bacteria which have lower growth rates and are therefore less affected by antibiotic drugs (Singh et al., 2009; Conlon et al., 2015). Moreover, no dosing regimen tested in this study, even if it exceeded the recommended PK/PD index values, was able to fully eradicate the planktonic bacteria co-existing with a biofilm, which could suggest that some planktonic bacteria were continuously released from the biofilm. As our study is the first one focusing on the biofilm in the HF, microscopy imaging will be further needed to investigate the distribution of the biofilm in the HF cartridge, which could be influenced, among others, by the shear forces in the extracapillary space. It should also be kept in mind that our system was characterized by an absence of the immune system and the presence of a rich medium – more favorable to bacterial growth -, that both limit the efficacy of antibiotic treatments compared to the in vivo situation. However, our in vitro results are in agreement with the reported lack of efficacy of systemic antibiotic treatments in patients for whom additional treatments, such as mechanical removal of biofilms or very high local antibiotic concentrations, are advised whenever possible (McConoughey et al., 2014; Wu et al., 2015).

In addition to efficacy, we assessed the ability of the combination to reduce the selection of resistant bacteria in planktonic and biofilm populations. The absence of resistance to vancomycin in this study was in accordance with other experiments conducted on S. aureus (LaPlante and Rybak, 2004). Conversely, bacteria (approximately 10−6 ) able to grow on agar supplemented with 6 µg/mL (sixfold MIC) of amikacin were systematically present in the planktonic and biofilm populations before drug exposure, implying that small proportions of such bacteria are spontaneously present in large populations, as previously reported (Ferro et al., 2015). Since similar proportions were also found at the end of the control experiments, it suggests that the growth and survival rates of less-susceptible and fully susceptible bacteria were similar in the absence of drugs. After 5 days of antibiotic exposure, the MIC of amikacin for these bacteria able to grow on agar supplemented with amikacin and termed "less-susceptible," never exceeded the resistance breakpoint (>16 µg/mL). These bacteria showed an intermediate amikacin-susceptibility with regard to the EUCAST breakpoints, implying that the administration of amikacin to patients infected by these bacteria would have an uncertain therapeutic effect (Rodloff et al., 2008), but it should also be stressed that the initial MIC of the tested strain was low (1 µg/mL). This suggests that the same selection phenomenon occurring on a strain with a two or four-fold higher MIC would lead to the selection of "true" resistant bacteria. The selection of less-susceptible bacteria, which represented the main population of planktonic bacteria and BEB after exposure for 5 days to amikacin in monotherapy, could be explained by an inducible mechanism of resistance, known as adaptive resistance, in which thickening of the cell wall results in less penetration of amikacin into the bacterial

# REFERENCES

Adamis, G., Papaioannou, M. G., Giamarellos-Bourboulis, E. J., Gargalianos, P., Kosmidis, J., and Giamarellou, H. (2004). Pharmacokinetic interactions of cell (Yuan et al., 2013). Interestingly, the addition of vancomycin to amikacin considerably reduced the proportions of these less-susceptible bacteria in both planktonic bacteria and BEB compared to amikacin alone, especially when vancomycin was administered in boluses. These results suggest that vancomycin was able to limit the growth of these bacteria less-susceptible to amikacin and prevent their selection. The vancomycin administered by CRI associated with amikacin seemed to limit the selection of less-susceptible bacteria to a lesser extent, but these differences require more thorough investigation.

# CONCLUSION

By studying planktonic bacteria and BEB in parallel and by mimicking the fluctuations in antibiotic concentrations over 5 days, as can occur in vivo after daily administrations, we demonstrated the increased efficacy of a combination of amikacin and vancomycin on planktonic bacteria but not on BEB. However, even though vancomycin did not increase the killing activity of amikacin on BEB, it reduced the selection of bacteria less-susceptible to amikacin, which could help to maintain the efficacy of this drug during treatments. Even if these results need to be further confirmed with clinically relevant strains of MSSA and MRSA, they highlight the importance of selecting combination therapies not only based on efficacy but also on resistance selection endpoints by taking into account the 2 coexisting populations of planktonic bacteria and BEB.

Equations:

Concentration HF =

(Concentration CR ∗ Volume CR)+ (Concentration ECS ∗ Volume ECS) Volume CR + Volume ECS (1)

With HF being the Hollow-Fiber, CR the Central Reservoir and ECS the Extra-Capillary Space.

# AUTHOR CONTRIBUTIONS

DB, AF, FW, FE, P-LT, and AB-M: substantial contributions to the conception or design of the work. DB, ML, AF, P-LT, and AB-M: acquisition, analysis, or interpretation of data for the work. DB, ML, FW, FE, P-LT, AB-M, and AF drafting the work or revising it critically for important intellectual content. Final approval of the version to be published. Agreement to be accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved.

ceftazidime, imipenem and aztreonam with amikacin in healthy volunteers. Int. J. Antimicrob. Agents 23, 144–149. doi: 10.1016/j.ijantimicag.2003.07.001

Aeschlimann, J. R., Allen, G. P., Hershberger, E., and Rybak, M. J. (2000). Activities of LY333328 and vancomycin administered alone or in combination with

gentamicin against three strains of vancomycin-intermediate Staphylococcus aureus in an in vitro pharmacodynamic infection model. Antimicrob. Agents Chemother. 44, 2991–2998. doi: 10.1128/AAC.44.11.2991-2998.2000

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Broussou, Lacroix, Toutain, Woehrlé, El Garch, Bousquet-Melou and Ferran. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# DNA Damage Repair and Drug Efflux as Potential Targets for Reversing Low or Intermediate Ciprofloxacin Resistance in E. coli K-12

Rasmus N. Klitgaard1 , Bimal Jana2 , Luca Guardabassi2 , Karen L. Nielsen3 and Anders Løbner-Olesen1 \*

1 Department of Biology, Section for Functional Genomics, University of Copenhagen, Copenhagen, Denmark, 2 Department of Veterinary and Animal Sciences, Section for Veterinary Clinical Microbiology, University of Copenhagen, Copenhagen, Denmark, 3 Department of Clinical Microbiology, Center for Diagnostics, Rigshospitalet, Copenhagen, Denmark

#### Edited by:

Sanna Sillankorva, University of Minho, Portugal

#### Reviewed by:

César de la Fuente, Massachusetts Institute of Technology, United States Munawar Sultana, University of Dhaka, Bangladesh Azucena Mora Gutiérrez, Universidade de Santiago de Compostela, Spain Catherine M. Logue, University of Georgia, United States

\*Correspondence:

Anders Løbner-Olesen<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Antimicrobials, Resistance and Chemotherapy, a section of the journal Frontiers in Microbiology

Received: 15 March 2018 Accepted: 11 June 2018 Published: 02 July 2018

#### Citation:

Klitgaard RN, Jana B, Guardabassi L, Nielsen KL and Løbner-Olesen A (2018) DNA Damage Repair and Drug Efflux as Potential Targets for Reversing Low or Intermediate Ciprofloxacin Resistance in E. coli K-12. Front. Microbiol. 9:1438. doi: 10.3389/fmicb.2018.01438 Ciprofloxacin is a potent antibacterial drug that is widely used in human clinical applications. As a consequence of its extensive use, resistance has emerged in almost all clinically relevant bacterial species. A mean to combat the observed ciprofloxacin resistance is by reversing it via co-administration of a potentiating compound, also known as a helper drug. Here, we report on the current advances in identifying ciprofloxacin helper drugs, and put them into perspective of our own findings. We searched for potential helper drug targets in Escherichia coli strains with different levels of ciprofloxacin resistance using transcriptomics i.e., RNAseq and by deletion of genes associated with hyper-susceptibility to ciprofloxacin. Differential gene expression analysis of the highly ciprofloxacin resistant uropathogenic E. coli strain, ST131 UR40, treated with a clinically relevant concentration of ciprofloxacin (2 µg/mL), showed that the transcriptome was unaffected. Conversely, genetic screening of 23 single gene deletions in the high-level ciprofloxacin resistant laboratory derived E. coli strain, LM693, led to a significant decrease in the minimal inhibitory concentration for several genes, including genes encoding the AcrAB-TolC efflux pump, SOS-response proteins and the global regulator Fis. In addition, deletion of acrA, tolC, recA, or recC rendered two E. coli strains with intermediate susceptibility to ciprofloxacin fully susceptible according to the CLSI recommended breakpoint. Our results corroborate the AcrAB-TolC efflux pump and the SOS response proteins, RecA and RecC, as potential targets for ciprofloxacin helper drugs in treatment of human bacterial infections caused by E. coli strains with intermediate sensitivity to ciprofloxacin.

Keywords: antibiotic resistance, ciprofloxacin, helper drugs, RNA-Seq, transcriptomics

# INTRODUCTION

Fluoroquinolones are some of the most prescribed antibacterial drugs in the world, commonly used for the treatment of urinary tract infections and sinusitis (Emmerson and Jones, 2003; Linder et al., 2005; Mitscher, 2005), but this has not always been the case. For the first two decades after the discovery of nalidixic acid in 1962, and its introduction into the clinic in 1964, the quinolones

**Abbreviations:** MIC, minimal inhibitory concentration; ST, sequence type.

were only used to treat uncomplicated urinary tract infections. This changed with the release of the second generation quinolones, including ciprofloxacin, which showed significant activity outside the urinary tract and against a broad spectrum of both Gram-negative and Gram-positive bacteria. Ciprofloxacin acts by binding to its targets, DNA gyrase and topoisomerase IV, inhibiting the native ability of these two enzymes to re-ligate double stranded DNA breaks, in turn leading to fragmentation of the chromosome. Due to its mechanism of action it is sometimes referred to as topoisomerase poison (Aldred et al., 2014). Inevitably, considering its extensive use and misuse, resistance toward ciprofloxacin has increased in almost all clinically relevant bacteria (Werner et al., 2011; Dalhoff, 2012). One method to overcome antibacterial resistance is by combinatorial treatment with a potentiating compound, also known as a helper drug. A helper drug is by definition non-antibacterial when administered alone, but it enhances the activity of the antibiotic when used in concert. The potentiating effect of a helper drug can be achieved by either direct inhibition of the resistance mechanism or by targeting endogenous cellular components and pathways like, cell membranes, efflux pumps and cellular repair systems. A classic example of targeting the resistance mechanism is the combination of amoxicillin and the β-lactamase inhibitor clavulanic acid (White et al., 2004). Highlevel ciprofloxacin resistance is primarily associated with multiple target site mutations in gyrA and parC, encoding subunits of the DNA gyrase and topoisomerase IV, respectively (Aldred et al., 2014). Since 1998 three different plasmid-mediated ciprofloxacin resistance mechanisms have been identified; (i) target protection (Qnr proteins), (ii) efflux pumps (QepA and OqxAB) and (iii) drug modification (AAC(60 )-Ib-cr acetyltransferase) (Rodríguez-Martínez et al., 2016).

# Potential Ciprofloxacin Helper Drug Targets

Studies of the endogenous cellular mechanisms involved in ciprofloxacin susceptibility and resistance evolution have revealed more than two dozen gene deletions that lead to increased ciprofloxacin susceptibility in wild-type Escherichia coli (Cirz et al., 2005; Tamae et al., 2008; Liu et al., 2010; Yamada et al., 2010). Thus, suggesting the gene products as potential ciprofloxacin helper drug targets. Recently, Tran et al. identified 23 single gene deletions that increased the ciprofloxacin susceptibility of a laboratory derived resistant E. coli strain. The most significant increase in ciprofloxacin susceptibility was observed for the deletion of SOS-response genes directly involved in DNA damage repair, and the genes encoding the AcrAB-TolC efflux pump (Tran et al., 2016). Recacha et al. recently showed that deletion of recA rendered a laboratory derived E. colistrain with intermediate sensitivity to ciprofloxacin clinically susceptible in vitro. In addition, the in vivo efficacy of ciprofloxacin against the same strain was significantly increased in a peritoneal sepsis murine model (Recacha et al., 2017). Thus, the current evidence suggests that targeting the repair of ciprofloxacin induced DNA damage or the efflux pump AcrAB-TolC are the most promising strategies for ciprofloxacin helper drugs. Here, we used a combined transcriptomic and genetic approach in an attempt to both identify novel helper drug targets, as well as further assess the potential of known helper drug targets in laboratory derived E. coli strains with different levels and mechanisms of ciprofloxacin resistance.

# MATERIALS AND METHODS

# Bacterial Strains and Plasmids

Strains LM693 and LM862 were obtained from Diarmaid Hughes from Uppsala University. LM693 is isogenic to the commonly used laboratory strain, MG1655, besides two gyrA mutations, S83L and D87N, and one parC mutation, S80I. LM862 is also isogenic to MG1655, but with one gyrA S83L mutation and one parC S80I mutation. ST131 UR40 has two gyrA mutations, S83L and D87, and two parC mutations, S80I and E84V, and carries aac-60 -Ib-cr on a plasmid (Cerquetti et al., 2010). The aac-60 -Ib-cr carrying plasmid pRNK1 (was constructed as follows: aac-60 -Ib-cr gene was amplified by PCR from ST131 UR40, using the following primers: GATCGGATCCATGAGCAACGCAAAAACAAAGTT AGGC and CATCGAATTCTTAGGCATCACTGCGTGTTCGC, and cloned into pMW119 (Nippon Gene, Toyama, Japan) using BamHI and EcoRI. The qnrS-carrying plasmid pRNK9 was constructed as follows: qnrS was amplified by PCR from the clinical E. coli isolate EC38 using the following primers: GATCGGATCCATGGAAACCTACAATCATACATAT CGGC and GATCAAGCTTTTAGTCAGGATAAACAACAAT ACCCAGTGC, and cloned into pMG25 using BamHI and HindIII (M. Mikkelsen and K. Gerdes, unpublished). pRNK1 (4796 bp) and pRNK9 (4723 bp) was then introduced in LM862 by electroporation. Strain EC38 was isolated from a patient with a urinary tract infection at Hvidovre Hospital, Denmark.

# Genetic Screening and MIC Tests

For the genetic screen, P1 phage lysates were prepared from the relevant Keio collection strains (Baba et al., 2006) and used for transduction into LM693 and LM862. All the transduced strains were verified by PCR. The ciprofloxacin MICs for LM693 and derived strains were determined using E-tests (0.002–32 µg/ml, BioMerieux) and according to the manufacturer's guidelines. The MICs for LM862 and derived strains were determined by broth micro-dilution using cation adjusted Mueller Hinton broth II with 1 mM and 10 µM IPTG for pRNK1 and pRNK9, respectively. The two different IPTG concentrations were used to obtain a ciprofloxacin MIC for LM862/pRNK1 and LM862/pRNK9 of 2 µg/mL i.e., within the CLSI intermediate susceptible range. The reference E. coli strain ATCC 25922 was used as standard in all MIC tests and the susceptibility was evaluated according to CLSI recommended breakpoints.

# Checkerboard Assay

All wells in a micro-titter plate were filled with 100 µl cation adjusted Mueller Hinton broth II (200 µL in the negative control wells). Copper phtalocyanine-3,40 ,400,4000-tetrasulfonic acid, was added to the first row, followed by serial dilution along the abscissa, leading to a start concentration of 100 µM. Hereafter ciprofloxacin was serial diluted along the ordinate, giving a start concentration of 2 and 64 µg/ml for LM862 and LM693, respectively. Hundred microliter diluted culture with an OD600 of 0.001 was then inoculated in each well and the plates were incubated at 37◦C for 24 h.

# RNA-Sequencing

fmicb-09-01438 June 28, 2018 Time: 17:56 # 3

Ciprofloxacin was added to a culture of ST131 UR40, which had been growing exponentially for more than six generations, to a final concentration of 2 µg/ml. Samples for RNA isolation were taken at 0 min (prior to ciprofloxacin addition) and 30 and 90 min after ciprofloxacin addition, which has previously been shown to be long enough to induce fragmentation of the E. coli chromosome (Charbon et al., 2014). Total RNA was isolated using a Thermo Scientific GeneJET RNA isolation kit. Dnase treated with TURBO DNA-free kit from Ambion. rRNA was depleted using an Illumina Ribo-zero rRNA removal kit, followed by RNA-Seq library prep using an Illumina TruSeq Stranded mRNA Library Prep Kit. Sequencing was performed on an Illumina Miseq with a Miseq reagent kit v3. (75 bp pairedend) from Illumina. Data analysis was performed in Rockhopper ver.2.03 (McClure et al., 2013). E. coli NA114 (ST131) (accession number: NC\_017644) was used as reference genome (Avasthi et al., 2011). The percentage of successfully aligned reads varied from 91 to 88% of the total read count. The sequencing data files and the Rockhopper results from the differential gene expression analysis are available from the Gene Expression Omnibus (GEO: GSE89507).

# RESULTS

# Identification of Helper Drug Targets by Genetic Screening

As mentioned in the introduction several single gene deletions are known to increase the ciprofloxacin susceptibility of E. coli. To further assess the helper drug target potential of these genes, 23 single gene deletions were introduced into the highlevel ciprofloxacin resistant E. coli strain LM693 (MIC of 24– 32 µg/ml) (Marcusson et al., 2009) and tested for hypersusceptibility toward ciprofloxacin (**Table 1**). LM693 is isogenic to the commonly used laboratory strain MG1655 besides two GyrA mutations; S83L and D87, and one ParC mutation; S80I. Even though nine of the mutant strains showed a three to four fold reduction in the MIC, none of them were found to be susceptible according to the CLSI MIC breakpoint for ciprofloxacin (≤1 µg/mL). Our results therefore indicate that none of the tested gene-knockouts identify valid helper drug targets in high-level ciprofloxacin resistant E. coli strains. However, they could potentially be used as helper drug targets in bacteria with intermediate susceptibility to ciprofloxacin. To create two strains with intermediate ciprofloxacin susceptibility, we constructed the plasmids pRNK1 and pRNK9 carrying the ciprofloxacin resistance determinants aac-60 -Ib-cr and qnrS, respectively. AAC-60 -Ib-cr inactivates ciprofloxacin by N-acetylation of the amino nitrogen of its piperazinyl substituent (Robicsek et al., 2006), while QnrS acts as a DNA mimic, binding to and protecting the gyrase from the action of ciprofloxacin (Rodríguez-Martínez et al., 2016). Introduction of pRNK1 and pRNK9 into strain LM862, which carries GyrA S83L and ParC S80I mutations, increased the MIC from 1 to 2 µg/ml, i.e., into the CLSI intermediate susceptible MIC range. We then evaluated the ability of seven of the most promising gene deletions described above to reduce the ciprofloxacin MIC of LM862/pRNK1 and LM862/pRNK9. Four of the gene deletions (acrA, tolC, recA, and recC) rendered both strains susceptible to ciprofloxacin (**Table 2**). To assess whether inhibition of RecA was an amenable strategy for potentiation of ciprofloxacin, synergy between ciprofloxacin and a RecA inhibitor, copper phtalocyanine-3,40 ,400,4000-tetrasulfonic acid (Alam et al., 2016), was tested by a checkerboard assay. However, we did not observe a reduction in the ciprofloxacin MICs for either LM693 or LM862.

# Identification of Helper Drug Targets by RNA Sequencing

The E. coli clonal group ST131 has become the predominant E. coli lineage isolated from human extra-intestinal infections and is currently regarded a global problem in hospitals and clinical practices (Nicolas-Chanoine et al., 2014). Two independent studies have shown that more than 90% of ESBL-producing

TABLE 1 | MIC values for the single gene deletions in LM693.

TABLE 2 | MIC values for the single gene deletions in LM862/pRNK1 and LM862/pRNK9.

ST131 isolates are also resistant to ciprofloxacin (Brisse et al., 2012; López-Cerero et al., 2014). Strain ST131 UR40 is resistant to high levels of ciprofloxacin due to GyrA mutations S83L and D87, and ParC mutations S80I and E84V (Cerquetti et al., 2010). Here we used RNA-Seq to map the transcriptomic changes during treatment of ST131 UR40 with a clinically relevant concentration of ciprofloxacin, 2 µg/ml, which is approximately equal to the maximum serum concentration following oral administration of 500 mg ciprofloxacin according to the FDA. The rationale behind this was to identify potential helper drug target genes that were upregulated upon ciprofloxacin exposure and thereby potentially involved in ciprofloxacin resistance. In contrast to the genetic screen, the RNA-Seq analysis would also reveal targets encoded by essential genes and non-coding RNA. The transcriptomic analysis did not show any non-ribosomal transcripts to be significantly upregulated in the presence of ciprofloxacin, i.e., with a false discovery rate of <1% and more than two-fold expression change.

# DISCUSSION

By utilizing a combination of "direct genetic screening" and differential gene expression analysis, we have attempted to identify potential genes suitable as targets for ciprofloxacin potentiating compounds. We did not find any genes to be significantly upregulated by ciprofloxacin, indicating that the transcriptome of ST131 UR40 was relatively unaffected by treatment with a sub-inhibitory and yet clinically relevant concentration of ciprofloxacin. The lack of an upregulation of the SOS response genes in the transcriptomic analysis suggests that the ciprofloxacin exposure did not cause sufficient DNA damage to induce a SOS response; hence it was not necessary for ST131 UR40 to upregulate any specific genes to cope with the presence of ciprofloxacin at a sub-inhibitory concentration.

The screening of selected mutant strains revealed a number of genes, which when deleted, lowered the MIC for ciprofloxacin significantly in LM693. These findings are in accordance with genes reported to contribute to high-level ciprofloxacin resistance by Tran et al. (2016). Treatment of bacteria with ciprofloxacin generates double stranded breaks in the DNA of the organism (Drlica et al., 2008), which in turn activates the SOS response. Seven of the tested gene deletions; recA, recC, recG, uvrD, xseAB, and ruvC, which all significantly reduced the MIC of LM693, are part of the SOS response and involved in DNA damage repair (Chase and Richardson, 1974; Kuzminov, 1993; Michel, 2005). Thus, deletion of any of these seven genes likely lowers the ability of the bacteria to cope with ciprofloxacin induced DNA damage. Deletion of genes encoding the AcrAB-TolC efflux pump, or the global regulator Fis (Factor for inversion stimulation) showed the largest decreases in MIC values for LM693. The Fis protein has been shown to repress the gyrA and gyrB promoters, thereby reducing the expression of the DNA gyrase (Schneider et al., 1999). Thus, deletion of fis likely increases DNA gyrase expression and the number of ciprofloxacin targets. As ciprofloxacin works as a topoisomerase poison, an increase in ciprofloxacin bound DNA gyrase could potentially lead to an increase in double stranded breaks, explaining the decrease in MIC for the fis deletion strain. The fis deletion did not have the same effect on the intermediate susceptible strains LM862/pRNK1 and LM862/pRNK9, which may be explained by the relatively higher affinity of ciprofloxacin for its target in LM862, compared to that of LM693. Thus, the increase in expression of the DNA gyrase might lead to an increase in ciprofloxacin-gyrase complexes, but if the ciprofloxacin induced DNA damage is already at a level, where the DNA repair mechanisms cannot keep up, the fis deletion does not have a dramatic effect on the MIC.

Individual deletions of acrA, acrB, or tolC genes encoding the AcrAB-TolC efflux pump had a large effect on the ciprofloxacin susceptibility of both LM693 and LM862 strains. This was not surprising as overexpression of the AcrAB-TolC efflux system has been connected to ciprofloxacin resistance numerous times (Mazzariol et al., 2000). The deletion of acrA or tolC in the LM862 strains lowered the MIC beneath the CLSI susceptible breakpoint indicating that AcrAB-TolC efflux system is a potential target for ciprofloxacin potentiating compounds in intermediate susceptible E. coli. A number of AcrAB-TolC inhibitors have been identified (Chevalier et al., 2004; Bohnert et al., 2013; Aparna et al., 2014; Opperman et al., 2014; Yilmaz et al., 2015), two of which have been shown to decrease the MIC of ciprofloxacin in susceptible E. coli strains (Opperman et al., 2014; Yilmaz et al., 2015), but none of them are used in clinical practice so far.

Inhibition of RecA and thereby of the SOS response has been proposed as a strategy to fight antibiotic resistance numerous times (Blázquez et al., 2012; Culyba et al., 2015; Alam et al., 2016). Our finding, that deletion of recA render intermediate susceptible strains of E. coli fully susceptible to ciprofloxacin is in accordance with recent observations by Recacha et al. (2017). Overall, this indicates that RecA could be a potential ciprofloxacin helper drug target.

Even though AcrAB-TolC or RecA deficiency rendered LM862/pRNK1 and LM862/pRNK9 susceptible to ciprofloxacin, the respective MICs were only two to four-folds lower than the susceptible MIC breakpoint. It therefore seems reasonable to assume that a given inhibitor should completely block the activity of either RecA or AcrAB-TolC in order for it to be an efficient helper drug. This hypothesis is backed by the failure of lowering the ciprofloxacin MIC of LM862 and LM693 with the RecA inhibitor phtalocyanine-3,40 ,400,4000-tetrasulfonic acid.

# CONCLUSION

The findings of this study and the evidence given in the literature, indicates that reversal of ciprofloxacin resistance in high-level resistant E. coli strains by the use of helper drugs does not appear to be plausible. Conversely, targeting RecA, RecC or the AcrAB-TolC efflux pump is a likely feasible strategy for reversing ciprofloxacin resistance in E. coli strains with intermediate susceptibility to ciprofloxacin. However, it should be noted that there is a discrepancy between the MIC breakpoint

for ciprofloxacin susceptibility proposed by the CLSI (≤1 µg/mL) and the EUCAST (≤0.25 µg/mL). Therefore, further in vivo studies are needed to asses if targeting either RecA, RecC, or AcrAB-TolC leads to a significant increase in the efficacy of ciprofloxacin against an intermediate susceptible E. coli strain.

# AUTHOR CONTRIBUTIONS

RK carried out all experimental work, designed the study, analyzed the data, and prepared the final manuscript. AL-O supervised all aspects of the study and helped prepare the final manuscript. BJ assisted and supervised the experimental part

## REFERENCES

of the RNA-seq. LG supervised and delivered the ST131 UR40 strain. KN performed genomic analyses and delivered the EC38 strain carrying the qnrS gene. All authors read and approved the final manuscript.

# FUNDING

We acknowledge the financial support from the University of Copenhagen Centre for Control of Antibiotic Resistance (UC-Care) and by the Center for Bacterial Stress Response and Persistence (BASP) funded by a grant from the Danish National Research Foundation (DNRF120).

a common aminoglycoside acetyltransferase. Nat. Med. 12, 83–88. doi: 10.1038/ nm1347

of an innovative antimicrobial agent. J. Antimicrob. Chemother. 53(Suppl. 1), i3–i20. doi: 10.1093/jac/dkh050

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Klitgaard, Jana, Guardabassi, Nielsen and Løbner-Olesen. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# In Vitro Antibacterial Activity of Teixobactin Derivatives on Clinically Relevant Bacterial Isolates

Estelle J. Ramchuran1 , Anou M. Somboro1 , Shimaa A. H. Abdel Monaim2 , Daniel G. Amoako1 , Raveen Parboosing3 , Hezekiel M. Kumalo4 , Nikhil Agrawal5 , Fernando Albericio2,6, Beatriz G. de La Torre5 and Linda A. Bester1 \*

1 Biomedical Resource Unit, School of Laboratory Medicine and Medical Sciences, College of Health Sciences, University of KwaZulu-Natal, Durban, South Africa, 2 Peptide Research Group, School of Chemistry and Physics, University of KwaZulu-Natal, Durban, South Africa, 3 Department of Virology, National Health Laboratory Service, University of KwaZulu-Natal, Durban, South Africa, 4 Discipline of Medical Biochemistry, School of Laboratory Medicine and Medical Science, University of KwaZulu-Natal, Durban, South Africa, 5 KRISP, College of Health Sciences, University of KwaZulu-Natal, Durban, South Africa, 6 CIBER-BBN, Networking Centre on Bioengineering, Biomaterials and Nanomedicine, and Department of Organic Chemistry, University of Barcelona, Barcelona, Spain

### Edited by:

Sanna Sillankorva, University of Minho, Portugal

#### Reviewed by:

Nagendran Tharmalingam, Alpert Medical School, United States Santi M. Mandal, Indian Institute of Technology Kharagpur, India

> \*Correspondence: Linda A. Bester<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Antimicrobials, Resistance and Chemotherapy, a section of the journal Frontiers in Microbiology

Received: 13 December 2017 Accepted: 20 June 2018 Published: 11 July 2018

#### Citation:

Ramchuran EJ, Somboro AM, Abdel Monaim SAH, Amoako DG, Parboosing R, Kumalo HM, Agrawal N, Albericio F, de La Torre BG and Bester LA (2018) In Vitro Antibacterial Activity of Teixobactin Derivatives on Clinically Relevant Bacterial Isolates. Front. Microbiol. 9:1535. doi: 10.3389/fmicb.2018.01535 Methicillin-resistant Staphylococcus aureus (MRSA) and vancomycin-resistant enterococcus (VRE) are included on the WHO high priority list of pathogens that require urgent intervention. Hence emphasis needs to be placed on developing novel class of molecules to tackle these pathogens. Teixobactin is a new class of antibiotic that has demonstrated antimicrobial activity against common bacteria. Here we examined the antimicrobial properties of three Teixobactin derivatives against clinically relevant bacterial isolates taken from South African patients. The minimum inhibitory concentration (MIC), the minimal bactericidal concentration (MBC), the effect of serum on MICs and the time-kill kinetics studies of our synthesized Teixobactin derivatives (3, 4, and 5) were ascertained following the CLSI 2017 guidelines and using the broth microdilution method. Haemolysis on red blood cells (RBCs) and cytotoxicity on peripheral blood mononuclear cells (PBMCs) were performed to determine the safety of these compounds. The MICs of 3, 4, and 5 against reference strains were 4–64 µg/ml, 2–64 µg/ml, and 0.5–64 µg/ml, respectively. The MICs observed for MRSA were (3) 32 µg/ml, (4) 2–4 µg/ml and (5) 2–4 µg/ml whilst those for VRE were (3) 8–16 µg/ml, (4) 4 µg/ml and (5) 2–16 µg/ml, respectively. In the presence of 50% human serum, there was no significant effect on the MICs. The compounds did not exhibit any effect on cell viability at their effective concentrations. Teixobactin derivatives (3, 4, and 5) inhibited bacterial growth in drug-resistant bacteria and hence emerge as potential antimicrobial agents. Molecular dynamic simulations suggested that the most dominant binding mode of Lys10-teixobactin (4) to lipid II is through the amide protons of the cycle, which is identical to data described in the literature for the natural teixobactin hence predicting the possibility of a similar mechanism of action.

Keywords: teixobactin derivatives, biological activity, antimicrobial agents, resistant bacteria, antimicrobial peptides, in silico analysis

# INTRODUCTION

fmicb-09-01535 July 11, 2018 Time: 15:9 # 2

The rate of antibiotic resistance is increasing faster than the development of new compounds for clinical practice. In an extremely short period, resistance to antibiotics has become a significant cause of disease and death globally (Penesyan et al., 2015; Brown and Wright, 2016; Hamilton and Wenlock, 2016). Limited success in collective research efforts to synthesize novel and efficient compounds has contributed to the drug-resistance scenario we are now facing and to the lack of new and efficient treatment options.

The first antibiotics were produced through screening soil microorganisms. However, by the 1960s, this limited resource of cultivable bacteria had been overexploited (Lewis, 2012). Synthetic approaches to produce antibiotics have been unable to replace this platform. Uncultured bacteria, which make up 99% of all species in external environments, emerge as a potent source of new antibiotics (Kaeberlein et al., 2002; Nichols et al., 2010; Fang et al., 2012).

Teixobactin (1, **Figure 1**) is a new class of antibiotic that was discovered through the screening of uncultured bacteria using i-Chip (isolation chip), a revolutionary method for bacterial culture (Ling et al., 2015; Piddock, 2015; von Nussbaum and Süssmuth, 2015). Teixobactin was identified as an effective agent against Gram-positive bacteria. It inhibits cell wall synthesis by binding to two lipid cell wall precursors, namely lipid II (peptidoglycan precursor) and lipid III (teichoic acid precursor) (Ling et al., 2015; Homma et al., 2016). Vancomycin also targets lipid II. However, taking into account that Teixobactin has been demonstrated to be active against vancomycinresistant enterococcus (VRE), its binding is through a different region compared to that of the Vancomycin. In this regard, biochemical assays have demonstrated that teixobactin binds the pyrophosphate and the first sugar moiety present in both, lipid II and lipid III (Ling et al., 2015).

Although much attention has shifted towards combating Gram-negative bacteria, there is still a need for compounds with novel mechanisms and low resistance profiles against Grampositive strains. In this regard, Texiobactin can satisfy this need and contribute to the treatment of resistant Gram-positive bacteria such as VRE and MRSA. In this study we sought to evaluate three novel derivatives of Teixobactin (3, 4, and 5) and determine their activity against clinically relevant Gram-positive resistant bacteria as well as against Gram-negative species.

# MATERIALS AND METHODS

## Antibiotics and Reagents

All the derivatives were dissolved in 5% DMSO. GIBCO RPMI-1640 cell culture media (with HEPES, L-glutamine and sodium pyruvate) was obtained from Life Technolgies (Carlsbad, CA, United States). Hyclone fetal bovine serum was purchased from GE Healthcare Life Sciences (Chicago, IL, United States). Phosphate Buffered Saline (PBS) was obtained from Lonza (Basel, Switzerland). Nunclon Delta Surface sterile microtiter plates (including the Edge 2.0 plate) were bought from Thermo Fisher Scientific (Waltham, MA, United States). Human serum from male AB plasma (sterile and filtered), antibiotics, antimycotic solution and all other reagents were obtained from Sigma (St. Louis, MO, United States).

# Bacterial Strains

Clinical isolates of MRSA and VRE were obtained from Lancet Laboratories, Durban, South Africa, with ethical approval BE394/15 from the Biomedical Research Ethical Committee of the University of KwaZulu-Natal. Four reference strains of bacteria, namely Escherichia coli ATCC 25922, Pseudomonas aeruginosa ATCC 27853, Bacillus subtilis ATCC 6051 and Staphylococcus aureus ATCC 29213 were obtained from the American Type Culture Collection (ATCC).

# Synthesis, Purification, and Characterization of Teixobactin Derivatives

Our group previously synthesized the Teixobactin derivatives (3, 4, and 5) (**Figure 1**) used in this study. Furthermore, they were chemically characterized by HPLC and MS and subjected to preliminary biological testing against two Gram-positive and two Gram-negative ATCC strains (Abdel Monaim et al., 2017).

# Minimum Inhibition Concentration (MIC) and Minimum Bactericidal Concentration (MBC) Determination

The MICs of the Teixobactin derivatives were determined using the broth microdilution method following the Clinical and Laboratory Standards Institute [CLSI], 2017 guidelines. Twofold dilutions of each compound solution were prepared using cation adjusted Mueller–Hinton Broth (CAMHB) in a microtiter plate. A 0.5 McFarland-standardized bacterial inoculum was used to prepare a total volume of 200 µl in each microtiter well. The plates were incubated at 37◦C for between 18 and 20 h. The MIC was determined as the lowest concentration at which no visible bacterial growth was observed. Control wells for bacteria and media were also included. Meropenem, vancomycin and ampicillin were used as standard control drugs. The plates containing VRE were incubated at 35◦C under aerobic conditions. The MBC was determined as the lowest concentration of the test compound that was able to produce a 99.9% decrease in viable bacterial cells on the agar plates. Control wells included the same amount of solvent used in dissolving the drug candidates, medium and bacteria.

# Effect of Human Serum on the MICs

The effect of serum on the MICs was determined in a similar way to the MIC method described above, but in this case 50% human serum: Mueller-Hinton broth was prepared.

# Time-Kill Kinetic Assays

Time-kill assays were performed following CLSI guidelines and previously described methods (Wang et al., 2015; Clinical Laboratory Standard Institute [CLSI], 2017; Zheng et al., 2017).

Overnight bacterial cell cultures were suspended in CAMHB and adjusted to an absorbance of approximately 106 CFU/ml. Varying concentrations of the test compounds were added to the inoculum suspensions, with final concentrations corresponding to 1x MIC, 2x MIC, and/or 4x MIC, and incubated at 37◦C. Aliquots were removed from the inoculum cultures after 0, 1, 2, 4, 6, 8, and 24 h of incubation. They were then serially diluted, plated on MH agar and incubated for 24 h at 37◦C. Bacterial cell viability was determined by colony count. The assays were performed in triplicate. Data was presented as mean and standard deviation of three independent replicates, analyzed with one-way ANOVA followed by Dunnett's test to determine the significance relative to the untreated bacteria (P < 0.05).

# Cell Culture

The Buffy coat used in this study was obtained from the South African National Blood Service (SANBS). The anonymised product is provided by the SANBS for research purposes, upon approval from their Ethics Committee (National Health Laboratory Service Clearance Certificate approval no: 2013/18). Aseptic techniques and appropriate biosafety precautions were observed.

# Haemolysis Assay on Red Blood Cells (RBCs)

The haemolysis assay was performed as previously described (Tramer et al., 2012; Jayamani et al., 2017), with modifications to allow for a 96-well microtiter plate format. Briefly, washed red blood cell pellet was re-suspended in PBS (to obtain a hematocrit of approximately 20%). Next, 10 µL of the cells was aliquoted into a 96-well microtiter plate containing 170 µL of PBS and lysed by addition of 20 µL of 1% TritonTM X-100 solution. After 30 min, the samples were spun at 3000 g for 5 min in an Orto Alresa Digicen 21R plate centrifuge. Absorbance was read at 405 nm in a Tecan SunriseTM plate reader.

Seven serial 5-fold dilutions of the compounds were then prepared in triplicate by adding 25 µL of the compound to 100 µL of PBS. Controls (i.e., 0% and 100% hemolysis samples) were included. Appropriately diluted RBCs (10 µL RBCs and 90 µL PBS per well) were added to the microtiter plate and incubated at 37◦C for 30 min and then spun at 3000 g for 5 min. The supernatants were then transferred to a fresh microtiter plate, and absorbance was read at 405 nm. The viability of the RBCs at each concentration of the compound was calculated as follows: % viability = 100 × [1 – [At/(A100 − A0)]] where At = mean absorbance of the test compound at a given concentration, A0 = mean absorbance of the untreated control, and A100 = mean absorbance of the sample lysed with TritonTM X-100. The results were represented graphically. The experiment was performed in triplicate (n = 3). The error bars indicate the standard deviation. One-way ANOVA followed by Dunnett's test was performed to determine the significance relative to the untreated RBC (P < 0.05; indicated by ∗ ).

TABLE 2 | Minimum inhibitory concentration of Teixobactin derivatives against methicillin-resistant Staphylococcus aureus (MRSA).

aETT, Endotracheal tube; CVP, Central venous catheter, –, Missing data.

TABLE 3 | Minimum inhibitory concentration of Teixobactin derivatives against vancomycin-resistant enterococci (VRE).

# Cytotoxicity on Peripheral Blood Mononuclear Cells (PBMCs)

The cytotoxicity assay was performed as previously described with slight modifications (Pannecouque et al., 2008; Pinto et al., 2011; Araújo et al., 2013; Azumah et al., 2016). Briefly, PBMCs (100,000 viable cells/well) were placed into the wells of a NunclonTM Delta Surface Edge 2.0 microtiter plate containing 100 µl of RPMI-1640 with 10% fetal bovine serum, 1% Antibiotic Antimycotic solution and 3% phytohemagglutinin. The cells were then incubated for 24 h at 37◦C and 5% CO2. Seven serial 5-fold dilutions of test compounds were prepared and transferred to the appropriate wells of the plate containing the cells. The plate was then incubated for 72 h at 37◦C and 5% CO2. Thereafter, 20 µl of MTT salt (7.5 mg/ml) was added to each well, and the plate was incubated for a further 4 h. Then 100 µl of the media was carefully removed from each well (avoiding agitation of the crystals) and replaced with 100 µl of solubilisation solution (containing acidified isopropanol and TritonTM X-100). The plate was then placed on a shaker for 30 min to facilitate dissolution of the crystals. Absorbance was read at 550 nm (background: 690 nm). The results were shown graphically. The experiment was performed in triplicate (n = 3). The error bars indicate the standard deviation. One-way ANOVA followed by Dunnett's test was performed to determine the significance relative to the untreated PMBC (P < 0.05; indicated by ∗ ).

# Molecular Dynamics Simulation

Structures of lipid II and teixobactin were downloaded from Automated Topology Builder (ATB) and Repository (Malde et al., 2011). In the teixobactin structure, the residue of enduracididine was replaced by Lys by molefacture program of VMD to get Lys10-teixobactin (4) structure (Humphrey et al., 1996). CHARMM General Force Field (CGenFF) parameters were used for simulation of both molecules (Vanommeslaeghe et al., 2010). The molecular dynamics simulation system contains one lipid II molecule, one Lys10-teixobactin (4) molecule, and 6767 water molecules. The TIP3P water model was used for the water molecules (Mark and Nilsson, 2001). The system was first energy minimized using the steepest descent algorithm (Bixon and Lifson, 1967), after which two sequential equilibrations were performed using canonical ensemble (NVT), followed by an isobaric-isothermic ensemble (NPT) for 100 picoseconds

(ps) each, and production simulation was performed using NPT ensemble for 100 nanoseconds (ns). The simulation was performed at 310 K temperature and 1 atm pressure, for temperature coupling velocity-rescale method and for pressure coupling the Parrinello-Rahman method was used (Parrinello and Rahman, 1981). The Particle Mesh Ewald method was used for long-range electrostatic interactions (Darden et al., 1993), with 10Å cut-off being used to calculate the VdW and shortrange coulombic interactions. MD simulation was performed using the GROMACS simulation package (Abraham et al., 2015). To identify the binding region of lipid II with Lys10-teixobactin (4) an in-house TCL script was used. Script counts the numbers of frames that each oxygens atoms of lipid II were within 3.5 Å of protons of Lys10-teixobactin (4) throughout the simulation time.

# RESULTS

# Teixobactin Derivatives

Teixobactin is an 11-amino acid "head to side-chain" cyclodepsipeptide (**Figure 1**) with a D-Thr as a bridge head that forms the ester with the carboxylic group of a L-Ile.  The tail is formed by two moieties of L-Ser, two moieties of L-Ile, D-allo-Ile and D-Gln ending with a N-Me-D-Phe. As Lallo-End was not commercially available, our group concentrated their efforts on synthesizing Arg10-Teixobactin (2, **Figure 1**), in which the L-allo-End is substituted by Arg (Jad et al., 2015; Parmar et al., 2016).

Arg10-Teixobactin (2), which has been converted from the parent Teixobactin analog, has slightly lower activity than Teixobactin. Our group had previously used a Lys-scanning strategy to prepare a small library of Teixobactin analogs containing more than one Lys residue—a residue that is absent in the natural structure (Abdel Monaim et al., 2016b, 2017). From this collection of peptides, three (3, 4, and 5, **Figure 1**) with good MICs against sensitive bacteria (ATCC strains) were selected for further in vitro evaluation in this study.

# Antimicrobial Activity of Teixobactin Derivatives and the Effects of Human Serum on the MICs

The antimicrobial activity of the three derivatives (3, 4, and 5) was examined by in vitro screening against drug-resistant and -sensitive bacteria using the broth micro-dilution method, following CSLI guidelines. The derivatives inhibited sensitive Gram-positive (ATCC strains) and resistant MRSA and VRE isolates (**Tables 1**–**3**). They demonstrated potent antimicrobial activity against Gram-positive bacteria as opposed to Gramnegative bacteria. Three conventional antibiotics (meropenem, vancomycin and ampicillin) were used as controls and exhibited activity against the drug-sensitive ATCC strains. The following

MIC50 were recorded for the derivatives: (3) 32 µg/ml, (4/5) 2 µg/ml for MRSA, as well as (3) 16 µg/ml and (4/5) 4 µg/ml for VRE. The derivatives yielded MICs as low as 2 µg/ml and 0.5 µg/ml for Gram-positive reference ATCC strains S. aureus and B. subtilis respectively. The MICs of the experimental compounds against susceptible Gram-negative bacteria were 32 µg/ml for E. coli and 64 µg/ml for P. aeruginosa. These compounds also inhibited drug-resistant clinical isolates of MRSA at concentrations of 32 µg/ml, 4 µg/ml and 2 µg/ml for 3, 5, and 4 respectively. Vancomycin, the current drug of choice for the treatment of MRSA, had an MIC of 1 µg/ml; while the MIC of ampicillin against MRSA was ≥512 µg/ml. Compounds 3, 4, and 5 showed MICs against VRE of 16 µg/ml, 8 µg/ml, and 4 µg/ml and respectively.

No significant effect of serum on the MICs was observed when the reference bacterial strains were tested with varying concentrations of the derivatives in the presence of 50% human serum; the values varied by only ±1 in fold dilutions. Compounds

Dunnett's test was performed to determine the significance relative to the RBC and PMBC (P < 0.05).

3, 4, and 5 demonstrated bactericidal activities, yielding a 99.9% decrease in viable cells on the agar plates at concentrations ≤ 4x MIC values. All the experiments were conducted in triplicate to confirm the outcomes.

# Time-Kill Kinetics

Time-kill kinetic assays were performed to determine whether the Teixobactin derivatives showed time-dependent or

TABLE 4 | Number of frames (>1000) that lipid I oxygens was within 3.5 Å of Lys10-teixobactin (4) protons.

aThe number of the O atoms are shown in the Figure 4B.

concentration-dependent properties, as well as whether their effects were bacteriostatic or bactericidal. The time-kill curves of compound 4 against Gram-positive S. aureus ATCC 29213 and B. subtilis ATCC 6051 are shown in **Figure 2**. The kinetics indicated time- and concentration- dependent bacterial killing for this compound, and the bactericidal effect was observed at a concentration of 2x and 4x MIC levels at 6 h, as well as at 1x MIC at 24 h against S. aureus and B. subtilis. Exposure of S. aureus and B. subtilis to 4 at 2x and 4x MIC resulted in a decrease in bacterial cell count greater than 3 log10 relative to the initial density from 6 and 4 h respectively, which was also indicative of a bactericidal effect. At a concentration of 1x MIC, compound 4 caused a significant reduction in log10 CFU 6 h after its addition (**Figure 2**).

# Haemolysis and Cytotoxicity

Haemolysis and cytotoxicity effects were evaluated by exposing RBCs and PBMCs to varying concentrations of the Teixobactin derivatives. The concentrations tested showed no cytotoxic effect on PBMCs or any hemolytic effect on erythrocytes. RBC and PBMC viability was above 90% at the highest concentration of the derivatives used in this study (64 µg/mL) (**Figure 3**).

# Molecular Dynamics Simulation: Binding of Lys10-Teixobactin (4) With Lipid II

To understand the binding modes of Lys10-teixobactin (4) with lipid II, 100 nanoseconds (ns) molecular dynamics simulations were performed. The data revealed different binding modes of Lipid II with Lys10-teixobactin (4) out of which pyrophosphate interaction with amide group proton of the cycle was the most dominant. To identify the interacting region of lipid II with Lys10-teixobactin (4), the number of frames that oxygens of lipid II is within 3.5 Å (hydrogen bonding distance) of proton atoms of Lys10-teixobactin (4) for whole simulations was calculated (**Table 4** indicates > 1000). As there are 10000 frames (10ps each), it was observed that four oxygens of pyrophosphate groups were formed the most interaction with Lys10-teixobactin (4) cycle amide group protons (**Figure 4**).

# DISCUSSION

Although antibiotic resistance in Gram-positive bacteria is increasing worldwide, as indicated by the WHO list of highpriority pathogens (i.e., VRE and MRSA), much attention has shifted to combating Gram-negative bacteria. Teixobactin has been demonstrated as effective against Gram-positive bacteria and no detectable resistance has been reported yet. The capacity of Teixobactin is attributed to it being structurally distinct from glycopeptides and it being the first member of a new class of lipid II binding antibiotics.

The MICs of compounds 4 and 5 for reference strains S. aureus and B. subtilis were between 0.5 and 4 µg/ml (**Table 1**) while for the MRSA isolates they were between 2 and 4 µg/ml. Compound 3 had an MIC of 32 µg/ml, a value that was much higher than that reported for the control antibiotic vancomycin (0.5–1 µg/ml). Other groups reported compound 3 to have a MIC of 4 µg/ml against MRSA (Jin et al., 2016; Parmar et al., 2017a; Schumacher et al., 2017). These results were echoed in the present study, as the MIC observed against MRSA for this compound was between 2 and 4 µg/ml.

The MBC reported by Ling et al. (2015) was 2x the MIC of Teixobactin. The bactericidal activity of Teixobactin and its derivatives against Gram-positive bacteria is superior to that of vancomycin, and these compounds retain excellent bactericidal activity against VRE (Ling et al., 2015). The strong bactericidal activity of Teixobactin and its derivatives is attributed to not

only inhibition of peptidoglycan synthesis but also the synergistic inhibition of cell wall teichoic acid synthesis. These derivatives show the same bactericidal activity as that observed for Teixobactin and the MIC/MBC ratios were ≤4 for all the three derivatives.

Time-kill kinetic assays were carried out with compound 4 as it showed the best MICs against S. aureus and B. subtilis. Complete bactericidal activity was observed at concentrations of 16 and 8 µg/ml at 4 h. Similar to observations of other Teixobactin derivatives, compounds 3, 4, and 5 had no cytotoxic or hemolytic effect in vitro. In the presence of 50% serum, there was no drastic change in the MICs (**Table 1**).

On the basis of our observations, we can conclude that human serum has no effect on the antibacterial activity of compounds 3, 4, and 5. These results are similar to those observed by Parmar et al. (2017a). The serum effect is essential as it aids in speculating the probable in vivo activity of the drug. These derivatives will possibly have low protein binding properties because they bind to multiple target sites, none of which are proteins. The present study confirms that Teixobactin derivatives 3, 4, and 5 are safe and can thus be considered potential treatment options against resistant bacterial infections (VRE and MRSA).

Interestingly, we observed that, at higher concentrations, 3, 4, and 5 were also active against Gram-negative bacteria (**Table 1**). This is a relevant observation given the low toxicity of these compounds. These derivatives may exert their activity against Gram-negative bacteria by disrupting the outer membrane layer. In conclusion, we have demonstrated the highly potent antimicrobial activity of three Teixobactin derivatives against clinically significant isolates of bacteria. Unlike vancomycin, these derivatives showed early stage killing kinetics.

Due to the lack of crystallography or NMR data of the complex lipid II-teixobactin, until now it has not been possible to establish the interaction of teixobactin residues with lipid II experimentally. However, Lewis and co-workers (Ling et al., 2015) have hypothesized that lipid II pyrophosphate group and N-acetylmuramic acid are essential for the binding to teixobactin. This was further supported by recent MD simulation study of Liu et al. (2017) that showed the importance of the participation

## REFERENCES

of the oxygens of pyrophosphate group of lipid II with amide protons of the teixobactin cycle. In the MD simulation study carried out herein, similar interactions were also observed with Lys10-teixobactin (4) and pyrophosphate of lipid II (**Table 4** and **Figure 4**). These data suggest that the most dominant binding mode of Lys10-teixobactin (4) to lipid II is through the amide protons of the cycle, which is identical to data described in the literature for the natural teixobactin hence predicting the possibility of a similar mechanism of action.

Given these promising results, further research should address the mechanism/s of action exerted by these compounds. The findings of this study will contribute to the development of other Teixobactin derivatives with high potent antimicrobial activity against resistant bacterial strains and to the development of novel peptide-based antimicrobial agents to tackle the global threat of drug resistance.

# AUTHOR CONTRIBUTIONS

ER, AS, SM, DA, BT, and FA conceived and designed the experiments. ER, AS, SM, DA, HK, and NA performed the experiments. ER, AS, DA, RP, HK, and NA analyzed the data. RP, BT, FA, and LB contributed to reagents, materials, and analysis tools. ER wrote the paper. All authors did a critical revision of the manuscript.

# FUNDING

This study was supported by College of Health Sciences, University of KwaZulu-Natal, Durban, South Africa, and the South African National Research Foundation (NRF) (Grant number: 103107).

# ACKNOWLEDGMENTS

We would like to thank the Center for High-Performance Computing (CHPC), Cape Town for supercomputing resources.

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Ramchuran, Somboro, Abdel Monaim, Amoako, Parboosing, Kumalo, Agrawal, Albericio, de La Torre and Bester. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Novel Polymyxin Combination With Antineoplastic Mitotane Improved the Bacterial Killing Against Polymyxin-Resistant Multidrug-Resistant Gram-Negative Pathogens

Thien B. Tran1,2, Jiping Wang1,2, Yohei Doi3 , Tony Velkov2,4, Phillip J. Bergen2,5 and Jian Li1 \*

#### Edited by:

Maria Olivia Pereira, University of Minho, Portugal

#### Reviewed by: Vishvanath Tiwari,

Central University of Rajasthan, India Govindan Rajamohan, Institute of Microbial Technology (CSIR), India

\*Correspondence:

Jian Li<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Antimicrobials, Resistance and Chemotherapy, a section of the journal Frontiers in Microbiology

Received: 26 October 2017 Accepted: 27 March 2018 Published: 12 April 2018

#### Citation:

Tran TB, Wang J, Doi Y, Velkov T, Bergen PJ and Li J (2018) Novel Polymyxin Combination With Antineoplastic Mitotane Improved the Bacterial Killing Against Polymyxin-Resistant Multidrug-Resistant Gram-Negative Pathogens. Front. Microbiol. 9:721. doi: 10.3389/fmicb.2018.00721 1 Monash Biomedicine Discovery Institute, Department of Microbiology, School of Biomedical Sciences, Faculty of Medicine, Nursing and Health Sciences, Monash University, Melbourne, VIC, Australia, 2 Drug Delivery, Disposition and Dynamics, Monash Institute of Pharmaceutical Sciences, Monash University, Melbourne, VIC, Australia, 3 Division of Infectious Diseases, Department of Medicine, University of Pittsburgh Medical Center, Pittsburgh, PA, United States, 4 Department of Pharmacology and Therapeutics, School of Biomedical Sciences, Faculty of Medicine, Dentistry and Health Sciences, The University of Melbourne, Melbourne, VIC, Australia, 5 Centre for Medicine Use and Safety, Monash Institute of Pharmaceutical Sciences, Monash University, Melbourne, VIC, Australia

Due to limited new antibiotics, polymyxins are increasingly used to treat multidrug-resistant (MDR) Gram-negative bacteria, in particular carbapenem-resistant Acinetobacter baumannii, Pseudomonas aeruginosa, and Klebsiella pneumoniae. Unfortunately, polymyxin monotherapy has led to the emergence of resistance. Polymyxin combination therapy has been demonstrated to improve bacterial killing and prevent the emergence of resistance. From a preliminary screening of an FDA drug library, we identified antineoplastic mitotane as a potential candidate for combination therapy with polymyxin B against polymyxin-resistant Gram-negative bacteria. Here, we demonstrated that the combination of polymyxin B with mitotane enhances the in vitro antimicrobial activity of polymyxin B against 10 strains of A. baumannii, P. aeruginosa, and K. pneumoniae, including polymyxin-resistant MDR clinical isolates. Time-kill studies showed that the combination of polymyxin B (2 mg/L) and mitotane (4 mg/L) provided superior bacterial killing against all strains during the first 6 h of treatment, compared to monotherapies, and prevented regrowth and emergence of polymyxin resistance in the polymyxin-susceptible isolates. Electron microscopy imaging revealed that the combination potentially affected cell division in A. baumannii. The enhanced antimicrobial activity of the combination was confirmed in a mouse burn infection model against a polymyxin-resistant A. baumannii isolate. As mitotane is hydrophobic, it was very likely that the synergistic killing of the combination resulted from that polymyxin B permeabilized the outer membrane of the Gram-negative bacteria and allowed mitotane to enter bacterial cells and exert its antimicrobial effect. These results have important implications for repositioning non-antibiotic drugs for antimicrobial purposes, which may expedite the discovery of novel therapies to combat the rapid emergence of antibiotic resistance.

Keywords: polymyxin, mitotane, repurposing, combination therapy, multidrug-resistance

# INTRODUCTION

fmicb-09-00721 April 10, 2018 Time: 15:46 # 2

The emergence of Gram-negative bacteria with resistance to multiple classes of antibiotics is causing serious problems for health care centers worldwide (Boucher et al., 2013). Infections caused by multidrug-resistant (MDR) Gram-negative bacteria not only have higher mortality rates (Harris et al., 2015) but also lead to more economic burden than infections caused by susceptible Gram-negative bacteria (Gandra et al., 2014). Among these MDR Gram-negative bacteria, carbapenem-resistant Acinetobacter baumannii has been identified as one of the most difficult-to-treat pathogens and is becoming increasingly problematic for critically ill patients and war-wounded soldiers (Davis et al., 2005; Gupta et al., 2006; Peleg et al., 2008; Centers for Disease Control and Prevention [CDC], 2013). A. baumannii possesses numerous mechanisms of carbapenem resistance (Tiwari et al., 2012a,b; Tiwari and Moganty, 2014; Roy et al., 2017; Verma et al., 2017), and can cause a wide range of infections including pneumonia, urinary tract and wound infections, bacteremia, and meningitis (Gupta et al., 2006; Maragakis and Perl, 2008; Tiwari et al., 2012a). More recently, the World Health Organization (WHO) has classified carbapenem-resistant A. baumannii, Pseudomonas aeruginosa, and Enterobacteriaceae as the top priorities for research and development of new antibiotics (Tacconelli and Magrini, 2017).

Due to the current lack of effective antibiotics against MDR Gram-negative bacteria, the polymyxins (colistin and polymyxin B) have been revived as antibiotics of last resort (Li et al., 2006; Nation et al., 2015). However, resistance to polymyxins is on the rise (Marchaim et al., 2011; Kim et al., 2014; Goli et al., 2016) and a growing body of evidence suggests that resistance to polymyxins can emerge with monotherapy (Tam et al., 2005; Tan et al., 2007; Bergen et al., 2011; Meletis et al., 2011; Deris et al., 2012; Hermes et al., 2013; Lee et al., 2013; Ly et al., 2015; Lenhard et al., 2017; Zhao et al., 2017). In vitro studies have revealed that polymyxin resistance may occur within 24 h after colistin or polymyxin B monotherapy (Bergen et al., 2011; Deris et al., 2012; Tran et al., 2016). The two main mechanisms of polymyxin resistance identified in Gram-negative bacteria are lipid A modifications and loss of LPS (Moffatt et al., 2010; Arroyo et al., 2011).

Unfortunately, the de novo drug discovery and development process is lengthy (usually 10–17 years) and has a low success rate (<10%; Ashburn and Thor, 2004). With limited new antibiotics in the pipeline, an approach to expedite the discovery process is through the repositioning of non-antibiotic FDAapproved drugs. This process can be as short as 3 years as these drugs have already passed the FDA safety requirements and have well-defined pharmacokinetics (Ashburn and Thor, 2004). In light of the dire resistance problem, we screened an FDA drugs' library to identify potential synergistic candidates with polymyxins for the treatment of MDR Gram-negative bacteria. Our screening identified FDA-approved antineoplactic mitotane as a highly potential non-antibiotic candidate for combination therapy with polymyxin B. In this study, we evaluated the in vitro antimicrobial activity of the combination of polymyxin B and mitotane against highly resistant clinical isolates of Gram-negative bacteria including carbapenemresistant A. baumannii, carbapenem-resistant P. aeruginosa, and New Delhi metallo-β-lactamase (NDM)-producing Klebsiella pneumoniae. Our findings highlight the potential of this novel polymyxin/non-antibiotic combination for treatment of these problematic Gram-negative "superbugs".

# MATERIALS AND METHODS

# Bacterial Isolates

Ten bacterial strains which included multidrug- and polymyxin-resistant isolates were examined in this study (**Table 1**). A. baumannii ATCC 17978, A. baumannii ATCC 19606, K. pneumoniae ATCC 13883, and P. aeruginosa ATCC 27853 were obtained from the American Type Culture Collection (Rockville, MD, United States). A. baumannii FADDI-AB225 (formally designated ATCC 17978-R2) is a polymyxin-resistant pmrB mutant (due to phosphoethanolamine-modified lipid A) derived from ATCC 17978 (Arroyo et al., 2011). A. baumannii FADDI-AB065 (formally designated ATCC 19606R) is a polymyxin-resistant, LPS-deficient, lpxA mutant derived from ATCC 19606 (Moffatt et al., 2010). Polymyxin-susceptible A. baumannii FADDI-AB180 (formally designated 2949) and lipid A modified (with phosphoethanolamine and galactosamine) polymyxin-resistant A. baumannii FADDI-AB181 (formally designated 2949A) are carbapenem-resistant MDR clinical isolates from the bronchoalveolar lavage fluid of a patient before and after colistin therapy (Pelletier et al., 2013). P. aeruginosa FADDI-PA070 is a non-mucoid, MDR (including carbapenemand polymyxin-resistant) clinical isolate from the sputum of a patient with cystic fibrosis (formally designated 19147 n/m; Bergen et al., 2011). K. pneumoniae FADDI-KP027 is a polymyxin-resistant, NDM-producing clinical isolate from the sputum of a patient with respiratory tract infection. Isolates were stored in tryptone soy broth (Oxoid) with 20% glycerol (Ajax Finechem, Seven Hills, NSW, Australia) in cryovials at −80◦C and subcultured onto nutrient agar plates (Media Preparation Unit, University of Melbourne, Melbourne, VIC, Australia) before use.

TABLE 1 | Minimum inhibitory concentrations (MICs) for polymyxin B and mitotane against bacterial isolates examined in this study.

MDR, multidrug-resistant: defined as non-susceptible to ≥1 treating agent in ≥3 antimicrobial categories (Magiorakos et al., 2012). PR, polymyxin resistant: defined as an MIC of ≥4 mg/L for Acinetobacter spp. and ≥8 mg/L for P. aeruginosa as per CLSI guideline (Clinical and Laboratory Standards Institute [CLSI], 2016); and >2 mg/L for Enterobacteriaceae as per EUCAST guidelines (The European Committee on Antimicrobial Susceptibility Testing [EUCAST], 2017); and mitotane breakpoints are not available. –, Not performed.

# Antimicrobial Agents and Susceptibility Testing

Polymyxin B (Beta Pharma, China; batch number 20120204) solutions were prepared in Milli-Q water (Millipore, North Ryde, NSW, Australia) and sterilized using a 0.20-µm cellulose acetate syringe filter (Millipore, Bedford, MA, United States). Mitotane (Sigma-Aldrich, Australia; lot number BCBG9480V) solutions were prepared in dimethyl sulfoxide (DMSO; Sigma-Aldrich, Australia). Stock solutions were stored at −20◦C for no longer than 1 month. The minimum inhibitory concentrations (MICs) to polymyxin B and mitotane were determined for all isolates in three replicates on separate days using broth microdilution with cation-adjusted Mueller–Hinton broth (CAMHB; Oxoid, England; 20–25 mg/L Ca2+ and 10–12.5 mg/L Mg2+) according to the Clinical and Laboratory Standards Institute guidelines (Clinical and Laboratory Standards Institute [CLSI], 2016). Stock solutions of polymyxin B were diluted to the desired concentrations in CAMHB, while mitotane was initially prepared in DMSO and subsequently in CAMHB to obtain the desired drug concentrations with 10% DMSO (v/v). The procedure to measure the MICs of polymyxin B and mitotane was adapted from our previous method (Tran et al., 2016). Briefly, 100 µL of the bacterial suspension (106 cfu/mL) was combined with 100 µL of the prepared polymyxin B solutions or 50 µL of CAMHB plus 50 µL of the prepared mitotane solutions in 96-well microtiter plates (Techno Plas, St Marys, SA, Australia). For mitotane MICs, the final concentration of 2.5% DMSO (v/v) was employed, as preliminary studies demonstrated that 2.5% DMSO (v/v) had no effect on the bacterial growth. The plates were incubated standing at 37◦C for 20 h and MICs were determined as the lowest drug concentrations that inhibited the visible growth of the bacteria. For polymyxin-resistant isolates, MICs of mitotane in the presence of 2 mg/L of polymyxin B were also determined. According to the CLSI guidelines, polymyxin B MIC is ≤2 mg/L for polymyxin-susceptible A. baumannii and P. aeruginosa, ≥4 mg/L for polymyxin-resistant A. baumannii, and ≥8 mg/L for polymyxin-resistant P. aeruginosa (Clinical and Laboratory Standards Institute [CLSI], 2016). For K. pneumoniae, breakpoints have not yet been established by the CLSI. Consequently, susceptibility to polymyxin B was extrapolated from the European Committee on Antimicrobial Susceptibility Testing (EUCAST) colistin breakpoints where susceptibility is defined as an MIC ≤2 mg/L and resistance an MIC of >2 mg/L (The European Committee on Antimicrobial Susceptibility Testing [EUCAST], 2017).

# Time-Kill Studies

Time-kill studies were conducted for all isolates based on our previously described method (Tran et al., 2016). Briefly, bacteria were grown overnight in 20 mL CAMHB. The overnight broth cultures were transferred to 20 mL of fresh CAMHB at ∼50–100-fold dilutions and incubated for an additional 3–4 h to generate log-phase culture at ∼0.55 McFarland standard. The log-phase cultures were transferred to 20 mL of fresh CAMHB at ∼100-fold dilution in borosilicate glass tubes for treatment to minimize loss of drug due to non-specific binding to the plastic. For the drug-containing tubes, polymyxin B, mitotane, or both compounds were added to achieve final concentrations of 2 mg/L for polymyxin B and 4 mg/L for mitotane (the minimum concentration of mitotane identified by broth microdilution assay to inhibit to growth of polymyxin-resistant isolates in the presence of 2 mg/L polymyxin B). The final concentration of 0.4% DMSO (v/v) was achieved for all treatments; 2.5% DMSO (v/v) had no effect on bacterial growth with 2 mg/L of polymyxin B (Tran et al., 2016). Samples (1 mL) were aseptically removed at 0, 0.5, 1, 2, 4, 6, and 24 h and inoculated onto nutrient agar plates for viable-cell counting. Colonies were counted after 24 h incubation at 37◦C using a ProtoCOL colony counter (Synbiosis, Cambridge, United Kingdom). The combination of polymyxin B and mitotane was considered synergistic if the bacterial killing was ≥2 log10 compared to the most active

monotherapy (Pillai et al., 2005). Changes to polymyxin B MICs were determined for all cultures that showed regrowth after 24 h to evaluate the emergence of polymyxin resistance.

# Phase Contrast, Scanning Electron, and Transmission Electron Microscopy

Phase contrast microscopy, scanning electron microscopy (SEM), and transmission electron microscopy (TEM) were employed to examine the effect of the polymyxin B/mitotane combination on the cellular morphology of polymyxinsusceptible A. baumannii ATCC 17978 and polymyxin-resistant A. baumannii FADDI-AB225. Bacteria were subcultured and treated with 2 mg/L polymyxin B, 4 mg/L mitotane, or both antibiotics for 2 h in CAMHB as per the time-kill studies. For phase contrast microscopy, 20 µL of each culture was used to prepare wet samples for instant observation on a phase contrast microscope. For the SEM and TEM studies, samples were transferred to 50-mL polypropylene tubes (Greiner Bio-One, Frickenhausen, Germany) and centrifuged at 3220 × g for 10 min three times. Between centrifugation steps, supernatants were discarded and bacterial pellets resuspended and washed in 1 mL phosphate buffered saline (PBS). Following the final centrifugation step, the supernatants were removed and bacterial pellets resuspended and fixed in 0.5 mL 2.5% glutaraldehyde in PBS. The tubes were left in a rocker shaker for 20 min at room temperature. Once fixed, tubes were centrifuged at 3220 × g for 10 min, the fixatives removed, and bacterial pellets washed twice in 1 mL PBS as above. Pellets were finally resuspended in 1 mL PBS, and SEM and TEM were conducted at the Department of Botany, University of Melbourne, Australia.

# Mouse Burn Wound Infection Model

A mouse burn wound infection model was employed to assess the in vivo antimicrobial activity of the polymyxin B/mitotane combination against polymyxin-resistant A. baumannii FADDI-AB225. Bacterial inoculums were prepared with early log-phase culture. After centrifugation at 3220 × g for 10 min, the supernatant was removed and bacterial cell pellets were suspended in 0.9% saline to approximately 109 cfu/mL. Bacterial samples (100 µL) were then loaded into 29-G 0.3-mL insulin syringes for inoculation of burn wounds. Drug solutions were prepared by initially dissolving mitotane in polyethylene glycol (PEG) 200 to ∼4,096 mg/L and polymyxin B in 0.9% saline to ∼1,536 mg/L. An equal amount of the two drug solutions was later combined to produce the combination solution with ∼2,048 mg/L mitotane and 768 mg/L polymyxin B. For mitotane monotherapy, mitotane solution was combined with an equal volume of 0.9% saline. For polymyxin B monotherapy, polymyxin B was combined with an equal volume of PEG 200. For solvent controls, equal volumes of blank PEG 200 and 0.9% saline were combined. Prior to infection, female NIH Swiss mice (6–10 weeks old, ∼30 g body weight) were sedated with isoflurane and anesthesia was maintained throughout the entire procedure. Hair from the mouse dorsal skin was removed and the local skin area was injected with 100 µL of Bupivacaine (Marcaine 0.5%). A burn wound was established with a hot iron bolt from boiling water and bacteria injected into the burn eschar. After 2 h, different treatments were applied topically by evenly spreading 200 µL of the drug solutions across the wounds of groups of four mice. This study included five groups of four mice comprising blank control (no treatment), solvent control, polymyxin B monotherapy, mitotane monotherapy, and the combination (polymyxin B and mitotane). Each wound of the treated groups received 154 µg of polymyxin B (0.5%, w/w), 410 µg of mitotane (1.4%, w/w), or both. Four hours after treatment, mice were sacrificed and the burn wound skin tissues and the muscle tissue (∼0.3 g) under the burn wounds were aseptically removed and placed separately into 8 mL of sterile saline in 50-mL Falcon tubes. Burn wound skin tissues were homogenized under sterile conditions and filtered using a filter bag (Bag Stomacher Filter Sterile, Pore Size 280 micrometer, 0.5 cm × 16 cm, Labtek Pty Ltd.). Filtrate (1 mL) was then transferred into a sterile test tube for serial dilution and 100 µL was cultured onto nutrient agar for viable counting. Viable counts were performed on the next day following overnight incubation at 37◦C. Statistical significance for the bacterial killing of different treatment groups was calculated with one-way ANOVA and Tukey's multiple comparisons (Tukey's HSD).

# RESULTS

# MICs of Polymyxin B and Mitotane Against Polymyxin-Susceptible and -Resistant Isolates of A. baumannii, P. aeruginosa, and K. pneumoniae

The polymyxin B and mitotane MICs against all 10 Gram-negative isolates are shown in **Table 1**. Additionally, **Table 1** shows the MICs of mitotane in the presence of 2 mg/L polymyxin B against the polymyxin-resistant isolates. Apart from A. baumannii FADDI-AB065, mitotane monotherapy had no antimicrobial activity at concentrations up to 128 mg/L. However, in the presence of 2 mg/L polymyxin B, 4 mg/L of mitotane was effective at inhibiting growth of five polymyxin-resistant isolates (**Table 1**).

The changes to the polymyxin B MICs of 10 examined isolates after overnight treatment with either polymyxin B monotherapy, mitotane monotherapy, or polymyxin B/mitotane combination are shown in **Table 2**. In the control group (overnight incubation in drug-free CAMHB), polymyxin B MICs of all isolates at 24 h were not affected as all values remained within two folds of the baseline MICs (European Committee for Antimicrobial Susceptibility Testing (EUCAST) of the European Society of Clinical Microbiology and Infectious Diseases (ESCMID), 2003). After treatment with polymyxin B monotherapy at 2 mg/L, polymyxin B MICs of the polymyxin-resistant isolates at 24 h remained unchanged. However, with the three polymyxin-susceptible isolates that showed regrowth at 24 h, polymyxin B MICs of the 24-h samples increased significantly (≥32 times). Following mitotane monotherapy at 4 mg/L, polymyxin B MICs remained unchanged for all polymyxin-susceptible isolates and

TABLE 2 | Changes in baseline polymyxin B MICs following overnight treatment with polymyxin B (PMB) monotherapy, mitotane (MIT) monotherapy, and polymyxin B/mitotane combination.

NG, no growth at 24 h.

three polymyxin-resistant isolates; the polymyxin B MIC of polymyxin-resistant A. baumannii FADDI-AB065 at 24 h could not be determined, as it was highly susceptible to mitotane and showed no regrowth after 24 h. Interestingly, the polymyxin B MIC of polymyxin-resistant A. baumannii FADDI-AB225 was reduced significantly (32-fold lower than the baseline) after 24-h exposure to mitotane. In the combination treatment group, the polymyxin B MICs did not change for all four polymyxin-resistant isolates that showed regrowth after 24 h.

# Time-Kill Results for Polymyxin B and Mitotane Against Polymyxin-Susceptible and -Resistant Isolates of A. baumannii, P. aeruginosa, and K. pneumoniae

Time-kill profiles for polymyxin B and mitotane mono- and combination therapy are shown in **Figure 1**. Against the five polymyxin-susceptible isolates, polymyxin B monotherapy (2 mg/L) showed effective bacterial killing within 6 h with a minimum of ∼3 log10 cfu/mL killing (FADDI-AB180) and ∼6 log10 cfu/mL killing for the remaining susceptible isolates; however, regrowth to control values occurred by 24 h with three isolates (**Figure 1A**). There was no bacterial killing of polymyxin-susceptible isolates with mitotane monotherapy (4 mg/L), with growth comparable to that of controls (**Figure 1A**). With the combination, bacterial counts for all five polymyxin-susceptible isolates were reduced to below the limit of detection within 0.5–1 h, with no viable colonies detected thereafter (**Figure 1A**). Against the five polymyxin-resistant isolates, 2 mg/L polymyxin B monotherapy was ineffective with growth paralleling that of the controls (**Figure 1B**). Similarly, mitotane monotherapy displayed no antimicrobial activity against four of the five isolates (**Figure 1B**). However, against A. baumannii FADDI-AB065 mitotane monotherapy reduced bacterial counts to below the level of detection within the first 0.5 h and prevented regrowth over 24 h. Combination treatment showed synergistic bacterial killing (i.e., >2 log10 reduction compared to the most active monotherapy) between 0.5 and 6 h with the remaining four isolates; interestingly, regrowth occurred at 24 h in all four cases and was close to control values in three cases (**Figure 1B**).

# Impact of Polymyxin B and Mitotane Treatment on the Cellular Morphology of Polymyxin-Susceptible and -Resistant A. baumannii

**Figure 2** shows phase contrast microscopy, SEM and TEM images of polymyxin-susceptible A. baumannii ATCC 17978 following treatment with polymyxin B (2 mg/L), mitotane (4 mg/L), or both. Phase contrast microscopy images showed that polymyxin B (**Figure 2B**) or mitotane (**Figure 2C**) monotherapy had minimal impacts on the overall morphology of the bacterial cells compared to the control group (**Figure 2A**); the average cell length remained approximately 2 µm in all cases. However, more clumps of cells were observed with polymyxin B monotherapy (**Figure 2B**). In combination (**Figure 2D**), polymyxin B and mitotane resulted in significantly shorter cells compared to the other groups with the average cell length reduced to approximately 1 µm. From SEM, polymyxin B monotherapy (**Figure 2F**) affected the integrity of the cell surface in polymyxin-susceptible A. baumannii. Without treatment (**Figure 2E**), the bacterial surface appeared even and smooth, while the surface became uneven and rough following treatment with polymyxin B (**Figure 2F**). Mitotane monotherapy (**Figure 2G**) and polymyxin B/mitotane combination therapy (**Figure 2H**) had minimal impacts on the bacterial surface, although the cell length was confirmed to be much shorter. TEM results reveal that polymyxin B monotherapy (**Figure 2J**) caused membrane blebbing. Compared to the control group (**Figure 2I**), treatment with mitotane monotherapy (**Figure 2K**) had little impact on the bacterial surface. Similar to polymyxin B monotherapy, membrane blebbing was also observed for the treatment with polymyxin B/mitotane combination (**Figure 2L**). Additionally, TEM images showed that bacterial cells treated with the polymyxin B/mitotane combination were much shorter in length and most appeared to be undergoing a cell division cycle, with evident chromosomal segregation.

indicated by the orange dotted line.

Phase contrast microscopy, SEM, and TEM images for polymyxin-resistant A. baumannii FADDI-AB225 treated with polymyxin B (2 mg/L), mitotane (4 mg/L), or both are shown in **Figure 3**. Similar to the results for polymyxin-susceptible A. baumannii ATCC 17978, phase contrast microscopy results showed no changes in bacterial size compared to the control group (**Figure 3A**) following treatment with polymyxin B (**Figure 3B**) and mitotane (**Figure 3C**) monotherapy, while the polymyxin B/mitotane combination (**Figure 3D**) led to a significant reduction in the cell length. For SEM, treatment with polymyxin B monotherapy (**Figure 3F**) did not affect the bacterial cell surface; however, the overall structure appeared distorted. Treatment with mitotane monotherapy (**Figure 3G**) affected the cell surface of polymyxin-resistant A. baumannii FADDI-AB225, as the surface was more uneven and rough compared to the control group (**Figure 3E**). Combination therapy (**Figure 3H**) did not affect the membrane surface, although it led to substantial shortening of the cells. For TEM, similar results to polymyxin-susceptible isolates were once again observed. Membrane blebbing was evident in bacteria treated only with polymyxin B (**Figure 3J**), but not in those treated only with mitotane (**Figure 3K**). With the polymyxin B/mitotane combination (**Figure 3L**), most cells were substantially shorter compared to the control group (**Figure 3I**) and appeared to be going through cell division. Unlike the polymyxin-susceptible isolate, no membrane blebbing was observed with the combination in the polymyxin-resistant isolate.

FIGURE 2 | Images from phase contrast microscopy (A–D), scanning electron microscopy (E–H), and transmission electron microscopy (I–L) for polymyxin-susceptible A. baumannii ATCC 17978 treated with 2 mg/L polymyxin B (B,F,J), 4 mg/L mitotane (C,G,K), or both (D,H,L). A, E, and I represent the control condition. Membrane blebs are indicated by red circles.

# In Vivo Antimicrobial Activity of Polymyxin B and Mitotane Against Polymyxin-Resistant A. baumannii FADDI-AB225 in a Mouse Burn Wound Infection Model

**Figure 4** shows the bacterial killing of polymyxin B (0.5%, w/w), mitotane (1.4%, w/w), and the polymyxin B/mitotane combination against polymyxin-resistant A. baumannii FADDI-AB225. One-way ANOVA showed significant difference between the means of all groups (p < 0.0001). There was no significant difference in the bacterial load between the blank control (i.e., no treatment) and solvent control groups (mean log10 cfu/wound difference, −0.33; Tukey's HSD, p > 0.05), indicating that the solvent possessed no major antimicrobial activity. Although this isolate was polymyxin-resistant, topical polymyxin B (0.5%, w/w) monotherapy significantly reduced the bacterial load (mean log10 cfu/wound difference, −1.44 vs. blank control; Tukey's HSD, p ≤ 0.0001). However, there was no significant reduction in the bacterial load (mean log10 cfu/wound difference, −0.11 vs. blank control; Tukey's HSD, p > 0.5) with topical mitotane (1.4%, w/w) alone (**Figure 4**). Importantly, both agents used in combination produced a further significant reduction in the bacterial load compared to polymyxin B monotherapy (mean log10 cfu/wound difference, −0.74; Tukey's HSD, p ≤ 0.01). Compared to the blank control group, the polymyxin B/mitotane combination resulted in a mean log10 cfu/wound difference of −2.19 (Tukey's HSD, p ≤ 0.0001).

# DISCUSSION

Given the rapid emergence of multidrug-resistance and the limited new effective antibiotics developed over the last two decades (Boucher et al., 2009, 2013), novel approaches for the treatment of MDR Gram-negative bacteria infections are urgently needed. This is the first study to investigate the potential utility of polymyxin B in combination with the FDA-approved antineoplastic mitotane to treat infections caused by polymyxin-resistant MDR Gram-negative pathogens. Mitotane is a derivative of the insecticide dichlorodiphenyl-trichloroethane and is currently used for the treatment of adrenocortical carcinoma (ACC) (Lalli, 2015). The precise mechanism of action of mitotane in ACC is not well understood, but it has been shown to inhibit the activity of sterol-O-acyl-transferase and induce endoplasmic reticulum (ER) stress in ACC cells (Sbiera et al., 2015). Our study is the first to demonstrate its potential application for the treatment of Gram-negative infections when combined with polymyxin B.

To ensure the applicability of the combination of polymyxin B and mitotane to a diverse population of problematic Gram-negative bacteria, three Gram-negative bacterial species

(A. baumannii, P. aeruginosa, and K. pneumoniae) were selected for the initial in vitro antimicrobial activity evaluation. Isolates selected included MDR, carbapenem-resistant, and polymyxin-resistant strains with known different mechanisms of polymyxin resistance. A. baumannii and P. aeruginosa were selected as they are frequently resistant to multiple classes of antibiotics and are currently considered by the WHO as two of the top bacterial "superbugs" requiring urgent antibiotic development (Tacconelli and Magrini, 2017). K. pneumoniae was also examined as it is also identified as a top bacterial "superbug" by the WHO due to the rapid emergence of carbapenem resistance (including NDM production) (Yong et al., 2009; Kumarasamy et al., 2010; Farzana et al., 2013). Concentrations of 2 mg/L for polymyxin B and 4 m/L for mitotane were examined as they were achievable in patients (Hermsen et al., 2011; Sandri et al., 2013).

control condition. Membrane blebs are indicated by red circles.

One of the major concerns surrounding the intravenous use of polymyxin B or colistin monotherapy for the treatment of infections caused by Gram-negative bacteria is the development of resistance via amplification of polymyxin-resistant subpopulations (Tam et al., 2005; Tan et al., 2007; Bergen et al., 2011; Meletis et al., 2011; Deris et al., 2012; Hermes et al., 2013; Lee et al., 2013; Ly et al., 2015; Lenhard et al., 2017; Zhao et al., 2017). Consequently, the use of antibiotic combination therapy represents a potential option to increase bacterial killing and prevent the emergence of polymyxin resistance as the combination may result in subpopulation or mechanistic synergy (Landersdorfer et al., 2013). Despite extensive bacterial killing by polymyxin B monotherapy against five polymyxin-susceptible isolates, regrowth with associated polymyxin resistance (the latter evident by significantly increased polymyxin B MICs compared to baseline values) subsequently occurred with three isolates (A. baumannii ATCC 19606, A. baumannii FADDI-AB180, and K. pneumoniae ATCC 13883; **Figure 1A**). When used as monotherapy, mitotane showed antimicrobial activity against only one isolate (**Figure 1B**). However, the combination of polymyxin B and mitotane significantly improved bacterial killing against the less susceptible isolates (i.e., those that were resistant to polymyxin B or mitotane monotherapy, or showed regrowth at 24 h; **Figures 1A,B**). The enhanced antimicrobial killing was indicated by the complete prevention of regrowth in all polymyxin-susceptible isolates after 24 h (**Figure 1A**) and >2 log10 cfu/mL reduction within the first 6-h treatment against the four polymyxin-resistant isolates compared to the more active monotherapy (**Figure 1B**). Although regrowth occurred in four of the five polymyxin-resistant isolates, the combination still enhanced initial bacterial killing which may assist with the bacterial clearance from the body. Since polymyxins are well known for their ability to permeabilize the outer membrane of Gram-negative bacteria (Salmelin et al., 2000; Tsubery et al., 2000; Sahalan and Dixon, 2008), a possible mechanism for the enhanced killing observed with the combination is

permeabilization of the outer membrane by polymyxin B leading to the entry of mitotane into the bacterial cell. Indeed, polymyxin B and its derivative polymyxin B nonapeptide had previously been shown to enhance the antimicrobial activity of hydrophobic antibiotics against Gram-negative bacteria and yeasts (Ofek et al., 1994; Pietschmann et al., 2009). Interestingly, mitotane monotherapy displayed substantial antimicrobial activity against LPS-deficient, polymyxin-resistant A. baumannii FADDI-AB065 (**Figure 1B**). LPS in the outer membrane of Gram-negative bacteria acts as a highly selective permeability barrier that protects bacteria from harmful substances (Nikaido, 2003). Consequently, it is possible that in the absence of LPS, mitotane was able to enter bacterial cells and exert its antimicrobial activity. Another notable finding is that mitotane monotherapy also lowered the polymyxin B MIC of polymyxin-resistant A. baumannii FADDI-AB225 (**Table 2**); however, it did not affect the polymyxin B MICs of the other polymyxin-resistant isolates. The mechanism for this phenomenon is currently unclear, although it may result from the expression of LPS variants by the different isolates. Coincidently, it has been reported that Moraxella catarrhalis and Salmonella typhimurium with deep rough-type LPS displayed higher susceptibility to hydrophobic antimicrobial agents (Tsujimoto et al., 1999). Further mechanistic studies are warranted.

According to the SEM imaging results, it is possible that the polymyxin resistance in A. baumannii FADDI-AB225 altered their surface interaction with mitotane, as the outer membrane appeared disrupted (uneven and rough) following mitotane monotherapy in A. baumannii FADDI-AB225 (**Figure 3G**), but not A. baumannii ATCC 17978 (**Figure 2G**). Both the SEM and TEM images showed disruptive changes to the outer membrane of polymyxin-susceptible A. baumannii ATCC 17978 following polymyxin B monotherapy (**Figures 2F,J**), which confirmed the known impact of polymyxin B on the outer membrane of Gramnegative bacteria. For the lipid A modified polymyxin-resistant A. baumannii FADDI-AB225, no disruptive effect on the surface membrane by polymyxin B monotherapy was observed with SEM (**Figure 3F**), most likely due to the modification of lipid A which resulted in minimal polymyxin B affinity. Membrane blebs, however, were still observed by TEM in polymyxinresistant A. baumannii FADDI-AB225 treated with 2 mg/L polymyxin B alone (**Figure 3J**), indicating that blebbing might not necessarily result in cell death, but was enough to allow the mitotane to enter and exert antibacterial effect. Although monotherapy with mitotane or polymyxin B appeared to impact the outer membrane of polymyxin-resistant and -susceptible A. baumannii, the combination impacted the overall structure of both strains leading to an extensive shortening in the length of the bacteria (**Figures 2**, **3**). SEM images showed a smooth membrane surface on the shortened bacterial cells, suggesting that the combination prevented the formation of the rough surface, which could be an adaptive response to polymyxin B or mitotane monotherapy. Numerous incompletely separated cells revealed by TEM images (**Figures 2L**, **3L**) suggest a possible impact on the bacterial DNA replication.

In our mouse burn wound infection study, the combination displayed effective antimicrobial activity against polymyxinresistant A. baumannii. The doses of 5 mg/kg for polymyxin B (subcutaneous median lethal dose in mice [LD50] 59 mg/kg) and 14 mg/kg for mitotane (oral LD50 > 4,000 mg/kg in mice, dermal LD50 not available) were selected, as they are safe in animals according to their material safety data sheets. Based on the available LD50 limits of polymyxin B and mitotane, it is likely that much higher doses of both drugs can be used for topical combination therapy. Given the lack of an optimized topical formulation, it is possible that the in vivo efficacy of the combination in the current study was underestimated. Nevertheless, the combination treatment was able to significantly reduce the number of polymyxin-resistant A. baumannii, compared to polymyxin B or mitotane monotherapy.

# CONCLUSION

Our study is the first to reveal the synergistic activity of mitotane, an FDA-approved non-antibiotic drug, in combination with polymyxin B against problematic Gram-negative bacteria. Importantly, the combination also prevented the emergence of polymyxin resistance. As mitotane is currently used in humans, its repositioning for antimicrobial purposes may be easier than discovering novel antibacterial compounds against Gram-negative "superbugs". The synergistic antibacterial killing of polymyxin B with mitotane in animals raises hopes for the potential repositioning of mitotane against MDR Gram-negative bacteria and further clinical investigations are warranted.

# ETHICS STATEMENT

fmicb-09-00721 April 10, 2018 Time: 15:46 # 10

This study was carried out in accordance with the recommendations of "Australian Code of Practice for the Care and Use of Animals for Scientific Purposes" and Monash Institute of Pharmaceutical Sciences Animal Ethics Committee. The protocol was approved by the Monash Institute of Pharmaceutical Sciences Animal Ethics Committee before the study started.

# AUTHOR CONTRIBUTIONS

TT carried out the main experiments, data analysis, and wrote the manuscript draft. JW participated in the animal study. PB

# REFERENCES

participated in in vitro studies' design. YD and TV participated in data analysis. JL designed the project and guided all experimental designs and data analysis. All authors participated in manuscript revision and read and approved the final manuscript.

# FUNDING

This study was supported by a research grant from the National Institute of Allergy and Infectious Diseases of the National Institutes of Health (R01AI111965) awarded to JL and TV. YD was supported in part by R01AI10 4895.

aeruginosa: profiling the time course of synergistic killing and prevention of resistance. J. Antimicrob. Chemother. 70, 1434–1442. doi: 10.1093/jac/dku567

**Conflict of Interest Statement:** The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institute of Allergy and Infectious Diseases or the National Institutes of Health. JL is an Australian National Health and Medical Research Council (NHMRC) Senior Research Fellow. TV is an Australian NHMRC Industry Career Development Research Fellow.

The other authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Tran, Wang, Doi, Velkov, Bergen and Li. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Low Concentrations of Vitamin C Reduce the Synthesis of Extracellular Polymers and Destabilize Bacterial Biofilms

Santosh Pandit1 , Vaishnavi Ravikumar1 , Alyaa M. Abdel-Haleem2,3 , Abderahmane Derouiche1 , V. R. S. S. Mokkapati1 , Carina Sihlbom4 , Katsuhiko Mineta2 , Takashi Gojobori2 , Xin Gao2 , Fredrik Westerlund1 and Ivan Mijakovic1,5 \*

1 Systems and Synthetic Biology Division, Department of Biology and Biological Engineering, Chalmers University of Technology, Gothenburg, Sweden, 2 Computational Bioscience Research Center, King Abdullah University of Science and Technology, Thuwal, Saudi Arabia, 3 Biological and Environmental Sciences and Engineering Division, King Abdullah University of Science and Technology, Thuwal, Saudi Arabia, 4 Proteomics Core Facility, Sahlgrenska Academy, University of Gothenburg, Gothenburg, Sweden, 5 Novo Nordisk Foundation Center for Biosustainability, Technical University of Denmark, Kongens Lyngby, Denmark

#### Edited by:

Mariana Henriques, University of Minho, Portugal

#### Reviewed by:

Vishvanath Tiwari, Central University of Rajasthan, India Ana Isabel Pelaez, Universidad de Oviedo Mieres, Spain

#### \*Correspondence:

Ivan Mijakovic<EMAIL_ADDRESS><EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Antimicrobials, Resistance and Chemotherapy, a section of the journal Frontiers in Microbiology

Received: 15 October 2017 Accepted: 13 December 2017 Published: 22 December 2017

#### Citation:

Pandit S, Ravikumar V, Abdel-Haleem AM, Derouiche A, Mokkapati VRSS, Sihlbom C, Mineta K, Gojobori T, Gao X, Westerlund F and Mijakovic I (2017) Low Concentrations of Vitamin C Reduce the Synthesis of Extracellular Polymers and Destabilize Bacterial Biofilms. Front. Microbiol. 8:2599. doi: 10.3389/fmicb.2017.02599 Extracellular polymeric substances (EPS) produced by bacteria form a matrix supporting the complex three-dimensional architecture of biofilms. This EPS matrix is primarily composed of polysaccharides, proteins and extracellular DNA. In addition to supporting the community structure, the EPS matrix protects bacterial biofilms from the environment. Specifically, it shields the bacterial cells inside the biofilm, by preventing antimicrobial agents from getting in contact with them, thereby reducing their killing effect. New strategies for disrupting the formation of the EPS matrix can therefore lead to a more efficient use of existing antimicrobials. Here we examined the mechanism of the known effect of vitamin C (sodium ascorbate) on enhancing the activity of various antibacterial agents. Our quantitative proteomics analysis shows that nonlethal concentrations of vitamin C inhibit bacterial quorum sensing and other regulatory mechanisms underpinning biofilm development. As a result, the EPS biosynthesis in reduced, and especially the polysaccharide component of the matrix is depleted. Once the EPS content is reduced beyond a critical point, bacterial cells get fully exposed to the medium. At this stage, the cells are more susceptible to killing, either by vitamin C-induced oxidative stress as reported here, or by other antimicrobials or treatments.

Keywords: biofilms, exopolymeric matrix, quantitative proteomics, Bacillus subtilis, vitamin C

# INTRODUCTION

Bacterial biofilms are culprits of various human infectious diseases, industrial corrosion and food contamination (Flemming et al., 2016). Bacteria within the biofilms synthesize a dense protective matrix composed of extracellular polymeric substances (EPS) (Branda et al., 2005). This matrix is mainly composed of polysaccharides, proteins and extracellular DNA (eDNA), whose continuous release leads to the establishment of a complex "mushroom-shaped" biofilm architecture (Branda et al., 2006; Barnes et al., 2012). Exopolysaccharides and proteins are the most abundant component of the biofilm matrix, defining its physico-chemical properties and morphology (Marvasi et al., 2010; Roy et al., 2017). Furthermore, the EPS serve as a food storage, which gets mobilized during extended nutrient depletion (Xiao et al., 2012). The structure of the EPS matrix varies considerably

among bacterial strains, and its composition is influenced by the local environment and nutrient availability.

Antibiotics are widely used to eradicate bacterial biofilms when treating infections. However, their prolonged use increases the risk of developing multi-resistant strains, and disrupts the ecology of the residential microflora (Cegelski et al., 2008). Hence an increasing interest for chemo-prophylactic agents, which can affect biofilm formation, and thereby reduce the time and dose of antibiotics treatments (Xavier et al., 2005; Cegelski et al., 2008). Vitamin C, a major dietary micronutrient, has been shown to exhibit bactericidal activity against mycobacteria (Vilcheze et al., 2013). However, this killing effect seems to be confined to mycobacteria, since vitamin C did not kill other opportunistic bacterial pathogens, such as Staphylococcus epidermidis, Staphylococcus aureus, Escherichia coli, and Pseudomonas aeruginosa (Khameneh et al., 2016). Interestingly, vitamin C has been reported to enhance the effect of antibiotics vs. a broad spectrum of bacteria via a synergistic effect, but the mechanism of this synergy remains unclear (Kallio et al., 2012; Khameneh et al., 2016). Similarly, vitamin C has been shown to enhance the killing effect of a physical bactericidal agent, cold atmospheric plasma, against biofilms of S. epidermidis, E. coli, and P. aeruginosa (Helgadóttir et al., 2017). In this study we set out to characterize the mechanism of this non-lethal synergistic effect of vitamin C, which enhances the effect of antibiotics and physical killing agents.

We performed an initial characterization with several bacterial strains, and different doses of vitamin C. Our conclusion was that while the low doses of vitamin C are harmless to the planktonic bacteria, they effectively destabilizes biofilms. We then focused on an in-depth quantitative analysis with Bacillus subtilis, a model organism for biofilm development (Vlamakis et al., 2013). Our findings, based on quantification of the biofilm EPS content and cell viability, quantitative proteome analyses and genome-scale metabolic modeling point to a vitamin C-dependent inhibition of the synthesis of polysaccharides that form the biofilm matrix. This proceeds via inhibition of the quorum sensing and other regulatory mechanisms, leading to repression of specific biosynthetic operons. Once the EPS content is reduced beyond a critical point, bacterial cells become exposed, and more susceptible to killing by any external factors.

# MATERIALS AND METHODS

# Bacterial Strains, Culture Media, and Reagents

Bacillus subtilis NCIB 3610, E. coli UTI89 and P. aeruginosa PAO1 were used in this study. LB (10 g of tryptone, 5 g of yeast extract and 5 g of NaCl per liter) or solid LB medium supplemented with 1.5% agar were used for the routine growth of all bacteria. Sodium ascorbate was purchased from Sigma–Aldrich.

# Bacterial Growth and Biofilm Formation

For growth analysis, an overnight bacterial culture was diluted 1:100 (1 × 107 CFU) in LB medium with 1% glycerol for B. subtilis, plain LB medium for P. aeruginosa and E. coli, with varying concentrations of sodium ascorbate (neutral pH form). The diluted cultures were incubated at 37◦C with continuous agitation (200 rpm) and the absorbance of the culture was measured periodically at 600 nm for 9 h with intervals of 1 h. B. subtilis biofilms were formed in LBGM medium (LB medium containing 1% glycerol; 1 mM MnSO4). 2–5 × 106 CFU/mL of bacterial culture was inoculated into 5 mL of LBGM medium and incubated at 37◦C for 24 h without agitation. E. coli and P. aeruginosa biofilms were formed on 24 well plates. 2–5 × 106 CFU/mL of an overnight bacterial culture was inoculated into a 24 well plate containing 2 mL of LB broth and incubated for 24 h without agitation.

# B. subtilis Biofilm Analysis

For the biofilm analysis, 24 h old biofilms, grown in the presence of various concentrations of sodium ascorbate, were removed and sonicated at 10 W for 30 s to homogenize the biofilm. The homogenized suspension (5 mL) was used to determine the biomass, colony forming units (CFU), polysaccharides, protein and eDNA. Briefly, for the determination of biomass, the homogenized suspension was washed three times (5000 g for 20 min) with sterile water, lyophilized and weighed. For the determination of viability, an aliquot (100 µL) from the homogenized suspension was diluted serially and plated on LB agar plates to count colonies. Water insoluble polysaccharide was extracted from the lyophilized sample by using 1 N sodium hydroxide (300 µL/mg biomass for three times) and quantified by using a phenol-sulfuric acid assay as described previously (Pandit et al., 2011). For protein quantification, biofilms were collected and homogenized in 1 N NaOH (300 µL/mg biomass for three times). The supernatant from the homogenized suspensions were collected after centrifugation (5000 g, 20 min) and protein content was quantified by using the Bradford assay. For quantification of eDNA in the EPS matrix, the filtered supernatant from the homogenized suspension was used. eDNA was extracted by using a DNA extraction kit (Thermo Fisher Scientific) and the quantity was measured using a nanodrop UV-Vis spectrophotometer (NanoDrop 2000, Thermo Scientific).

# Fluorescence Microscopy Analysis

The effect of sodium ascorbate on B. subtilis biofilms was analyzed by simultaneous labeling of the bacterial cells and the polysaccharides in the biofilm. Briefly, 10 µg/mL of Alexa flour R 633-labeled wheat germ agglutinin conjugate (absorbance/fluorescence emission maxima 632/647 nm; Molecular Probes Inc., Eugene, OR, United States) and 50 µg/mL of Concanavalin A, Tetramethylrhodamine conjugate (555/580 nm; Molecular Probes) was added to the culture medium during biofilm formation. The toxicity of these fluorescence probe toward the bacterial cells in biofilms was evaluated by comparison of viability and biomass with control samples. After 24 h, the biofilms were exposed to 2.5 µm of SYTO R 9 green-fluorescent nucleic acid stain (480/500 nm; Molecular Probes) for 30 min. The stained biofilms were transferred to a glass slide and laser scanning confocal microscope imaging of the biofilms was performed using an LSM 710 NLO (Carl Zeiss) equipped with argon-ion and helium

neon lasers. Three independent experiments were performed and image stacks from five sites per experiment were collected (n = 15). EPS biovolume was quantified from confocal stacks by COMSTAT (Heydorn et al., 2000). Biovolume is defined as the volume of the biomass (µm3 ) divided by substratum area (µm2 ). For the detection of reactive oxygen species (ROS) in biofilm cells, 24 h old B. subtilis biofilms grown with or without presence of vitamin C were stained with DAPI and CellRox deep ROS sensor (Life Technologies) as described previously (Durmus et al., 2013). Briefly, biofilms were stained with 5 µM of CellRox deep red stain for 30 min, washed with sterile water and counter- stained with DAPI for 20 min. The stained biofilms were visualized by fluorescence microscopy. To visualize the live/dead cells in B. subtilis biofilms grown with and without presence of vitamin C, biofilms were stained with 6.0 µM SYTO 9 and 30 µM propidium iodide (LIVE/DEAD BacLight bacterial viability kit L13152, Invitrogen, Molecular Probes, Inc., Eugene, OR, United States). Imaging was performed with a fluorescence microscope (Axio Imager 2. Carl Zeiss, Zena, Germany).

# Assay for Biofilm Formation of E. coli and P. aeruginosa

For the biofilm formation assay, 24 h old biofilms of E. coli and P. aeruginosa were rinsed three times with sterile water to remove the loosely adherent bacteria and dried for 30 min at room temperature. Dried biofilms were then stained with 1% crystal violet for 5 min without agitation. All the biofilms were washed at least five times with sterile water to remove the excess stain and dried for 1 h at room temperature. Absolute ethanol (1 mL) was added to the dried stained biofilm and agitated vigorously for 15 min to dissolve the stain. Optical density was measured at 600 nm.

# Proteome Analysis

All experiments for MS analysis were carried out in biological triplicates. 24 h biofilms of B. subtilis were collected and centrifuged to obtain a cell pellet. Cell lysis was performed by re-suspending the cell pellets in an SDS lysis buffer containing 4% SDS in 100 mM triethylammonium bicarbonate pH 8.5, 5 mM β-glycerophosphate, 5 mM sodium fluoride, 5 mM sodium orthovanadate and 10 mM ethylenediaminetetraacetic acid, along with a protease inhibitor cocktail (Roche). The cell extracts were boiled at 90◦C for 10 min followed by sonication. The cell debris was removed by centrifugation at 13400 rpm for 30 min and the crude protein extracts were cleaned up by chloroform/methanol precipitation. Dried protein pellets were dissolved in denaturation buffer containing 8 M urea in 10 mM Tris-HCl pH 8.0. The protein lysate was reduced with 1 mM dithiothreitol and alkylated with 5.5 mM iodoacetamide in the dark, for 1 h each at room temperature. Proteins were then subjected to overnight digestion with an endoproteinase Trypsin (1:100, w/w; PierceTM). The reaction was stopped by acidification with 10% trifluoroacetic acid and stage-tipped before injecting the samples into the mass spectrometer (Ishihama et al., 2006). Samples were analyzed on an Q Exactive mass spectrometer coupled to an Easy-nLC 1200 (both Thermo Fisher Scientific, Inc., Waltham, MA, United States). Chromatographic separation was performed using an in-house constructed pre-column (45 mm × 0.075 mm I.D) and analytical (200 mm × 0.075 mm I.D.) column set up packed with 3 µm Reprosil-Pur C18-AQ particles (Dr. Maisch GmbH, Ammerbuch, Germany). Peptides were injected onto the column with solvent A (0.2% formic acid in water) at a flow rate of 300 nL/min and 500 bars. Peptides were then eluted using a segmented gradient of 7–27% B-solvent (80% acetonitrile with 0.2% formic acid) over 45 min, 27–40% B over 5 min, 40–100% B over 5 min with a final hold at 100% B for 10 min. The mass spectrometer was operated on a data-dependent mode. Survey full-scans for the MS spectra were recorded between 400 and 1600 Thompson at a resolution of 70,000 with a target value of 1e6 charges in the Orbitrap mass analyzer. The top 10 most intense peaks from the survey scans of doubly or multiply charged precursor ions were selected for fragmentation with higher-energy collisional dissociation (HCD) with a target value of 1e5 in the Orbitrap mass analyzer in each scan cycle. Dynamic exclusion was set for 30 s. Triplicate injections (technical replicates) were carried out for each of the samples for label free quantitation (LFQ).

# Data Processing and Analysis

Acquired MS spectra were processed with the MaxQuant software suite (version <IP_ADDRESS>) (Cox et al., 2009), integrated with an Andromeda search engine. Database search was performed against a target-decoy database of B. subtilis 168 downloaded from UniProt (taxonomy ID 1423), containing 4,195 protein entries, and additionally including also 248 commonly observed laboratory contaminant proteins. Endoprotease Trypsin/P was set as the protease with a maximum missed cleavage of two. Carbamidomethylation (Cys) was set as a fixed modification. Label free quantification was enabled with a minimum ratio count of two. A false discovery rate of 1% was applied at the peptide and protein level individually for filtering identifications. Initial mass tolerance was set to 20 ppm. In case of the main search, the peptide mass tolerance of precursor and the fragment ions were set to 4.5 and 20 ppm, respectively. Downstream bioinformatics analysis was performed using Perseus version <IP_ADDRESS> (Tyanova et al., 2016). Grouping of proteins with similar expression profiles was achieved by hierarchical clustering analysis. Log10 transformation of mean LFQ intensities of proteins was performed for all the tested conditions. Missing values were replaced from the normal distribution via imputation. Hierarchical clustering was performed on Z-score transformed values using Euclidean as a distance measure and Average linkage cluster analysis. Significance B (p ≤ 0.05) was calculated to identify significantly regulated proteins in each of the ascorbate treatment conditions relative to the control.

# Reconstruction of Context-Specific Models Using Proteomics Data

Log10 LFQ protein intensities were used to generate sodium ascorbate concentration specific models by mapping the protein intensities to the genome-scale metabolic model of B. subtilis

(Bs-iYO844) (Oh et al., 2007). Log10 LFQ protein intensities from biological replicates (three replicates for each sodium ascorbate concentration and 2 for the control) were averaged and used to constrain the fluxes in the associated reaction using Gene Inactivity Moderated by Metabolism and Expression (GIMME) (Becker and Palsson, 2008). GIMME was run using 90% of the objective function threshold and 50th percentile of proteins expression level. Since properly constrained reactions do not demonstrate uniform distributions of feasible steadystate fluxes, the range and distribution of feasible metabolic flux for each reaction were determined by using Markov Chain Monte Carlo (MCMC) sampling (Lewis et al., 2012). To do this, a large number of feasible sets of metabolic fluxes were randomly moved within the solution space until they were well mixed, thereby sampling the entire solution space (Lewis et al., 2012). This sampling process yielded a distribution of feasible steady-state fluxes for each reaction. Sampling was done using gpSampler with default settings from the COBRA toolbox (Schellenberger et al., 2011) using Gurobi (Gurobi Optimization, Inc., Houston, TX, United States) and MATLAB R (The MathWorks Inc., Natick, MA, United States). Averaged sampled predicted flux distributions for each reaction at each sodium ascorbate concentration were compared to those from the control model in order to identify reactions (and their associated genes) that have significantly altered flux rates upon adding sodium ascorbate to B. subtilis.

# Statistical Analysis

The data are presented as the mean ± standard deviation. Intergroup differences were estimated by one-way analysis of variance (ANOVA), followed by a post hoc multiple comparison (Tukey) test to compare the multiple means. Differences between values were considered to be statistically significant when the P-value was <0.05.

# RESULTS

# Vitamin C Does Not Affect Bacterial Growth in a Liquid Medium, but Inhibits Biofilm Formation

Five–ten millimeter doses of vitamin C were previously reported to completely exterminate mycobacteria (Vilcheze et al., 2013). By contrast, it has been reported that vitamin C is not bactericidal toward opportunistic pathogens such as E. coli and P. aeruginosa, but it renders them more susceptible to antibiotics and some physical treatments (Kallio et al., 2012; Khameneh et al., 2016; Helgadóttir et al., 2017). To clarify this effect of vitamin C on non-mycobacterial species, we used the opportunistic pathogens E. coli and P. aeruginosa. Since we previously hypothesized that the synergistic effects of vitamin C may be related to biofilms (Helgadóttir et al., 2017), we included also B. subtilis, the model organism for biofilm development (Vlamakis et al., 2013). We exposed these bacterial strains to a concentration range of vitamin C from 10 to 40 mM (sodium ascorbate, neutral pH form), assessing both the survival and growth in liquid media and biofilms. Up to 40 mM vitamin C did not significantly inhibit the planktonic growth of B. subtilis, E. coli, or P. aeruginosa (**Supplementary Figure S1**). However, in the same concentration range, biofilm formation was impaired for all three species (**Figure 1** and **Supplementary Figure S2**). Since the vitamin C effect was most pronounced on biofilms, we focused on B. subtilis for an in-depth study of the mechanism behind this effect. B. subtilis biofilm is the most robust and easiest to analyze in terms of structure and composition, and the mechanisms leading to its formation are well characterized (Vlamakis et al., 2013). The normal B. subtilis pellicle (control, 0 mM) exhibits wrinkled and folded architecture (**Figure 1C**). Vitamin C treatment abolished this wrinkled architecture and visibly attenuated the pellicle in a concentration-dependent manner (**Figure 1C**). This effect was accompanied by a linear decrease in biofilm biomass (**Figure 1A**). However, the viability of B. subtilis in the pellicle was not significantly affected, suggesting that the reduced biofilm biomass could be due to loss of EPS and not the loss of cells.

# Low Concentrations of Vitamin C Do Not Kill B. subtilis but Deplete the EPS Matrix

Next, we examined in details the content of various biofilm components in response to the same concentration range of vitamin C. All three major components of the biofilm matrix, polysaccharides, proteins and eDNA, were reduced in the presence of vitamin C (**Figures 2A–C**). The reduction in protein content vs. vitamin C concentration showed a linear regression coefficient of only 0.88, but the trend of exopolysaccharide and DNA reduction was more closely correlated to increasing concentration of vitamin C and the loss of pellicle biomass (**Supplementary Figure S3**). The polysaccharide content and bacterial bio-volume in the B. subtilis biofilm were examined by laser scanning confocal microscopy, in the same concentration range of vitamin C (**Figures 2E–G**). EPS account for over 40% of the mass of the B. subtilis biofilm, but their identification and characterization is not complete. Roux et al. (2015) identified that poly n-acetylglucosamine is the major polysaccharide component of the B. subtilis biofilm matrix (Roux et al., 2015). We therefore used fluorescence probes to visualize EPS components: Alexa flour, WGA conjugate, for n-acetylglucosamine and ConA, Tetramethylrhodamine conjugate, as an unspecific EPS binder for proteins and other polysaccharides. We first established that these probes had no effect on biofilm formation and exhibited no toxicity to B. subtilis cells (**Supplementary Figure S4**). The n-acetylglucosamine (stained in red) occupied a significant part of the biovolume in the untreated sample (**Figure 2G**). With increasing concentrations of vitamin C, the overall thickness of the biofilm decreased, and the content of the poly n-acetylglucosamine (red) and the unspecific EPS matrix (blue) decreased as well (**Figure 2G**). The decreasing pattern of NAG with vitamin C treatment was consistent with the total polysaccharides content observed by colorimetric assay, where significant inhibition was observed with ≥20 mM of concentration. Meanwhile, the bacterial bio-volume remained constant up to 30 mM vitamin C. The decreasing pattern of bacterial bio-volume was

not consistent with other results because vitamin C treatment is mainly affecting the ECM production but not the growth and viability of bacteria at lower concentrations as shown in **Figure 1**. Viability of cells was examined by live/dead staining (**Figure 3A**) where it was observed that, at low concentration of vitamin C (10 mM) the cell survival rate was similar to that of untreated control biofilms. By contrast, a significant number of cells were dead in biofilms treated with a higher concentration of vitamin C (40 mM). The killing of cells at the higher concentration of vitamin C coincided with higher levels of detectable oxidative stress (**Figure 3B**). This confirmed that vitamin C effect on biofilms takes place in two stages: at concentrations of up to 30 mM the cell viability is preserved, but there is a loss of the EPS, primarily exopolysaccharides. Above 30 mM vitamin C, the bacterial cells start dying.

# Stage 1: Low Concentrations of Vitamin C Inhibit Exopolysaccharide Synthesis, Stage 2: High Concentrations of Vitamin C Induce Lethal Oxidative Stress

Since vitamin C seemed to inhibit B. subtilis biofilm formation in two stages: reduction of EPS components at up to 30 mM (**Figure 2**), and killing of cells at 30 mM and higher by inducing the oxidative stress (**Figures 1B**, **3**), we performed an in-depth label-free quantitative proteome analysis of vitamin-C treated biofilms in this critical concentration range. A total of 2056 B. subtilis proteins were identified, of which 1373 were quantified (**Supplementary Table S1**). Three biological replicates showed a high degree of correlation (Pearson correlation coefficient ≥ 0.9) (**Supplementary Figure S5**). Hierarchical clustering analysis was employed for grouping similar expression profiles of proteins (**Figures 4A–C**). Differentially regulated proteins grouped in four clusters (**Figure 4B**). The majority grouped in clusters 1 (expression reduced upon the addition of vitamin C) and 4 (expression enhanced upon the addition of vitamin C). Clusters 2 and 3 showed variation across the range of vitamin C concentrations. All proteins falling in the four different clusters are listed in the **Supplementary Table S1**. Proteins for which the expression levels were most strongly affected by vitamin C treatment (p ≤ 0.05) were identified by plotting log2 transformed label-free quantification (LFQ) ratios against log10 transformed LFQ intensities (**Figure 4C**). Around 100 proteins were found in this category. Among these, at lower vitamin C concentrations, many proteins directly involved in exopolysaccharides synthesis, export and biofilm formation were depleted: notably PtkA, SlrR, SpeA, EpsC, EpsD, EpsE, EpsH, EpsI, EpsO, TuaD, Ugd (Kobayashi, 2008; Gerwig et al., 2014; Mijakovic and Deutscher, 2015). This correlated with disrupted expression of the key regulators controlling synthesis or activity of these proteins, such as ComA, RepC, Spo0A, YmdB, KinC, FloT, and PtkA (Lazazzera et al., 1999; Diethmaier et al., 2011; Yepes et al., 2012). By contrast, at higher vitamin C concentrations, oxidative stress associated proteins (Antelmann et al., 2000; Grimaud et al., 2001; Towe et al., 2007) were strongly overexpressed: MsrA, MsrB, MhqA, MhqD, AzoR2, and RocA. Individual roles of these proteins in biofilm production and oxidative stress response are reviewed in detail in the discussion section. It was evident from this dataset that vitamin C treatment provoked a two-stage global

FIGURE 2 | Biochemical and confocal laser scanning microscopy analysis of B. subtilis biofilm grown in the presence of vitamin C. (A) Content of polysaccharide, (B) content of protein and (C) eDNA concentration in a 24 h old biofilm. (D) Bacterial biovolume, (E) biovolume of poly n-acetylglucosamine and (F) biovolume of unspecific EPS matrix (proteins and polysaccharides) in a 24 h biofilm. (G) Representative 3-D architecture of a 24 h old B. subtilis biofilm grown in the presence of vitamin C (green: bacteria; red: n-acetylglucosamine; blue: unspecific EPS matrix). All data (A–F) are mean values ± standard deviation from three biological replicates. Values marked by the same superscripts in panels (A–F) are not significantly different from each other (P > 0.05).

rearrangement of the cellular proteome, which correlated well to our previous observations on EPS content and cell viability. At low concentrations of vitamin C, biosynthetic pathways leading to exopolysaccharide synthesis and export were down-regulated, explaining the observed depletion of biofilm EPS. At higher vitamin C concentrations, the cell started expressing proteins to cope with excessive oxidative stress, which correlates to cell death and loss of the bacterial bio-volume in the biofilm. To assess the specific impact of this global proteome adaptation on redistribution of metabolic fluxes, we used the quantitative proteomics data to generate vitamin C concentration-specific genome-scale metabolic models, by mapping the protein intensities to the available B. subtilis model Bs-iYO844 (Oh et al., 2007) (**Supplementary Figure S6**). This enabled us to identify several metabolic pathways with flux redistribution provoked by vitamin C, which corroborate our findings (**Figure 5**). Notably, the model guided analysis showed that pyrroline-5-carboxylate dehydrogenase (P5CDH)/RocA had a significantly upregulated flux in the presence of vitamin C (**Figure 5**), which indicates that the cells are trying to neutralize ROS.

# DISCUSSION