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Originally published In Press as doi:10.1074/jbc.M709443200 on December 17, 2007

J. Biol. Chem., Vol. 283, Issue 8, 4690-4698, February 22, 2008
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A Balancing Act between Net Uptake of Water during Dihydrofolate Binding and Net Release of Water upon NADPH Binding in R67 Dihydrofolate Reductase*Formula

Shaileja Chopra, Russell M. Dooling, Caroline Glyn Horner, and Elizabeth E. Howell1

From the Department of Biochemistry, Cellular, and Molecular Biology, University of Tennessee, Knoxville, Tennessee 37996-0840

Received for publication, November 16, 2007 , and in revised form, December 13, 2007.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
R67 dihydrofolate reductase (DHFR) catalyzes the reduction of dihydrofolate (DHF) to tetrahydrofolate using NADPH as a cofactor. This enzyme is a homotetramer possessing 222 symmetry, and a single active site pore traverses the length of the protein. A promiscuous binding surface can accommodate either DHF or NADPH, thus two nonproductive complexes can form (2NADPH or 2DHF) as well as a productive complex (NADPH·DHF). The role of water in binding was monitored using a number of different osmolytes. From isothermal titration calorimetry (ITC) studies, binding of NADPH is accompanied by the net release of 38 water molecules. In contrast, from both steady state kinetics and ITC studies, binding of DHF is accompanied by the net uptake of water. Although different osmolytes have similar effects on NADPH binding, variable results are observed when DHF binding is probed. Sensitivity to water activity can also be probed by an in vivo selection using the antibacterial drug, trimethoprim, where the water content of the media is decreased by increasing concentrations of sorbitol. The ability of wild type and mutant clones of R67 DHFR to allow host Escherichia coli to grow in the presence of trimethoprim plus added sorbitol parallels the catalytic efficiency of the DHFR clones, indicating water content strongly correlates with the in vivo function of R67 DHFR.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
R67 dihydrofolate reductase, a type II DHFR,2 catalyzes the NADPH-dependent reduction of dihydrofolate (DHF) to tetrahydrofolate. The gene for this enzyme is carried by an R-plasmid, and its presence confers resistance to the antibiotic drug, trimethoprim (TMP). Trimethoprim is a competitive inhibitor of chromosomal DHFR with a picomolar Ki (1). Although R67 DHFR is not a good catalyst, it is not inhibited by TMP very well, and thus it allows cell growth in the presence of this antibacterial drug (2, 3). R67 DHFR shares no homology in sequence or structure with chromosomal DHFR and has been proposed to be a good model of a primitive enzyme (4).

R67 DHFR is a homotetramer with a single active site pore; the overall structure possesses 222 symmetry as seen in Fig. 1 (5). The symmetry of the active site results in overlapping binding sites for substrate, DHF, and cofactor, NADPH. This can be observed as R67 DHFR binds a total of two ligands as follows: either two NADPH molecules or two folate/DHF molecules or one NADPH plus one folate/DHF molecule (6). The first two complexes are dead-end (binary) complexes, whereas the third is the productive ternary complex. Because of the 222 symmetry, binding to either ligand is unlikely to be optimal.

The active site pore of R67 DHFR is unusual in its hourglass shape as well as its large size (2938 Å3 for apoR67 DHFR with hydrogens added, calculated by CASTp (4, 7)). Because of this large volume, DHF and NADPH cannot occupy all the space in the pore and must use water to mediate some contacts with the protein.

How do substrate and cofactor bind to R67 DHFR? From NMR, crystallography, and docking studies, the pteridine ring of DHF/folate binds at the center of the pore, but the para-aminobenzoylglutamate tail of dihydrofolate/folate is disordered (5, 8, 9). Interactions have been predicted between symmetry-related Lys-32 residues (at either edge of the pore) and the glutamate tail of substrate. Also, increasing concentrations of sucrose or trehalose (10) increase Km (DHF) as well as Kd (DHF). As polyols can have a variety of effects, including perturbation of solvent structure, a possibility is that these compounds affect the water structure and, in turn, DHF binding.

If water is involved in an interaction, perturbation of water content should affect binding. Thus we probed the role of water in DHF/folate binding to R67 DHFR, using osmolytes with different properties to determine whether the primary effect was because of water or some other variable.

To provide a complete picture, the effects of osmolytes on the interaction of NADPH with R67 DHFR were also monitored. Previous crystallography and NMR studies (8, 9, 11, 12) show NADPH binds in an extended conformation, with numerous specific interactions, including H-bonds between the nicotinamide carboxamide of NADP+ with backbone NH and O atoms from Ile-68. Ionic interactions also form between the nicotinamide phosphate and the adenosyl-2'-phosphate with symmetry-related Lys-32 residues (13–16).

Because of the 222 symmetry of the active site pore, DHF and NADPH bind to the same promiscuous surface. Using various osmolytes, this study finds water release accompanies NADPH binding, whereas water uptake accompanies DHF binding.


Figure 1
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  		FIGURE 1.
The secondary structure of tetrameric R67 DHFR (Protein Data Bank code 1VIE). The top panel shows the four monomers in green, pink, ochre, and violet. The active site pore faces the viewer. The dimer-dimer interfaces occur on the top (green and violet) and bottom (pink and ochre) of the structure. The bottom panel is related to the top panel by a 90° rotation along the y axis, followed by slicing through the structure to show the pore. A Connolly surface displays the positions of key residues: Lys-32 (orange), Ile-68 (yellow), Gln-67 (blue), and Tyr-69 (sea green). Arrows on opposite sides of the pore indicate entry of the ligands. In this ternary complex structure (8), NADP+ enters from the left and DHF from the right (8); however, binding at symmetry-related sites can also occur. The color code for the atoms is carbon, green; nitrogen, blue; oxygen, red; phosphorus, magenta; and hydrogen, white. The pteridine ring of folate is fixed near the center of the pore; however, the para-aminobenzoylglutamate tail is disordered. One potential tail conformer was built in the structure and is shown in yellow. Alternate tail positions are suggested by the circular arrow.

 

   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Protein Purification—High yields of R67 DHFR were obtained as described previously (17). Briefly, ammonium sulfate precipitation and ion-exchange column chromatography were used to purify the protein to homogeneity. Purified samples were dialyzed against distilled, deionized H2O and then lyophilized. Protein concentrations were determined with a biuret assay (18).

Steady State Kinetics—Steady state kinetic data were obtained using a PerkinElmer Life Sciences {lambda}35 spectrophotometer interfaced with an IBM personal computer as described previously (19). MTA (MTA buffer: 100 mM Tris, 50 mM MES, 50 mM acetic acid polybuffer plus 1 mM β-mercaptoethanol) polybuffer was used. This buffering system maintains a constant ionic strength from pH 4.5 to 9.5 (20). The steady state kinetic rates were measured at 30 °C by the addition of substrate and cofactor, followed by the addition of enzyme to initiate the reaction. To obtain kcat and Km values, the concentration of NADPH was held constant at a subsaturating level, whereas the concentration of DHF was varied. This process was repeated using four additional subsaturating concentrations of NADPH. The data were fit globally to the nonlinear bisubstrate Michaelis-Menten equation utilizing SAS (statistical analysis software (21, 22)). The NLINEK macro for use in SAS is available on line. For those conditions where Km (NADPH) did not change, saturating concentrations of NADPH were used, and DHF concentrations varied to obtain kcat and Km (DHF) values.

Water Activity Measurements—A Westcor 5500 vapor pressure osmometer was used to obtain the osmolality of the solutions. This value was converted into water activity using Equation 1,

Formula 1(Eq. 1)
where aH2O is the water activity (23).

The binding of DHF to R67 DHFR·NADPH in a buffer containing neutral osmolytes (S) can be described by Equation 2,

Formula 2(Eq. 2)
where {nu}H2O and {nu}S are the stoichiometric coefficients of water and osmolyte, respectively. From Wyman (24), it can be calculated that ln Ka (or ln kcat/Km (DHF)) is related to ln as according to the following: {nu}H2O and {nu}S, the stoichiometric coefficients of water and osmolyte, as shown in Equation 3,

Formula 3(Eq. 3)
Various groups have found that low molecular weight osmolytes are preferentially excluded from the surface of proteins (25–27). This observation suggests that the value of {nu}H2O would contribute more significantly to Equation 3 than {nu}S. This scenario can be tested by use of osmolytes from different classes, which differentially associate with proteins (and presumably ligands as well (25, 26)).

Isothermal Titration Calorimetry—Affinities, stoichiometries, as well as {Delta}H values associated with binding were determined using isothermal titration calorimetry (ITC) as described previously (6). Measurements were performed on a VP-ITC microcalorimeter from MicroCal interfaced to a Gateway personal computer for data acquisition and analysis. Origin version 5 scientific software was used to analyze the data. This instrument has been described previously (28). R67 DHFR concentrations typically ranged from 60 to 150 µM in MTA buffer (pH 7). For titrations with osmolyte present, MTA buffer plus osmolyte was used in the reference cell. The "c value" (=[Ptotal]/Kd) for binding was from 3 to 127, within the suggested range of 1–1000 (28).

The heat capacity ({Delta}Cp) or the change in {Delta}H as a function of temperature is described by Equation 4,

Formula 4(Eq. 4)
Heat capacity changes were obtained for NADPH binding to R67 DHFR and also DHF binding to R67 DHFR·NADP+ using the temperature range 278–303 K.

Escherichia coli Growth Conditions—The ability of the E. coli strain DH5{alpha} to grow on M9 minimal media (29) containing 0.02% casamino acids with 20 µg of TMP/ml was assessed. Another layer of screening added increasing concentrations of sorbitol to the media. The water activity of the plates was measured using an Aqualab meter (Decagon Devices) and converted into osmolality using Equation 1.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Role of Water in kcat/Km(DHF)—To garner more information concerning how DHF is bound to R67 DHFR, the perturbation associated with addition of varying osmolytes was studied. This approach monitors any change in the number of water and/or solute molecules associated with two different states, free DHF and the enzyme·NADPH binary complex versus the enzyme·NADPH·DHF transition state structure (23, 30–34). However, alterations in solvent viscosity, solution dielectric constant, or volume exclusion could also affect binding upon osmolyte addition. To determine whether the previously observed effect of sucrose on DHF binding (10) arises solely from perturbations on water activity, osmolytes with different characteristics were used. For example, addition of glycine betaine or sucrose can both affect water activity, yet these molecules either increase or decrease the dielectric constant of the solution, respectively (30, 35, 36). If both compounds show similar results, then effects on the dielectric constant are not involved. To test the role of volume exclusion on binding, PEGs of increasing size are typically used. The osmotically active volume is the volume accessible to water but inaccessible to osmolyte. Here, the osmotically active volume depends on the size of the osmolyte, with larger osmolytes detecting changes in larger volumes. Osmotic stress occurs as the solution must compensate for this exclusion (30, 31). A last possibility is the direct interaction of the osmolyte with either the ligand or protein (37, 38). If similar results are observed with chemically different osmolytes, this option appears unlikely (25, 26).

With this background in mind, the steady state kinetic behavior of R67 DHFR was monitored in the presence of the neutral osmolytes glycerol, ethylene glycol, trimethylamine N-oxide (TMAO), dimethyl sulfoxide (Me2SO), glycine betaine, and sucrose. Little to no effect (<1.5-fold) is observed on the catalytic rate constant, kcat, whereas the Km for DHF increases in the presence of all these osmolytes. A possible decrease in Km for NADPH is also observed. If a linear relationship is observed in plots of either osmolality or ln water activity versus ln kcat/Km (DHF), then effects on water are involved (30, 38, 40, 41). Fig. 2 shows the linear relationships associated with plots of osmolality versus ln kcat/Km (DHF). As a precaution, we also plotted effects on solution viscosity or solution dielectric and overlapping data were not observed. These figures are available as supplemental Figs. 1 and 2. Another control experiment monitored the effect of osmolytes on the pH-dependent dissociation of tetramer to two protonated dimers. Addition of 10% ethylene glycol, 12.5% Me2SO, or 20% PEG400 did not greatly alter the pH titration data (experimental design in supplemental material as well as supplemental Fig. 3), indicating that osmolyte addition minimally perturbs the oligomeric state of R67 DHFR.


Figure 2
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  		FIGURE 2.
A plot of osmolality versus ln kcat/Km(DHF). Steady state kinetic data were obtained at saturating [NADPH] in the presence of various osmolytes. Michaelis-Menten plots were then used to calculate kcat and Km (DHF) values. The units of kcat/Km (DHF) are s–1 M–1. Data for buffer (gray circle), glycerol (solid circle), ethylene glycol (open star), TMAO (inverted open triangle), sucrose (open square), Me2SO (open circle), glycine betaine (open triangle), and PEG400 (checkerboard) are shown. Lines through the various data sets are also included. The slopes of these plots were converted to {Delta}nW using the relationship: d ln kcat/Km (DHF)/d[osmolal] =–{Delta}nW/55.6. These values are reported in Table 1. These values can also be obtained from a plot of ln kcat/Km (DHF) versus ln water activity.

 
From Equation 3, the slope of a plot of ln water activity versus ln kcat/Km (DHF) describes the change in the number of water molecules, {Delta}nW, involved in DHF binding to the enzyme·NADPH complex and going to the transition state. The slopes associated with the individual osmolytes for this type of plot are given in Table 1. The slopes associated with TMAO, glycerol and ethylene glycol are low, around 16–25, and the data overlap reasonably well. A fit to these three combined data sets yields a slope of 17 ± 2. However, the slopes for the glycine betaine, Me2SO, and sucrose data are higher, ranging from 40 to 60. Variable slopes are common in osmolality studies, but their origin is not clear (37, 38, 41–54). This issue will be broached under the "Discussion."


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  		TABLE 1
Osmolyte information and slopes ({Delta}nW) associated with plots of In aH20 versus In kcat/Km (DHF) as determined by steady state kinetics

 
Ka Determination—Because Km values can contain information on binding as well as additional rate constants, we also inquired whether the Ka for DHF was altered by osmolyte addition. Initial isothermal titration calorimetry (ITC) experiments monitored binding of DHF to enzyme·NADP+, which results in a reasonable mimic of the catalytic ternary complex. Increasing concentrations of betaine and sucrose weaken binding of DHF as can be seen in Fig. 3A, which plots ln Ka versus osmolality. Using the relationship ln Ka/d[osmolal] =–{Delta}nW/55.6, the slope of the combined betaine and sucrose data were converted to a {Delta}nW of 29 ± 3. This value is lower than the effect of sucrose and betaine on kcat/Km (DHF) and suggests that Km may contain kinetic terms. We also present the data as a plot of total heat (Qtotal) versus [DHF] at increasing concentrations of betaine (see Fig. 4A).

Although the data for betaine and sucrose overlap in Fig. 3A, steeper slopes were observed using PEG400 ({Delta}nW = 77 ± 11), indicating DHF binding remains sensitive to osmolyte identity. We also performed titrations with Me2SO, glycerol, and ethylene glycol, and at low cosolvent concentrations, decreases in Ka were observed. However, at higher osmolyte concentrations, good fits were not obtained; thus these data are not presented.

From Fig. 4A, as the osmolyte concentration increases, the observed enthalpy value becomes smaller. Less negative {Delta}H values associated with water release have been observed previously as well as more negative {Delta}H values associated with water uptake (55, 56). Water-mediated enthalpy-entropy compensation may be the mechanism by which changes in enthalpy occur (57–60).

We additionally monitored osmolyte effects on folate binding to enzyme (i.e. formation of the enzyme·2folate complex). Large effects are observed, particularly on the enthalpy change, making fitting difficult. Thus the data are presented in Fig. 4B as a plot of Qtotal versus [folate].

To determine whether osmolytes affect NADPH binding, several ITC titrations were performed in varying concentrations of betaine (0–20%). Fig. 4C shows a plot of Qtotal versus [NADPH]. An opposite trend compared with DHF/folate binding is noted in these plots, with increasing osmolyte concentrations tightening binding. Similar trends are noted when ethylene glycol, Me2SO, or PEG400 are used. The data were fit to a sequential sites model, and a plot of ln Ka1 versus osmolality is shown in Fig. 3B. All the data overlay reasonably well. Converting the slope of this plot to {Delta}nW yields a value of –38 ± 6. This result indicates that release of water accompanies binding of NADPH, in contrast to water uptake upon binding of DHF. These opposite results are surprising as R67 DHFR possesses 222 symmetry and DHF shares a related binding site with NADPH. Also of note, we find that different osmolytes provide overlapping data when NADPH binding is probed whereas addition of different solutes yields variable slopes for DHF binding.


Figure 3
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  		FIGURE 3.
A plot of ln Ka values versus osmolality. Ka values were obtained by ITC. A contains data for DHF binding to R67 DHFR·NADP+, whereas B contains data for NADPH binding to free enzyme. Only the first Ka values (units of M–1) for the latter data set are shown. Total heats for some of the data sets are shown in Fig. 4. Data for buffer (gray circle), ethylene glycol (open circle), sucrose (open square), Me2SO (open circle), glycine betaine (open triangle), and PEG400 (checkerboard) are shown. It is clear that the DHF and NADPH binding data possess opposite slopes.

 
Is Osmolality Important for R67 DHFR Function in Vivo?—Depending on the media and the type of bacteria, cells typically adapt to osmotic stress by internally producing betaine, potassium glutamate, proline, trehalose, or other osmolytes (61–65). E. coli can tolerate over a hundredfold variance in external osmolality, from as low as 0.015 osM (66) up to ~1.9 osM (67, 68) in minimal medium. Our above studies indicate that increasing osmolality negatively affects the ability of R67 DHFR to bind DHF and proceed to the transition state. Does this have any impact with respect to function of R67 DHFR in bacteria? We addressed this question by adding increasing concentrations of sorbitol (69, 70) to M9 minimal media containing TMP and asked whether this osmolyte impacted growth of E. coli strain DH5{alpha}, which has been transformed with wild type or mutant R67 DHFR clones. This approach appeared feasible as well as unique because few reports describe uptake of water upon binding, and those that do may involve conformational changes (36, 56, 71–76). (Note: This enzyme is a homotetramer; when a single residue is mentioned, all four related residues are implied.)


Figure 4
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  		FIGURE 4.
Plots of total heat (Qtotal) versus ligand concentration. A shows three data sets for DHF binding to R67 DHFR·NADP+. The data describing binding in buffer are given by open circle points, binding in 10% betaine by open square points and binding in 20% betaine by open triangle points. B shows three data sets for folate binding to apoenzyme to form enzyme·2folate using the same symbols. C shows three data sets for NADPH binding to apoenzyme to form enzyme·2NADPH, with binding in buffer shown by open circle points, binding in 5% betaine by open square points, and binding in 20% betaine by open triangle points.

 
A first set of control plates evaluated the ability of R67 DHFR clones to provide TMP resistance, whereas the second set monitored the effect of increasing concentrations of sorbitol on cell growth. DH5{alpha} did not grow on minimal M9 control plates containing 20 µg of TMP/ml as this antibacterial agent selectively inhibits chromosomal DHFR. Transformation of pUC8 carrying the wild type (WT) R67 DHFR gene into DH5{alpha} allows growth on this selective media. Transformation of mutant R67 DHFR clones into DH5{alpha} allowed a range of growth patterns on TMP plates. The WT gene and mutants with high kcat/Km (DHF) values allowed confluent growth overnight. In contrast, the Y69L mutant (with a lower catalytic efficiency) shows growth after 3 days. The K32M mutant has a very low catalytic efficiency and low protein expression levels; it is unable to rescue DH5{alpha} from TMP-selective pressure. In the second set of control plates, all cultures were streaked on M9 minimal agar containing increasing concentrations of sorbitol to determine the effect of osmolality on cell growth. Cells showed confluent growth overnight until 1.95 osmol was reached, at which point 2 days were required for growth. Higher concentrations of sorbitol were unable to support growth.

Next, we asked whether sorbitol could impact the ability of DH5{alpha} carrying various R67 DHFR clones to grow on minimal M9 plates containing 20 µg of TMP/ml. The number of days required to obtain visible growth on these plates at 37 °C is listed in supplemental Table S1. The WT R67 DHFR clone allows growth on all sorbitol conditions until the osmolyte concentration becomes too high (or cell water content becomes too low (42)) and cell growth stops. The Q67H mutant, which has a reasonable catalytic efficiency, but displays substrate and cofactor inhibition (77), allows growth up to 1.81 osmol conditions. The I68M and Y69L mutants, with lower kcat/Km (DHF) values, allow growth up to 1.44 and 0.81 osmol conditions, respectively. Because DHFR activity is required to restore folate end product prototrophy and as cell growth parallels to some degree the efficiency of the mutants, it appears that lowering the intracellular water activity by sorbitol addition confers another level to the selection process. To our knowledge, this is the first time osmolyte addition has been successfully used as a selection tool to probe for the correlation between in vitro water uptake and in vivo function.

Heat Capacity—Because water uptake has been shown to be involved in DHF binding and water release associated with NADPH binding, we asked whether water reorganization might contribute to the binding enthalpy. As Chervenak and Toone (57) have shown that water reorganization correlates with heat capacity values, we monitored the change in enthalpy as a function of temperature. For NADPH binding, the Kd value decreases, accompanied by a {Delta}H change. The data for {Delta}H for the first NADPH site are shown in Fig. 5A. The slope ({Delta}Cp) is –178 ± 15 cal/K mol. (There are changes in {Delta}H for the second NADPH site as well, but the errors are larger.) We note that NADPH binding to R67 DHFR occurs without release or uptake of protons, as does binding of DHF to enzyme·NADP+, so protonation effects do not contribute to the {Delta}H change (14).


Figure 5
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  		FIGURE 5.
Heat capacity plots. A shows the temperature dependence of {Delta}H (solid circle), {Delta}G (open square), and –T{Delta}S (open triangle) for DHF binding to enzyme·NADP+. B shows the temperature dependence of these thermodynamic parameters for NADPH binding to free enzyme, yielding DHFR·2NADPH. The thermodynamic changes for binding of the first NADPH molecule are shown. A similar plot (with lower {Delta}Cp) is seen describing binding of the second NAPDH molecule, with a higher error (data not shown). Best fit values are given in the text.

 
When binding of DHF to enzyme·NADP+ is monitored, changes in both Kd and {Delta}H are observed. This plot is depicted in Fig. 5B, and heat capacity of –199 ± 16 cal/K mol is calculated. Both {Delta}Cp values for DHF and NADPH binding are small, negative, and within error of each other. Because a number of the interactions between NADPH and protein or DHF and protein are similar (8, 15, 78, 79), the convergence of the {Delta}Cp values may describe at some level the ability of symmetry-related sites to accommodate the two different ligands.

From Fig. 5A, NADPH binding is enthalpy- and entropy-driven and the entropy term is positive (favorable) until 281 K (8 °C). Above this temperature, the entropy term becomes negative, and binding becomes enthalpy-driven. A similar scenario occurs for DHF binding, although the point at which {Delta}S changes sign occurs at a higher temperature, 291 K (18 °C). The changes in {Delta}G are much smaller than those in {Delta}H and T{Delta}S, consistent with enthalpy-entropy compensation. Dunitz (58) has proposed that enthalpy-entropy compensation commonly occurs in weak interactions. Also this type of compensation is a hallmark of water involvement (80, 81) as calculations indicate that the {Delta}H value associated with cavity formation for accommodating a solvent molecule is exactly balanced by the entropy of the cavity (82, 83). Thus, contributions to {Delta}H and {Delta}S can occur but will not necessarily show up in {Delta}G as {Delta}Gsolvation equals zero (={Delta}Hsolvation – T{Delta}Ssolvation). Similar results have been observed previously in R67 DHFR upon titrating folate into R67 DHFR·NADPH in the presence of increasing salt concentrations. A titration in the {Delta}H value was observed but not in {Delta}G (15). Solvent-accessible area calculations have been performed and are provided in the supplemental material.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
When binding of NADPH and DHF to R67 DHFR is compared using an osmotic stress approach, two differences are noted. First, NADPH binding is accompanied by water release, whereas DHF binding requires water uptake. Second, NADPH binding is insensitive to osmolyte identity, whereas DHF binding responds differently to various classes of osmolytes. These differences are surprising given that NADPH and DHF bind at symmetry-related sites in R67 DHFR. These two issues are intertwined and are discussed below. To our knowledge, this is the first description of different responses to co-solvents for ligand binding at a promiscuous site.

Variable {Delta}nW Values—Because the slope values describing kcat/Km (DHF) vary in Fig. 2, several potential scenarios could be occurring. The first possibility is that the different slopes arise by preferential binding of the osmolyte upon the protein surface (37, 42). From Equation 3, if an osmolyte is preferentially bound by the protein, then the net change in water molecules associated with ligand binding will be overestimated if water is taken up (or underestimated if water is released). In other words, the slope can be generated by any combination of numbers for {nu}H2O and {nu}solute with the caveat that {nu}solute is a positive value (37). For example, a "{Delta}nW"of +17 could be generated by 17H2O + 0solute or 11H2O + 6solute or 5H2O + 12solute, etc.

At first glance, binding of solutes does seem possible as R67 DHFR possesses a promiscuous binding surface and could be imagined to weakly bind other molecules as well. However, NADPH binding serves as an internal control. If osmolytes do bind and provide a range of slopes for the DHF-binding site, why isn't the same phenomenon observed for NADPH? In other words, because DHF and NADPH bind at symmetry-related sites, we might expect both slopes in Fig. 2 and Fig. 3B to be sensitive to osmolyte identity. At 1–2 osmol, Fig. 2 shows the kcat/Km (DHF) values clearly separating with different osmolytes, whereas at ~2 osmol in Fig. 3B, the Ka values describing NADPH binding overlap.

A second option that could describe the variable slopes is crowding or volume exclusion (43–45). This term describes the volume of the protein probed plus the volume swept out by a sphere with radius r rolled over the surface. Because of different radii, the excluded volume varies for each cosolvent with larger osmolytes probing larger surface features. PEGs of varying sizes were used to analyze this possibility. The average radius of gyration for PEG400 is 8.1 Å; however, its radius increases because of hydration; the radii probed vary from 9 to 15 Å, depending on the macromolecule studied (84). This range of values contrasts with a maximal pore radius in R67 DHFR of 12 Å (5). When PEG400, PEG3350, and PEG8000 are used to probe DHF binding, the slopes of the ln aH2O versus ln kcat/Km(DHF) plot increase to 78, 145, and 353, respectively (Table 1). The increase in slope indicates the PEGs are probing larger surface features as the osmolyte becomes too big to fit in the active site pore. Supplemental Fig. 3 shows a plot of ln molar volume (Vmol) versus the slopes listed in Table 1, indicating crowding plays some role, particularly with larger molecular weight PEGs. However, if crowding causes the variable slopes associated with DHF binding in Fig. 2, then one might expect crowding to affect NADPH binding in a similar fashion. Because this is not observed, crowding is probably not the origin of the behavior.

Another possibility is that conformational changes occur in the protein and/or ligands such that release of water upon ligand binding is masked by water uptake associated with the conformational change, which results in a larger surface area. Examples of this phenomenon include glucose binding to hexokinase (85), aspartate to aspartate transcarbamoylase (71), DNA to repressors (30), etc. However NMR studies of R67 DHFR indicate little change in the generalized order parameter upon binding NADP+ (<Sfree2> = 0.89, <Sbound2> 0.86), consistent with a fairly rigid structure (12). Small changes may occur when the apo and ternary complex structures are compared, as introduction of asymmetry, particularly at Gln-67 and Tyr-69 residues, is observed (8).

Another origin of the different slope effects could lie in the fact that DHF binds to an enzyme·NADPH complex, whereas NADPH binds to the apoenzyme. However, binding of a second NADPH molecule to enzyme·NADPH still shows water release and overlapping data with varying osmolytes ({Delta}nW =–20 ± 8; data not shown), and binding of two folate molecules to the free enzyme shows very large effects (Fig. 4B). Both of these observations suggest it is the identity of the ligands rather than the position in the binding sequence (i.e. first versus second molecule) that causes the different behavior.

Because NADPH and DHF compete for binding to related sites in R67 DHFR, it seems likely that osmolyte sensitivity is either reporting on differences in how the ligands bind, differences in solvation of the two ligands (86, 87), differences in solvent that may be induced by osmolyte addition, and/or differences in protein solvation levels (88, 89). For example, for the latter possibility, Dzingeleski and Wolfenden (72) saw an uptake of 9 water molecules associated with substrate and inhibitor binding in adenosine deaminase. They proposed that a hydrated form of this enzyme is in equilibrium with a dehydrated form and that only the hydrated form can bind substrate. A similar mechanism could exist in R67 with NADPH preferentially binding to the dehydrated form and DHF preferentially interacting with the hydrated form. At the moment, we are unable to differentiate between the models proposed in this paragraph.

Correlation of in Vitro Results with in Vivo Function?—Given the uncertainty in interpreting the variable slopes associated with DHF binding, we note that what is important is that the results of our in vitro experiments correlate with the in vivo efficacy of R67 DHFR. For example, if we consider the Y69L mutant clone, its catalytic efficiency and intracellular protein concentration appear barely sufficient to allow DH5{alpha} to grow in the presence of TMP. This is not surprising as its Km (DHF) equals 180 µM (90); however, the normal intracellular DHF concentration in E. coli is ~300 µM (91, 92); thus it is likely working under kcat/Km conditions. From Fig. 2, these are exactly the conditions where we might expect to observe the greatest osmolyte effect. If betaine is one of the intracellular osmolytes produced in response to osmotic stress (42), there should be a large inhibitory effect on R67 DHFR efficiency. Thus addition of sorbitol increases the osmotic pressure, cellular water content decreases, and the Y69L mutant is no longer able to support growth on TMP plates.

Extrapolating from our in vitro studies to in vivo conditions, we also might expect increasing osmolyte concentrations to tighten NADPH binding to both sites, potentially leading to inhibition by the two NADPH complex (see Fig. 5C). Concurrently, for the two substrate complex, we would predict that it is less likely to form in the cell (Fig. 5B). As we have shown, formation of the transition state is compromised upon addition of osmolytes. This sensitivity to water content suggests R67 DHFR is not very efficient in vivo, supporting our previous hypothesis that R67 DHFR is a primitive enzyme (4).

Conclusion—R67 DHFR is an unusual enzyme that uses symmetry-related sites to bind two different ligands. NADPH interacts with R67 DHFR through specific contacts, and a unique bound conformation exists (8). This structural description is consistent with results from this study, where removing surface water by osmotic stress results in tighter binding of NADPH. In contrast, DHF utilizes specific contacts for binding of its pteridine ring (8); however, its para-aminobenzoylglutamate tail of dihydrofolate/folate is disordered. This different mode of binding may correlate with its different response to osmolyte addition. Because NADPH serves as an internal control, whatever effect causes the different behavior associated with DHF binding must lie in either the free ligand, free protein, or in the different mode of binding in the complexes. Clearly interfacial water contributes to binding plasticity in R67 DHFR and can provide hydrogen bonds either alone or in networks between ligand and protein (93–97). In other words, these osmolyte studies indicate that NADPH and R67 DHFR interact more directly, utilizing more protein contacts (i.e. a "dry interface" according to Janin (98)), whereas the DHF interaction with protein uses more water contacts ("wet interface"). This view proposes water is intimately involved in binding specificity as well as affinity in R67 DHFR.


   FOOTNOTES
 
* This work was supported by National Science Foundation Grant MCB-0445728. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

Formula The on-line version of this article (available at http://www.jbc.org) contains supplemental text, equations, references, Figs. 1–4, and Table S1. Back

1 To whom correspondence should be addressed: Dept. of Biochemistry, Cellular and Molecular Biology, University of Tennessee, Knoxville, TN 37996-0840. Tel.: 865-974-4507; Fax: 865-974-6306; E-mail: lzh{at}utk.edu.

2 The abbreviations used are: DHFR, dihydrofolate reductase; DHF, dihydrofolate; ITC, isothermal titration calorimetry; MES, 4-morpholineethanesulfonic acid; PEG, polyethylene glycol; TMAO, trimethylamine N-oxide; TMP, trimethoprim. Back


   ACKNOWLEDGMENTS
 
We thank Peter Mazur, Robert Rosenberg (University of North Carolina), and Mike Fried (University of Kentucky) for use of their water vapor osmometers. David Golden generously allowed us to use his Aqualab water activity meter. We also thank Jeremy Smith, Todd Reynolds, Mike Fried, Adrian Parsegian (National Institutes of Health), Nina Sidorova (National Institutes of Health), and Don Rau (National Institutes of Health) for helpful discussions and Engin Serpersu, Can Ozen, and Cynthia Peterson for critical reading of the manuscript. We also appreciate Don Nguyen, who performed the Qtotal analysis of the ITC data.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

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