Structural basis of rifampin inactivation by rifampin

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Structural basis of rifampin inactivation by rifampin phosphotransferase Xiaofeng Qia,b, Wei Lina,1, Miaolian Maa, Chengyuan Wanga,b, Yang Hea,b, Nisha Heb,c, Jing Gaod, Hu Zhoud, Youli Xiaoc, Yong Wangc, and Peng Zhanga,2 a National Key Laboratory of Plant Molecular Genetics, Chinese Academy of Sciences Center for Excellence in Molecular Plant Sciences, Institute of Plant Physiology and Ecology, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences, Shanghai 200032, China; bUniversity of Chinese Academy of Sciences, Beijing 100039, China; cChinese Academy of Sciences Key Laboratory of Synthetic Biology, Chinese Academy of Sciences Center for Excellence in Molecular Plant Sciences, Institute of Plant Physiology and Ecology, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences, Shanghai 200032, China; and dChinese Academy of Sciences Key Laboratory of Receptor Research, Shanghai Institute of Materia Medica, Chinese Academy of Sciences, Shanghai 201203, China

Rifampin (RIF) is a first-line drug used for the treatment of tuberculosis and other bacterial infections. Various RIF resistance mechanisms have been reported, and recently an RIF-inactivation enzyme, RIF phosphotransferase (RPH), was reported to phosphorylate RIF at its C21 hydroxyl at the cost of ATP. However, the underlying molecular mechanism remained unknown. Here, we solve the structures of RPH from Listeria monocytogenes (LmRPH) in different conformations. LmRPH comprises three domains: an ATP-binding domain (AD), an RIF-binding domain (RD), and a catalytic His-containing domain (HD). Structural analyses reveal that the C-terminal HD can swing between the AD and RD, like a toggle switch, to transfer phosphate. In addition to its catalytic role, the HD can bind to the AD and induce conformational changes that stabilize ATP binding, and the binding of the HD to the RD is required for the formation of the RIF-binding pocket. A line of hydrophobic residues forms the RIF-binding pocket and interacts with the 1-amino, 2-naphthol, 4-sulfonic acid and naphthol moieties of RIF. The R group of RIF points toward the outside of the pocket, explaining the low substrate selectivity of RPH. Four residues near the C21 hydroxyl of RIF, His825, Arg666, Lys670, and Gln337, were found to play essential roles in the phosphorylation of RIF; among these the His825 residue may function as the phosphate acceptor and donor. Our study reveals the molecular mechanism of RIF phosphorylation catalyzed by RPH and will guide the development of a new generation of rifamycins.

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antibiotic resistance rifampin phosphotransferase molecular mechanism toggle switch

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the target, the RNAP β-subunit; these mutations significantly decrease the binding of rifamycins and thus neutralize the antibiotic activity (10). Another prevalent resistance strategy adopted by bacteria is modification of the rifamycins, such as ADP ribosylation, glycosylation, and phosphorylation (11–13). These covalent modifications occur on the critical hydroxyls of the 1-amino, 2-naphthol, 4-sulfonic acid (ansa) chain of rifamycins and thus make rifamycins unable to fit into the binding pocket on RNAP. Additional resistance mechanisms have been reported also (14–16). Antibiotic resistance is a great threat to the treatment of infectious disease, and understanding the molecular mechanisms of resistance no doubt will help guide the development of a new generation of drugs (17, 18). A number of studies have been carried out to understand rifamycin resistance caused by RNAP mutations (1, 19). However, the proteins and mechanisms involved in the covalent modifications of rifamycins remain largely unknown. Recently, an antibiotic-resistance protein family, RIF phosphotransferase (RPH), was found to inactivate RIF by phosphorylating it at the hydroxyl attached to the C21 of its ansa chain. RPHs in heterologous bacteria are able to inactivate diverse Significance Rifampin phosphotransferases (RPH) belong to a recently identified antibiotic-resistance protein family that inactivates rifampin, the first-line drug against tuberculosis, by phosphorylation. By determining the structures of RPH from Listeria monocytogenes (LmRPH) in different conformations, we reveal a toggle-switch mechanism of the LmRPH catalytic process in which the C-terminal His domain swings between the ATPbinding domain and the rifampin-binding domain to transfer phosphate from ATP to rifampin. These structures explain the low substrate selectivity of RPH for the rifamycins. The residues involved in rifampin phosphorylation are identified also. The structural mechanism revealed in this study will guide the development of a new generation of rifamycins.

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ifamycins are a group of natural or semisynthetic antibiotics used for treating a broad repertoire of bacterial infections. These compounds bind directly to the β-subunit of bacterial RNA polymerase (RNAP) at a highly conserved region, blocking the exit tunnel for RNA elongation and thus inhibiting the process of transcription (1). The first member of the rifamycins to be described, rifamycin B, was extracted from the soil actinomycete Amycolatopsis mediterranei (2). The natural product had modest antibiotic activity, but semisynthetic derivatives of the rifamycin family have proven highly successful in the clinic (3). The best-known member of the rifamycin family, rifampin (RIF), was introduced to the clinic in 1968; it is highly effective against Mycobacterium tuberculosis and greatly shortens the duration of tuberculosis therapy (4). At present, RIF continues to be a first-line drug for the treatment of tuberculosis (5). Through the years additional derivatives have been developed to treat a wider range of bacterial infections (3); for example, rifalazil serves as an effective antibiotic against Chlamydia-based persistent infections (6), and rifaximin is used to treat travelers’ diarrhea and irritable bowel syndrome (7, 8). Extensive use of rifamycins has led to the development of bacterial resistances (9). In M. tuberculosis and other mycobacteria the most common resistance mechanisms are point mutations of

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Author contributions: X.Q. and P.Z. designed research; X.Q., W.L., M.M., C.W., Y.H., N.H., J.G., and H.Z. performed research; X.Q., W.L., M.M., C.W., Y.H., N.H., J.G., H.Z., Y.X., Y.W., and P.Z. analyzed data; and P.Z. wrote the paper. The authors declare no conflict of interest. This article is a PNAS Direct Submission. A.S. is a guest editor invited by the Editorial Board. Data deposition: The structural factors and coordinates reported in this paper have been deposited in the Protein Data Bank (PDB) [PDB ID codes 5HV1 (LmRPH–ANP–RIF), 5HV2 (LmRPHG527Y–apo), 5HV3 (LmRPHG527Y–ANP), and 5HV6 (LmRPH–AD)]. 1

Present address: Waksman Institute of Microbiology, Rutgers, The State University of New Jersey, Piscataway, NJ 08854.

2

To whom correspondence should be addressed. Email: [email protected].

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10. 1073/pnas.1523614113/-/DCSupplemental.

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Edited by Alexander Serganov, New York University, New York, NY, and accepted by the Editorial Board March 1, 2016 (received for review November 30, 2015)

at the cost of ATP and confers the bacteria with high-level resistance to RIF.

Fig. 1. Activity of LmRPH. (A) In vitro LmRPH reaction products analyzed by HPLC with the RIF detection program. (B) Identification of the two peaks in A by LC-MS. (Left) Peak 1. (Right) Peak 2. (C) The samples in A were analyzed by HPLC with a nucleotide-detection program. (D) E. coli growth assay. Bacteria transformed with LmRPH or vector were cultured in solid LB medium complemented with 0, 10, 100, or 1,000 μg/mL RIF.

clinically used rifamycins with great efficiency (13). Bioinformatic analyses suggest that RPHs are widespread in both pathogenic and nonpathogenic bacteria. The RPH protein contains three domains (listed from the N terminus to the C terminus): the ATP-binding domain (AD), the RIF-binding domain (RD), and the His domain (HD), which contains a conserved His residue essential for phosphate transfer. This architecture is similar to that of phosphoenolpyruvate (PEP) synthase, which also contains three domains, an ATP-binding domain, a catalytic His domain, and a pyruvate-binding domain and catalyzes the reversible conversion of ATP, water, and pyruvate to AMP, inorganic phosphate (Pi), and PEP (20). Apart from this information, little is known about RPHs. Here we report the crystal structures of RPH from Listeria monocytogenes (LmRPH) in different catalytic conformations. Structural and functional analyses reveal the molecular basis of substrate binding, phosphate transfer, and RIF phosphorylation by LmRPH. This study identifies the molecular mechanism of RIF phosphorylation and will guide strategies to overcome RPHmediated rifamycin resistance. Results Characterization of LmRPH. The gene encoding RPH from LmRPH was cloned, expressed in Escherichia coli, and purified. The enzymatic activity of the recombinant LmRPH was tested in a reaction system containing the substrates RIF and ATP, and the products were separated by HPLC. As the reaction proceeded, the amount of RIF gradually decreased, accompanied by the increase of a subsequent product peak (Fig. 1A), which was identified as phosphorylated RIF (RIF-P) by LC-MS (Fig. 1B and Fig. S1). The other substrate, ATP, was converted into AMP rather than ADP until RIF was phosphorylated completely (Fig. 1C). To examine ability of LmRPH to inactivate RIF in vivo, E. coli BL21 (DE3) cells transformed with pQE80L-LmRPH were cultured on solid LB medium containing a gradient of RIF concentrations. The results show that E. coli growth is strongly inhibited by 10 μg/mL RIF, but the introduction of LmRPH at concentrations greater than 1,000 μg/mL confers resistance to RIF (Fig. 1D). These data suggest that LmRPH catalyzes the conversion of RIF to RIF-P 3804 | www.pnas.org/cgi/doi/10.1073/pnas.1523614113

Structures of LmRPH at Different Conformations. The LmRPH protein was purified further using gel filtration before crystallization, and the two major peaks (peaks 1 and 2) observed were both confirmed to be LmRPH proteins with similar molecular radius/mass by dynamic light scattering (DLS) (Fig. S2). This result indicates that LmRPH might have different conformations in solution. However, we could obtain diffractable crystals only with LmRPH protein from peak 1. The LmRPH structure was solved in an AMP–PNP (ANP)-, Mg2+- and RIF-bound state (LmRPH–ANP–RIF) by the single-wavelength anomalous dispersion (SAD) method. The overall structure adopts a saddle-like shape, with the AD (residues 1–315) and the RD (residues 323–748) forming two flaps of the saddle. The C-terminal HD (residues 771–867) binds to the RD from the concave side of the “saddle” (Fig. 2A). The ATP analog, ANP, binds in a cleft of the AD from the concave side of the saddle, and RIF binds in a pocket of the RD from the convex side. The two substrate-binding sites are about 49 Å apart, leaving ample room for the HD to play an indispensable role in catalysis. Intriguingly, the HD is linked to the RD by a long, flexible linker (residues 749–770) through which the HD might swing between the AD and RD to transfer phosphate from ATP to RIF. In the LmRPH–ANP–RIF structure, the HD contacts the RD mainly through hydrophobic interactions (Fig. S3). We introduced mutations at this interface to disrupt these interactions and found that LmRPH proteins containing these mutations had gel-filtration profiles different from those of wild-type proteins (Fig. 2B), i.e., two major peaks for wild-type proteins vs. one major peak for mutants. Accordingly, LmRPH-G527A, LmRPHG527S, and LmRPH-G527Y mutants have much decreased or no RIF-phosphorylation activity in vitro and reduced or no RIF resistance in vivo (Fig. 2 C and D). These data suggest that, instead of two conformations, the LmRPH-G527A, LmRPHG527S, and LmRPH-G527Y mutants tend to adopt one conformation in solution. Indeed, the structures of LmRPHG527Y in both the apo form (LmRPHG527Y–apo) and the ANP-bound form (LmRPHG527Y–ANP) were in a conformation different from that of the LmRPH–ANP–RIF structure (Fig. 2 E and F), with the HD binding to the AD from the concave side. Notably, even though the interactions between the HD and the RD and between the HD and the AD have been observed in different LmRPH conformations, we could not quantify the interaction affinities between the individually purified HD and RD or AD in isothermal titration calorimetry (ITC) experiments; the interactions are too weak to be detected by ITC (Fig. S4), suggesting that the interactions between the HD and the RD and between the HD and the AD are dynamic. Our structural data demonstrate that the HD can swing between the RD and the AD, as is required for LmRPH catalysis, more specifically, for the transfer of phosphate from ATP to RIF. ATP Binding with the AD Is Stabilized by the HD. Although the LmRPHG527Y–ANP and LmRPH–ANP–RIF structures adopt different conformations, both can bind with ANP, prompting us to determine their ATP-binding affinities. The results show that the ATP-binding affinity of LmRPHG527Y (Kd = 0.43 μM) is higher than that of the wild-type protein (Kd = 1.11 μM), but the separated AD itself cannot bind with ATP (the affinity is too low to be detected by ITC) (Fig. 3A), suggesting that the HD may contribute to the ATP binding. To resolve this mystery, we solved the structure of the AD in apo state (LmRPH–AD) and compared it with the LmRPHG527Y–apo structure (Fig. 3B). The AD contains two subdomains, subdomain I (residues 1–183) and subdomain II (residues 190–315), which are connected by a flexible linker (L13, residues 184–189) to form a hinge-like conformation. The binding of the HD with the AD induces significant conformational changes Qi et al.

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Fig. 2. Overall structures of LmRPH at different states. (A) Structure of wild-type LmRPH in complex with RIF, ANP, and Mg2+. The AD, RD, and HD are colored lemon, light blue, and orange, respectively. ANP and RIF are shown as sticks and are colored green and magenta, respectively. Mg2+ is shown as a sphere. (B) Gel-filtration profiles of wild-type LmRPH (red) and the G527A (green), G527S (cyan), and G527Y (blue) mutants. (C) In vitro catalytic activity of Gly527 mutants detected by HPLC. The reaction time of G527A, G527S, and G527Y is 1 h, and that of wild-type LmRPH is 5 min. Proteins were used at 0.5 mg/mL. (D) E. coli growth assay for Gly527 mutants. Bacteria transformed with wild-type LmRPH, vector, G527A, G527S, or G527Y were cultured in solid LB medium complemented with 0, 10, 40, or 80 μg/mL RIF. (E) Structure of LmRPHG527Y in apo form. (F) Structure of LmRPHG527Y in complex with ANP and Mg2+. Color codes in E and F are as A.

in both subdomain I and II: helices α4, α5, and α8 of subdomain I undergo a dramatic shift toward the HD, leading to hydrophobic interactions between α8 (subdomain I) and α31 (HD), and helix α9 unwinds to bind with the HD, also through hydrophobic interactions (Fig. 3B). As a result, the conformations of subdomains I and II of the AD are stabilized by the binding of the HD, as is the ATP-binding cleft between these two subdomains. These findings explain why the HD is required for tight binding of ATP. In the LmRPHG527Y mutant, the HD is restricted from binding with the RD; therefore the binding affinity of LmRPHG527Y is higher than that of the wild-type protein (Fig. 3A). The binding of ANP to the cleft of the AD results in further conformational changes in the surrounding structural elements, as can be seen clearly by comparing the LmRPHG527Y–apo and LmRPHG527Y–ANP structures (Fig. 3C). After ANP binding, β3–β4, which adopts a loop conformation in the LmRPHG527Y– apo structure, forms a five-stranded antiparallel β-sheet with β1–β2–β5, as do β10–β11 in subdomain II. In addition, loop L9 Qi et al.

(residues 123–134) from subdomain I, which is disordered in the absence of ANP, can be seen clearly after ANP binding. ANP binding also induces a conversion of the α9 from subdomain II. The formation of L9 and α9 after ANP binding generates steric repulsions of the HD, thereby weakening the interaction between the AD and at HD, as reflected by the poor electron density of the HD in the LmRPHG527Y–ANP structure (Fig. S5). The structural rearrangements of the AD described above accommodate the tight binding of ANP to the cleft through a number of conserved residues in addition to an Mg2+ (Fig. 3D). Specifically, the adenine ring of ANP forms three hydrogen bonds with the guanidine group of Arg117, the carbonyl oxygen of Gln184, and the side chain of Gln183; the 2′-hydroxyl group of the ANP ribose forms a hydrogen bond with the side chain of Glu297; the α, γ-phosphates of ANP form hydrogen-bonding interactions with residues Arg117, Thr136, Lys22, Arg311, and Gly132; and the β, γ-phosphates of ANP are coordinated with residues Glu297 and Gln309 through the Mg2+. The importance PNAS | April 5, 2016 | vol. 113 | no. 14 | 3805

Fig. 3. ATP-binding site. (A) ATP-binding affinity of the AD (green isotherm), wild-type LmRPH (blue isotherm), and the G527Y mutant (red isotherm) measured by ITC. (B) Conformational changes of the AD induced by HD binding. The LmRPH–AD structure (gray) is superposed with the AD of the LmRPHG527Y–apo structure (lemon). The L13 loop connecting subdomains I and II is highlighted in red. The interaction interfaces between the AD and HD (orange) are shown in zoom-in views, and residues constituting the interface are shown with side chains. (C) Conformational changes induced in the AD and HD by ANP binding. The AD (lemon) and HD (orange) of the LmRPHG527Y–apo structure are superposed with those of the LmRPHG527Y-ANP structure (light blue). Structural elements undergoing conformational changes after ANP binding are colored in red. (D) Residues constituting the ATP-binding site. ANP (green) and residues (lemon) are shown as sticks, and Mg2+ is shown as a lemon sphere. Coordination and hydrogen bonds are shown as dashed lines. (E) In vitro catalytic activity of ATP-binding site mutants detected by HPLC. The amounts of enzymes used in the assays of the K22A, R117A, E297A, T136A, Q309A, and R311A mutants are 10× those used in assays of wild-type LmRPH. (F) E. coli growth assay for ATP-binding site mutants. Bacteria transformed with wild-type LmRPH, vector, or mutants were cultured in solid LB medium complemented with 0, 10, 40, 80, or 320 μg/mL RIF.

of these residues was validated by an in vitro enzymatic activity assay. The results show that the activity of Q183A is slightly lower than that of the wild-type LmRPH, the activities of T136A, Q309A, and R311A mutants are significantly reduced, and those of other mutants (K22A, R117A, and E297A) are extremely low (Fig. 3E). Accordingly, the RIF-resistance levels of these mutants are reduced to different extents, except for the Q183A mutant, in which resistance is comparable to that in wild-type LmRPH (Fig. 3F). Both the RD and HD Are Involved in Rif Binding. The structure of the RD can be divided further into three subdomains: subdomain I (α12–16, 28–30, and β14–18), II (α17–20 and 26–27), and III (α21–25) (Fig. 4A). Searches of the Protein Data Bank failed to identify any entry that is structurally homologous to the RD, suggesting that the RD represents a previously unidentified structural fold related to RIF binding. In the LmRPH–ANP–RIF structure, the HD binds with all three subdomains of the RD from the concave side and forms the RIF-binding pocket with subdomains I and II of RD. Distinct from the AD ATPbinding cleft, which faces the concave side of LmRPH, the opening of the RIF-binding pocket faces the convex side (Figs. 2A and 4A). Structural comparison of LmRPH–ANP–RIF with LmRPHG527Y-ANP reveals significant conformational changes at the RIF-binding pocket (Fig. 4B). Specifically, the binding of the HD with the RD pushes away α14–α16 and connecting loops of the RD, leading to a rearrangement of the surrounding inward-facing residues that create an RIF-binding pocket (Fig. 4 B–D). These structural observations suggest that both the RD and HD are involved in RIF binding. Consistently, we found that the RD alone is not sufficient to bind with RIF, but the RD and HD together can bind RIF with high affinity (Kd = 79.4 μM) (Fig. 4E). (The binding affinity of RIF with full-length LmRPH 3806 | www.pnas.org/cgi/doi/10.1073/pnas.1523614113

changes over time; therefore we used the RD and HD for the detection of RIF binding.). The RIF-binding pocket is comprised mainly of hydrophobic residues. Residues Val333, Met359, and Val368 constitute a hydrophobic patch and contact the naphthol ring of RIF through van der Waals forces; residues Ile331, Ile370, Ile394, Met383, Leu387, Met823, Met491, Met488, Leu478, and Met673 stabilize the ansa chain of RIF through hydrophobic interactions (Fig. 4 F and G). Mutations V333A, V368A, M383A, or M673A increase the size of the pocket and reduce the phosphorylation activity of LmRPH, whereas V333W or V368W causes steric conflict and almost abolishes the activity (Fig. 4H). The R group of RIF points toward the opening of the pocket and packs against residues Pro356 and Phe479; replacement of either of these two residues with alanine has only minor effects on the phosphorylation activity and RIF binding (Fig. 4F and Table S1). This finding likely explains why RPH can phosphorylate various members of the rifamycin family that differ primarily at the R group (13). Phosphorylation of RIF. The LmRPH–ANP–RIF complex structure allows us to examine the phosphorylation site of RIF. The previously identified phosphorylation site of RIF, C21 hydroxyl, is about 6.7 Å away from residue His825 of the HD (Fig. 5A). A water molecule between His825 and C21 hydroxyl forms a hydrogen bond with the C21 hydroxyl. When we modeled the product RIF-P into the structure (Fig. 5B), we found that the phosphate group of RIF-P could form four hydrogen bonds with Lys670, Arg666, and Gln337 and that the distance between RIF-P and residue His825 is about 4 Å. These structural observations suggest that the HD residue His825 and residues Lys670, Arg666, and Gln337 are involved in RIF phosphorylation. To verify this possibility, we mutated these four residues to alanine and Qi et al.

determined their activities. The results show that the H825A, R666A, and K670A mutants lose the ability to phosphorylate RIF both in vitro and in vivo, and Q337A has significantly reduced activity (Fig. 5 C and D). Notably, residue His825 is highly conserved among RPHs, reminiscent of the catalytic His residue in PEP synthase (20). Using the Phos-tag SDS/PAGE experiment, we found that LmRPH is phosphorylated in the presence of ATP but the H825A mutant is not (Fig. 5E). The phosphorylation of residue His825 is confirmed in an MS analysis (Fig. S6). These results suggest that this conserved His residue also may function as a phosphate acceptor and donor in the phosphorylation of RIF. In

addition, positively charged residues often function as catalytic bases to abstract a proton at the reaction centers of phosphatetransfer enzymes and other enzymes, including MAPK, phosphothreonine lyase, IMP dehydrogenase, pectate/pectin lyases, fumarate reductase, and L-aspartate oxidase (21–23), suggesting that Lys670 and Arg666 might be candidate residues for the catalytic bases of LmRPH. Discussion In this work we captured two major conformational states of the LmRPH catalytic process: a conformation in which the HD binds

Fig. 5. Catalytic center of RIF phosphorylation. (A) Catalytic site of RIF phosphorylation. Residues His825, Lys670, Arg666, and Gln337 and RIF are shown as sticks; the water molecule is shown as a red sphere. Distances between the water molecule and surrounding residues are shown as dashed lines. (B) Catalytic site with a modeled RIF-P. Distances between the phosphate group and residues are shown as dashed lines. (C) In vitro catalytic activity of H825A, K670A, R666A, and Q337A detected by HPLC. (D) E. coli growth assay of the mutants in C. Bacteria transformed with wild-type LmRPH, vector, or mutants were cultured in solid LB medium complemented with 0 or 10 μg/mL RIF. (E) Phosphorylation analysis of wild-type LmRPH and the H825A mutant using Phos-tag SDS PAGE. LmRPH-P, phosphorylated LmRPH.

Qi et al.

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Fig. 4. The RIF-binding pocket. (A) Structure of the RD (in LmRPH–ANP–RIF). The gray dashed lines separate three subdomains (I, II, and III) of the RD. RIF is shown as magenta sticks. (B) Conformational changes of the RD induced by HD binding. The RD (light blue) and HD (orange) of the LmRPH–ANP–RIF structure was superposed with that of the LmRPHG527Y–apo structure (gray). α14–16, which undergo conformational changes after HD binding, are highlighted in red. (C) Surface view of the RIF-binding pocket in the LmRPHG527Y–apo structure. (D) Surface view of the RIF-binding pocket in the LmRPH–ANP–RIF structure. The RD, HD, and α14–16 are colored light blue, orange, and red, respectively. (E) The RIF-binding affinity of the RD and RD–HD measured by ITC. Binding isotherms for RD and RD–HD are colored green and red, respectively. (F) Residues at the RIF-binding site. RIF (magenta) and interacting residues (light blue) are shown as sticks. (G) Chemical structure of RIF. The naphthol ring and ansa chain of RIF are shown in pink and blue, respectively, and the R group is outlined by a dashed box. (H) In vitro catalytic activity of RIF-binding mutants detected by HPLC.

the pyruvate-binding domain to transfer phosphate from ATP to pyruvate (Fig. S7) (24–26). Based on the structure of RIF bound to the Thermus aquaticus RNAP core enzyme, the C21 hydroxyl of RIF points toward the inside of the RIF-binding pocket and forms hydrogen-bonding interactions with nearby residues (1). Phosphorylation of this hydroxyl may lead to steric clash, thereby weakening or abolishing the binding of RIF to RNAP and ultimately resulting in resistance to RIF. RPHs are widespread among Bacillales, Actinomycetales, and Clostridiales, which include many human pathogens. Searches of the pathogenic bacterial genomes of Bacillus anthracis, Enterococcus faecalis, Nocardia brasiliensis, and Listeria monocytogenes all reveal RPH genes, which may limit the clinical use of rifamycins against these organisms. Our mechanistic study of LmRPH provides feasible strategies, such as developing highaffinity RPH inhibitors or new RIF derivatives that are not susceptible to RPH, to overcome RPH-mediated resistance. Materials and Methods

Fig. 6.

Catalytic process of LmRPH-mediated RIF phosphorylation.

to the AD, and a conformation in which the HD binds to the RD. Structural-based analysis confirmed that the HD functions as a toggle switch, swinging between the two distant domains. When binding to the AD, the HD facilitates ATP binding and hydrolysis, grabbing a phosphate by residue His825. Then the HD swings over to the RD, facilitating RIF binding and initiating RIF phosphorylation. The dynamic nature of the LmRPH protein enables the smooth transition between these two conformational states (Fig. 6). This mechanism resembles that of three-domain pyruvate orthophosphate dikinase (PPDK) enzymes in which the His domain swivels between the nucleotide-binding domain and 1. Campbell EA, et al. (2001) Structural mechanism for rifampicin inhibition of bacterial rna polymerase. Cell 104(6):901–912. 2. Sensi P, Margalith P, Timbal MT (1959) Rifomycin, a new antibiotic; preliminary report. Farmaco, Sci 14(2):146–147. 3. Aristoff PA, Garcia GA, Kirchhoff PD, Showalter HD (2010) Rifamycins–obstacles and opportunities. Tuberculosis (Edinb) 90(2):94–118. 4. Sensi P (1983) History of the development of rifampin. Rev Infect Dis 5(Suppl 3): S402–S406. 5. Getahun H, Matteelli A, Chaisson RE, Raviglione M (2015) Latent Mycobacterium tuberculosis infection. N Engl J Med 372(22):2127–2135. 6. Rothstein DM, van Duzer J, Sternlicht A, Gilman SC (2007) Rifalazil and other benzoxazinorifamycins in the treatment of chlamydia-based persistent infections. Arch Pharm (Weinheim) 340(10):517–529. 7. Huang DB, DuPont HL (2005) Rifaximin–a novel antimicrobial for enteric infections. J Infect 50(2):97–106. 8. Schoenfeld P, et al. (2014) Safety and tolerability of rifaximin for the treatment of irritable bowel syndrome without constipation: A pooled analysis of randomised, double-blind, placebo-controlled trials. Aliment Pharmacol Ther 39(10):1161–1168. 9. Dorman SE, Chaisson RE (2007) From magic bullets back to the magic mountain: The rise of extensively drug-resistant tuberculosis. Nat Med 13(3):295–298. 10. Goldstein BP (2014) Resistance to rifampicin: A review. J Antibiot (Tokyo) 67(9): 625–630. 11. Baysarowich J, et al. (2008) Rifamycin antibiotic resistance by ADP-ribosylation: Structure and diversity of Arr. Proc Natl Acad Sci USA 105(12):4886–4891. 12. Spanogiannopoulos P, Thaker M, Koteva K, Waglechner N, Wright GD (2012) Characterization of a rifampin-inactivating glycosyltransferase from a screen of environmental actinomycetes. Antimicrob Agents Chemother 56(10):5061–5069. 13. Spanogiannopoulos P, Waglechner N, Koteva K, Wright GD (2014) A rifamycin inactivating phosphotransferase family shared by environmental and pathogenic bacteria. Proc Natl Acad Sci USA 111(19):7102–7107. 14. Tupin A, et al. (2010) Resistance to rifampicin: At the crossroads between ecological, genomic and medical concerns. Int J Antimicrob Agents 35(6):519–523. 15. Wright GD (2005) Bacterial resistance to antibiotics: Enzymatic degradation and modification. Adv Drug Deliv Rev 57(10):1451–1470.

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See SI Materials and Methods for details. In general, LmRPH protein was expressed in E. coli and purified to homogeneity for crystallization. All data were collected and processed with HKL3000 (27). The structures were determined using programs in Phenix (28), and structural models were built with Coot (29). The products of the enzymatic assay were detected with HPLC/MS. The substrate-binding affinity was measured with ITC. Data collection and refinement statistics are summarized in Table S2. ACKNOWLEDGMENTS. We thank the staff members at the BL19U beamline of the National Center for Protein Science Shanghai and the BL17U beamline of the Shanghai Synchrotron Radiation Facility for technical assistance in data collection and the staff at the core facility center of the Institute of Plant Physiology and Ecology for MS experiments and analysis. This work was supported by National Natural Science Foundation of China Grant 31322016 and National Program on Key Basic Research Projects Grant 2015CB910900 and by funding from the National Key Laboratory of Plant Molecular Genetics, CAS Center for Excellence in Molecular Plant Sciences, Institute of Plant Physiology and Ecology, Shanghai Institutes for Biological Sciences, CAS.

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