extfieldsofstudy: Chemistry Medicine

s2fieldsofstudy: Chemistry Biology

text:
Activation mechanism of the CO sensor CooA. Mutational and resonance Raman spectroscopic studies.

CooA is a CO-dependent heme protein transcription factor of the bacterium Rhodospirillum rubrum. CO binding to its heme causes CooA to bind DNA and activate expression of genes for CO metabolism. To understand the nature of CO activation, several CooA mutational variants have been studied by resonance Raman spectroscopy, in vivo activity measurements, and DNA binding assays. Analysis of the Fe-C and C-O stretching Raman spectroscopy bands permits the conclusion that when CO displaces the Pro2 heme ligand, the protein forms a hydrophobic pocket in which the C-helix residues Gly117, Leu116, and Ile113 are close to the bound CO. The displaced Pro2 terminus is expelled from this pocket, unless the pH is raised above the pKa, in which case the terminus remains in H-bond contact. The pKa for this transition is 8.6, two units below that of aqueous proline, reflecting the hydrophobic nature of the pocket. The proximal Fe-His bond in Fe[II]CooA is as strong as it is in myoglobin but is weakened by CO binding, an effect attributable to loss of an H-bond from the proximal His77 ligand to the adjacent Asn42 side chain. A structural model is proposed for the position of the CO-bound heme in the active form of CooA, which has implications for the mechanism of CO activation.

CooA is a member of the emerging class of heme sensor proteins that modulate biological activity in response to fluctuations in the concentration of the gaseous molecules CO, NO, or O 2 (1,2). Binding of these diatomic ligands to the heme induces the biological response. The mechanisms whereby the protein responds to the binding event are of great current interest.
CooA is isolated from the bacterium Rhodospirillum rubrum, which grows anaerobically on CO as sole energy source (3). CooA regulates the expression of genes, termed coo for CO oxidation, whose products are associated with CO metabolism. The protein is a homodimer (222 amino acids per monomer) containing two b-type hemes that reversibly bind CO. The CO binding enables the protein to bind a specific DNA sequence and induce transcription of the coo genes. CooA contains DNAbinding and signal domains analogous to those found in the catabolite gene activator protein CAP 1 (or CRP) (1, 2) (Fig. 1). The heme is bound within the signal domain, where it is ligated by a histidine (His 77 ) side chain (although this ligand is replaced by the Cys 75 side chain when the Fe[II] is oxidized to Fe[III] (4)). The sixth Fe [II] ligand is unprecedented; it is the N-terminal proline residue (Pro 2 ) from the opposite chain, which is displaced by CO binding (5).
The structure of CO-bound CooA is unknown, but on the basis of the CAP structure (6) it is reasonable to assume that the DNA-binding domains are brought into proper position for DNA binding by movement of the C-helix, which runs the length of the protein and passes close to the heme (1,2,7). It has been shown that a repositioning of the C helices at the dimer interface is a major communication pathway between the hemes and the DNA-binding domains (8), but the basis for this repositioning upon CO binding remains unclear. To elucidate the nature of the CO-bound form of CooA and the mechanism of its CO activation, we have prepared a series of CooA variants by site-directed mutagenesis and examined in vivo activity, specific DNA binding, and the Fe-CO and C-O stretching resonance Raman (RR) bands. The residues chosen for mutagenesis are in the region of the heme groups, as defined by the Fe[II]CooA structure. Fig. 2 gives a close up view of this region.
The data permit us to identify the approximate position of the heme-CO in CooA and propose a specific model for how that positioning affects CooA activation and leads to the C-helix repositioning. An important aspect of the active form involves the breakage of a critical interaction that stabilizes the inactive structure: an H-bond from the His 77 proximal ligand to the side chain of Asn 42 .
These structural inferences also provide a basis for understanding the cooperative binding and complex kinetics of CooA. 2 Finally, we propose a structural model for the heme movement that is consistent with the data and will serve as a working hypothesis for future work.

EXPERIMENTAL PROCEDURES
Strains, Plasmids, and in Vivo Assays-The construction of strains overexpressing WT CooA and CooA variants in an Escherichia coli background having a ␤-galactosidase reporter system in the chromosome was described previously (8), and in vivo activities were quanti-tated using the standard protocol (10). All the site-directed and regionrandomized CooA mutations were constructed in a pEXT20-based expression plasmid, which provides tight control of CooA expression (11).
CooA Purification-The purification of WT CooA and most CooA variants was performed with our standard method as described previously (12). Some CooA variants such as C75S, L116K, and M124R CooA were purified with modified methods, which are also described elsewhere (12)(13)(14). The purity of WT CooA and CooA variants was estimated to be Ͼ95% based on SDS-PAGE (in the case of L116K CooA, ϳ90%). The heme content of CooA preparations was estimated using the extinction coefficient of Fe[III] WT CooA (3) or by a modified reduced pyridine-hemochromogen method (3), and protein concentration was measured using the BCA assay (Pierce).
In Vitro DNA Binding Assays-In vitro DNA binding assays of WT CooA and CooA variants were performed using the method described elsewhere (11). As a fluorescence probe, a 26-bp target DNA containing PcooF was labeled with Texas Red on one end of the duplex and used at the concentration of 6.4 nM. Salmon sperm DNA was used as the nonspecific DNA competitor. Dissociation constants (K d ) were calculated by fitting of the binding data to a nonlinear equation with correction of the fluorescence quenching as described elsewhere (15).
Sample Preparation-Purified CooA was diluted in buffer (25 mM Mops/0.1 M NaCl, pH 7.4) to a heme concentration of ϳ20 M. Other pH values were investigated using 0.1 M Tris, pH 7-9, 0.1 M glycine, pH 9.5-10, and ϳ0.1 M NaOH, pH [11][12]. CooA samples were purged with N 2 for 20 min and then with CO for ϳ3 min, followed by reduction of the Fe[III] to the Fe[II] form with sodium dithionite (final concentration ϳ63 mM). Reduction and CO binding were monitored by changes in the absorption spectra.
RR Spectroscopy-RR spectra were obtained with excitation wavelengths of 406.7 and 568.1 nm from a Kr ϩ laser (2080-RS; Spectra Physics) and 441.7 nm from a He-Cd (Liconix) laser in a 270°backscattering sample geometry. Photodissociation of the bound CO was minimized by using low laser power (ϳ5 milliwatts at the sample) and by focusing it with a cylindrical lens onto a spinning sample. The scattered light was collected and focused onto a single spectrograph (Chromex) equipped with a notch filter (Kaiser Optical) and a CCD detector (Roper Scientific) operating at 77 K. Spectra were calibrated with dimethyl formamide and dimethyl sulfoxide-d 6 .

RESULTS
The goal of this study has been to understand the nature of the heme environment of CO-bound CooA and use this information to better understand the mechanism of CooA activation. RR and activity data are gathered and compared in Table  I. We note at the outset that spectroscopic probes, which report on local structural features, are not expected to track the degree of activity, which measures the biochemical competency of the global protein conformation. It is the interplay between the two sets of data that inform our thinking about activation mechanism.
The results of this study are organized in the following manner. 1) There are actually two different forms of CooA-CO, with different C-O stretching frequencies, as revealed by reanalysis of RR spectra. This observation has an effect on the interpretation of RR data in this manuscript and resolves a discrepancy in RR data reported previously. 2) Backbonding analysis of the RR data reveals that the Fe-His proximal bond is weakened in the major form of CooA-CO. Mutational data point to breaking of the His 77 -Asn 42 H-bond as the cause of this weakening and is consistent with a significant role for this process in CooA activation. 3) The bound CO is certainly in close proximity to Ile 113 , Leu 116 , and Gly 117 , which has implications for the movement of the heme and the protein upon CO binding. This proximity was revealed by the RR analysis of mutational variants substituted at these positions that show striking perturbations and is generally consistent with functional characterization of these variants. 4) RR analysis of pH effects of WT CooA, and of variants containing deletions at the N terminus that includes Pro 2 , suggest that CO displacement of Pro 2 leads to its expulsion from the hydrophobic CO binding pocket, because of its positive charge when protonated. The previously reported depression of the Fe[II]CooA 11 heme RR band is found, via the effects of variants, to stem from the strong ligand field of the Pro 2 ligand.
Structurally Distinct Populations of CO-bound CooA-We have previously reported RR spectra of CooA-CO showing a Fe-CO stretching band (FeC) at 487 cm Ϫ1 and an unusually high CO stretching band (CO) at 1982 cm Ϫ1 (16). In contrast, Uchida et al. (17) reported CO ϭ 1969 cm Ϫ1 for their CooA-CO preparation although they reported the same FeC value as ours, 487 cm Ϫ1 . To resolve this discrepancy we reexamined the CO and FeC regions of the RR spectrum at high amplification ( Fig. 3), using 13 CO shifts to identify the bands definitively. FeC was measured at 487 cm Ϫ1 , as before, and we also reproduced the 1982-cm Ϫ1 CO band. However the newly measured CO band has a pronounced shoulder at 1962 cm Ϫ1 . Upon 13 CO substitution, this 1962-cm Ϫ1 band shifted appropriately, indicating a second CO band.
The 487-cm Ϫ1 FeC band is a single band. Deconvolution from the 465-cm Ϫ1 band of excess dithionite (Fig. 3) yields a bandwidth, 14.5 cm Ϫ1 , which is less than that of the CO adduct of myoglobin (data not shown), 18 cm Ϫ1 , ruling out multiple FeC contributions in CooA-CO. Therefore, we conclude that the molecules having CO ϭ 1982 and 1962 cm Ϫ1 both have the same FeC ϭ 487 cm Ϫ1 .
The CO reported by Uchida et al. (17), 1969 cm Ϫ1 , is closer to the lower of the two values we observe, but their CooA-CO preparation also had a fraction with CO ϭ 1979 cm Ϫ1 ; this is the frequency reported in picosecond FTIR experiments (18) for CooA-CO molecules undergoing photolysis. Thus, CooA-CO has two populations of molecules, having distinct CO values (1962 and 1982 cm Ϫ1 in our hands), but the same FeC value, 487 cm Ϫ1 . The relative size of the populations appears to be preparation-dependent. In our preparation the 1962-cm Ϫ1 fraction is small; the species is labeled CooA-COЈ. We suggest that the species represented by 1982 cm Ϫ1 is the active form and that CooA-COЈ is a form that has not rearranged to the active, DNA binding structure. There may be an equilibrium between active and inactive forms, or else a fraction of the molecules are stuck in the inactive configuration because of an energy barrier of some type.
Conformational Change of CooA upon CO Binding Weakens the Fe-His 77 Bond by Breaking the His 77 -Asn 42 H-bond-The anomalous CO of the major form of CooA-CO, 1982 cm Ϫ1 , is produced by a weakened Fe-His 77 bond. This can be seen in a plot of FeC against CO (Fig. 4). Because of backbonding, FeC and CO are negatively correlated in heme proteins (19,20), as illustrated for a set of Mb variants in Fig. 4 (solid line). Changes in the polarity of the binding pocket augment or diminish backbonding and shift the points along the line (19,20). In addition, the correlation is displaced upward when the Fe-proximal ligand bond is weakened, because of diminished bonding competition between the proximal ligand and the CO (19,21). The upper dashed line in Fig. 4 represents data for Fe[II] porphyrin CO adducts without any trans ligand (21).
When the CooA-CO values are plotted on the graph, the minority form, CooA-COЈ, is seen to fall near Mb variants with hydrophobic pockets, in which the distal histidine is replaced by hydrophobic residues (cluster of points marked ϪH64 in Fig.  4). However, the distinctly higher CO of the dominant form, CooA-CO, places it above the Mb line by 7 cm Ϫ1 , implying a weakened bond from the Fe to the proximal ligand, His 77 in CooA (see Fig. 2).  This displacement from the Mb backbonding line was missed in our earlier correlation (16), because a more varied group of heme proteins was used, having greater scatter of the points. The high CO was instead attributed to negative polarity in the CO binding pocket (16). However, examination of the pocket region, defined by the mutagenesis results to be described below, fails to reveal any candidate residue with a negative charge or even a lone pair of electrons that could provide a negative dipole. In principle, peptide dipoles (22) could interact with the bound CO, but this is unlikely, because the residues that form the binding pocket are part of a helix.
The inference of a weakened Fe-His 77 bond in CooA-CO is strongly supported by the picosecond RR study by Uchida et al. (23), which identified the Fe-His stretching band at 211 cm Ϫ1 in the immediate photoproduct of CooA-CO. This band, a direct measure of the Fe-His bond strength, is enhanced in five coordinate Fe[II] hemes, and is found at 220 cm Ϫ1 in deoxyMb (24). The distinctly lower value found by Uchida et al. (23) shows that the Fe-His 77 bond is weaker than in Mb for the immediate photoproduct, consistent with a weaker bond in the CO adduct itself. The minority form, CooA-COЈ, which lies on the Mb backbonding line (Fig. 4), does not have a weakened Fe-His 77 bond and is expected to have an Mb-like photoproduct; however, its Fe-His would escape detection because of the low population.
How strong is the Fe-His 77 bond in Fe[II]CooA itself? Because Fe[II]CooA is six-coordinate, the Fe-His vibration is not enhanced (25), and its frequency cannot be measured. However, we took advantage of a significant five-coordinate fraction in the G117I variant of Fe[II]CooA (7), in which the bulky Ile side chain causes partial dissociation of the Pro 2 ligand (see below). The five-coordinate heme Soret absorption band is redshifted from the six-coordinate band, allowing us to use laser wavelength tuning (442 nm) to enhance the five-coordinate RR signals (Fig. 5). Selection of the five-coordinate species is confirmed by the position of the strong 4 band at 1355 cm Ϫ1 , the standard five-coordinate frequency (25). The Fe-His band is seen at 220 cm Ϫ1 , the same frequency as in deoxyMb. Thus the Fe-His bond is as strong in G117I variant of Fe[II]CooA as in Mb. We infer that the Fe-His bond is similarly strong in WT Fe[II]CooA (there is no reason that steric displacement of Pro 2 should make the proximal Fe-His connection stronger) and that it weakens significantly in the major form of CooA-CO, as well as its immediate photoproduct. At later times, the photoproduct Fe-His position should increase to 220 cm Ϫ1 as the protein relaxes to the Fe[II]CooA structure, but this process cannot be followed in photolysis experiments because of rapid geminate rebinding of CO (18,23).
These results establish that CO binding weakens the Fe-His 77 bond. We therefore asked what could be the mechanism responsible for this property of CooA. It has been known that H-bond donation increases the anionic character of the imidazole ring and strengthens the Fe-His bond (26). For example, the Fe-His frequency is much higher, 240 cm Ϫ1 , in five-coordinate Fe[II] forms of the peroxidases that have a negative Asp side chain in position to accept the proximal His H-bond (24). In Mb the proximal His is H-bonded to a neutral OH group of a Ser residue, as well as to a backbone carbonyl (Fig. 4, inset).
In the Fe[II]CooA structure (Fig. 2), the H-bond acceptor is the amide side chain carbonyl of Asn 42 (Fig. 2) (21). The cluster of Mb points in the middle of the diagram (ϪH64) is from variants in which the distal histidine is replaced with hydrophobic residues. The point for the minor WT CooA-CO (X) lies in the cluster, whereas the major WT CooA-CO and the CO-bound form of L116K CooA variant, as well as the high pH form of WT CooA-CO, lie between the two lines, indicating that the Fe-His bond is weaker in CooA-CO than in MbCO.
With regard to the minor CooA-COЈ form, we note that the low frequency shoulder on the N42A CO band is attenuated relative to the wild-type 1962-cm Ϫ1 shoulder. Curve resolution gives a higher frequency, 1966 cm Ϫ1 for the N42A shoulder, suggesting a weakened H-bond in the N42A CooA-COЈ structure.
Although Asn 42 interacts with His 77 in the Fe[II] form and not the CO-bound form, the nature of the residue is of some importance to activation. N42A, N42D, and N42K CooA all displayed poorer DNA affinity in the presence of CO (and poorer ␤-galactosidase activity) than did WT CooA (Table I), suggesting some additional role for this residue in the active form. The basis for that effect is unknown. Note that the activity assays that measure DNA affinity are somewhat easier to interpret than the in vivo assays reporting ␤-galactosidase activity, because the former only measures DNA binding, whereas the latter measures interaction with RNA polymerase and has other complications (8).
These results indicate that an element of the activation mechanism is the perturbation of the His-Fe bond. The data FIG. 5. RR spectrum of G117I CooA variant in the Fe[II] form, obtained with 442-nm excitation to enhance the five-coordinate high spin fraction, whose Soret band is at ϳ440 nm (7). The 1355-cm Ϫ1 4 band position is diagnostic for five-coordinate high spin heme (23), and a strong Fe-His stretching band is seen at 220 cm Ϫ1 . The spectrum closely resembles that of deoxyMb (21). and arguments below strongly suggest that the His-Fe bond is perturbed by displacement of the heme upon CO binding, which is critical for CooA activation.
Hydrophobic C-helix Residues, but Not Pro 2 , Form the CO Pocket of CooA That Is Necessary for Activation-The distal heme pocket has an important role in ligand discrimination in many heme proteins (27). CooA activation is specific to CO, and an understanding of the CO binding pocket should be valuable in understanding both specificity and the activation in response to CO. We have already shown that hydrophobic residues at positions 113 and 116 are critical for normal CO activation (28) and that Gly 117 is also necessary for this response (7). However, the positioning of these residues with respect to the bound CO was not investigated. We therefore measured the RR spectra of a variety of CooA variants that were either novel in terms of activity and might therefore be altered in their sensing of the bound CO (Table I) Table I, neither replacement of the Pro 2 terminus by tyrosine (P2Y CooA) nor deletion of the two penultimate residues (⌬P3R4 CooA) showed noticeable effect on the FeC or CO band. This implies that Pro 2 in WT CooA does not form the CO pocket and is consistent with the previous observation that Pro 2 is not essential for activation of CooA by CO (11). We note, however, that the DNA binding affinities of these two Pro 2 variants are 5-10-fold poorer than WT CooA, suggesting a minor role for the displaced Pro 2 in the active form of CooA. Properties of the Pro 2 variants are further considered below.
Because Pro 2 is not near the bound CO, then the structure of Fe[II]CooA suggests that the most probable candidates for CO pocket residues are on the C-helix, with Ile 113 , Leu 116 , and Gly 117 being particularly attractive (1,28). The RR analysis of CooA variants altered at these positions establishes significant influences on FeC and CO consistent with a positioning of these residues close to the bound CO (see Fig. 7 and Table I). When these residues are replaced by H-bond donors, histidine and lysine FeC shifts up, and CO shifts down from the position in WT CooA, as expected for a positive polar interaction. 3 The effect is particularly marked for the Gly 117 position. G117H CooA displayed an 11-cm Ϫ1 upshift of FeC and an 18-cm Ϫ1 downshift of CO. These shifts are opposite in sign and comparable in magnitude to those observed when the distal histidine of Mb is replaced by hydrophobic residues (19,20). G117S CooA also showed a sizeable, though more modest effect on the RR spectrum, shifting FeC up by 7 cm Ϫ1 and CO down 12 cm Ϫ1 . These results indicate that the newly introduced His 117 or Ser 117 (albeit to a smaller extent) lie sufficiently close to the CO to exert a positive potential, via H-bond donation. However, the introduced His 117 in G117H CooA is not well ordered and exerts a heterogeneous effect on the CO as evidenced by broad CO and FeC bands (Fig. 7). Consistent with the polarity interpretation, introduction of steric hindrance at 117 position (G117I CooA) had very little effect on FeC and CO (Table I), because the CO environment remains non-polar. The importance of Gly 117 to CooA activation has already been described (7) and is reflected in the dramatically reduced ac-tivity of Gly 117 variants seen in Table I. Because Gly 117 is critical for activity, and its substitution perturbs the RR spectra, it would be tempting to conclude that its role is to contact the bound CO. Although this hypothesis cannot be discounted, it is also possible that Gly 117 is an important determinant of the C-helix structure per se and simply happens to be near the bound CO.
At position 113, the effect of mutation on RR spectra is smaller, as FeC shifts up only 3 cm Ϫ1 and CO down 11 cm Ϫ1 in I113H CooA (see Fig. 7 and Table I). Interestingly, essentially the same effect was seen in I113A and I113D CooA. A possible interpretation is that in these cases water molecules may be brought into contact with the CO, filling the cavity that results from substituting the smaller Ala residue, or attracted to the negatively charged Asp side chain. These results suggest that Ile 113 is close to the bound CO but not as close as Gly 117 . In the case of I113Y CooA, the RR spectrum gives a slight indication of negative polarity, shifting FeC down 2 cm Ϫ1 and CO up 3 cm Ϫ1 . This effect could result from proximity to the electron cloud of the Tyr ring to the CO. Substitutions at position 113 typically have modest effects on activity, as reported previously (28) and shown in Table I. The exception is I113D, and we have already shown that hydrophilic residues at positions 113 and 116 severely perturb activation (28). It is our working hypothesis that the effect is because the hydrophobic pocket is important for eliciting the proper conformational change in the C-helices upon displacement of the Pro 2 ligand by CO.
At position 116, Lys and Phe substitutions were tested. Other possible H-bond donors were not examined, because they 3 An alternative interpretation is that these donors ligate the iron in place of Pro 2 and that CO displaces His 77 instead. However, there is no reason to suppose that distal replacements would labilize the Fe-His 77 bond. When His 77 itself is replaced by Tyr, the FeC/CO values (Table  I) are significantly different, and there is a five-coordinate CO-bound fraction, implying displacement of both ligands (16). have been shown to accumulate heme-containing CooA very poorly, presumably because of disruption of the hydrophobic pocket (28). L116K CooA variant was chosen, because it showed very high CO-independent DNA binding activity (comparable with that of wild-type CooA in the presence of CO) and diminished CO-dependent DNA binding activities (see Ref. 14 and Table I). As shown in Fig. 7, the introduction of Lys at position 116 exerted an almost identical effect on the RR spectrum as did G117H, shifting FeC by 13 cm Ϫ1 up and CO by 18 cm Ϫ1 down (Table I). This result is consistent with the hypothesis that the L116K substitution, like G117H, introduces positive polarity into what is otherwise a non-polar environment around the CO. This CooA variant also showed broad CO and FeC bands. CooA-CO activity is impaired by the L116K substitution but is still considerable, indicating a modest perturbation of the active structure. The high activity of L116K CooA in the absence of CO has already been analyzed, and the result strongly implies that the substituted Lys residue serves as a ligand in both the Fe[III] and Fe[II] forms. We have suggested that this ligation leads to a similar helix repositioning as that seen in WT CooA when bound to CO (14). The absence of significant FeC or CO shifts in L116F is again consistent with maintenance of a hydrophobic environment for the bound CO.
Importantly, in the G117H and L116K CooA variants, the FeC/CO point shifts up and to the left in the backbonding plot (Fig. 4, only L116K is shown, for clarity) but remains displaced from the Mb line by the same amount as WT CooA-CO. This behavior indicates that the Fe-His 77 bond is unaffected by the distal residue replacement, although backbonding is enhanced by the distal polarity. CO binding weakens the Fe-His 77 bond in these variants, just as in WT CooA-CO.
It should be noted that these observations apply only to the major population of CooA-CO molecules as described at the beginning of the "Results." It is impossible to tell how the substitutions affect the minor CooA-COЈ fraction because of its weak signal and the breadth of the perturbed bands. However, a non-polar binding pocket is also indicated for the minor WT CooA-CO by its position in the FeC/CO correlation (Fig. 4), close to those of Mb variants with non-polar pockets.
In summary, the RR data establish that Gly 117 , Leu 116 , and Ile 113 are all in or near the CO binding pocket. However, the data also indicate that substitutions of the Ile 113 and Leu 116 side chains are much more permissive with respect to CO activation of CooA than are those at Gly 117 . A possible interpretation is that the CO directly contacts the Gly 117 and that any steric hindrance destabilizes the DNA binding conformation of the protein. However, it is also possible that Gly 117 induces a critical bend in the C-helix, which is disrupted by substitutions.
We also examined other candidate residues for their effect on RR spectra. These included C-helix variants M124R and W110L and B-helix variants E99L and I95W (the B-helix is the short helix on the "bottom" of CooA as shown in Fig. 1). M124R (13) was chosen because of its significant impact on CO-sensing function of CooA. The others were chosen because of their potential proximity to the heme iron. As shown in Table I, all these variants except M124R had little effect on RR spectra, suggesting that these residues are not near the bound CO and consistent with the pocket being formed primarily by Gly 117 , Leu 116 , and Ile 113 .
The M124R replacement has little effect on the CooA-CO activity (Table I) but produces a double peaked CO band; one component is at the WT CooA position, 1982 cm Ϫ1 , whereas the other is ϳ17 cm Ϫ1 lower. The FeC band is lowered slightly and is broadened (see Table I and Fig. 7). Although Met 124 is fur-ther from the heme iron than are Gly 117 , Leu 116 , and Ile 113 , it is possible that the long Arg side chain reaches the vicinity of the bound CO in a fraction of the CooA-CO molecules, producing the downshifted CO component. Modeling of the introduced arginine in Fe[II]CooA (13) suggested two favored conformations of the side chain, the guanidine cation interacting either with the seven-propionate of the heme or with the backbone carbonyl of Ser 78 . It is possible that in the M124R-CO structure one of the Arg conformers interacts instead with the CO.
The Strongly Basic Pro 2 Ligand Is Expelled from the Hydrophobic CO Pocket in the CO-bound Form of CooA at Physiological pH-Extensive mutational studies have shown that Pro 2 is not critical for the response of CooA to CO (11). Nevertheless, the Pro 2 ligand is a central aspect of cooperativity in CO binding by CooA. 2 Because the global conformational change of CooA upon CO binding is initiated by the displacement of the Pro 2 ligand by CO, it would be informative to trace the fate of the displaced Pro 2 . We have shown above that Pro 2 does not appear to be near the bound CO at normal pH, but when the solution pH was increased above 7, the FeC and CO of WT CooA-CO were affected significantly (Fig. 8). The FeC band at 487 cm Ϫ1 was replaced by one at 497 cm Ϫ1 whereas the CO band at 1982 cm Ϫ1 was replaced by one at 1964 cm Ϫ1 . Although the latter frequency is close to that of CooA-COЈ, the FeC frequency is distinctly higher than in CooA-COЈ. Thus the change at high pH is not simply an increase in the CooA-COЈ fraction, but instead a new species is formed in basic solution.
The high pH FeC and CO values are similar to those displayed by the L116K and G117H variants (see Fig. 7 and Table I). The FeC/CO point remains well above the MbCO backbonding correlation (Fig. 4), indicating little change in the Fe-His 77 bond. We infer that the effect of raising the pH is to bring a H-bond donor into proximity with the bound CO. As in L116K and G117H, the high pH CO and FeC bands are broadened, suggesting heterogeneity in the H-bond donor position.
What might this H-bond donor be? Key evidence comes from the ⌬P3R4 CooA variant, whose FeC and CO bands are unperturbed when the pH is raised, up to 11 (Fig. 8). We conclude that the H-bond donor is the N-terminal Pro 2 , interacting with the bound CO via its secondary amine NH group. When the two penultimate residues, Pro 3 and Arg 4 , are removed, the N terminus is constrained from interacting with the CO.
The Pro 2 interaction in WT CooA requires pH elevation, presumably because the secondary amine becomes protonated when it is displaced from the Fe[II] at pH 7. Because the CO binding pocket is hydrophobic, the protonated N terminus would be expelled from the binding site, because it is positively charged, just as the distal histidine is expelled from the MbCO binding pocket upon protonation at low pH (29,30). At high pH, the displaced Pro 2 , being neutral, can remain in contact with the CO and enhance backbonding, just as the distal histidine does in Mb.
What is the pK a of the displaced Pro 2 ? In aqueous solution the pK a of the Pro amine is 10.5, but it should be lowered in CooA-CO because of the energy cost of expelling Pro 2 from the hydrophobic pocket. In Mb, the pK a of the distal His is 4.3 (29,30), nearly three units lower than the aqueous His pK a . We examined the pH dependence of WT CooA-CO by decomposing the FeC band into 487-and 497-cm Ϫ1 components and plotting their intensities against the solution pH (Fig. 9). The resulting titration curves yield a pK a of 8.6, two units lower than the aqueous pK a of proline. If the assignment of the pK a to Pro 2 is correct, then the energy cost for expelling the distal H-bond donor is about two-thirds as high in CooA as it is in Mb.
We note that the absorption spectrum is not sensitive to the pK a 8.6 process. The Soret band position, which is 422 nm at pH 7, only shifts significantly when the pH is raised to values greater than 10; at pH 12 the band maximum is 419 nm. We found that the RR spectral changes could not be fully reversed for solutions whose pH was raised to 10 or higher, indicating some irreversible change in the protein structure. However the Soret band shift could be reversed, even from pH 12, showing that there is a different physical mechanism for perturbation of the absorption spectrum.
These results indicate that Pro 2 does not lie near the bound CO under physiological conditions, and the role of Pro 2 in the active form of CooA is uncertain. The DNA affinity of the CO-bound forms of P2Y and ⌬P3R4 CooA is 5-to 10-fold poorer than that of WT CooA, implying some small perturbation of the active structure when the N terminus is altered. However the primary roles of Pro 2 appear to be to maintain the inactive form of CooA in the absence of CO (8) and to provide a mechanism for the cooperative binding of CO when it is present. 2 The ligating nitrogen atom of Pro 2 is part of a strongly basic secondary amine and is expected to be a strong donor to the Fe[II]. Evidence for strong donation is seen in the RR spectrum of Fe[II]CooA, which reveals a depressed value, 1532 cm Ϫ1 , for the 11 porphyrin ring stretching vibration (25) (Fig. 10, 568-nm excitation was used to enhance 11 via resonance with the heme Q bands (25)). This mode is known to be sensitive to the strength of the axial ligand field in low spin Fe[II] porphyrin complexes. For example, the band shifts from 1547 to 1533 cm Ϫ1 when methionine is replaced by lysine in Fe[II] cytochrome c (31) and from 1539 to 1527 cm Ϫ1 upon deprotonation of the imidazole complexes of microperoxidase or of bisimidazole Fe[II] protoporphyrin (32,33). The sensitivity to imidazole deprotonation was the basis of our earlier suggestion (16) that His 77 might be deprotonated or strongly H-bonded in Fe[II]CooA. However, His 77 is now revealed by the crystal structure to have a normal H-bond with Asn 42 , and the position of 11 is unaffected when Asn 42 is replaced by the non-H-bonding Ala (Fig. 10).
We conclude that the 11 depression is unrelated to His 77 but instead reflects the strong ligand field of Pro 2 . This interpretation is supported by the observation (Fig. 10) that 11 shifts up, by 4 cm Ϫ1 , in the P2Y and ⌬P3R4 variants. When Pro 2 is replaced by Tyr, the secondary amine is replaced by a primary amine at the terminus, producing a somewhat weaker ligand field. In the ⌬P3R4 variant, the penultimate pair of residues is deleted, shortening the N terminus. The Fe[II] protein remains FIG. 8. pH dependence of the FeC and CO bands for the CO adducts of WT CooA and of the ⌬P3R4 CooA variant. The shift in FeC for WT CooA is also seen in the position of the ϳ1860-cm Ϫ1 band assigned to a combination band with 4 (1371 cm Ϫ1 , the strongest Soret-excited RR band). Another band, at 2049 cm Ϫ1 , is because of a combination of 4 with 7 (677 cm Ϫ1 , another strong fundamental), which is unaffected by pH (and whose intensity indicates the weakness of resonance enhancement for the CO band). The asterisks mark the excess dithionite band.
FIG. 9. pH titration curves of WT CooA-CO. For the analysis, FeC band envelopes of RR spectra were deconvoluted into 487-and 497-cm Ϫ1 components and then the fractional intensity of each band was plotted against the solution pH. The solid curves represent best fits to single proton release and uptake processes having a pK a of 8.6. mainly six-coordinate, but a small fraction of the heme is fivecoordinate, as judged by a shoulder on the 4 porphyrin band at the expected five-coordinate position, 1355 cm Ϫ1 (25). Pro 2 might remain the ligand in ⌬P3R4, but the shortening of the polypeptide chain would strain the Pro 2 -Fe bond, again weakening the field. This interpretation is consistent with the very rapid binding of CO to the ⌬P3R4 variant. 2

DISCUSSION
The objective of this study was to gain insight into the nature of the CO-bound form of CooA and its implications for the conformational change that allows CooA-CO to bind its DNA target. RR signals arising from the heme and its ligands were examined for selected variants, whose activity in vivo and in a DNA binding assay were determined. It is not expected that spectroscopic and activity perturbations should be correlated. On the contrary, it is an important feature of this study that quite different aspects of the CooA molecule are sensed by the spectroscopic and activity probes. Full activity assures that the protein has the correct conformation to bind its DNA target; alterations in many parts of the molecule can destabilize the correct conformation. The RR signals are sensitive to the structure of the heme and its ligands. It is this information that we seek to integrate into a model of how the binding of CO at the heme initiates the conformation change from inactive to active protein.
Two principal findings are central to our model. 1) The Chelix residues Ile 113 , Gly 117 , and Leu 116 become part of the CO binding pocket in CooA-CO. This is revealed by the spectral perturbations observed when their side chains are replaced with H-bond donors. This result implies repositioning of the hemes relative to the C-helices, because all three residues are at substantial distances from the heme iron in the Fe[II]CooA crystal structure (1).
Measured Fe-C ␣ distances are 9.4 (10.2) Å for Leu 116 , 13.0 (7.0) Å for Gly 117 , and 11.2 (9.3) Å for Ile 113 . The first number refers to the residue on the same subunit as the heme, whereas the second number refers to the residue on the opposite subunit. Although not required crystallographically this part of the structure is nearly 2-fold symmetric; the Fe-C ␣ distances from the second heme are within 0.1 Å of those from the first. It is notable that only in the case of Leu 116 does the closer residue reside on the same subunit as the heme. For both Ile 113 and Gly 117 , the residue from the opposite subunit is much closer than is the residue from the same subunit. Thus a simultane-ous rearrangement of both hemes and both C-helices is suggested to bring all three residues close enough to allow H-bond donor replacements to interact with the bound CO. 2) The His 77 -Asn 42 H-bond, which is evident in the Fe[II]CooA crystal structure, appears to be broken upon CO binding. The strongest evidence is that the CO and FeC RR bands are unaffected when Asn 42 is replaced by the non-H-bonding residue Ala (or by any other residue), despite the known sensitivity of these bands to the H-bond status of the proximal histidine ligand in heme-CO adducts (19,20). Moreover, the position of the CO and FeC on the backbonding correlation reveal that the Fe-His bond is weaker in CooA-CO than in MbCO These are the findings that flow from the present study. Together with the Fe[II]CooA crystal structure, they lead to a model for the initial motions leading to activation of CooA. The proposed motions are shown as dashed arrows in Fig. 2.
Upon binding CO, the heme is suggested to slide upward, bringing the FeCO unit closer to the side chain of Leu 116 on the same subunit. As discussed above, the L116K variant is active in the absence of CO, suggesting that a Lys at this position induces the same heme displacement by displacing Pro 2 as the distal ligand. We modeled a Lys side chain in the orientation of the Leu 116 side chain and measured a 4.0-Å separation from the iron to the amine nitrogen. Thus a ϳ2.0-Å displacement of the heme group would be required to form a bond. This displacement provides an explanation for loss of the His 77 -Asn 42 H-bond, because His 77 remains bound to the heme and is displaced with it. It is likely that this heme displacement is also involved in the His 77 /Cys 75 ligand switch that is known to occur The hydrophobic pocket consists of the indicated residues, as well as Val 44 , Ser 78 , Leu 112 , and Ile 113 , which are either obscured in this view or have been omitted to show the heme ligands Cys 75 , His 77 , and Pro 2 . The irregularly shaped orange structure designates one 53-Å 3 cavity, which is proposed to expand to allow the heme to insert further into the hydrophobic pocket in the CooA Fe[III] and Fe[II]-CO states. Structure and cavity analyses were generated using the Swiss Pdb Viewer, version 3.7 program (9) (www.expasy.org/spdbv/); the figure was rendered using POV-Ray, version 3.1g software (www.povray.org/). upon oxidation to Fe[III]CooA (4), because the shifted heme position would accommodate Cys 75 ligation. Similar heme movement and ligand switching has been described for cytochrome cd 1 (35). Although WT Fe[III]CooA is not active, there are a number of mutations that induce activity in the Fe[III] form of the protein (13).
It is further suggested that when the heme is displaced upward, it is approached more closely by the C-helix from the second subunit, whose Gly 117 and Ile 113 residues help form the hydrophobic binding pocket for the CO. A modest motion of the second C-helix would permit H-bond donors at these positions to perturb the CO, whereas a much larger rearrangement of the first C-helix would be required to produce this effect. Thus the overall motion that initiates the conformation change to the active CooA-CO structure is viewed as a clockwise rotation of both hemes and both C-helices, in the plane of Fig. 2.
Some CooA molecules do not undergo this motion. The minority CooA-COЈ fraction represented by the 1962-cm Ϫ1 CO shoulder appears to have an intact His 77 -Asn 42 H-bond (the FeC/CO point falls on the Mb backbonding line), indicating that the heme is undisplaced from its position in the Fe[II]CooA structure. Presumably these molecules are not active in binding DNA. We do not know whether they are in a dynamic equilibrium with the active molecules or else are prevented from undergoing activation by some kind of energy barrier.
What is the driving force for the proposed heme and C-helix displacements? We propose that hydrophobic interactions are responsible (28), perhaps abetted by the electronic reorganization of the heme that results from a acceptor ligand (CO) replacing a donor ligand (the N terminus). Examination of the Fe[II]CooA crystal structure shows the heme to be partially exposed to solvent, whereas it would become more buried by sliding into the cavity (Fig. 11). Hydrophobic interactions would be enhanced by the suggested displacement of the opposite C-helix toward the heme. The hydrophobic character of the CO binding pocket is evidenced by the ϳtwo-unit pK a reduction of the displaced Pro 2 , relative to the expected aqueous pK a . CONCLUSIONS RR spectroscopy, combined with activity measurements, on variants of CooA has provided evidence about the nature of the CO adduct. Three C-helix residues, Ile 113 , Leu 116 , and Gly 117 , are shown to be close to the bound CO, forming a hydrophobic pocket. The displaced Pro 2 ligand is expelled from this pocket, unless the amine is unprotonated; the pK a for this process is 8.6. The RR evidence points strongly to breaking of the proximal His 77 -Asn 42 H-bond when CO binds. On the basis of the Fe[II]CooA crystal structure, these observations lead to the proposal that when CO binds, the heme is displaced into an adjacent cavity and is approached by the C-helix of the opposite subunit. These motions are suggested to trigger the protein conformation change required for DNA binding.


text:
Mimic Aldehydes and Inhibit Liver Alcohol Dehydrogenases and Ethanol Metabolism

Title Footnotes: * This work was supported by United States Public Health Service Grant AA00279. 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. The atomic coordinates and structure factors (code 1P1R) have been deposited in the Protein Data Bank, Research Collaboratory for Structural Bioinformatics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org).

Vertebrate alcohol dehydrogenases (EC 1.1.1.1, ADH) 1 are involved in the metabolism of relatively non-polar alcohols and aldehydes (1). The five different human enzymes have broad substrate specificity, oxidizing primary and secondary alcohols and reducing their corresponding carbonyl compounds. Specific inhibitors of these dehydrogenases would be useful for defining substrate specificity, for studying the physiological roles of the enzymes and for preventing the oxidation of methanol and ethylene glycol to toxic products (2)(3)(4)(5)(6).
Formamides and sulfoxides are analogues of the carbonyl substrates, are bound preferentially to the enzyme-NADH complex, and are potent uncompetitive inhibitors against varied concentrations of alcohol (7)(8)(9)(10)(11)(12)(13)(14)(15). Uncompetitive inhibitors could be especially useful for controlling alcohol metabolism since they are effective even when the concentrations of alcohol are saturating. Previous studies identified some potent uncompetitive inhibitors for human (14,15), horse (10,11,14,15), monkey (11), and rat (10,11) liver alcohol dehydrogenases. We now extend the studies with 14 new branched-chain and chiral formamides in order to explore active site topologies and to make more effective inhibitors for the mouse (MmADH1) and human (HsADH1C*2) enzymes. For comparison, we also studied the potency of five sulfoxides and two 4-substituted pyrazoles with purified MmADH1.
We also studied the inhibition by five compounds of ethanol metabolism in mice as a model animal. We used numerically integrated rate equations that describe the non-linear progress curves and provide good estimates of inhibition constants in vivo. The results provide information for optimizing the sizes of inhibitors for maximal efficacy.
We determined by x-ray crystallography the structure of the complex of horse liver ADH with NADH and a potent, chiral formamide inhibitor, which binds as an analogue of an aldehyde. The structure shows, for the first time in a ternary complex of a dehydrogenase with NADH, that the reduced nicotinamide ring is puckered, as expected for the transition state for hydrogen transfer. The structure also provides a basis for interpreting the structure-function relationships of the inhibitors.

EXPERIMENTAL PROCEDURES
Materials-Native crystalline horse liver alcohol dehydrogenase (EE isoenzyme), LiNAD ϩ and Na 2 NADH were purchased from Roche Applied Science. The plasmid for the expression of HsADH1C*2 was obtained from Dr. Jan-Olov Höög (Karolinska Institutet, Stockholm, Sweden) and that of MmADH1 from Dr. Gregg Duester (Burnham Institute, La Jolla, CA). Amines and other reagents were obtained from Lancaster and Aldrich Chemical Co. Some formamides and sulfoxides were prepared previously (11,14).
Synthesis of Formamides-The formamides were generally prepared (unless stated otherwise) by refluxing the amine with 88% HCOOH, removing excess HCOOH and H 2 O under reduced pressure, and distilling the product. N-1-Cyclohexylethyl and N-1-(4-methylphenyl)-ethyl formamides were solids and were recrystallized from ethyl acetate/ hexane or dichloromethane. Final yields were 50 -70%. Compounds were characterized using a Varian 500 MHz or a Bruker AC 300 NMR spectrometer. Multiple resonances are due to the cis-trans (Z,E) isomerism of the formamide group. The major geometrical isomer was cis (ϳ70 -80%). Mass spectra (MS or high resolution, HRMS) were obtained using a Micromass Autospec M spectrometer. The structure of each compound was confirmed by three different analytical methods.  13  N-1-Ethylhexylformamide-As for the first stage of the Leukart reaction, ammonium formate and 3-octanone (4:1 molar ratio) were heated (140 -150°C) until the mixture became homogeneous. Water and some 3-octanone distilled over. The ketone was returned to the reaction vessel, and the mixture was heated at 155-160°C for 4 h. The cooled reaction mixture was washed twice with water, and the product was distilled. 1  Protein Purification-Human alcohol dehydrogenase 1C*2 was prepared essentially as described (16). Mouse ADH1 was purified using the following procedure, which gave better yields than the general procedure suggested by Qiagen. Mouse ADH1 was expressed in Escherichia coli M15 (pREP4) from the full-length cDNA linked to a His 6 tag in the pQE-30 plasmid (Qiagen, Valencia, CA). The E. coli cells were grown at 37°C on plates and in starter cultures with LB media containing 100 g/ml of ampicillin and 25 g/ml of kanamycin and used to inoculate 9-liter cultures of LB without antibiotics. At an optical density of 0.6 -0.8 at 600 nm, expression was induced with 0.5 mM isopropyl thiogalactoside. Cells were allowed to grow at 30°C for 16 h, collected by centrifugation and stored at Ϫ20°C overnight. Cell paste was thawed and suspended in 3 ml of 50 mM Tris-HCl, 0.25 mM EDTA, pH 8.0, per gram of cell paste. Subsequent steps were at 0 -4°C. Lysozyme (1 mg/ml, total volume) was added, and the suspension was stirred for 30 min. The cells were sonicated for 12 cycles of 30 s each with intermittent cooling. Protamine sulfate was added to a final concentration of 0.2%, and the mixture was centrifuged after stirring for 30 min. The supernatant was applied to a DEAE Sepharose Fast Flow column (100 ml), and the enzyme was eluted with 50 mM Tris-HCl buffer, pH 8.0. The fractions with ADH activity were concentrated with an Amicon concentrator with a YM30 membrane under 40 psi N 2 while the buffer was changed to 50 mM sodium phosphate buffer containing 300 mM sodium chloride and 10 mM imidazole, pH 8.0. The enzyme was loaded onto a 15 ml nickel nitrilotriacetate-affinity column, which was washed with 50 mM sodium phosphate, 300 mM sodium chloride, and 20 mM imidazole buffer, pH 8.0, until the effluent had negligible protein. The column was developed with a 200-ml linear gradient from 20 to 250 mM imidazole. Fractions with ADH activity were concentrated with an Amicon concentrator with a YM30 membrane under 40 psi N 2 while the buffer was changed to 50 mM Tris-HCl and 0.25 mM EDTA buffer, pH 8.0, in order to remove imidazole. The enzyme was again loaded onto a DEAE column (15 ml), eluted with the 50 mM Tris-HCl buffer, and concentrated and stored at 4°C.
About 100 mg of protein was obtained from 9 L of culture. Electrophoretic analysis using polyacrylamide gel with sodium dodecyl sulfate showed a single band at the position expected for an ADH subunit (40 kDa). The ratio of tyrosine to tryptophan predicted from the sequence (3:2) was confirmed with an UV-absorption spectrum of the protein in 0.1 M NaOH (17). The absorption at 280 for a 1 mg/ml solution was estimated to be 0.37 at neutral pH from the spectrum in alkali. The turnover number was determined to be 0.66 s Ϫ1 and 1.07 s Ϫ1 based on a standard assay (18) at 25°C and 37°C, respectively.
X-ray Crystallography-Crystals of wild-type EqADH complexed with NADH and (R)-N-1-methylhexylformamide were prepared as described by the general procedure used previously (19). Recrystallized horse liver enzyme from Roche Applied Science (about 0.5 ml of 9.4 mg/ml) was dialyzed at 4°C against 10 ml of 50 mM ammonium N-[Tris(hydroxymethyl)methyl]-2-aminoethanesulfonate, pH 6.7 (measured at 25°C) with 1 mM NADH and 5 mM (R)-N-1-methylhexyl-formamide, and the concentration of 2-methyl-2,4-pentanediol was raised over 24 days to 21%, when crystals formed. The concentration of diol was finally raised to 25% over 17 more days. X-ray data were collected at 100 K at the IMCA-CAT beam line at the Advanced Photon Source at Argonne National Laboratories with a wavelength of 1.000 Å, a crystal to detector distance of 140 nm, an exposure time of 1 s, and an oscillation angle of 0.3 Å, for a total of 600 images. The data were processed with DEN20 and SCALEPACK (20). The structure was solved by molecular replacement using AMORE (21) and the coordinates for the refined wild-type horse ADH-NAD ϩ -2,3,4,5,6-pentaflourobenzyl alcohol complex (PDB entry 1hld) as a model. The structure was refined by cycles of restrained refinement using REFMAC (22) and model building with the program O (23). The initial electron density maps were calculated without including the formamide, but after one cycle of refinement, the positions for the formamides in all four subunits were readily apparent. Initially, the dictionary used for NADH maintained planarity of the nicotinamide ring, but the electron density maps clearly showed that the ring was not planar. Therefore, during the final cycles of refinement, the dictionary for NADH was modified to remove the restraints on planarity and torsion angles, as would be appropriate for NADH.
Biological Evaluation-The effects of the inhibitors on ethanol metabolism in mice were studied by the following procedure. Fed, male Swiss-Webster mice (20 -25 g, obtained from Harlan-Sprague, Indianapolis, IN) were administered intraperitoneal injections of the inhibitor in a dose of 0 to about 1 mmol/kg of body weight. The solutions of inhibitor were 0.1 M or lower in physiological saline. Ethanol (4 M in physiological saline) was administered 20 -30 min later, by intraperitoneal injection at 65 mmol/kg of body weight. Blood samples (10 l) were drawn from the tail at intervals timed from the ethanol injection. The blood alcohol concentration was determined by gas chromatography as previously described (10).

Evaluation of Inhibitors with Purified
Enzymes-We determined the inhibition constants for the series of N-substituted formamides, sulfoxides, and pyrazoles by initial velocity studies with varied concentrations of the inhibitor and a substrate appropriate for each purified alcohol dehydrogenase. Formamides produce uncompetitive inhibition against varied concentrations of ethanol and competitive inhibition against varied concentrations of cyclohexanone (Fig. 1). Previous studies showed that the K i values are not dependent on the substrate used and that the inhibition constants determined for the forward or reverse reaction were about (within 2-fold) the same (10, 13-15). The simplest kinetic explanation for the inhibition results is the Ordered Bi Bi mechanism where the inhibitor binds only to the enzyme-NADH complex. For this mechanism, the K i values represent true dissociation constants since the concentrations of coenzyme were saturating, and dissociation of NADH is rate-limiting for turnover (10).
The inhibition constants for EqADH, MmADH1, and HsADH1C*2 are presented in Table I. The K i values for most of the formamides ranged from about 1 to 100 M, indicating potent inhibition. These enzymes have a few differences of amino acid residues in the active sites and thus different specificities, which are reflected in K i values that can differ by 10-fold or more among the enzymes. It is significant, for instance, that the linear N-n-propyl-and N-n-butylformamides (1, 2) bind more tightly to EqADH and MmADH1 than to HsADH, whereas the secondary formamides that are branched at the one position (7, 8, 9, 10, 11, 12, 13, 14, and 20) bind more tightly to MmADH1 and HsADH than to EqADH. (R)-N-1-Methylhexylformamide (7), its heptyl analogue (10) and N-1,5dimethylhexylformamide (11) are particularly potent inhibitors of the human and mouse enzymes, with K i values less than 1 M. The methylated cyclohexylformamides are also better inhibitors of MmADH1 and HsADH than of EqADH. It appears that MmADH1 and HsADH have more room in the active site than does EqADH.
We also determined inhibition constants for sulfoxides and pyrazoles with MmADH1. The sulfoxides bind to the E-NADH complex and are uncompetitive inhibitors against varied concentrations of alcohol, whereas the pyrazoles bind to the E-NAD ϩ complex and are competitive inhibitors. The values for MmADH1 are similar to those observed for EqADH, HsADH1C*2, and rat ADH (10,11,15). The 3-substituted thiolane 1-oxides (alkyl derivatives of the 5-membered tetramethylene sulfoxide ring) are potent inhibitors of all of the enzymes, whereas the methyl alkyl sulfoxides are much less potent. The 4-substituted pyrazoles are also potent inhibitors.
X-ray Crystallography-The structure of EqADH complexed with NADH and (R)-N-1-methylhexylformamide was determined at 1.57 Å resolution. The x-ray data collection and refinement statistics are summarized in Table II. The asymmetric unit contained two molecules of enzyme; all four subunits have very similar structures, which are very similar to those of other ADH complexes, and are in the closed conformation (13,19,24). The location of the formamide is well-defined in the electron density maps (Fig. 2A). The results show how the enzyme binds the aldehyde analogue and accommodates the hexyl group.
The geometry of binding suggests that the formamide mimics the Michaelis complex with NADH and aldehyde (Fig. 2B). Significantly, the nicotinamide ring of the NADH-(R)-N-1methylhexylformamide complex is in a boat conformation where the N1 and C4 atoms are not in the same plane as atoms C2, C3, C5, and C6. The puckering of the nicotinamide ring can be described by the angle ␣N1 between the planes defined by atoms C2-N1-C6 and C2-C3-C6, angle ␣C4 between the planes defined by atoms C3-C4-C5 and C2-C3-C6, and the twist (distortion of the boat) calculated as the distance in angstroms between C5 and the plane defined by atoms C2-C3-C6 (25). For the formamide complex, ␣N1 is 12 Ϯ 2°, ␣C4 is 18 Ϯ 4°, and twist is 1.2 Ϯ 0.7 ⌭, mean values for the four subunits.
Biological Evaluation in Mice-The effects of five selected inhibitors on the rate of metabolism of ethanol were studied with mice, and the progress curves were fitted to an overall mechanism that provides in vivo inhibition constants and rate constants for the time-dependent elimination of the inhibitor. (In this context, we use the term "elimination" to include the metabolism and excretion of the alcohol and inhibitors.) The results presented in Fig. 3 show that N-1-methylhexylformamide significantly decreases the rate of elimination of ethanol. In the absence of inhibitor, the kinetics are zero-order, which is due to the saturation of the alcohol metabolizing system (i.e. it is at maximum velocity) and is typical of animals (26). In the presence of inhibitor, the time required for complete elimina-tion of alcohol is substantially longer, showing that inhibitors are active in vivo. However, the progress curves are not linear, probably due to elimination of the inhibitor and decreased efficacy over time. Thus, elimination rates and in vivo K i values are not easily calculated. Previous studies used estimates of the initial velocity of ethanol elimination in order to obtain K i values (10 -12). Advances in computing now provide a more robust method of analysis.
Numerical integration was used to fit the full progress curves to a mechanism that describes the overall process (27,28), and the apparent in vivo K i values and the rates of elimination of the inhibitors were determined. The mechanism used for the simulation is given in Fig. 4. The Ordered Bi Bi reaction of alcohol dehydrogenase is modified by including the binding of the inhibitor (I) to the enzyme-NADH complex to form E-NADH-I, which accounts for the uncompetitive inhibition against alcohol. The elimination of inhibitor (I 3 J) by a first order process is expressed as an additional reaction. Previous studies that directly determined the concentration of inhibitor in blood of rats gave a first order rate constant for isobutyramide of 0.042 min Ϫ1 (12) and for tetramethylene sulfoxide of 0.075 min Ϫ1 (10). Although ADH is the major enzyme system involved in ethanol metabolism (ϳ90%), other metabolic pathways such as a cytochrome P450 (CYP2E1) and excretion can account for elimination of ethanol when the ADH inhibitor is TABLE I Inhibition of purified liver alcohol dehydrogenases by formamides, sulfoxides, and pyrazoles The inhibition constants were determined in 83 mM potassium phosphate, 40 mM KCl, and 0.25 mM EDTA buffer, pH 7.3, at 37°C by measuring the change in absorbance of NADH at 340 nm. Buffers for the determinations cited from the literature were 46 mM sodium phosphate, pH 7.0 and 25°C. Substrates for the enzymes were chosen for optimal activity. The coenzyme concentrations were fixed at high, saturating levels (2.0 mM NAD ϩ or 0.2 mM NADH) while the concentrations of substrates and inhibitors were varied over at least a 3-fold range. Some 20 -25 data points were collected for each inhibitor. The data were fitted to the equations for competitive or uncompetitive inhibition with the appropriate computer programs (50). Standard errors are usually less than 20% of the values. ND, not determined.  saturating. In rats, non-ADH pathways were estimated to contribute about 10% of the total rate and to have an apparent K m value for ethanol of 21 mM (12). Sophisticated isotopic methods also show that alcohol dehydrogenase accounts for the metabolism of at least 89% of the ethanol (29,30). Thus, the reaction of enzyme F is added to the overall mechanism. Many of the rate constants for the mechanism in Fig. 4 can be estimated from kinetic constants that have been determined for liver alcohol dehydrogenases (31,32). Since ADH is typically saturated with substrates during ethanol metabolism, the turnover number (V max ) of ADH controls the rate of ethanol metabolism, and this is usually controlled by the rate-limiting release of NADH, k ϩ4 , which is the only sensitive rate constant for the ADH reaction. The maximum rate of ethanol elimina- FIG. 3. Inhibition of ethanol metabolism in mice by N-1-meth-ylhexylformamide. Mice were given varied doses of formamide and 10 min later a dose of 65 mmol/kg ethanol. The blood alcohol concentration was determined at various times. A total of five animals were used, with different concentrations of the inhibitor: 0 (q), 100 (f), 300 (ࡗ), 600 (OE) micromoles/kg. All the data were fitted together using the mechanism given in Fig. 4 and all rate constants fixed except for k Ϫ5 and k ϩ6 . The concentration of E (ADH) was set at 0.15 M, enzyme F at 0.1 M, NAD ϩ at a fixed steady-state level of 0.5 mM, NADH at a fixed steady-state level of 2 M, and the initial concentration of ethanol at 95 mM. The concentration of acetaldehyde is negligible due to rapid oxidation by aldehyde dehydrogenases (34). Fixed rate constants were as follows:  tion is the product of enzyme concentration and k ϩ4 . The concentration of ADH is calculated from the rate of ethanol elimination in the absence of inhibitor. (The concentration of ADH is calculated for the whole body mass and is consistent with measurements based on ADH activity in the liver and the fraction of body mass that is liver (33, 34)). Similarly, the concentration of enzyme F and its turnover number (k ϩ8 ) are set to account for 10% of the overall rate of ethanol elimination at the dose of ethanol used.
The data in Fig. 3 are described well by the mechanism in Fig. 4, and the estimates of the fitted parameters are good ( Table III). All of the inhibitors are effective in vivo. The rates of elimination of the inhibitors determined from these data are comparable to those determined from analysis of the remaining compounds in blood levels during the ethanol elimination (10,12). However, the in vivo K i values are typically about an order of magnitude larger than the ones determined in vitro with isolated enzyme. Previous studies showed that inhibition by tetramethylene sulfoxide, isobutyramide, isovaleramide, and 4-methylpyrazole also was characterized by larger K i values in vivo as compared with in vitro (10,12). The differences are not due to the different units being used (mol/liter in vitro; mol/kg in vivo) since the animal has about 70% body water (0.7 ml/g) as determined from the concentration of ethanol at zero time (mol/liter) as compared with the dose of ethanol (mol/kg). The differences do not appear to be due to an incorrect mechanism as previous studies showed that the types of inhibition by these compounds are essentially the same in vitro and in vivo (10,12). The differences are not due to metabolism of the inhibitors as the analysis of progress curves accounts for the elimination, and initial velocity analysis gives similar results. A simple explanation is that the bioavailability of the inhibitors in the animals reduces the effective concentration of free inhibitor and thereby increases the apparent K i values. If the compounds were preferentially absorbed in extrahepatic tissues or membranes, the concentration of free inhibitor in the liver cytosol would be reduced.
Potent inhibition in vivo requires a compromise in lipophilicity that is optimized for affinity for ADH and bioavailability. Increasing the size of the aliphatic side chain, from isopropyl to methylhexyl for instance, decreases the in vitro K i , but has little effect on the in vivo K i (Table III). Similarly, 3-butylthiolane 1-oxide is a better inhibitor in vivo than the more hydrophobic 3-phenyl and 3-hexyl derivatives or the less hydrophobic tetramethylene sulfoxide and 3-methylthiolane 1-oxide (11).
Toxicity of the formamides was not directly determined, but during the studies on the inhibition of ethanol metabolism, the mice survived for at least 1 day after the experiments with the highest doses (in mmol/kg) being 0.36 for N-isopropylformamide, 0.32 for N-isopentylformamide, 0.30 for N-1-methyl-hexylformamide, 0.40 for N-cyclohexylformamide, and 0.36 for 3-butylthiolane 1-oxide. However, at doses of 0.72 mmol/kg of N-isopentylformamide, 0.80 of N-cyclohexylformamide, 0.60 of N-1-methylheptylformamide, and 0.72 of 3-butylthiolane 1-oxide, toxicity was suspected because the rates of ethanol metabolism were depressed more than expected from the studies with lower doses of inhibitor. Furthermore, a dose of 1.8 mmol/kg of N-1-methylhexylformamide was lethal within 30 min.
The results show that the short chain N-isopropylformamide is about as potent in vivo as the longer chain compounds. The relatively slow rate of elimination of N-isopropylformamide is a potential advantage as compared with the other inhibitors. N-Cyclohexylformamide and 3-butylthiolane 1-oxide are also potent inhibitors, but their rates of elimination are faster.

DISCUSSION
Structure of the Michaelis Complex Analogue-The structure of the enzyme-NADH-(R)-N-1-methylhexylformamide complex appears to resemble the ground state for the NADH-aldehyde complex. The structural features are relevant for understanding the catalytic mechanism. The formamide oxygen is ligated to the catalytic zinc at a distance of 2.2 Å (Fig. 2A). The oxygen forms a hydrogen bond to the hydroxyl group of Ser-48, which is connected to the 2Ј-hydroxyl group of the nicotinamide ribose and eventually to His-51, which is on the surface of the enzyme (19). This system appears to relay a proton between substrate and His-51, which acts as an acid/base catalyst (19,(35)(36)(37). The inhibitor binds with a cis conformation, which produces a potential cation-interaction between the amide N-H and the benzene ring of Phe-93 of the enzyme. The formyl group of the formamide is oriented as would be expected for direct transfer of hydrogen from the nicotinamide ring of the coenzyme to the re face of the carbonyl carbon (Fig. 2B). The distance between the carbonyl carbon of the formamide and C4N of NADH is 3.6 Å, which is reasonable for the ground state for hydride transfer.
The puckering of the nicotinamide ring of NADH has not yet been observed in any other structure of a complex of a dehydrogenase with NADH and a substrate or substrate analogue. A twisted boat conformation has been observed for a complex of EqADH with NADH and a bound water, but the relevance of the complex for the catalytic mechanism is not clear (25). The nicotinamide ring is also puckered in the complex with NAD ϩ and pyrazole, which form a partial covalent bond between C4 of the nicotinamide ring and a N from pyrazole and may mimic the transition state of the reaction (38). Molecular dynamics simulations on the native ADH-NADH-benzaldehyde complex show fluctuations of the protein cause the nicotinamide ring of NADH to bend, with angles for the C4 deviation being as large as 20° (39). Calculations show that puckering of the dihydronicotinamide ring, to form a quasi-boat conformation, decreases the transition state energy for hydrogen transfer of the pseudoaxial hydrogen at C4 (40,41). The structure of the complex with NADH and the formamide suggests that the ground state can have a puckered nicotinamide ring.
Structure-Function Relationships-These studies provide data for the design of potential therapeutic agents and for correlating structure and function of the liver alcohol dehydrogenases. The inhibition constants determined in this study are thermodynamic dissociation constants that reflect the energetics of multiple binding interactions. Thus, the binding of the inhibitors is a measure of substrate specificity.
In order to explain the results in Table I, we docked models of the various inhibitors into the active sites of the enzymecoenzyme complexes. We used the structures of EqADH with NADH and (R)-N-1-methylhexylformamide or N-cyclohexylformamide (1LDY.PDB, Ref. 13). Structures of the mouse and human enzymes complexed with these formamides are not available, but there is a structure of the human liver alcohol dehydrogenase with coenzyme (1HTO.PDB, Ref. 42), and a model of the mouse enzyme was constructed based on the structure of EqADH (1HLD.PDB). These three enzymes share 84 -89% sequence identity, and the amino acid residues lining the substrate binding sites are identical, except at positions 110 and 141. As compared with Phe-110 and Leu-141 in the horse enzyme, the mouse enzyme has Ile-141, and the human enzyme has Tyr-110 and Val-141 (43). The substitutions at residue-141 should allow the substrate binding sites in the mouse and human enzymes to accommodate somewhat larger ligands. There are also other differences in the amino acid residues that make contact with the residues in the active site, and these may affect specificity through indirect effects on conformation and enzyme flexibility. The amino acid residues in the substrate binding site are all aliphatic or aromatic ( Fig. 2A). Only the catalytic zinc and Ser-48, which interact with the O of the substrate are polar. Thus, we expect that tight binding is associated with good van der Waals contacts or hydrophobic interactions, a lack of voids that would accommodate water molecules, and the absence of steric conflicts. We expect that an inhibitor that fills the space should bind optimally. Of course, the binding modes can vary due to flexible amino acid side chains that adapt to the ligand structure and optimize interactions (24).
The modeling shows that all of the formamides can fit, without bad steric contacts, into the substrate-binding sites, sometimes with small changes in positions of the amino acid side chains. However, it does not appear that binding affinity simply reflects hydrophobic interactions, as there are only modest changes in affinity as the size of the inhibitor (number of carbons) is increased (Table I). For instance, increasing the chain length from N-isopropyl (4) to (R)-N-1-methyhexyl (7) slightly decreased affinity for the horse enzyme, whereas affinities for the mouse and human enzymes were increased by about an order of magnitude. In general, the mouse and human enzymes have higher affinities for the larger molecules, which might be attributed to the substitutions at residue 141 (42). Consistent with this explanation is that the N-1-ethylhexyl and N-1-ethylheptyl formamides (13,14) bind less well than the N-1-methyl derivatives (9, 10) to the horse enzyme whereas they bind more tightly to the mouse and human enzymes. However, our modeling suggests that the N-1-ethyl derivatives can be accommodated in the horse enzyme in a position similar to the one observed for (R)-N-1-methylhexylformamide. Previous studies also suggested that the human enzyme could accommodate larger N,N-disubstituted, bicyclic formamides (14), and we conclude that the human enzyme has more flexibility to adapt to larger ligands than does the horse enzyme. In contrast, the N-n-propyl (1) and butyl (2) formamides bind more weakly to the human enzyme than to the horse enzyme, and this may be explained by the increased volume in the human enzyme that allows a water molecule to infiltrate and disrupt hydrophobic interactions. The factors that control binding affinity are subtle and require further studies to provide quantitative explanations.
In principle, enzymes are stereoselective for binding of inhibitors, and the horse enzyme binds the 1S isomers of 3-butylthiolane 1-oxide at least 10-fold more tightly than the 1R isomers (24). In contrast, the binding of the R and S isomers of N-1-methylhexylformamide shows a moderate preference for the R isomer. Both isomers can be modeled into the binding sites without producing bad steric contacts. Similarly, the isomers of the other chiral inhibitors (5, 6, 10, 11, 13, 14, 16, and 17, studied as racemic mixtures) could fit into the active sites. Adding methyl groups to the cyclohexyl rings (16, 17) did not improve binding. Further work is required to define the stereochemical preferences of the active site.
As previously determined for the horse and human enzymes, the methyl alkyl sulfoxides are less potent inhibitors than the cyclic thiolane 1-oxides, for which structures of the complexes with alcohol dehydrogenase have been determined by x-ray crystallography (24). Structures for the horse and human enzymes with pyrazole and 4-iodopyrazoles have been determined (38,44,45,46). The especially high affinity of the pyrazoles is due to the formation of a partial covalent bond, mimicking the transition state, with the bound NAD ϩ (38).
Biological Relevance-Liver alcohol dehydrogenase is the predominant rate-determining factor in the metabolism of ethanol and therefore a good target for inhibitory drugs (2,10,47). Since the Class I enzymes can detoxify various biogenic and dietary alcohols and aldehydes, we directed our studies to these enzymes. Recent gene knockout studies in mice provide compelling information that the ADH1 is the major enzyme involved in ethanol metabolism (48). Mice lacking a functional ADH1 gene have a significantly decreased rate of elimination of an intoxicating dose of ethanol. Isopropylformamide, cyclohexylformamide and 3-butylthiolane 1-oxide are good inhibitors of mouse ADH1 in vitro and are effective in vivo. These results support the conclusion that ADH1 is a major factor in ethanol metabolism and the proposal that the inhibitors are relatively specific.
These compounds are relatively non-toxic. The LD 50 for Ncyclohexylformamide is 2.5 mmol/kg (14), and that for 3-butylthiolane 1-oxide is 1.4 mmol/kg (11). Since the effectiveness of inhibition also depends upon the rates of metabolism of the inhibitors, the results on the apparent rate of elimination can be used to devise protocols for repetitive dosing. The formamides and 3-substituted thiolane 1-oxides are almost as potent as the competitive inhibitor 4-methylpyrazole, which is being used to treat methanol poisoning, but they are uncompetitive inhibitors against varied concentrations of ethanol and thus are effective even at saturating concentrations of alcohol. These compounds should be useful for studies on the metabolism of various alcohols and for treatment of poisoning due to methanol or ethylene glycol.


Synthesis, Crystal Structure, Stacking Effect and Antibacterial Studies of a Novel Quaternary Copper (II) Complex with Quinolone

The structural properties of a new copper (II) complex [Cu2(cip)2(bpy)2(pip)]·6H2O (bpy=2,2’-bipyridyl, cip=1-cyclopropyl-6-4-oxo-1,4-dihydroquinoline-3-carboxylic acid, pip= piperazinyl anion), obtained during the synthesis of the copper complex with ciprofloxacin (cpf), has been investigated. The complex crystallizes in the triclinic system, space group P-1. The cell dimensions are: a=6.874(2) Å, b= 10.761(3) Å, c= 17.969(5) Å, α=80.071(6)°, β= 85.253(6)°, γ=79.109(6)°，V=1284.5(6) Å3, and Z=2. The Cu (II) ion displays a five-coordinate square pyramidal coordination with two nitrogen donors from bpy, the 4-keto and 3-carboxylate oxygen donors of cip, and the third nitrogen atom of the pip anion occupying the fifth site. There is a stack effect between cip ring and bpy ring from another molecule, where the stacking distance is about 3.5 Å. The results of elemental analysis and FT-IR measurement are also included. Both ligand and complex were assayed against gram-positive and gram-negative bacteria by the doubling dilutions method. The complex shows the same minimal inhibitory concentration (MIC) against S. Aureus and E. Coli bacteria as the corresponding ligand.

Introduction
Many drugs possess modified pharmacological and toxicological properties when administered in the form of metallic complexes. Probably the most widely studied cation in this respect is Cu 2+ , since a host of low-molecular-weight copper complexes have been proven beneficial against several diseases such as turberculosis, rheumatoid, gastric ulcers, and cancers [1][2][3][4]. There has been a tremendous growth of drugs from quinolone family, which began with the discovery of nalidixic acid (Hnal) some 40 years ago. Since then, the exponential growth of this family has produced more than ten thousand analogues [5]. The coordination chemistry of these drugs with metal ions of biological and pharmaceutical importance is of considerable interest. Ciprofloxacin (cpf, Figure 1) was considered the best of the third generation quinolone family. There have been several reports about the synthesis and crystal structure of metal complexes with cpf [6][7][8][9]. Recently, we have found that during the synthesis of a copper complex containing the fluoroquinolone ligand, the cpf ligand was interestingly changed into the cip ligand ( Figure 2). The purity of the cpf used was determined by the Xinchang Pharmaceutical Factory and its structure was characterized by FT-IR, elemental analysis, 1 H-NMR, 13 C-NMR and MS. The crystal structure of this copper complex was also characterized and is reported in this paper.

(pip)]·6H 2 O [10]
The crystal data and refinement parameters of the title complex are summarized in Table 1 (see  Experimental). Figures 3,4 and 5 show the coordination of the metal and the atomic numbering scheme for the complex, and the crystal packing in the unit cell, respectively. The copper (II) atom is coordinated to the keto and the carboxylic acid oxygen of the cip to form a six-membered ring. The copper atom displays a five-coordinate square pyramidal coordination with two nitrogen donors from 2,2'-bpy, the 4-keto and 3-carboxylate oxygen donors, and one nitrogen atom of pip anion occupying the fifth site. The complex is dinuclear copper(II) complex.  [9,[12][13][14]. In the present study, we find that the Cu-O distances of 1.922(6)(acid) and 1.924(7)(keto) fall in the middle of 1.913～1.936 Å, as concluded by Wallis [12].
The dihedral angle between the two ligands in the complex [Cu 2 (cip) 2 (bpy) 2 (pip)]·6H 2 O is about 8°. There is a stack effect between cip rings from two neighbor molecules, and the distance is about 3.40 Å. Also there is a stack effect between cip ring and bpy ring from another molecule with a distance of about 3.4～3.6 Å. The observed stacking in these molecules are indicative of a π～π interaction at a distance that is similar with those encountered in the base stacking of DNA [15]. Also, this distance is encountered in some compounds like [Cu(phen)(cnx)(H 2 O)]NO 3 ·H 2 O [16], where the stacking distance between the cnx ring and phen ring from another molecule is about 3.5 Å, and in some intercalated compounds like 5-iodouridylyl-(3'-5')-adenosine:ethidium [17], where the stacking distances are: 3.3 Å (adenine-ethidium) and 3.4 Å (uracyl-ethidium), respectively. It is important to notice that the arrangement in the crystal packing is dependent on the environment of the metal ion, and this should be relevant in any proposal of intercalation processes. The data suggests that this type of compound could act as an intercalating agent and also that the mechanism of action should be by a metal-quinolone complex. Supporting evidence of this idea is given by the Shen's model [18], which proposes a direct interaction between the quinolones and DNA [19]. Figures 6 and 7 show the IR spectra of cpf and the complex, respectively. Comparing the main IR frequencies of copper complex with that of cpf, the following was found: (1) there were two very strong absorption peaks in the spectrum of the ligand cpf, i.e., 1707.6 cm -1 and 1627.1 cm -1 ; (2) the band at 1707.6 cm -1 , due to the carboxylic group, was not detected in the spectrum of the complex, indicating that this moiety participates in the bonding to the metal ion [20]; (3) however, the technique doesn't permit a definitive conclusion about the participation of the ketone group in the bonding to the metal, the corresponding band was recorded at 1627.1 cm -1 in the spectrum of cip, and a band close to this position (1635.4 cm -1 ) in the spectrum of the complex could be due to either the ketone group or to the carboxylic group bonded to the metal ion.

FT-IR Spectroscopy
If this band is due to the ketone group, the antisymmetric and symmetric modes of the carboxylic group would account for the bands at 1589.6 cm -1 and 1546.4 cm -1 (not labelled in Figure 7), respectively. However, if the ketone group participated in the bonding to the metal we would expect a shift of its stretching band towards lower wavenumbers, thus corresponding to the band recorded at 1589.6 cm -1 , and the bands at 1635.4 and 1473.1 cm -1 would correspond to the carboxylic group. According to the crystal structure, this conclusion is reasonable. We should conclude that FT-IR spectroscopy, by itself, doesn't permit a definitive answer as to the way the ligand is bonded to the metal cation. Macias [21] and Wang [22] have studied the IR spectra of some metal quinolone complexes and gave similar explanations.

Antibacterial Studies
Both ligand and complex were assayed against gram-positive and gram-negative bacteria in order to determine if the spectrum of activity of the complex was changed with respect to that of the ligand. Table 2 shows the results of the in vitro antimicrobial activity of the compounds. The complex shows the same minimal inhibitory concentration as the corresponding ligand against S. Aureus and E. Coli bacteria, and lower MIC against M. Lutens and P. Aeruginosa than that of ligand.

Possible synthetic reaction
During the reaction, the piperazine ring on ciprofloxacin was substituted by a chloride ion in the NaOH solution, and the chloride ion was from starting material cpf·HCl. Figure 8 shows the possible synthetic reactions taking place. Further experimental research is required to confirm the intermediate products, cip and pip, in this reaction process.

Chemicals, analyses and physical measurements
Cpf·HCl (pure powder, 99.9% ) was kindly donated by Xinchang Pharmaceutical Factory. All other chemicals used were of analytical reagent grade. The analysis of carbon, hydrogen, and nitrogen was carried out on a Carlo-Erba 1110 elemental analyzer. FT-IR spectra were recorded using KBr pellets on a Nicolet FT-IR spectrometer.
Synthesis of [Cu 2 (cip) 2 (bpy) 2 (pip)]·6H 2 O Bpy (0.5 mmol) dissolved in ethanol (10mL) was added to an aqueous solution (10 mL) containing Cu(NO 3 ) 2 ·3H 2 O (0.5 mmol). To this mixture was then added a solution prepared by dissolving cpf·HCl (1mmol) in water (100 mL) containing NaOH (2 mmol); the pH was adjusted to 7.0～8.0 with HNO 3 and NaOH. The resulting blue solution was slowly evaporated at room temperature, and blue crystals suitable for X-ray structure determination were formed finally after a period of 2 months. Anal. Calc.

Crystallographic Data and Structure Determination
A Bruker Smart CCD X-ray diffractometer with graphic monochromated Mo-Kα radiation and a 12kW rotating anode generator was used for the determination of the unit cell and data collection. The + data were collected using the θ and φ scan technique to a maximum θ value of 25.50°. The linear absorption coefficient, µ, for Mo-Kα radiation was 12.13 cm -1 . An empirical absorption correction based on azimuthal scans of several reflections was applied which resulted in transmission factors ranging from 0.77 to 1.00. The data were corrected for Lorentz and polarization effects. A correction for secondary extinction was applied (coefficient = 3.46614×10 -7 ). The structure was solved by direct methods using SHELXS86 and expanded using Fourier techniques. The non-hydrogen atoms were refined anisotropically. Hydrogen atoms were included but not refined. The final cycle of full-matrix least-squares refinement was based on 1853 observed reflections (I>2.00σ (I)) and 362 variable parameters and converged with unweighted and weighted agreement factors of R =║F 0 │-│Fc║ /│F 0 │=0.0776 and R W = [ω(│F 0 │-│Fc│) 2 /ωF 0 2 ] 1/2 = 0.2235. The weighting scheme, ω=1/(σ 2 (F 0 )).
Neutral atom scattering factors were taken from Cromer and Waber [11]. Anomalous dispersion effects were included in F calc . All calculations were performed using the teXsan crystallographic software package of Molecular Structure Corporation.

Antibacterial Susceptibility Tests
The minimal inhibitory concentration (MIC) for the 4 bacteria strains listed in Table 2 was measured by the agar incorporation method in plastic Petri dishes containing Mueller Hinton agar (10mL) and incorporating the antimicrobials in concentrations of 0.015 to 64 µg/mL in doubling dilutions, following the method of The National Center for Drug Screening (Shanghai, P.R. China).