Towards understanding the catalytic mechanism of M.EcoP15I [EcoP15I MTase (DNA methyltransferase); an adenine methyltransferase], we investigated the role of histidine residues in catalysis. M.EcoP15I, when incubated with DEPC (diethyl pyrocarbonate), a histidine-specific reagent, shows a time- and concentration-dependent inactivation of methylation of DNA containing its recognition sequence of 5′-CAGCAG-3′. The loss of enzyme activity was accompanied by an increase in absorbance at 240 nm. A difference spectrum of modified versus native enzyme shows the formation of N-carbethoxyhistidine that is diminished by hydroxylamine. This, along with other experiments, strongly suggests that the inactivation of the enzyme by DEPC was specific for histidine residues. Substrate protection experiments show that pre-incubating the methylase with DNA was able to protect the enzyme from DEPC inactivation. Site-directed mutagenesis experiments in which the 15 histidine residues in the enzyme were replaced individually with alanine corroborated the chemical modification studies and established the importance of His-335 in the methylase activity. No gross structural differences were detected between the native and H335A mutant MTases, as evident from CD spectra, native PAGE pattern or on gel filtration chromatography. Replacement of histidine with alanine residue at position 335 results in a mutant enzyme that is catalytically inactive and binds to DNA more tightly than the wild-type enzyme. Thus we have shown in the present study, through a combination of chemical modification and site-directed mutagenesis experiments, that His-335 plays an essential role in DNA methylation catalysed by M.EcoP15I.
DNA methylation is catalysed by a diverse group of enzymes that are uniformly dependent on AdoMet (S-adenosyl-L-methionine) as a methyl donor . M.EcoP15I [EcoP15I MTase (DNA methyltransferase)] adds a methyl group to the second adenine in the recognition sequence 5′-CAGCAG-3′ in the presence of AdoMet . The enzyme is a part of the type III R-M (restriction modification) system. The type III R-M system contains two subunits: the Res subunit encoded by the res gene and the Mod subunit encoded by the mod gene. Although the Mod subunit alone catalyses the methylation reaction, both the Res and Mod subunits are necessary for DNA cleavage, where ATP hydrolysis is required. Cleavage by M.EcoP15I requires two sites in opposite orientation and occurs 25–27 bp downstream from one of the recognition sequences in an ATP- and Mg2+-dependent reaction . It is an N6-adenine methyltransferase and, like all N6-adenine methyltransferases, M.EcoP15I contains two highly conserved sequences: FXGXG (motif I) at position 444–448 and DPPY (motif IV) at position 125–128. While mutations in motif I completely abolished AdoMet binding but left target DNA recognition unaltered, mutations in motif IV resulted in loss of enzyme activity, but AdoMet and DNA binding were not affected . By employing chemical modification using thiol-directed agents and site-directed mutagenesis, it was demonstrated that Cys-344 in M.EcoP15I was necessary for enzyme activity and played an essential role in DNA binding . We showed that when M.EcoP15I binds to its recognition sequence, both the adenine bases in the recognition site appear to be structurally distorted, supporting the proposed base flipping mechanism for this enzyme . To achieve catalysis, the enzyme requires magnesium ions. Using a Fenton chemistry affinity cleavage assay, we located the magnesium-binding-like motif to amino acids 355–377 of M.EcoP15I . We demonstrated that the acidic amino acid residues of the region 355–377 in M.EcoP15I are important for the critical positioning of magnesium ions for catalysis . Little is known about the chemical mechanism of the methylation reaction catalysed by M.EcoP15I; in particular, the identity of critical amino acids and the nature of the active centre have not been investigated so far.
Chemical modification and the pH dependency of kinetic parameters are two widely used techniques for the identification of amino acids essential for catalysis and binding to substrate in enzyme-catalysed reactions. For instance, the bell-shaped pH–activity curve for the M.EcoRI exhibits a maximum at 7.8, with two pKa values at pH 7.0 and pH 8.4, and the most likely candidate is a histidine residue. Everett and Reich  demonstrated by using a histidine-residue-specific reagent, DEPC (diethyl pyrocarbonate), that M.EcoRI activity was completely inhibited and that this loss of activity was due to the modification of a critical histidine residue.
To begin the identification of catalytic and/or substrate-binding regions within M.EcoP15I and to further advance our understanding of how DNA recognition and catalysis might be coupled, we conducted biochemical and molecular studies on the role of histidine residues in its activity. In the present study, we have exchanged all 15 histidine residues of M.EcoP15I for alanine residues by PCR-based site-directed mutagenesis and characterized the variants expressed in Escherichia coli with respect to their DNA methylation activity and biochemical properties.
MATERIALS AND METHODS
Bacterial strains and plasmid vectors
E. coli strain JM109 (hsdR, recA) was used as a host for propagating all plasmids used in the present study. E. coli strain DH5α was used as a host for preparing pUC19. The DNA constructs derived from pUC19 were used for overexpression and purification of mutant M.EcoP15I. WT (wild-type) M.EcoP15I was expressed in E. coli BL21(DE3)pLysS cells by transforming with plasmid pDN8 .
Enzymes and chemicals
All reagents were of analytical or ultrapure grade. AdoMet was procured from Sigma. [methyl-3H]AdoMet (80 Ci/mmol) was purchased from GE Healthcare Lifescience Asia Pacific and from NEN Life Science Products. [γ-32P]ATP (5000 Ci/mmol) was purchased from Bhabha Atomic Research Centre (Mumbai, India). Magnesium chloride, BSA, ampicillin, Hepes, PEI (polyethyleneimine), DEPC, hydroxylamine and Coomassie Brilliant Blue R-250 were procured from Sigma Chemical Co. 2AP (2-aminopurine)-substituted oligonucleotides were a gift from Geoff Wilson (New England Biolabs). R.EcoP15I was purified as described earlier .
General recombinant techniques
Restriction enzymes, Klenow fragment of DNA polymerase, T4 DNA ligase and T4 polynucleotide kinase were purchased and used according to manufacturer's recommendations. Ligations, transformations and DNA electrophoresis were performed as described in . Plasmid DNA pUC19 or pGEM3Zf(–) was prepared as described in .
Oligonucleotides and radiolabelling
Basic procedures of labelling of oligonucleotides were performed as described in . All oligonucleotides showed a purity of > 95%. Concentrations of oligonucleotides were determined by UV absorbance at 260 nm by using the sum of the molar absorbance coefficients (ϵ) of the individual bases. The oligonucleotides used in the mutagenesis reactions are listed in Table 1.
|Histidine residue||Mutated to||Restriction site created||Primer and its sequence||Sensitivity/resistance to R.EcoP15I|
|Histidine residue||Mutated to||Restriction site created||Primer and its sequence||Sensitivity/resistance to R.EcoP15I|
Purification of WT M.EcoP15I and mutant M.EcoP15I
Plasmids pUC19-M.EcoP15I, pUC19-M.EcoP15I-H171A and pUC19-M.EcoP15I-H335A were transformed separately into E. coli DH5α cells. WT M.EcoP15I and mutant M.EcoP15I were purified by the method of Rao et al.  to near homogeneity. Peak fractions from the heparin–Sepharose column containing the enzyme were pooled and concentrated by dialysing against 20 mM Tris/HCl (pH 8.0) buffer containing 0.1 mM EDTA, 7 mM 2-mercaptoethanol, 40 mM NaCl and 50% (v/v) glycerol. The purity of the WT and mutant enzymes was greater than 99% on SDS/PAGE . Protein concentration was determined by the method of Bradford . The protein was dialysed against 10 mM potassium phosphate buffer (pH 7.0) containing 0.1 mM EDTA, 40 mM NaCl, 7 mM 2-mercaptoethanol and 50% glycerol for all the DEPC modification experiments just before use.
Assays of M.EcoP15I
In vitro methylation activity
MTase activity was monitored by incorporation of tritiated methyl groups into pUC19 DNA and the specific activity of the enzyme was measured as described in .
Sensitivity to restriction endonuclease
Plasmid DNA, carrying mutant MTases, was isolated using the alkaline lysis method  and then digested with EcoP15I restriction enzyme. Approx. 2 μg of plasmid DNA was digested with EcoP15I restriction enzyme for 60 min at 37 °C, followed by proteinase K/SDS treatment at 65 °C for 20 min. The products were analysed by 0.8% (w/v) agarose-gel electrophoresis in the presence of ethidium bromide.
Chemical modification of M.EcoP15I with DEPC
Since DEPC hydrolyses rapidly, stock solutions in anhydrous ethanol were prepared fresh before each experiment. From the stock solution, dilutions were done in 10 mM potassium phosphate buffer (pH 7.0). The pH of the modification reaction was 7.0, since the reaction is specific for unprotonated histidine residues between pH 5.5 and 7.5 . DEPC concentration was determined by reaction with imidazole and monitoring the increase in the absorbance at 240 nm using a molar absorbance coefficient (ϵ) of 3200 M−1·cm−1 for the formation of the reaction product N-carbethoxyimidazole. M.EcoP15I (6.6 μM) in 10 mM potassium phosphate buffer (pH 7.0) was incubated with DEPC (0–1 mM) at 25 °C. The final concentration of ethanol in reaction mixtures never exceeded 4% (v/v) of the total volume and was shown not to have any effect on enzymatic activity. At various time points, aliquots were removed, and residual M.EcoP15I activity was measured as described in , except that the enzyme assay mixture additionally contained 5 mM imidazole to quench unchanged DEPC. The rate of the modification reaction can be followed as the rate of loss of activity of the enzyme.
Under conditions where the modifier is far in excess of the enzyme, the loss of activity declines at a pseudo-first-order rate, hence:
where Vt is the velocity at time t and V0 is velocity at zero time. A linear plot of the natural logarithm of percentage residual activity remaining against time gives a straight line whose slope kapp is the pseudo-first-order rate constant at the particular modifier concentration. The pseudo-first-order rate constant, kapp, is a function of the modifier. The logarithm of the kapp obtained at different concentrations of the modifier, when plotted against the logarithm of the modifier concentration, gives a linear plot with the slope that specifies the number of residues modified (n).
where K is the y intercept that gives the rate of inactivation of the enzyme. The modification of histidine residues was also monitored spectrophotometrically by observing the increase in absorbance at 240 nm, and the number of modified residues (N-carbethoxyhistidine) was determined as described above. The control containing the same components but without DEPC was used to blank the absorbance. The stoichiometry of histidine modification was correlated with enzyme activity by monitoring a parallel experiment for time-dependent loss of activity. To examine substrate protection against DEPC inactivation, M.EcoP15I was pre-incubated with and without DNA or AdoMet in 10 mM potassium phosphate buffer (pH 7.0) for 10 min at 4 °C prior to inactivation with DEPC. At different time points, aliquots were withdrawn and the residual methylation activity was determined as described in .
Hydroxylamine treatment of inactivated enzyme
Hydroxylamine solution was prepared by dissolving the solid reagent in 10 mM potassium phosphate buffer, followed by titration to pH 7.0 with potassium hydroxide. Re-activation of DEPC-inactivated enzyme with hydroxylamine was assessed by incubating M.EcoP15I (6.6 μM) in 10 mM potassium phosphate buffer (pH 7.0) with 1 mM DEPC for 9 min at 25 °C until enzyme activity decreased to 10% of its original activity. The reaction was rapidly quenched with 5 mM imidazole (pH 7.0). Hydroxylamine was then added to a final concentration of 250 mM for absorbance change study and 350 mM for assaying enzyme activity. Aliquots were removed every hour and the residual enzyme activity was determined. Hydroxylamine had no measurable effect on the activity of the unmodified enzyme.
Site-directed mutagenesis was performed using the PCR-based technique  to replace all the 15 histidine residues individually. Single mutations were introduced into the M.EcoP15I gene by using the two-stage megaprimer PCR method. PCR reactions were carried out with Phusion DNA polymerase (Finnzymes). For each histidine-to-alanine substitution, appropriate forward primer and mutagenic primer were used in the first stage. In the first round of PCR, oligonucleotide primers (Table 1) and template plasmid pDN8 were used to amplify a DNA fragment, which was used as a megaprimer in the second round of PCR. The full-length PCR product was obtained in the second round of PCR by extension of megaprimer. The PCR product obtained was purified, digested with DpnI restriction enzyme to cleave the methylated template DNA and was transformed into DH5α E. coli strain and plated in LB (Luria–Bertani) agar medium containing ampicillin (100 μg/ml). The mutagenic primers were designed in such a way to change the respective histidine to alanine and to create a type II restriction enzyme site (see Table 1). Hence, the resultant plasmids could be screened easily. The resultant plasmids were used for expression and purification of mutant M.EcoP15I. The transformants were checked for desired mutation by amplifying DNA fragments of interest. The mutants were scored by the restriction site created. All mutations were confirmed by dideoxy sequencing of the region derived from the PCR fragment.
EMSAs (electrophoretic mobility-shift assays)
Duplex A (Table 1) was used for specific DNA binding. Non-denaturing polyacrylamide gels were used for the EMSAs. The oligonucleotide duplex labelled with [γ-32P]ATP was incubated for 10 min on ice in binding buffer (50 mM potassium phosphate, pH 7.0, 1 mM EDTA, 10 mM MgCl2, 20 mM NaCl, 7 mM 2-mercaptoethanol and 100 μM sinefungin) and protein. Reaction volumes were typically 20 μl. These were subjected to 6% PAGE in TBE (Tris/borate/EDTA; 1×TBE=45 mM Tris/borate and 1 mM EDTA). Electrophoresis was performed at 6 °C and 100 V for 3–5 h, depending on the separation required. Protein–DNA complexes formed were visualized by a phosphoimager.
CD spectral analysis
CD measurements were recorded on a Jasco J810 polarimeter between 200 and 250 nm in a quartz cuvette of 1 mm path length. All experiments were performed at 25 °C in 10 mM potassium phosphate buffer (pH 7.0). The protein solution was incubated for 10 min in the cuvette in a final volume of 400 μl prior to recording the spectrum. M.EcoP15I spectra were recorded at a protein concentration of 6.6 μM in 10 mM potassium phosphate buffer (pH 7.0). The observed ellipticities were converted into mean residue ellipticity [θMRE], by using the following equation: [θMRE]=[θ]obs(MRW)/10cl, where θobs is the observed ellipticity in degrees, MRW is the mean residue molecular mass based on a molecular mass of 150 kDa, c is the protein concentration in mg/ml and l is the path length of the cell in centimetres. Each experimental spectrum represents the best fit of at least three determinations.
Purification of AdoMet
Unlabelled AdoMet as supplied by Sigma was approx. 80% pure. In order to rule out any experimental artefacts due to these impurities, AdoMet was purified using cation-exchange chromatography . Briefly, AdoMet was dissolved in 50 mM HCl and applied to an SP-Sepharose (sulfopropyl–Sepharose) column equilibrated with 10 mM HCl. The column was washed with 50 mM HCl, followed by 150 mM HCl. AdoMet was eluted with 500 mM HCl.
Steady-state fluorescence emission spectra of 2AP-containing oligonucleotide samples were measured as described in . All samples were incubated until equilibrium was established under the particular set of conditions before measuring the steady-state fluorescence intensities. All fluorescence emission spectra and fluorescence intensities from titrations were corrected for protein tryptophan fluorescence by subtraction of control spectra and control titrations. In addition, fluorescence data were corrected for variable background emission of the solutions. Each spectrum recorded was an average of three scans. Sequences of the oligonucleotides used to measure steady-state fluorescence emission spectra are given in Table 1 (Duplex B). Titrations of duplex oligodeoxynucleotides (250 nM) with M.EcoP15I were performed in 50 mM Tris/HCl buffer (pH 8.0) containing 20 mM NaCl in the presence of 10 mM MgCl2.
Protein fluorescence emission spectra were measured for WT and mutant M.EcoP15I on a PerkinElmer spectrofluorimeter LS 55 using a 1 cm quartz cuvette at 25 °C. The emission spectra were recorded over a wavelength of 300–400 nm with a λex of 280 nm. The slit width of 10 nm was used for both excitation and emission. Enzymes were allowed to equilibrate for 2 min in methylation buffer [10 mM Tris/HCl (pH 8.0) containing 20 mM NaCl] before measurements were made. Small aliquots of cofactor (final concentration 0.5–4.5 μM) were added to the reaction containing WT or mutant MTase (500 nM) and spectra were recorded. The binding of AdoMet to MTase resulted in quenching of tryptophan fluorescence. Each spectrum recorded was an average of three scans.
SPR (surface plasmon resonance) studies
The binding kinetics of M.EcoP15I and M.EcoP15I H335A mutant with DNA was determined by SPR spectroscopy using the BIAcore2000 optical biosensor (GE Healthcare Lifescience, Uppsala, Sweden). The 5′-biotinylated double-stranded oligonucleotides with 5′-CAGCAG-3′ recognition sequence (Duplex C, Table 1) were immobilized on to a streptavidin-coated SA chip (GE Healthcare Lifescience) as per the manufacturer's recommendations. The binding reactions were carried out in a continuous flow of buffer A (10 mM Hepes buffer, pH 8, containing 20 mM NaCl, 1 mM EDTA and 0.05% surfactant P-20) and buffer B (10 mM Hepes buffer, pH 8, containing 100 mM NaCl, 1 mM EDTA and 0.05% surfactant P-20) at a flow rate of 10 μl/min. The required protein concentration was made by diluting with the running buffer (buffer A or B), and cofactors including sinefungin (40 μM) and MgCl2 (10 mM) were added to the protein prior to running on the DNA surface. The surface was regenerated by passing 5 μl of 0.05% SDS, followed by 10 μl of running buffer for further binding reactions. One of the four surfaces not having the biotinylated oligonucleotide was used as a negative control. The binding data were analysed using a 1:1 Langmuir binding model in BIAcore evaluation software, version 3.0.
RESULTS AND DISCUSSION
Inactivation of M.EcoP15I
When M.EcoP15I was treated with DEPC, its methylase activity was inhibited in a dose-dependent manner. To study the effect of DEPC on the activity of M.EcoP15I, the enzyme was first dialysed against 10 mM potassium phosphate buffer (pH 7.0) containing 0.1 mM EDTA, 40 mM NaCl, 7 mM 2-mercaptoethanol and 50% glycerol. Inactivation kinetics was carried out with freshly dialysed enzyme at 25 °C with 0, 0.1, 0.25, 0.5, 0.75 and 1 mM DEPC for 9 min. The modification reaction was arrested by transfer of the reaction mixture into assay buffer containing 5 mM imidazole. Both time- and concentration-dependent inactivations of the enzyme were observed with excess DEPC dissolved in 8% (v/v) ethanol at near neutral pH values (Figure 1A).
Kinetics of inactivation of M.EcoP15I by DEPC
The results obtained from the enzyme-inactivation experiments at various concentrations of DEPC are presented as a plot of the natural logarithm of percentage activity against time (Figure 1A). The linearity of these plots indicates that under the conditions used the process approximates to first-order kinetics at all DEPC concentrations used. The linearity of this plot also indicated that under the inactivation conditions used no reversible DEPC–enzyme complex was formed before inactivation. Imidazole (5 mM) and ethanol (0.02%) did not show any effect on enzyme activity (results not shown). The inactivation curves showed that at 1 mM DEPC, there was almost complete inhibition of the enzyme activity (Figure 1A). The apparent first-order rate constant kapp was calculated from the slopes of the lines obtained from the first-order plot. These values were plotted against log[DEPC] and the slope of this line gives the number of histidine residues modified (Figure 1B). For M.EcoP15I, the value obtained was 1.5, which suggested that only a single/single class of histidine residues were modified.
UV difference spectrum of DEPC-modified M.EcoP15I versus native M.EcoP15I
DEPC is known to modify histidine residues by the addition of an ethyl group to the imidazole ring to yield N-carbethoxyhistidine derivatives . Although DEPC is used for modification of histidine residues, it can also react with tyrosine and lysine residues. The UV difference spectrum of DEPC-modified M.EcoP15I versus native M.EcoP15I at different time points is depicted in Figure 2(A). After 5 min of incubation, the spectrum showed a peak at 240 nm characteristic of the formation of N-carbethoxyhistidine. Further, the absence of a decrease in absorption at 278 nm, which is characteristic of the formation of o-carbethoxytyrosine, excludes the possibility of tyrosine modification.
Correlation between inactivation and histidine modification
Correlation between the number of histidine residues modified and the loss of catalytic activity
To quantify the number of histidine residues that reacted with DEPC, the absorbance at 240 nm was followed as a function of time. The absorbance of M.EcoP15I at 240 nm, where changes in absorbance are observed with the modification of histidine residues, increased in a dose-dependent manner with DEPC treatment (Figure 2B). The DEPC–histidine adduct, in a stoichiometry of 1:1, absorbs strongly in the near-UV region. The number of histidine residues modified can be estimated from the absorbance change at 240 nm [a molar absorbance coefficient (ϵ) of 3200 M−1·cm−1] upon addition of DEPC. Figure 2(B) represents the correlation between the extent of enzyme inactivation and the extent of histidine modification over an incubation period of 70 min with DEPC. It is clear from the Figure that after 5 min of incubation with DEPC, when four histidine residues are modified, there was almost complete loss of activity.
Hydroxylamine treatment of DEPC-modified enzyme
DEPC modifies different nucleophiles such as amine, alcohol, thiol, imidazole and guanidine groups, producing carbethoxyl derivatives. Hydroxylamine removes the ethoxyformyl group from modified histidine and tyrosine residues, but not from the more stable ethoxyformylcysteine and ethoxyformyl-lysine residues . Restoration of the activity of modified protein after treatment with hydroxylamine should be accompanied by a parallel decrease in the absorbance at 240 nm. If lysine side chains are modified, then hydroxylamine is unable to reverse the effect caused by DEPC. M.EcoP15I (6.6 μM) was incubated with 1 mM DEPC in 10 mM potassium phosphate buffer (pH 7.0) for 9 min. The change in the absorbance against control enzyme was monitored. Hydroxylamine (250 mM) was added and the decrease in absorbance was monitored after 1, 2 and 3 h. Clearly, there was a decrease in the absorbance at 240 nm at each time point (Figure 3).
Reaction of DEPC-inactivated M.EcoP15I with hydroxylamine
Similarly the experiment was repeated, but with 350 mM hydroxylamine. Aliquots were withdrawn at 1, 2 and 3 h, dialysed against 10 mM potassium phosphate buffer (pH 7.0) and assayed for activity. The aliquots withdrawn after 1, 2 and 3 h treatment restored the activity to 22, 47 and 80% respectively compared with the modified enzyme (results not shown). Thus hydroxylamine treatment caused a decrease in absorbance at 240 nm accompanied by restoration of activity. This clearly established that tyrosine or lysine residues were not affected by DEPC treatment. While the inactivation process was very fast at a relatively low concentration of the modifier, the enzyme re-activation proceeded somewhat more slowly. The reversibility of inactivation indicates that excess of modifier was not used and that the inactivation did not result from denaturation of the enzyme. The result obtained with hydroxylamine also indicates that the ethoxyformylhistidine did not react further with DEPC, resulting in the cleavage of the imidazole ring. Otherwise, the loss of activity would have been irreversible with hydroxylamine.
CD spectra of the native and DEPC-modified M.EcoP15I
One possible interpretation of the loss of activity could be that the chemical modification of the reactive surface-accessible histidine residues may have altered the conformation of M.EcoP15I or impeded substrate accessibility to the active site and thereby affected MTase activity. It is also possible that the reactive histidine residues are not in the active site, but rather they might be necessary for stability. CD spectra of the native M.EcoP15I were compared with that of the DEPC-modified M.EcoP15I. The far-UV CD spectra, specifically the minima at 208 and 221 nm, demonstrate the characteristics of an α-helical secondary structure that showed no change after modification with DEPC. This suggests that inactivation is not due to alterations of the α-helix. No significant changes in the CD spectrum of the modified enzyme at other wavelengths were observed, providing further evidence for unaltered protein structure after modification with DEPC (Figure 4A).
Effect of DEPC on protein conformation and DNA binding properties
Binding of unmodified and DEPC-modified M.EcoP15I to DNA
We next investigated whether DEPC-modified M.EcoP15I could bind to DNA. To monitor the binding of unmodified and DEPC-modified M.EcoP15I to DNA, a fixed concentration of 30-mer oligonucleotide containing the recognition sequence 5′-CAGCAG-3′ and varying concentrations of unmodified and DEPC-modified M.EcoP15I were used. The binding was carried out in the presence of 100 μM sinefungin and 10 mM MgCl2. Increasing unmodified protein concentration increased the binding of M.EcoP15I to duplex (lanes 4 and 5), whereas there was no complex formation with DEPC-modified M.EcoP15I (lanes 7 and 8) (Figure 4B). This suggested that a histidine residue(s) is possibly involved in DNA binding.
Protection against DEPC inactivation of M.EcoP15I by substrates
The protection offered by the substrate is proof of the fact that the inactivation is not due to conformational changes or other changes that occur far from the active site. The binding of substrates to an enzyme often protects the active site from chemical modification. To confirm that the DEPC-accessible histidine residues participated in the M.EcoP15I activity, active M.EcoP15I (5.32 μM) was pre-incubated with pUC19 DNA (0.72 μM) in 10 mM potassium phosphate buffer (pH 7.0), and treated with various concentrations of DEPC. M.EcoP15I was then diluted with assay buffer, and the remaining methylase activity was determined. As shown in Figure 5, pre-incubation of M.EcoP15I with DNA substantially inhibited the inactivation of M.EcoP15I by DEPC. That is, more than 75% of the methylase activity remained even after treatment with 0.2 mM DEPC. In contrast, pre-incubation of the methylase with increasing concentrations of AdoMet (0.2–1.5 μM) did not protect the enzyme against inactivation by DEPC (results not shown). These results indicated that at least one DEPC-sensitive histidine residue of M.EcoP15I was involved in the binding of DNA or in M.EcoP15I's catalytic activity. The simultaneous presence of both substrates did not provide further protection than DNA alone against DEPC inactivation (results not shown).
Protection against DEPC inactivation of M.EcoP15I
Generation of M.EcoP15I variants by site-directed mutagenesis
From the results presented above, it is evident that DNA can protect ∼80% of the enzyme from DEPC inactivation and suggests that modification of a single histidine residue is sufficient to inactivate the enzyme. In an effort to corroborate the biochemical studies and determine which of the histidine residues were critical for the MTase activity, we performed site-directed mutagenesis to replace individually each of the 15 histidine residues of M.EcoP15I with alanine residue. By replacing with alanine in each case, it was possible to introduce a convenient restriction site, thus allowing screening of mutants (Table 1).
Activities of mutant M.EcoP15I
In order to assess the modification phenotype of the M.EcoP15I mutants, we first carried out an in vitro restriction assay. Plasmid DNAs from cells expressing the WT or mutant MTases were isolated, and their sensitivity to digestion by the cognate M.EcoP15I restriction enzyme (R.EcoP15I) was determined. Plasmids harbouring an active methylase would result in methylation of all M.EcoP15I sites in vivo, and thereby be resistant to subsequent in vitro cleavage by R.EcoP15I. It is clear from Figure 6 that the plasmid DNAs carrying the mutants M.EcoP15I-H171A and M.EcoP15I-H335A were not protected from cleavage by R.EcoP15I, indicating these to be inactive methylases. All other plasmid DNAs carrying the mutation at other histidine residues were protected from cleavage by R.EcoP15I, indicating these methylases to be functionally active (Figure 6).
Digestion of pUC19 plasmid encoding WT and mutant methyltransferase with R.EcoP15I
The purified M.EcoP15I and mutant M.EcoP15I-H335A enzyme were analysed on native polyacrylamide gel (Figure 7A) and by Western blotting (Figure 7B) for alterations in the electrophoretic mobilities, and no apparent changes were detected. In vitro methylation activity measurements clearly showed that purified preparation of H171A was almost as active as the WT enzyme, while H335A mutant enzyme was inactive (Figure 7C). Increasing the amount of H335A mutant enzyme (up to 10 μg) in the standard assay conditions led to no enhancement of methylation activity. The results of the in vitro methylation assay corroborated the in vivo restriction assay, except for the H171A mutant. The discrepancy in the results obtained by in vivo and in vitro assays of methylation for H171A mutant enzyme is not clear at the moment.
Characterization of WT and M.EcoP15I-H335A
Methylation activity and substrate binding properties of M.EcoP15I-H335A
In vitro methylation assays (described above), which measured the ability to transfer 3H-labelled methyl group from [methyl-3H]AdoMet, clearly showed that the mutant M.EcoP15I-H335A was inactive. We reasoned that the loss of activity of M.EcoP15I-H335A could be due to conformational changes in the protein structure, or to altered substrate binding properties. The effect of placing an alanine residue at position 335 in M.EcoP15I was determined by measuring (for WT and mutant enzymes) the secondary structures using CD spectroscopy. Gross conformational changes could not be detected between WT and mutant enzymes (results not shown). As we detected no major conformational changes from the substitution, it is likely that the changes observed resulted from a direct effect of the substitution rather than from some more generalized change in MTase structure.
To test if the ability of binding to DNA and AdoMet was affected because of the substitution, binding properties of the WT and mutant MTases were investigated. Gel shift assays were performed using a 30-mer oligonucleotide containing a single M.EcoP15I recognition site. End-labelled oligonucleotide was titrated with increasing concentrations of WT and mutant MTases separately. The complex formed was resolved on native polyacrylamide gels and quantified using a phosphoimager. As can be seen from Figure 8(A), both the WT and mutant M.EcoP15I-H335A enzymes bind to DNA, but with seemingly different affinities. The replacement of His-335 by an alanine residue has another interesting consequence. The complex formed with the mutant enzyme has faster mobility, indicating that the complex is more compact than the complex formed in the case of WT enzyme. Gel shift assays performed using methylated 31-mer oligonucleotides (CAGCA*G, with ‘A*’ indicating a methylated adenine) revealed that the mutant enzyme bound slightly more tightly than the WT enzyme (results not shown). Increased migration in a non-denaturing gel could be indicative of (i) a change in charge on the mutant protein, (ii) a change in the shape of the protein on interacting with DNA and (iii) a mutation resulting in the dissociation of subunits. To determine the size and subunit structure of M.EcoP15I-H335A in solution, gel filtration chromatography was performed. Both WT and H335 mutant enzymes are eluted as single peaks at a position corresponding to a globular protein with a molecular mass of 150 kDa, suggesting that these proteins exist as dimers in solution under native conditions (results not shown). It is clear from these results that mutation of histidine to alanine at position 335 did not affect the oligomeric status of the enzyme. In the absence of DNA, the extent of migration of both the proteins on a non-denaturing gel is the same, thus ruling out the effect of any charge difference on the protein (Figure 7A). It is thus likely that the mutant MTase upon binding to DNA acquires a compact structure, resulting in faster mobility. Extended electrophoresis in a non-denaturing gel revealed differences between a ternary MTase–DNA–AdoMet complex and a binary MTase–DNA complex in the case of M.EcoKI . These results suggest that the loss in enzyme activity of this mutant MTase is not due to its inability to bind to DNA, but, possibly because of tighter binding of the mutant enzyme to DNA, the enzyme thereby may no longer be available to methylate further.
DNA binding properties of WT and M.EcoP15I-H335A
2AP, an analogue of adenine, is a strongly fluorescent adenine isomer whose fluorescence is quenched within duplex DNA, largely due to intra-strand base stacking in its environment. Fluorescence intensity changes caused by the binding of a protein to DNA are indicative of a change in the environment of 2AP . The base-flipping mechanism postulated for many MTases suggests that if 2AP is located at an appropriate position, its fluorescence would be dramatically enhanced as it would be swung out of the strongly quenched DNA double-helical environment into the enzyme catalytic site. Such enhancements of fluorescence emission intensity have been observed for enzymes known to use base flipping from the availability of crystal structures such as HhaI, HaeIII and TaqI MTases . 2AP is not only a probe for base flipping, but also probes other aspects of DNA structure and dynamics that can also be altered by protein binding. The fluorescence intensity changes of DNA containing 2AP caused by the binding of adenine and cytosine MTases has been used as a probe for base flipping . To investigate base flipping, if any, fluorescence spectra of DNA containing 2AP modification (Duplex B; Table 1) was measured in the absence and presence of the enzyme. We had earlier shown that irradiation of an oligonucleotide containing 2AP at the target position instead of an adenine, at 320 nm, produced a strong fluorescence emission spectra with a λmax of 370 nm [6,7]. Annealing of this oligonucleotide with the complementary strand (formation of duplex DNA) resulted in a 2-fold decrease in fluorescence intensity at 370 nm [6,7]. When M.EcoP15I (2.5 μM) was incubated with the duplex, a 10–12-fold increase in 2AP fluorescence was observed (Figure 8B). The fluctuations in fluorescence intensity appeared to be dependent on the preparation of the substrate and the relative enzyme activity. Next, 2AP fluorescence measurements were carried out in the presence of M.EcoP15I-H335A. The addition of mutant MTase to the 2AP-containing duplex DNA showed significant enhancement of the fluorescence of 2AP (Figure 8B) when compared with WT M.EcoP15I, which corroborates our previous observation (Figure 8A) about tight binding to DNA by the mutant enzyme. These observations would suggest that a step after base flipping is affected in this mutant enzyme. One possible reason could be that replacement of histidine with alanine at this position causes conformational change, probably altering the alignment of critical amino acids at the active site required for the further step(s) (post base-flipping step) in the methylation reaction scheme.
The ability of the M.EcoP15I-H335A mutant MTase to bind AdoMet was assessed by a fluorescence quenching experiment. Both WT and mutant MTases (500 nM) were titrated with increasing amounts of AdoMet (0.5–80 μM) and the fluorescence emission spectra were recorded between 300 and 400 nm. Addition of AdoMet to M.EcoP15I WT and mutant MTases resulted in significant quenching of the native fluorescence (Figures 9A and 9B). A simple interpretation of cofactor binding and DNA binding would be that the mutant MTase binds DNA and AdoMet, flips the target base, but is still inactive.
Tryptophan fluorescence quenching analysis of WT and M.EcoP15I-H335A with increasing concentrations of AdoMet
SPR spectroscopy was used to determine the kinetics of DNA binding by M.EcoP15I MTase and M.EcoP15I-H335A mutant MTase. We have monitored the extent of binding of M.EcoP15I MTase and M.EcoP15I-H335A mutant MTase to DNA by using fixed concentrations of 35-mer oligonucleotide containing the recognition sequence 5′-CAGCAG-3′ (Duplex C, Table 1) and varying concentrations of MTase. The binding was carried out in the presence of cofactors, including 40 μM sinefungin and 10 mM MgCl2. The association and dissociation of the proteins from DNA were monitored by changes in the resonance due to the change in mass on the sensor surface. The kinetic constants were determined by subjecting the sensorgrams of association and dissociation phases to global analysis using BIAevaluation software 3.0. The global fitting analyses both association and dissociation data for all concentrations simultaneously using a 1:1 Langmuir binding model. Interaction studies of M.EcoP15I MTase and M.EcoP15I-H335A mutant MTase with DNA in SPR spectroscopy revealed significant differences in Kd values. A Kd value of 1.38×10−7 M for WT M.EcoP15I (in 20 mM NaCl) and a Kd value of 6.75×10−5 M at 100 mM NaCl concentration revealed the lower affinity at high concentrations of salt (Figures 10A and 10B). This correlates well with the previously determined DNA binding properties of the WT M.EcoP15I MTase . In contrast, Kd values of 4.51×10−8 M (20 mM NaCl) and 1.49×10−6 M (100 mM NaCl) for H335A enzyme were obtained (Figures 10C and 10D), which are 10-fold lower than the Kd values obtained for the WT M.EcoP15I MTase. This indicates a higher affinity of the mutant MTase for DNA, corroborating the gel shift assay described previously (Figure 8A).
Kinetic analysis of interaction of WT and M.EcoP15I-H335A with double-stranded DNA
It is interesting to note that similar observations were reported by Wyszynski et al.  and Mi and Roberts  in the case of M.EcoRII and M.HhaI respectively. Replacement of a catalytic cysteine residue by a glycine residue in these cytosine MTases resulted in mutant enzymes that bind abnormally tightly to DNA. As a result, these mutations were cytotoxic to E. coli cells expressing the mutant MTases [24,25]. In the case of M.HhaI, the glycine replacement lowered the Kd of the enzyme for its DNA substrate approx. 10-fold .
The main objective of the present study was to identify histidine residues that are essential for catalytic activity of the M.EcoP15I. Histidine residues are known to occur in the active sites of several enzymes where they act as the general base or general acid. The active histidine residue may act as a proton acceptor or donor in protein–ligand interactions, play a role in charge–relay interactions between amino acids at the active site and may be involved in conformational changes associated with substrate binding or oligomerization of protein chains. In order to determine whether histidine residues are essential for the activity of M.EcoP15I, we have chemically modified the enzyme using the histidine-specific reagent DEPC. We have shown here that the methylase activity of M.EcoP15I is inhibited by DEPC treatment, suggesting that histidine residues are essential for the methylase activity. There are 15 histidine residues in M.EcoP15I, distributed throughout the coding region. We have compared the primary structures of 11 N6-adenine methyltransferases belonging to the β-group using the ClustalW program. His-171 and His-241 were the only two residues in M.EcoP15I that aligned with exocyclic amino MTases. His-171 lies in motif V and His-241 lies in motif VII . These are believed to be involved in folding of the catalytic region. In order to determine which of the histidine residues are important for catalytic function of the methylase, we have exchanged all the 15 histidine residues in M.EcoP15I. The results of our analysis imply that the effects on DNA methylase activity that were observed with the H335A mutant enzyme is probably because the enzyme bound tightly to DNA and, therefore, was no longer available to methylate other substrate molecules. Of course, we cannot exclude subtle structural effects of the amino acid substitution that indirectly impair catalytic activity. It is interesting to recall that in one of our earlier studies, we demonstrated that the cysteine residue at position 344 in M.EcoP15I is a critical amino acid involved in DNA binding . Replacing the cysteine residue at position 344 resulted in an enzyme that does not bind DNA, while replacing the histidine at position 335 resulted in a mutant MTase that bound tightly to DNA. Both these amino acids are in the putative TRD (target recognition domain) region of the MTase.
We are aware of the fact that neither chemical modification nor site-directed mutagenesis experiments can provide unequivocal evidence for catalytic involvement of a particular amino acid residue. This is only possible with reference to detailed structural information. In the absence of such information, experiments of the kind described here may help in obtaining only an idea of possible mechanisms. In conclusion, we have shown through a combination of chemical modification and site-directed mutagenesis that histidine at position 335 is an essential amino acid in M.EcoP15I.
P. S. J. acknowledges the DBT (Department of Biotechnology), Indian Institute of Science, Bangalore, India, for a postdoctoral fellowship. We thank Ms Arathi and Ms Chiranjeevi for expert technical assistance. Financial support from DBT Program Support to the Indian Institute of Science is acknowledged.