoxynucleotides, which are derived from the recognition sequence (for abbreviations of duplex oligodeoxynucleotides see Table 1). In G2GC the target cytosine, in 2CGC its 5'-neighbour and in GC2G its 3'-neighbour, is replaced with the fluorescent base 2-aminopurine. Since the recognition sequence of M.HhaI is palindromic, M.HhaI can, in principle, bind in two orientations. In order to direct binding of M.HhaI in the orientation, in which 2-aminopurine (G2GC) or cytosine (2CGC and GC2C) is targeted, the complementary strand in these duplexes contained 5-methylcytosine at the other target position. Such a preference for single binding orientation was observed in cocrystals of M.HhaI with hemimethylated DNA (53).
Analogous hemimethylated duplex oligodeoxynucleotides were used for the fluorescence measurements with M.TaqI, which catalyses the methylation of the adenine within the ds 5'-TCGA-3' DNA sequence. In the duplex oligodeoxynucleotides TCG2 and TC2A the target adenine and its 5'-neighbour are replaced by 2-aminopurine, whereas in TCGA2 2-aminopurine occupies the 3'-neighbour position outside the recognition sequence. In order to direct binding in the desired orientation the complementary strands in these duplex oligodeoxynucleotides contained N6-methyladenine at the other target position.
The various duplex oligodeoxynucleotides containing 2-amino-purine showed quite different fluorescence intensities (Table 2). In general, the fluorescence intensities of duplex oligodeoxynucleotides, in which the 2-aminopurine base can form a Watson-Crick-like base pair with thymine, are lower than those having 2-aminopurine in a mismatch position. Apart from base pairing, the 2-aminopurine fluorescence in these duplex oligodeoxynucleotides is also quite sensitive to the sequence context.
Binding of M.HhaI and M.TaqI to the various duplex oligodeoxynucleotides was analysed in gel mobility shift assays. The results obtained with M.HhaI are shown in Figure 1. Duplex oligodeoxynucleotides were incubated with M.HhaI and the free duplex oligodeoxynucleotides (lower bands) separated from the M.HhaI-bound duplexes (upper bands) by non-denaturing gel electrophoresis. Under these conditions M.HhaI completely bound to G2GC, partly to the specific duplex oligodeoxynucleotide GCGC and no binding was observed with 2CGC and GC2G (lanes 1-4). However, addition of S-adenosyl-l-homocysteine (AdoHcy) to the incubation mixture resulted in nearly complete binding of all duplex oligodeoxynucleotides (lanes 5-8). Binding of M.TaqI to specific duplex oligodeoxynucleotide TCGA and the various duplex oligodeoxynucleotides containing 2-aminopurine is demonstrated in Figure 2. The electrophoretic mobilities of the duplex oligodeoxynucleotides alone are shown in lanes 1, 3, 5 and 7. Addition of M.TaqI (30 nM) prior to electrophoresis resulted in nearly complete formation of complexes for all four duplex oligodeoxynucleotides even in the absence of AdoHcy (upper bands in lanes 2, 4, 6 and 8). Gel mobility shift experiments with different M.TaqI concentrations showed that with increasing M.TaqI concentrations the ratio between complexes with lower mobility and complexes with higher mobility changed in favour of the complexes with lower mobility (data not shown). These results suggest that the complexes with lower mobility may represent complexes in which multiple M.TaqI molecules are bound to the duplex oligodeoxynucleotides.
Fluorescence spectra of G2GC, which places the 2-aminopurine fluorophor at the target site of M.HhaI, are shown in Figure 3. The 2-aminopurine fluorescence intensity of G2GC is very low at the concentration used in this study (curve a), but can be readily observed at higher levels (curve b). This is due to a very strong quenching of the fluorophor, which mainly results from stacking interactions with the neighbouring bases. The degree of fluorescence quenching in G2GC was determined by digestion with DNase I and phosphodiesterase. Enzymatic degradation of the duplex and release of the free 2-aminopurine-2'-deoxynucleotide (curve d) leads to a 100-fold increase of the fluorescence intensity at 381 nm. Upon addition of saturating amounts of M.HhaI to G2GC the 2-aminopurine fluorescence intensity at 381 nm increases 54-fold (Fig. 3, curve c and Table 2). Furthermore, the shape of the fluorescence emission spectrum in the presence of M.HhaI is changed compared with the duplex spectrum, but resembles that of the free 2-aminopurine-2'-deoxynucleotide. This can be seen in Figure 3 by comparing the fluorescence emission spectrum of G2GC at 10 µM concentration (curve b) with those of G2GC at 250 nM concentration in the presence of M.HhaI and after enzymatic digestion (curves c and d, respectively). The emission spectrum of G2GC in the presence of M.HhaI shows a maximum at 359 nm, which is very close to the maximum of the free 2-aminopurine-2'-deoxynucleotide at 360 nm. Figure 4A shows a titration curve of G2GC with M.HhaI. The fluorescence signal reaches saturation within the concentration range used. However, because of the high concentration of G2GC (250 nM) used no accurate dissociation constant, Kd, could be determined in this experiment. Titrations with 20 nM G2GC (Fig. 4B) yielded Kd of 3.9 ± 1.1 and 2.9 ± 0.9 nM in the absence and presence of AdoHcy, respectively, which agrees well with the values determined in gel mobility shift assays (S. Serva and S. Klimasauskas, unpublished data).AB
Titrations of duplex oligodeoxynucleotides containing 2-aminopurine with M.TaqI are shown in Figure 5, and the relative fluorescence changes are listed in Table 2. Addition of saturating amounts of M.TaqI to the duplex oligodeoxynucleotide TCG2, which carries the 2-aminopurine fluorophor at the target site of M.TaqI, results in a 13-fold increase of the fluorescence intensity. The titration data were fitted to the real solution of the quadratic binding equation for one binding site and resulted in a Kd of 120 ± 30 nM. This relatively weak binding is in conflict with the apparent tighter binding in the gel mobility shift assay (Fig. 2). Therefore, binding of M.TaqI to TCG2 was measured in a third independent assay. The intrinsic tryptophan fluorescence of M.TaqI increases upon binding of M.TaqI to specific DNA. Titration of M.TaqI with TCG2 at an excitation wavelength of 295 nm and an emission wavelength of 340 nm also results in a fluorescence increase (data not shown), which is highly dominated by the protein tryptophan fluorescence. This titration yielded a Kd of 120 ± 16 nM, which agrees very well with the Kd determined by 2-aminopurine fluorescence titration. Given the similarities of the Kd determined in the different fluorescence solution assays, we think that the gel mobility shift assay indicates too tight binding for M.TaqI. Fluorescence emission spectra of TCG2 at 10-fold higher concentrations (2.5 µM) in the absence and the presence of saturating amounts of M.TaqI are shown in Figure 6. The shape of the emission spectrum of TCG2 in the presence of M.TaqI (curve a) differs from the duplex spectrum (curve d), but resembles that of the free 2-aminopurine-2'-deoxynucleotide in Figure 3 (curve a). This is very similar to the result obtained for G2GC in the presence of M.HhaI. However, the emission spectrum of the M.TaqI-TCG2 complex has a maximum of 357 nm and is slightly blue-shifted compared with the emission maximum of the free 2-aminopurine-2'-deoxynucleotide at 360 nm.
We have used the C5-cytosine DNA methyltransferase M.HhaI as a paradigm for a DNA base flipping enzyme. It was demonstrated by X-ray crystallography of a series of ternary complexes between M.HhaI, various versions of substrate DNA and the product cofactor AdoHcy that the enzyme flips its target cytosine out of the DNA and into its active site (1,53-55). In addition, there is indirect evidence that base flipping does also occur in the binary M.HhaI-DNA complex. It was shown, that M.HhaI catalyses exchange of the proton at the 5-position of cytosine with solvent in the absence of cofactor (56). Base flipping in the binary complex is also supported by the observation that M.HhaI binds more tightly to duplex oligodeoxynucleotides containing thymine, uracil (20) and even adenine or guanine replacements for the target cytosine (21). This enhanced binding of M.HhaI to such duplexes is attributed to the lower energy required for opening a mismatched base pair upon formation of the binary complex. Recently, base flipping in the binary complex has been observed by 19F NMR spectroscopy (57). 2-Aminopurine also forms a mismatch with guanine in the G2GC duplex, and it is therefore not surprising that M.HhaI binds strongly to this duplex oligodeoxynucleotide in the gel shift assay.
We would like to thank Jack Benner and Cheryl Baxa from New England Biolabs for providing the M.TaqI expression vector and Nathalie Bleimling for the purification of M.TaqI. We also would like to thank Roger Goody for his continuous support and Bernard Connolly for carefully reading the manuscript. This work was supported by a grant from the Deutsche Forschungsgemeinschaft and from the Volkswagen-Stiftung.
Nucleic Acids Research
Pages
Introduction
Materials And Methods
Oligodeoxynucleotide synthesis, purification and evaluation
DNA methyltransferases
Gel mobility shift assays
Steady-state fluorescence measurements
Fluorescence data processing
Results
Duplex oligodeoxynucleotides used
Binding of M.HhaI and M.TaqI to duplex oligodeoxynucleotides
Fluorescence studies with M.HhaI
Fluorescence studies with M.TaqI
Discussion
Acknowledgements
References
REFERENCES
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R. K. Neely, D. Daujotyte, S. Grazulis, S. W. Magennis, D. T. F. Dryden, S. Klimasauskas, and A. C. Jones Time-resolved fluorescence of 2-aminopurine as a probe of base flipping in M.HhaI-DNA complexes Nucleic Acids Res., December 9, 2005; 33(22): 6953 - 6960. [Abstract] [Full Text] [PDF]
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T.-J. Su, M. R. Tock, S. U. Egelhaaf, W. C. K. Poon, and D. T. F. Dryden DNA bending by M.EcoKI methyltransferase is coupled to nucleotide flipping Nucleic Acids Res., June 7, 2005; 33(10): 3235 - 3244. [Abstract] [Full Text] [PDF]
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N. B. Gaied, N. Glasser, N. Ramalanjaona, H. Beltz, P. Wolff, R. Marquet, A. Burger, and Y. Mély 8-vinyl-deoxyadenosine, an alternative fluorescent nucleoside analog to 2'-deoxyribosyl-2-aminopurine with improved properties Nucleic Acids Res., February 17, 2005; 33(3): 1031 - 1039. [Abstract] [Full Text] [PDF]
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E. Merkiene and S. Klimasauskas Probing a rate-limiting step by mutational perturbation of AdoMet binding in the HhaI methyltransferase Nucleic Acids Res., January 13, 2005; 33(1): 307 - 315. [Abstract] [Full Text] [PDF]
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R. A. Estabrook, R. Lipson, B. Hopkins, and N. Reich The Coupling of Tight DNA Binding and Base Flipping: IDENTIFICATION OF A CONSERVED STRUCTURAL MOTIF IN BASE FLIPPING ENZYMES J. Biol. Chem., July 23, 2004; 279(30): 31419 - 31428. [Abstract] [Full Text] [PDF]
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T.-J. Su, B. A. Connolly, C. Darlington, R. Mallin, and D. T. F. Dryden Unusual 2-aminopurine fluorescence from a complex of DNA and the EcoKI methyltransferase Nucleic Acids Res., April 23, 2004; 32(7): 2223 - 2230. [Abstract] [Full Text] [PDF]
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E. G. Malygin, W. M. Lindstrom Jr., V. V. Zinoviev, A. A. Evdokimov, S. L. Schlagman, N. O. Reich, and S. Hattman Bacteriophage T4Dam (DNA-(Adenine-N6)-methyltransferase): EVIDENCE FOR TWO DISTINCT STAGES OF METHYLATION UNDER SINGLE TURNOVER CONDITIONS J. Biol. Chem., October 24, 2003; 278(43): 41749 - 41755. [Abstract] [Full Text] [PDF]
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A. M. Kallman, M. Sahlin, and M. Ohman ADAR2 A->I editing: site selectivity and editing efficiency are separate events Nucleic Acids Res., August 15, 2003; 31(16): 4874 - 4881. [Abstract] [Full Text] [PDF]
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 D. Daujotyte, G. Vilkaitis, L. Manelyte, J. Skalicky, T. Szyperski, and S. Klimasauskas Solubility engineering of the HhaI methyltransferase Protein Eng. Des. Sel., April 1, 2003; 16(4): 295 - 301. [Abstract] [Full Text] [PDF]
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K. S. Christine, A. W. MacFarlane IV, K. Yang, and R. J. Stanley Cyclobutylpyrimidine Dimer Base Flipping by DNA Photolyase J. Biol. Chem., October 4, 2002; 277(41): 38339 - 38344. [Abstract] [Full Text] [PDF]
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 Z. E. R. Newby, E. Y. Lau, and T. C. Bruice A theoretical examination of the factors controlling the catalytic efficiency of the DNA-(adenine-N6)-methyltransferase from Thermus aquaticus PNAS, June 11, 2002; 99(12): 7922 - 7927. [Abstract] [Full Text] [PDF]
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X. Cheng and R. J. Roberts AdoMet-dependent methylation, DNA methyltransferases and base flipping Nucleic Acids Res., September 15, 2001; 29(18): 3784 - 3795. [Abstract] [Full Text] [PDF]
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E. G. Malygin, A. A. Evdokimov, V. V. Zinoviev, L. G. Ovechkina, W. M. Lindstrom, N. O. Reich, S. L. Schlagman, and S. Hattman A dual role for substrate S-adenosyl-L-methionine in the methylation reaction with bacteriophage T4 Dam DNA-[N6-adenine]-methyltransferase Nucleic Acids Res., June 1, 2001; 29(11): 2361 - 2369. [Abstract] [Full Text] [PDF]
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M. Rist and J. Marino Association of an RNA kissing complex analyzed using 2-aminopurine fluorescence Nucleic Acids Res., June 1, 2001; 29(11): 2401 - 2408. [Abstract] [Full Text] [PDF]
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S. S. Szegedi, N. O. Reich, and R. I. Gumport Substrate binding in vitro and kinetics of RsrI [N6-adenine] DNA methyltransferase Nucleic Acids Res., October 15, 2000; 28(20): 3962 - 3971. [Abstract] [Full Text] [PDF]
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I. Pompizi, A. Haberli, and C. J. Leumann Oligodeoxynucleotides containing conformationally constrained abasic sites: a UV and fluorescence spectroscopic investigation on duplex stability and structure Nucleic Acids Res., July 15, 2000; 28(14): 2702 - 2708. [Abstract] [Full Text] [PDF]
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W. M. Lindstrom Jr., J. Flynn, and N. O. Reich Reconciling Structure and Function in HhaI DNA Cytosine-C-5 Methyltransferase J. Biol. Chem., February 18, 2000; 275(7): 4912 - 4919. [Abstract] [Full Text] [PDF]
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A. Jeltsch, F. Christ, M. Fatemi, and M. Roth On the Substrate Specificity of DNA Methyltransferases. ADENINE-N6 DNA METHYLTRANSFERASES ALSO MODIFY CYTOSINE RESIDUES AT POSITION N4 J. Biol. Chem., July 9, 1999; 274(28): 19538 - 19544. [Abstract] [Full Text] [PDF]
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B. Holz, N. Dank, J. E. Eickhoff, G. Lipps, G. Krauss, and E. Weinhold Identification of the Binding Site for the Extrahelical Target Base in N6-Adenine DNA Methyltransferases by Photo-cross-linking with Duplex Oligodeoxyribonucleotides Containing 5-Iodouracil at the Target Position J. Biol. Chem., May 21, 1999; 274(21): 15066 - 15072. [Abstract] [Full Text] [PDF]
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G. Vilkaitis, A. Dong, E. Weinhold, X. Cheng, and S. Klimasauskas Functional Roles of the Conserved Threonine 250 in the Target Recognition Domain of HhaI DNA Methyltransferase J. Biol. Chem., December 1, 2000; 275(49): 38722 - 38730. [Abstract] [Full Text] [PDF]
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G. Vilkaitis, E. Merkiene, S. Serva, E. Weinhold, and S. Klimasauskas The Mechanism of DNA Cytosine-5 Methylation. KINETIC AND MUTATIONAL DISSECTION OF HhaI METHYLTRANSFERASE J. Biol. Chem., June 8, 2001; 276(24): 20924 - 20934. [Abstract] [Full Text] [PDF]
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H. Y. Yi-Brunozzi, O. M. Stephens, and P. A. Beal Conformational Changes That Occur during an RNA-editing Adenosine Deamination Reaction J. Biol. Chem., October 5, 2001; 276(41): 37827 - 37833. [Abstract] [Full Text] [PDF]
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A. A. Evdokimov, V. V. Zinoviev, E. G. Malygin, S. L. Schlagman, and S. Hattman Bacteriophage T4 Dam DNA-[N6-adenine]Methyltransferase. KINETIC EVIDENCE FOR A CATALYTICALLY ESSENTIAL CONFORMATIONAL CHANGE IN THE TERNARY COMPLEX J. Biol. Chem., January 4, 2002; 277(1): 279 - 286. [Abstract] [Full Text] [PDF]
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