(Received for publication, October 3, 1996)
From the Department of Biochemistry, University of Tennessee, Knoxville, Tennessee 37996-0840
R67 dihydrofolate reductase (DHFR) is encoded by an R-plasmid, and expression of this enzyme in bacteria confers resistance to the antibacterial drug, trimethoprim. This DHFR variant is not homologous in either sequence or structure with chromosomal DHFRs. The crystal structure of tetrameric R67 DHFR indicates a single active site pore that traverses the length of the molecule (Narayana, N., Matthews, D. A., Howell, E. E., and Xuong, N.-H. (1995) Nat. Struct. Biol. 2, 1018-1025). A pH profile of enzyme activity in R67 DHFR displays an acidic pKa that is protein concentration-dependent. This pKa describes dissociation of active tetramer into two relatively inactive dimers upon protonation of His-62 and the symmetry-related His-162, His-262, and His-362 residues at the dimer-dimer interfaces. Construction of an H62C mutation results in stabilization of the active tetramer via disulfide bond formation at the dimer-dimer interfaces. The oxidized, tetrameric form of H62C R67 DHFR is quite active at pH 7, and a pH profile displays increasing activity at low pH. These results indicate protonated dihydrofolate (pKa = 2.59) is the productive substrate and that R67 DHFR does not possess a proton donor.
Dihydrofolate reductase (DHFR,1 EC 1.5.12.3) reduces dihydrofolate (DHF) to tetrahydrofolate using NADPH as a cofactor. DHFR is important in folate metabolism as the reaction product, tetrahydrofolate, is required for the synthesis of thymidylate, purine nucleosides, methionine, and other metabolic intermediates. The chromosomally encoded DHFR from Escherichia coli utilizes a general acid to facilitate catalysis (1) and has specific binding sites for both substrate (DHF) and cofactor (NADPH) (2). It has also been designated a highly evolved enzyme with a calculated efficiency of 0.15 (3). Hydride transfer rates are faster than the rate-determining step, which is release of product, tetrahydrofolate (4). In addition, ab initio quantum mechanical calculations suggest protein-mediated electronic polarization of bound DHF and NADPH may help lower energy barriers and aid catalytic function (5-7). Additionally, an overlap between the binding sites for NADP+ and 5-deazafolate in human DHFR has been proposed to facilitate catalysis by compressing the distance between C-6 of DHF and C-4 of NADPH to a separation that is optimal for hydride transfer (8). These studies all indicate that chromosomal DHFR has developed various mechanisms to maximize catalytic efficiency.
A different DHFR has emerged recently due to use of trimethoprim as a clinical drug to treat numerous bacterial infections. Trimethoprim is an active site-directed inhibitor of chromosomal DHFR. Resistance to trimethoprim has been correlated with the production of novel DHFRs encoded by R-plasmids. Type II R-plasmid-encoded R67 DHFR is unrelated genetically to chromosomal DHFRs (9, 10).
To compare this novel R-plasmid-encoded DHFR with its chromosomal counterpart, several crystal structures have been determined. A crystal structure of dimeric R67 DHFR was reported by Matthews et al. (11). More recently, the active homotetramer has been crystallized both as an apoenzyme and in a binary complex with folate (12). Difference Fourier maps describing bound folate indicate the active site is a pore traversing the length of the molecule and that residues from each monomer contribute to the single active site. The center of the pore possesses exact 222 symmetry, indicating R67 DHFR binds DHF and NADPH in an unusual manner. The difference Fourier map for bound folate shows the pteridine ring in the center of the pore. The electron density can be fit by two asymmetric binding sites each present at 1/4 occupancy (12). Due to steric constraints, a maximum of two ligands can bind concurrently (13). A productive model for catalysis proposes binding of DHF in half the pore and binding of cofactor in the other half. The pteridine ring of dihydrofolate and the nicotinamide ring of NADPH encounter each other at the center of the pore. This model of catalysis suggests R67 DHFR uses related sites (due to 222 symmetry) for binding of ligands and each half-pore accommodates either DHF or NADPH. However, DHF and NADPH are bound in slightly different orientations.
The recent evolutionary origin of R67 DHFR coupled with this unusual binding scheme hints that R67 DHFR may not be a very efficient catalyst. To probe the efficiency of the R67 DHFR reaction further and contrast it with the E. coli chromosomal DHFR, a series of experiments have been designed to investigate the pH dependence of the R67 DHFR reaction. As indicated above, a general acid has been proposed to facilitate catalysis in E. coli chromosomal DHFR (1). In a D27S2 mutant of E. coli chromosomal DHFR, the proton donor, Asp-27, was removed by site-directed mutagenesis. The D27S DHFR partially compensated by binding pre-protonated DHF (pKa = 2.59); this was reflected in an increased kcat at low pH where protonated dihydrofolate predominates. Alternately, a role for Asp-27 in tautomerization of bound substrate (14) or in alteration of the pKa of N-5 for bound substrate (15) has been proposed. When the crystal structure of R67 DHFR is examined for potential proton donors, the only residue identified in the active site pore is Tyr-69,3 as well as its symmetry-related residues Tyr-169, Tyr-269, and Tyr-369. Thus we pose the question, does R67 DHFR utilize a proton donor to facilitate catalysis as does chromosomal DHFR?
During a series of experiments to answer the above question, we found that the ionization describing reduction of protonated dihydrofolate is masked by dissociation of tetrameric R67 DHFR to dimers. Thus we additionally ask, what are the relative activities of dimeric and tetrameric R67 DHFR? And can the tetramer be stabilized by site-directed mutagenesis to introduce disulfide bonds across the dimer-dimer interfaces? This paper addresses these questions.
The H62C
mutation was constructed by standard site-directed mutagenesis
techniques using a synthetic R67 DHFR gene (16) and the following
oligonucleotide: 5-TGAGCCCGGGCAAGCCTCAGA-3
.
The entire mutant H62C R67 DHFR gene was sequenced to confirm that no additional mutations were present. This mutant gene conferred trimethoprim resistance upon E. coli, indicating that an active DHFR was produced.
H62C R67 DHFR was purified according to the protocol used to purify wild type (wt) R67 DHFR except that E. coli cells were lysed by sonication and reducing agents were omitted from buffers (16). 5 mM EDTA was included in the buffers to minimize air-oxidation of sulfhydryl groups. Purification steps included G75 molecular sieving, DEAE-Fractogel and DEAE-Sephacel chromatography, and a Mono-Q column on a Pharmacia FPLC system.
Steady State KineticsSteady state kinetic data were
obtained with a Perkin-Elmer 3a spectrophotometer interfaced with an
IBM PS2 according to Howell et al. (17). The computer
program UVSL3 (Softways, Moreno Valley, CA) was used in data collection
and analysis. Assays were performed at 30 °C in a polybuffer
containing 50 mM acetic acid + 50 mM MES + 100 mM Tris + 10 mM
-mercaptoethanol (MTA
buffer). This buffering system maintains a constant ionic strength
between pH 4.5 and 9.5 (18). For the wt R67 DHFR pH profile, the enzyme concentration was held constant at all pH values unless noted. Enzyme
was diluted and incubated in the cuvette for 5 min prior to initiation
of the assay by addition of 110 µM dihydrofolate and 64 µM NADPH. Non-enzymic hydrolysis rates were measured and subtracted from enzymic rates.
Formation of NADPD, used for measuring isotope effects, involved the reaction of alcohol dehydrogenase from Leuconostoc mesenteroides (Boehringer Mannheim) on 1,1-dideuterioethanol (MSD Isotopes) and NADP+ (Sigma). This reaction was coupled with NADP+ aldehyde dehydrogenase (Sigma) to allow the reaction to approach completion (17, 19). The dehydrogenases were removed from NADPD by ultrafiltration through a YM-10 membrane. Chromatography on a DEAE Fractogel column (0-0.4 M KCl gradient) removed any unreacted NADP+. NADPD was subsequently desalted by chromatography on a Bio-Gel P-2 column and lyophilized.
A model to describe the behavior of R67 DHFR as a function of pH is
shown in Scheme I, where T is tetrameric R67 DHFR;
DHn is protonated dimer; Ka and
Ka are ionization constants;
Kd, Ks, and
Ks are binding constants; and
Koverall describes the linked dissociation and ionization of tetramer to 2 DHn (20).
Scheme I.
From Scheme I, an equation can be derived to describe the concentration of DHn.
![]() |
(Eq. 1) |
![]() |
The concentration of T·NADPH·HDHF can be calculated as shown below.
![]() |
(Eq. 2) |
![]() |
![]() |
(Eq. 3) |
For H62C R67 DHFR, Km values for NADPH and DHF were obtained by varying both DHF (4-25 µM) and cofactor (8-80 µM) concentrations at subsaturating levels; primary and secondary plots were calculated according to Cleland (23). The pH profiles for kcat and kcat/Km were fit to Equation 4,
![]() |
(Eq. 4) |
Circular dichroism spectra of wt and H62C R67 DHFRs in 10 mM KH2PO4 at pH 7.0 and 5.0 were obtained at 22 °C using a Jasco J720 spectropolarimeter. Cell pathlength was 2 mm. Ten spectra were acquired per sample using 1-nm steps and 2-s integrations, and an averaged spectrum calculated. An essentially flat buffer baseline scan was then subtracted from the average protein scan. The CD data are described as the mean residue ellipticity by taking 111 g/mol as the mean residue molecular weight.
Gel filtration using a Superose 12 HR10-30 column on a Pharmacia FPLC
was performed according to Nichols et al. (2). Reducing and
non-reducing Ellman's titrations were done according to Creighton (25)
and Riddles et al. (26). Concentrations were determined using the following extinction coefficients: 28,000 M1 cm
1 at 282 nm for DHF (27);
6220 M
1 cm
1 at 340 nm for NADPH
(28). The molar extinction coefficient used to assess DHFR reduction of
DHF was 12,300 M
1 cm
1 (29).
Enzyme concentrations were determined by biuret assays and all proteins
were homogeneous according to SDS-polyacrylamide gel
electrophoresis.
The disulfide bond stability was assayed according to Sauer et al. (30). Oxidized H62C R67 DHFR was mixed with 8.8 mM oxidized dithiothreitol (Sigma) and varying concentrations of reduced dithiothreitol (0.05-0.1 mM; Life Technologies, Inc.) and incubated at room temperature for 24 and 48 h. The reactions were quenched by addition of 0.3 volume of 1 M iodoacetic acid. All experiments were performed under a nitrogen atmosphere in TE buffer (10 mM Tris, 1 mM EDTA, pH 8). The dimer/tetramer concentrations in the quenched samples were analyzed using a Superose 12 HR10-30 column on a Pharmacia FPLC. Peak areas were measured by weighing on a balance.
The only potential
proton donating group in the active site pore is Tyr-69 as well as its
symmetry-related residues, Tyr-169, Tyr-269, and Tyr-369. To determine
if tetrameric R67 DHFR uses Tyr-69 as a proton donor, we examined a pH
profile of R67 DHFR activity. If Tyr-69 were a general acid
participating in catalysis, we would expect to observe a half-bell
profile displaying a pKa of pH 9.5-10.5,
corresponding to ionization of tyrosine (assuming no effects due to
environment). On the other hand, if R67 DHFR does not use Tyr-69 as a
general acid, we would expect to see increasing activity at low pH as
the pKa for protonation of dihydrofolate at N-5 is
approached (pKa = 2.59; Ref. 21), similar to the pH
profile exhibited by D27S E. coli chromosomal DHFR (1).
Instead of either of these patterns, a pH profile of
kcat displays a bell shape as shown in Fig.
1. From the pH profile at high pH, we can conclude that
if Tyr-69 is acting as a proton donor, its contribution to catalysis
must be minimal since a titration is not observed up to pH 9.88. Activity measurements above this pH were not performed due to the high
enzyme concentrations needed. Because of the size of the active site
pore (~18 Å wide by ~24 Å long) and consequent solvent
accessibility, it seems unlikely that the pKa of
Tyr-69 would be perturbed.
The apparent pKa value of 6.15 (Ptot = 166 nM dimer) observed in Fig. 1 suggests histidine may be ionizing and affecting enzyme activity. A single histidine (His-62) occurs per monomer at the dimer-dimer interfaces. From the 222 symmetry of the tetrameric crystal structure, 2 histidines appear per dimer-dimer interface (i.e.. His-62 and His-362 at one interface, and His-162 and His-262 at the second symmetry-related interface). Protonation of His-62 (pKa 6.84) has previously been found to destabilize tetrameric R67 DHFR and favor dimer formation (20). As the active site pore is lost when tetramer dissociates to dimer, decreased activity results.
Since dissociation of tetrameric R67 DHFR to 2 dimers is a bimolecular reaction, the ionization describing this process should be protein concentration-dependent. To test this, the activity associated with three protein concentrations was monitored as a function of pH. As seen in Fig. 1, the basic portion of the bell-shaped curves overlays fairly well. However the acidic titration clearly shows a protein concentration dependence, consistent with dissociation of active tetramer to less active dimers.
Engineering a Stable TetramerSince protonation of His62 is
linked to dissociation of active tetramer into inactive or partially
active dimers, any potential activity increase at low pH due to
protonation of dihydrofolate in solution (pKa = 2.59) will be masked. To investigate whether R67 DHFR utilizes
protonated or nonprotonated DHF as the productive substrate, we
constructed a stable tetramer to eliminate the T + 2nH+ 2DHn ionization from the pH profile.
The distance between the -carbons of histidines 62 and 362 (or 162 and 262) at the dimer-dimer interface is 9.6 Å in R67 DHFR. The
-carbon distance is 7.8 Å. Sowdhamini et al. (31) have
proposed that CA-CA distances of
6.5 Å and CB-CB distances of
4.5
Å indicate a good potential for disulfide bond formation. However,
Matsumura and Matthews (32) have constructed disulfide bridges in T4
lysozyme with CA-CA distances of 8.1 Å. While the distances in R67
DHFR are slightly longer, a H62C substitution was modeled using INSIGHT
(Biosym Technologies) and a disulfide bond generated. The model
appeared feasible, thus a H62C mutation was constructed in the R67 DHFR
gene as described under "Materials and Methods."
A majority of purified
H62C R67 DHFR is oxidized and tetrameric as determined by molecular
sieving studies and Ellman's titrations. A reduced, dimeric component
can also be observed, which upon extended incubation produces stable,
oxidized tetramer. This result indicates the H62C mutation has greatly
increased the Kd describing the tetramer 2 dimers equilibrium, most likely due to the unfavorable distance between
cysteines for disulfide bond formation. Previously, the
Kd for wt R67 DHFR was estimated to be
50
nM by sedimentation equilibrium and 9.72 nM by
fitting of protein concentration-dependent pH titration of
fluorescence (20). The relatively small dimer-dimer interface areas
indicate stacking and hydrogen bond formation between His-62 and
His-362 and between His-162 and His-262 are a major determinant of
association state.
Disulfide bond formation between H62C R67 DHFR dimers does occur
spontaneously in vitro, albeit slowly. Numerous procedures were initially investigated to facilitate disulfide bond formation, for
example disulfide exchange (33, 34) or mercuric ion cross-bridging (35). However, the easiest and most efficient method found was to
incubate dimeric H62C R67 DHFR at high protein concentrations (15
mg/ml), pH 8.8, 4 °C for >1 week. Since the Kd
for tetramer formation in H62C R67 DHFR is elevated, high protein concentrations were necessary to facilitate tetramer formation and
subsequent disulfide cross-linking. Generation of active, cross-linked
tetramer was readily monitored by either increased enzyme activity
and/or by molecular sieving chromatography and confirmed by Ellman's
titrations (see below). From the chromatography results, two peaks are
observed during elution from a Superose 12 HR10-30 column (Pharmacia
FPLC), indicating tetrameric and dimeric species are not in rapid
equilibrium in H62C R67 DHFR. With increasing time (days
weeks),
the first peak increases in area. The first peak was identified as
active tetramer (S-S cross-linked) and the second peak as reduced dimer
by Ellman's titrations, Kav values (and thus
predicted molecular weights) and by the first peak possessing virtually
all the enzyme activity. To separate active tetramer from dimer,
ion-exchange chromatography on a Mono-Q column with a 0.05-0.07
M KCl gradient in 10 mM Tris, pH 8 was utilized
(Pharmacia FPLC). Purified dimer was maintained in buffer + 1 mM dithiothreitol. Alternatively, molecular sieving chromatography on G-75 Sephadex at pH 5 allows separation of reduced, dimer from oxidized tetramer.
Ellman's titrations were performed to verify the formation of
disulfide bonds in tetrameric H62C R67 DHFR. A single sulfhydryl per
monomer, Cys-47, occurs on the surface of R67 DHFR. Non-reducing Ellman's titrations yield 0.78 and 0.99 free sulfhydryls/monomer for
wt and oxidized, tetrameric H62C R67 DHFRs, respectively (average of
2 determinations). In contrast, reducing Ellman's titrations yield
1.1 and 2.2 sulfhydryls/monomer (average of 2 determinations) for wt
and oxidized, tetrameric H62C R67 DHFR, respectively. These values
clearly confirm a stable tetrameric species has been constructed by
disulfide cross-linking.
As shown in Fig. 2, a comparison of elution position for
wt R67 DHFR from a Superose 12 HR 10-30 column at pH 8 and 5 shows a
significant change, consistent with dissociation of tetramer to 2 dimers (20). In contrast, molecular sieving studies of active,
tetrameric H62C R67 DHFR at pHs 8 and 5 show changes in elution
position that are the same within experimental error. From a plot of
Kav (= (elution volume void
volume)/(total bed volume
void volume)) versus
molecular weight standards, the molecular mass of H62C R67 DHFR at pH 8 and 5 was calculated as 42,000. For comparison, the measured molecular
mass of wt R67 DHFR using molecular sieving techniques is also 42,000 at pH 8.0 (20). These values are higher than a value of 33,720 calculated from the amino acid sequence. Aberrant molecular weight
estimates are not unusual, however, as both the shape and size of the
molecule affect elution position (36).
Circular dichroism studies were performed to assess the effects of the
H62C mutation on the secondary structure of R67 DHFR (Fig.
3). The spectra for oxidized, tetrameric H62C R67 DHFR
at pH 5 and 7 correlate well with the spectrum of wt, tetrameric R67
DHFR at pH 7.0. However, the spectra for reduced, dimeric H62C R67 DHFR
at both pH 5 and 7 show significant differences from the spectrum for
wt, dimeric R67 DHFR at pH 5.0. Since dimeric H62C R67 DHFR possesses
minimal activity (see below) and is stably folded (37), the change in
CD signal must reflect a change in protein conformation/environment.
Since His-62 is near Trp-38 at the dimer-dimer interface and His-62 is
protonated at pH 5 in wt R67 DHFR, perhaps the H62C substitution
affects the contribution of Trp-38 to the CD signal. Woody (38)
indicates aromatic side chains can make detectable contributions to the
far-UV CD. In contrast to the spectra for wt R67 DHFR, the spectra for
oxidized and reduced H62C R67 DHFR do not show pH-dependent
changes since they are locked in their respective tetrameric and
dimeric forms.
Activity of Tetrameric H62C R67 DHFR
Oxidized, tetrameric
H62C R67 DHFR is quite active with kcat = 74 ± 0.9 min1,
Km(DHF) = 29 ± 0.6 µM and Km(NADPH) = 34 ± 3.0 µM at pH 7.0. For comparison, wt R67 DHFR values
are kcat = 78 min
1,
Km(DHF) = 5.8 µM and
Km(NADPH) = 3.0 µM at pH
7.0. Km values increase 5-11-fold, while
kcat remains unaffected. One additional
difference associated with the H62C mutation is the observation that
DHF inhibition now occurs at high concentrations (
40 µM
at pH 7.0). Substrate inhibition is not observed in wt R67 DHFR;
however, it is consistent with nonproductive binding of 2 DHF
molecules, which blocks formation of the productive R67
DHFR·DHF·NADPH ternary complex
(13).4
The availability of a pH stable tetramer allows us the unique
opportunity of addressing the proton donation mechanism in R67 DHFR. A
pH profile for tetrameric H62C R67 DHFR shows increasing activity at
low pH (Fig. 4). This behavior is reminiscent of the pH
profile for the mutant D27S E. coli chromosomal DHFR (1) and
supports a model for catalysis where R67 DHFR uses protonated dihydrofolate as a substrate. Catalytic activity increases at low pH as
the pKa for N-5 in dihydrofolate is approached. The
values obtained for fitting the kcat profile
data to Equation 4 with a fixed pKa of 2.59 for
protonated DHF (21) are a pH-independent kcat
value of 197,000 ± 43,000 min1 and a slope of
0.64 ± 0.03. Setting the pKa is necessary for
fitting, since a plateau has not yet been reached and values for
kcat at lower pH levels are difficult to obtain
due to hydrolysis of NADPH.
At pH 4.95, kcat increases to 2500 ± 110 min1, Km(DHF) increases to
180 ± 11 µM, and
Km(NADPH) increases to 260 ± 25 µM in oxidized, tetrameric H62C R67 DHFR. The best fit
values obtained for the
kcat/Km(DHF)
profile data are a pH-independent
kcat/Km(DHF)
value of 260 ± 53 min
1
µM
1 and a slope of 0.29 ± 0.014.
To confirm that hydride transfer is rate-limiting throughout this pH range for H62C R67 DHFR, NADPD isotope effects were measured. At pH 5.0, DV (= kcat using NADPH/kcat using NADPD) = 3.6 ± 0.45 and at pH 7.0, DV = 3.3 ± 0.33. These results indicate hydride transfer is fully rate-determining from pH 5-7. DV/Km(DHF) at pHs 5 and 7 are 3.8 ± 0.46 and 2.7 ± 0.34, respectively.
Activity of Dimeric H62C R67 DHFRThe activity of dimeric
H62C R67 DHFR was assessed under reducing conditions. At pH 7.0, kcat is 2.0 ± 0.12 min1,
Km(DHF) = 36 ± 4 µM
and Km(NADPH) = 74 ± 6 µM. These values compare well with previously determined
values for diethyl pyrocarbonate (DEPC)-modified wt R67 DHFR (20). DEPC
modification of histidines 62, 162, 262, and 362 resulted in a
stabilized dimer and allowed setting of an upper activity limit for
dimeric R67 DHFR with kcat,
Km(DHF) and
Km(NADPH) values of 3.1 min
1, 40 µM, and 72 µM,
respectively (20). The upper limit qualification was due to partial
modification of only 2.7 histidines/tetramer. Loss of substantial
enzyme activity in both dimeric H62C R67 DHFR and DEPC-modified, wt,
dimeric, R67 DHFR can be understood in the context of loss of the
active site pore upon dissociation of tetrameric R67 DHFR to 2 dimers.
From these experiments however, we cannot exclude the possibility of
transient tetramer formation induced by ligand binding.
To assess the stability of the H62C intersubunit disulfide bonds, the overall Kd for their reduction/oxidation was monitored. The reduction/oxidation reaction is as follows,
![]() |
![]() |
![]() |
(Eq. 5) |
Two lines of evidence indicate R67 DHFR does not possess a general acid in its active site pore. First, the crystal structure does not show a general acid in the active site pore, except for perhaps Tyr-69. Since a pH profile of kcat for R67 DHFR does not display a pKa up to pH 9.88, Tyr-69 must not play a major role in catalysis. Additionally, the altered pH profile for oxidized, tetrameric H62C R67 DHFR displays increasing activity at low pH, consistent with increasing concentrations of protonated DHF as the pKa for protonation of N-5 in DHF is approached. This model for catalysis contrasts with mechanisms proposed for chromosomally encoded DHFRs, which possess a proton donor to facilitate catalysis.
A Kinetic MechanismBased on the pH profiles for both wt and
H62C R67 DHFR, a simple model describing R67 DHFR catalysis is shown in
Scheme I, where T is tetramer and DHn is protonated dimer. In
this model, R67 DHFR uses protonated dihydrofolate (HDHF) as substrate; binding of unprotonated dihydrofolate is unproductive unless
protonation occurs after binding (Ka). While
random addition of substrate and cofactor might be expected from the
symmetry of the crystal structure (12), our recent binding studies (13) indicate a total of two ligands can bind per pore and a preferred binding pathway occurs where NADPH is bound initially, followed by DHF.
Binding of 2 NADPH molecules can occur but is disfavored due to
negative binding cooperativity, while binding of two DHF molecules is
disfavored due to positive binding cooperativity (microscopic
Kd values of 250 and 4.4 µM). These
observed cooperativities arise from ligand-ligand interactions which
minimize nonproductive binding of two identical ligands. A preferred
binding pathway involving two heteroligands predominates and is
enhanced by positive cooperativity between bound NADPH and DHF. While
these binding studies indicate a more complicated mechanism involving two nonproductive homoligand complexes, a simple, but predominate mechanism was used to fit the pH profile data in Fig. 1, as the fit can
only yield two variables with reasonable error.
From Scheme I, an equation can be derived for fitting the
pH-dependent kinetic data in Fig. 1 (,
,
). It is
not clear whether the pKa of bound protonated
substrate (HDHF) is equivalent to the pKa of free
substrate or whether the Km values for HDHF and DHF
differ. Since fitting to Equation 3 cannot yield estimates for
Ka,
Ks,
Ka, and Ks as well as for
kcat(max) and Koverall,
for fitting purposes we assumed Ka =
Ka and Ks =
Ks. If either Ks or
Ka are perturbed, then the fitted values would
change accordingly. For the fit below, Ks was held
constant at 5.8 µM, the Kd for NADPH was set at 2.5 µM (13) and the pKa
utilized for DHF protonation was 2.59 (21).
All three data sets in Fig. 1 were fit globally to Equation 3 using
NONLIN and best fit values are: n = 1, Koverall = 1.24 × 1010
M (1.14 × 10
10 to 1.34 × 10
10, 67% confidence interval), and
kcat(max) = 2.91 × 106
min
1 (2.77 × 106 to 3.05 × 106 min
1, 67% confidence interval). A value
of n = 1 indicates the addition of only one
proton/dimer; this could occur if a hydrogen bond already exists
between a protonated histidine and a second symmetry-related, nonprotonated histidine at each dimer-dimer interface.
The pH-independent kcat values predicted for wt
and H62C R67 DHFRs show a 15-fold difference (2,910,000 min1
versus 197,300 min
1, respectively). The
origin of this difference likely lies in the fits for these two
different data sets, as the slope describing protonation of free and
bound DHF was allowed to vary for the H62C R67 DHFR data but was
constrained to 1 for the wt R67 DHFR data. If the slope
(i.e. the number of H+ adding to DHF and
E·NADPH·DHF in Scheme I) is treated as a variable and the wt R67
DHFR data refit globally, the fit values obtained are
Koverall = 2.66 × 10
10
M (2.53 × 10
10 to 2.77 × 10
10, 67% confidence interval),
kcat(max) = 93,000 min
1
(77,200-110,000, 67% confidence interval) and slope = 0.794 (0.784-0.805, 67% confidence interval; fit not shown). The use of a
non-integral slope for both wt and H62C R67 DHFR data results in only a
2-fold difference in kcat(max) values,
consistent with a similar mechanism associated with each DHFR variant.
A non-integral slope likely reflects minor effects of additional
ionizations on the catalytic activity and/or protein conformation.
As indicated in the Introduction, E. coli chromosomal DHFR appears to be a well evolved enzyme. In striking contrast to catalytic function in chromosomal DHFRs, our kinetic results indicate that R67 DHFR is not very efficient as it does not possess a proton donor. Also, in a model of a productive ternary complex based on the crystal structure and difference Fourier maps as well as solution binding studies, either DHF or NADPH can occupy the same half pore, although with different orientations (12, 13). These observations lead us to wonder what factors R67 DHFR utilizes to promote hydride transfer. It is interesting to speculate that the elements most likely involved in facilitating catalysis are: binding interactions for the substrate and cofactor, a cavity effect that may affect dielectric constants in the pore; and third, an orientation effect that restricts accessible conformations (39, 40).
A mutant D27S E. coli chromosomal DHFR was previously
constructed using site-directed mutagenesis techniques to determine the
effect on catalysis of removing the proton donor (1). An increase in
activity at low pH was also observed. When the catalytic efficiencies
(kcat/Km) for R67 DHFR and
D27S E. coli chromosomal DHFR are compared at pH 7.0, R67
DHFR is 30 times more efficient, as kcat for the
D27S E. coli chromosomal DHFR at pH 7.0 is 24.6 min1 and Km(DHF) = 56 µM. The relative agreement of rates (25 versus
74 min
1) in D27S chromosomal E. coli DHFR and
R67 DHFR may indicate a limit to the activity feasible without a proton
donor present.
We thank Enrico DiGiammarino and Chuck Linn for their assistance in several experiments. We also thank Cynthia Peterson and Dan Roberts for their helpful discussions and reading of the manuscript.