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Originally published In Press as doi:10.1074/jbc.M409573200 on September 27, 2004

J. Biol. Chem., Vol. 279, Issue 50, 52346-52352, December 10, 2004
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TetX Is a Flavin-dependent Monooxygenase Conferring Resistance to Tetracycline Antibiotics*{boxs}

Wangrong Yang{ddagger}, Ian F. Moore{ddagger}, Kalinka P. Koteva{ddagger}, David C. Bareich{ddagger}, Donald W. Hughes§, and Gerard D. Wright{ddagger}

From the {ddagger}Antimicrobial Research Center, Department of Biochemistry and Biomedical Sciences and the §Department of Chemistry, McMaster University, Hamilton, Ontario L8N 3Z5, Canada

Received for publication, August 19, 2004 , and in revised form, September 23, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The tetracycline antibiotics block microbial translation and constitute an important group of antimicrobial agents that find broad clinical utility. Resistance to this class of antibiotics is primarily the result of active efflux or ribosomal protection; however, a novel mechanism of resistance has been reported to be oxygen-dependent destruction of the drugs catalyzed by the enzyme TetX. Paradoxically, the tetX genes have been identified on transposable elements found in anaerobic bacteria of the genus Bacteroides. Overexpression of recombinant TetX in Escherichia coli followed by protein purification revealed a stoichiometric complex with flavin adenine dinucleotide. Reconstitution of in vitro enzyme activity demonstrated a broad tetracycline antibiotic spectrum and a requirement for molecular oxygen and NADPH in antibiotic degradation. The tetracycline products of TetX activity were unstable at neutral pH, but mass spectral and NMR characterization under acidic conditions supported initial monohydroxylation at position 11a followed by intramolecular cyclization and non-enzymatic breakdown to other undefined products. TetX is therefore a FAD-dependent monooxygenase. The enzyme not only catalyzed efficient degradation of a broad range of tetracycline analogues but also conferred resistance to these antibiotics in vivo. This is the first molecular characterization of an antibiotic-inactivating monooxygenase, the origins of which may lie in environmental bacteria.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Tetracyclines represent one of the most successful classes of antibiotics used in the past 50 years. Since the first identification of chlortetracycline in 1948 from extracts of Streptomyces aureofaciens, numerous analogues, both natural and semisynthetic, have found clinical use (Fig. 1). This class of antibiotic has been a mainstay in the treatment of bacterial infections, being highly prized for a broad spectrum of antimicrobial activity, oral availability, and low cost. Tetracyclines are also widely used in agriculture to treat bacterial infections of plants (1), in aquaculture (2), and in animal growth (3). Given this intensive use, it is not surprising that tetracycline resistance has emerged over the past decades as a significant issue in the use of this class of antibiotic (for reviews, see Refs. 4-7).



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  		FIG. 1.
Tetracycline structure and nomenclature. A, structures of tetracycline antibiotics used in this study. B, numbering scheme of tetracycline antibiotics.

 
Tetracycline resistance in the clinic manifests itself through two primary mechanisms, active efflux and ribosomal protection. Efflux proteins, in particular, are major sources of resistance, responsible for the bulk of clinical failures of this class. These proteins belong to the major facilitator family of integral membrane efflux enzymes that couple the energetically unfavorable movement of tetracycline with the proton motive force to pump tetracyclines out of the cell against the concentration gradient. Ribosomal protection mechanisms, on the other hand, harness soluble structural homologues of elongation factors to destabilize the interaction between tetracyclines and their cellular target, the ribosome. In neither efflux nor ribosomal protection mechanisms, however, is the concentration of tetracycline in the environment altered.

In contrast, resistance to other antibiotics such as the {beta}-lactams and the aminoglycosides occurs primarily via the destruction or covalent modification of the antibiotics, which effectively decreases the local concentration of antibiotic. Such a mechanism was unknown for the tetracyclines except for a series of reports over a decade ago describing a gene, tetX, that encoded a putative 388-amino acid NADPH-requiring enzyme that was associated with tetracycline resistance (8-10). Amino acid sequence analysis revealed putative FAD-binding and monooxygenase fold domains (Fig. 2). The tetX gene was identified in transposons Tn4351 (10) and Tn4400 (9) harbored by the obligate anaerobe Bacteroides fragilis. Transfer of this gene to aerobically growing Escherichia coli uncovered a cryptic tetracycline resistance activity that was associated with destruction of the antibiotic and a commensurate darkening of the growth medium (9, 10). Preliminary biochemical studies using crude cell-free extracts revealed that both oxygen and NADPH were required for tetracycline resistance activity (11). More recently, two orthologues of the original gene, tetX1 and tetX2, were identified in another Bacteroides transposon, CTnDOT (12). The predicted TetX2 is 99% identical to the original TetX, whereas TetX1 is an N-terminal truncate (359 amino acids) with 66% identity to the other proteins lacking the FAD-binding domain (Fig. 2).



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  		FIG. 2.
Architecture of TetX orthologues from B. fragilis. The N-terminal domain 1, which is lacking in TetX1, is a predicted flavinbinding domain (Pfam FAD3), whereas the C-terminal domain 2 is a signature Pfam monooxygenase fold.

 
We report here the overexpression of TetX in E. coli and characterize it as a broad spectrum tetracycline-degrading enzyme operating by an unprecedented mechanism. TetX is a flavin-dependent monooxygenase that regioselectively hydroxylates the tetracycline substrate, resulting in an unstable compound that undergoes non-enzymatic decomposition.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Reagents—Tetracyclines, NADP+, NADPH, and glucose-6-phosphate were from Sigma. Glucose-6-phosphate dehydrogenase was from Roche Diagnostics.

Expression of Recombinant TetX Proteins—Plasmids encoding various tetX genes were the generous gifts of A. Salyers and N. Shoemaker, University of Wisconsin. Plasmid pDB1 was created by digestion of plasmid pBS2 (11) with HindIII and EcoRI, followed by ligation of the tetX-containing fragment into pUC18. Overexpression constructs of the tetX, tetX1, and tetX2 genes were amplified by PCR using the oligonucleotide primers listed in Supplementary Table I and using the appropriate plasmids as templates. PCR was performed on a Progeny 96-well thermocycler with 95 °C 1 min, 52 °C 1 min, 72 °C 1.5 min and was repeated for 30 cycles. The PCR products were excised from a 0.8% agarose gel, extracted by Qiagen QIAquick gel extraction kit, digested with NdeI and HindIII, and ligated into plasmid pET28 (Novagen) digested with the same restriction enzymes, generating fusion constructs that give an N-terminal His6-tagged protein for ease of purification. Plasmids were used to transform E. coli BL21(DE3) and were selected by kanamycin resistance. The absence of adventitious mutations during amplification was confirmed by complete gene sequencing.

A single colony of E. coli BL21(DE3) containing the appropriate plasmid constructs was used to inoculate 25 ml of Luria broth supplemented with 50 µg/ml kanamycin and incubated at 37 °C and 250 rpm for 12-16 h. Ten ml of this culture were used to inoculate 1 liter of Luria Bertani medium supplemented with 50 µg/ml kanamycin. The cultures were grown at 37 °C, 250 rpm to an A600 of ~0.6, followed by the addition of sterile isopropyl-{beta}-D-thiogalactopyrandoside to 1 mM. The culture was then incubated overnight with 250 rpm shaking at 16 °C.

Cells were collected by centrifugation at 8000 rpm for 5 min, resuspended in 10 ml of 0.1 mM dithiothreitol, 1 mM phenylmethanesulfonyl fluoride, 1 mM EDTA, 20 mM HEPES, pH 8.0, and lyzed by three passes through a French pressure cell at a maximum pressure of 20,000 psi. The cell lysate was clarified by centrifugation at 15,000 rpm for 15 min.

The supernatant was applied to a 1-ml nickel-agarose column equilibrated with 20 mM HEPES, pH 8.0. Enzyme was eluted by application of a linear gradient with 20 mM HEPES + 250 mM imidazole. Fractions were analyzed by electrophoresis through 11% sodium dodecylsulfate polyacrylamide gels to assess purity of the protein. If required, an additional chromatographic step consisting of application of the pooled fractions onto a Mono Q column equilibrated with 20 mM Tris-HCl, pH 8.0, and elution with a gradient to 1 M NaCl. Purified TetX and TetX2 (but not TetX1) were yellow in color, and all proteins were stored in 20 mM HEPES, pH 8.0. A 1-liter culture yielded 6 mg of pure protein (Supplementary Fig. S20).

Analysis of Flavin Cofactor Content—Purified TetX2 was boiled to denature the protein and briefly centrifuged to remove the precipitate. A sample of the supernatant was applied to a C18 column (10 units, 250 x 22 mm; Alltech Econosil) equilibrated with 5 mM ammonium acetate (pH 6.0). The bound flavin was separated by a linear gradient of 5 mM ammonium acetate (pH 6.0) to 100% methanol in 20 min with a flow rate of 1 ml/min while monitoring the absorbance at 451 nm. Commercial FMN, FAD, and riboflavin served as standards.

Spectrophotometric Assay of TetX Activity—Each 100-µl reaction in a 96-well microtitre plate included 1 mM NADPH and up to 3 mM tetracycline substrate in 25 mM TAPS, pH 8.5. The decrease in absorbance at 340 nm (NADPH oxidation) upon antibiotic inactivation or the change in the absorbance of oxytetracycline at 400 nm ({epsilon}400 = 1080 M-1 cm-1) was monitored using a Molecular Devices SpectraMax Plus microtitre plate reader.

Steady state kinetic parameters were determined by fitting initial rate (v) data to the standard Michaelis-Menten equation using the Grafit 4 software (13), v = kcat[Eo][S]/([S] + Km) where Eo is the total enzyme concentration.

Microbiological Assay—The effects of TetX activity on the antimicrobial properties of tetracyclines were assessed by a microbiological disk assay. Inactivation reactions contained 3 mM oxytetracycline, 1 mM NADP+, 40 mM glucose-6-phosphate, 0.3 unit of glucose-6-phosphate dehydrogenase, and 10 µg of purified TetX2, 25 mM TAPS, pH 8.5, in a total volume of 0.1 ml. A 15-µl aliquot was applied to a sterile filter paper disc (5 mm) and air dried for 45-60 min. The disk was then placed on a tryptic soy agar plate inoculated with an overnight culture of tetracycline-sensitive Micrococcus luteus diluted to an A625 nm of 0.008-0.01 and incubated at 30 °C for 48 h.

HPLC Separation of Products of Tetracycline Inactivation—The products of tetracycline inactivation were separated by reverse phase high performance liquid chromatography (HPLC)1 using a Dionex Acclaim 120 C18 column (3 µm 120 Å, 4.6 x 150 mm). The column was equilibrated with H2O plus 0.05% trifluoroacetic acid and tetracyclines, and the products of inactivation were eluted using a linear gradient to 95% CH3CN plus 0.05% trifluoroacetic acid over 14 min at a flow rate of 1 ml/min.

NMR Analysis of Oxytetracycline and Inactivation Products—A solution of 4 mM NADP+, 40 mM glucose-6-phosphate, and 20 units of glucose-6-phosphate dehydrogenase in 25 mM TAPS, pH 8.5, was incubated at 37 °C for 20 min to generate NADPH. MgCl2 (1 mM) was added, followed by 1.7 mg of TetX2 and 5 mg of oxytetracycline. The total reaction volume was 5 ml, and the mixture was incubated at room temperature. The progress of the reaction was monitored by reverse phase HPLC. Following completion of the reaction, concentrated HCl was added to a final concentration of 0.1 M. The crude reaction mixture was applied to a C-18 Sep-Pak mini column equilibrated in water. The product of TetX2 catalysis, P1 was eluted with water and the purity of the sample verified by HPLC. A total of four reactions were run, and purified P1 was combined and lyophilized. A final purification by preparative scale HPLC was performed prior to NMR analysis. The lyophilized product was dissolved in 0.1 M DCl/D2O and the 1H and 13C NMR spectra were recorded on a Brucker AV 600 instrument.

Mass Spectrometry—Mass spectrometry of the products of enzymatic reactions was performed on an Applied Biosystems Q Trap liquid chromatography mass spectrometry system. High resolution mass spectrometry was performed by Dr. K. Green at the McMaster Regional Center for Mass Spectrometry on a Waters-Micromass Global Ultima Quadrupole time of flight mass spectrometer.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
TetX Is a Flavoprotein—We first subcloned tetX into pUC18, creating the plasmid pDB1. This gene conferred tetracycline resistance to E. coli W3110 and resulted in discoloration of the medium associated with antibiotic inactivation (Fig. 3). To obtain purified proteins in sufficient quantity for molecular studies, we prepared separate constructs of N-terminal His6-tagged TetX, TetX1, and TetX2 in E. coli under control of a T7 promoter. Only the constructs expressing TetX and TetX2 conferred tetracycline resistance. Purified TetX and TetX2, but not the truncated TetX1, were visibly yellow in solution, indicative of bound flavin cofactor (see below). TetX1 is therefore an inactive protein, and its presence in the CTnDOT transposon may be a relic of an incomplete gene duplication event. We selected the TetX2-expressing construct for all further studies.



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  		FIG. 3.
Effect of TetX on tetracycline in liquid culture. E. coli W3110 bearing the tetX gene on plasmid pDB1 when grown in the presence of tetracycline turns the medium black and is associated with tetracycline degradation.

 
Purification of TetX2 gave a protein of the predicted 44 kDa by SDS-polyacrylamide gel electrophoresis, and analytical gel filtration was consistent with a monomeric protein of the correct size. Solutions of purified TetX2 were yellow in color, and the UV-visible spectrum was consistent with the presence of a flavin cofactor with absorbance maxima at 366 and 445 nm (Fig. 4). Denaturation of the enzyme followed by reverse phase HPLC of the soluble material identified FAD as the bound cofactor. The FAD-TetX2 stoichiometry was found to vary between batches (~60->80%), and addition of exogenous FAD to 2 µM was found to stabilize activity. Therefore, this coenzyme was typically added to purified protein during storage in the dark.



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  		FIG. 4.
UV-visible spectrum of purified TetX2. The absorbance maxima at 366 and 445 nm shown in the inset are indicative of flavin content.

 
TetX Requires NADPH and O2 for Enzyme Activity—Primary sequence analysis and preliminary studies by Salyers and coworkers (11) using cell extracts predicted that TetX might be an NADP+-requiring oxidoreductase. We confirmed with purified enzyme that NADPH was essential for oxytetracycline degradation activity and could not be substituted by NADH. Furthermore, degassing of solutions followed by incubation of the reaction in an N2-only atmosphere resulted in no inactivation of oxytetracycline (not shown), demonstrating that O2 was also essential for oxytetracycline inactivation, and consistent with the observation that only aerobically grown cultures showed tetracycline resistance (10). A survey of the influence of initial rate on pH demonstrated that the enzyme showed maximal activity at pH 8.5.

TetX Catalyzes the Inactivation of a Broad Spectrum of Tetracycline Antibiotics—The tetracycline inactivation activity of TetX2 was established by a series of biochemical assays including UV-visible spectroscopy and reverse phase HPLC. Tetracyclines show two absorption maxima, one at 260 nm and another at 363 nm. The {beta}-tricarbonyl chromophore (ring A) is responsible for the 260 nm absorbance, whereas the aryl {beta}-diketone chromophore (rings B-D) is responsible for the 363 nm absorbance and the yellow color of tetracyclines. Incubation of tetracyclines and TetX2 under assay conditions (presence of NADPH and O2, pH 8.5) resulted in the disappearance of the absorbance maximum at 363 nm, a more modest decrease in the absorbance maximum at 260 nm, and a broad low intensity increase in the absorbance at wavelengths greater than 430 nm (Fig. 5).



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  		FIG. 5.
UV-visible spectrum of oxytetracycline degradation catalyzed by TetX2. The sample cuvette contained purified TetX2, oxytetracycline, and an NADPH regenerating system, whereas the reference cuvette did not contain TetX2 (see "Experimental Procedures" for details). Each scan was taken at 20-s intervals.

 
These changes in absorbance as a result of enzyme action provided a means to continuously monitor the progress of the reaction. Because NADPH has a maximum absorbance at 340 nm, the absorbances of both substrates overlap in the 360 nm region. Therefore, the absorbance at 400 nm ({epsilon}400 of oxytetracycline is 1080 M-1 cm-1) (See Fig. 5 for spectrum) was chosen to monitor the enzyme activity in continuous assays in 96-well microtitre plates.

The progress of the TetX2-catalyzed reaction could also be monitored by the disappearance of the antibiotic substrate (S) and the appearance of product peaks by reverse phase HPLC (Fig. 6). Using oxytetracycline (Fig. 1) as a model substrate, the antibiotic was converted to two products, P1 and P2 (Fig. 6). The temporal separation between the appearance of P1 and P2 implied that P2 was derived from P1 (Fig. 6). Further analysis of the conversion of P1 to P2 revealed that this process was enzyme-independent and accelerated at neutral pH but was slower at acidic pH (data not shown). Therefore, the exclusive product of TetX2 activity is P1.



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  		FIG. 6.
Reverse phase HPLC separation of oxytetracycline and TetX2-catalyzed products. A, decrease in the oxytetracycline peak at 363 nm resulting from disruption of the aryl {beta}-diketone chromophore upon TetX2-catalyzed hydroxylation. B, absorbance at 260 nm reflecting the aromatic (Ring D) region of the substrates and products.

 
TetX2-catalyzed modification of oxytetracycline was associated with loss of antibiotic activity; therefore, TetX has been rigorously established as a tetracycline-inactivating enzyme. Purified P1 did not show any antimicrobial activity in our assays. However, because we demonstrated that this compound is unstable at neutral pH, we cannot conclusively state that P1 is devoid of antimicrobial activity itself or whether downstream products such as P2 are the inactive compounds. We do know that initial modification of antibiotic catalyzed by TetX2 is essential for conversion of tetracycline into the inactive metabolites that eventually result in the black pigment seen in cell-free extracts that are characteristic of the presence of this enzyme. Preliminary studies indicate that this black pigment is a high molecular weight polymer of undefined structure.2

TetX2 catalyzed the efficient inactivation of a variety of tetracycline antibiotics (Tables I and II). The enzyme showed a maximum 5-fold discrimination between these substrates in the steady state, including natural products such as tetracycline, oxytetracycline, and demeclocycline as well as semisynthetic compounds such as minocycline and doxycycline (Table I).


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  		TABLE I
Steady state kinetic parameters for the inactivation of tetracycline analogues by TetX2 Kinetic values are shown ± S.E. of fit of the data to the Michealis-Menten equation.

 


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  		TABLE II
MIC values of tetracycline antibiotics with E. coli W3110/pDB1 Plasmid pDB1 contains the tetX gene in a pUC18 background. Values for pUC18 alone were <2 µg/ml.

 
TetX2 Is a Monooxygenase That Oxidizes Oxytetracycline— Liquid chromatography mass spectrometry analysis of the first product peak of oxytetracycline degradation, P1 (Fig. 6B), gave an m/z value of 477 in positive ion mode, equivalent to the addition of 16 Da to oxytetracycline (mass 461). This is consistent with monooxidation of the tetracycle. The oxidized product P1 is unstable under the conditions (pH 8.5) of the enzymatic reaction. However, it was found that decomposition of P1 could be prevented by acidifying the product reaction mixture to pH 1. Therefore, purified P1 was kept in 0.1 M HCl, and characterization by NMR was performed in 0.1 M DCl/D2O. High resolution positive ion electrospray ionization mass spectrometry performed on purified P1 gave an m/z of 477.1502. The expected mass of positively charged P1 corresponding to C22H25N2O10+ is 477.1509.

The Structure of P1—To determine the identity of P1 the 1H, 13C, correlated spectroscopy, HSQC, HMBC, and nuclear Overhauser effect spectra of P1 were determined and compared with the corresponding spectra of oxytetracycline determined under identical conditions (see Supplementary information for complete spectra, Figs. S1-S19 and Tables S2-S5 for complete assignments). The one-dimensional 1H NMR spectra of oxytetracycline and P1 are shown in Fig. 7. There are the same number of proton resonances observed in the 1H NMR spectra of both oxytetracycline and P1. The aromatic region of the 1H NMR spectrum of P1 (H8 (7.60 ppm), H7 (7.12 ppm), H9 (7.04 ppm)) shows no change in the observed coupling pattern or significant changes in chemical shift when compared with oxytetracycline. This is significant because a large number of flavin monooxygenases hydroxylate activated aromatic rings similar to ring D of oxytetracycline. The methine protons of P1 show significant changes in chemical shift and coupling compared with oxytetracycline. In oxytetracycline (0.1 M DCl/D2O), the four non-aromatic methine protons are observable as a doublet at 4.30 ppm (H4, J4,4a = 1.4 Hz), a doublet of doublets at 3.87 ppm (H5, J5,4a = 11.4 Hz, J5,5a = 8.3 Hz), a doublet at 2.89 ppm (H5a), and a doublet of doublets at 2.87 ppm (H4a). In P1, four methine proton signals are observed as a doublet at 4.16 ppm (J = 8.9 Hz), a doublet at 3.97 ppm (J = 1.4 Hz), a doublet of doublets at 3.67 ppm (J = 8.9, 1.4 Hz), and a singlet at 2.85 ppm. The correlated spectroscopy spectrum of P1 confirms the coupling interaction between the doublet of doublets at 3.67 ppm and the doublets at 4.16 and 3.97 ppm (Supplementary Figs. S5-S7). The collapse of one doublet to a singlet and the loss of one of the doublet of doublets from the oxytetracycline spectrum mean the coupling interaction of either H5a or H4 has been removed from the system of methine protons in P1. Either the C5a-H5a or C4-H4 bond is broken and a new C-H bond is formed somewhere else in the molecule during the conversion of oxytetracycline to P1 or the dihedral angle between H5a or H4 and their adjacent proton is 90°. A nuclear Overhauser effect difference spectrum of oxytetracycline obtained by saturating the protons of the methyl group at carbon 6 (6-CH3, 1.72 ppm) reveals enhancement of the resonances of H7 (7.14 ppm), H5 (3.876 ppm), and H5a (2.90 ppm) (Supplementary Fig. S8). By saturating the protons of 6-CH3 (1.45 ppm) in P1, enhancements of H7 (7.12 ppm) and the methine signals at 4.16 and 2.85 ppm are observed (Supplementary Fig. S9). Because the same number of enhancements are observed in both oxytetracycline and P1, a change in bonding around 6-CH3 is unlikely and suggests the resonance for H5a is the singlet at 2.85 ppm.



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  		FIG. 7.
The 600 MHz 1H NMR spectra of P1 and oxytetracycline in 0.1 M DCl/D2O. The aromatic region of P1 shows no change compared with oxytetracycline. The non-aromatic region shows significant changes in chemical shift and coupling of the methine protons of P1.

 
The 13C NMR spectrum of P1 contains 21 signals, the same number observed for oxytetracycline (Supplementary Figs. S10-S12). In the 13C NMR spectrum of oxytetracycline there are five carbonyl resonances assigned to C11, C12, C1, C3 (shown as enol tautomers), and CONH2 (14). In P1 there are only four, suggesting one of the keto/enol carbons has undergone a hybridization change. Concomitant with the loss of a carbonyl resonance is the appearance of a new signal at 102.29 ppm in the 13C NMR spectrum of P1.

The two-dimensional HSQC and HMBC proton-carbon correlation spectra help to assign the 13C resonances and identify the site of hydroxylation. As expected, the HSQC spectrum of P1 shows ten proton-carbon correlations (Supplementary Fig. S13). These correlations fix three of the six aromatic carbons, C8 (137.40 ppm), C9 (117.68 ppm), and C7 (114.75 ppm) (Supplementary Fig. S14). The 6-CH3 carbon resonance is 17.23 ppm, and the two carbon resonances of the dimethyl amine group are 43.37 and 41.91 ppm (Supplementary Fig. S15). The four methine carbon resonances are 69.27, 66.33, 59.77, and 43.37 ppm.

The HMBC spectrum of P1 contains several 2- and 3-bond proton-carbon correlations (Supplementary Figs. S16-S19). The protons of 6-CH3 are expected to show three correlations, a 2-bond coupling to C6 and two 3-bond couplings to C6a and C5a. All three couplings are observed in the HMBC spectra, allowing the following assignments, C6a (147.59 ppm), C6 (79.63 ppm), and C5a (59.77 ppm) (Supplementary Fig. S16). The singlet at 2.85 ppm in the 1H NMR spectrum can now be assigned to H5a because the HSQC spectrum shows a correlation between C5a (59.77 ppm) and the singlet at 2.85 ppm. Taken with the nuclear Overhauser effect data, this indicates that in P1 the dihedral angle between H5a and H5 is close to 90°, explaining the lack of observed coupling between them. The remaining proton resonances can then be assigned as follows, H5, 4.16 ppm (JH5,H4a = 8.9 Hz); H4, 3.97 ppm, (JH4,H4a = 1.8 Hz); and H4a, 3.67 ppm (JH4a,H5 = 8.9 Hz, JH4aH4 = 1.8 Hz). Using the 1-bond correlations in the HSQC spectra, the remaining methine carbons can be assigned as follows, C4 (69.27 ppm), C5 (66.33 ppm), and C4a (43.37 ppm).

Proton H5a is expected to show three 2-bond 1H-13C couplings to C11a, C5, and C6 and four 3-bond couplings to C6a, C4a, C11, and C12. Five correlations are observed in the HMBC spectrum, 199.51, 102.29, 85.22, and the already assigned 66.33 (C5) and 43.37 ppm (C4a) (Supplementary Figs. S17, S18). Correlations to the already assigned C6a (147.59 ppm) and C6 (79.63 ppm) are not observed. The three resonances at 199.51, 102.29, and 85.22 ppm therefore correspond to C11, C12, and C11a. These are significant chemical shift changes compared with oxytetracycline where C11 is 193.50, C12 170.05, and C11a 104.24 ppm. C11a and C12 have undergone a hybridization change because their signals have shifted upfield.

Flavin-dependent hydroxylases act on activated double bonds. The most widely studied reactions are those where hydroxylation of an aromatic substrate occurs. A typical example is phenol hydroxylase where the substrate is hydroxylated ortho to the hydroxyl group. An intermediate {alpha}-hydroxy ketone is formed during the reaction that loses a proton in a subsequent step to regenerate the aromatic ring. The aromatic ring of oxytetracycline is unchanged upon transformation to P1. The enol tautomer of a {beta}-diketone can be considered an activated double bond. Hydroxylation of C11a of the {beta}-diketone would generate an {alpha}-hydroxy ketone at C11a-C12. Unlike the substrates for aromatic hydroxylases, there is no proton {alpha} to the carbonyl group at C11a to deprotonate and reform the double bond of the enol. C11a changes hybridization from sp2 to sp3 during hydroxylation and is consistent with the change in chemical shift from 104.24 to 85.22 ppm. The formation of an {alpha}-hydroxy ketone is expected to shift the C12 resonance downfield, not upfield, from 170.05 to 102.29 ppm. However, there is a unique property of 6-hydroxytetracyclines that explains the shift of the C12 resonance upfield. Non-enzymatic oxidation reactions of tetracyclines at C11a are known. In 1963 Blackwood et al. (15) prepared several C11a-halogenated derivatives of various tetracyclines. These included 11a-fluorotetracycline, 11a-fluorooxytetracycline, 11a-chlorotetracycline, and 11a-chlorooxytetracycline. They observed that the 11a-halogenated tetracycline derivatives with a hydroxyl group at C6 (like oxytetracycline) formed an intramolecular 6,12-hemiketal with the carbonyl group at C12. The 11a-halogenated tetracycline derivatives without a hydroxyl group at C6, 11a-fluoro-6-demethyl-6-deoxytetracycline and 11a-fluorodeoxy-oxytetracycline did not form an intramolecular hemiketal, and a free ketone group at C12 was observed in the IR spectra of these compounds. We therefore propose that TetX2 hydroxylates C11a of oxytetracycline and that the ketone initially formed at C12 undergoes rapid conversion to the 6,12-hemiketal. This is consistent with a chemical shift value of 102.29 ppm for C12. Formation of the intramolecular hemiketal explains the acid stability of P1 and accounts for the loss of a carbonyl signal in the NMR. Also, formation of the intramolecular bridge adds a degree of rigidity to the molecule. Comparison of MM2 energyminimized structures of 11a-hydroxy-oxytetracycline with and without the 6,12-hemiketal shows that formation of the intramolecular bond changes the dihedral angle between H5a,H5 from 145° to 87°. This is consistent with the lack of coupling observed between H and H5a and H5 1H in the NMR spectrum.

The remaining 13C assignments are as follows, C11 (199.51 ppm), C1 (194.01 ppm), C3 (186.54 ppm), CONH2 (171.18 ppm), C10 (161.12 ppm), C10a (113.19 ppm), C2 (98.96 ppm), and C12a (74.10 ppm). These carbons do not show any significant chemical shift difference from oxytetracycline.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The enzymes TetX and TetX2 are FAD-requiring monooxygenases that inactivate a broad selection of tetracycline antibiotics. The presence of this prosthetic group suggested that tetracycline inactivation could be the result of reductive electron transfer or hydroxylation reactions. We demonstrated that TetX requires both NADPH and molecular oxygen to inactivate tetracycline, consistent with a role as a monooxygenase. These results predict NADPH reduction of the flavin cofactor to FADH2, followed by reaction of O2 with the resulting electronrich isoaloxazine to form a reactive FAD-4a-hydroperoxide. Our work has demonstrated that tetracycline inactivation is triggered by regiospecific hydroxylation at C11a (Scheme 1), which could proceed via initial epoxidation of the C11a-C12 enol. The absence of the predicted carbonyl signature of 11a-hydroxy-oxytetracycline in the 13C NMR spectrum suggests that the acid-stabilized product P1 is the 6-12-hemiketal.



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  		SCHEME I
 
We were unable to directly assess the antibiotic activity of P1 as this compound spontaneously decomposed to downstream inactive products. However, hydroxylation at C11a would likely impact the Mg2+-chelating properties of tetracycline, which are a requirement for ribosome binding (16). Liquid chromatography mass spectrometry analysis of the HPLC peak P2 revealed loss of a molecule of water; however, we were unable to purify and study this compound as this peak contained more than one unstable compound (not shown). The expression of the hydroxylase TetX in E. coli aerobically grown in the presence of tetracycline results in formation of an amorphous black pigment, indicative of a complex polymeric structure. Therefore, initial hydroxylation catalyzed by this enzyme precipitates a cascade of molecular events that result in tetracycline deactivation. This is the first example, to our knowledge, of a flavin monooxygenase catalyzing antibiotic inactivation.

TetX is capable of recognizing and inactivating a broad range of tetracycline antibiotics of both natural and semisynthetic origin. In vitro analysis of specificity as measured by kcat/Km using purified enzymes showed only a 6-fold difference between the best substrate (oxytetracycline) and the poorest (minocycline). Correspondingly, minocycline is also the poorest substrate for the enzyme in vivo with a MIC of 8 µg/ml, whereas the presence of the enzyme confers resistance to 256 µg/ml oxytetracycline. The correlation is, however, not as complete as tetracycline, which is comparable with minocycline as a substrate of the purified enzyme; nonetheless, it is robustly resisted in vivo. The disparity in MIC is therefore not reflected adequately in disparity of similar magnitude in steady state kinetic parameters. The molecular basis for this observation is obscure at present, but the MIC data may also reflect additional downstream processing and inactivation of tetracycline into multiple products.

The paradoxical discovery of the tetX genes, which we have unambiguously shown encode oxygen-requiring monooxygenases, in the obligate anaerobe B. fragilis speaks to the extensive exchange of genetic material in the microbial world. The tetX genes were all localized on Bacteroides transposons that mobilize genes for exchange. The G+C content of the tetX genes is marginally lower (37%) than the B. fragilis genome (42%), but it is within the reported values of other bacteria of the same genus (11). BLAST search reveals the presence of highly homologous gene products (E values <10-42) of TetX in the sequenced genomes of the aerobic soil bacteria Cytophaga hutchinsonii (phylogenetically related to the Bacteroides), Streptomyces coelicolor, and Streptomyces avermitilis, suggesting that this family of enzymes is widespread in the environment. Aromatic polyketide natural products that resemble tetracyclines, for example the anthracyclines such as daunorubicin, actinorhodin, granaticin, and many others, are produced by numerous Streptomyces. TetX-like monooxygenases that are present in these bacteria may reflect the density of such molecules in the environment and the requirement to oxidatively modify them, perhaps not always as a means of detoxification but also in biosynthesis. Although the G+C content of tetX genes isolated from Bacteroides does not approach that of the Streptomyces (>70%), it is similar to the G+C content of Cytophaga (33-42%). Like other antibiotics such as the aminoglycosides (17) and the glycopeptides (18), the origin of this mechanism of tetracycline resistance may be the environment.


   FOOTNOTES
 
This article is dedicated to Professor Christopher Walsh on the occasion of his 60th birthday.

* This work was supported by the Natural Sciences and Engineering Council of Canada and by a Canada Research Chair in Antibiotic Biochemistry (to G. D. W.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{boxs} The on-line version of this article (available at http://www.jbc.org) contains 20 supplementary figures and 5 tables. Back

To whom correspondence should be addressed. Tel.: 905-525-9140 (ext. 22454); Fax: 905-522-9033; E-mail: wrightge{at}mcmaster.ca.

1 The abbreviations used are: HPLC, high performance liquid chromatography; MIC, minimum inhibitory concentration; TAPS, N-tris-(hydroxymethyl)methyl-3-amino-propanesulfonic acid; HSQC, heteronuclear single quantum coherence; HMBC, heteronuclear multiple bond coherence. Back

2 G. D. Wright and D. C. Bareich, unpublished results. Back


   ACKNOWLEDGMENTS
 
We thank A. Salyers and N. Shoemaker for gifts of tetX plasmids.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

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