Article13 March 2007Open Access Gyrase inhibitors induce an oxidative damage cellular death pathway in Escherichia coli Daniel J Dwyer Daniel J Dwyer Program in Molecular Biology, Cell Biology and Biochemistry, Boston University, Boston, MA, USA Center for BioDynamics and Center for Advanced Biotechnology, Boston University, Boston, MA, USA Search for more papers by this author Michael A Kohanski Michael A Kohanski Center for BioDynamics and Center for Advanced Biotechnology, Boston University, Boston, MA, USA Department of Biomedical Engineering, Boston University, Boston, MA, USA Boston University School of Medicine, Boston, MA, USA Search for more papers by this author Boris Hayete Boris Hayete Center for BioDynamics and Center for Advanced Biotechnology, Boston University, Boston, MA, USA Bioinformatics Program, Boston University, Boston, MA, USA Search for more papers by this author James J Collins Corresponding Author James J Collins Program in Molecular Biology, Cell Biology and Biochemistry, Boston University, Boston, MA, USA Center for BioDynamics and Center for Advanced Biotechnology, Boston University, Boston, MA, USA Department of Biomedical Engineering, Boston University, Boston, MA, USA Bioinformatics Program, Boston University, Boston, MA, USA Search for more papers by this author Daniel J Dwyer Daniel J Dwyer Program in Molecular Biology, Cell Biology and Biochemistry, Boston University, Boston, MA, USA Center for BioDynamics and Center for Advanced Biotechnology, Boston University, Boston, MA, USA Search for more papers by this author Michael A Kohanski Michael A Kohanski Center for BioDynamics and Center for Advanced Biotechnology, Boston University, Boston, MA, USA Department of Biomedical Engineering, Boston University, Boston, MA, USA Boston University School of Medicine, Boston, MA, USA Search for more papers by this author Boris Hayete Boris Hayete Center for BioDynamics and Center for Advanced Biotechnology, Boston University, Boston, MA, USA Bioinformatics Program, Boston University, Boston, MA, USA Search for more papers by this author James J Collins Corresponding Author James J Collins Program in Molecular Biology, Cell Biology and Biochemistry, Boston University, Boston, MA, USA Center for BioDynamics and Center for Advanced Biotechnology, Boston University, Boston, MA, USA Department of Biomedical Engineering, Boston University, Boston, MA, USA Bioinformatics Program, Boston University, Boston, MA, USA Search for more papers by this author Author Information Daniel J Dwyer1,2,‡, Michael A Kohanski2,3,4,‡, Boris Hayete2,5 and James J Collins 1,2,3,5 1Program in Molecular Biology, Cell Biology and Biochemistry, Boston University, Boston, MA, USA 2Center for BioDynamics and Center for Advanced Biotechnology, Boston University, Boston, MA, USA 3Department of Biomedical Engineering, Boston University, Boston, MA, USA 4Boston University School of Medicine, Boston, MA, USA 5Bioinformatics Program, Boston University, Boston, MA, USA ‡These authors contributed equally to this work *Corresponding author. Center for BioDynamics and Department of Biomedical Engineering, Boston University, 44 Cummington Street, Boston, MA 02215, USA. Tel.: +617 353 0390; Fax: +617 353 5462; E-mail: [email protected] Molecular Systems Biology (2007)3:91https://doi.org/10.1038/msb4100135 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions Figures & Info Modulation of bacterial chromosomal supercoiling is a function of DNA gyrase-catalyzed strand breakage and rejoining. This reaction is exploited by both antibiotic and proteic gyrase inhibitors, which trap the gyrase molecule at the DNA cleavage stage. Owing to this interaction, double-stranded DNA breaks are introduced and replication machinery is arrested at blocked replication forks. This immediately results in bacteriostasis and ultimately induces cell death. Here we demonstrate, through a series of phenotypic and gene expression analyses, that superoxide and hydroxyl radical oxidative species are generated following gyrase poisoning and play an important role in cell killing by gyrase inhibitors. We show that superoxide-mediated oxidation of iron–sulfur clusters promotes a breakdown of iron regulatory dynamics; in turn, iron misregulation drives the generation of highly destructive hydroxyl radicals via the Fenton reaction. Importantly, our data reveal that blockage of hydroxyl radical formation increases the survival of gyrase-poisoned cells. Together, this series of biochemical reactions appears to compose a maladaptive response, that serves to amplify the primary effect of gyrase inhibition by oxidatively damaging DNA, proteins and lipids. Synopsis The topological state of the Escherichia coli chromosome is actively maintained by DNA topoisomerases, which enzymatically modulate the degree of DNA supercoiling by catalyzing strand breakage and rejoining reactions (Champoux, 2001). This activity is critical to the processes of DNA replication (by promoting progression of the replication fork) and RNA transcription (by promoting local melting during initiation); additionally, topoisomerases play important roles in the decatenation of linked DNA and straightening of knotted DNA (Wang, 1996). Perhaps the most well-characterized member of the DNA topoisomerase family is topoisomerase II or DNA gyrase (Gellert et al, 1976; Cozzarelli, 1980). Gyrase is responsible for the introduction of biologically essential negative DNA supercoils and accomplishes this task, in part, by inducing double-stranded DNA breaks in an ATP-dependent reaction (Reece and Maxwell, 1991). DNA gyrase, and more specifically the reaction it catalyzes, is the target of both synthetic quinolone antibiotic- and natural proteic-based inhibition (Bernard and Couturier, 1992; Chen et al, 1996). The interaction between gyrase inhibitors and DNA-bound gyrase results in the formation of a stable open cleavage complex which sterically prohibits the passage of DNA and RNA polymerases, while generating widespread DNA damage (Critchlow et al, 1997; Drlica and Zhao, 1997). The combinatorial effect of replication fork stalling and double-stranded DNA break accumulation is rapid entry into bacteriostasis, and ultimately is considered sufficient to induce cell death. In this work, we performed a series of phenotypic and gene expression studies on E. coli treated with quinolone or norfloxacin or expressing the peptide toxin, CcdB, in order to identify genetic and biochemical events that contribute to gyrase inhibitor-induced cell death. Taking a systems biology approach, we enriched our microarray-derived transcriptome data using biological pathway classifications and transcription factor regulatory connections (Ashburner et al, 2000; Boyle et al, 2004; Camon et al, 2004; Salgado et al, 2006; Faith et al, 2007) to distinguish significantly changing functional units from background gene expression patterns (highlighted in Figure 2). As anticipated, this analysis showed that expression of the DNA damage response and repair program was highly upregulated soon after gyrase inhibition. Unexpectedly, however, our systems-level approach revealed that genes associated with iron uptake, iron–sulfur cluster synthesis and oxidative stress also exhibited significantly increased expression upon norfloxacin treatment and CcdB expression. The results of our gene expression analysis were then used to guide follow-up experiments that explored these varied responses to gyrase poisoning. Using the fluorescent reporter dye, HPF (Setsukinai et al, 2003), we demonstrate that gyrase inhibition by either norfloxacin or CcdB induces the formation of highly deleterious hydroxyl radicals, which oxidatively damage DNA, proteins and membrane lipids. Importantly, using the iron chelator, o-phenanthroline, we show that hydroxyl radical generation occurs through the Fenton reaction (in which hydrogen peroxide is reduced by ferrous iron; Imlay, 2003), and that blockage of the Fenton reaction leads to an increase in the survival of gyrase-inhibited E. coli. To determine the Fenton-reactive iron source contributing to gyrase inhibitor-mediated cell death, we employed a variety of promoter-reporter gene fusions (including constructs that sense changes in the cellular superoxide response, iron regulation and iron–sulfur cluster synthesis) and performed growth studies of several single-gene knockout strains. The results of these experiments highlighted critical steps in the formation of hydroxyl radicals—first, superoxide is indeed generated following inhibition of DNA gyrase; second, a breakdown of iron regulatory dynamics, likely related to iron–sulfur cluster oxidation by superoxide, occurs upon gyrase poisoning; third, and perhaps most importantly, the number and redox status of iron–sulfur clusters present intracellularly is critical to hydroxyl radical generation and thus, gyrase inhibitor-induced cell death. Together, these data have provided for the construction of an oxidative damage cell death pathway model (Figure 8). This series of events appears to constitute a maladaptive response to gyrase inhibition by E. coli, growing in an oxygen-rich environment, which amplifies the primary effect of gyrase poisons like norfloxacin and CcdB. These results may provide new targets for novel antibacterial therapies with enhanced bactericidal activity. Introduction Bacterial chromosomal topology is maintained by the activities of topoisomerase I, topoisomerase IV and DNA gyrase (topoisomerase II) (Champoux, 2001). The gyrase enzyme is responsible for the introduction of negative DNA supercoils in an ATP-dependent manner, and participates in the processes of replication, transcription, repair, recombination and decatenation (Gellert et al, 1976; Cozzarelli, 1980; Wang, 1996). Gyrase is composed of two subunits, gyrA and gyrB, and complexes with its DNA substrate as an A2B2 tetramer (Mizuuchi et al, 1978; Sugino et al, 1980). GyrA catalyzes the concomitant double-stranded breakage and rejoining of DNA phosphodiester bonds following binding and hydrolysis of ATP by GyrB (Gellert, 1981; Reece and Maxwell, 1991). The interaction between gyrase inhibitors and the GyrA subunit converts DNA gyrase into a lesion-inducing agent, which results in widespread generation of double-stranded DNA breaks (Maki et al, 1992; Bernard et al, 1993; Chen et al, 1996). DNA damage formation rapidly induces cell division arrest, initially driving entry into bacteriostasis (Hanawalt, 1966; Jaffe et al, 1985; Drlica and Zhao, 1997). In time, the stable DNA–GyrA–inhibitor complex sterically inhibits replication and transcription by establishing a roadblock to DNA and RNA polymerases, respectively, at stalled replication forks (Kreuzer and Cozzarelli, 1979; Bernard and Couturier, 1992; Willmott et al, 1994; Cox et al, 2000). This deadly barrier impedes lesion repair while preventing new DNA synthesis, eventually killing the bacterium (Critchlow et al, 1997; Drlica and Zhao, 1997; Couturier et al, 1998). In this work, we performed phenotypic, genetic and microarray analyses on gyrase-inhibited Escherichia coli to identify additional contributors to cell death resulting from gyrase poisoning. Taking a systems biology approach, we enriched our gene expression data using transcription factor and functional pathway classifications. This analysis expectedly revealed that the expression of DNA damage response and repair genes was markedly upregulated following inhibition of DNA gyrase and the generation of DNA lesions. Surprisingly, however, we also observed significantly upregulated expression of genes related to superoxide stress, iron–sulfur (Fe–S) cluster synthesis and iron uptake and utilization. The unfortunate byproduct of increased aerobic respiratory activity and oxidative phosphorylation is the increased formation of the reactive oxygen species (ROS), superoxide (Imlay and Fridovich, 1991). Although superoxide itself presents no additional direct threat to DNA, cytosolic dehydratase proteins containing Fe–S clusters are highly susceptible to oxidative attack (Liochev and Fridovich, 1999; Imlay, 2003, 2006). Various methods have been used to show that superoxide-mediated decomposition of Fe–S clusters, following shifts between anaerobic and aerobic growth states, leads to cytoplasmic release of ferrous (II) iron (Keyer and Imlay, 1996; Liochev, 1996). Elevated intracellular concentrations of ferrous iron have been shown to mediate DNA damage either directly (Henle et al, 1999; Rai et al, 2001) or via the formation of additional oxidative molecules (Touati, 2000; Imlay, 2003). Hydroxyl radicals, a highly destructive ROS, are the product of the Fenton reaction (Imlay et al, 1988; Imlay and Linn, 1988), in which ferrous iron mediates the reduction of hydrogen peroxide. These biologically threatening oxidative species are capable of inducing a wide array of mutagenic single-base substitutions and DNA adducts (Aruoma et al, 1989; McBride et al, 1991; Balasubramanian et al, 1998; Nunoshiba et al, 1999), as well as damaging peptides and lipids (Farr and Kogoma, 1991). Interestingly, the generation of hydroxyl radicals following DNA damage has been shown in higher order systems, including mammalian cells, and is considered as one of the hallmark features of apoptosis (Jacobson, 1996; Simon et al, 2000). Here, we provide direct evidence that hydroxyl radicals are generated in bacteria following both antibiotic- and peptide-based gyrase poisoning. Our results show that hydroxyl radical production, upon gyrase inhibition, is directly related to the presence and status of Fe–S cluster-containing proteins. In turn, our data suggest that cycling of Fe–S clusters between oxidized and reduced states contributes to a breakdown in endogenous iron regulatory dynamics and global iron management. We thus propose that ROS contribute significantly to bacterial cell death following gyrase inhibition and that the associated series of biochemical reactions represents a maladaptive response to aerobic growth which promotes terminal cell fate. Results Phenotypic response to DNA gyrase inhibition We initially sought to characterize the in vivo response of the wild-type E. coli strain BW25113 (Wanner, 1983) to gyrase inhibition and to identify additional contributors to cell death following gyrase poisoning. To accomplish this, we employed the synthetic quinolone antibiotic, norfloxacin, and the natural peptide toxin, CcdB (a component of the F plasmid-encoded CcdB–CcdA toxin–antitoxin system; Miki et al, 1992). Upon treatment with norfloxacin, we observed an immediate 2-log reduction in colony forming units (CFU/ml) (Figure 1A). These values steadily declined for the remainder of our experiments. Induction of CcdB expression similarly resulted in consistent CFU/ml reduction over the first 3 h of our experiments (Figure 1A). Wild-type cells expressing CcdB, however, routinely reached their minimum density at the 3 h time point, and then began a sustained increase in cell density that continued over the remainder of the experiment. Figure 1.Phenotypic response to gyrase inhibition. (A) Log change in colony forming units per ml (CFU/ml) of wild-type, BW25113 E. coli cells (mean±s.d.). Untreated (black triangles, solid line) and uninduced (harboring CcdB plasmid; black squares, dashed line) cell growth, respectively, was compared with the growth of norfloxacin-treated (250 ng/ml; gray triangles, solid line) and CcdB-expressing (gray squares, dashed line) cultures. (B, C) Induction of DNA damage by gyrase inhibitors. To confirm the occurrence of DNA damage following gyrase poisoning, we employed an engineered promoter-GFP reporter gene construct, pL(lexO)-GFP. LexA cleavage, and thus DNA lesion formation, could be examined at single-cell resolution by measuring green fluorescent protein expression using a flow cytometer. Shown are representative fluorescence population distributions of wild-type cultures (B) treated with norfloxacin, or (C) expressing CcdB, before (time zero, gray) and after (3 h (orange) and 6 h (red)) gyrase inhibition. The respective insets show representative control fluorescence measurements of uninhibited wild-type cultures at time zero (gray) and 6 h (black). Download figure Download PowerPoint Although these results show that, in our system, CcdB is not as efficient as norfloxacin in its ability to induce cellular death in BW25113 cells, our data suggested that both gyrase inhibitors rapidly induce DNA lesion formation. To confirm this hypothesis, we employed an engineered, DNA damage-inducible reporter gene construct that relies upon LexA repression for tight regulation of gfp transcription; fluorescence output is thus reflective of RecA-stimulated autocleavage of LexA following DNA damage recognition (Little, 1991). As anticipated, gyrase poisoning of wild-type bacteria resulted in significant GFP expression from this construct (Figures 1B and C). Norfloxacin treatment uniformly induced high levels of fluorescence (Figure 1B), demonstrating the efficiency and highlighting the irreversibility with which the quinolone traps gyrase and stimulates the formation of DNA breaks. We also observed a large shift in fluorescence upon expression of CcdB, indicative of widespread DNA damage (Figure 1C). In line with our CcdB+ growth data (and the relative inefficiency of CcdB-mediated cell killing compared with norfloxacin in our system), the broad DNA damage-related fluorescence distribution exhibited by CcdB+ cells suggests that DNA damage is being corrected by endogenous repair systems. Taken together, these phenotypic data confirm that norfloxacin and CcdB are both potent inhibitors of DNA gyrase that promote the rapid generation of DNA lesions. Gene expression response to DNA gyrase inhibition We next examined the transcriptional response of wild-type E. coli cells treated with norfloxacin or expressing CcdB, with the goal of identifying potential secondary contributors to cell death. Analysis of our time-course microarray data revealed that >800 genes exhibited statistically significant upregulation or downregulation at one or more time points as a function of each respective inhibitor (Supplementary Table 1); significance (z-score) was determined on a gene by gene basis, by comparison of mean expression levels to a large (∼500) compendium of E. coli microarray data collected under a wide array of conditions (see Materials and methods and Supplementary information for details). Taking a systems biology approach, we first applied gene ontology (GO) (Ashburner et al, 2000; Camon et al, 2004) classifications to categorize this gene list by biological function, and then enriched these data using GO::TermFinder (Boyle et al, 2004), which enabled us to determine the significance of these classifications (Supplementary Figures 1 and 2; Supplementary Table 2). We next employed the RegulonDB database (Salgado et al, 2006) of known transcription factor connections, in conjunction with a published map of transcriptional regulation (Faith et al, 2007), to focus on transcription factors whose targets were significantly enriched in our data set (Supplementary Table 3). These measures thus allowed us to group genes based on regulatory and functional pathway classifications and to identify coordinated responses to gyrase inhibition. As expected, LexA, which coordinates the expression of the SOS DNA damage response program, was found to be among the most overrepresented transcriptional regulators following norfloxacin treatment and CcdB expression; this result served to further validate our growth data and DNA damage reporter results. As highlighted in Figure 2, DNA damage-inducible genes comprised one of the largest and most strongly perturbed functional groups in our study. This again is consistent with activation of both the SOS response pathway and the Din (damage-inducible) family of genes (Witkin, 1976; McEntee, 1977; Kenyon and Walker, 1980; Little and Mount, 1982; Walker, 1984; Courcelle and Hanawalt, 2003; Friedberg et al, 2005). The strongest responding among these repair-related genes were error-prone DNA polymerases IV (DinP) and V (composed of UmuC and UmuD) (Elledge and Walker, 1983; Sutton and Walker, 2001). Figure 2.Transcriptome response to gyrase inhibition. Highlighted is a portion of the functionally enriched gene expression response of norfloxacin-treated or CcdB-expressing wild-type cells. Relative weighted z-scores (a measure of standard deviation) were calculated for each gene, based on comparison to mean expression values derived from a large (∼500) database of microarray data; these values were then normalized by subtracting the corresponding uninduced sample z-scores. Using biochemical pathway and transcription factor regulatory classifications, we identified significantly upregulated functional units that responded in a coordinated manner. For each functional unit, genes that exhibited a weighted z-score ⩾2 standard deviations are shown; scale is shown on left. This analysis is described in greater detail in Materials and methods and Supplementary information. Additionally, all gyrase inhibitor microarray results can be found in Supplementary information. Download figure Download PowerPoint Interestingly, we also observed statistically significant enrichment for the IscR (iron–sulfur cluster regulator) transcription factor as a function of each perturbation. IscR has been shown to autoregulate expression of the iscRUSA operon of genes, which are primarily involved in de novo Fe–S cluster assembly. The repair of heavily oxidized Fe–S clusters is believed to require the activity of the Isc protein family, and the derepression of the iscRUSA operon following oxidant stress has been previously explored (Schwartz et al, 2000; Djaman et al, 2004). Along these lines, additional analysis of our microarray results revealed that the oxidative stress response regulator SoxS was also significantly enriched in our data set. Treatment of E. coli with either norfloxacin or CcdB induced the superoxide response operon, soxRS (Figure 2). Transcription of soxS is positively regulated by SoxR and is stimulated upon SoxR activation, the result of superoxide-induced (yet reversible) oxidation of SoxR's Fe–S cluster (Gaudu et al, 1997; Hidalgo et al, 1998). The SoxS transcription factor is known to activate expression of the manganese-superoxide dismutase, sodA, considered as the first responder in the protection of biomolecules from superoxide-induced damage (Hopkin et al, 1992). SodA, like SoxS, was among our list of significantly upregulated genes following norfloxacin treatment or CcdB expression (Figure 2). That damage to Fe–S clusters was occurring as a function of gyrase poisoning was further supported by the observation that each component of the iscRSUA operon exhibited highly increased expression upon inhibition of gyrase (Figure 2). SoxS has also been shown to activate expression of fur, a ferrous (II) iron-binding transcription factor that regulates the expression of iron uptake, utilization and homeostasis genes (Escolar et al, 1999; Andrews et al, 2003). As highlighted in Figure 2, a number of Fur-regulated iron uptake-related genes were among those genes found to be significantly upregulated by norfloxacin or CcdB. Specifically, we observed increased expression of siderophore-based iron acquisition systems, including components of the Fe3+-siderophore receptor/permease Fec, Fep and Fhu families, as well as the energy transducing TonB–ExbBD complex, which provides the energy required for active siderophore transport. Gyrase inhibition induces hydroxyl radical formation In the Fenton reaction, free ferrous iron reacts with hydrogen peroxide to generate highly destructive hydroxyl radicals (Imlay, 2003). Superoxide, via oxidative damage to Fe–S clusters and iron leaching, has been shown to play a critical role in the generation of hydroxyl radicals. Based on our microarray results and previous studies, we hypothesized that production of hydroxyl radicals may be the cytotoxic end product of a surge in superoxide and Fe–S cluster cycling following gyrase inhibition. To detect the generation of hydroxyl radicals upon gyrase poisoning by norfloxacin or CcdB, we employed the fluorescent reporter dye, 3′-(p-hydroxyphenyl) fluorescein (HPF) (Setsukinai et al, 2003), which is oxidized by hydroxyl radicals with high specificity. We detected a significant increase in hydroxyl radical-induced HPF fluorescence at both 3 and 6 h post-treatment of wild-type E. coli with norfloxacin (Figure 3A). Similarly, expression of CcdB in wild-type E. coli also resulted in a large HPF fluorescence increase at both 3 and 6 h post-induction (Figure 3B). Importantly, untreated cultures did not show any detectable increase in HPF fluorescence for the duration of our experiments. These data show that inhibition of DNA gyrase and the generation of double-stranded DNA breaks lead to environmental changes that promote the formation of deleterious hydroxyl radicals. Figure 3.Generation of hydroxyl radicals following gyrase inhibition. Formation of oxidatively damaging hydroxyl radicals was measured using the highly specific fluorescent dye, HPF. Hydroxyl radical generation was observed following both (A) norfloxacin treatment and (B) CcdB expression. Representative flow cytometer-measured fluorescence population distributions of wild-type cells before (time zero, gray) and after (3 h (light blue) and 6 h (blue)) gyrase inhibition are shown. The respective insets show representative control fluorescence population distributions of uninhibited wild-type cultures at time zero (gray) and 6 h (black). Download figure Download PowerPoint To demonstrate that generation of hydroxyl radicals was related to the Fenton reaction and an available pool of reactive iron, we repeated our HPF fluorescence measurements in the presence of the iron chelator, o-phenanthroline. This chelator has been previously used to block the effects of the Fenton reaction in bacteria exposed to hydrogen peroxide (Imlay et al, 1988; Asad and Leitao, 1991) and to uncover the role of free radical damage in mitochondria (Yang et al, 1958; de Mello Filho and Meneghini, 1985). As anticipated, o-phenanthroline inhibited the formation of hydroxyl radicals following gyrase inhibition by norfloxacin (Figure 4A) or CcdB (Figure 4B) for the duration of the respective experiments. Figure 4.Iron chelation prevents hydroxyl radical formation and reduces gyrase inhibitor-mediated cell death. Application of the iron chelator, o-phenanthroline, suppresses the formation of hydroxyl radicals and reduces cell death following gyrase inhibition in wild-type E. coli cells. (A, B) Using the fluorescent reporter dye, HPF, we monitored the effect of o-phenanthroline addition on hydroxyl radical formation in (A) norfloxacin-treated and (B) CcdB-expressing wild-type cells. Representative flow cytometer-measured fluorescence population distributions of wild-type cells before (time zero, gray) and after (6 h (green)) gyrase inhibition, in the presence of the iron chelator are shown. For direct comparison, fluorescent population distributions of gyrase-inhibited cells, at 6 h, in the absence of chelator, are also shown (blue). (C) Log change in colony forming units per ml (CFU/ml) of wild-type cells (mean±s.d.). Cells were grown in the presence of o-phenanthroline and treated with norfloxacin (250 ng/ml, green triangles, solid line) or induced to express CcdB (green squares, dashed line). These results were compared with the growth of cells treated with norfloxacin (gray triangles, solid line) or expressing CcdB (gray squares, dashed line) alone. As controls, normal growth (black triangles, solid line) and growth in the presence of o-phenanthroline alone (purple triangles, solid line) are shown. Download figure Download PowerPoint To determine the effect of hydroxyl radical-induced oxidative damage on the survival of gyrase-inhibited E. coli, we performed growth studies in the presence of Fenton reaction-neutralizing o-phenanthroline. Addition of the chelator to cultures treated with norfloxacin significantly inhibited cell killing, essentially rendering the drug bacteriostatic (Figure 4C). While the attenuation of cell death in CcdB-expressing cells was not as dramatic, we did observe a greater than 0.5-log increase in CFU/ml before recovery around 2 h (Figure 4C). Our findings thus provide strong evidence for the involvement of hydroxyl radicals in gyrase inhibitor-induced cell death. Finally, to show that the observed generation of hydroxyl radicals following norfloxacin application was not related to direct redox cycling of the antibiotic, we examined hydroxyl radical formation and survival in two gyrA mutant E. coli strains; these strains, RFS289 (gyrA111) (Schleif, 1972) and KL317 (gyrA17) (Norkin, 1970), are both known to be resistant to the quinolone, nalidixic acid. Treatment with norfloxacin of both mutant strains did not result in loss of cellular viability, proving that these specific gyrA mutations likewise confer resistance to norfloxacin (Supplementary Figure 3). More importantly, we did not observe an increase in hydroxyl radical formation following norfloxacin treatment of these strains (Supplementary Figure 3). These results clearly show that norfloxacin-mediated hydroxyl radical formation occurs only when cell death is observed and as a consequence of the interaction between gyrase and the gyrase inhibitor. Iron misregulation following gyrase inhibition As our hydroxyl radical-sensitive dye experiments revealed,
