Combinatorial exploration of the catalytic site of a drug-resistant dihydrofolate reductase: creating alternative functional configurations

Andreea R. Schmitzer1, François Lépine2 and Joelle N. Pelletier1,3

1Département de Chimie, Université de Montréal, C.P. 6128, Succursale Centre-ville, H3C 3J7, Montréal and 2INRS–Institut Armand-Frappier, 531 Boulevard des Prairies, H7V 1B7, Laval, Québec, Canada

3 To whom correspondence should be addressed. E-mail: joelle.pelletier{at}umontreal.ca


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
We have applied a global approach to enzyme active site exploration, where multiple mutations were introduced combinatorially at the active site of Type II R67 dihydrofolate reductase (R67 DHFR), creating numerous new active site environments within a constant framework. By this approach, we combinatorially modified all 16 principal amino acids that constitute the active site of this enzyme. This approach is fundamentally different from active site point mutation in that the native active site context is no longer accounted for. Among the 1536 combinatorially mutated active site variants of R67 DHFR we created, we selected and kinetically characterized three variants with highly altered active site compositions. We determined that they are of high fitness, as defined by a complex function consisting jointly of catalytic activity and resistance to trimethoprim. The kcat and KM values were similar to those for the native enzyme. The favourable {Delta}({Delta}G) values obtained (ranging from –0.72 to –1.08 kcal/mol) suggest that, despite their complex mutational pattern, no fundamental change in the catalytic mechanism has occurred. We illustrate that combinatorial active site mutagenesis can allow for the creation of compensatory mutations that could not be predicted and thus provides a route for more extensive exploration of functional sequence space than is allowed by point mutation.

Keywords: active site/combinatorial mutagenesis/fitness landscape/R67 dihydrofolate reductase/tetramer


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
Combinatorial mutagenesis and directed evolution approaches (Lutz and Patrick, 2004Go) are increasingly applied to modify enzymes both with respect to industrial applications and towards gaining a better understanding of enzyme catalysis. Previously, three key active site residues of the enzyme glycinamide ribonucleotide transformylase had been combinatorially mutated (Warren and Benkovic, 1997Go). While none of these residues was strictly essential for activity, that study demonstrated that only one mutation at a time could be tolerated, illustrating the difficult nature of active site modification. Here, we examined a more ‘primitive’ enzyme (Narayana et al., 1995Go), the bacterial R67 dihydrofolate reductase (DHFR) enzyme was identified as a result of its intrinsic resistance to the widely administered antibiotic trimethoprim (TMP) (Narayana et al., 1995Go). Dihydrofolate reductases (DHFR, EC 1.5.1.3) catalyse the reduction of dihydrofolate (DHF) to tetrahydrofolate (THF) by facilitating protonation at N5 of DHF and transfer of a hydride ion from NADPH to C6 of DHF (Scheme 1). As a result of its importance in key biosynthetic processes, chromosomally encoded DHFR has long been the target of antibiotic drugs such as TMP. It has been suggested that bacterial R67 DHFR has evolved recently in direct response to selective TMP pressure. As the catalytic efficiency of R67 DHFR is ~100 times lower than that of Escherichia coli chromosomal DHFR, R67 DHFR appears to have evolved as a poor mimic of the chromosomal enzyme. An important distinguishing feature is that R67 DHFR does not appear to possess a proton donor at the active site, as opposed to its chromosomal counterpart (Howell et al., 1986Go), which may account for its lower catalytic efficiency. However, R67 DHFR presents the crucial advantage of being intrinsically resistant to TMP. By providing discrimination between DHF and TMP, R67 DHFR provides the bacterial host with resistance to concentrations of TMP at which the chromosomal DHFR is sensitive.



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Scheme 1. (A) The substrates of DHFR for the reduction of 7,8-dihydrofolate (DHF) to 5,6,7,8-tetrahydrofolate (THF) using NADPH as the hydride-donating cofactor. DHF must be protonated at N5 for hydride transfer at C6 to occur. (B) Structure of trimethoprim (TMP), an inhibitor of chromosomal DHFR. R67 DHFR is intrinsically TMP-resistant.

 
Of particular interest to our combinatorial active site exploration, the R67 DHFR polypeptide folds and assembles as a tetramer, creating an active site pore (Figure 1). The active site residues are related by the 222 symmetry associated with the structure. However, the restricted volume of the cavity dictates that binding of a single substrate/cofactor pair (DHF/NADPH) can occur at a time, occupying only one of the four equivalent, symmetry-related active sites at any given time (Strader et al., 2001Go). The three ‘unoccupied’ sites are involved in binding both DHF and NADPH by additional contacts with the ligands (Narayana et al., 1995Go). Thus, the 16 principal active site residues are encoded by four residues per monomer: residues 66 to 69. As a result of the small yet symmetrical active site architecture, equivalent residues can play as many as four distinct roles toward substrate recognition, turnover and resistance to TMP. Any active site mutation can be expected to have a large cumulative effect since one mutation can result in four functional consequences. It follows that important evolutionary constraints must be imposed upon these active site residues. Not unexpectedly, point mutations at this active site are generally deleterious to function (Martinez et al., 1996Go; Park et al., 1997aGo; Strader et al., 2001Go). Despite this complexity, we demonstrate that mutation of a large number of neighbouring residues at a time yields heavily mutated variants of R67 DHFR that are functional.



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Fig. 1. Structure of R67 DHFR. (A) Crystal structure of the wild-type tetramer (1VIE pdb file) (Narayana et al., 1995Go). The four monomers (A–D) are illustrated in red, blue, green and yellow, respectively. Active site residues 66–69 are shown in ball-and-stick representation. The dimerization interface involves hydrophobic contacts (in grey) and the tetramerization interface involves acidic residues [His62, in pink (Dam et al., 2000Go)]; neither interface is proximal to the active site residues of interest. (B) Space-filled charge map of the modelled ternary R67 DHFR:DHF:NADPH complex, constructed as described under Materials and methods: negative charges are in red and positive charges are in blue. Residues 66–69 are represented in pink to highlight their contribution to the active site surface. (C) Space-filled view of the interior of the modelled ternary complex: relative positioning of DHF (green) and NADPH (yellow) at the interior of the active site. Two monomers are shown (monomers A and C), as described in the pictogram. Residues 66–69 are represented in translucent pink, such that their side chains can be distinguished. The images were generated using WebLabViewerPro software (version 3.7).

 
We have undertaken the combinatorial sequence exploration of all the 16 principal amino acids that constitute the active site of this enzyme. This combinatorial strategy of enzyme active site exploration should be generally applicable and rapidly provides information on specific active site residues and on the general active site environment.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
Reagents

All reagents were of the highest available purity. Synthetic oligonucleotides were obtained from AlphaDNA (Montréal, Canada) when unmodified, from Integrated DNA Technologies (Coralville, USA) when degenerate or from Li-Cor Biotechnology Division when dye-labelled for sequencing. 7,8-Dihydrofolate (DHF) was prepared as described previously (Blakley, 1960Go) or was purchased from Aldrich. Restriction enzymes and DNA-modifying enzymes were obtained from New England Biolabs and MBI Fermentas.

Molecular modelling

Energy minimizations and molecular dynamics simulations were carried out using the Discover module of InsightII (Accelrys) with the CFF91 and AMBER force fields using {varepsilon} = 80. Energy minimizations were typically computed with 10 000 iterations of an adjusted basis Newton–Raphson algorithm until an r.m.s.d. of 0.01 kcal/mol · Å was attained.

We modelled NADPH binding to R67 DHFR. Precise alignment between hydride donor (NADPH) and acceptor (DHF) have been shown to be critical determinants of the rate of hydride transfer in the chromosomal E.coli DHFR (Miller and Benkovic, 1998Go). The atom set of the folate-bound monomer (1VIF.PDB) was aligned with each of the four corresponding units in the free tetramer (1VIE.PDB); only one of the four identical folate molecules (on monomer A) was retained in the folate : R67 DHFR complex. It should be noted that the p-aminobenzoate-{gamma}-glutamate tail (PABA-Glu) of DHF was not observed in the crystal structure, indicating disorder (Narayana et al., 1995Go). To reproduce the same distance and angle between DHF and NADPH as in the chromosomal E.coli DHFR, we superimposed the pteridine ring of folate from the E.coli ternary complex (7DFR.PDB) with the pteridine ring in the folate:R67 DHFR complex. The entire folate molecule (heavy atoms) from the E.coli complex, including the PABA-Glu tail, was retained in the structure. It was modified to DHF, the hydrogen atoms were added and the DHF was energy minimized using the Steepest Descent (1000 steps) and the Conjugate Gradient (until convergence of 0.01 kcal/mol · Å) algorithms, yielding the tetrameric R67 DHFR:DHF binary complex. The nicotinamide portion of NADPH was positioned relative to the pteridine ring, in accordance with the E.coli DHFR ternary complex coordinates. A manual torsion was applied (nicotinamide N1–ribose C1 = –71.26°) to avoid steric clash of the NADPH 2',5'-adenine dinucleotide phosphate (2',5'-ADP) tail with the protein. The position of the pteridine ring was frozen relative to monomer A and the nicotinamide ring of NADPH relative to the pteridine ring, to ensure the retention of the same distance and angle between the pteridine ring and the nicotinamide ring as in the E.coli DHFR (Miller and Benkovic, 1998Go). A molecular dynamics simulation was undertaken. The system was heated to 300 K over 1 ps and kept at that temperature for a 1 ps equilibration and a 10 ps (or longer) dynamics simulation. The system was then energy minimized. In this way, DHF (including the PABA-Glu tail) and NADPH (including the 2',5'-ADP tail) were positioned at the active site (Figure 1B and C).

When mutated forms of R67 DHFR were modelled, the mutations were introduced in the 1VIF.PDB file using the Biopolymer module. We performed an energy minimization of 1000 iterations of an adjusted basis Newton–Raphson algorithm, during which the protein backbone was fixed in order to maintain the global conformation of the protein. In all molecular modelling studies we did not consider the N-terminal (MRGSHHHHHHGIH) and C-terminal (ELGTPGRPAAKLN) tails.

Bacterial strains and plasmids

The R67 DHFR wild-type (WT) gene in pET19b plasmid was a generous gift from Dr A.Blondel (Institut Pasteur, France). Escherichia coli BL21 containing plasmid pRep4 (Qiagen, Mississauga, Ontario, Canada, lac Iq) was used for expression. Ampicillin (Amp) was used at 100 µg/ml and kanamycin (Kan) at 50 µg/ml for maintenance of plasmids pQE32 and pRep4, respectively, while isopropyl ß-D-1-thiogalactopyranoside (IPTG) was used at 100 µM to induce protein expression. TMP was used at 70 µg/ml during selection.

Site-directed mutagenesis

The 234 bp WT R67 DHFR gene was the template for mutagenic PCR (Cadwell and Joyce, 1994Go). Primers 5' CACGGATCCATATGGAACGAAGTAGCAATGA (forward) and 5' ACACACGAGCTCGTTGATGCGTTCCAACGCCGCAACAGGAWRVNWMTYANYTGAGCCTGGGTGAGCCT (reverse) introduced the desired mutations and BamHI and SacI restriction sites (underlined). PCR was carried out using 1 µg of template R67 DHFR DNA, 0.2 mM of each dNTP, 1x PCR buffer (MBI Fermentas), 2.5 mM MgCl2, 1.5 µM of each primer and 2.5 U Taq polymerase (MBI Fermentas) in a total volume of 100 ml. Each reaction cycle consisted of a 2 min denaturation at 94°C followed by annealing at 52.8°C for 30 s and extension at 72°C for 1 min, for 30 cycles. A final extension step of 3 min ended the program. The BamHI–SacI-digested PCR fragment was isolated on 2% agarose gel. The appropriate band was excised and the DNA extracted. The BamHI–SacI-digested PCR fragment was cloned into pQE32 (Qiagen). The resulting library was electroporated into E.coli BL21/pRep4. The transformants were plated on 1.2% agar Terrific Broth medium (TB) (Tartof and Hobbs, 1987Go) containing Amp and Kan and were incubated at 37°C for 16 h to allow colony formation. The resulting constructs encode mutated R67 DHFR containing an N-terminal six-histidine tag (MRGSHHHHHHGIH) to be used in enzyme purification and a 13-residue C-terminal tail (ELGTPGRPAAKLN) which was introduced as a linker to allow the ulterior genetic duplication of the mutated library.

Screening and selection

Each of 5000 colonies from the TB plate was picked in 100 µl of minimal M9c medium. The quality of the library was verified by sequencing the entire gene for 20 clones before selection. DNA sequencing was performed using a Thermo Sequenase Cycle Sequencing kit (USB) with a LiCor IR2 sequencer. After gentle agitation, 2.5 µl were spotted on to TB plates containing Amp and Kan and also on to M9c minimal medium plates containing Amp, Kan, IPTG and 70 µg/ml TMP [selective minimal medium (Sambrook et al., 1989Go)]. Escherichia coli chromosomal DHFR is inhibited by 0.1 µg/ml TMP. The cells were allowed to grow at 37°C for 14 h on TB plates or 36 h on M9c plates, at which time full-size colonies were obtained.

Serial rounds of selection We subjected variants of interest to serial rounds of selection by first plating each variant on selective minimal medium. Individual colonies were then picked and the plasmids were isolated, retransformed into fresh cells and plated again on selective minimal medium.

The quality of the library was verified by sequencing the entire gene for 20 clones before selection. Using Nln(1 – 1/n) = ln(1 – P), where N is the number of clones sequenced, 1/n is the fractional proportion of each codon at each residue and P is the probability of finding any given codon among those sequenced, the theoretical codon representation was compared to the observed codon representation. By applying {lambda} = n[1 – (1/n)]m where n is the number of theoretical library members and m is the number of clones experimentally screened, we determined {lambda}, the number of theoretical library members not sampled (Ross, 1996Go).

Protein overexpression and purification

All purification procedures were carried out at 4°C. Wild-type (WT) R67 DHFR and variants of interest were over-expressed by inoculation of 100 ml of TB medium + Amp + Kan with 1 ml of overnight culture. At OD600 = 0.8, IPTG was added. After 3 h, cells were harvested by centrifugation at 3500 g and resuspended in 3 volumes of cold lysis buffer [50 mM KH2PO4–K2HPO4 pH 8.0, 0.5 mM phenyl methyl sulfonyl fluoride (PMSF)]. The cells were disrupted by sonication for 2 x 30 s at 2 min intervals and centrifuged for 20 min at 3800 g. The supernatant was applied at 1 ml/min to a 1 x 10 cm column of DEAE-Sepharose Fast Flow (Sigma) previously equilibrated with wash buffer (10 mM KH2PO4–K2HPO4 pH 7.6, 1 mM EDTA, 20 mM ß-mercaptoethanol). The column was washed with 2 volumes of wash buffer, brought up to 0.1 M KCl in 10 min followed by a linear gradient elution from 0.1 to 0.3 M KCl in 60 min at 1 ml/min. Fractions exhibiting DHFR activity (Horecker, 1948Go) were analysed by tricine–SDS–PAGE (Schägger and Jagow, 1987Go) and samples showing ≥60% purity were selected for kinetic characterization. Enzyme purity was quantitated by densitometric analysis of the scanned gel with the ImageJ software (public domain Java image processing program). The protein concentration was determined by the Bradford assay (Bradford, 1976Go).

Denaturating purification Following overexpression, cells were lysed as described above except in 100 mM KH2PO4–K2HPO4, 10 mM Tris–Cl buffer containing 8 M urea, pH 8.0, at room temperature. The supernatant was loaded on to a 1 ml Ni-NTA column (Qiagen). The column was washed with urea buffer at pH 6.3 and the enzyme was eluted with the urea buffer at pH 4.5, according to the manufacturer's protocol.

Mass spectral characterization of R67 DHFR variants

The partially purified enzymes (1 ml), obtained under native conditions, were dialysed against 8 l of 18.2 M{Omega} Millipore water overnight. Samples were diluted to 1 µg/ml in water and further purified on C4 ZipTip (Millipore). Samples were eluted in 10 µl of 60% (v/v) acetonitrile with 0.1% (v/v) trifluoroacetic acid, transferred into a nanoelectrospray needle (Micromass) and analysed using a Micromass Quattro II mass spectrometer. The source temperature was 80°C. A potential of 0.7 kV was applied to the needle and 30 V were applied as the cone voltage. Argon at a pressure of 2 x 10–3 mTorr was the collision gas and the collision energy was 33 eV. Data processing was performed using the Mass-Lynx Windows NT system (Micromass).

Size-exclusion chromatography

Size-exclusion chromatography was carried out at 4°C and pH 7.0 using a Pharmacia FPLC and a 1.5 x 28 cm Superose 12 column equilibrated in 50 mM MES–100 mM Tris–50 mM acetic acid–10 mM ß-mercaptoethanol (MTA) buffer. The flow rate was 0.75 ml/min. Standard curves at pH 7.0 were produced by plotting the logarithm of the molecular mass of protein standards (aprotinin, lysozyme, carbonic anhydrase, chicken serum albumin, bovine serum albumin) and fluorescein versus Kav. This allowed the determination of the molecular mass and thus the oligomeric state of each R67 DHFR active mutant. WT R67 DHFR (with N- and C-terminal tails) was used as a positive control. Kav is defined as

where VE is the elution volume, VV is the void volume and VB is the bed volume of the column matrix.

Kinetic characterization

Spectrophotometric enzyme activity assays (Horecker, 1948Go) were performed using purified enzyme (60–95% pure, purified under native conditions). The negative control was a mock purification using untransformed cells. Assays were performed in the presence of 50 nM TMP (14.5 pg/ml) for inhibition of any E.coli chromosomal DHFR that may have copurified, although controls showed only trace activity from this source. The extinction coefficients used were: 28 400 M–1 cm–1 at 282 nm for DHF (Blakley, 1960Go), 6220 M–1 cm–1 at 340 nm for NADPH (Horecker and Kornberg, 1948Go) and 12 300 M–1 cm–1 (Strader, 2001Go) at 340 nm to determine product formation. Enzyme concentration was at least 100-fold lower than the limiting substrate concentration. Initial rates were measured under conditions where <15% substrate conversion to product had occurred. Unless specified otherwise, standard fixed concentrations were 50 µM NADPH and 50 µM DHF. The concentration of a single component (either NADPH or DHF) varied from ~0.1 x KM to ~10 x KM while keeping all others constant. Assays were carried out in triplicate using enzyme from independent purifications. The data were fitted to the Michaelis–Menten equation using non-linear regression analysis with the software GraphPad (version 3.01).

Transition state stabilization by the binding energy The changes in substrate binding energy at the transition states for WT and mutant enzymes (Wilkinson et al., 1983Go) were calculated as


    Results and discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
We used a semi-rational approach to introduce a sub-set of the possible mutations at all the principal active site residues of R67 DHFR. In order to select the active site residues of interest for combinatorial mutagenesis, we needed to define the residues in contact with the substrate and cofactor. The structure of NADPH-bound R67 DHFR has not been resolved. The structural information available for R67 DHFR consists of the coordinates of the free (apoenzyme) tetramer and a binary complex with two folates bound (1VIE and 1VIF at the Protein Data Bank) (Narayana et al., 1995Go). More recent binding studies indicate that, whereas two DHF or two NADPH molecules can bind to the enzyme (non-productive complexes), the productive ternary complex stoichiometry is 1:1:1 DHF:NADPH:R67 DHFR (Bradrick et al., 1996aGo).

A model of the ternary folate:NADPH: R67 DHFR complex obtained by DOCK and SLIDE docking methods has recently been reported (Howell et al., 2001Go). The head-to-tail positioning of the two ligands with favourable juxtaposition of the reactive rings near the centre of the cavity is predicted in this model. We performed a modelling study of the ternary complex using distinct methodologies and obtained very similar results (Figure 1B and C), further validating the model of the ternary complex. Briefly, our model is based on the hypothesis that the relative position of the reactive rings of DHF and NADPH (pteridine and nicotinamide, respectively; Scheme 1) is similar to that determined in the crystal structure of the folate:NADPH:E.coli chromosomal DHFR ternary complex (Miller and Benkovic, 1998Go). Our approach made use of the crystal-structure coordinates of R67-bound folate to model bound DHF. Using InsightII software (Accelrys), a constraint was applied to position the NMN moiety of NADPH relative to the pteridine ring in accordance with their coordinates in the E.coli DHFR, as detailed under Materials and methods and an energy minimization was performed on the ternary complex (Figure 1B and C). Visual inspection of the model created in the previously reported docking study (Howell et al., 2001Go), our structure-based model and additional biochemical data (Nichols et al., 1993Go; Park et al., 1997aGo,bGo; Strader et al., 2001Go; Smiley et al., 2002Go) were used to define the residues to mutate in our exploration of ligand binding and reactivity in R67 DHFR.

Substrate and cofactor binding: residues near the reactive substrate atoms

R67 DHFR is active as a homotetramer (monomers A, B, C and D; Figure 1A), possessing a central pore (Narayana et al., 1995Go). On the basis of the available data, we defined the residues within the central cavity having contacts with either ligand and most likely to promote reactivity, residues 66–69 of the four monomers, as the 16 ‘principal active site residues’. These residues form the majority of the active site surface and are proximal to the site of hydride-transfer (Figure 1B and C) and were thus selected for mutagenesis. Residues near the centre of symmetry interact substantially with each ligand in pairs (Gln67 and Ile68), while residues further away from the symmetry operator form individual binding surfaces (Val66 and Tyr69) (Narayana et al., 1995Go).

Based on previous results from other research groups (Howell et al., 1986Go; Dam et al., 2000Go; West et al., 2000Go), we selected a number of active site functionalities for modification, namely the potential contribution of a proton donor to catalysis, dielectric environment and transition state stabilization. There is no evident proton donor in the active site pore of R67 DHFR. The active site residue Tyr69 is unlikely to fulfil this role as none of the four symmetrical hydroxyl groups appears to lie proximal to the pteroyl ring of DHF. In the modelled ternary complex (Howell et al., 2001Go; this study), residue 67, which protrudes the most into the active site cavity and is proximal to the site of hydride transfer, could advantageously carry an acidic proton. In an effort to increase the catalytic efficiency of R67 DHFR, we replaced Gln67 by potential proton donors (Glu and Asp, to mimic the proton-donating Glu of chromosomal DHFR); we also encoded the conservative mutation Gln to Asn to determine the effect of decreasing the side-chain length. Potential proton donors were also encoded at positions 66, 68 and 69 (Table I) although these side-chains do not project into the active site cavity as does residue 67.


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Table I. Mutations introduced combinatorially in R67 DHFR

 
In the chromosomal DHFR, charged active site residues increase the tautomerization state of DHF, promoting substrate binding and hydride transfer (Miller and Benkovic, 1998Go). We introduced mutations that are predicted to alter the dielectric environment of the active site: Val66Asp, Gln67Lys, Ile68Lys, Ile68Arg and Tyr69Phe. Moreover, the side chain of residue 68 may play a role in solvent exclusion (Howell et al., 2001Go). By introducing bulky residues at this position, we may decrease the size of the active site pore and favour the approach between the cofactor and the substrate. We also encoded a variety of hydrophobic residues (Ala, Val, Ile at position 66; Ile, Phe, Leu, Trp at position 68; Leu, Phe at position 69) to promote binding of the DHF pteridine ring and the NADPH NMN moiety, which should bind in a hydrophobic environment.

It should be noted that there is no evidence from the models of the ternary complex implicating residues outside of the pocket defined by residues 66–69 in specific contact with the pteroyl ring of the substrate or the nicotinamide moiety of the cofactor (Brito et al., 1991Go; Nichols et al., 1993Go; Howell et al., 2001Go; Strader et al., 2001Go; Hicks et al., 2004Go; this study). Consequently, we did not include any amino acids outside the active site cavity in our library design. This restriction has the benefit of reducing the likelihood of adversely affecting oligomerization, as the residues involved in the dimerization and the tetramerization interfaces are not located in the vicinity of the active site (Novak et al., 1983Go; Dam et al., 2000Go; West et al., 2000Go; Mejean et al., 2001Go) (Figure 1).

Library design and evaluation of library quality

We designed the combinatorial mutation of all the principal active site residues, 66–69. This yields mutations at 16 positions within the tetrameric active site. Because oligomerization occurs in vivo, homotetrameric mutants will be obtained rather than mixed active sites. Our long-term goal is the creation of heterotetrameric mutants, by genetic fusion of mutated monomeric libraries. In order to restrict the size of a future, combinatorial heterotetrameric library such that adequate sampling will be possible, we created a restricted monomer library of 1536 combinatorially mutated enzyme variants of 28 possible individual mutations spanning residues 66–69 (eight amino acids encoded at position 66 + four at position 67 + 12 at position 68 + four at position 69, Table I). A schematic description of the sequence space covered by the library is provided in Figure 2. Relatively few conservative mutations were encoded as our goal was to create a complex set of widely differing active site environments. We excluded the WT sequence to avoid its domination during selection. The mutation of Gln67 exclusively to alternate amino acids presented the greatest interest as a result of its prominent position in the active site cavity (Figure 1A). Thus, the native residues were allowed only at positions 66, 68 and 69. We included a safeguard against possible contamination by native sequence by inserting a tracer in the mutagenic oligonucleotides under the form of a silent mutation at codon 74 [CTT (Leu) to TTG (Leu)].



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Fig. 2. Schematic representation of the sequence space covered by the 1536 R67 DHFR variants created, as described in the text. The central column represents WT R67 DHFR; column height (z axis) is a non-quantitative representation of fitness. Mutational distance is measured in the (x, y) plane where each concentric circle is one mutation away from the WT R67 DHFR. The library encodes four point mutants (Gln67Asp, Gln67Glu, Gln67Lys and Gln67Asn ~0.2% of the clones) of no detectable fitness, shown as black circles on the first concentric ring. Double mutants where two positions (including 67) are mutated occur at a frequency of ~5% (second ring). Triple and quadruple mutants (third and fourth rings, respectively) occur in 34% and in almost 60%, respectively, of all variants. Two triple (M10; M36) and one quadruple mutant (M8_30) of native-like fitness were identified and characterized (Table II). Variants NS15 and NS19 were identified prior to selection and have no detectable activity.

 

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Table II. Steady-state kinetic parameters for R67 DHFR variants at pH 7.0 and 300 K

 
Following mutagenic PCR and insertion into an inducible expression vector, a plasmid library containing 5 x 106 variants was obtained. DNA sequencing revealed no deviation from expected compositions at degenerate codon positions from 20 clones picked prior to selection, confirming that library construction was successful (Table III, available as Supplementary data at PEDS Online). There was no bias toward the allowed WT codon for residues 66, 68 and 69. No unintentional mutations or frame shifts were observed.

Library selection yields highly mutated, functional variants

Growth of E.coli expressing R67 DHFR in the presence of TMP is an efficient means of selecting for active and TMP-resistant R67 DHFR variants. Five thousand clones [~96% coverage of the 1536 theoretical variants (Ross, 1996Go)] obtained on non-selective TB plates were picked for inoculation of equal aliquots on TB plates and on selective minimal medium plates with TMP. This selection strategy can exclude active variants that are not resistant to TMP; subsequent structure–function analysis considers this limitation. We verified that the WT enzyme allows colony formation under these conditions. Library selection at TMP concentrations ranging from 1 to 70 µg/ml yielded no observable change in the selection stringency. It was not possible to select at lower TMP concentrations (i.e. at lower stringency) in this system as the endogenous E.coli DHFR would no longer be efficiently inhibited. A similar selection strategy was previously applied by Strader et al. (2001)Go, using a pUC8-based plasmid in E.coli SK383 with 20 µg/ml TMP in a modified version of TB medium.

Selection yielded four clones that encode three functional variants (Table II, Figure 3: M10, M36 and M8_30, where the latter was identified twice). The selected sequences carry three (M10 and M36) or four (M8_30) mutations per monomer, yielding 12 or 16 mutations per tetrameric active site. Screening a greater amino acid diversity at these active site positions could yield a larger number of functional variants with sequences as divergent again as those we have identified.



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Fig. 3. Sequences of the functional variants. (A) Sequence alignment of the expressed His-6-WT R67 DHFR and the mutated, functional variants as confirmed by mass spectrometry. The features of residues 66–69 are colour coded as indicated in the legend. (B) Molecular replacement of the side chains of the active site residues allowing visualization of the side-chain changes in the backdrop of the native structure. The images were generated using 1VIF.PDB file and the Biopolymer module of InsightII, followed by an energy minimization.

 
DNA sequencing revealed no trace of WT after four serial rounds of selection. That the activity recorded cannot be attributed to a cellular acquisition of TMP resistance was established by retransforming the mutant plasmids after each round and plating on selective medium. No unexpected changes were observed in their DNA sequence, confirming that growth results from plasmid-encoded R67 DHFR activity. During over-expression, DNA was extracted post-induction and sequenced. No sequence variations were observed. The expression levels and solubility of the active variants were similar to the WT although the expression of mutant M10 was 2- to 3-fold weaker, excluding better expression as a mechanism for increased activity in vivo (data not shown).

Mass spectral and kinetic characterization of the selected variants confirms the identity of highly active, multiply mutated variants

In order to confirm the identity of the selected variants, the mass of the hexahistidine-tagged WT (11315 Da) was determined by positive nano-electrospray (ESI) mass spectrometry (MS) and by MS/MS (data not shown). This proved to be very informative as the mass of the mutants under the same ionization conditions indicated that N-terminal cleavage of the non-native, 13-residue His-tag sequence had occurred (Figure 3A). These results are consistent with faster electrophoretic migration of the mutants (results not shown) and with the inefficient binding of the mutants on Ni-NTA agarose compared with the WT (results not shown). For this reason, enzyme purification using classical methods under native conditions was required. No peaks corresponding to the WT protein were detected in the mass spectra of the mutants, although the ESI-MS approach was sensitive enough to detect trace WT protein in a different mutant where the WT band could not be detected by electrophoresis on tricine gel but was suggested by traces in the DNA sequencing (data not shown). Similar N-terminal cleavage has been observed for an active site R67 DHFR point mutant, with no effect on activity (VerBerkmoes et al., 2002Go).

The kinetic parameters obtained for the N-terminally hexahistidine-tagged WT R67 DHFR containing the silent mutation at position 74 and a 13-residue C-terminal tail (Table II) were essentially identical with those previously reported for native enzyme (Dam et al., 2000Go). This is consistent with previous reports of terminal modifications of R67 DHFR (Reece et al., 1991Go; Bradrick et al., 1996bGo) where native-like activity was observed, confirming the structural tolerance of R67 DHFR to concomitant N- and C-terminal modifications. This tolerance may result from the termini protruding from the outer face of the tetrameric structure while the active site is buried in the central cavity. Hence, although the selected variants lack 13 non-native residues at the N-terminus relative to the WT (Figure 3), the deletion should have no observable effect on kinetic parameters (Reece et al., 1991Go). The selected variants were at least as active as the WT (Table II), with kcat increased by 1.5–3.9-fold and KM (DHF) values decreased by 1.4 to 3-fold. The catalytic efficiencies (kcat/KM) were 3.4 to 6.2-fold that of native R67 DHFR. Hence the heavily mutated active site environments of mutants M10, M36 and M8_30 allow for native-like kinetic characteristics. It is likely that the selection conditions applied are responsible for the catalytic efficiencies being at least as high as the native enzyme. However, the available data do not allow us to correlate further the in vitro kinetic characteristics with the event of in vivo selection.

We did not identify point mutants or double mutants with sufficient catalytic efficiency and TMP resistance to promote bacterial propagation under selective conditions. While our sampling of point mutants was very limited (four possibilities tested; Table I; Figure 2) and biased, since it was performed uniquely at position 67, previous characterization of point mutants at position 67 showed kcat decreases of 13- and 60-fold (Q67C and Q67H, respectively) (Strader et al., 2001Go). If the four point mutants we created (Q67D, Q67E, Q67K and Q67N) affect kcat in such a way, the resulting fitness may be insufficient to support bacterial propagation in our selection system.

The functional mutants are tetrameric

We verified whether the changes in catalytic efficiency described above might be due to alterations in the oligomeric state of the functional mutated R67 DHFRs. The elution patterns on a Superose 12 size-exclusion column were determined at pH 7.0 for mutants M36 and M8_30. Their apparent molecular masses were within the range expected for the tetrameric state of the masses determined by MS, confirming that the mutant enzymes assemble predominantly as homotetramers (Table II). No significant dimer or monomer peaks were observed. Mutant M10 was not analysed as its slightly lower expression level resulted in a concentration, once purified, too low to obtain a readily interpretable signal. The WT R67 DHFR with the N- and C-terminal tails also eluted at the molecular mass corresponding to its tetrameric state. These data suggest that the mutations, located at the inner face of the tetrameric active site pore, do not perturb the dimerization and the tetramerization interfaces (Figure 1A) and that catalysis occurs within heavily mutated tetrameric active sites.

Important active site differences reveal strong functional coupling

In order to identify amino acid determinants that are essential to the maintenance of catalytic activity and TMP resistance, we compared the obtained active site sequences with the WT sequence. The primary sequence alignment of the active variants (Figure 3A) was observed. While mutant M36 has conserved Ile68 and mutant M10 has conserved Tyr69, relative to the WT, no amino acid is strictly essential for function. This is clearly indicated by the highly functional, quadruple mutant M8_30.

In the absence of sequence identity at any position, we attempted to identify similarities between the amino acids selected at each position for the three functional variants and the WT (Figure 3A). At position 66, neither size nor hydrophobicity was constant as small (Gly or Ser) or medium (Val or Ile), hydrophilic (Ser) or hydrophobic (Val or Ile) residues were selected, from the residues encoded (Ala, Asn, Asp, Gly, Ile, Ser, Thr or Val; WT = Val). At position 67, where hydrophilic residues Asn, Asp, Glu and Lys were encoded, Asn, Glu or Lys were selected (WT = Gln). Thus, the most conservative change to Asn was not specifically enriched. Position 67 can thus carry a charge, either positive (Lys) or negative (Glu) or no charge (Gln or Asn). At position 68 where a large variety of residues was encoded (Asn, Arg, Cys, Ile, Leu, Lys, Met, Phe, Ser, Thr, Trp or Tyr; WT = Ile), aliphatic residues (Ile or Leu) or Arg were selected. Although, at first glance, Arg stands out as an outlier, its long aliphatic chain may provide it with properties similar to an aliphatic residue. Thus, a residue with important aliphatic character may be favoured at position 68. Finally, at position 69, His (twice) or the WT Tyr were selected whereas Phe and Leu were not. Aromaticity may play a role at this position, although lighter sampling of non-aromatic residues at this position (Leu69 was encoded in 384 among the 1536 variants tested) precludes a definite functional assignment. It may be that the additional polarity or volume conferred by the side chain of Tyr or His relative to Phe contribute to function although there are insufficient data to conclude on this point. Overall, there are no obvious conservational patterns nor retention of WT residues at the four positions varied.

Mutating individual active site residues is frequently deleterious to function, resulting in a large effect on fitness (Voigt et al., 2000Go; Aita et al., 2001Go). As a result, the fitness landscape describing the active site sequence space of most enzymes is considered to be rugged (Voigt et al., 2001aGo,bGo). The shape of a fitness landscape consists of peaks identifying sequences of high fitness and valleys for those of low fitness. The underlying cause of landscape ruggedness at active sites is ‘functional coupling’ between residues, which defines the degree to which residues must be altered simultaneously when their individual mutation results in non-additive effects on function. The absence of simple or conservative mutational patterns strongly supports a high degree of functional coupling between the active site residues of R67 DHFR, particularly with respect to position 67, which was mutated in all variants we created. Further evidence of functional coupling is provided by the fact that, for each of the three functional variants, multiple point mutants exist but their function was insufficient to result in selection. For example, variant M36 (Figure 3) encodes Ser66, Lys67, Ile68 and His69. The library contained seven point mutants of this sequence at position 66, three at position 67, 11 at position 68 and three at position 69, none of which was selected (Table I). Thus, in the context of the environment created by Ser66, Lys67 and Ile68, the residue Tyr69 (for example) does not provide a functional active site environment as the variant was not selected. Since we obtained almost full library coverage during the selection process, almost all (~96%) mutants were tested for function, justifying analysis of the sequences not selected. We obtained a point mutant (NS19: Ile68Lys relative to M36, where NS stands for ‘non-selected’) and a double mutant (NS15) of variant M36 while sequencing the library prior to selection (Table III). Variant NS19 is at the same time a double mutant of variant M8_30. NS19 is not functional according to in vivo and in vitro assays, indicating that point mutations to the functional variants can lead to a dramatic loss of activity. This is analogous to the great perturbations that have been observed upon analysis of certain active site point mutations of R67 DHFR (Strader et al., 2001Go). NS19 and NS15 were inactive in vitro even in the absence of TMP, indicating a greater functional perturbation of the point mutation than only the loss of TMP resistance. Thus, the multiple point mutants of each functional variant were not selected although the library encoded numerous conservative mutations of the selected variants. This suggests that the active site residues in R67 DHFR, whether in the context of the WT enzyme or in functional variants, profit from a high degree of functional coupling where the destructive nature of a point mutation must be balanced by additional, compensatory mutations. The homotetrameric nature of the active site is likely to be at the source of this effect as mutation of one residue per monomer results in four mutations at the homotetrameric active site. The four mutations can have as many as four distinct effects on function, since the substrate-bound homotetramer is no longer symmetrical. Thus, point mutations have a high potential for disrupting function.

In terms of the fitness landscape describing R67 DHFR function, our results illustrate that walking from the WT to the selected active site environments requires traversing sequence space with function too low to be detected in our system, between very sharply-defined points of high fitness (Figure 2). Thus, all non-selected variants constitute the flat ‘plain’ of the landscape; their catalytic activity is insufficient and/or they are TMP-sensitive, such that they are too weakly functional to be selected. The non-selected variants NS-15 and NS-19 (Table II) belong to this population. The three selected, functional variants stand as isolated pillars among their non-selected though closely related neighbours. Their representation as pillars rather than tapered peaks is accurate in that, within the sequence space we explored, any step away from the top of a pillar (or any mutation away from a functional sequence) results in a sharp drop in activity, down to the level of the ‘plain’ of undetectable function, rather than a gradual descent toward lower activity as would be the case for simple, additive mutations.

Compensatory mutations provide native-like transition-state stabilization

Transition state stabilization in R67 DHFR may be achieved by forcing C4 of the NADPH nicotinamide ring proximal to C6 of the DHF pteroyl ring as in the case of chromosomal DHFR (Miller and Benkovic, 1998Go). Visual inspection of the modelled structure of the native enzyme suggests that the backbone of Ile68 may participate in this role via H-bonding with the nicotinamide ring and in solvent exclusion (Figure 1C). From the large variety encoded at position 68, bulky residues were selected (Figure 3: Ile, Leu or Arg) that may position the backbone so as to favour the approach between cofactor and substrate.

For the three functional variants, the changes in the transition state energy relative to the WT [{Delta}({Delta}G)] are slightly negative (Table II), indicating little change in transition state stabilization between the WT and the active variants. Since no side-chain is conserved among the selected variants, this supports a model for catalysis where the transition state is mainly controlled by the good relative positioning of the substrate and the cofactor at the active site rather than by direct participation of specific side-chains to catalysis. These results contrast with the positive {Delta}({Delta}G) values determined previously for a variety of weakly active point mutants (Strader et al., 2001Go) of R67 DHFR. We have also identified important deviations from simple additivity in the effect of each mutation on activity. Specifically, the point mutants Ile68Leu and Tyr69His which have previously been reported (Strader et al., 2001Go) result in {Delta}({Delta}G) values of +1.2 and +3.44 kcal/mol, respectively (Strader et al., 2001Go; Table II), while variant M8_30 containing both mutations (as well as two more) has a {Delta}({Delta}G) of –0.97 kcal/mol (Table II). Variant M36 also encodes the mutation Tyr69His with two further mutations and has a {Delta}({Delta}G) of –1.08 kcal/mol. Phe was not selected at position 69 in the context of additional mutations although the point mutant Tyr69Phe shows only a modest reduction in catalytic efficiency relative to the WT (Strader et al., 2001Go; Table II). These results highlight the compensatory nature of the neighbouring mutations and provide evidence of strong functional coupling between the active site residues of WT or functional variants of R67 DHFR.

There is no direct evidence of proton donation occurring at the active site cavity

It has been suggested that the similar turnover rates at pH 7 for native R67 DHFR and a mutant of E.coli chromosomal DHFR (Asp27Ser) that cannot protonate DHF [25 vs 74 min–1, respectively (Howell et al., 1987Go)] may indicate the upper boundary of DHFR activity in the absence of a proton donor and may reflect rate-limiting diffusion and binding of protonated DHF as substrate (Holland et al., 1993Go). The increased activity of the native R67 DHFR at low pH suggests that pre-protonated DHF acts as its actual substrate (Holland et al., 1993Go; Strader et al., 2001Go). Since the pKa of DHF N5 is 2.59, then under standard reaction conditions at pH 7 the DHF/DHF-H+ ratio is 2.5 x 104. If protonated DHF is the actual substrate, the KM measured for DHF is an apparent KM for protonated DHF; the actual KM for protonated DHF is obtained by dividing KM (apparent) by 2.5 x 104, yielding KMs in the nM range for the WT and the functional variants. Considering this, the second-order rate constant (kcat/KM) for the WT and the functional variants then is in the range 1010 M–1 s–1, which surpasses the limit of diffusion control (Fersht, 1985Go; Jencks, 1987Go). Considering the ‘primitive’ nature of this enzyme, which is unlikely to achieve catalytic perfection, these data suggest that native R67 DHFR may provide rate-limiting protonation.

In order to overcome the limited potential for protonation of DHF in solution, we encoded potential proton donors in a variety of active site environments (Table I), particularly at position 67 (Asp and Glu) because its side chain protrudes into the active site cavity (Figure 1A). Surprisingly, at position 67, the non-conservative lysine was selected once as were asparagine and glutamate (Table II). It is striking that each of the functional variants encodes at least one potential proton-donating residue (Lys, Arg, His, Tyr, Glu). The catalytic efficiency of all functional variants was similar, suggesting that any protonation step would need to occur with roughly the same kinetics if it were rate limiting. Variant M36, with Lys67, displays a catalytic efficiency in the same range as the WT, indicating that the active site can accommodate four positive charges although the length of the side chain (~8 Å when fully extended) could allow the aliphatic portion to line the hydrophobic wall, with the charged amines near the mouths of the channel rather than at the centre of the active site pore. Hence DHF may actually be protonated within the enzyme, in the WT and in these variants. If protonation occurs within the active site cavity, we must consider that the nature and the position of the proton-donating residue are different in each variant, implying a unique proton-donating mechanism in each case. However, protonation could occur as DHF enters the active site channel, prior to taking up its position near the centre of the active site, in which case the active site mutations would not be expected to alter this step. The data provided here and in previous experiments do not allow us to conclude on this point.

The dielectric environment is partially, but not strictly, conserved in the selected variants

To probe the impact of the dielectric environment on binding specificity (DHF vs TMP) and catalysis, we introduced mutations that are predicted to alter the dielectric environment of the active site. We performed molecular replacement of the side chains of residues 66–69 (Figure 3B) to allow a visual assessment of the volume and polarity of the residues composing the new tetrameric active sites. While these active site representations are simple energy minimisations of molecular replacements in WT R67 DHFR and do not provide structural data, they provide a view of the chemical composition of the functional active sites. As qualitatively assessed from the nature of the active site mutations (Figure 3B), the three functional variants appear to exhibit a similar degree of active site hydrophobicity and may be moderately more hydrophilic than the WT R67 DHFR. A more detailed molecular modeling study of the hydrophobic active site cavities will be presented elsewhere (A.R.Schmitzer and J.N.Pelletier, submitted).

Conservation of the general hydropathy is likely to contribute to excluding the hydrophilic TMP, thus allowing survival under selective conditions. However, Figure 3A and B also illustrates that the chemical nature and location of polar and non-polar residues within the cavity can vary widely. At position 66, a variety of small hydrophobic and hydrophilic amino acids were encoded. Ser, Ile and Gly were selected at position 66 in the functional variants. Val66Ser, Val66Ile, Val66Ala and Val66Thr point mutants were previously identified as fully active mutants (Martinez et al., 1996Go; Hicks et al., 2004Go), indicating a good structural and functional tolerance at this position.

To maintain potential van der Waals interactions between Tyr69 and the ribose hydroxyl groups, we encoded mostly aromatic residues at position 69. The selected variants encode the native Tyr (1x) or His (2x) whereas Phe and Leu were not selected. While Tyr and His are both aromatic containing polar atoms, sampling was too light to conclude on any functional contribution of their polar atoms.

In each new variant, a charged residue has been selected (at each monomer): positively charged Lys67 (M36) and Arg68 (M10) or negatively charged Glu67 (M8_30). The structure of the WT R67 DHFR suggests that the side chain at position 68 does not protrude deeply into the cavity and its mutation should therefore have a smaller effect on the dielectric environment than charges at position 67 (Figure 1A). Again, the similar kcat values of the three variants and the WT R67 DHFR suggest that the rate-limiting step is unchanged and that there is no important change in catalytic mechanism. Thus, the mechanism of R67 DHFR is compatible with the absence of charge (WT) or the presence of a total of four charges (positive or negative) in the active site cavity; furthermore, molecular replacement suggests that these charges are located at different subsites of the active site pore (Figure 3B). Hence the important differences in the primary sequences of the WT R67 DHFR and the three functional variants, taken together with their similar kinetic properties and state of oligomerization, allow us to conclude that the reduction of dihydrofolate by NADPH at the active site pore can be conducted with native-like efficiency by variants with significantly differing chemical composition.

Function is supported by an important cavity effect

From the variety of side chains encoded in the functional variants (Figure 3), it appears that the cavity shape need not perfectly imitate that of the WT in order to be catalytically competent and TMP resistant. These results are consistent with the proposal that the active site cavity is a ‘binding hot spot’ (Strader et al., 2001Go); the selected functional variants represent alternative versions of this binding hot spot. Our data provide strong evidence supporting the hypothesis that the global active site environment of R67 DHFR is more important than the presence of any specific residue (Hicks et al., 2004Go). Binding and catalysis are not induced by specific interactions from particular active site residue side chains as each active site residue of the native enzyme can be substituted. The results provide new evidence that catalysis in the WT enzyme, as in the active variants, is promoted by a good relative positioning of protonated DHF and NADPH, a cavity effect that may affect dielectric constants in the pore and an orientation effect that restricts accessible conformations. These results are consistent with recent binding and kinetic analyses of active site point mutations of R67 DHFR suggesting that binding enthalpy is an important contributor to catalysis in this enzyme, that is proposed to involve the formation of productive interligand interactions which can proceed to the transition state (Hicks et al., 2004Go).

In conclusion, by creating combinatorial active site mutations in R67 DHFR, we selected novel active site configurations that fulfil the chemical and steric requirements of catalysis and TMP resistance. There is no simple pattern that connects the ensemble of functional sequences, suggesting that the fitness landscape of this enzyme is rugged, characterized by sharp peaks of high fitness that rapidly fall to low function upon mutation. A high number of compensatory mutations were shown to be required to regain function, highlighting the importance of a combinatorial approach to active site mutation. It is important to note that a significant fraction of the mutants (three out of 1536) created yielded novel functional active site patterns. The relative ease with which new functional patterns were identified underscores the simple requirements for catalysis in this enzyme. The importance of the cavity effect in reactivity is consistent with the binding plasticity that has previously been described for R67 DHFR, where broad substrate specificity (Holland et al., 1993Go) and moderate reducing cofactor tolerance have been observed (Smith and Burchall, 1983Go). The strategy of combinatorially exploring a restricted area of sequence space centred about an active site will be generally applicable, even in the case of rugged fitness landscapes where point mutations are generally deleterious. The strategy is readily carried out and can rapidly provide information about the minimal chemical requirements for enzyme function.


    Acknowledgments
 
This work was supported by NSERC Canada (Grant No. 227853-02). A.R.S. held fellowships from NCI Canada and the Association des Universités Francophones.


    References
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 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
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Received September 16, 2004; revised November 18, 2004; accepted November 19, 2004.

Edited by Jacques Fastrez