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Comparison of crystal structures of human type 3 3-hydroxysteroid dehydrogenase reveals an "induced-fit" mechanism and a conserved basic motif involved in the binding of androgen

Oncology and Molecular Endocrinology Research Center, Laval University Medical Center (CHUL) and Laval University, Québec, G1V 4G2, Canada

Reprint requests to: Rock Breton, Centre de Recherche en Endocrinologie Moléculaire et Oncologique, Centre Hospitalier de l’Université Laval, 2705, boul. Laurier, Ste-Foy, QC, G1V 4G2, Canada; e-mail: rock.breton{at}crchul.ulaval.ca; fax: (418) 654-2761.

(RECEIVED January 11, 2005; FINAL REVISION March 17, 2005; ACCEPTED March 18, 2005)

	Abstract

TOP Abstract Introduction Results and Discussion Materials and methods References

 
The aldo-keto reductase (AKR) human type 3 3-hydroxysteroid dehydrogenase (h3–HSD3, AKR1C2) plays a crucial role in the regulation of the intracellular concentrations of testosterone and 5-dihydrotestosterone (5-DHT), two steroids directly linked to the etiology and the progression of many prostate diseases and cancer. This enzyme also binds many structurally different molecules such as 4-hydroxynonenal, polycyclic aromatic hydrocarbons, and indanone. To understand the mechanism underlying the plasticity of its substrate-binding site, we solved the binary complex structure of h3–HSD3-NADP(H) at 1.9 Å resolution. During the refinement process, we found acetate and citrate molecules deeply engulfed in the steroid-binding cavity. Superimposition of this structure with the h3–HSD3-NADP(H)-testosterone/acetate ternary complex structure reveals that one of themobile loops forming the binding cavity operates a slight contraction movement against the citrate molecule while the side chains of many residues undergo numerous conformational changes, probably to create an optimal binding site for the citrate. These structural changes, which altogether cause a reduction of the substrate-binding cavity volume (from 776 ų in the presence of testosterone/acetate to 704 ų in the acetate/citratecomplex), are reminiscent of the "induced-fit" mechanism previously proposed for the aldose reductase, another member of the AKR superfamily. We also found that the replacement of residues Arg₃₀₁ and Arg₃₀₄, localized near the steroid-binding cavity, significantly affects the 3–HSD activity of this enzyme toward 5-DHT and completely abolishes its 17–HSD activity on 4-dione. All these results have thus been used to reevaluate the binding mode of this enzyme for androgens.

Keywords: aldo-keto reductase; hydroxysteroid dehydrogenase; crystal structure; induced-fit mechanism

Abbreviations: h3–HSD3, human 3-hydroxysteroid dehydrogenase type 3 • h20–HSD, human 20-hydroxysteroid dehydrogenase • h3– HSD1, human 3-hydroxysteroid dehydrogenase type 1 • 17–HSD, 17-hydroxysteroid dehydrogenase • NADP(H), reduce form of nicotinamide adenine dinucleotide phosphate • GST, glutathione sulfotransferase • PDB, Protein Data Bank • AKR, aldo-keto reductase • progesterone, 5-pregnen- 3,20-dione • 4-dione, 4-androsten-3,20-dione • PAH, polycyclic aromatic hydrocarbon • EDTA, ethylene diamine tetraacetic acid.

Article and publication are at http://www.proteinscience.org/cgi/doi/10.1110/ps.051353205.

	Introduction

TOP Abstract Introduction Results and Discussion Materials and methods References

 
Hydroxysteroid dehydrogenases (HSDs), which form a subgroup in the aldo-keto reductase (AKR) superfamily (AKR1C), are well known to selectively catalyze the oxydo-reduction of hydroxyl/ketone groups found at specific positions on steroid molecules (Dufort et al. 1996, 1999, 2001; Zhang et al. 2000) (Fig. 1A). Among the members of the AKR1C subgroup, human type 3 3–HSD (h3–HSD3; AKR1C2; EC 1.1.1.213 [EC] ) is able to exert its activity on structurally different steroids, such as androgens (C19-steroids) and progestins (C21-steroids). In addition to steroids (Dufort et al. 1996, 2001; Penning et al. 2000), this enzyme is also able to bind/transform numerous molecules with various shapes, such as acetate (Nahoum et al. 2001), prostaglandins (Lovering et al. 2004), polycyclic aromatic hydrocarbons (PAHs) (Palackal et al. 2001, 2002), 1-indanol (Matsuura et al. 1997), 9, 10-phenanthrenequinone (Penning et al. 1984), and 4-hydroxy-2- nonenal (Burczynski et al. 2001) (Fig. 1B). Because all these structurally different molecules bind in the ligand-binding cavity, mainly via hydrophobic contacts, h3– HSD3 must adapt the structure of its binding cavity in order to form an optimal binding site for each of them. This capacity to alter the orientation of numerous side chains surrounding the ligand-binding site, which was first described for the yeast hexokinase, is named the "induced-fit" mechanism (DelaFuente and Sols 1970; DelaFuente et al. 1970). The induced-fit mechanism is also known for members of the AKR superfamily and has been well studied, notably in the case of the aldose reductase, in a strategy for the design of inhibitors (Klebe et al. 2004; Sotriffer et al. 2004).

View larger version (33K): [in this window] [in a new window]   Figure 1. Schematic representation of enzymatic properties of HSDs member of the AKR superfamily. (A) Reactions catalyzed by h3–HSD3 on 4-dione and DHT. (B) Overview of the molecules that bind and/or are transformed by HSDs member of the AKR superfamily (the figure has been created with SwissPDBViewer and rendered with Povray; Kaplan and Littlejohn 2001). (C) Orbital steering constraints proposed by Heredia et al. (2003). Here, the optimal distance from the catalytic Tyr residue, the angle of the hydride with the target ketone (in parentheses), and the distance of the C4 of the pyridine head of the cofactor are indicated (created with Molscript and POVScript; Kraulis 1991; Fenn et al. 2003).

Considering these recent findings—the pronounced induced-fit adaptations shown for at least one member of the AKR enzyme family and the precise localization of the ketone group to be reduced inside the active site—and taking advantage of ternary complex crystal structures we have determined so far for many AKR1C enzymes (Nahoum et al. 2001; Couture et al. 2003, 2004), we decided to revisit all these structures, especially those for which the steroid substrate was found in the steroid-binding cavity but at a nonproductive position (Nahoum et al. 2001; Couture et al. 2003, 2004). We wished to investigate some pending issues: (1) the mechanism by which h3–HSD3 binds/transforms numerous structurally different molecules, which implies an ability to discriminate various ligands from different steroid hormone classes; (2) the orientation adopted by a steroid in a productive complex; and (3) the geometry of the residues that surround the ligand in the productive complexes.

Here we report the crystal structure of h3–HSD3 in binary complex with NADP(H), a complex in which a citrate and an acetate molecule, added during the crystallization step, were found deeply bound inside the steroid-binding cavity. Comparison with the testosterone/acetate ternary complex structure reveals that loop A undergoes a slight contraction toward the citrate. Moreover, numerous residues found in the cavity, namely Val₅₄, Val₁₂₈, Ile₁₂₉, Trp₂₂₇, Leu₃₀₆, and Leu₃₀₈, alter the orientation of their side chains, narrowing the cavity. All these structural changes are reminiscent of the induced-fit mechanism previously proposed for the aldose reductase (Sotriffer et al. 2004). We also found that, in order to respect the orbital steering constraint, the optimal position of the ketone of a steroid during the reduction reaction is somewhat similar to that of the acetate molecule found in the active site. Manual positioning of 4-dione and DHT in an ideal productive ternary complex revealed that the nucleus of the steroid must be oriented toward the C-terminal domain (loop C) of the protein. To verify this, we proceeded to a leucine-scanning mutagenesis study on many residues of loop C, and found that the replacement of residue Arg₃₀₁ or Arg₃₀₄, located 10 Å from the catalytic Tyr₅₅, significantly affects the activity of this enzyme. These results suggest that residues involved in the formation of a mature binding cavity in h3–HSD3 differ according to the nature of the steroid substrate, the position of the ketone group on the steroid nucleus, and the activity exerted by this enzyme.

	Results and Discussion

TOP Abstract Introduction Results and Discussion Materials and methods References

 
Overall structure of the h3–HSD3 binary complex
The crystal used to determine the binary complex structure belonged to the R32 space group and contained two molecules per asymmetric unit (monomers A and B). The crystal structure of the h3–HSD3/NADPH binary complex was refined to a crystallographic R-factor of 17.4% (R_free=19.9%) at 1.9 Å of resolution. No major differences were found when both monomers were compared (root-mean-square deviation, calculated for 327 C atoms is 0.36 Å). The model obtained was of good quality since more than 99.7% of the dihedral angles of the main chain were found in the allowed region of the Ramachandran plot (Table 1). Residue Ser₂₂₁ was found in the disallowed region due to a hydrogen bond between its nitrogen and the pyrophosphate group of NADPH. The protein was folded as a triose phosphate isomerase barrel motif [(/)₈-barrel], in which eight strands forming the cylindrical core of the barrel are surrounded by eight helices. The first 17 residues are folded as a two-strands anti-parallel beta-sheet that closes the bottom of the barrel. Finally, three large loops, named A (Phe₁₁₈ to Asp₁₄₃), B (Ser₂₁₇ to Asp₂₃₈), and C (Leu₂₉₉ to Tyr₃₂₃) are situated in a manner that creates a funnel-shaped pocket at the carboxy end of the -strands, the top of the barrel (Fig. 2A).

View larger version (80K): [in this window] [in a new window]   Figure 2. Structure of h3–HSD3 in binary complex with NADP(H) crystallized in the presence of citrate and acetate salts. (A) Overall view of a superposition of the h3–HSD3 complex structures with testosterone (RCSB PDB entry, 1J96 [PDB] ; black) and with citrate and acetate (monomer A) (RCSB PDB entry, 1XJB; gray). (B) Close-up stereo view of the citrate binding cavity of h3–HSD3. The 2(Fo–Fc) electron density map for citrate (dark gray) and acetate (light gray) are respectively contoured at 0.9 and 1.0 level. (C) Schematic representation of interactions made by citrate and acetate (thick lines) within the binding cavity. Only residues that interact with ligands are shown. The dashed lines and the number in parentheses represent the hydrogen bonds and the H-bond distances, respectively. (D) Stereo view of the superposition of the h3–HSD3 complex structures with citrate and acetate (monomer A; white) and with testosterone (gray). Only residues affecting the size of the steroid-binding cavity are depicted for both complexes.

Steroid-binding site
During the refinement process, two unexpected positive peaks appeared inside the steroid-binding cavity of monomer A. Citrate and acetate molecules, added to the mother liquor during the crystallization step, were fitted into the F_o–F_c difference Fourier map and refined with the rest of the model (Fig. 2B). These two molecules were surrounded by residues Tyr₂₄, Ala₂₅, Val₅₄, Tyr₅₅, Trp₈₆, His₁₁₇, Ile₁₂₉, Asn₁₆₇, Tyr₂₁₆, His₂₂₂, Glu₂₂₄, Trp₂₂₇, Leu₃₀₆, Leu₃₀₈, Ile₃₁₀, and Phe₃₁₁ (Fig. 2C). Together, these residues form a narrow cavity where the acetate and the citrate are tightly bound. At the bottom of the cavity, the acetate molecule occupies the catalytic binding site of the h3–HSD3 enzyme with the C=O of its carboxyl group mimicking the ketone group of a steroid substrate. Indeed, one of the oxygen atoms of this carboxylate group is involved in a hydrogen bond network that includes the OH group of Tyr₅₅ (2.7 Å), the N atom of His₁₁₇ (2.7 Å), in addition to the N of Lys₈₄ (located at 3.1 Å from the tyrosine hydroxyl group), while the carbon atom is located at 3.3 Å from the C-4 of the nicotinamide head of the cofactor, the hydride donor atom. The presence of an acetate molecule well stabilized inside the catalytic binding site of an enzyme of the AKR superfamily has been reported before (Nahoum et al. 2001; Lovering et al. 2004), but this is the first time it is seen orientated so as to interact with the catalytic residues (Tyr₅₅ and His₁₁₇) and, at the same time, participate in the ligand stabilization, here the citrate molecule. Indeed, one of the oxygen atoms of the acetate molecule carboxylate group is equidistant (2.5 Å) from two oxygen atoms (O2 and O7) of the citrate molecule with which it could share a hydrogen atom (Fig. 2C). In addition, two other oxygen atoms of the citrate molecule are involved in hydrogen bonds with residues of the h3–HSD3 enzyme: O2 with the side chain (N²) of residue His₂₂₂ (2.6 Å) and O5 with the hydroxyl group of Tyr₂₄ (2.7 Å) and with the nitrogen atom of the indole ring of Trp₂₂₇ (2.8 Å).

The numerous potential hydrogen bonds that stabilize the acetate and citrate in the substrate-binding cavity of h3–HSD3 led us to think that these two molecules could probably interfere in the binding of the steroid, which normally forms only one hydrogen bond with the enzyme, and thus could be potential inhibitors of h3–HSD3. We thus tested the capacity of the acetate and citrate to inhibit the 3-reduction of DHT and found that none of them, alone or together, could measurably inhibit the h3–HSD3 activity (data not shown). This result suggests that weaker and less specific forces such as van der Waals bonds or hydrophobic interactions, which are well known to mediate the binding of steroids and other structurally different ligands by h3–HSD3, are probably much more important than anticipated, provided of course that the plasticity of the substrate pocket is sufficient to insure the steric complementarity between the enzyme and its ligands.

Citrate bound in a steroid-binding cavity suggests a ligand induced-fit mechanism
The side chains of residues Tyr₅₅ and His₁₁₇ form an anionic binding site that could bind molecules bearing a negative charge. This, combined with the fact that the substrate-binding site is funnel shaped and largely open on the protein surface, probably explains why human 3– HSD3 can bind and transform numerous molecules that vary in length and shape. Indeed, it has been shown that this enzyme can bind acetate (Nahoum et al. 2001), citrate (this paper), indanol (Matsuura et al. 1997), steroids (Dufort et al. 1996; Penning et al. 2000), and PAHs (Burczynski et al. 1998). To understand the mechanism underlying this plasticity we have superposed and compared our model (h3–HSD3/NADP(H)/Citrate/Acetate) with another h3–HSD3 complex structure (h3–HSD3/NADP(H)/Testosterone/Acetate). We first observed that a short segment of loop A, between residues Pro₁₂₄ and Phe₁₃₉, makes a movement against the citrate molecule that slightly closes the top of the binding cavity (Fig. 2A). Loop A is one of the three loops involved in the formation of the testosterone-binding cavity in rat and human 3–HSD enzymes. It has been previously suggested for HSDs of the AKR superfamily that this loop plays a key role in the recognition, binding, and transformation of their substrates since a chimera rat 3–HSD having the loop A found in human 20–HSD sequence acquires 17- and 20–HSD activities on testosterone and progesterone, respectively (Ma and Penning 1999). Among conformational changes observed in this segment, the backbone of residues Val₁₂₈ and Ile₁₂₉ moves 0.8 Å toward the citrate, contributing to a reduction of the cavity opening. These two residues are fairly important since they directly interact with and participate in the binding of testosterone and 20-OHProg by h3–HSD3 and h20–HSD, respectively (Nahoum et al. 2001; Couture et al. 2003). In addition, previous studies on h20–HSD enzymes revealed that the side chain of another residue of loop A, Glu₁₂₇, rotates upon binding of the steroid and slightly closes the top of the cavity, preserving the hydrophobic environment of the steroid-binding site (Couture et al. 2003). All these observations support the importance of the role played by loop A in the formation of the ligand-binding cavity and in the plasticity of this site. In comparison with loop A, the main chains of loops B and C do not undergo a significant structural shift (Fig. 2A) but close inspection of the super-imposed binding site regions reveals that some residues that belong to these loops (Trp₂₂₇, Leu₃₀₆, and Leu₃₀₈) or which delineate the cavity (Val₅₄) make important conformational changes and become closer to the citrate molecule (Fig. 2D).

Compared with the model of h3–HSD3 in ternary complex with testosterone/acetate, the side chain of residue Val₅₄ in the present model undergoes a rotation of 180° around its C, placing its C atom toward the binding cavity (Fig. 2D), probably to stabilize the citrate molecule. Interestingly, the importance of this residue for the selectivity of HSDs of the AKR superfamily has been previously demonstrated by site-directed mutagenesis experiments (Matsuura et al. 1997; Couture et al. 2003, 2004). For example, Matsuura et al. (1997) demonstrated that the replacement of residue Val₅₄ by a leucine (residue found in the h20–HSD sequence) confers to the mutated h3–HSD3 almost the same catalytic properties as h20–HSD. Now, our structural results tend to demonstrate that residue Val₅₄, through its capacity to adopt different conformations, is also important for the plasticity of the ligand-binding cavity of h3–HSD3.

We also observed that the indole ring of residue Trp₂₂₇ moves toward the citrate molecule, where it is stabilized through a hydrogen bond between the O5 and the nitrogen atom of its indole ring (Fig. 2D). In the ternary complex structure of h3–HSD3 with testosterone, this residue makes a -stacking interaction with cycles B and C of testosterone and greatly contributes to the formation of a mature steroid-binding cavity (Nahoum et al. 2001). In r3–HSD, the replacement of Trp₂₂₇ has been shown to affect the capacity of this enzyme to bind and transform testosterone and progesterone (Heredia et al. 2004) and, more importantly, its replacement can affect the activity on decalone and 9,10-phenanthrenequinone (Fig. 1B). Combined with the h3–HSD3/NADP(H)/Acetate/Citrate ternary complex structure study, these results suggest that upon the binding of a non-steroid-shaped ligand, this residue is of great importance for the optimal formation of a mature cavity and for the plasticity of this enzyme.

Finally, the side chains of residues Leu₃₀₆ and Leu₃₀₈ also rotate 80° around their C toward the citrate molecule (Fig. 2D). This "leucine-flip" thus seems to play a role in the plasticity of this cavity. Similarly, a recent study on aldose reductase has pointed out the Leu₃₀₀, a residue found in loop C (also called specificity pocket), as an important factor in the modulation of the shape of this cavity. It appears that the conformation adopted by the side chain of this residue with the backbone shift observed for residues Ala₂₉₉ and Leu₃₀₀ are directly related to the shape of the bound molecule (Sotriffer et al. 2004). Our finding indicating that the conformational change undergone by Leu₃₀₆ and Leu₃₀₈ reduced the size of the binding cavity of h3–HSD3 upon binding of the citrate is thus in total agreement with the model proposed for the aldose reductase, even if the contraction movement observed here does not affect the backbone position of these two residues.

To ascertain that the size of the h3–HSD3 substrate cavity really changes upon binding of its structurally different substrates, and thus confirm our first visual finding, we decided to precisely measure the volume of this surface groove in the presence of the acetate/citrate (monomer A) and compare it with that of the same enzyme in complex with testosterone/acetate (Nahoum et al. 2001) (RCSB PDB entry, 1J96 [PDB] ) and ursodeoxycholate (Jin et al. 2001) (RCSB PDB entry, 1IHI [PDB] ). We thus found that the substrate cavity volume was directly related to the size of the bound ligand. Indeed, the cavity volume for the acetate/citrate (Fig. 3A) was 704 ų compared to 776 ų for testosterone/acetate (Fig. 3B), in agreement with the size of these molecules (236 ų vs. 333 ų). Interestingly, the cavity volume in the ursodeoxycholate complex (734 ų) is smaller than the testosterone cavity, although ursodeoxycholate (362 ų) is slightly bigger (Fig. 3C). This apparent contradiction could be easily explained by the difference in the binding mode adopted by these two steroidal substrates. The ursodeoxycholate is bound in such a manner that its long and thin 17-(1-methyl-3- carboxypropyl) arm is positioned close to the catalytic site of the enzyme, pushing almost half of the steroidal nucleus (cycle A and a large part of cycle B) out of the cavity (Fig. 3D). In this position, the ursodeoxycholate is in fact a "smaller" ligand than testosterone (248 ų vs. 277 ų), which is almost totally inside the cavity.

View larger version (116K): [in this window] [in a new window]   Figure 3. Variation in the surface of the steroid-binding cavity of h3–HSD3 in complex with structurally different ligands. Surfaces of the steroid-binding cavity of h3–HSD3 in complex with (A) acetate/citrate, (B) testosterone/acetate (Nahoum et al. 2001), and (C) ursodeoxycholate (Jin et al. 2001) have been calculated with the program VOIDOO (Kleywegt 1994). (D) Superposition of various ligands (testosterone, blue, and ursodeoxycholate, green) in complex with the h3–HSD3 enzyme to compare their mode of binding. Acetate and citrate molecules, which are profoundly engulfed inside the steroid-binding cavity, are not visible from this angle. Figures have been rendered with Pov-Ray (Kaplan and Littlejohn 2001).

What is the real productive ternary complex?
Residues found in loops A, B, and C are of great importance in the plasticity of the steroid-binding cavity of h3–HSD3, a characteristic that allows this enzyme to bind molecules with various shapes and structures. In addition, these loops must optimally orient the incoming molecules to promote their transformation. Unfortunately, each HSD member of the AKR superfamily for which the ternary complex structure has been solved shows its steroid substrate in a nonproductive position, i.e., in a position (or in an orientation) that does not permit the oxido-reduction reaction to take place (Bennett et al. 1997; Nahoum et al. 2001; Couture et al. 2004). According to recent reports, HSDs of the AKR superfamily would transform steroids through an orbital steering mechanism where the optimal distance and angle between the carbon atom of the substrate ketone group and C4 of the nicotinamide would be around 3.0 Å and 100°, respectively (Heredia et al. 2003). Close inspection of the h3–HSD3 structure with the citrate and acetate molecules bound in the substrate cavity (monomer A) reveals that this optimal position corresponds exactly to the carbon atom of the carboxylate group of the acetate molecule.

To better understand how h3–HSD3 can bind androgen in a productive complex, we manually modeled steroid molecules (4-dione and DHT) in a position that respects the orbital steering constraints. When these two structurally similar steroids were modeled with their reactive group in an optimal position (the C3 for the DHT and C17 for the 4-dione), their steroid nuclei were found in different positions (Fig. 4). For DHT, the nucleus of the steroid was roughly directed toward loop C and clashed with the indole ring of Trp₂₂₇, a residue belonging to loop B and that is highly conserved among AKR members. Interestingly, we had previously found that Trp₂₂₇, with many other residues of loop B, can undergo significant conformational changes to promote the binding of the steroid in the crystal structure of rb20–HSD in ternary complex with testosterone (Couture et al. 2004). In fact, we have found that Trp₂₂₇ with some other residues of loop B that point toward the steroid-binding cavity in the rb20–HSD binary complex, probably thus maintaining the hydrophobic character of the steroid cavity, are at the surface of the protein and exposed to the solvent upon binding of testosterone. This relaxation movement results in an enlargement of the steroid-binding cavity and in the mobilization of some residues belonging to the C-terminal part of the protein (loop C) to create the mature steroid-binding site (Couture et al. 2004). Crystal structure determination of r3–HSD in ternary complex with testosterone also reveals that the indole ring of Trp₂₂₇ can rotate by 180° around its C, a movement that drastically changes the shape of the cavity and allows DHT to bind in a productive complex.

View larger version (24K): [in this window] [in a new window]   Figure 4. Stereo view of the hypothetic orientation of two different androgens for their optimal reduction by h3–HSD3. The target ketone group of 4-dione (light gray) and DHT (dark gray) were manually superimposed on the acetate molecule. Only residues in proximity of the steroid are depicted. Figure was created with POVScript (Fenn et al. 2003) and rendered with Pov-Ray (Kaplan and Littlejohn 2001).

3–HSD1

A C-terminal basic motif, a concerted way to bind androgen?
We thus found that, in order to respect the orbital steering constraints, the steroid should be maintained approximately parallel to the pyridine head of the cofactor with the body of the steroid pointing toward the C-terminal part of the protein. To evaluate the role of residues of this domain in steroid discrimination and binding, we undertook a systematic leucine scanning mutagenesis study. We found that the replacement of two residues, Arg₃₀₁ and Arg₃₀₄, leads to a drastic effect on the capacity of h3– HSD3 to transform androgen. Indeed, when the kinetic constants for the reduction of DHT were compared with the wild-type h3–HSD3 (Table 2), we found that K_m for the 3-reduction of DHT was barely affected, whereas k_cat was 60- and 227-fold lower for the Arg₃₀₁Leu and Arg₃₀₄Leu mutants, respectively. For the 17-reduction of 4-dione, we found that the replacement of Arg₃₀₁ affects the K_m and the k_cat values by 3.8- and 7.3-fold, respectively. More strikingly, we found that the 17-activity on 4-dione was totally abolished for the Arg₃₀₄Leu mutant enzyme. No activity was detectable even by using radio-activity, the most sensitive technique to detect the transformation of a steroid (Luu-The et al. 1995).

View this table: [in this window] [in a new window]   Table 2. Steady-state kinetic parameters for steroid reduction catalyzed by homogeneous recombinant h3-HSD3s

View this table: [in this window] [in a new window]   Table 3. Effect of mutations on the binding of NADPH by homogeneous recombinant h3-HSD3s

View this table: [in this window] [in a new window]   Table 4. Transformation of 1-indanone by homogeneous recombinant h3-HSD3s

View larger version (36K): [in this window] [in a new window]   Figure 5. Sequence alignment of the C-terminal end of 12 HSDs member of the AKR superfamily. The alignment of the C-terminal sequences (residues 291 to 323 or 322 for AKR1C9) was done with CLUSTALW. Residues 301 and 304 in the human 3–HSD type 3 sequence are boxed while corresponding residues in other AKR members are indicated in bold characters. Abbreviated names for HSDs are AKR1C1, human 20–HSD; AKR1C2, human 3–HSD type 3; AKR1C3, human 17–HSD type 5; AKR1C4, human 3–HSD type 1; AKR1C5, rabbit 20–HSD; AKR1C6, mouse 17–HSD; AKR1C7, bovine prostaglandin F synthase 1; AKR1C8, rat 20–HSD; AKR1C9, rat 3–HSD; AKR1C10, frog rho-crystallin; AKR1C11, prostaglandin F synthase 2; and AKR1C18, mouse 20–HSD.

These observations also have important physiological relevance since, in the prostate tissues, h3–HSD3 is able to inactivate the most potent androgen, DHT, into a weak androgen, 3-diol, while this enzyme is also involved in the formation of testosterone, a potent androgen, through its 17-reduction activity on 4-dione. If the residues involved in the binding of DHT and 4-dione differ, it is tempting to speculate that it would be feasible to synthesize an inhibitor specific for the 17-reduction of 4-dione that does not affect, or very weakly affects, the 3-inactivation of DHT. In this regard, it will be essential to take into account that some residues of the h3–HSD3 binding cavity involved in the binding and maintaining of the steroid substrates can undergo significant conformational changes, in part dictated by the shape of the steroid itself. Similar to the "ligand induced-fit mechanism" reported for the aldose reductase, this binding site adaptation is of major importance in the efficiency of inhibitors targeted against this enzyme.

	Materials and methods

TOP Abstract Introduction Results and Discussion Materials and methods References

 
Material
NADPH, 1-indanone, and other chemical products were purchased from Sigma-Aldrich Canada. Unlabeled steroids and [¹⁴C]-labeled steroids were purchased from Steraloids and PerkinElmer, respectively.

Site-directed mutagenesis
Arg₃₀₁Leu and Arg₃₀₄Leu mutations were created using Quick- Change Site-Directed Mutagenesis Kit (Stratagene) with these forward primers: Arg₃₀₁Leu, AAAGCCATAGATGGCC TAAACCTAAATGTGCGATATTTGACCCTTGATAT and Arg₃₀₄Leu, CATAGATGGCCTAAACAGAAATGTGCTAT ATTTGACCCTTGATATTTTTGATGGCC. To confirm the presence of the desired mutation and to ascertain that no other mutations had occurred, the complete coding region of the mutated cDNA was sequenced with a T7 sequencing kit (Amersham Bioscience).

Overexpression and purification of wild-type and mutant h3–HSD3
Briefly, the cDNA encoding h3–HSD3 was subcloned in a pGEX vector (Amersham Bioscience), overproduced in E. coli BL21(DE3)pLysS as a fusion protein with glutathione-S transferase (GST) and purified by a combination of affinity, anionic, and size-exclusion chromatographies, yielding about 5 mg of a highly purified enzyme preparation per 100 mL of cell culture. Mutated h3–HSD3s were overproduced and purified to homogeneity as described for the wild-type enzyme.

Determination of kinetic parameters
Reduction reactions on [¹⁴C]-steroids were performed as previously described, with minor modifications (Luu-The et al. 1995; Couture et al. 2002). Kinetic parameters (K_m and k_cat/K_m) were calculated using the EnzFitter program (Biosoft). Because some HSDs of the AKR superfamily have been shown to lose their activity when they are purified to homogeneity (Dufort et al. 1999), a kinetic characterization was performed with DHT and 4-dione, two natural ligands of this enzyme, to verify that the purification steps did not affect the enzyme activity. Within the experimental error, K_m values obtained for the 3-reduction of the DHT (5.7 ± 0.8 µM) and for the 17-reduction of 4-dione (10.5 ± 1.8 µM) (Table 2) were very similar to those published earlier (1.4 ± 0.66 µM and 9.73 ± 1.8 µM [Penning et al. 2000; Dufort et al. 2001]), indicating that our h3–HSD3 preparation was suitable for subsequent experiments.

Reduction of 1-indanone was measured at room temperature in a 50-µL system containing 100 µM 1-indanone in 2% (v/v) ethanol, 200 µM NADPH, and 100 mM KH₂PO₄–K₂HPO₄ (pH 7.5). Reactions were started when 10 µg of homogenous recombinant type 3 h3–HSDs were added to the system in which 1-indanone transformation was monitored with a Beckman DU-7400 spectrophotometer by measuring the rate of change of absorbance of NADPH ( = 6270 cm^–1 M^–1) in a 1-cm light path.

Fluorescence
Measurements of the fluorescence of wild-type, Arg₃₀₁Leu, and Arg₃₀₄Leu enzymes were conducted on an SLM 8000 fluorometer. Scans of the fluorescence emission spectra were performed at 25°C in a 200-µL microcell containing 2 µM of the purified enzyme, 50 mM Na₂HPO₄/NaH₂PO₄ buffer (pH 7.5), 20% glycerol, 1 mM EDTA, and variable concentrations of NADPH (0–4000 nM). Tryptophan fluorescence was detected using 290 nm as the excitation wavelength, while the emission was scanned from 300 to 600 nm (at 120 nm/min). Emission and excitation bandpasses were both set to 4 nm.

Crystallization
A homogenous preparation of h3–HSD3 enzyme freshly purified was used for crystallization experiments using the hanging-drop vapor diffusion method at 4°C. Crystals were obtained by mixing equal volumes of h3–HSD3 concentrated to 14 mg/mL and of mother liquor (0.1 M NaCitrate at pH 6.5, 0.1 M NH₄Ac, and 24%–30% PEG4000). Crystal used for X-ray diffraction experiments was directly soaked into the cryopreserving solution (20% of ethylene glycol added to the mother liquor) and flash-cooled at 100 K in a nitrogen gas stream.

Structure determination and refinement
X-ray diffraction oscillation images of h3–HSD3/NADPH binary complex were recorded on an R-AXIS IIc image-plate detector mounted on a Rigaku RU-200 rotating-anode generator equipped with focusing mirrors (Rigaku MSC). Data were integrated and scaled using the December 2000 version of the XDS program (Kabsch 1993) and the crystal structure was directly determined by performing a rigid body refinement using the Refmac program (Murshudov et al. 1997). The h3–HSD3 structure (PDB code 1J96 [PDB] ; Nahoum et al. 2001) was used as a model and molecular replacement solution was calculated with two molecules per asymmetric unit. Crystallographic statistics are shown in Table 1. The refinement procedure was performed using CNS (Brunger et al. 1998) and Refmac (Murshudov et al. 1997) programs. The initial model issued from rigid body refinement was submitted to a cycle of simulated annealing at 3000 K followed by energy minimization cycles. After electronic density maps calculations and manual rebuilding using O (Jones et al. 1991), NADPH was added to the model. Atomic models were then refined by simple energy minimization followed by isotropic B-factors refinement (restrained, individual B-factor refinement) and corrected by manual rebuilding. Missing parts of the model, sulfate ion, -mercaptoethanol, ethylene glycol, citric acid, acetate, and water molecules were progressively added during the refinement procedure. Finally, the quality of the model was verified with PROCHECK (Table 1) (Laskowski et al. 1993).

The substrate cavity volumes of h3–HSD3 in the presence of its different ligands were measured with the program VOIDOO (Kleywegt 1994) using program default parameters. The volume values obtained this way were finally corrected using the equation given by Chakravarty et al. (2002).

Accession numbers
Atomic coordinates of this binary complex have been deposited in the Protein Data Bank with the accession code 1XJB.

	Footnotes

 
¹ Present addresses: Department of Biological Chemistry, University of Michigan, 5416 Medical Science Building 1, 1301 Catherine Street, Ann Arbor, MI 48109-0606, USA;

² Synchrotron Soleil, L’Orme des Merisiers, Saint-Aubin BP 48, 91192 Gif-sur-Yvette CEDEX, France.

	Acknowledgments

 
This work was supported by the Medical Research Council (MRC) of Canada and Endorecherche Inc. Jean-François Couture and Pierre-Luc Côté are the recipients, respectively, of a doctoral scholarship provided by the Laval University Foundation, Hydro-Québec and a master’s degree scholarship provided by the Natural Sciences and Engineering Research Council of Canada. The authors thank Sylvie Méthot for careful reading of the manuscript.

	References

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