Solubility engineering of the HhaI methyltransferase

Dalia Daujotyte1, Giedrius Vilkaitis1, Laura Manelyte1, Jack Skalicky2,3, Thomas Szyperski2,4 and Saulius Klimasauskas1,4

1 Laboratory of Biological DNA Modification, Institute of Biotechnology, LT-2028 Vilnius, Lithuania and 2 Department of Chemistry, University at Buffalo, The State University of New York, Buffalo, NY 14260, USA 3 Present address: NHMFL/Florida State University, Tallahasee,FL 32310, USA

4 To whom correspondence should be addressed. E-mail: szypersk{at}chem.buffalo.edu; klimasau{at}ibt.lt


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
DNA methylation is involved in epigenetic control of numerous cellular processes in eukaryotes, however, many mechanistic aspects of this phenomenon are not yet understood. A bacterial prototype cytosine-C5 methyltransferase, M.HhaI, serves as a paradigm system for structural and mechanistic studies of biological DNA methylation, but further analysis of the 37 kDa protein is hampered by its insufficient solubility (0.15 mM). To overcome this problem, three hydrophobic patches on the surface of M.HhaI that are not involved in substrate interactions were subjected to site-specific mutagenesis. Residues M51 or V213 were substituted by polar amino acids of a similar size, and/or the C-terminal tetrapeptide FKPY was replaced by a single glycine residue ({Delta}324G). Two out of six mutants, {Delta}324G and V213S/{Delta}324G, showed improved solubility in initial analyses and were purified to homogeneity using a newly developed procedure. Biochemical studies of the engineered methyltransferases showed that the deletion mutant {Delta}324G retained identical DNA binding, base flipping and catalytic properties as the wild-type enzyme. In contrast, the engineered enzyme showed (i) a significantly increased solubility (>0.35 mM), (ii) high-quality 2D-[15N,1H] TROSY NMR spectra, and (iii) 15N spin relaxation times evidencing the presence of a monomeric well-folded protein in solution.

Keywords: DNA cytosine methyltransferase/fluorescence spectroscopy/NMR/protein engineering/protein solubility


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The post-replicative modification of cytosine and adenine residues in DNA plays an important role in both prokaryotes and eukaryotes. DNA methylation is performed by DNA methyltransferases (MTases) using the methyl group donor S-adenosyl-L-methionine. In higher organisms, methylation at the C5 position of cytosine is solely observed. DNA methylation is involved in controlling many cellular processes such as gene repression, X-chromosome inactivation, genome imprinting and replication timing (Vertino, 1999Go). 5-Methylcytosine is a key determinant of epigenetic regulation and is essential for normal development of animal and plant species. Aberrations in cytosine-5 methylation correlate with human genetic disease and, therefore, the MTases are potent candidate targets for developing new therapies (Szyf, 1998Go). In prokaryotes, DNA MTases often serve as components of restriction-modification systems (Dryden, 1999Go). Their role in regulation of certain essential genes in pathogenic bacteria has led to their use as targets for developing novel antibiotic drugs (Wahnon et al., 2001Go).

M.HhaI is a DNA cytosine-5 methyltransferase from bacterium Haemophilus haemolyticus, which recognizes the tetranucleotide sequence 5'-GCGC-3' and methylates the inner cytosine (bold face). With 327 residues (37 kDa), it is one of the smallest representatives of a homologous family of enzymes (Cheng and Roberts, 2001Go). The HhaI MTase became a paradigm for structural studies after the discovery of an unusual reaction intermediate, whereby an enzyme completely flips its target nucleotide out of the DNA helix and into the catalytic site for methylation (Klimasauskas et al., 1994Go). Numerous subsequent studies showed that DNA base flipping is a common and crucial event in enzymatic DNA modification and DNA repair (Roberts and Cheng, 1998Go; Hollis et al., 2000Go; Mol et al., 2000Go; Goedecke et al., 2001Go).

Substantial efforts have since been devoted to elucidate the mechanisms of base flipping by DNA modifying enzymes. The HhaI MTase has been extensively examined by employing a variety of methods including mutagenesis, kinetic analysis, fluorescence and NMR spectroscopy, isothermal calorimetry and X-ray crystallography (Wang et al., 2000Go; Cheng and Roberts, 2001Go; Vilkaitis et al., 2001Go; Swaminathan et al., 2002Go; Varnai and Lavery, 2002Go; Zhou et al., 2002Go). A series of crystal structures for M.HhaI•DNA complexes revealed valuable details of interactions at atomic resolution. However, they all showed the target base in its final position outside the helix (O’Gara et al., 1998Go; Vilkaitis et al., 2000Go). 19F-NMR studies in solution identified important base-flipping intermediates (Klimasauskas et al., 1998Go), but technical limitations (see below) precluded thorough structural characterization by NMR. Crucial aspects of the reaction mechanism still remain obscure and to advance understanding of the base-flipping process requires structural characterization of initial and intermediate conformers of the reaction complex. To obtain further insights into the mechanism of base flipping we decided to initiate an NMR structural study of macromolecular interactions and dynamics of the HhaI methyltransferase–DNA system in solution. However, our extensive experience, including previous 19F-NMR studies (Klimasauskas et al., 1998Go), revealed an insufficient solubility of the enzyme under reaction conditions (0.15 mM in a buffer containing 50 mM NaCl). Increased salt concentration or other additives help somewhat to maintain the HhaI MTase in solution, but variation of the ionic strength also interferes with the enzyme’s interaction with DNA. Moreover, owing to poor behavior at high concentrations, in certain cases problems were encountered when studying MTase–DNA interactions with isothermal calorimetry, stopped-flow fluorescence or even gel shift analysis (see below).

Here we report the structure-based rational design of functional M.HhaI mutants with enhanced solubility. Three solvent-accessible hydrophobic positions of the protein were chosen for engineering and six mutants were prepared and analyzed. One of the engineered MTases showed significantly enhanced solubility (>0.35 mM) with full retention of kinetic and catalytic properties of the WT enzyme. The modified variant exhibits high-quality 2D NMR spectra and thus represents the first DNA methyltransferase suitable for detailed structural studies employing NMR spectroscopy and other techniques.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Mutagenesis

Restriction endonucleases, DNA modification enzymes and kits were obtained from MBI Fermentas. Oligonucleotides for site-specific mutagenesis (mutated codons are shown in bold face and newly created diagnostic restriction endonuclease sites are underlined) were as follows:

  1. 5'-TTCACCAAAATTCTTCTCATATACTTC-3' (a new diagnostic site for R.MboII)
  2. 5'-TCTTGGTTTGTCATCGATAAATCTTTTCTATC-3' (a new diagnostic site for R.Bsu15I)
  3. 5'-TTAGCCATTTAATGATGAACCAATG-3'
  4. 5'-TTTTCGCAATGATCTCAATATTC-3'

The pHH553 plasmid, which is a derivative of pHSHW-5 (Klimasauskas et al., 1991Go), carries an IPTG-inducible overexpression vector for M.HhaI (S.Klimasauskas and G.Vilkaitis, unpublished results). The pTZ-HE phagemid was constructed by inserting the 970 bp HincII fragment of pHH553 into the HincII-linearized vector pTZ19R. Mutations M51K and V213S were introduced using the Kunkel method (Sambrook and Russell, 2001Go). The single-stranded uracil-substituted pTZ-HE DNA was annealed with primers 1 or 2 and the complementary strand was synthesized using T7 DNA polymerase and circularized by ligation. The resulting dsDNA was transformed into the Escherichia coli ER1727 strain (Kumar et al., 1992Go). Transformants containing a diagnostic site of the restriction endonuclease MboII (for M51K mutation) or Bsu15I (for V213S mutation) were selected and sequenced through the regions concerned. Full-length MTase genes were restored by transferring the recombinant fragments into the pHH553 vector. For this purpose the 225 bp Eco81I–MunI fragment (M51K mutation) or the 117 bp CpoI–Ecl136I fragments (V213S mutation) were ligated with the 4.2 kb pHH553 Eco81I–MunI or CpoI–Ecl136I fragment, respectively. Following transformation into E.coli ER1727, the desired clones were selected by analyzing individual plasmid DNAs for the appearance of the diagnostic restriction endonuclease sites.

The {Delta}324G deletion mutant was constructed by PCR using pHH553 template and primers 3 and 4. The PCR fragment was hydrolyzed with R.BpiI, gel-purified and ligated into the linearized pHH553 vector, which was prepared by hydrolysis with R.HindIII, end-blunting with T4 polymerase followed by digestion with R.BpiI.

The double mutants M51K/{Delta}324G and V213S/{Delta}324G and the triple mutant M51K/V213S/{Delta}324G were produced by recombining fragments containing the corresponding mutations through proper restriction endonuclease sites. All mutations were confirmed by nucleotide sequencing of the regions concerned.

Methylation activity of mutant proteins in vivo was tested as described previously (Vilkaitis et al., 2000Go).

Protein expression and solubility analysis

WT (wild-type) HhaI and mutant proteins were expressed in E.coli strain ER1727 and the cells were harvested as described previously (Kumar et al., 1992Go; Vilkaitis et al., 2000Go). A 5 g amount of cell paste was resuspended in 20 ml of 10 mM HEPES pH 7.5, 5 mM EDTA, 10% glycerol and 10 mM 2-mercaptoethanol and lysed by ultrasonic disintegration. The suspension was centrifuged at 20 000 g for 20 min. The pellet was extracted twice with 10 mM K-PO4 pH 7.4, 5 mM EDTA, 10% glycerol, 10 mM 2-mercaptoethanol and 0.4 M NaCl (2 ml/g cell paste) and centrifuged at 20 000 g for 20 min. Both supernatants (after cell lysis and extraction with NaCl) and the pellet were analyzed by SDS–PAGE followed by staining with Coomassie Brilliant Blue R-250.

Protein purification

WT HhaI MTase was expressed and purified as described previously (Vilkaitis et al., 2000Go). {Delta}324G HhaI and V213S/{Delta}324G mutants were purified according to a newly developed scheme as follows (all purification procedures were performed at 4 °C). The harvested cells were resuspended in a lysis buffer (10 mM K-PO4 pH 7.4, 5 mM EDTA, 10 mM 2-mercaptoethanol and 1 mM PMSF, 4 ml/g cell paste), lysed by ultrasonic disintegration and centrifuged at 20 000 g for 20 min. The supernatant was applied to a DEAE-Sepharose FF (Pharmacia) column. M.HhaI eluted in the flow-through, which was directly loaded on to a heparin-Sepharose FF (Pharmacia) column (1.6x10 cm). After washing with a 3-fold column volume of equilibration buffer (10 mM K-PO4 pH 7.4, 5 mM EDTA, 10 mM 2-mercaptoethanol) the protein was eluted with a 10-fold column volume of a 0–1.0 M NaCl linear gradient in the equilibration buffer. Fractions were analyzed by SDS–PAGE and in vitro methylation activity as described previously (Kumar et al., 1992Go) and those containing the HhaI MTase band were pooled and dialyzed overnight against equilibration buffer containing 25 mM NaCl. The dialyzate was applied to an S-Sepharose FF (Pharmacia) column (1.6x8 cm). The column was washed with equilibration buffer containing 25 mM NaCl and the proteins were eluted with a 10-fold column volume of a 0.025–0.5 M NaCl linear gradient in the equilibration buffer. Fractions containing MTase were pooled, concentrated by dialysis against storage buffer (10 mM K-PO4 pH 7.4, 5 mM EDTA, 10 mM 2-mercaptoethanol, 100 mM NaCl and 50% glycerol) and stored at -20°C.

The 15N-labeled HhaI MTases were prepared by cultivating the cells in an M9 medium containing ampicillin and tetracycline and supplemented with His, Met and Trp (Sigma, 20 µg/ml each). 15NH4Cl (1 g/l, Martek Biosciences) was used as the sole nitrogen source. Protein expression and purification were performed as described above.

The purified WT and mutant MTases appeared as sole bands after SDS–PAGE followed by staining with Coomassie Brilliant Blue R-250. Protein concentration was determined by measuring A280 (theoretical extinction coefficient for WT MTase 25 610 M-1 cm-1 and for mutant proteins 24 330 M-1 cm-1). The protein concentration for WT MTase was verified by active site titration using fluorescence spectroscopy assay (see below) and was in excellent agreement with the A280 measurements.

Electrophoretic gel mobility shift assays

We used 37-mer duplex DNA oligonucleotides in this study (obtained gel-purified from MBI Fermentas): top strand, 5'-GACTGGTACAGTATCAGGCGCTGACCCACAACATCCG-3' (GCGC); lower strand, 5'-TCGGATGTTGTGGGTCAGMGCCTGATACTGTACCAGT-3' (GMGC), where M = 5-methylcytosine. Oligonucleotides were 5'-labeled using a DNA labeling kit (MBI Fermentas) and DNA duplexes were prepared as described previously (Serva et al., 1998Go).

Dissociation constants were estimated by titrating 100 pM oligonucleotide with increasing concentrations (0.1–10 nM) of the protein in binding buffer (10 mM K-PO4 pH 7.4, 5 mM EDTA, 2 mM 2-mercaptoethanol and 10% glycerol) containing 50 mM NaCl. Reactions of 20 µl were incubated at room temperature for 20 min and then 10 µl samples were loaded on to 8% polyacrylamide gel (acrylamide:bisacrylamide = 19:1) and fractionated by electrophoresis in 45 mM Tris–borate, 1 mM EDTA at 10 V/cm for 1.5 h. Autoradiography and data analysis were performed as described previously (Vilkaitis et al., 2001Go).

Fluorescence spectroscopy study of DNA base flipping

The following 37-mer duplex DNA oligonucleotides were obtained (gel-purified) from MBI Fermentas: 5'-GACTGGTACAGTATCAGGPGCTGACCCACAACATCCG-3' (GPGC) and 5'-GACTGGTACAGTATCAGGAGCTGACCCACAACATCCG-3' (GAGC), where P = 2-aminopurine, M.HhaI recognition sequence is underlined and the target base is in bold face. DNA duplexes were prepared by annealing appropriate strands as described previously (Holz et al., 1998Go). Fluorescence experiments were performed at 22°C on a Perkin-Elmer LS50-B luminescence spectrometer equipped with a xenon lamp and a 4 mm rectangular quartz cell with a mixer. Fluorescence intensity scans were measured at an excitation wavelength {lambda}Ex of 320 nm and an emission wavelength {lambda}Em of 370 nm as described previously (Holz et al., 1998Go). The 100 nM GPGC/GMGC duplex was titrated with 50–1000 nM MTase by adding appropriate amounts of the protein in reaction buffer (10 mM Tris–HCl pH 7.4, 0.5 mM EDTA, 50 mM NaCl, 2 mM 2-mercaptoethanol, 0.2 mg/ml BSA) containing 100 nM GPGC/GMGC. Three scans were averaged for each spectrum. Control spectra were recorded under identical conditions except that GAGC/GMGC substrate was used instead of the fluorescent DNA. Emission and excitation spectra with saturating protein concentrations were recorded using 100 nM substrate and 150 nM protein in the reaction buffer without BSA. Emission spectra were measured at {lambda}Ex 320 nm with excitation and emission bandwidths of 2.5 and 5 nm, respectively. Excitation spectra were measured at {lambda}Em 370 nm with excitation and emission bandwidths of 5 and 10 nm, respectively. Control measurements were performed using the GAGC/GMGC duplex. Final results were obtained by subtracting the control spectra to eliminate tryptophan fluorescence and the Raman effect as described previously (Vilkaitis et al., 2000Go).

Stopped-flow experiments were performed on an Applied Photophysics SX.18MV-R Stopped-flow Reaction Analyzer equipped with a 150 W xenon lamp. {lambda}Ex was 315 nm (monochromator bandwidth 4.2 nm) and emission was recorded using a 360 nm cut-off filter. The reaction was initiated by rapidly mixing 75 nM GPGC/GMGC oligonucleotide with 37.5–1200 nM M.HhaI (final concentrations) in reaction buffer (10 mM Tris–HCl pH 7.4, 0.5 mM EDTA, 50 mM NaCl, 2 mM 2-mercaptoethanol, 0.2 mg/ml BSA). Progress curves were collected for each protein concentration during the first 10 s. Multiple time courses (5–6 runs) were averaged and analyzed as described previously (Vilkaitis et al., 2000Go).

Steady-state kinetics

Methylation reactions were assayed as described previously, product formation being traced using the radioactive substrate [methyl-3H]AdoMet (84 Ci/mmol, Amersham) (Wu and Santi, 1987Go; Vilkaitis et al., 2000Go). Typically, reactions were performed at 37°C in methylation buffer (50 mM Tris–HCl pH 7.4, 0.5 mM EDTA, 2 mM 2-mercaptoethanol, 0.2 mg/ml BSA) containing poly(dG–dC)•poly(dG–dC) (Sigma) and [methyl-3H]AdoMet as the substrates. KmAdoMet was estimated at constant 50 nM DNA and variable 2.5–500 nM [methyl-3H]AdoMet. Similarly, KmDNA reactions contained 100 nM [methyl-3H]AdoMet and 0.1–50 nM DNA. KmAdoMet and KmDNA were measured using 50 pM MTase with 10 min incubation or 5 pM MTase and 5 min incubation, respectively (Wu and Santi, 1987Go). The reactions were quenched by adding HCl to 0.5 M concentration. Subsequent procedures including 3H radioactivity measurements and data analysis were carried out as described previously (Vilkaitis et al., 2001Go).

NMR spectroscopy

NMR spectra (see Table IIIGo) were recorded on a Varian INOVA 750 spectrometer at 25°C. Uniformly 15N-labeled WT and engineered {Delta}324G HhaI MTase were dissolved in 95% H2O–5% D2O (10 mM K-PO4, 50 mM NaCl, 0.1 mM EDTA, pH 7.4) at concentrations of 0.15 and 0.25 mM, respectively.


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Table III. NMR spectroscopy of the WT and engineered HhaI methyltransferases
 
Average T1 and T1{rho} 15N spin relaxation times were measured using previously described radiofrequency (r.f.) pulse schemes (Szyperski et al., 1993Go; Klimasauskas et al., 1998Go) extended for suppression of cross-correlated relaxation. The 15N r.f. carrier was set to 121 p.p.m. and a 3 kHz continuous-wave spin-lock r.f. frequency was used for the T1{rho} measurement. The first free induction decays were recorded at T1 delays of 8, 29, 95, 148, 213, 310, 379, 480, 592, 716, 852 and 1000 ms and T1{rho} delays of 8, 16, 24, 32, 40, 48, 64, 76, 84 ms. The total integral of the 1HN resonances located between 8 and 10 p.p.m. was determined for each delay, which largely excludes the resonances of the more flexibly disordered side chain amides. A single exponential function was fitted to the experimental data yielding T1{rho}av and T1av. The correlation time of the overall rotational tumbling {tau}c was then obtained from the T1av/ T1{rho}av ratio (Kay et al., 1989Go) using the program DASHA (Orekhov et al., 1995Go).


    Results
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 Materials and methods
 Results
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 References
 
Selection of target residues

An abundance of solvent-exposed hydrophobic residues may lead to intermolecular aggregation and low solubility of a protein in aqueous solutions. Rational protein engineering is often used to improve the solubility of such proteins by deleting non-essential apolar surface residues or replacing them with polar ones (Jenkins et al., 1995Go; Das and Georgiadis, 2001Go; Gesell et al., 2001Go; Malissard and Berger, 2001Go). To identify regions of hydrophobicity on the solvent-accessible surface, we analyzed the available crystal structures for the binary M.HhaI•AdoMet (Cheng et al., 1993Go) and the ternary M.HhaI•DNA•AdoHcy complexes (Klimasauskas et al., 1994Go). A preliminary list of targets was produced by identifying apolar residues that were more than 30% solvent accessible in both complexes as determined with Swiss PDB Viewer v.3.51 software. After excluding all Pro and Gly residues, which may be important owing to their special role in shaping the backbone conformation and all residues that may affect the motions of the catalytic loop, substrate interactions or catalysis (F92, F102, W41, Y254), we came up with three major target epitopes on the surface of the protein. In two cases, we chose to replace single aliphatic residues with polar ones of a similar size: M51 for K and V213 for S. The third epitope comprised the terminal tetrapeptide FKPY, which is seen as a protruding extension sticking out of the globule of the protein in both crystal structures. This ‘tail’ is located remote from any functionally important site, which encouraged us to eliminate the whole C-terminal tetrapeptide by replacing it with a single glycine residue (variant {Delta}324G) (Figure 1Go). Altogether, six mutant versions were prepared by site-directed mutagenesis: the three ‘single’ mutants (M51K, V213S and {Delta}324G), two double mutants (M51K/{Delta}324G and V213S/{Delta}324G) and one triple mutant (M51K/V213S/{Delta}324G).



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Fig. 1. Structure of the HhaI MTase in complex with DNA substrate and AdoHcy (PDB entry 5MHT). (A) Protein (spacefill) is shown in light gray, with hydrophobic surface residues in gray and target positions for mutagenesis in black; DNA is shown as sticks. (B) C{alpha}-trace of two symmetry-related molecules in the crystal lattice showing extensive intermolecular contacts through the C-terminal ‘tails’ (heavy) as indicated by arrows.

 
Analysis and purification of mutant proteins

The mutant HhaI MTases were expressed in E.coli cells carrying an IPTG-inducible expression vector. An initial assessment of the catalytic activity in vivo was performed by challenging their respective plasmids with the Hin6I restriction endonuclease (Vilkaitis et al., 2000Go). The activity of the endonuclease is fully abolished if the targets GCGC sites are methylated by M.HhaI. Our analysis revealed that all mutant MTases exhibited full protection as the WT enzyme (not shown). Although this assay is not very sensitive with respect to moderate differences in catalytic efficiency, this result suggests that none of the introduced mutations leads to significant structural or functional alterations in M.HhaI.

All six variants were then subjected to a solubility test. This experiment was conducted by measuring the distribution of the proteins between pellet and supernatant in crude cell extracts at different salt conditions. As previously demonstrated (Kumar et al., 1992Go), the wild-type M.HhaI remains associated with the disrupted cell debris after centrifugation at low salt conditions, but can be back-extracted by treatment with a buffer containing 0.4 M NaCl. Remarkably, our analysis showed that two engineered MTases, {Delta}324G and V213S/{Delta}324G, were soluble even at these low salt conditions, whereas all other variants, including the WT enzyme, were not (Figure 2Go and Table IGo). We also found that the M51K and V213S single mutants along with WT can be resolubilized in 0.4 M NaCl, whereas the M51K/{Delta}324G and M51K/V213S/{Delta}324G mutants were insoluble in both low and high ionic strength buffers. Therefore, we chose the {Delta}324G and V213S/{Delta}324G mutant MTases for further analysis.



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Fig. 2. Solubility of HhaI MTase variants in crude cell extracts. Cells expressing the six mutants and the WT protein were lysed by ultrasonic disintegration and centrifuged in a low ionic strength buffer. The supernatant (A) and pellet (B) fractions were analyzed by SDS–PAGE as described in Materials and methods. The appearance of the {Delta}324G and V213S/{Delta}324G mutants in supernatant but not the pellet indicates their complete solubility under low salt conditions.

 

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Table I. Solubility analysis of the HhaI methyltransferase variants. Distribution of protein between supernatant (S) and pellet (P) in crude extract at different ionic strengths
 
The WT protein (Kumar et al., 1992Go) and its numerous mutants can be readily purified by exploiting the selective back-extraction of protein from pelleted material. For obvious reasons, this procedure is no longer applicable for mutants with enhanced solubility. Therefore, we developed a new purification procedure, which is in part based on a previously reported scheme (Wu and Santi, 1988Go) and involves three chromatographic steps. After cell lysis and centrifugation the protein is fractionated using DEAE-Sepharose, heparin-Sepharose and S-Sepharose. Using this method, the {Delta}324G and V213S/{Delta}324G M.HhaI mutants were purified to homogeneity with a yield of ~0.8 mg of AdoMet-free MTase per gram of cell paste (5 mg/l of culture).

DNA binding and base flipping studies

The DNA binding capabilities of the two mutant and WT enzymes were compared using a gel mobility shift assay. A 5'-32P-labeled 37-mer duplex oligonucleotide containing hemimethylated single target site GCGC/GMGC was titrated with increasing amounts of enzyme and analyzed under non-denaturing conditions (Vilkaitis et al., 2001Go). Quantitative analysis of the binding isotherms showed that enzyme–DNA interactions under these conditions were identical: all three enzymes exhibited dissociation constants (Kd) of 0.5 nM (data not shown). One can note some protein aggregation at the high end of titration curves (at >100 nM protein) in the case of WT MTase, manifested as the appearance of low-mobility radioactive bands.

The base flipping potential of the mutant MTases was examined using the similar 37-mer hemimethylated duplex oligonucleotide GPGC/GMGC containing 2-aminopurine (P) as a target base. 2-Aminopurine can be selectively excited at 310–320 nm and detected by monitoring emission at 370 nm, however, fluorescence of this base is highly quenched in the DNA helix owing to base stacking interactions. Binding of M.HhaI induces a 50-fold enhancement of fluorescence intensity owing to enzyme-mediated flipping of the target base out of the DNA helix (Holz et al., 1998Go). Titration of the duplex with the proteins showed no significant differences between the WT and mutant MTases. A slightly smaller fluorescence intensity amplitude was achieved upon binding of a V213S/{Delta}324G HhaI MTase (Figure 3AGo), but excitation (not shown) and emission spectra (Figure 3BGo) appeared identical, within error. These results suggested that the two mutants and the WT MTase behave in a similar manner with respect to both the extent and the environment of the base in the flipped-out state (Vilkaitis et al., 2000Go).



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Fig. 3. Fluorescence study of target base flipping by WT and modified HhaI MTases. (A) Titration of 100 nM GPGC/GMGC oligonucleotide with increasing amounts of WT ({circ}), {Delta}324G HhaI (•) and V213S/{Delta}324G HhaI ({triangledown}) MTases. The total fluorescence intensity increase was measured and calculated as described in Materials and methods. (B) Corrected 2-aminopurine excitation spectra of 100 nM M.HhaI•GPGC/GMGC complexes: dashed line, WT; solid line, {Delta}324G; dashed-dotted line, V213S/{Delta}324G.

 
Real-time kinetics of M.HhaI interaction with the 37-mer fluorescent DNA in solution was studied using a stopped-flow technique. Stopped-flow fluorescence measurements were performed with constant fluorophore and varied protein concentrations. The fluorescence intensity signal was recorded for 10 s after mixing. Detailed analysis and interpretation of interactions between M.HhaI and the DNA duplex have been presented elsewhere (Vilkaitis et al., 2000Go). Briefly, the first two exponentials that are observed in the time window of 1–50 ms (Figure 4AGo) represent the fast steps of (i) binding–flipping and (ii) subsequent rearrangement of the flipped-out base. Both mutants and the WT enzyme showed very similar behavior on the millisecond time scale as derived from both qualitative comparison of the fluorescence traces (Figure 4AGo) and quantitative analysis (not shown) involving global fitting of data to a two-step reversible binding mechanism (Vilkaitis et al., 2000Go). Here we would like to focus on the third phase in the reaction profile, which includes the time range from 1 to 10 s. One can see that a slight change in fluorescence is observed in the case of WT M.HhaI at increasing protein concentrations (>0.5 µM), but is detectable for neither mutant (Figure 4BGo). In other words, both mutants, which lack the C-terminal hydrophobic tail, exhibit a simpler kinetic behavior at high protein concentrations. The complex kinetic behavior of the WT protein apparently derives from its tendency to aggregate at high protein concentrations (see discussion below). Interactions of M.HhaI with DNA may lead to changes in the multimeric (aggregation) state of the protein on the second time scale, which would manifest itself as a slight change in the steady-state fluorescence intensity.



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Fig. 4. Transient kinetic analysis of target base flipping by the WT (bottom) and modified {Delta}324G (top) M.HhaI. Fluorescence intensity progress curves at {lambda}Ex = 315 nm, {lambda}Em > 360 nm were collected after rapid mixing of 75 nM GPGC/GMGC with 300, 600 and 1200 nM MTase (final concentrations, traces bottom to top) at 25°C in a stopped-flow apparatus. (A) Fluorescence changes associated with the fast steps of DNA binding, base flipping and conformational rearrangement (Vilkaitis et al., 2000Go) are observed in the time frame 1–50 ms (log time base) for both proteins. (B) Additional change in fluorescence (due to intermolecular protein interactions) is observed on the second time scale for WT M.HhaI (linear time base).

 
Steady-state kinetic analysis

Multiple-turnover kinetic parameters KmAdoMet, KmDNA and kcat were determined for both mutant {Delta}324G and V213S/{Delta}324G HhaI MTases along with the WT enzyme using poly(dG–dC) and [methyl-3H]AdoMet as substrates (Wu and Santi, 1987Go; Vilkaitis et al., 2000Go). The concentration of DNA or AdoMet was independently varied and the measured initial velocities were fitted by non-linear regression analysis to the Michaelis–Menten equation to obtain kcat and Km values for each substrate. The control parameters for the WT M.HhaI (kcat = 0.018 s-1, KmAdoMet = 20 nM, KmDNA = 0.4 nM) were in good agreement with those obtained previously [0.022 s-1, 15 nM and 2.3 nM, respectively (Wu and Santi, 1987Go) and 0.02 s-1, 13 nM and 0.9 nM, respectively (Vilkaitis et al., 2000Go)].

The kinetic constants of the mutants were also fairly uniform (Table IIGo). Both kcat and KmAdoMet were nearly identical with those of the WT M.HhaI. Only KmDNA of the V213S/{Delta}324G mutant was 2-fold higher compared with the WT and {Delta}324G M.HhaI enzymes. The observed results suggest that the deletion of the four C-terminal residues does not alter the steady-state kinetic properties of the HhaI MTase, but the additional mutation at V213 leads to a slight change in MTase–DNA interactions. Therefore, the least modified soluble variant {Delta}324G was selected for further studies.


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Table II. Steady-state kinetic parameters of the WT and mutant HhaI methyltransferases
 
NMR spectroscopy

The WT and engineered {Delta}324G HhaI MTases were characterized by 1D 1H and heteronuclear 2D-[15N,1H] NMR spectroscopy (Table IIIGo). Uniformly 15N-labeled proteins were prepared and purified to homogeneity for that purpose. Comparison of 2D-[15N,1H] TROSY spectra (Pervushin et al., 1997Go) recorded for WT and {Delta}324G variants showed an impressive improvement of spectral quality for the engineered variant (Figure 5Go). Furthermore, the concentration of the engineered protein could be readily increased to 0.35 mM as opposed to 0.15 mM for the WT enzyme.



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Fig. 5. 2D-[15N,1H] TROSY spectra (750 MHz 1H resonance frequency) recorded for (A) WT and (B) engineered {Delta}324G HhaI MTases at concentrations of 150 and 250 µM, respectively. The poor signal-to-noise ratios of the cross peaks in the WT spectrum indicates line broadening due to protein–protein multimer formation (see text). For the engineered protein (B), high dispersion of cross peaks and average 15N spin relaxation times indicate the presence of a well-folded monomeric protein in solution.

 
Assuming isotropic rotational reorientation of the engineered {Delta}324G HhaI MTase, we then determined the correlation time for the overall rotational tumbling, {tau}c, from average backbone 15N spin T1 and T1{rho} relaxation times (Kay et al., 1989Go; Szyperski et al., 1993Go). The obtained T1 (1.9 ± 0.1 s) and T1{rho} (32 ± 1.5 ms) yield a {tau}c of about 25 ns for the engineered protein at 25°C. This value suggests that the 36 kDa {Delta}324G HhaI MTase does not aggregate in solution and T1{rho} (15N) relaxation times around 30 ms evidence that high-quality multidimensional NMR spectra can be recorded for future structural studies.

Long-term stability of the 0.35 mM {Delta}324G HhaI MTase complex with a 12-mer DNA duplex in solution was monitored by 1D 1H-NMR over a period of 2 weeks. No precipitation or loss of NMR signal was registered (data not shown). To verify the stability of the protein in the absence of DNA, the fluorescence titration analysis (as shown in Figure 3AGo) was performed following incubation of the 0.1 mM protein at ambient conditions. Our measurements showed no more than a few percent difference in the base flipping potential of the {Delta}324G MTase during the time span from 1 to 18 days (not shown). Retention of the measured parameters by the engineered protein under conditions pertinent to prolonged data collection demonstrates its suitability for structural NMR studies.


    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
With a molecular weight of 37 kDa, the solubility limit of 0.15 mM at 0.05 M NaCl presented an obvious obstacle for NMR-based structural studies of the HhaI MTase (Klimasauskas et al., 1998Go). Moreover, our experiments indicated poor behavior of the protein even at this concentration, with weak dispersion of resonances in 2D-TROSY spectra (Figure 5Go). Taking into account its tendency to produce smears in gel shift analyses (not shown) and complicated kinetic behavior at high protein concentrations (Figure 4Go), it is clear that the WT MTase is prone to intermolecular aggregation. Here we show that the major determinant of limited solubility and aggregation is the C-terminal tetrapeptide. This hydrophobic tail protrudes off the globule of the bilobal protein (Figure 1AGo) and appears to mediate intermolecular contacts in the crystal lattice of the ternary complex (Figure 1BGo). It is therefore possible that in a similar manner this hydrophobic tail promotes intermolecular interactions in solution leading to aggregation and low solubility. In retrospect, it is interesting that the WT protein can be maintained under certain conditions at concentrations (5–8 mg/ml) sufficient for successful crystallization of binary M.HhaI•AdoMet (Cheng et al., 1993Go; O’Gara et al., 1998Go) and ternary M.HhaI•DNA•AdoHcy complexes (Klimasauskas et al., 1994Go; O’Gara et al., 1996Go; Vilkaitis et al., 2000Go). Although it is widely believed that macromolecular aggregates in solution impede crystallization (McPherson, 1989Go), M.HhaI appears as an exception to the rule. One possible explanation to this phenomenon is that the C-terminal tail of M.HhaI is relatively rigid as it retains a similar fold in different crystal forms obtained (Cheng et al., 1993Go; Klimasauskas et al., 1994Go). Therefore, although promoting intermolecular association and macroscopic aggregation, the tail does not lead to structural heterogeneity at the monomer level and thus does not preclude the formation of an ordered crystal latice.

Removal of the C-terminal tetrapeptide leads to a significantly higher solubility of the protein. Consistent with a remote location of the tail with respect to the known functional sites in the enzyme, no functional role for it was detected in vitro. However, it cannot be excluded that this epitope is important for a yet unknown in vivo function, such as targeting to a predetermined cellular site (i.e. cell wall). Individual mutations at the two other sites had no detectable effect on the solubility of the protein (Table IGo). The V213S replacement had a slight effect on enzymatic parameters (2-fold increase in KmDNA) of the MTase (Table IIGo), which could probably be explained by the proximity of this residue to the DNA binding site (Figure 1AGo). Unexpectedly, the M51K mutation resulted in a lower solubility when combined with the C-terminal deletion in the protein (Table IGo, Figure 2Go). The reason for the observed ‘lack of additivity’ in the contribution from the M51K mutation is not clear.

In conclusion, this paper describes the first successful design, preparation and characterization of a highly soluble, functionally active variant of a DNA methyltransferase. The modifed HhaI MTase can be prepared in high yield (5 mg per liter of culture) as a pure isotope-labeled protein. Importantly, the modified variant showed the desired behavior during preliminary NMR experiments, indicating the presence of a well-folded and stable monomeric protein in solution. This now paves the way to detailed structural studies of this model cytosine-5 MTase and its interactions with the ligands employing NMR spectroscopy, stopped-flow fluorescence, isothermal calorimetry and other methods to further our understanding of the mechanism of this physiologically important enzymatic reaction in DNA.


    Acknowledgments
 
This work was supported in part by a NATO collaborative linkage grant (CLG 975806), grants from the State Science and Study Foundation of Lithuania (projects V-004 and V-023) and a start-up fund of the University at Buffalo, SUNY. The authors are grateful to the Howard Hughes Medical Institute, Volkswagen-Stiftung and MBI Fermentas for continued support.


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Cheng,X. and Roberts,R.J. (2001) Nucleic Acids Res., 29, 3784–3795.[Abstract/Free Full Text]

Cheng,X., Kumar,S., Posfai,J., Pflugrath,J.W. and Roberts,R.J. (1993) Cell, 74, 299–307.[ISI][Medline]

Das,D. and Georgiadis,M.M. (2001) Protein Sci., 10, 1936–1941.[Abstract/Free Full Text]

Dryden,D.T.F. (1999) S-Adenosylmethionine-dependent Methyltransferases: Structures and Functions. World Scientific, Singapore, pp. 283–340.

Gesell,J.J., Liu,D., Madison,V.S., Hesson,T., Wang,Y., Weber,P.C. and Wyss,D. (2001) Protein Eng., 14, 573–582.[Abstract/Free Full Text]

Goedecke,K., Pignot,M., Goody,R.S., Scheidig,A.J. and Weinhold,E. (2001) Nature Struct. Biol., 8, 121–125.[CrossRef][ISI][Medline]

Hollis,T., Ichikawa,Y. and Ellenberger,T. (2000) EMBO J., 19, 758–766.[Abstract/Free Full Text]

Holz,B., Klimasauskas,S., Serva,S. and Weinhold,E. (1998) Nucleic Acids Res., 26, 1076–1083.[Abstract/Free Full Text]

Jenkins,T.M., Hickman,A.B., Dyda,F., Ghirlando,R., Davies,D.R. and Craigie,R. (1995) Proc. Natl Acad. Sci. USA, 92, 6057–6061.[Abstract/Free Full Text]

Kay,L.E., Torchia,D.A. and Bax,A. (1989) Biochemistry, 28, 8972–8979.[ISI][Medline]

Klimasauskas,S., Nelson,J.L. and Roberts,R.J. (1991) Nucleic Acids Res., 19, 6183–6190.[Abstract]

Klimasauskas,S., Kumar,S., Roberts,R.J. and Cheng,X. (1994) Cell, 76, 357–369.[ISI][Medline]

Klimasauskas,S., Szyperski,T., Serva,S. and Wüthrich,K. (1998) EMBO J., 17, 317–324.[Abstract/Free Full Text]

Kumar,S., Cheng,X., Pflugrath,J.W. and Roberts,R.J. (1992) Biochemistry, 31, 8648–8653.[ISI][Medline]

Malissard,M. and Berger,E.G. (2001) Eur. J. Biochem., 268, 4352–4358.[Abstract/Free Full Text]

McPherson,A. (1989) Preparation and Analysis of Protein Crystals. Robert E. Krieger, Florida, p. 371.

Mol,C.D., Izumi,T., Mitra,S. and Tainer,J.A. (2000) Nature, 403, 451–456.[CrossRef][ISI][Medline]

O’Gara,M., Klimasauskas,S., Roberts,R.J. and Cheng,X. (1996) J. Mol. Biol., 261, 634–645.[CrossRef][ISI][Medline]

O’Gara,M., Horton,J.R., Roberts,R.J. and Cheng,X. (1998) Nature Struct. Biol., 5, 872–877.[CrossRef][ISI][Medline]

Orekhov,V.Y., Nolde,D.E., Golovanov,A.P., Korzhnev,D.M. and Arseniev,A.S. (1995) Appl. Magn. Reson., 9, 581–588.[ISI]

Pervushin,K., Riek,R., Wider,G. and Wüthrich,K. (1997) Proc. Natl Acad. Sci. USA, 94, 12366–12371.[Abstract/Free Full Text]

Roberts,R.J. and Cheng,X. (1998) Annu. Rev. Biochem., 67, 181–198.[CrossRef][ISI][Medline]

Sambrook,J. and Russell,D.W. (2001) Molecular Cloning, 3rd edn, Vol. 2. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, pp. 13.1–13.91.

Serva,S., Weinhold,E., Roberts,R.J. and Klimasauskas,S. (1998) Nucleic Acids Res., 26, 3473–3479.[Abstract/Free Full Text]

Swaminathan,C.P., Sankpal,U.T., Rao,D.N. and Surolia,A. (2002) J. Biol. Chem., 277, 4042–4049.[Abstract/Free Full Text]

Szyf,M. (1998) Cancer Metastasis Rev., 17, 219–231.[CrossRef][ISI][Medline]

Szyperski,T., Luginbühl,P., Otting,G., Güntert,P. and Wüthrich,K. (1993) J. Biomol. NMR, 3, 151–164.[ISI][Medline]

Varnai,P. and Lavery,R. (2002) J. Am. Chem. Soc., 124, 7272–7273.

Vertino,P.M. (1999 S-Adenosylmethionine-dependent Methyltransferases: Structures and Functions. World Scientific, Singapore, pp. 341–372.

Vilkaitis,G., Dong,A., Weinhold,E., Cheng,X. and Klimasauskas,S. (2000) J. Biol. Chem., 275, 38722–38730.[Abstract/Free Full Text]

Vilkaitis,G., Merkiene,E., Serva,S., Weinhold,E. and Klimasauskas,S. (2001) J. Biol. Chem., 276, 20924–20934.[Abstract/Free Full Text]

Wahnon,D.C., Shier,V.K. and Benkovic,S.J. (2001) J. Am. Chem. Soc., 123, 976–977.[CrossRef][ISI][Medline]

Wang,P., Nicklaus,M.C., Marquez,V.E., Brank,A.S., Christman,J.K., Banavali,N.K. and MacKerell,A.D.,Jr. (2000) J. Am. Chem. Soc., 122, 12422–12434.[CrossRef][ISI]

Wu,J.C. and Santi,D.V. (1987) J. Biol. Chem., 262, 4778–4786.[Abstract/Free Full Text]

Wu,J.C. and Santi,D.V. (1988) Nucleic Acids Res., 16, 703–717.[Abstract]

Zhou,L., Cheng,X., Connoly,B.A., Dickman,M.J., Hurd,P.J. and Hornby,D.P. (2002) J. Mol. Biol., 321, 591–599.[CrossRef][ISI][Medline]

Received September 6, 2002; revised January 20, 2003; accepted February 5, 2003.





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