C10-Benzoate Esters of Anhydrotetracycline Inhibit Tetracycline Destructases and Recover Tetracycline Antibacterial Activity

Abstract

Tetracyclines (TCs) are an important class of antibiotics threatened by enzymatic inactivation. These tetracycline-inactivating enzymes, also known as tetracycline destructases (TDases), are a subfamily of class A flavin monooxygenases (FMOs) that catalyze hydroxyl group transfer and oxygen insertion (Baeyer–Villiger type) reactions on TC substrate scaffolds. Semisynthetic modification of TCs (e.g., tigecycline, omadacycline, eravacycline, and sarecycline) has proven effective in evading certain resistance mechanisms, such as ribosomal protection and efflux, but does not protect against TDase-mediated resistance. Here, we report the design, synthesis, and evaluation of a new series of 22 semisynthetic TDase inhibitors that explore D-ring substitution of anhydrotetracycline (aTC) including 14 C10-benzoate ester and eight C9-benzamides. Overall, the C10-benzoate esters displayed enhanced bioactivity and water solubility compared to the corresponding C9-benzamides featuring the same heterocyclic aryl side chains. The C10-benzoate ester derivatives of aTC were prepared in a high-yield one-step synthesis without the need for protecting groups. The C10-esters are water-soluble, stable toward hydrolysis, and display dose-dependent rescue of tetracycline antibiotic activity in E. coli expressing two types of tetracycline destructases, represented by TetX7 (Type 1) and Tet50 (Type 2). The best inhibitors recovered tetracycline antibiotic activity at concentrations as low as 2 μM, producing synergistic scores <0.5 in the fractional inhibitory concentration index (FICI) against TDase-expressing strains of E. coli and clinical P. aeruginosa. The C10-benzoate ester derivatives of aTC reported here are promising new leads for the development of tetracycline drug combination therapies to overcome TDase-mediated antibiotic resistance.

Tetracyclines (TCs) are essential antibacterial agents used for treating a wide range of clinical infections caused by aerobic, anaerobic, Gram-positive, and Gram-negative pathogens.¹⁻³ Third generation TCs including tigecycline (Tig), eravacycline, and omadacycline are considered drugs of last resort for treating multidrug resistant (MDR) pathogens such as MDR Acinetobacter baumannii.⁴ These third-generation TCs overcome traditional clinical resistance mechanisms including efflux pumps and ribosome protection proteins,⁵ but a new resistance mechanism is emerging in the form of tetracycline-inactivating enzymes known as tetracycline destructases (TDases). TDases have been found within inducible antibiotic resistance operons in MDR bacterial pathogens including A. baumannii, Pseudomonas aeruginosa, Legionella longbeachae, and Mycobacterium abscessus.⁶⁻¹⁰ TDases can inactivate all generations of the TC antibiotic family and threaten the clinical viability of tetracycline drugs of last resort.^11,12

We and others have characterized the dissemination of TDase genes in the environment, including hospital settings.¹² The occurrence of TDases is increasing in clinical pathogens and there is soon to be a serious clinical need for TDase inhibitors, much like the standard of care established for beta-lactam antibiotic and beta-lactamase inhibitor combination therapies.^13,14 We have reported that anhydrotetracycline (aTC) and its analogs are potent broad-spectrum inhibitors of the two major classes of TDases (Type 1 and 2) found in bacterial pathogens (Figure 1).^9,15 However, aTC is cytotoxic^16,17 and certain Type 1 TDases are capable of oxidizing aTC,¹⁸ which led us to develop a series of bivalent TDase inhibitors based on the fusion of aTC and benzamide linked biomimetics of nicotinamide placed at the C9-position of the D-ring.¹⁹ These bivalent inhibitors are competitive with both TC and NADPH substrates to block the initial reduction of FAD and prevent turnover of both the inhibitor and the substrate. This strategy results in more efficient TDase inhibition and preserves the inhibitor scaffold by blocking inhibitor oxidation. In our initial work, we explored two types of linkages at the C9-position of aTC: benzamides and benzylamines. The benzamides are more potent inhibitors but have limited water solubility and poor cell permeability consistent with the observed weak rescue effect in E. coli when dosed in combination with TC antibiotics.¹⁹ Here, we report that C10-ester TDase inhibitors are fully water-soluble at all concentrations tested, and have improved whole cell activity compared to C9-amides for the rescue of growth inhibition by TC and Tig against E. coli, P. aeruginosa, and Mycobacterium smegmatis expressing Type 1 (TetX or TetX7) and Type 2 (Tet50) TDases.

Figure 1.

Summary of three aTC-based inhibitor types for TDases: (1) aTC analogs; (2) C9-benzamides/benzylamines; (3) C10-benzoate esters (this work). aTC analogs compete directly with TC for TDase binding to inhibit the coupled path for TC oxidation and directing the TDase down the uncoupled path for release of H₂O₂. C9-benzamides/benzylamines are bivalent inhibitors that compete with TC and NADPH for TDase binding to block FAD reduction. C10-benzoate esters display mixed inhibition patterns with some analogs inhibiting FAD reduction and other analogs supporting the uncoupled release of H₂O₂.

Results and Discussion

Design and Synthesis of C10-Benzoate Inhibitors

To improve the water solubility and cell permeability of aTC-based inhibitors, we explored modifications of the aTC scaffold at different positions. While synthesizing C9-amides from C9-amino aTC, we observed the formation of an additional product that we hypothesized to be the C10-ester which spontaneously rearranges to make the more thermodynamically stable C9-amide product (Figure S1). To test this hypothesis, we treated aTC with benzoic acids in the presence of a coupling reagent (HATU) under basic conditions (DIPEA) in DMF solvent. We observed rapid (<15 min), clean, and near complete conversion of aTC (1) to the corresponding C10-esters 2–15 (Scheme 1). We confirmed the chemoselective esterification of the C10-hydroxyl group through complete structural elucidation via multidimensional NMR analysis including COSY, TOCSY, HMBC, and HSQC (see Supporting Information). We observed a consistent downfield shift of the C9 hydrogen in the ¹H NMR spectrum of the C10 esters relative to aTC due to the exposure of the C9–H to the deshielding effects of the C10 ester carbonyl. In aTC, the chemical shift order of the D-ring hydrogens appears as C8 (t, 7.6 ppm) > C9 (d, 7.5 ppm) > C7 (d, 6.9 ppm). In C10 ester 2, for example, the chemical shift order of the D-ring hydrogens appears as C9 (d, 8.1 ppm) > C8 (t, 7.8 ppm) > C7 (d, 7.4 ppm). We observed a similar pattern for all the C10 ester derivatives. The yields after purification by prep-HPLC ranged from 21 to 95%. This allowed for the rapid synthesis of 14 analogs derived from substituted benzoic acid and heterocyclic aromatic acids to explore the potential of C10 esters as inhibitors of TDases. For direct comparison, we synthesized an additional 8 aTC analogs as C9-amides 16–23 using the same aromatic acids as C10-esters 7–13 and 15. While amide coupling with C9-NH₂-aTC is also a one-step synthesis, the yields for this reaction compared to C10-esterification are significantly lower ranging from 8 to 32%. Further, the preparation of C9-NH₂-aTC requires two additional synthetic steps starting from the commercially available aTC.¹⁹ We describe below our evaluation of the C10-ester derivatives as promising lead compounds for the development of effective TDase inhibitors as potential combination therapies with TC antibiotics.

Scheme 1. Synthesis of C10-Benzoate Ester and C9-Benzamide aTC Analogs.

Compound Stability and Cytotoxicity

To compare the toxicity of our C10-ester inhibitors to aTC which is known to display cytotoxicity,²⁰ we performed assays for cell viability using human renal proximal tubule epithelial cells (contracted through Eurofins, St. Louis, MO). Relative to the positive control staurosporine (IC₅₀ ∼ 0.1–0.3 μg/mL), none of the compounds showed significant toxicity below 30 μM (15 μg/mL which provides a viable therapeutic window based on the observed TC rescue at 2 μg/mL inhibitor (Figure S2 and Table S1). We also tested TC, aTC, and compounds 8, 10–15, and 19 for hemolysis in human red blood cells. No hemolysis was observed at concentrations up to 500 μM (∼250 μg/mL) (Figure S3). We considered the possibility that the C10-esters are prodrugs that release potentially toxic aTC upon hydrolysis, either esterase-catalyzed or nonenzymatic hydrolysis. We performed hydrolytic stability studies on compounds 7 and 8 in bacterial cell lysates (E. coli). The positive control, p-NO₂-phenyl acetate, caused a time- and lysate-dependent increase in A₄₀₅ as expected.²¹ Compounds 7 and 8 were stable in the cell lysate according to the LC–MS analysis and the appearance of free aTC was not observed (Figure S4). A p-NO₂-phenyl benzoate control compound was also used to model the stability of a general benzoate ester. Like benzoate esters 7 and 8, the p-NO₂-phenyl benzoate control was stable in the cell lysate, suggesting that unlike acetate esters the benzoate esters are not vulnerable to hydrolysis by bacterial esterases.

C10-Benzoate Inhibitors Bind TDases with Mixed Binding Modes

We initially explored the ability of the synthetic C10-esters and C9-amides to inhibit TetX7 and Tet50 in vitro using our standard NADPH/TC consumption assay.¹⁹ These experiments gave mixed results indicating that some of the compounds appear to be competitive inhibitors while others promote the consumption of NADPH making it difficult to extrapolate a reliable IC₅₀ value by monitoring for NADPH or TC consumption over time. We have observed that ligands including aTC and TC can bind to Type 1 and 2 TDases via multiple binding modes.^9,18 This sampling of binding modes may confound the in vitro optical absorbance assays resulting in a mixed interpretation of inhibition vs enzymatic turnover (consumption of NADPH and/or TC). Alternatively, overlapping absorbance spectra for the substrate, inhibitor, and oxidation products of the substrate and inhibitor could create interfering signals. To avoid these issues with optical absorbance-based assays and to determine whether our C10-esters protect TCs from TDase-catalyzed degradation, we monitored an in vitro TDase (TetX7) reaction by LC–MS to quantify the concentration of TC over time in the presence or absence of C10-ester 11 as a representative inhibitor (Figure S5). We observed that increasing concentrations of C10-ester 11 reduced the rate of TC oxidation over time suggesting that C10-ester 11 is a dose dependent inhibitor of TetX7.

We next investigated the mechanism of TDase inhibition by C10-esters 3–14 using optical absorbance to monitor the consumption of NADPH (λ_max = 340 nm) and the inhibitor (λ_max = 450 nm). We also tested the reactions for peroxide formation to determine whether the FAD is reduced by NADPH, leading to the formation of a peroxyflavin intermediate that decomposes to release hydrogen peroxide in an uncoupled reaction (Figures 2 and S6–S8). These in vitro assays support that the heterocycle-containing inhibitors 7–14 stimulate time-dependent NADPH consumption and hydrogen peroxide production, while the substituted benzene derivatives 3–6 appear to inhibit NADPH consumption similarly to the previously reported C9-aTC derivatives.¹⁹ We recently reported that aTC increases the rate of NADPH consumption in a dose dependent manner for Tet(X7) (K_app = 2.1 ± 0.3 μM, V_max = 0.043 ± 0.002 min^–1) and Tet(50) (K_app = 2.0 ± 0.3 μM, V_max = 0.24 ± 0.01 min^–1).¹⁹ Similarly, we analyzed the apparent steady-state kinetics of NADPH consumption for TetX7 and Tet50 in the presence of increasing concentrations inhibitors 7 and 8 by monitoring optical absorbance at 340 nm. These results show a similar dose-dependent increase in NADPH consumption for TetX7 + 7 (K_app = 26 ± 11 μM, V_max = 0.03 ± 0.01 min^–1), Tet50 + 7 (K_app = 37 ± 5 μM, V_max = 0.19 ± 0.01 min^–1), TetX7 + 8 (K_app = 110 ± 60 μM, V_max = 0.05 ± 0.02 min^–1), and Tet50 + 8 (K_app = 67 ± 22 μM, V_max = 0.13 ± 0.02 min^–1) (Figure S9). However, the A₄₅₀ did not change significantly for any of the inhibitors tested, indicating that the core aTC scaffold of the C10-esters is not being oxidized by TetX7 or Tet50. Additionally, we did observe the clear formation of a [M+O+H]⁺ oxidation product derived from inhibitor 9 which was detected via LC–MS analysis of the reaction mixture. This could be due to oxidation of the C10-ester aryl side chain which preserves the conjugated chromophore of the aTC derivatives responsible for the absorbance at 450 nm (Figure 2). Taken together, these results support that the C10-benzoate ester derivatives of aTC bind TDases with low μM affinity and inhibit TDase activity through a competitive binding mode for the TC and/or NADPH binding sites depending on the identity of the aryl ester side chain. The C10-esters could act as competitive inhibitors where oxidation of the aryl side chain protects the aTC core and allows the oxidation product to retain binding affinity for TDases. Similarly, aTC has been shown to undergo slow oxidation, presumably at C11a, by Type I TDases including TetX7, yet aTC is still effective at rescuing TC activity in TDase-expressing cells either because the aTC oxidation product retains inhibition or because of favorable differences in the K_m/k_cat of aTC compared to TC toward TDases. The mixed results among the C10-ester inhibitors related to stimulating or slowing NADPH consumption might be caused by a sampling of different inhibitor binding modes, which we have shown can influence the positioning of the dynamic FAD cofactor either into the active site (active state) or exposed to solvent outside of the active site (inactive state).¹⁹

Figure 2.

In vitro evaluation of inhibitor 9 against TetX7 and Tet50 for NADPH and inhibitor consumption. (A,B) shows optical absorbance over time (minutes) after treatment with inhibitor 9. (C,D) Top panels show molecule ion counts over time for the inhibitor [M + H]⁺ and the corresponding oxidation product [M+16]⁺ supporting the rapid oxidation of the inhibitor scaffold. Bottom panels show the production of hydrogen peroxide over time using a colorimetric assay.

Cocrystal Structure Shows C10-Ester Inhibitor in the Substrate Binding Mode

To explore the inhibitor binding mode, we solved the structure of TetX7 cocrystallized with inhibitor 13 at 3.3 Å (Figure 3A and Table S2). The inhibitor-bound TetX7 structure was similar to the inhibitor-free TetX7 structure PDB: 6WG9(12) with a root-mean-square deviation (r.m.s.d.) of 0.35 Å (Figure S10). After modeling TetX7 and FAD in the electron density map, a large piece of unmodeled positive density was found next to the isoalloxazine group of the bound FAD cofactor (Figure 3B). Inhibitor 13 was therefore modeled in with the A-ring component of inhibitor 13 pointing into the substrate-binding pocket and the D-ring component of inhibitor 13 pointing outside, providing space to accommodate the C10-ester present on the D-ring. The aTC component of inhibitor 13 fit well into the map, but no electron density was observed on the C10-ester aryl side chain after refinement, suggesting flexibility. The binding mode of the bound inhibitor 13 was similar to that of aTC bound to TetX6 PDB: 8ER0 (Figure 3C).¹⁸

Figure 3.

Structure of TetX7 in complex with inhibitor 13 (PDB 9DRH). (A) Overall view of TetX7 structure bound with FAD and inhibitor 13. Tet(X7) is represented in ribbon and ligands are showed as stick model with carbon in gray (FAD) and yellow (inhibitor 13), oxygen in red, nitrogen in blue, phosphorus in orange. (B) Inhibitor 13 molecule is enclosed in a difference Fo – Fc electron density cage (green) with 2.5 σ level. (C) Substrate binding site of Tet(X7). Structure of Tet(X7)-inhibitor 13 was superposed on aTC:TetX6 PDB: 8ER0 (in dark gray color).

Both aTC and analog 13 can be oxidized by TetX7 while maintaining the inhibition of TC oxidation. Based on LC–MS and optical absorbance spectra, we believe that aTC is oxidized at C11a, whereas analog 13 is oxidized at a position that does not disrupt the conjugation of the pi-electron system in the core of the aTC scaffold (Figures 2 and S6–S8). The core aTC scaffold of inhibitor 13 forms contacts with the conserved active site residues Q203 and R224, but the C10-pyrimidine-2-carboxylate side chain was too dynamic for observation in the X-ray crystal structure (Figure 3C). Despite the proven stability of the C10-esters toward various buffers and cell lysates (Figure S4), we cannot rule out dynamic in crystallo hydrolysis and/or selective trapping of the hydrolysis product aTC during the cocrystallization process. Therefore, we used molecular docking with AutoDock Vina to model the aryl ester side chain to show that there is ample space in the open TetX7 active site to accommodate this group in multiple conformations. In this binding conformation, the C10-benzoate side chain is too far away from the FAD C4a position, while the two known TC oxidation sites, C11a and C1, are still within striking distance of an FAD C4a-peroxy group. In the aTC:TetX6 structure the distances are 5.7 Å for C11a-C4a and 6.0 Å for C1–C4a. In the new structure of inhibitor 13:TetX7 reported here the distances are 5.3 Å for C11a–C4a and 6.7 Å for C1–C4a. It is noteworthy that the C11a position in the two structures are offset by 1.1 Å, and the C10 atom in the analog 13:TetX7 structure overlays best with the C11a position in the aTC:TetX6 structure. Further, the C10-ester is an electron withdrawing group and may act to decrease the nucleophilicity of the conjugated C11a enol carbon, increasing the transition state energy for C11a hydroxylation by a C4a-peroxy flavin. This subtle shift in conformation and redox potential may account for the observation that aTC, but not the C10-ester aTC analogs, are oxidized at C11a by TetX7 and other Type 1 TDases. The crystal structure of inhibitor 13 bound to TetX7 suggests that the C10-benzoate ester derivatives of aTC still bind tightly to the Type 1 TDase active site and accommodate a wide variety of C10 ester substituents because the placement of this side chain pointing into the spacious opening of the solvent exposed active site.

Tetracycline Rescue in Resistant E. coli, P. aeruginosa, and M. smegmatis

We next evaluated the ability of the compounds to rescue TC antibacterial activity against E. coli cells expressing plasmid-encoded TetX7 or Tet50. The measured MIC value of TC against E. coli expressing TetX7 and Tet50 is >8 μg/mL,⁹ so we performed combined MIC studies using variable inhibitor concentrations with a fixed TC concentration of 8 μg/mL to screen for inhibitors that could potentially have synergistic effects with TC against TetX7 and Tet50. Hence, the recorded MIC values reflect the concentration of inhibitor needed to rescue TC antibacterial activity at 8 μg/mL. We first tested for the inherent antibacterial activity of the inhibitors by performing standard MICs on the inhibitor series. We found that inhibitors 2–6, 8, and 16–23 displayed no growth inhibitory activity alone at concentrations >128 μg/mL. Inhibitors 7 and 9–15 inhibited growth to some extent, with MIC values ranging from 8 to 64 μg/mL. The inhibitors rescued the activity of TC to varying degrees, with MICs as low as 2 μg/mL for compounds 7, 10, 11, and 15 when combined with a fixed concentration of TC at 8 μg/mL (Table 1). These data are consistent with TDase inhibitors rescuing TC activity, because even inhibitors that lack inherent antibacterial activity (compound 8, combined MIC = 4 μg/mL) have similar rescue activity as inhibitors that are antibacterial (compound 7, combined MIC = 2 μg/mL).

Table 1. Minimum Inhibitory Concentration (MIC in μg/mL) against E. coli DH5αZ1 Expressing TetX7 or Tet50 from pZE24 Plasmid in the Absence or Presence of 8 μg/mL TC.

In our previous work, we used growth rate analysis to reveal synergy between C9-benzamide aTC derivatives and TC against E. coli expressing TetX7 or Tet50.¹⁹ The growth rate analysis enabled us to observe synergy even when growth was only partially inhibited. We performed this type of growth rate analysis for the C10-ester aTC derivatives 3 and 6 and observed complete growth inhibition of E. coli at 128 μg/mL inhibitor and 8 μg/mL TC (Figure S11). Next, we used checkerboard MIC assays to determine the fractional inhibitory concentration index (FICI) against strains of E. coli, P. aeruginosa, and M. smegmatis.²² We selected three inhibitors representing compounds with or without inherent antibacterial activity against E. coli (Table 1): antibacterial aTC (1), antibacterial C10-ester (7), and nonantibacterial C10-ester (8). Interestingly, the only structural difference between inhibitors 7 and 8 is the position of the nitrogen in the pyridine ring of the aryl acid side chain. We determined FICI scores for inhibitors 1, 7, and 8 in combination with TC or Tig against a panel of bacteria representing laboratory strains (E. coli DH5αZ1; M. smegmatis mc²155) and clinical isolates (P. aeruginosa PA27853 and PA-3)¹² with varying degrees and mechanisms of TC resistance (Table 2; Figure 4; see Tables S3–S5 for information on strains). aTC (1) was synergistic with TC and Tig against P. aeruginosa PA-3 (FICI ≤ 0.5) and was indifferent or additive against all other strains tested. Inhibitor 7 was synergistic with Tig against P. aeruginosa PA-3 (FICI ≤ 0.5) and was indifferent or additive against all other strains tested. Inhibitor 8 was synergistic with E. coli DH5αZ1 expressing TetX7 (FICI = 0.31–0.56) or Tet50 (FICI = 0.265–0.625) and was indifferent or additive against all other strains tested. Inhibitor 8 had a borderline synergistic effect with Tig against P. aeruginosa PA-3, but since this inhibitor lacks inherent antibacterial activity, we were not able to distinguish the FICI = 0.5 as a reflection of compound synergy. All three inhibitors produced additive FICI values ≤1 with borderline synergy in some cases against M. smegmatis mc²155 expressing TetX as a fusion with hemagglutinin (Figure S12), although the visual scoring of the growth phenotypes showed synergy when partial growth inhibition was considered (Figure S13).

Table 2. Fractional Inhibitory Concentration Index (FICI) of aTC, Inhibitor 7, and Inhibitor 8 against E. coli, P. aeruginosa, and M. smegmatis.

strain	drug combination	FICI	effect	fold increase in TC MIC
E. coli DH5αZ1+pZE24	tetracycline – inhibitor 7	1.125	indifferent	NA
	tetracycline – inhibitor 8	1	indifferent	NA
E. coli DH5αZ1+pZE24-TetX7	tetracycline – inhibitor 7	0.625	additive	2
	tetracycline – inhibitor 8	0.375	synergetic	8
E. coli DH5αZ1+pZE24-Tet50	tetracycline – inhibitor 7	0.625	additive	2
	tetracycline – inhibitor 8	0.265	synergetic	4
E. coli DH5αZ1	tetracycline–aTC	>1	indifferent	NA
	tetracycline – inhibitor 7	0.75	additive	4
	tetracycline – inhibitor 8	>1	indifferent	NA
	tigecycline–aTC	>1	indifferent	NA
	tigecycline – inhibitor 7	>1	indifferent	NA
	tigecycline – inhibitor 8	>1	indifferent	NA
P. aeruginosa PA 27853	tetracycline–aTC	0.75	additive	2
	tetracycline – inhibitor 7	>1	indifferent	NA
	tetracycline – inhibitor 8	>1	indifferent	NA
	tigecycline–aTC	0.75	additive	4
	tigecycline – inhibitor 7	>1	indifferent	NA
	tigecycline – inhibitor 8	>1	indifferent	NA
P. aeruginosaPa3̅	tetracycline–aTC	0.375	synergetic	4
	tetracycline – inhibitor 7	1	indifferent	NA
	tetracycline – inhibitor 8	>1	indifferent	NA
	tigecycline–aTC	0.375	synergetic	4
	tigecycline – inhibitor 7	0.5	synergetic	4
	tigecycline – inhibitor 8	0.507	additive	2
M. smegmatis mc2155	tetracycline–aTC	1	indifferent	2
	tetracycline – inhibitor 7	0.75–1	additive/indifferent	2
	tetracycline – inhibitor 8	0.75–1	additive/indifferent	2

Figure 4.

Fractional Inhibitory Concentration Index (FICI) of aTC, inhibitor 7, and inhibitor 8 against E. coli, P. aeruginosa, and M. smegmatis. Graph depicts the plotted MIC values for inhibitor concentration (μg/mL; y-axis) versus tetracycline or tigecycline concentration (μg/mL; x-axis). Triangles represent synergy. Each MIC value was determined from three independent biological replicates.

Conclusions

We synthesized and characterized inhibitors of TDase enzymes via the fusion of a known inhibitor, aTC, and structural mimics of nicotinic acid joined via ester linkages to the C10-phenolate on the D-ring of aTC. While many semisynthetic modifications of tetracyclines have been reported,^1,23,24 esterification of C10 has not been explored in depth. We found only one report of C10 esterification of tetracyclines within the patent literature.²⁵ This is not surprising given that the C10 phenol group is essential for tetracycline Mg²⁺ chelation and ribosome binding, so there is little motivation to explore SAR at this position for the purpose of inhibiting protein translation.²⁶ For this reason, we do not associate the inherent antibacterial activity of our C10-ester and C9-amide aTC analogs to the inhibition of protein synthesis. aTC is known to disrupt bacterial cell membranes,¹⁷ so it is possible that this mechanism contributes to the observed antibacterial activity and ability of the C10-ester analogs to permeate the bacterial cell envelope and inhibit intracellular TDases. Given that there is growing exploration in the use of TC derivatives in other therapeutic areas, including inflammatory diseases,^27,28 our simple synthetic method for direct C10 esterification could be broadly applicable in analog production aimed at these drug development efforts.

Our prior work on aTC-based TDase inhibitors demonstrated that C9-benzamides and C9-benzylamines are potent bivalent TDase inhibitors that compete with both TC and NADPH for TDase binding.¹⁹ We designed C10-benzoate ester aTC analogs with this potential for bivalent binding but discovered that these analogs show mixed competitive inhibition with respect to TC and NADPH depending on the nature of the aryl ester side chain. The analog 13:TetX7 cocrystal structure showed that the C10-benzoate ester inhibitor occupies the substrate binding mode. The C10-benzoate side chain is disordered, and the ligand orientation indicates free rotation of this group in the solvent exposed region of the large opening to the active site cavity. Based on molecular modeling, the placement of benzamides at the C9 position of the aTC D-ring allows for the orientation of the D-ring into the active site to interact with the FAD in the “out” conformation.¹⁹ However, at the C10-position, the benzoate ester moiety is sterically occluded from accessing this binding orientation and forces the C10-ester aTC analogs into the substrate binding mode. In this orientation, a wide range of C10-functionalization should be tolerated along with added substituents at the C8 and C9 positions of the aTC D-ring.

Knowing that the inhibitor mechanism and TDase binding mode can guide structure-based drug design, we ultimately want to drive inhibitor optimization by whole cell efficacy toward the rescue of TC growth inhibitory activity against TDase expressing pathogens. This is the most exciting aspect of the C10-ester aTC analogs, which are more active than C9-benzamides bearing the same aryl acid side chain. We attribute this improved activity to increased water solubility, cell permeability, and inhibition of TDases. The most distinct SAR pattern to emerge was that C10-ester derivatives derived from N-containing heterocyclic aryl acids were the most active compounds in the series. In total, compared to 3/11 C9-benzamides, 11/14 C10-benzoates were active in rescuing TC antibacterial activity based on MIC determination (Table 1). The most potent 9/11 C10-benzoates were derived from N-containing pyridine, pyrimidine, or pyrazine aryl acids. Within the pyridine series, the picolinic and isonicotinic acid analogs 7 and 9, respectively, both showed inherent antibacterial activity while nicotinic acid analog 8 was able to rescue TC activity against both Type 1 and 2 TDases without showing any inherent antibacterial activity; the latter is a true reflection of compound synergy via TDase inhibition. It is noteworthy that both picolinic and isonicotinic acid alone possess antimicrobial activity while nicotinic acid is nontoxic to all cell types.²⁹⁻³¹ Picolinic acid is a natural metabolite derived from the catabolism of tryptophan, a metal chelator, and a membrane uncoupler.²⁹ Isonicotinic acid is the core scaffold of the anti-TB drug isoniazid, which inhibits the enoyl-acyl carrier protein reductase InhA resulting in reduced fatty acid biosynthesis.³² The hydrolysis of picolinic acid esters is promoted by the presence of divalent metal cations such as Co²⁺, Ni²⁺, Zn²⁺, and Cu²⁺,³³ suggesting that the enhanced antibacterial activity of picolinate ester 7 (E. coli MIC = 16 μg/mL) relative to isonicotinate ester 9 (E. coli MIC = 64 μg/mL) may be due to metal cation-promoted intracellular hydrolysis; however, we note that both inhibitors 7 and 8 were stable toward hydrolysis in E. coli cell lysates (Figure S4).

It remains unclear if it is more advantageous to employ antibacterial or nonantibacterial TDase inhibitors since both inhibitors 7 and 8 are capable acting synergistically with TC antibiotics against tetracycline-resistant bacteria. We note that for beta-lactam antibiotics, beta-lactamase inhibitors are typically nonantibacterial and hence do not apply a separate selection force for resistance.³⁴ We also recognize that certain combinations of antibacterial agents can reduce the occurrence of spontaneous resistance.³⁵ All of the pyrimidine and pyrazine-derived esters 10–15 were antibacterial alone against E. coli and strongly rescued TC activity (Table 1). Pyrimidine and pyrazine carboxylic acids are known inhibitors of fatty acid synthesis.³⁶ Further, pyrazinamide is a prodrug for pyrazinoic acid used to treat TB.³⁷ We note that pyrimidine and pyrazine-derived esters 10–15 were generally less cytotoxic in the human renal proximal tubule epithelial cell viability assay (Figure S2; Table S1). The heterocyclic aryl acid side chain might provide a means to systematically improve TDase binding affinities and redox potentials by exploring pi-pi stabilization energies with the FAD isoalloxazine ring, which has been rigorously explored in FAD-binding RNA aptamers and synthetic flavin model systems.^38,39 While other TDase inhibitor types have been reported,⁴⁰⁻⁴² we favor aTC based inhibitors due to the proven in vivo efficacy of this general pharmacophore⁴³ and the ease of analog synthesis through our high-yield one step C10 esterification protocol. While further mechanistic studies and optimization of the C10-ester series are merited, this study supports that C10-ester derivatives of aTC are attractive water-soluble candidates for combination therapy with TC antibiotics to overcome TDase-mediated resistance.

Experimental Procedures

Synthesis of C10-Benzoate Ester aTC Derivatives (Compounds 2–15)

The corresponding benzoic acid (0.0199 g, 0.162 mmol, 1.00 equiv) and DIPEA (0.045 mL, 0.260 mmol, 1.60 equiv) were dissolved in 3.5 mL of anhydrous DMF (DriSolv) in a vial and charged with HATU (0.0617 g, 0.162 mmol, 1.00 equiv). The clear solution turned pale yellow and was stirred at room temperature for 10 min before the addition of compound aTC (HCl salt). The reaction solution was stirred for 5 min at room temperature and concentrated under reduced pressure via rotary evaporation. The residue was dissolved in MeOH, filtered through a 0.45 μm PTFE syringe filter, and purified by RP-C18 prep-HPLC to provide the desired products 2–15 as the corresponding formic acid salts. Compound purity, including epimeric ratio, was assessed by LCMS and NMR as reported in the Supporting Information. All compounds tested in biological assays were determined to be >95% pure from detectable organic contaminants as a reported ratio of C4-epimers. See the Supporting Information for synthetic procedures and compound characterization data. All compounds were fully soluble at concentrations tested.

Synthesis of C9-Benzamide-aTC Derivatives (Compounds 16–23)

C9-substituted aTC derivatives were synthesized following a general protocol previously described by our group.¹⁹ Compound purity, including epimeric ratio, was assessed by LCMS and NMR as reported in the Supporting Information. All compounds tested in biological assays were determined to be >95% pure from detectable organic contaminants as a reported ratio of C4-epimers. See the Supporting Information for synthetic procedures and compound characterization data. All compounds were fully soluble at concentrations tested.

Cloning, Expression, and Purification of TDases

All genes encoding TDases used in this study were cloned and inserted into pET28b(+) vectors (Novagen) as previously described (BamHI and NdeI restriction sites) and transformed into BL21-Star (DE3) competent cells (Life Technologies).¹² The cells were cultured at 37 °C in lysogeny broth (LB) containing kanamycin (Kan) at 0.05 mg/mL (final concentration); once the culture reached an OD600 of ∼0.6, the cells were cooled to 0 °C in an ice water bath. Protein expression was induced by the addition of 1 mM IPTG (final concentration), and the cells were grown at 15 °C for 12–15 h. To harvest protein, the induced cells were pelleted by centrifugation at 4000 rpm for 15 min (4 °C) and resuspended in cold 40 mL of lysis buffer (50 mM K₂HPO₄, 500 mM NaCl, 20 mM imidazole, 10% glycerol, 5 mM BME, pH 8.0) containing SIGMAFAST© protease inhibitor (Millipore-Sigma). The cell suspensions were flash frozen in liquid nitrogen and stored at–80 °C. The frozen cell suspensions were thawed and mechanically lysed using an Avestin EmulsiFlex-C5 cell disruptor, and the resulting lysate was clarified via ultracentrifugation at 45,000 rpm for 35 min at 4 °C. The clarified supernatant was transferred to a fritted column containing washed and equilibrated Ni-NTA resin and incubated for 30–45 min with gentle rocking. The resin was then washed with lysis buffer (2 × 40 mL), and the protein was eluted from the resin with elution buffer (5 × 10 mL elutions, 50 mM K₂HPO₄, 500 mM NaCl, 5 mM BME, 300 mM imidazole, 10% glycerol, pH 8.0). Fractions containing the desired proteins (as determined by SDS-PAGE analysis) were combined, and transferred to a 10,000 molecular weight cutoff (MWCO) Snakeskin dialysis tubing (ThermoScientific) and equilibrated in dialysis buffer (50 mM K₂HPO₄ pH 8.0, 150 mM NaCl, 1 mM DTT) overnight. The dialyzed protein solutions were concentrated using a 30,000 MWCO Amicon centrifugal filter (Millipore-Sigma), and the concentrated protein solution was flash frozen as beads in liquid nitrogen (50 μL portions) and stored at −80 °C.

In Vitro Characterization of TDases

TDase reactions were prepared in 100 mM TAPS buffer (pH 8.5) with 252 μM NADPH and 5.04 mM MgCl₂, 40 μM substrate and 0.4 μM TDase enzyme (TetX7 and Tet50) (all concentrations represent final working concentrations). Reaction progress was monitored by optical absorbance spectroscopy (280–550 nm, 1 nm and 5 min intervals) over 2 h. Twenty μL aliquot of of reaction mixture was removed at 0, 5, 30, 60, 90, and 120 min time points and used for colorimetric detection of hydrogen peroxide formation performed using an aqueous Pierce Quantitative Peroxide Assay Kit (ThermoScientific). Each 20 μL aliquot (performed in triplicate on three separate aliquots for each time point) was added to a 96-well plate containing 200 μL of working reagent (prepared according to specifications for Pierce Quantitative Peroxide Assay kit). The plate was incubated for at least 20 min at room temp and observed for color change.

Cytotoxicity

All cytotoxicity studies were performed by Eurofins Panlabs (St. Charles, MO). Cell viability of Human Renal Proximal Tubule Epithelial Cells was determined using CellTiter-Glo after 48 h incubation at 37 °C.⁴⁴ The percent of control was calculated using the formula: Control (%) = (compound/T1)*100 where “compound” is the individual reading in the presence of the test compound and T1 is the mean reading in the absence of the test compound. The percent of inhibition is calculated by subtracting the percent of control from 100. The IC₅₀ value (concentration causing a half-maximal inhibition of the control value) was determined by nonlinear regression analysis of the concentration–response curve using the Hill equation. The positive control was staurosporine and all tests were performed in duplicate as independent trials.

Measuring Growth Rate

Overnight cultures were grown in fresh MH-II + KAN50 broth supplemented with 1 mM IPTG to exponential phase (OD600 = 0.3–0.8), then diluted to OD600 = 0.1, and inoculated into each 96-well panel at a 1:1 ratio. To avoid edge effects only the interior wells included cells, and the exterior wells were filled with cell-free broth. Plates were sealed with Breathe-Easy membranes (Sigma-Aldrich) then incubated at 37 °C with continuous shaking and OD600 measurements taken every 5 min for 20 h using a Synergy H1 plate reader (BioTek). Maximal growth rate was calculated from this plate reader data using GrowthRateR (https://github.com/kevinsblake/GrowthRateR). This function log-transforms growth curves and generates a rolling regression with a shifting window of 1 h, such that the maximum slope of any of the regressions is the maximal growth rate.

Antibiotic Susceptibility Testing – Minimum Inhibitory Concentration (MIC)

Inhibitor screening by antibiotic susceptibility tests (AST) were performed with the microbroth dilution method, following CLSI guidelines⁴⁵ and as previously described.^19,46 MIC panels were prepared in 96-well flat-bottom microplates (Corning) by 2-fold serial dilution of inhibitors in cation-adjusted MH-II broth (BD) supplemented with 50 μg/mL kanamycin (KAN50), 1 mM IPTG, and with or without 16 μg/mL of tetracycline (TC). The panels were stored at −80 °C before use. Single colonies of E. coli DH5αZ1 containing TetX7, Tet50, or empty vector in pZE24 expression system were grown in MH-II broth with KAN50 overnight at 37 °C. On the day of the experiment, MIC panels were thawed at room temperature. Overnight cultures were subcultured in fresh MH-II broth with KAN50 and 1 mM IPTG, grown to exponential phase (OD600 of 0.3–0.8), then diluted in MH-II broth with KAN50 and 1 mM IPTG. The diluted cells were inoculated into the MIC panel at a 1:1 ratio, resulting in a final concentration of ∼5 × 10⁵ CFU/mL cells in each well. The panels were incubated at 37 °C for 20 h, then scored by visual inspection. Potential synergistic effects between inhibitors and TC were evaluated by comparing the MIC of inhibitors alone to the MIC of inhibitors in combination with TC. Each test was performed in triplicate, with no-antibiotic and no-cell control wells.

Antibiotic Susceptibility Testing – Fractional Inhibitory Concentration (FICI) Index

The synergistic effects and fractional inhibitory concentration index (FICI) between tetracycline family antibiotics and inhibitors were assessed using a checkerboard assay.²² FIC panels were prepared in 96-well flat-bottom microplates by performing 2-fold serial dilutions of tetracyclines and inhibitors in cation-adjusted MH-II broth, supplemented with or without KAN50 and 1 mM IPTG. Column 1 and row H contained only the inhibitors or tetracyclines, respectively. The procedures described in the MIC section were followed to prepare and inoculate cells into each well. For E. coli DH5αZ1 strains with TetX7 and Tet50, FIC panels included 1 mM IPTG and KAN50. For Pa3̅ and PA27853, FIC panels were prepared without 1 mM IPTG and KAN50. After cell inoculation, the panels were incubated at 37 °C for 20 h and then scored by visual inspection. FICIs were calculated using the formula: FICI = (A/MIC_A) + (B/MIC_B), where A and B represent the MICs of drugs tested in combination. MIC_A and MIC_B represent the MICs of drugs when tested alone. Each test was repeated in triplicate and included controls.

Bacterial Strains, Plasmids, and Oligonucleotides Used

All bacterial strains, plasmids, and oligonucleotides used in this study are listed in Tables S3–S5. The E. coli strains utilized in this study were same as the ones previously described.⁴⁶ In brief, TetX7 and Tet50 sequences were cloned into the KpnI and MluI sites of the pZE24 plasmid (Expressys) and transformed into E. coli DH5αZ1 (Expressys). The Pseudomonas aeruginosa strains Pa3̅ and ATCC 27853 were the same as those described in a previous study,¹² with Pa3̅ containing chromosomal TetX7 genes and ATCC 27853 serving as the P. aeruginosa control lacking TetX7.

Production of M. smegmatis TetX^T249K-HA Expressing Cells

B. fragilis TDase TetX with a C-terminal HA tag (TetX-HA) gene was cloned into the pCLS16 vector using InFusion master mix (see Tables S4 and S5) and transformed into E. coli DH5α cells. Sequence of vector was confirmed using Plasmidsaurus sequencing. TetX-HA pCLS16 vector or empty pCLS16 vector were then transformed into M. smegmatis mc²155 cells and expression of gene was confirmed via HA Western blot (Figure S12). E. coli strains were grown at 37 °C in LB broth supplemented with antibiotics when necessary. M. smegmatis strains were grown at 37 °C in LB broth supplemented with 0.5% dextrose, 0.5% glycerol, and 0.05% Tween-80, and supplemented with antibiotics when necessary (see Table S3).

Cloning, Expression, and Purification of TetX for Crystallization

The sequence of TetX7 (GenBank CP025402.1) was cloned and inserted into pET28 with 6His tagged to the N-terminus.¹² The plasmid was transformed into BL21(DE3) (ThermoFisher) and the cells were grown at 37 °C in LB broth supplemented with 50 μg/mL kanamycin. Expression was induced at OD600 ∼1 with a final concentration of 1 mM IPTG. The cells were harvested 4 h after induction at 37 °C. The cell pellet was resuspended in 25 mM Tris, pH 7.4, 0.3 M NaCl, 5 mM DTT and supplemented with Pierce protease inhibitor cocktail (ThermoFisher) and DNase I. The cell suspension was then lysed by sonication for 5 min on ice. Cell debris was removed by centrifugation at 39,000×g for 20 min at 4 °C. The supernatant was loaded into Excel Ni-NTA (Cytiva) equilibrated with 25 mM Tris, pH 7.4, 0.3 M NaCl, 5 mM DTT, 30 mM imidazole. The column was then washed with the same buffer. TDase protein was eluted using 25 mM Tris, pH 7.4, 0.3 M NaCl, 5 mM DTT, 150 mM imidazole. Fractions were concentrated and injected into a Superdex 75 Increase column (Cytiva) equilibrated with 20 mM Tris, pH 8.0, 100 mM NaCl, 5 mM DTT for size exclusion chromatography. The fractions were again concentrated and stored at −80 °C.

Crystallization, Data Collection, and Structure Determination of the TetX7-Inhibitor 13 Complex

The protein of TetX7 (25 μL of 12 mg/mL) was mixed with 1 μL of inhibitor 13 (100 mM in DMSO) and incubated on ice for 30 min before setting up crystallization trays. Crystals of TetX7-inhibitor 13 were grown using hanging-drop vapor diffusion at 18 °C. The complex mixture (12 mg/mL) was mixed with 0.1 M HEPES, pH 7.5, 0.2 M ammonium sulfate, 16% PEG 4000 and 10% isopropanol at a 1:1 ratio for crystallization. The crystals were cryoprotected in a well solution supplemented with 30% PEG 400 prior to flash freezing in liquid nitrogen. X-ray diffraction experiments were carried out at 100 K at the ALS 5.0.2 beamline at the Advanced Light Source at Lawrence Berkeley National Laboratory. The diffraction data were processed with XDS.⁴⁷ The structure was determined by molecular replacement in PHASER⁴⁸ using TetX7 PDB: 6WG9 as the search model.¹² Iterative model building using COOT,⁴⁹ and refinement collection and refinement using PHENIX⁵⁰ led to the current model for (Rwork/Rfree of 26.23/29.46). The data collection and refinement statistics are shown in Table S2. The structure has been deposited in the Protein Data Bank under accession code 9DRH.

Glossary

Abbreviations

aTC

anhydrotetracycline

FAD

flavin adenine dinucleotide oxidized form

FMO

flavin monooxygenase

KMO

kynurenine-3-monooxygenase

NADPH

nicotinamide adenine dinucleotide phosphate reduced form

NADP⁺

nicotinamide adenine dinucleotide phosphate oxidized form

TC

tetracycline

TDase

tetracycline destructase

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsinfecdis.4c00912.

• Experimental methods, synthetic procedures, supplementary figures, supplementary tables, plasmid sequences, NMR spectra, and LC–MS chromatograms (PDF)

Author Contributions

E.E.W., T.A.W., G.D., and N.H.T. conceived and designed the experiments in this study. E.E.W., K.V.J., and X.L. synthesized all TDase inhibitors. E.E.W., K.V.J., and R.L. performed all in vitro TDase assays. Y.P.X., E.E.W., and K.S.B. performed all whole cell studies. W.KT. crystallized and collected X-ray data. H.B. and C.L.S. designed and constructed M. smegmatis strains expressing TetX-HA. E.E.W. and T.A.W. wrote the manuscript, created figures, and compiled the Supporting Information documents with input and approval from all authors.

This work is supported in part by the National Institute of Allergy and Infectious Diseases (NIAID) of the National Institutes of Health (NIH) through grant 2U01AI123394 awarded to G.D. and T.A.W. at Washington University in St. Louis. N.H.T. is supported by the Intramural Research Program of the NIAID of the NIH. K.S.B. is supported by the National Institute of Diabetes and Digestive and Kidney Diseases (T32-DK007130; PI: N. Davidson). E.E.W. is supported by the National Science Foundation through Graduate Research Fellowship (DGE-2139839). High Resolution Mass Spectrometry data collection was supported by NIH grant 8P41GM103422a. The content is solely the responsibility of the authors and does not necessarily represent the official views of the funding agencies. Access to ALS beamlines was provided through South East Regional Collaborative Access Team (SER-CAT). SER-CAT is supported by its member institutions, equipment grants (S10_RR25528, S10_RR028976, and S10_OD027000) from the National Institutes of Health, and funding from the Georgia Research Alliance. The Berkeley Center for Structural Biology is supported by the Howard Hughes Medical Institute, Participating Research Team members, and the National Institutes of Health, National Institute of General Medical Sciences, ALS-ENABLE grant P30 GM124169. The Advanced Light Source (ALS) is a Department of Energy Office of Science User Facility under Contract No. DE-AC02–05CH11231. This work was also supported by the Intramural Research Program of the Division of Intramural Research, National Institute of Allergy and Infectious Diseases, National Institutes of Health. This study used the Office of Cyber Infrastructure and Computational Biology (OCICB) High Performance Computing (HPC) cluster at the National Institute of Allergy and Infectious Diseases (NIAID), Bethesda, MD.

The authors declare the following competing financial interest(s): A provisional patent application (63/687,889) describing these inhibitors has been filed through the Washington University in St. Louis Office of Technology Management.