Semisynthetic analogues of anhydrotetracycline as inhibitors of tetracycline destructase enzymes

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Semisynthetic Analogues of Anhydrotetracycline as Inhibitors of

Tetracycline Destructase Enzymes

Jana L. Markley,^†Luting Fang,^†Andrew J. Gasparrini,^‡Chanez T. Symister,^†Hirdesh Kumar,^§

Niraj H. Tolia,*^,§,∇Gautam Dantas,*^{,‡,∥,⊥,#,∇}and Timothy A. Wencewicz*^,†,∇

^†Department of Chemistry and ^#Department of Biomedical Engineering, Washington University in St. Louis, One Brookings Drive,

St. Louis, Missouri 63130, United States

^‡The Edison Family Center for Genome Sciences & Systems Biology, Washington University School of Medicine, 4513 Clayton

Ave., Campus Box 8510, St. Louis, Missouri 63108, United States

^§Laboratory of Malaria Immunology and Vaccinology, National Institute of Allergy and Infectious Diseases, National Institutes of

Health, 9000 Rockville Pike, BG 29B Rm 4NN08, Bethesda, Maryland 20814, United States

^∥Department of Pathology and Immunology, Washington University School of Medicine, St. Louis, Missouri 63110, United States

^⊥Department of Molecular Microbiology, Washington University School of Medicine, 4515 McKinley Avenue, ﬁfth Floor, Room

5314, St. Louis, Missouri 63110, United States

S

* Supporting Information

ABSTRACT: The synthesis and biological evaluation of semi-

synthetic anhydrotetracycline analogues as small molecule

inhibitors of tetracycline-inactivating enzymes are reported.

Inhibitor potency was found to vary as a function of enzyme

(major) and substrate-inhibitor pair (minor), and anhydrotetracy-

cline analogue stability to enzymatic and nonenzymatic degrada-

tion in solution contributes to their ability to rescue tetracycline

activity in whole cell Escherichia coli expressing tetracycline

destructase enzymes. Taken collectively, these results provide the

framework for the rational design of next-generation inhibitor

libraries en route to a viable and proactive adjuvant approach to

combat the enzymatic degradation of tetracycline antibiotics.

KEYWORDS: tetracyclines, anhydrotetracycline, tetracycline destructases, Tet(X), antibiotic resistance, inactivating enzymes,

enzymology, antibiotic adjuvants, tigecycline, eravacycline, omadacycline

facilitate intra- and extra-cellular antibiotic clearanceoften

ince the isolation of chlortetracycline (2, aureomycin)

1

S_{from Streptomyces aureofaciens in 1948, the tetracycline}

through the enzymatic, irreversible inactivation of antibiotic

scaﬀoldsfrequently pervade resistance landscapes as the

family of broad-spectrum antibiotics has served as essential

most eﬃcient means of achieving resistance.^24,25Historically,

medicines for the treatment of bacterial infections in hospital

and agricultural settings (Figure 1).²⁻⁸Driven by challenges

the enzymatic inactivation of β-lactam antibiotics has been

well-studied,²⁶⁻²⁸and strategies aimed at combatting this

with stability, toxicity, and rising antibiotic resistance, the

resistance using an adjuvant approachwhere the antibiotic is

development of more eﬀective, semisynthetic tetracycline

coadministered with a small molecule inhibitor of the

inactivating enzymehave emerged as fundamentally useful

tools for the rescue of β-lactam antibiotics in the clinic.²⁹⁻³²

With the discovery and characterization of 10 tetracycline-

inactivating enzymes with varying resistance proﬁles,^33,34the

development of small molecule inhibitors of tetracycline

destructase enzymes stands at the forefront of strategies

aimed at combatting the imminent clinical emergence of this

resistance mechanism in multidrug resistant infections. We

herein report preliminary ﬁndings focused on understanding

variants has led to the introduction of next-generation

tetracycline antibiotics tailored to overcome emerging

resistance mechanisms.⁹⁻¹²In this regard, the majority of

current treatment strategies employ the use of second-

generation C6-deoxy-tetracyclines (i.e., doxycycline and

minocycline), which were developed to overcome eﬄux and

stability issues,⁹and third-generation glycylcyclines (tigecy-

cline,^13,14eravacycline,^15,16and omadacycline¹⁷), which were

designed to evade eﬄux and ribosomal protection^9,18and are

used as last-resort treatments for multidrug resistant infections

(Figure 1).¹⁹⁻²¹While the most common, clinically relevant

resistance mechanisms for tetracycline antibiotics include eﬄux

and ribosomal protection,^9,22,23those mechanisms that

Received: December 9, 2018

A

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Figure 1. Tetracycline development and parallel emergence of resistance mechanisms.

Figure 2. Introduction to the tetracycline destructase family of FMO enzymes and structure of the ﬁrst inhibitor, anhydrotetracycline (5). (A)

Phylogenetic tree [aligned with Clustal Omega and viewed using iTOL software]. (B) X-ray crystal structure of chlortetracycline bound to Tet(X)

(PDB ID 2y6r). (C) X-ray crystal structure of chlortetracycline bound to Tet(50) (PDB ID 5tui).

the factors that inﬂuence inhibitor potency and stability en

route to the development of viable adjuvant approaches to

counter tetracycline resistance by enzymatic inactivation.

Tetracycline-inactivating enzymes, including the most

studied tetracycline destructase, Tet(X),³³and the subse-

quently identiﬁed enzymes Tet(47)−Tet(56),³⁴are Class A

ﬂavin-dependent monooxygenase enzymes conﬁrmed to confer

tetracycline resistance by the nonreversible functionalization of

the tetracycline scaﬀolds (Figure 2A). Gut-derived Tet(X) and

soil-derived Tet(47)−Tet(56) possess unique three-dimen-

sional (3D) structures, which directly contribute to the

observed variation in phenotypic tetracycline resistance proﬁles

across enzyme clades (Figure 2B,C).³⁵⁻³⁷In general,

tetracycline destructase enzymes are composed of at least

three functional domains: a substrate-binding domain, a ﬂavin

adenine dinucleotide (FAD)-binding domain, and a C-terminal

α helix that stabilizes the association of the two. The presence

of a second C-terminal α helix, termed the “Gatekeeper” helix,

was also observed for the soil-derived tetracycline destructases

[Tet(47)−Tet(56)] and is thought to facilitate substrate

recognition and binding.³⁷

A variety of substrate binding modes have been observed for

Tet(X) and the tetracycline destructases. A search for

competitive inhibitors identiﬁed anhydrotetracycline (aTC,

5), a tetracycline biosynthetic precursor, as a potential broad-

spectrum inhibitor (Figures 1, 2).³⁷aTC showed dose-

dependent and potent inhibition of tetracycline destructases

in vitro and rescued tetracycline antibiotic activity against

Escherichia coli overexpressing the resistance enzymes on an

inducible plasmid. The crystal structure of aTC bound to

Tet50 revealed a novel inhibitor binding mode that pushes the

FAD cofactor out of the active site to stabilize an inactive

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Figure 3. Michaelis−Menten kinetics of tetracycline destructase degradation of ﬁrst generation tetracyclines. (A) Representative Michaelis−

Menten plot of tetracycline destructase degradation of oxytetracycline. (B) Representative optical absorbance kinetic plots for the degradation of

oxytetracycline by tetracycline destructase enzymes [as observed at 400 nm for Tet(50) and Tet(X)_3; 380 nm for Tet(X)]. (C) Apparent K_m,

k_app, and catalytic eﬃciencies for the tetracycline destructase-mediated degradation of tetracycline, chlortetracycline, demeclocycline, and

oxytetracycline. Error bars represent standard deviation for two independent trials.

enzyme conformation.³⁷Upon the basis of these preliminary

results, we crafted two hypotheses with regard to tetracycline

destructase inhibition. Because of the variability observed in

phenotypic resistance proﬁles between tetracycline destructase

enzymes and phylogenetic clades, we hypothesized that

inhibitor potency would also vary as a function of enzyme

and inhibitor-substrate pairing; thus, a library of inhibitors may

be required to preserve the viability and eﬀectiveness of an

adjuvant approach. This has proven to be the case with β-

lactam adjuvants, where multiple generations of inhibitors are

required to cover the diverse families of β-lactamase resistance

enzymes (classes A−D) present in the clinic.²⁹In addition, we

proposed that aTC, in particular, could serve as a privileged

scaﬀold about which to design inhibitor libraries. Thus, we

herein report the generation and biological evaluation of four

semisynthetic derivatives of anhydrotetracycline as potential

inhibitors of tetracycline destructase enzymes. To identify the

factors aﬀecting the inhibition of tetracycline-inactivating

enzymes, we assessed the inhibitory activity of the aTC

analogue library, in reference to aTC, against the degradation

of ﬁrst-generation tetracyclines by three representative

tetracycline destructase enzymes (Figure 1). Taken collec-

tively, these results highlight the factors that inﬂuence inhibitor

potency and stability and provide the framework for the

rational design of next-generation inhibitor libraries.

tetracycline (1), chlortetracycline (2), demeclocycline (3), and

oxytetracycline (4). Representative Michaelis−Menten plots

for the enzymatic degradation of oxytetracycline are shown in

Figure 3A.

Consistent with previous reports,³⁷micromolar (μM)

apparent binding aﬃnities (K_m), ranging from 2 μM to 13

μM, were observed across all enzyme−substrate combinations

(Figure 3). Apparent rate (k_app) of tetracycline degradation was

highest for Tet(X)_3, followed closely by Tet(50), and last by

Tet(X); this variation in apparent rate is highlighted in the

deviation in shape and slope of the raw plots of enzyme-

dependent tetracycline degradation, observed as the change in

the absorbance at 400 nm over time (Figure 3B). Conversely,

apparent catalytic eﬃciency (k_app/K_m) was highest for Tet(50)

over Tet(X)_3due to a 2−10 fold diﬀerence in apparent K_m.

This trend is consistent with the hypothesis that Gatekeeper

helix-facilitated substrate recognition results in an increase in

substrate speciﬁcity and turnover for the soil-derived

tetracycline destructases (Figure 2).³⁷The second, C-terminal

gatekeeper helix is notably absent in X-ray crystal structures of

gut-derived, canonical tetracycline-inactivating enzyme, Tet(X)

(Figure 2),³⁵and the presence or absence of a similar helix in

Tet(X)_3which clusters closely with Tet(X)is currently

unknown. Similar apparent binding aﬃnities (K_m) were

observed for phylogenetically clustered Tet(X)_3 and Tet(X),

though ﬁve- to eight-fold diﬀerences in apparent rate results in

drastically diﬀerent catalytic eﬃciencies for the two gut-derived

enzymes (Figure 3C). The paradoxical functional similarities of

Tet(X)_3 to both Tet(50) in apparent rate and Tet(X) in

binding aﬃnity and resistance phenotype, vide infra, has

garnered interest in the unique facets of its three-dimensional

structure that allows for accelerated turnover and broad

substrate scope; eﬀorts to resolve an X-ray crystal structure of

Tet(X)_3 are currently ongoing in our laboratories and will be

reported in due course. Taken collectively, the variability in

binding aﬃnity and catalytic eﬃciency highlights both enzyme-

to-enzyme and substrate-to-substrate diﬀerences across the

tetracycline destructase family of enzymes. In this light, we

hypothesized this same variability would manifest in inhibitor

RESULTS AND DISCUSSION

■

Michaelis−Menten Kinetics Highlight Enzyme Diﬀer-

ences. Three representative tetracycline destructase enzymes

were chosen based upon observed phenotypic resistance

proﬁles and phylogenetic clustering.³⁴These enzymessoil-

derived Tet(50) and gut-derived Tet(X) and Tet(X)_3

[GenBank KU547176.1]were recombinantly expressed and

puriﬁed from BL21-Star (DE3) competent E. coli. For each

enzyme, the in vitro enzyme-dependent inactivation of ﬁrst-

generation tetracyclines was characterized using an optical

absorbance kinetic assay developed in our laboratory.^34,37

Apparent Michaelis−Menten kinetic parameters were deter-

mined from the enzyme- and time-dependent degradation of

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Figure 4. In vitro aTC inhibition of tetracycline destructase degradation of ﬁrst-generation tetracycline antibiotics as observed via an optical

absorbance kinetic assay. (A) aTc inhibition of Tet(50) degradation of tetracyclines. (B) aTc inhibition of Tet(X) degradation of tetracyclines. (C)

aTc inhibition of Tet(X)_3 degradation of tetracyclines. (D) Apparent IC₅₀for aTC inhibition (denoted for each substrate and enzyme). Error

bars represent standard deviation for three independent trials. All data points possess error bars, though some are not visible at the plotted scale.

Scheme 1. Semisynthetic Strategies toward aTC Analogues 6−9

potency ﬂuctuation as a function of enzyme and inhibitor-

substrate pairing.

maximal inhibitory concentrations (IC₅₀s) for the aTC

inhibition of Tet(50) were higher than those observed for

Tet(X) [5- to 10-fold], with the most potent inhibition

observed for Tet(X)_3. Surprisingly, the apparent IC₅₀s

observed for the aTC inhibition of tetracycline destructase-

mediated degradation of tetracyclines varied modestly as a

function of inhibitor-substrate pair within the context of a

single enzyme (Figure 4D). However, the half-maximal

inhibitory concentrations of aTC inhibition of CTc were

aTC Inhibitory Activity Varies as a Function of

Enzyme and Antibiotic Pair. To evaluate the hypothesis

that inhibitor potency will vary as a function of enzyme and

antibiotic pair, we assessed the in vitro inhibitory activity of

aTC against the tetracycline destructase-mediated degradation

of ﬁrst-generation tetracycline antibiotics, the results of which

are displayed in Figure 4A−D. In general, the apparent half-

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Figure 5. In vitro inhibition of tetracycline destructase degradation of ﬁrst-generation tetracycline antibiotics as observed via optical absorbance

kinetic assay. Inhibitory activity of aTC library against (A) Tet(50) degradation of tetracycline, (B) Tet(X) degradation of tetracycline, (C) Tet(X)

_3 degradation of tetracycline, (D) Tet(50) degradation of chlortetracycline, (E) Tet(X) degradation of chlortetracycline, and (F) Tet(X)_3

degradation of chlortetracycline. Error bars represent standard deviation for three independent trials. All data points possess error bars, though

some are not visible at the plotted scale.

notably higher than those observed for the enzymatic

degradation of the other ﬁrst-generation tetracyclines. In

addition, the IC₅₀associated with aTC inhibition of the

Tet(X) degradation of oxytetracycline was over an order of

magnitude lower than other aTC-tetracycline pairs for the gut-

derived enzyme. It is unclear what factors contribute to this

eﬀect; however, the combination of lower binding aﬃnity

(higher K_m, Figure 3C) and the polyhydroxylated nature of

oxytetracycline may allow for more favorable inhibitor

competition for the substrate binding pocketsince oxy-

tetracycline may appear more Tet(X) product-like than other

tetracycline substrates, vide infra.³³

Analysis of Lineweaver−Burk plots of the aTC inhibition of

tetracycline destructase-mediated degradation of tetracycline

supports the idea of a mixed competitive/noncompetitive

inhibition model (Supporting Information Figure 2). Taken

collectively, these results suggest that aTC inhibition involves

more than a contest of competitive binding and catalytic

eﬃciency, which is consistent with multicomponent enzyme

processes and substrate/inhibitor binding mode ﬂexibility

observed for the tetracycline destructase enzymes.^36,38−42

Consequently, because inhibition model ambiguity and

binding mode ﬂexibility can complicate broad computational

docking to direct the rational design of inhibitors, direct

modiﬁcation of the aTC scaﬀold may be the most eﬃcient way

to aid in the generation of larger inhibitor libraries by

determining empirical inhibitor structure−activity relationships

(SAR).

Semisynthesis of aTC Analogues. Inspired by the

seminal work of Nelson and co-workers, as well as Gmeiner

and co-workers, involving semisynthetic strategies to tetracy-

cline and anhydrotetracycline analogues,^43,44we synthesized

four aTC analogues from parent tetracyclines to evaluate the

potential for aTC to serve as a privileged scaﬀold for the

development of tetracycline-inactivating enzyme inhibitors

(Scheme 1). Acid-catalyzed dehydration of C6-hydroxy-

tetracyclines chlortetracycline (CTc, 2) and demeclocycline

(Dem, 3) provided the corresponding anhydrotetracycline

variants, anhydrochlortetracycline (aCTc, 6) and anhydrode-

meclocycline (aDem, 7) in quantitative crude yields and

excellent isolated yields (C18-silica gel, reverse-phase prepa-

rative high-performance liquid chromatography (HPLC)).

Correspondingly, electrophilic aromatic substitution of anhy-

drotetracycline (aTC, 5) with either N-iodosuccinimide (NIS)

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Figure 6. (A) Whole cell inhibition of E. coli expressing tetracycline destructase enzymes including calculated FICI and observed fold change

enhancements. (B) Working model of the inhibition of tetracycline destructase enzymes by aTC-like small molecules (competitive inhibitor vs

sacriﬁcial substrate).

or molecular bromine (in acid) aﬀorded C7-iodoanhydrote-

tracycline (7-I-aTC, 8) and C9−Br-anhydrotetracycline (9-Br-

aTC, 9) in excellent crude and isolated yields. These brightly

colored solids were storable at low temperatures (−20 °C),

away from light, with little decomposition observed over six

month periods (by high-performance liquid chromatography−

mass spectrometry, LCMS), and stability and longevity

improved when the compounds were stored at low temper-

ature under argon atmosphere. However, triturations with

methyl-tert-butyl ether (MTBE) of analogues with trace

impurities, followed by ﬁltration, allowed for reisolation of

greater than 90−95% purity material as determined by LCMS

and NMR.

Biological Evaluation of aTC Analogue Library. With a

scalable synthetic route in hand, we evaluated the ability of the

aTC analogues to inhibit tetracycline-destructase enzymes via

an in vitro optical absorbance kinetic assay and referenced the

results to those obtained for known inhibitor, aTC, in parallel

scenarios (Figure 5). In general, inhibitor potency varied in an

enzyme-dependent manner that was consistent with what was

previously observed with aTC, vide supra. Apparent half-

maximal inhibitory concentrations (IC₅₀s) were highest for the

Tet(50)-mediated degradation of tetracycline and chlortetra-

cycline (Figure 5A,D), followed by Tet(X) (Figure 5B,E) and

Tet(X)_3 (Figure 5C,F), respectively. Inhibitor potency

decreased (2- to 10-fold) when inhibitors were coadministered

with CTc (Figure 5D−F) over tetracycline (Figure 5A−C),

and chlorinated aTC analogues aCTc and aDem performed

marginally better than aTC when coadministered with

tetracycline, suggesting little signiﬁcant cooperativity/syner-

gism of structurally similar inhibitor-substrate pairs. In general,

halogenation of the D-ring improved in vitro inhibitory activity,

though the enhancement observed for C7-chlorination is more

pronounced across all enzyme-antibiotic combinations. Re-

moval of the C6-methyl group was well tolerated for the

inhibition of Tet(X) and Tet(X)-homologue, Tet(X)_3;

however, aDem performed poorly against tetracycline

destructase Tet(50). Proposed reasoning for this phenomenon

is discussed later in this report, vide infra.

With in vitro inhibitor potencies established, we next tested

the ability of the aTC analogues to rescue tetracycline activity

in whole cell inhibition assays of E. coli expressing tetracycline

destructase enzymes. By varying concentrations of the

tetracycline-inhibitor combinations in a checkboard broth

microdilution antibiotic susceptibility assay, we were able to

identify the lowest concentration of inhibitor that results in at

least a fourfold change in the minimum inhibitory concen-

tration (MIC) of the tetracycline alone; highlights of the

checkerboard assay are shown in Figure 6A. Severalnot all

tetracycline-inhibitor pairs showed enhanced antibiotic activity,

when tetracycline substrates were coadministered with small

doses of various inhibitors (four- to eight-fold enhancement

over antibiotic alone, Figure 6A). Notably, though aTC was, in

general, the least potent of the inhibitor library, it performs

welland on par with the chlorinated analogues, aCTc and

aDemin the whole cell rescue of tetracycline activity. It is

important to note that, while some of the aTC analogues

possess baseline antibiotic activity alone, antibiotic synergistic

killing and destructase inhibition may not be mutually

exclusive (Figure 6A), and it is not entirely clear how much

of a contribution is made by each mechanism for each

combination. The calculated fractional inhibitory concentra-

tion index (FICI) is provided for reference.⁴⁵However, the

enhancements in MIC observed for several anhydrotetracy-

cline-variant/antibiotic combinations are modest yet promis-

ing, and potent in vitro destructase inhibition by each of the

aTC variantsin combination with these resultssuggests

that whole cell adjuvant activity could be optimized in the

future second generation synthesis of larger aTC-like inhibitor

libraries.

While 9-Br-aTC and 7-I-aTC possess potent in vitro

destructase inhibitory activity, the analogues showed limited

activity in whole cell inhibition studies, potentially due to

increased instability in solution over time and/or increased

reactivity toward enzymatic degradation. In particular, 7-I-aTC

showed no inhibitory activity with any enzyme−antibiotic

combination against whole cell E. coli expressing destructase

enzymes. We hypothesized that the instability of 7-I-aTC was

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Figure 7. Tet(X)-mediated degradation of aTC and aTC analogues. (A−F) Each panel depicts the degradation of the denoted aTC as observed via

optical absorbance spectroscopy and plots of monitored extracted mass counts from LCMS. Each represents a reaction containing puriﬁed Tet(X)

enzyme (or none, in the case of the control), aTc analogue, and an NADPH regenerating system [including MgCl₂]. The plots represent extracted

ion counts normalized to an internal standard (Fmoc-alanine) and depicted as a percent of the total ion count [aTC+aTC−OH].

contributing to this phenomenon, as the nonenzymatic

degradation of 7-I-aTc in solution was observed by LCMS

(overnight) and during extended ¹³C NMR experiments.

Because the o/p-substitution of phenols with heavy halogens is

known to increase the rate of nonenzymatic photooxida-

tion,⁴⁶⁻⁴⁹the incorporation of D-ring halogens to future

inhibitor libraries may be limited to chlorination and

ﬂuorination to preserve inhibitor stability. However, the Br-

and I-substituents present in 7-I-aTC and 9-Br-aTC may serve

as useful functional handles to access more structurally diverse

and stable inhibitor scaﬀolds. While we propose that disparities

in in vitro inhibitor potency and whole cell rescue of

tetracycline activity are largely due to problems with inhibitor

stability, vide infra, the potent in vitro inhibition of tetracycline

destructase enzymes suggests structural modiﬁcations that

promote inhibitor stability and maintain potency could

improve whole cell performance.

formations via a series of complex and dynamic conformational

changes involving a mobile ﬂavin cofactor.^33,34,38,39While the

precise sequence of events is currently unknown, the proposed

degradation process involves substrate recognition and binding

to FAD-OUT enzyme conformation I (Figure 6B), followed by

rapid ﬂavin reduction (FAD to FADH₂) by NADPH. Through

discrete conformational changes, the reduced ﬂavin cofactor is

pushed toward the newly bound substrate and reacts with

molecular oxygen to generate the reactive C4a-hydroperoxy-

ﬂavin in the solvent-protected FAD-IN conformation. The

hydroperoxyﬂavin then reacts with the tetracycline sub-

strate,^33,34,36,39and another series of conformational changes

allows for release of the degradation product and dehydration

of the resultant C4a-hydroxyﬂavin to regenerate FAD-OUT

conformer I.

Upon the basis of the ﬁndings presented herein, in

combination with previous studies,^33,34,37we developed a

working model for inhibition of tetracycline destructase

enzymes in the context of the proposed degradation process

(Figure 6B). In the event that inhibition occurs competitively,

inhibitor could bind to substrate unbound enzyme and exclude

Working Inhibition Model. The tetracycline destructase

enzymes are class A ﬂavin-monooxygenase (FMO) enzymes

that catalyze nicotinamide adenine dinucleotide phosphate

(NADPH)- and oxygen-dependent, multicomponent trans-

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Figure 8. Tet(X)-mediated degradation of oxytetracycline as a model for the degradation of aTC-like sacriﬁcial substrates.

Figure 9. aDem inhibition of tetracycline destructase-mediated degradation of ﬁrst-generation tetracyclines. (A) aDem inhibition of tetracycline

destructase degradation of tetracycline. (B) aDem inhibition of tetracycline destructase degradation of demeclocycline. (C) Apparent IC₅₀for the in

vitro inhibition of tetracycline and demeclocycline degradation by tetracycline destructase enzymes with aTC and aDem. (D) Michaelis−Menten

plot of dose-dependent aDem acceleration of Tet(50) consumption of NADPH, apparent K_m, and calculated catalytic eﬃciency. Error bars

represent standard deviation for two to three independent trials. All data points possess error bars, though some are not visible at the plotted scale.

accommodation of substrate into the active site; this inhibited

state (III) could be a stalled state (nonproductive), or the

inhibitor could react with the enzyme as a sacriﬁcial substrate

(moving through inhibited state IV and multiple successive

inhibited states en route to the enzymatic degradation of the

inhibitor, Figure 6B). Alternatively, inhibition could occur

noncompetitively, where inhibitor binds to substrate-bound

enzyme complex (Intermediate II or successive substrate-

bound functional enzyme states) and restrains the conforma-

tional ﬂexibility of the enzyme to impede productive turnover.

Because degradation is a complex, multicomponent process

involving a number of discrete enzyme conformational

changes, none of the multiple inhibited states derived from

substrate-bound and substrate-unbound functional enzyme

states can implicitly be excluded. Therefore, to assess the

potential of the generated aTC analogues to undergo

enzymatic degradation as sacriﬁcial substrates, we chose to

evaluate the in vitro degradation of the inhibitor library by

canonical tetracycline-inactivating enzyme, Tet(X), which is

known to degrade aTC, albeit slowly.³⁴

aTC Analogues as Substrates for Destructase

Enzymes. Using Tet(X) as a model system, we assessed the

potential for the aTC analogues to serve as sacriﬁcial substrates

via an in vitro, broad-scan optical absorbance kinetic assay

coupled to LCMS, as previously reported for the Tet(X)-

mediated degradation of aTC.³⁴The results of the Tet(X)-

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Figure 10. (A) X-ray crystal structure of aTC bound to Tet(50) [PDB ID: 5TUF] and corresponding binding mode identiﬁer; (B) time- and

aDem-dependent degradation of NADPH by Tet(50) observed using a broad-scan optical absorbance kinetic assay; (C) hydrogen peroxide

colorimetric detection experiments, from left to right: NADPH (no enzyme) control, no enzyme control (NADPH + aDem), Tet(50) + NADPH

[no aDem] reaction mixture, and Tet(50) + NADPH + aDem reaction mixture.

mediated aTC analogue degradation assays are summarized

below (Figure 7). We conﬁrmed that aTC is a substrate for

Tet(X), indicated by the time- and enzyme-dependent

decrease in the 440 nm absorption band and aTC extracted

mass (LCMS) and used this result as a positive control for the

enzymatic degradation of the aTC scaﬀold. In general, aTC

analogue stability increased with D-ring halogenation, and

chlorination provided the most “protection” against enzymatic

degradation. Moreover, observed stability tracks with degree of

electron deﬁciency and the electron-withdrawing nature of the

added substituent (i.e., aCTc is more stable than 9-Br-aTC,

which is more stable than 7-I-aTC and aTC, respectively). We

hypothesize that this stability is due to decreased nucleophil-

icity of the C,D-ring aromatic framework, which would be

fundamentally important to an enzyme-mediated electrophilic

hydroxylation event similar to that determined by Wright and

co-workers in 2004 for Tet(X) degradation of oxytetracycline

(Figure 8).³³

Anhydrodemeclocycline, a Special Case. As detailed

above, chlorinated aTC analogue anhydrodemeclocycline

(aDem, 7) possesses potent in vitro inhibitory activity against

the Tet(X)- and Tet(X)_3-mediated degradation of tetracy-

cline antibiotics (apparent IC₅₀s 1−4 μM, Figure 9A−C) and

rescues tetracycline activity in whole cell E. coli expressing

these enzymes (Figure 6A); however, 7 was found to possess

poor in vitro inhibitory activity against the enzymatic

degradation of both tetracycline and demeclocycline by soil-

derived tetracycline destructase Tet(50). Moreover, when

aDem was exposed to Tet(50) in the presence of NADPH and

absence of tetracycline in an optical absorbance kinetic assay, a

steady, observable decrease in the absorbance at 400 nm was

observed, suggesting that aDem itself was a substrate for

Tet(50). To expand upon this observation, we determined

kinetic parameters for the aDem dose-dependent response,

similar to the Michaelis−Menten parameters previously

described (Figure 9D); however, LCMS-coupled experiments

for the reaction showed no measurable decrease in aDem

extracted mass over time (Supporting Information Figure 3)

suggesting that the decrease in absorbance at 400 nm was the

result of rapid, aDem-dependent Tet(50)-mediated consump-

tion of NADPH (broad absorbance band at 340 nm) and not

the enzymatic degradation of aDem 7.

The previously reported X-ray crystal structure of aTC

bound to Tet(50) revealed a unique inhibitor binding mode,

42

denoted Mode I_A,D

,

resulting from the noncovalent

interaction of aTC with residues in both the substrate binding

domain and the FAD binding domains of the enzyme (Figure

10A).³⁷In particular, the C6-methyl substituent on aTC

occupies a small hydrophobic pocket between lysine 198,

lysine 205, and methionine 222 of the substrate binding

domain. Because this substituent is notably absent in aDem, we

hypothesized that binding mode ﬂexibility previously observed

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Figure 11. (A) Michaelis−Menten plot and apparent kinetic parameters (K_m, k_app, and catalytic eﬃciencies) for the Tet(X)-mediated degradation

of tigecycline. (B) Inhibitory activity of aTC and chlorinated aTC analogs against Tet(X) degradation of tigecycline. (C) Last-generation

tetracycline antibiotics tigecycline and eravacycline. Error bars represent standard deviation for three independent trials. All data points possess

error bars, though some are not visible at the plotted scale.

for tetracycline-inactivating enzymes could allow for the

accommodation of aDem in a unique, nonreactive binding

mode to promote the NADPH-dependent reduction of the

mobile ﬂavin element without providing a productive pathway

for the degradation of enzyme-bound aDem. Moreover,

because the addition of aDem promotes the Tet(50)-mediated

consumption of NADPH without resulting in the degradation

of aDem itself, we hypothesized that the formation of a reactive

hydroperoxyﬂavin cofactor in the absence of a viable substrate

would lead to the release of hydrogen peroxide generated from

the nonspeciﬁc oxidation of water. The NADPH-dependent

formation of hydrogen peroxide from FMOs has been reported

previously.^50,51Thus, we characterized the aDem-promoted,

Tet(50)-mediated consumption of NADPH using a broad-scan

optical absorbance kinetic assay (Figure 10B) and conﬁrmed

the formation of hydrogen peroxide using a colorimetric

detection method (Pierce Quantitative Peroxide Assay,

ThermoScientiﬁc, Figure 10C). The nonspeciﬁc formation of

hydrogen peroxide was not unique to aDem, as it was also

observed in the Tet(50)-mediated degradation of tetracycline

(see Supporting Information Figures 4 and 5), conﬁrming that

“ligand” binding to the substrate-binding domain promotes the

consumption of NADPH and reduction of the mobile ﬂavin

cofactor (as is canonical with Class A FMO enzymes).³⁸This

mode of inhibition is under further investigation in our lab and

might prove to be eﬀective against pathogens expressing

tetracycline destructases, as a method of inducing oxidative

stress by stimulating ﬂavin reduction and release of hydrogen

peroxide inside the cell.

systems for tetracycline-inactivating enzyme activity; however,

with the advent of tigecycline, eravacycline, and omadacycline,

the enzymatic degradation of last-generation tetracyclines is

fundamentally important to study, since new resistance

mechanisms, including antibiotic inactivation, are certain to

emerge upon widespread antibiotic deployment. The third-

generation tetracyclines were strategically designed to over-

come resistance due to eﬄux and ribosome protection, making

antibiotic inactivation a likely candidate for future clinical

resistance. Previous reports have identiﬁed that Tet(X) can

degrade tigecycline and eravacycline, albeit slowly, to achieve

resistance to these last-generation tetracyclines·⁵⁵⁻⁵⁷Resist-

ance to tigecycline was not observed with the soil-derived

tetracycline destructase enzymes, including Tet(50), presum-

ably because the presence of the “Gatekeeper helix” excludes

productive accommodation of tetracyclines with bulky D-ring

substituents (Figure 2C).^34,37Building upon previous reports,

we conﬁrmed and characterized the Tet(X)-mediated degra-

dation of tigecycline and determined Michaelis−Menten

kinetic parameters using an optical absorbance kinetic assay

(Figure 11A). After identifying the most promising inhibitor

candidates from previously described in vitro and whole cell

inhibition assays, we evaluated the in vitro inhibitory activity of

aTC, aCTc, and aDem against the Tet(X)-mediated

degradation of tigecycline and found all to be potently

inhibitory (apparent IC₅₀s from ∼0.4 to 1.5 μM). Unfortu-

nately, in our hands, whole cell E. coli expressing Tet(X)

displays minimal resistance to tigecycline (twofold increase

over empty vector controls); thus, it was diﬃcult to identify

inhibition proﬁles from variations in the limited resistance

response. However, the potent in vitro inhibition of aTC,

aCTc, and aDem against Tet(X) degradation of tigecycline are

promising preliminary results for further development of

adjuvant approaches to combat the enzymatic degradation of

last-generation tetracyclines. Moreover, because of the func-

tional similarities and phylogenetic clustering of the gut-

derived enzymes, we hypothesize that Tet(X)_3 may possess

similar abilities to degrade last-generation tetracyclines with

velocities more amenable to both in vitro and whole cell

inhibition assaysthough full characterization of Tet(X)_3

Inhibiting Tet(X) Degradation of Third Generation

Tetracycline, Tigecycline. The rise of multidrug-resistant

superinfections has cultivated a renaissance for tetracycline

antibiotics as last-resort treatments.¹⁹In 2005, the ﬁrst

member of the third-generation tetracyclines tigecycline was

Food and Drug Administration (FDA)-approved for the

treatment of skin and intra-abdominal infections and pneumo-

nia.⁵²Earlier this year, two additional third-generation

tetracyclines, eravacycline and omadacycline, were FDA-

approved for similar treatment strategies (Figure 11C).^53,54

In this report, we used ﬁrst-generation tetracyclines as model

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Figure 12. Toward extended library synthesis of aTC-like inhibitors of tetracycline destructase enzymes. (A) An adjuvant approach to combat

enzymatic inactivation of tetracycline antibiotics and summary of preliminary structure−activity information. (B) Surface view of X-ray crystal

structure of Tet(X) bound to tigecycline (PDB ID 4a6n) highlights importance of open active site to accommodate bulky D-ring substituents,

suggesting further use of glycylcycline antibiotics may drive selective pressure of tetracycline destructase-involved resistance mechanisms.

was somewhat beyond the scope of this report. Studies focused

on the resistance proﬁle and microbial evolution of

tetracycline-inactivating enzyme Tet(X)_3 are currently on-

going in our laboratories and will be reported in due course.

EXPERIMENTAL SECTION

■

General Methods. Unless stated, all synthetic reactions

were performed under inert, argon atmosphere, and all in vitro

kinetic assays were prepared actively open to air (in

nondegassed solvents). All solventsincluding deuterated

NMR solventsand reagent chemicals used in preparation

or analysis of the aTC analogue library were obtained

commercially and used without further puriﬁcation. IR

spectroscopy was performed on a Bruker Alpha FTIR machine

with a Pt-ATR diamond, and IR data were analyzed using

Bruker OPUS 7.5. Melting points were observed using a Stuart

SMP10 digital melting point apparatus. NMR spectra were

obtained on a Varian Unity-Plus 300 MHz, Varian Unity-Inova

500 MHz, or Agilent PremiumCompact+ 600 MHz

spectrometer. All free induction decay ﬁles (FIDs) were

processed using Mestrenova version 11.0.4 software. Chemical

shifts (δ) are reported in parts per million (ppm) and

referenced to residual nondeuterated solvent. Coupling

constants (J) are reported in hertz (Hz). High-resolution

mass spectrometry data were obtained at the Danforth Plant

Science Center (DPSC) in St. Louis, MO, by direct infusion

using an Advion Nanomate Triversa robot into a Thermo-

Fisher Scientiﬁc Q-Exactive mass spectrometer, and mass

spectra were recorded in positive ion mode from m/z 150−500

and a resolution setting of 140 000 (at m/z 200). In vitro

degradation experiments monitored by optical absorbance

spectroscopy were performed on an Agilent Cary 50 UV−

visible spectrophotometer. In vitro degradation experiments

monitored by LCMS were performed using an Agilent 6130

single quadrupole instrument with G1313 autosampler, G1315

diode array detector, and 1200 series solvent module and

separated using a Phenomenex Gemini C18 column, 50 × 2

mm (5 μm) with guard column cassette and a linear gradient

of 0% acetonitrile and 0.1% formic acid to 95% acetonitrile and

0.1% formic acid over 20 min at a ﬂow rate of 0.5 mL/min

before analysis by electrospray ionization (ESI+). Whole cell

assays were performed using Difco BBL Mueller-Hinton broth

CONCLUSION

■

In conclusion, the synthesis and biological evaluation of aTC-

like small molecule inhibitors of tetracycline-inactivating

enzymes are reported (Figure 12A). The four analogues

were screened for inhibitory activity against the enzymatic

degradation of tetracycline antibiotics by three representative

tetracycline destructase enzymes via both in vitro and whole

cell-based inhibition assays. All synthesized analogues were

found to possess in vitro inhibitory activity to some degree, and

inhibitor potency was found to vary largely as a function of

enzyme and moderately as a function of inhibitor−substrate

pairing within the context of a single enzyme. The addition of

electron-withdrawing groups to the D-ring of aTC was found

to improve both the enzymatic and nonenzymatic stability of

the aTC analogues, and potent in vitro inhibitory activity of

this small library shows promise for the rational design of

larger tetracycline-inactivating enzyme inhibitor libraries.

Notably, aTC and chlorinated analogues aCTc and aDem

were found to inhibit the Tet(X)-mediated degradation of last-

generation tetracycline, tigecycline. Further development of

small molecule inhibitors of glycylcycline-inactivating enzymes

like Tet(X), with open active sites that can accommodate large

D-ring substituents on tetracycline substrates (Figure 12B),³⁵

are fundamentally important to establishing viable adjuvant

approaches that combat the imminent emergence of this

resistance mechanism in multidrug-resistant infections. Eﬀorts

aimed at improving inhibitor stability while maintaining

potency are currently ongoing in our laboratories and will be

reported in due course.

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in Costar 96-well plates at 37 °C. End-point growth was

assayed at OD₆₀₀using a Synergy H1 plate reader (BioTek,

Inc.).

condenser, was added chlortetracycline hydrochloride (50 mg,

0.10 mmol) and 6 N HCl in methanol (5 mL) under argon

atmosphere. The reaction was heated to 60 °C and allowed to

stir at 60 °C for 1.5 h (monitored by LCMS). When the

reaction was complete, the reaction was concentrated under

reduced pressure to provide crude product (50 mg, 0.10 mmol,

100% crude yield) as an orange solid [clean by NMR].

Puriﬁcation by preparative HPLC (Si−C18 reverse phase

column, gradient 0−95% CH₃CN/H₂O with 0.1% formic acid,

t_R= 16 min) provided (4S,4aS,12aS)-7-chloro-4-(dimethyla-

mino)-3,10,11,12a-tetrahydroxy-6-methyl-1,12-dioxo-

1,4,4a,5,12,12a-hexahydro-tetracene-2-carboxamide, which was

reconstituted as the hydrochloride salt to provide the title

compound as an orange solid (44 mg, 0.089 mmol, 91% yield).

FTIR (neat) 3306, 3042, 1621, 1583, 1565, 1531, 1464, 1375,

1217, 1137, 1058, 820, 560 cm⁻¹; mp 211−212 °C

(decomposed); ¹H NMR (500 MHz, deuterated dimethyl

sulfoxide (DMSO-d₆)) δ 9.68 (s, 1H), 9.22 (s, 1H), 7.66 (d, J

= 8.5 Hz, 1H), 6.88 (d, J = 8.5 Hz, 1H), 4.44 (d, J = 4.0 Hz,

1H), 3.46 (ddt, J = 18.0, 14.2, 4.9 Hz, 2H), 3.15 (dd, J = 16.8,

8.9 Hz, 1H), 2.93 (s, 6H), 2.63 (s, 3H); ¹³C NMR (126 MHz,

DMSO-d₆) δ 199.7, 192.7, 187.4, 172.1, 163.2, 157.6, 136.2,

135.9, 134.1, 121.5, 119.3, 114.6, 111.8, 109.2, 97.6, 76.2, 66.9,

42.9 (2C), 35.8, 29.6, 19.2; high-resolution mass spectrometry

(HRMS) (time-of-ﬂight (TOF) MS ES+) calcd for

C₂₂H₂₂ClN₂O₇[M + H]⁺461.1116; found 461.1115.

(4S,4aS,12aS)-7-Chloro-4-(dimethylamino)-3,10,11,12a-

tetrahydroxy-1,12-dioxo-1,4,4a,5,12,12a-hexahydrotetra-

cene-2-carboxamide hydrochloride (aDem, 7). To a clean,

dry round-bottom ﬂask, equipped with stirbar and reﬂux

condenser, was added demeclocycline hydrochloride (50 mg,

0.10 mmol) and 6 N HCl in methanol (5 mL) under argon

atmosphere. The reaction was heated to 70 °C and allowed to

stir at 60 °C for 3 h (monitored by LCMS). When the reaction

was complete, the reaction was concentrated under reduced

pressure to provide crude product (54 mg, 0.11 mmol, quant.

yield) as a yellow-orange solid (clean by NMR). Puriﬁcation

by preparative HPLC (Si−C18 reverse phase column, gradient

0−95% CH₃CN/H₂O with 0.1% formic acid, t_R= 15 min) to

provide (4S,4aS,12aS)-7-chloro-4-(dimethylamino)-

3,10,11,12a-tetrahydroxy-1,12-dioxo-1,4,4a,5,12,12a-hexahy-

drotetracene-2-carboxamide, which was reconstituted as the

hydrochloride salt to provide the title compound as a light

orange solid (37 mg, 0.077 mmol, 77% yield). FTIR (neat)

3301, 3076, 1619, 1567, 1533, 1444, 1375, 1353, 1202, 1188,

1071, 993, 820, 685 cm⁻¹; mp 213−214 °C (decomposed); ¹H

NMR (500 MHz, DMSO-d₆) δ 9.69 (s, 1H), 9.29 (s, 1H), 7.68

(d, J = 8.4 Hz, 1H), 7.39 (s, 1H), 6.92 (d, J = 8.5 Hz, 1H),

4.46 (d, J = 6.5 Hz, 1H), 3.58 (dd, J = 17.3, 4.0 Hz, 1H), 3.53−

3.39 (m, 2H), 2.89 (s, 6H); ¹³C NMR (126 MHz, DMSO-d₆)

δ 198.3, 193.1, 187.2, 172.2, 165.2, 157.2, 135.9, 135.2, 132.7,

119.2, 113.9, 113.5, 111.3, 109.7, 97.5, 76.8, 66.4, 42.3, 41.6,

37.2, 29.5; HRMS (TOF MS ES+) calcd for C₂₁H₂₀ClN₂O₇

[M + H]⁺447.0959; found 447.0958.

Cloning, Expression, and Puriﬁcation of Tetracycline-

Destructase Enzymes. All genes corresponding to the

tetracycline destructases^34,37used in this report (for Tet(X)

_3, see Supporting Information Table 1) were cloned into

pET28b(+) vectors (Novagen) as previously described

(BamHI and NdeI restriction sites)^34,37and transformed into

BL21-Star (DE3) competent cells (Life Technologies). Cells

were cultured at 37 °C in lysogeny broth (LB) containing

kanamycin (0.03 mg/mL); once the uninduced culture

reached an OD₆₀₀of 0.6, the cells were cooled to 0 °C,

induced with 1 mM IPTG, and allowed to grow at 15 °C for

12−15 h (harvest OD₆₀₀varied by tetracycline destructase

expressed, but on average, harvest OD₆₀₀< 4.5 resulted in

greater isolated enzyme yield). To harvest, 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 2-mercaptoethanol, pH 8.0) containing SIGMAFAST

protease inhibitor. The cells were transferred to falcon tubes,

ﬂash frozen in liquid nitrogen, and stored at −80 °C. To

harvest, the cells were thawed and mechanically lysed using an

Avestin EmulsiFlex-C5 cell disruptor, and the resultant lysate

was centrifuged at 45 000 rpm for 35 min. The supernatant was

transferred to a column containing prewashed Ni-NTA resin

and incubated for 30−45 min; at which point, the resin was

washed with lysis buﬀer (2 × 40 mL), and the protein was

eluted from the resin with fractions of elution buﬀer (5 × 10

mL, 50 mM K₂HPO₄, 500 mM NaCl, 5 mM β-

mercaptoethanol, 300 mM imidazole, 10% glycerol, pH 8.0).

The fractions were combined in 10 000 molecular weight

cutoﬀ (MWCO) Snakeskin dialysis tubing (ThermoScientiﬁc)

and soaked in buﬀer (50 mM K₂HPO₄pH 8.0, 150 mM NaCl,

1 mM dithiothreitol (DTT)) overnight to minimize imidazole

concentration. To isolate the desired protein, the dialyzed

solution was concentrated using a 30 000 MWCO Amicon

centrifugal ﬁlter (Millipore-Sigma), and concentrated protein

solution was ﬂash frozen in liquid nitrogen (50 μL portions)

and stored at −80 °C.

Kinetic Characterization of Tetracycline Inactivation.

Kinetic characterization of tetracycline inactivation was

achieved in a manner similar to previously reported

procedures.³⁷In brief, reaction samples were prepared in

[tris(hydroxymethyl)methylamino]propanesulfonic acid

(TAPS) buﬀer (100 mM, pH 8.5) with 504 μM NADPH,

5.04 mM MgCl₂, varying concentrations of tetracycline

substrate (typically, 0−40 μM), and 0.4 μM enzyme. After

the addition of enzyme, the reactions (in duplicate or

triplicate) were mixed, manually by pipet, and the reaction

was monitored continuously in a single frame by optical

absorbance spectroscopy (absorbance at 380 nm, Carey UV−

visible spectrophotometer) for 3−4 min. Initial enzyme

velocities were determined by linear regression using Agilent

Cary WinUV Software over the linear range of the reaction,

and the velocities were ﬁtted to the Michaelis−Menten

equation⁵⁸⁻⁶⁰using GraphPad Prism 6.

(4S,4aS,12aS)-4-(Dimethylamino)-3,10,11,12a-tetrahy-

droxy-7-iodo-6-methyl-1,12-dioxo-1,4,4a,5,12,12a-hexahy-

drotetracene-2-carboxamide hydrochloride (7-I-aTC, 8).

Procedure A−To a clean, dry round-bottom ﬂask, equipped

with stirbar, was added anhydrotetracycline hydrochloride

(100 mg, 0.216 mmol) and methanol (2.2 mL) under argon

atmosphere. The ﬂask was cooled to −10 °C, and N-

iodosuccinimide (58.3 mg, 0.259 mmol) was added in one

portion. The reaction was allowed to warm to room 0 °C over

Synthesis and Characterization of aTC Analogues 5−

8. (4S,4aS,12aS)-7-Chloro-4-(dimethylamino)-3,10,11,12a-

tetrahydroxy-6-methyl-1,12-dioxo-1,4,4a,5,12,12a-hexahy-

drotetracene-2-carboxamide Hydrochloride (aCTc, 6). To a

clean, dry round-bottom ﬂask, equipped with stirbar and reﬂux

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2 h, then stirred at 0 °C for 1 h (monitored by LCMS). When

the reaction was complete, the reaction was diluted with

methanol (to 10 mL total volume) and immediately puriﬁed

by preparative HPLC (Si−C18 reverse phase column, gradient

0−95% CH₃CN/H₂O with 0.1% formic acid, t_R= 16 min) to

provide (4S,4aS,12aS)-4-(dimethylamino)-3,10,11,12a-tetrahy-

droxy-7-iodo-6-methyl-1,12-dioxo-1,4,4a,5,12,12a-hexahydro-

tetracene-2-carboxamide, which was reconstituted as the

hydrochloride salt to provide the title compound as an orange

solid (70.0 mg, 0.119 mmol, 55% yield).

presence of varying concentrations of chosen inhibitor.

Reaction samples were prepared in 100 mM TAPS buﬀer

(pH 8.5) with 504 μM NADPH, 5.04 mM MgCl₂, 25.3 μM

tetracycline substrate, varying concentrations of inhibitor

(μΜ), and 0.4 μM enzyme. After the addition of enzyme,

the reactions (in triplicate) were mixed manually by pipet, and

the reaction was monitored, continuously in a single frame, by

optical absorbance spectroscopy (absorbance at 380 or 400

nm, Carey UV−visible spectrophotometer) for 4 min. Initial

enzyme velocities were determined by linear regression using

Agilent Cary WinUV Software over the linear range of the

reaction. The velocities were plotted against the logarithm of

inhibitor concentration, and IC₅₀values were determined using

nonlinear regression analysis in Graphpad Prism 6. Plus/minus

error values were determined using linear regression analysis of

initial velocities versus concentrations of inhibitor in Graphpad

Prism 6. Each set of experiments was accompanied by a variety

of controls, including a no-enzyme control (NADPH + Tet +

inhibitor)which was used to simulate full enzyme inhibition

and assigned to inhibitor concentration of 1 × 10¹⁵, and a no-

inhibitor control (NADPH + Tet + enzyme)which was

assigned an inhibitor concentration of 1 × 10⁻¹⁵. A no-

substrate control (NADPH + inhibitor + Tet) was also

performed to identify competitive background signals from the

enzymatic degradation of the inhibitor itself. For all inhibitor−

enzyme combinations (except for Tet(50)−aDem), the initial

velocities of the no-substrate controls were negligible.

Checkerboard Whole Cell Inhibition Assay. Substrates

and inhibitors were dissolved DMSO before being diluted to

working concentrations in cation-adjusted Mueller−Hinton

broth supplemented with 50 μg/mL kanamycin. A twofold

dilution series of each drug was made independently across 8

rows of a 96-well master plate before 100 μL of each drug

dilution series was combined into a 96-well culture plate

(Costar), with rows included for no-drug and no-inocula

controls. The plates were inoculated with ∼1 μL of tetracycline

destructase expressing E. coli MegaX (Invitrogen) diluted to

OD₆₀₀0.1 using a sterile 96-pin replicator (Scinomix). Plates

were sealed with Breathe-Easy membranes (Sigma-Aldrich)

and incubated at 37 °C with shaking at 220 rpm. End-point

growth was assayed at OD₆₀₀at 20 and 36 h of growth using a

Synergy H1 plate reader (BioTek, Inc.). Three independent

replicates were performed for each strain/drug combination.

Highlighted MIC data were reﬁned from a complete raw data

set to identify mixtures resulting in the largest MIC fold change

(at least fourfold) with the least amount of inhibitor (fold

change/inhibitor dose; see Figure 6A and Supporting

Information Tables 2a −c). Synergy of inhibitor and tetracy-

cline combinations was determined using the fractional

inhibitory concentration index (FICI) method⁴⁵

Procedure B−To a clean, dry round-bottom ﬂask, equipped

with stirbar, was added anhydrotetracycline hydrochloride

(250 mg, 0.540 mmol) and methanol (25 mL) under argon

atmosphere. The ﬂask was cooled to 0 °C, and solid N-

iodosuccinimide (0.134 g) was added, in one portion. The

reaction stirred at 0 °C for 1 h (monitored by LCMS), and the

reaction was concentrated under reduced pressure (no

heating) to yield a crude brown solid. The solid was triturated

with tert-butylmethyl ether (TBME) for 30 min protected from

light; ﬁltration provided the title product (0.3178 g, 0.540

mmol, quantitative crude yield) as a green-brown solid. FTIR

(neat) 3307, 3078, 1660, 1615, 1556, 1396, 1377, 1321, 1228,

1

1131, 1075, 1058, 810, 702, 618 cm⁻¹; mp 188−189 °C; H

NMR (500 MHz, DMSO-d₆) δ 9.62 (s, 1H), 9.23 (s, 1H), 8.01

(d, J = 8.9 Hz, 1H), 7.31 (d, J = 8.9 Hz, 1H), 4.38 (s, 1H),

3.57−3.36 (m, 3H), 3.16 (s, 1H), 2.89 (s, 6H), 2.56 (s, 1H),

2.40 (s, 3H); ¹³C NMR (126 MHz, DMSO-d₆) δ 199.5, 192.9,

187.2, 179.3, 163.7, 156.2, 141.4, 138.5, 130.5, 121.6, 117.2,

112.2, 108.7, 97.4, 79.6, 76.3, 67.0, 42.1 (2C), 35.8, 29.5, 14.1;

HRMS (TOF MS ES+) calcd for C₂₂H₂₂IN₂O₇[M + H]⁺

553.0472; found 553.0469.

(4S,4aS,12aS)-9-Bromo-4-(dimethylamino)-3,10,11,12a-

tetrahydroxy-6-methyl-1,12-dioxo-1,4,4a,5,12,12a-hexahy-

drotetracene-2-carboxamide hydrochloride (9-Br-aTC, 9).

To a clean, dry round-bottom ﬂask, equipped with a stirbar

and argon inlet, was added liquid bromine (13 μL, 0.2592

mmol, 1.2 equiv), acetic acid (10 mL), and triﬂuoroacetic acid

(2 μL, 0.0216 mmol, 0.1 equiv). The mixture was heated to 50

°C (over 20 min), at which point anhydrotetracycline

hydrochloride (100 mg, 0.216 mmol) was added in one

portion. The reaction mixture stirred at 50 °C for 30 min;

then, the reaction was cooled to room temperature and stirred

at room temperature for 4 h (monitored by LCMS). After the

reaction was complete, the crude mixture was concentrated

under reduced pressure to provide the crude product, which

was reconstituted with aqueous HCl to provide the title

compound (0.1165 g, 0.215 mmol, quantitative crude yield) as

an orange solid HCl salt. FTIR (neat) 3295, 3078, 1618, 1553,

1393, 1373, 1317, 1224, 1124, 1058, 699, 620 cm⁻¹; mp 203−

1

204 °C (decomposed); H NMR (500 MHz, DMSO-d₆) δ

9.64 (s, 1H), 9.22 (s, 1H), 7.84 (d, J = 9.0 Hz, 1H), 7.42 (d, J

= 9.1 Hz, 1H), 4.44 (d, J = 4.0 Hz, 1H), 3.56 (dd, J = 17.7, 4.9

Hz, 1H), 3.46 (dd, J = 9.9, 4.7 Hz, 1H), 3.07 (d, J = 14.5 Hz,

1H), 2.91 (s, 6H), 2.39 (s, 3H); ¹³C NMR (126 MHz, DMSO-

d₆) δ 200.6, 192.7, 187.0, 172.1, 161.8, 153.4, 137.7, 135.8,

131.6, 122.3, 116.7, 112.6, 109.3, 104.5, 97.3, 76.4, 67.0, 42.1

(2C), 35.3, 21.1, 14.0; HRMS (TOF MS ES+) calcd for

C₂₂H₂₂BrN₂O₇[M + H]⁺505.0610; found 505.0610.

In Vitro Characterization of aTC and aTC Analogue

Inhibition. Half-maximal inhibitory concentrations (IC₅₀) for

the aTC and aTC analogue inhibition of Tet(50), Tet(X), and

Tet(X)_3 were determined from the nonlinear regression

analysis of initial velocities of tetracycline degradation in the

MICA_combo

MICA_alone

MICB

combo

MICB_alone

FICI =

+

(1)

where FICI > 1 indicates antagonism, FICI = 1 indicates

additivity, and FICI < 1 indicates synergy.

Kinetic Characterization of aTC and aTC Analogue

Degradation by Tet(X). The kinetic characterization of the

degradation of aTC and aTC analogues by Tet(X) was

monitored by optical absorbance spectroscopy (Carey UV−

visible spectrophotometer) coupled to LCMS detection

(Agilent 6130 single quadrupole instrument with G1313

autosampler, G1315 diode array detector, and 1200 series

M

DOI: 10.1021/acsinfecdis.8b00349

ACS Infect. Dis. XXXX, XXX, XXX−XXX

ACS Infectious Diseases

Article

solvent module and separated using a Phenomenex Gemini

C18 column, 50 × 2 mm (5 μm) with guard column cassette

and a linear gradient of 0% acetonitrile and 0.1% formic acid to

95% acetonitrile and 0.1% formic acid over 20 min at a ﬂow

rate of 0.5 mL/min before analysis by electrospray ionization

(ESI+)). Reactions (in duplicate) were prepared in 100 mM

TAPS buﬀer (pH 8.5) with an NADPH regenerating system

(40 mM glucose-6-phosphate, 4 mM NADP⁺, 1 mM MgCl₂, 4

U/mL glucose-6-phosphate dehydrogenase), 28.0 μM sub-

strate (aTC or corresponding analogue), and 0.24 μM enzyme.

Reaction progress was monitored by optical absorbance

spectroscopy (280−550 nm, 1 nm and 30 min intervals)

over 3.5 h, where 150 μL of reaction sample was removed at 30

min intervals and quenched with 600 μL volumes of quench

solution (1:1 acetonitrile/0.25 M aqueous HCl). The

quenched samples were centrifuged (5000 rpm, room

temperature) for 5 min, and 600 μL of supernatant was

transferred to an LCMS-compatible vial containing Fmoc-

alanine internal standard (2.21 μM ﬁnal concentration) and

analyzed by LCMS (reverse-phase HPLC, C18-silica, gradient

0−95% CH₃CN/H₂O, 0.5 mL/min ﬂow rate). Substrate

masses [M + H]⁺and hydroxylated product masses [M−OH

+H]⁺were extracted from the crude mass chromatogram and

normalized to the internal standard [M + H]⁺counts. No

enzyme controls were performed for each aTC analogue

screened and showed no signiﬁcant nonenzymatic degradation

over the course of the observable reaction. Degradation of 7-I-

aTC and 9-Br-aTC at extended solution times (overnight)

showed a decrease in LCMS extracted ion counts for both

analogues, suggesting some nonenzymatic degradation over

longer reaction times.

consumption of NADPH by Tet(50), images for the

qualitative colorimetric detection of Tet- and aDem-

promoted hydrogen peroxide formation by Tet(50), and

NMR spectra of synthesized compounds 6−9 (PDF)

AUTHOR INFORMATION

Corresponding Authors

*E-mail: wencewicz@wustl.edu. (T.A.W.)

*E-mail: dantas@wustl.edu. (G.D.)

*E-mail: niraj.tolia@nih.gov. (N.H.T.)

■

ORCID

Jana L. Markley: 0000-0001-6855-7161

Andrew J. Gasparrini: 0000-0002-4551-4633

Hirdesh Kumar: 0000-0002-8488-3001

Niraj H. Tolia: 0000-0002-2689-1337

Gautam Dantas: 0000-0003-0455-8370

Timothy A. Wencewicz: 0000-0002-5839-6672

Author Contributions

^∇These authors jointly supervised this work. The manuscript

was written through the collaborative contributions of all

authors. All authors have given approval to the ﬁnal version of

the manuscript.

Funding

This research is supported by the National Institute of Allergy

and Infectious Diseases (NIAID-NIH R01-123394). N.T. is

supported by the Intramural Research Program of the National

Institute of Allergy and Infectious Diseases, National Institutes

of Health. J.L.M. is supported by the W. M. Keck Program in

Molecular Medicine.

Notes

Qualitative Detection of aDem-promoted Hydrogen

Peroxide Formation by Tet(50). Qualitative colorimetric

detection of aDem-promoted hydrogen peroxide formation by

Tet(50) was performed using an aqueous Pierce Quantitative

Peroxide Assay kit (ThermoScientiﬁc). Reaction samples were

prepared in 100 mM TAPS buﬀer (pH 8.5) with 252 μM

NADPH, 2.52 mM MgCl₂, 25 μM substrate (either aDem or

Tet), and 0.4 μM enzyme. After the addition of enzyme, the

reaction was mixed manually by pipet, and the reaction was

monitored by optical absorbance spectroscopy (280−550 nm,

1 nm and 0.1 min scan intervals) over 8 min. At 8 min, 100 μL

of reaction solution was added to a detection Eppendorf

containing 1000 μL of working reagent (prepared according to

speciﬁcations for Pierce Quantitative Peroxide Assay kit). The

detection Eppendorf was incubated for 20 min at room

temperature to result in the observed color changes reported in

the main text (see Figure 10C and Supporting Information

Figure 5).

The authors declare no competing ﬁnancial interest.

ACKNOWLEDGMENTS

■

The authors would like to thank Washington Univ. in St. Louis

(WUSTL), Washington Univ. School of Medicine, and the

National Institutes of Health for their support of this research

and our programs. In particular, N.T. would like to thank the

Intramural Research Program of the National Institute of

Allergy and Infectious Diseases, National Institutes of Health,

for the support of his program. We would also like to

acknowledge J. Kao and M. Singh (WUSTL, Dept. of

Chemistry) for their assistance with NMR experiments and

B. Evans (Danforth Plant Science Center, St. Louis, MO) and

his team for their assistance in acquiring high-resolution mass

spectra for all synthesized compounds. In addition, J.L.M.

would like to acknowledge the WM Keck Postdoctoral

Program in Molecular Medicine for funding support of her

postdoctoral fellowship.

ASSOCIATED CONTENT

■

ABBREVIATIONS

■

S

* Supporting Information

Tet, tetracycline; CTc, chlortetracycline; Dem, demeclocy-

cline; Oxy, oxytetracycline; aTC, anhydrotetracycline; aCTc,

anhydrochlortetracycline; aDem, anhydrodemeclocycline; 7-I-

aTC, 7-iodoanhydrotetracycline; 9-Br-aTC, 9-bromoanhydro-

tetracycline; FMO, ﬂavin-dependent monooxygenases;

NADPH, nicotinamide adenine dinucleotide phosphate

reduced form; FAD, ﬂavin adenine dinucleotide; HPLC,

high-performance liquid chromatography; LCMS, liquid

chromatography−mass spectrometry; NMR, nuclear magnetic

resonance spectroscopy; IR, infrared spectroscopy; mp,

melting point

The Supporting Information is available free of charge on the

ACS Publications website at DOI: 10.1021/acsinfec-

dis.8b00349.

Relevant strains, sequences, plasmids, and primers, SDS-

page gel images of puriﬁed enzyme, Lineweaver−Burk

plots, methods for the aTC inhibition of the tetracycline

destructase-mediated degradation of tetracycline, ex-

panded data tables for whole cell inhibition assays of E.

coli expressing tetracycline destructase enzymes, in vitro

characterization of the aDem- and Tet-promoted

N

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ACS Infect. Dis. XXXX, XXX, XXX−XXX

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