Antifungal Azoles as Tetracycline Resistance Modifiers in Staphylococcus aureus

ABSTRACT

Staphylococcus aureus has developed resistance to antimicrobials since their first use. The S. aureus major facilitator superfamily (MFS) efflux pump Tet(K) contributes to resistance to tetracyclines. The efflux pump diminishes antibiotic accumulation, and biofilm hampers the diffusion of antibiotics. None of the currently known compounds have been approved as efflux pump inhibitors (EPIs) for clinical use. In the current study, we screened clinically approved drugs for possible Tet(K) efflux pump inhibition. By performing in silico docking followed by in vitro checkerboard assays, we identified five azoles (the fungal ergosterol synthesis inhibitors) showing putative EPI-like potential with a fractional inhibitory concentration index of ≤0.5, indicating synergism. The functionality of the azoles was confirmed using ethidium bromide (EtBr) accumulation and efflux inhibition assays. In time-kill kinetics, the combination treatment with butoconazole engendered a marked increase in the bactericidal capacity of tetracycline. When assessing the off-target effects of the azoles, we observed no disruption of bacterial membrane permeability and polarization. Finally, the combination of azoles with tetracycline led to a significant eradication of preformed mature biofilms. This study demonstrates that azoles can be repurposed as putative Tet(K) EPIs and to reduce biofilm formation at clinically relevant concentrations.

IMPORTANCE Staphylococcus aureus uses efflux pumps to transport antibiotics out of the cell and thus increases the dosage at which it endures antibiotics. Also, efflux pumps play a role in biofilm formation by the excretion of extracellular matrix molecules. One way to combat these pathogens may be to reduce the activity of efflux pumps and thereby increase pathogen sensitivity to existing antibiotics. We describe the in silico-based screen of clinically approved drugs that identified antifungal azoles inhibiting Tet(K), a pump that belongs to the major facilitator superfamily, and showed that these compounds bind to and block the activity of the Tet(K) pump. Azoles enhanced the susceptibility of tetracycline against S. aureus and its methicillin-resistant strains. The combination of azoles with tetracycline led to a significant reduction in preformed biofilms. Repurposing approved drugs may help solve the classical toxicity issues related to efflux pump inhibitors.

INTRODUCTION

Antibiotic-resistant superbugs are no longer only a laboratory concern but have become a high-end global health concern. Staphylococcus aureus is a commensal bacterium as well as a predominant human pathogen. S. aureus is the most conspicuous wound pathogen, causing both acute and chronic skin and soft tissue infections (SSTI) upon the formation of robust biofilms (1). According to a study, the estimated number of emergency department visits for SSTI in the United States increased from 1.2 million in 1993 to 3.4 million in 2005 (2). The epidemiology of S. aureus infections worsened by virtue of two predominant shifts—first, an increase in the number of health care-associated infections, and second, an epidemic of community-associated SSTI driven by the manifestation of multiple resistance determinants in bacteria (3).

Out of various determinants of resistance, efflux pumps are at the forefront to endorse the resistance phenotype and constitute between 6% and 18% of transporters present in bacterial species (4, 5). Efflux pumps are also known to play a vital role in biofilm formation by extruding quorum sensing signals and molecules constructing extracellular matrix and helping with bacterial adhesion and pathogenesis (6–8). Furthermore, several previously known efflux pump inhibitors (EPIs) are evidenced to decrease biofilm formation (9–11). Considering the prevalence of tetracycline resistance in methicillin-resistant S. aureus (MRSA), to date, 29 different tetracycline resistance genes have been characterized, of which 18 code for efflux pumps (12). The efflux proteins of the major facilitator superfamily (MFS) that largely contribute to tetracycline resistance in both Gram-positive and Gram-negative bacteria are from Tet family, more particularly Tet(K) in Gram-positive bacteria. The Gram-positive genes tet(K), tetL, and tetA(P) and the Gram-negative genes tetA, tetC, tetD, tetE, tetG, and tetH encode efflux proteins which confer resistance to tetracycline but not minocycline and tigecycline (13, 14). In a study, it was found that approximately 90% of MRSA, 70% of multidrug-resistant (MDR) Enterococcus faecalis, and 60% of MDR Streptococcus pneumoniae are tetracycline resistant (15). Tet proteins are 46-kDa membrane-bound efflux proteins with 12 (Gram-negative) or 14 (Gram-positive) hydrophobic membrane-spanning regions separated by a short central hydrophobic region of amino acids. What is noteworthy is that resistance determinants of tet are primarily present on small transmissible plasmids, which are occasionally integrated into the chromosomes of staphylococci and thereby promote acquired resistance in bacteria (13).

The rate of spread of resistance is expeditious like a furious storm; having said that, there was a drastic decline in the number of antibiotics being discovered and approved—from 29 in the 1980s and 23 in the 1990s to only 9 antibiotics in the last decade (16). Considering the slow discovery rate of new antibiotic solutions, the only source of assistance is to contemplate restoring the efficacy of previously approved antibiotics.

EPIs work as adjuvants along with the “old” antibiotics and help in the reversal of intrinsic and acquired resistance in bacteria. Many EPIs have been discovered (17–19), but none of the antibiotic-EPI duos is approved clinically due to their unnecessary toxicities. One of the strategies is to explore already approved drugs as adjuvants with the known antibiotics; repurposing these drugs can be highly beneficial, as their pharmacokinetics and pharmacodynamics profiles are already known. Developing inhibitors of the Tet(K) efflux pump is a promising strategy with which to reverse the multidrug resistance and break the back of the biofilm-forming potential of bacteria.

To find efficient EPIs, we screened a Food and Drug Administration (FDA)-approved library of ∼2,200 drugs (www.drugbank.ca) against the Tet(K) efflux pump protein by performing in silico docking studies. We shortlisted 200 best hits based on the criterion of high docking score, performed in vitro screening assays, and discovered a series of azoles as EPIs. Azoles are known antifungal agents that act by inhibition of the ergosterol biosynthesis pathway (20). Here, we report the novel use of already approved azoles as EPIs combined with tetracycline as a solution to combat S. aureus.

RESULTS

Virtual screening and molecular docking to the active site of Tet(K).

The FDA-approved compound library was docked against the entire surface of the Tet(K) homology model. The SiteMap analysis suggests five potential druggable sites for ligand binding, with site scores between 0.56 and 0.96. The binding site with a site score of 0.96 corresponding to the central cavity of Tet(K) efflux pump was selected for grid generation. The molecular docking was performed using the Glide Xtra-Precision (XP) module as described in the Materials and Methods section. The docked structures were further used for binding free-energy calculations using the molecular mechanics energies combined with the generalized born and surface area continuum solvation (MM-GBSA) module. Molecular docking of about 2,200 FDA-approved drugs at the central active site binding cleft resulted in many potential hits. Many azoles were among the top hits showing high docking scores and favorable binding energies. The hydrophobic, hydrophilic, and pi-pi interactions are the major interactions that stabilize the binding of azoles with Tet(K) aromatic amino acid residues (Table 1). Among azoles, butoconazole showed the highest docking score of −8.066 and MM-GBSA-based binding energy score of −53.98, while clotrimazole exhibited the lowest docking score of −4.465 and MM-GBSA-based binding energy score of −25.33. Clotrimazole and tetracycline (see Fig. S1 in the supplemental material) docked at a different binding site than the other four azoles. Both bind deeper inside the binding cleft of the Tet(K) pump. The high docking and MM-GBSA scores suggest that these drugs may have favorable interactions and may inhibit Tet(K) activity. The docking pose and the ligand-protein interactions for all five hits are shown in Fig. 1.

TABLE 1.

Binding energy of the lead compounds to the homology model of Tet(K) efflux pump

FIG 1.

Interaction of butoconazole (A), econazole (B), miconazole (C), tioconazole (D), and clotrimazole with the active site of Tet(K) (E). Cartoon (ball and stick) representation of Tet(K) showing azoles docked in the active site cleft (left). 3D electrostatic potential map (middle). Ligand interaction diagram in 2D representation showing several interactions involved in Tet(K)-azole binding (right).

Azoles do not interfere with the primitive growth kinetics of bacteria.

Bacterial growth kinetics is an autocatalytic reaction, which entails that the growth rate is directly proportional to the concentration of cells. The MICs of clotrimazole, tioconazole, miconazole, econazole, and butoconazole were 100 μM, 100 μM, 50 μM, 100 μM, and 50 μM, respectively, against the S. aureus XU212 strain. Ideally, as the EPIs were used at subinhibitory concentrations, it should not interfere with the growth kinetics of the bacteria. The growth kinetics of S. aureus XU212 was assessed in the presence of azoles, and as expected, none of the compounds caused any alteration in the growth pattern of bacteria. They did not exhibit any difference from the control group. This finding indicates that in a nutrient-rich environment, the subinhibitory concentrations of azoles may not affect S. aureus growth (Fig. 2A).

FIG 2.

(A) Growth kinetics of Staphylococcus aureus XU212 in CA-MHB under subinhibitory concentrations (1/4× MIC) of the azoles. OD₆₀₀ values are the means of three independent experiments. The control (untreated) set was included to monitor normal bacterial growth. (B) Tetracycline-azole synergy test using agar-well diffusion assay on S. aureus XU212. The agar wells containing azoles individually did not produce any zone of inhibition; however, tetracycline alone displayed a zone of inhibition of only 9 mm. The combination of tetracycline with all the azoles was bactericidal, as indicated by a significant increment in the zone of growth inhibition. The images are representatives of two independent experiments. Abbreviations: TET, Tetracycline; CLO, clotrimazole; TIO, tioconazole; MIC, miconazole; ECO, econazole; BUT, butoconazole. Shown are effects of subinhibitory concentrations (1/4× MIC) of clotrimazole (C), tioconazole (D), miconazole (E), econazole (F), and butoconazole (G) on the real-time accumulation of EtBr in a concentration-dependent manner; the positive-control reserpine (H) was included for comparison. The results presented correspond to the average of three independent assays ± SD.

Azoles potentiate the antimicrobial activity of tetracycline against tetracycline-resistant S. aureus.

To evaluate the possible synergistic activity, we performed a checkerboard synergy assay of a series of azoles in combination with tetracycline on S. aureus XU212 possessing the Tet(K) efflux pump. The MIC of azoles and tetracycline were determined on various strains (Tables 2 and 3; see Table S2 in the supplemental material), and the reduced concentration of tetracycline in the presence of azoles needed to inhibit bacterial growth was determined. The results indicate that all the azoles at subinhibitory concentrations (1/4× MIC), i.e., clotrimazole (25 μM), tioconazole (25 μM), miconazole (12.5 μM), econazole (25 μM), and butoconazole (12.5 μM), were able to modulate the MIC of tetracycline by 32-, 32-, 64-, 32-, and 64-fold, respectively (Table 2), suggesting butoconazole and miconazole are the most effective Tet(K) inhibitors/EPI. Moreover, the fractional inhibitory concentration index (FICI) of all EPIs was ≤0.5, which indicates “synergy.” The fold reduction in MIC of tetracycline in the presence of azoles was more significant than that of reserpine and chlorpromazine (both display 4- and 2-fold reduction, respectively). The concentration-dependent potentiation effect was observed with all the azoles. For the validation of synergism due to efflux pump inhibition, we also performed the assay on the wild-type strain of S. aureus ATCC 29213 [tetracycline-susceptible S. aureus strain with a basal level expression of Tet(K) efflux pump]. None of the azoles demonstrated any significant modulation in the MIC of tetracycline, indicating the Tet(K) efflux pump as the putative target for azoles. Moreover, we determined the synergistic activity of the combination against tetracycline-resistant clinical strains of S. aureus. The combination of azoles with tetracycline modulated the MIC of tetracycline by 8- to 64-fold, thereby increasing the susceptibility of bacteria to the tetracycline (Table 3). We also performed a checkerboard synergy assay of azoles in combination with a panel of antibiotics belonging to different classes on S. aureus strains expressing three different efflux pumps, i.e., Tet(K) (XU212), NorA (SA1199B), and MsrA (RN4220). Besides the previously observed effect on tetracycline susceptibility (Table 3), no further synergy was observed, with antibiotics belonging to other structural families when tested against the Tet(K)-expressing strain. Moreover, no synergism was observed in strains expressing either NorA or MsrA, when the azoles were combined with antibiotics that are substrates of these efflux pumps. Finally, it is worth mentioning that azoles reduced ethidium bromide (EtBr; a common, nonspecific substrate of efflux pumps) MIC in the strain expressing Tet(K) and not in the ones expressing either NorA or MsrA (see Table S3 to S5 in the supplemental material). Altogether, these results suggest that the target of azoles is Tet(K) and neither NorA nor MsrA are inhibited by these compounds.

TABLE 2.

MIC of tetracycline for S. aureus XU212^a in combination with different azoles

Compound	MIC	Concn of azoles in μM (μg/ml)	MIC of tetracycline in presence of azoles (μg/ml)	Fold reduction in tetracycline MIC	FICI
Tetracycline	256 μg/ml
Clotrimazole	100 μM	25 (8.6)	8	32	0.281
		12.5 (4.3)	8	32	0.156
		6.25 (2.15)	32	8	0.188
		3.125 (1.07)	128	2	0.531
		1.563 (0.54)	256	1
Tioconazole	100 μM	25 (9.6)	8	32	0.281
		12.5 (4.8)	16	16	0.188
		6.25 (2.4)	32	8	0.188
		3.125 (1.2)	32	8	0.156
		1.563 (0.6)	128	2	0.516
Miconazole	50 μM	12.5 (5.98)	4	64	0.266
		6.25 (2.99)	8	32	0.156
		3.125 (1.49)	16	16	0.125
		1.563 (0.75)	32	8	0.156
		0.7812 (0.37)	64	4	0.266
Econazole	100 μM	25 (11.11)	8	32	0.281
		12.5 (5.55)	16	16	0.188
		6.25 (2.77)	32	8	0.188
		3.125 (1.38)	128	2	0.531
		1.563 (0.69)	256	1
Butoconazole	50 μM	12.5 (5.93)	4	64	0.266
		6.25 (2.96)	4	64	0.141
		3.125 (1.48)	8	32	0.094
		1.563 (0.74)	16	16	0.094
		0.781 (0.37)	64	4	0.266
Reserpine	>210 μM	25 μM (15.21)	64	4	0.5
Chlorpromazine	180 μM	32 μM (11.37)	128	2	0.677

^aTet(K) positive.

TABLE 3.

MIC of tetracycline for S. aureus strains in combination with different azoles

Compound by S. aureus strain	MIC of azoles (μM)	Concn of azoles (μM)	MIC of tetracycline (μg/ml)	Fold reduction in tetracycline MIC	FICI
MRSA1 (clinical isolate)
Tetracycline			32
+Clotrimazole	12.5	3.125	16	2	0.75
+Tioconazole	12.5	3.125	8	4	0.5
+Miconazole	3.125	0.78	8	4	0.5
+Econazole	6.25	1.56	16	2	0.75
+Butoconazole	6.25	1.56	4	8	0.375
ATCC 33591 (MRSA)
Tetracycline			256
+Clotrimazole	12.5	3.125	32	8	0.375
+Tioconazole	12.5	3.125	16	16	0.3125
+Miconazole	3.125	0.78	16	16	0.3125
+Econazole	6.25	1.56	64	4	0.5
+Butoconazole	3.125	0.78	8	32	0.281
GMCH 6188 (clinical isolate)
Tetracycline			256
+Clotrimazole	12.5	3.125	32	8	0.375
+Tioconazole	6.25	1.56	16	16	0.3125
+Miconazole	3.125	0.78	16	16	0.3125
+Econazole	6.25	1.56	16	16	0.3125
+Butoconazole	6.25	1.56	4	64	0.265
ATCC BAA-39 (MDR)
Tetracycline			256
+Clotrimazole	12.5	3.125	32	8	0.375
+Tioconazole	12.5	3.125	32	8	0.375
+Miconazole	3.125	0.78	16	16	0.3125
+Econazole	6.25	1.56	32	8	0.375
+Butoconazole	6.25	1.56	8	32	0.281
ATCC 29213 (wild-type quality-control strain)
Tetracycline			0.5
+Clotrimazole	25	6.25	0.25	2	0.75
+Tioconazole	12.5	3.125	0.25	2	0.75
+Miconazole	6.25	1.56	0.125	4	0.5
+Econazole	6.25	1.56	0.25	2	0.75
+Butoconazole	3.125	0.78	0.125	4	0.5

Furthermore, we compared the effect of azoles upon their addition to tetracycline in a qualitative agar-well diffusion assay. As expected, there was no zone of growth inhibition for the azoles (clotrimazole, tioconazole, miconazole, econazole, and butoconazole) alone at subinhibitory concentrations (1/4× MIC). At the same time, tetracycline alone showed a zone of inhibition having diameter of ∼9 mm. In the combination, zone diameter increased significantly up to ∼19 to 21 mm, indicating the potentiation of the bactericidal activity of tetracycline in the presence of azoles (Fig. 2B).

Azoles inhibited EtBr efflux.

The checkerboard synergy assay results are an indirect measure of the potential efflux activity of the strains and the effect of azoles on this activity. To validate the efflux inhibitory activity of the azoles, we performed the real-time EtBr (which is a DNA binding dye and a broad efflux pump substrate) (21) accumulation assay on S. aureus XU212. The relative final fluorescence (RFF) values were calculated for each compound. The RFF value is a measure of the compound’s effectiveness in the inhibition of the EtBr efflux by comparing the final fluorescence of the treated versus untreated cells (22). An RFF of >1 indicates an enhanced accumulation of EtBr. From the results, it is clear that all the azoles, i.e., clotrimazole (25 μM), tioconazole (25 μM), miconazole (12.5 μM), econazole (25 μM), and butoconazole (12.5 μM), caused significantly increased accumulation of EtBr, with RFF values of 5.67, 8.864, 9.085, 5.956, and 11.403, respectively, compared with reserpine (RFF, 3.892) (Table 4). A concentration-dependent EtBr accumulation effect was observed for azoles (Fig. 2C to H). The increase in accumulation of dye indicates the partial or complete cessation of extrusion of dye through inhibition of the Tet(K) efflux pump.

TABLE 4.

RFF values based on the accumulation of EtBr for the S. aureus XU212 in the presence of the azoles

Compound	Concn used (μM)	Relative final fluorescencea
Clotrimazole	25	5.670 ± 0.067
	12.5	3.576 ± 0.405
	6.25	2.663 ± 0.243
Tioconazole	25	8.864 ± 0.225
	12.5	6.609 ± 0.268
	6.25	4.687 ± 0.353
Miconazole	12.5	9.085 ± 0.310
	6.25	5.987 ± 0.373
	3.125	3.796 ± 0.342
Econazole	25	5.956 ± 0.168
	12.5	3.389 ± 0.095
	6.25	2.30 ± 0.010
Butoconazole	12.5	11.403 ± 0.203
	6.25	11.362 ± 0.248
	3.125	8.859 ± 0.237
Reserpine	25	3.892 ± 0.455

^aThe results correspond to the average of three independent experiments ± SD.

Additionally, the ability of azoles to directly inhibit the efflux of EtBr from S. aureus XU212 was evaluated by using the fluorescence assay. The presence of the Tet(K) efflux pump resulted in a rapid decrease in fluorescence, indicating the extrusion of EtBr by the pump. The addition of azoles (clotrimazole, tioconazole, miconazole, econazole, and butoconazole) at subinhibitory concentrations (1/4× MIC) significantly slowed the extrusion of dye compared with that of the control in S. aureus XU212. The presence of glucose reduced the efflux pump inhibitory activity of compounds because glucose reenergizes the cells, promoting active efflux (23). The results indicate the potent efflux pump inhibitory activity of the azoles (Fig. 3A to F).

FIG 3.

Effect of subinhibitory concentrations (1/4× MIC) of clotrimazole (A), tioconazole (B), miconazole (C), econazole (D), and butoconazole (E) on the efflux inhibition of EtBr in the presence and absence of glucose; the positive-control reserpine (F) was included for comparison. The results presented correspond to the average of three independent assays ± SD.

Azoles inhibit tetracycline efflux from the bacterial cell.

For the more realistic validation of the Tet(K) efflux pump inhibition, we performed a tetracycline efflux inhibition assay in the presence of azoles on S. aureus XU212 and S. aureus ATCC 29213. The uptake of tetracycline can be easily monitored by fluorescence measurement when it enters the cell (24). In this assay, reserpine was included as a positive control because of its known efflux pump inhibitory activity (18, 25). The results clearly depict a significant decrease in the extrusion of tetracycline in the presence of azoles (clotrimazole, tioconazole, miconazole, econazole, and butoconazole) compared with that of reserpine and cells without any treatment (Fig. 4A to F). Although, the presence of glucose reduced the efflux pump inhibitory activity of azoles because glucose reenergizes the cells, promoting active tetracycline efflux.

FIG 4.

(A to F) Efflux inhibition of tetracycline in S. aureus XU212, when the cells were treated with clotrimazole, tioconazole, miconazole, econazole, butoconazole, and reserpine at subinhibitory concentrations (1/4× MIC). The change in fluorescence was monitored in the presence and absence of glucose (4%). The tetracycline fluorescence was recorded at 535 nm by exciting at a wavelength of 405 nm. The results presented correspond to the average of two independent experiments with three repeats ± SD.

Effect of butoconazole on time-kill kinetics of tetracycline.

The time-dependent killing of S. aureus XU212 was examined for 24 hours to assess the bactericidal effect of the combination of tetracycline with butoconazole (best hit). Tetracycline was used at 1× MIC (256 μg/ml) alone as well as in combination with butoconazole at 1/4× MIC (12.5 μM). As expected, butoconazole alone at 12.5 μM did not show any bactericidal activity. Moderate killing was observed for tetracycline alone (256 μg/ml) in 8 h; however, significant regrowth was observed after 24 h. A drastic enhancement in the bactericidal activity of the combination of tetracycline with butoconazole was observed in 8 h compared with tetracycline alone. Even after regrowth, the combination maintained the difference of 2.5 to 3 log₁₀ CFUs below the initial log₁₀ CFUs at 0 h (Fig. 5A). Henceforth, the combination of tetracycline and butoconazole could be a potential anti-infective therapy.

FIG 5.

(A) Time-kill curves of S. aureus XU212 showing the bactericidal effect of tetracycline (256 μg/ml) in combination with butoconazole (12.5 μM). Each time point represents the mean log₁₀ CFU/ml ± SD of three readings. (B) Shows the postantibiotic effect induced by tetracycline (1× MIC) alone and in combination with butoconazole (1/4× MIC) for 2 h in S. aureus XU212. Time 0 h corresponds to the beginning of growth monitoring immediately after compound removal and cell resuspension in fresh CA-MHB. The value of PAE corresponds to the delay undergone by the treated culture with respect to the untreated control in reaching an OD value of one-half of the final OD. The assays were repeated three times independently, and results were presented as mean ± SD. (C) Bacterial membrane permeability is measured with fluorescent PI dye upon exposure to the azoles at 1× MIC. Increased fluorescence corresponds to cells with the permeabilized membranes. Paenibacillin (10 μM) was used as a positive control since it destabilizes the Gram-positive bacterial membrane. The drug-free control was included to monitor change in fluorescence. (D) The azoles do not depolarize the bacterial membrane. Shown is the change in fluorescence of S. aureus XU212 cells using a DiSC3(5) assay. Data are presented as a change in fluorescence before and after the addition of the azoles and valinomycin at 1× MIC. (E) S. aureus XU212 was exposed to the azoles at 1/4× MIC during 4 h. The ATP levels were quantified using a luciferin-luciferase bioluminescence detection assay. CCCP and valinomycin were included for comparison. The results presented correspond to the mean of three independent assays ± SD. Results were considered highly significant when the P value was <0.001 (***).

Butoconazole prolonged the postantibiotic effect (PAE) of tetracycline.

According to a study (26), an antimicrobial agent causes PAE when, immediately after its removal, it brings about a growth delay of at least 0.5 h on a bacterial culture. A log-phase culture of S. aureus XU212 was exposed to 1× MIC of tetracycline in the presence and absence of butoconazole (12.5 μM) for 2 h. The PAE of tetracycline alone was found to be 2.15 h, while the combination with butoconazole further increased the PAE of tetracycline by 2.30 h. The enhanced PAE of the combination will help in the sustenance of the widely spaced dosing intervals and determine the optimum dosing frequency of the drugs (Fig. 5B).

Azoles do not function by altering membrane permeability.

To eliminate the possibility of synergy due to an off-target effect, the outer membrane permeabilization using fluorescent propidium iodide (PI) dye on S. aureus XU212 was determined. PI is a membrane-impermeable fluorescent stain, and it binds double-stranded nucleic acids. The increase in fluorescence indicates altered membrane permeability (27). For all the azoles (clotrimazole, tioconazole, miconazole, econazole, and butoconazole), no significant increase in fluorescence was observed, revealing no membrane permeabilization, while paenibacillin (positive control) (28) demonstrated a drastic increase in fluorescence compared with that of the control (without treatment) (Fig. 5C).

Azoles cause nix membrane depolarization.

We used 3, 3′-dipropylthiadicarbocyanine iodide [DiSC₃(5)], a fluorescent molecular probe for the membrane depolarization assay, against S. aureus XU212. The fluorescence of DiSC₃(5) decreases as the dye partitions to the surface of polarized cells; depolarization prevents partitioning and can release bound dye into the media (29). All azoles (clotrimazole, tioconazole, miconazole, econazole, and butoconazole) had a minimal effect on bacterial membranes, as they allowed for stabilization of the quenched dye. In contrast, the positive-control valinomycin-treated cell membranes displayed a substantial increase in fluorescence, indicating membrane depolarization (Fig. 5D). Azole-treated cells displayed essentially no change in fluorescence, revealing little to no depolarization compared with the known ionophore valinomycin.

Azoles do not contribute to bacterial ATP depletion.

Destabilization of membrane functions can impair respiratory chain functions and consequently reduce ATP levels (22). To preclude the likelihood of ATP depletion by the EPIs as the mechanism behind efflux inhibition, we evaluated alteration in the ATP synthesis levels after 4 h in the presence of azoles (clotrimazole, tioconazole, miconazole, econazole, and butoconazole). Carbonyl cyanide 3-chlorophenylhydrazone (CCCP) and valinomycin were included as a positive control due to their energy source-dissipating nature (30, 31). Fortunately, no depletion in the bacterial ATP levels was observed in the presence of azoles, while CCCP and valinomycin demonstrated significant disruption of ATP synthesis compared with the control (without treatment) (Fig. 5E).

Preeminent potency of the tetracycline-butoconazole combination in biofilm disruption.

Considering the strong potentiation of tetracycline activity engendered by azoles, we further sought to determine the effect of compounds on eradicating the preformed biofilm of S. aureus XU212. In the crystal violet (CV) assay, treatment with azoles (clotrimazole, tioconazole, miconazole, econazole, and butoconazole) alone at subinhibitory concentrations (1/4× MIC) resulted in a negligible impact on the biofilm biomass (see Fig. S2A in the supplemental material). Similarly, tetracycline treatment alone (128 μg/ml) did not affect the biofilm biomass. In contrast, the combination of tetracycline and all the azoles resulted in a significant decrease in the biofilm biomass. The combination of azoles, i.e., clotrimazole, tioconazole, miconazole, econazole, and butoconazole, along with tetracycline, decreased the biofilm biomass by 51% ± 5.87%, 49% ± 16.22%, 57% ± 3.42%, 50% ± 1.38%, and 63% ± 4.36%, respectively, compared with the untreated control (Fig. 6A).

FIG 6.

(A) Influence on the biofilm eradication ability of tetracycline (128 μg/ml) alone or in combination with the azoles at subinhibitory concentrations (1/4× MIC); the crystal violet staining assessed the biomass of S. aureus XU212 after exposure to tetracycline and the tetracycline-azole combination for 24 h. The percent values represent the amount of biofilm formation with respect to drug-free control. The experiments were carried out in three biological repeats, and results correspond to average ± SD. Results were considered significant when the P value was <0.05 (*) and highly significant when the P value was <0.001 (***). (B and C) Quantification of the effects of the azoles at subinhibitory concentrations (1/4× MIC) in combination with tetracycline at 1/2× MIC and synergistic MICs on mature biofilm; the viable bacterial cells were determined using the MTT assay after exposure for 24 h. The percent values represent the number of live cells with respect to the drug-free control. The experiments were carried out in three biological repeats, and results correspond to average ± SD. Results were considered significant when the P value was <0.01 (**) and highly significant when the P value was <0.001 (***). (D) Effect of tetracycline (128 μg/ml) alone or in combination with butoconazole (12.5 μM) on biofilm eradication assessed by confocal laser scanning microscopy (40×); static biofilms after exposure for 24 h were stained with SYTO9. S. aureus XU212 within biofilm on glass carriers display green fluorescence. The images are representatives of two independent experiments.

These results made us curious to determine the effect of the combination on the viability of cells in the biofilm biomass, and we performed a 3-(4, 5-dimethylthiazol-2-yl)-2, 5-diphenyl tetrazolium bromide (MTT) colorimetric assay. Unlike the previously used CV staining method, the MTT assay does not stain polysaccharides, DNA, proteins, and other biological molecules within the biofilm. Only live bacteria in the biofilms are stained in the MTT assay by measuring the metabolic activity of an individual bacterial cell. There is an excellent correlation between formazan concentrations (absorbance at 570 nm) and bacterial viability (32). The treatment with azoles alone at subinhibitory concentrations (1/4× MIC) resulted in a negligible impact on the viability of cells within the mature biofilm (Fig. S2B). The tetracycline treatment alone (128 μg/ml) had a significant impact, decreasing the preformed biofilm viability by 45% ± 5.66%. Simultaneously, the combination of tetracycline and all the azoles resulted in a substantial increase in biofilm disruption. The combination of azoles, i.e., clotrimazole, tioconazole, miconazole, econazole, and butoconazole, along with tetracycline, increased the bactericidal activity in preformed biofilm by 44% ± 0.52%, 46% ± 0.36%, 45% ± 3.09%, 48% ± 0.53%, and 48% ± 0.5%, respectively, compared with the tetracycline alone (Fig. 6B). We also performed the MTT assay with a combination of tetracycline at a synergistic concentration (4 and 8 μg/ml) along with azoles at subinhibitory concentrations. Interestingly, the combination reduced the viable cell population by ≥36% compared with tetracycline alone (26% ± 6.01% at 8 μg/ml and 20% ± 5.73% at 4 μg/ml) (Fig. 6C).

These assays served as a proof of concept in accordance with the role of EPIs in the potentiation of biocidal as well as biofilm disruption activity of tetracycline. Furthermore, we performed confocal microscopy to investigate the effect of butoconazole (best EPI) and tetracycline on biofilm structure. The regions of biofilm with green fluorescence indicate SYTO9 staining and represent viable cells. The biofilm of the untreated control showed sturdy architecture, and for tetracycline, a little disruption was observed. In contrast, the combination of butoconazole (12.5 μM) in combination with tetracycline (128 μg/ml) caused significant disintegration of the structure of biofilm as well as a reduction in the viable cell population (Fig. 6D).

DISCUSSION

The ever-growing multidrug resistance in bacteria has taken us to threatening situations such that bacterial infections sound like the worst nightmare. Sophisticated resistance mechanisms have evolved for bacteria, including powerful efflux pumps that extrude a wide range of antibacterial substrates (33). The broad specificities of multidrug efflux systems advise that their upregulation might result in the efflux of intracellular concentrations of antibiotics, influencing their clinical efficacy. Considering the dwindling antibiotic pipeline, the revival of previously discarded antibiotics can pave the path for the journey to tackle antimicrobial resistance, possibly by employing EPIs as adjuvant therapies. A limited structural homology between bacterial and mammalian efflux pumps (34) makes them a promising chemotherapy target.

The NorA efflux pump is one of the most studied ones in S. aureus (35), and given the fact that it is overexpressed in a considerable percentage of clinical strains, the past focus has been directed toward the investigation of NorA EPIs only. In addition to the NorA pump, there are some efflux pumps in S. aureus specific to a particular family of antibiotics; an example is Tet(K), which effluxes tetracycline (18). Apart from ribosomal protection, the Tet(K) efflux system is the foremost cause of the emergence of tetracycline-resistant MRSA strains (36). Very few Tet(K) EPIs have been reported (37, 38), and unfortunately, none of them is both FDA approved and used at clinically relevant concentrations. Thus, we set out to screen already approved drugs for potential Tet(K) efflux pump inhibition; however, our approach was previously validated by docking a virtual compound library of FDA-approved drugs to a homology model of the NorA efflux pump (39).

This study integrated an approach that combined homology modeling, molecular docking, and in vitro screening to discover potential new inhibitors of Tet(K). The initial screening using in silico docking (∼2,200 compounds) followed by potentiating modeling (200 compounds) utilized in this study allowed for the identification of five azoles, i.e., clotrimazole, tioconazole, miconazole, econazole, and butoconazole, that increased the effectiveness of tetracycline. The aromatic component of the azoles and their conformational poses seem to be essential contributors to stabilizing the interaction with the binding core of the Tet(K) protein. The major Tet(K) aromatic amino acids involved in pi-pi interactions are Tyr281, Phe62, and Phe12. Our results may highlight the possibility that the azoles act competitively and block the extrusion of substrate (antibiotics) molecules. Based on MM-GBSA-based binding energy calculations, the azoles have Tet(K) binding potential in the order butoconazole > miconazole > tioconazole > econazole > clotrimazole. Indeed, we observed EPI activity of these azoles in a similar order in our experiments, further validating our in silico studies. Furthermore, details of the mode of binding the azoles to Tet(K) presented in this study provide new insight into the potential molecular mechanism of MFS transporters.

The absence of intrinsic antibacterial activity makes these azoles advantageous for use as EPIs. Thus, according to one study (40), EPIs should be devoid of any antimicrobial activity because bacteria may lead to the development of resistance mechanisms against them. Next, the azoles enhanced the intrinsic susceptibility of S. aureus to tetracycline by severalfold in a dose-dependent manner. It was reported earlier that a decrease in the MICs by at least 4-fold concerning their original values in the presence of EPI was considered an indication of efflux inhibition (41, 42). Notably, in support of our approach, it has been shown that the concurrent treatment with an antibiotic and potentiator that blocks the resistance mechanism of that antibiotic but which lacks antibacterial property itself can lead to diminished resistance development (42). In total, the azoles demonstrated synergism with tetracycline and reduced its MIC below or up to the clinical resistance breakpoint against Tet(K)-expressing as well as clinical strains of S. aureus. The lack of synergism with nontetracycline antibiotics on efflux pump-overexpressing (NorA and MsrA) strains supports the specificity of the azoles as putative Tet(K) EPIs. In addition, by testing tetracycline-azole synergy with the agar-well diffusion assay, the zone of inhibition diameters of the tetracycline were enlarged by more than 10 mm. This increase indicates the ability of azoles to potentially reverse resistance and inhibit Tet(K) efflux pump that can extrude tetracycline, facilitating the treatment of tetracycline-resistant S. aureus infections. The antibiotic-EPI disc synergy test has been used in many studies to investigate the ability of some compounds to inhibit the efflux pump and reverse the resistance (43). Also, we think that azoles could be used to detect the Tet(K) efflux pump in S. aureus in clinical or laboratory settings.

The next step was to test the ability of azoles to inhibit the extrusion of the fluorescent dye EtBr in efflux assays. The accumulation and efflux of EtBr are good indicators of the involvement of efflux pumps in the resistance mechanism, particularly in Gram-positive bacteria such as S. aureus (17). The fluorescence-based EtBr efflux accumulation and inhibition studies of Tet(K)-possessing S. aureus cells revealed reduced efflux and enhanced uptake in the presence of azoles. Although EtBr is a common and nonspecific substrate of efflux pumps, azoles reduced the MIC of EtBr as well as inhibited its efflux in the strain expressing Tet(K). This observed phenomenon may be attributed to the Tet(K) inhibitory activity of azoles or due to non-EPI-like effects, which need to be investigated in the future. The uptake of tetracycline was investigated in the presence of azoles to test whether or not Tet(K) efflux inhibition activity of azoles could increase the accumulation of the antibiotic in the bacterial cell. Tetracycline is a naturally fluorescent antibiotic, and increased accumulation is direct evidence of increased concentration of antibiotics inside the cell (24). The results suggested that azoles indeed increased the concentration of tetracycline inside the S. aureus cells. These results increased our confidence and provided more realistic validation in the specificity of azoles discovered here.

In time-kill kinetics, synergy was stated as a difference of 2 log₁₀ CFU/ml between the combination and its most active counterpart, and the number of survivors in the combination must be ≥2 log₁₀ CFU/ml beneath the starting inoculum (44). We observed ≥3-log₁₀ reductions in CFU between the butoconazole-tetracycline combination and tetracycline alone, and the butoconazole-tetracycline combination exhibited synergy. Furthermore, butoconazole enhanced the PAE of tetracycline by 2.45 h (106% increment). This observation may have important therapeutic implications if an EPI endowed with PAE-causing activity is intended to be coadministered with antibiotics. Thus, theoretically, an EPI, which is used as an antibiotic potentiator, would not need to be present to ensure the effectiveness of the treatment, as long as the target organism retains sensitization to the antibiotic.

Many EPIs had been discovered earlier, but none of them is used clinically due to their off-target effects contributing to superfluous toxicities (22, 45). Disruption of the bacterial cell membrane causes efflux inhibition through a secondary effect of membrane depolarization, leading to inhibitory activity alone and the common identification of false-positive, nonspecific EPIs (46). Regarding our lead azoles, at least at the concentrations tested, we observed little to no membrane destabilization for S. aureus XU212. It has been previously reported that miconazole causes membrane damage by the K⁺ ion release (47). Unlike the previous reports, no such effect was observed in the presence of miconazole, perhaps due to the lower concentration tested and variation in bacterial strains. The loss of membrane integrity is closely related to the loss of the cell capacity to synthesize ATP (48). Nevertheless, due to similarities between the bacterial and the mammalian electron transport chain, the effects of azoles on ATP production need to be evaluated. We observed no depletion in the bacterial total ATP levels in the presence of EPIs. Conclusively, the synergistic activity exhibited by the azoles is not an outcome of any off-target effects but is indicative of the specificity and true EPI likeness of the drugs.

Active efflux has been reported to be linked with increased resistance in bacterial biofilms (49). Biofilms have a slow rate of cell growth and metabolism, contributing to an elevated level of resistance because most currently used antibiotics have been discovered by targeting primary metabolic pathways (for example, protein synthesis) or cell division. The synergistic action of EPIs was reported to abolish the formation of biofilms (40, 50, 51). These ideas prompted the assessment of the role of butoconazole in the eradication of biofilms, a major clinical problem in the current era of anti-infective drug discovery (52). The influence of the Tet(K) efflux pump on biofilm formation is still largely unknown. Because butoconazole alone did not show any effect on biofilm eradication, we presume that the effect of butoconazole on mature biofilms may not be due to direct disruption of biofilm but rather due to inhibition of Tet(K), leading to an accumulation of tetracycline. All the azoles discovered in this study, including butoconazole, are used as topical agents to treat candidiasis (53). Interestingly, biofilm formations by S. aureus isolates have been found in human skin lesions (54). Henceforth, the combination of tetracycline and azoles can be a potential anti-infective therapy, more so for the worsened S. aureus superficial infections involving robust biofilms.

In conclusion, the azoles are novel inhibitors of bacterial Tet(K) efflux pumps. Such inhibitors play a potential role in drug combinations by retaining higher antibiotic concentrations in bacteria and biofilm-associated infections.

MATERIALS AND METHODS

Chemicals, bacterial strains, and growth conditions.

All the chemicals, antibiotics, and dyes used in the study were purchased from Sigma-Aldrich Chemical Co. (St. Louis, MO, USA) unless mentioned otherwise. The tetracycline-resistant clinical isolate S. aureus XU212 has Tet(K) efflux pump expression (25). The MRSA strain ATCC 33591 and the control strain ATCC 29213 were purchased from HiMedia (India). The S. aureus ATCC BAA-39 strain with the MDR phenotype was procured from Chroma Chemie (India). The tetracycline-resistant S. aureus clinical isolates MRSA 1 and GMCH 6188 were collected from Government Medical College and Hospital, Chandigarh, India. The strains S. aureus SA-1199B (NorA overexpressed) and S. aureus RN4220-MsrA (transformed with pSK265, containing MsrA efflux gene) were also used in the study. All the strains were grown under standard culture conditions at 37°C in BBL cation-adjusted Mueller-Hinton broth (CA-MHB; BD, USA) unless specified otherwise. For CFU counting, Mueller-Hinton agar (MHA) (HiMedia) was used.

Homology modeling of Tet(K) and molecular docking studies.

The three-dimensional structure of the Tet(K) protein was modeled using the I-TASSER online server (55). The Tet(K) efflux pump receptor was prepared for docking using the Protein Preparation Wizard (56) by including hydrogen atoms followed by energy minimization using the OLPS3 force field with 0.3-Å default root mean square deviation (RMSD) (57). The processed Tet(K) model was then used to locate potential binding sites using the SiteMap module (58). The potential binding site with a high site score (>0.9) was used for docking grid generation using the Receptor Grid Generation module by keeping the grid box dimension at 25 Å. The FDA-approved drug library was downloaded from the DrugBank database (www.drugbank.ca) and prepared for docking study using the LigPrep module (59). The molecular docking studies were performed using XP mode in the Glide module of Schrodinger suite 2019, and the potential compounds were sorted based on their docking score. The binding energy scores of ligands and receptors were calculated using the MM-GBSA module in the Schrodinger software suite. The docked Tet(K) and drug complex were separated and loaded as receptor or ligand, respectively, in the MM-GBSA module. The MM-GBSA-based analysis includes five energy calculation methods, including optimization of free ligand, optimization of free receptor, optimization complex, ligands from minimized complex, and receptors from the minimized complex (60). All the molecular graphics figures were prepared using PyMOL (61).

MIC determination.

MICs were determined using the broth-microdilution assay in CA-MHB as per the Clinical and Laboratory Standards Institute (CLSI) guidelines (62). The assay was conducted in 96-well polystyrene tissue culture plates (Corning, USA), and the wells containing 2-fold serial dilutions of the compounds were inoculated with 5 × 10⁵ CFU/ml of the bacterial suspension. The plates were incubated for 18 h at 37°C, and the minimum concentration with no visual turbidity was regarded as the MIC of the compound.

Determination of growth kinetics.

For the growth kinetics assay, a freshly grown culture of S. aureus XU212 was prepared up to a final density of 5 × 10⁵ CFU/ml and cocultured with subinhibitory concentrations of azoles. The final concentrations of clotrimazole, tioconazole, miconazole, econazole, and butoconazole used in this experiment were 25 μM, 25 μM, 12.5 μM, 25 μM, and 12.5 μM, respectively. At the same time, the control group (without treatment) was included for comparison. The cell growth densities based on the absorbance at 600 nm in CA-MHB were measured at every 1-h interval from 0 h to 21 h.

Checkerboard synergy assay.

To determine the synergistic activity of the azoles with tetracycline, we performed the broth microdilution checkerboard synergy assay. The wells of 96-well polystyrene tissue culture plates were loaded with 2-fold serial dilutions of the antibiotics down the assay plate along with various concentrations of compounds across the assay plate. The assay plates were loaded with 100 μl of 5 × 10⁵ CFU/ml of bacterial inocula and incubated for 18 h at 37°C. FICI values were determined according to the following formula:

$$\mathrm{FICI}={\mathrm{FICI}}_{\mathrm{A}}+{\mathrm{FICI}}_{\mathrm{B}}$$

$${\mathrm{FICI}}_{\mathrm{A}}=\mathrm{MIC\; \hspace{0.17em}of\; \hspace{0.17em}compound\; \hspace{0.17em}A\; \hspace{0.17em}in\; \hspace{0.17em}combination/MIC\; \hspace{0.17em}of\; \hspace{0.17em}compound\; \hspace{0.17em}A\; \hspace{0.17em}alone}$$

$${\mathrm{FICI}}_{\mathrm{B}}=\mathrm{MIC\; \hspace{0.17em}of\; \hspace{0.17em}compound\; \hspace{0.17em}B\; \hspace{0.17em}in\; \hspace{0.17em}combination/MIC\; \hspace{0.17em}of\; \hspace{0.17em}compound\; \hspace{0.17em}B\; \hspace{0.17em}alone}$$

The FICI value of ≤0.5 was considered synergy, whereas 0.5 < FICI ≤ 4.0 was an “indifference” effect, and the FICI value of >4.0 was regarded as an “antagonistic” effect (63).

Agar well diffusion assay.

For this method, the test microorganism (S. aureus XU212) was grown up to an optical density at 600 nm (OD₆₀₀) of 0.3 and seeded onto MHA plates at the final inoculum of 10⁵ CFU/ml. The plates inoculated with the test organism had 8-mm wells cut with a sterile stainless steel cork borer. The individual wells were loaded with a 50-μl volume of the azoles (1/4× MIC) and tetracycline alone and a combination of both. The plates were incubated for 24 h at 37°C and were assessed for the zone of growth inhibition. The zone of clearance around the wells was measured with the HiAntibiotic Zonescale.

EtBr accumulation assay.

The fluorometric estimation of EtBr accumulation was performed as described previously (17), with some modifications. The strains were grown to the log phase in CA-MHB and then centrifuged at 16,060 × g for 15 min. The pellet was washed two times and resuspended in uptake buffer (110 mM NaCl, 7 mM KCl, 50 mM NH₄Cl, 0.4 mM Na₂HPO₄, 52 mM Tris base; pH 7.5). The culture with an OD₆₀₀ of 0.3 was treated with EtBr (4 μg/ml) and azoles at subinhibitory concentrations. The gain of fluorescence was recorded for 45 min at excitation and an emission wavelength of 530 nm and 600 nm, respectively, in a microplate reader (BioTek, USA). The RFF index was calculated according to the following formula: RFF = (RFF_{treated at 45 min} − RFF_{untreated at 45 min})/RFF_{untreated at 45 min} (22). Additionally, concentration-dependent time-based accumulation curves were plotted.

Tetracycline efflux inhibition assay.

The experiment was performed as described previously with slight modifications (24). Briefly, the freshly grown bacterial culture (mid-log phase) was centrifuged at 9,000 × g and then washed twice in 10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer (pH 7.2). Finally, the bacteria were resuspended to 10⁸ CFU/ml supplemented with tetracycline (1/4× MIC) and azoles (1/4× MIC) and incubated for 2 h at room temperature (25°C). After 2 h, bacteria were pelleted, washed once, and resuspended in HEPES buffer. Furthermore, the bacterial suspension was loaded into the wells of 96-well black flat-bottom plates. The change in fluorescence was monitored in the presence and absence of glucose (4%). In the control experiment, DMSO was added instead of azoles. The emission of tetracycline was recorded at 535 nm by exciting at a wavelength of 405 nm for a time period of 25 min (5-min interval). A slow decrease in the tetracycline fluorescence intensity was indicated by the decreased extrusion of tetracycline from the bacterial cells.

EtBr efflux inhibition assay.

The fluorometric estimation of the efflux inhibitory potential of azoles was performed by a method described earlier (23), with some modifications. The bacterial cultures were grown at 200 rpm and 37°C until the OD₆₀₀ of 1.0. The culture was pelleted at 16,060 × g for 10 min, then washed twice, and diluted to OD₆₀₀ of 0.4 (without glucose) in phosphate-buffered saline (PBS; pH 7.4). The cells were loaded with EtBr (4 μg/ml) and azoles at subinhibitory concentrations such that maximum accumulation occurred (no glucose; incubated at 25°C for 60 min). When maximum accumulation was reached (after 60 min), bacteria were pelleted at 16,060 × g for 15 min and then resuspended in PBS. Finally, the aliquots were transferred to 96-well black tissue culture plates (Corning, USA) with and without glucose, and fluorescence was recorded for 45 min (9-min time interval) at excitation and emission wavelengths of 530 nm and 600 nm, respectively, in a microplate reader (BioTek, USA).

Time-kill kinetics.

For time-kill kinetics, S. aureus XU212 was incubated at 37°C and grown to mid-log phase (OD₆₀₀ of 0.3). The final inoculum was adjusted to 10⁵ CFU/ml, and cells were treated with butoconazole (12.5 μM), tetracycline (256 μg/ml), and the combination of both. The cells were incubated for 24 h at 37°C with shaking. The culture without any treatment served as a control. At different time points, 100 μl of culture was withdrawn, and different dilutions were used to spread onto MHA plates. The colonies were counted, and CFU/ml was assessed. The CFU/ml was plotted against different time points for each set.

Determination of PAE by turbidimetry.

The suppression of bacterial growth that continues after a short exposure to antibiotics has been defined as PAE (26). Inoculum from a fresh culture of S. aureus XU212 was grown in CA-MHB to mid-exponential phase (OD₆₀₀ ≈ 0.3). This culture was incubated with tetracycline (256 μg/ml) alone and combined with butoconazole (12.5 μM) for 2 h at 37°C, with shaking. The suspensions were diluted 50 times to eliminate the drug carryover, and a 250-μl aliquot from the suspension was dispensed in triplicates into a 96-well flat-bottom microtiter plate (Cole-Parmer, USA). The untreated cellular suspension was regarded as the control for the calculation of PAE. PAE was calculated according to the formula PAE = T₅₀ − C₅₀, where T₅₀ and C₅₀ are the time in hours required for the drug-treated and untreated cultures, respectively, to reach a value of OD₆₀₀ corresponding to 50% of the final absorbance reached by an untreated control (64). The cellular concentration of the control set was adjusted in accordance with treated suspension prior to growth resumption in order to minimize the difference in the inoculum.

Membrane permeabilization assay.

The membrane permeabilization assay was performed using PI according to the manufacturer’s instructions, with some modifications. The bacterial culture S. aureus XU212 was grown to log phase in CA-MHB and washed twice in 0.85% saline at 10,000 × g. The OD₆₇₀ was adjusted to 0.4. Furthermore, the cells were incubated in the presence of azoles (1× MIC) and paenibacillin (20 μg/ml) for 1 h at 37°C. The treated culture was centrifuged at 10,000 × g to remove the effect of azoles and resuspended in saline. The PI at a final concentration of 30 μM was added to 96-well polystyrene black flat-bottom plates containing cells treated with azoles. The fluorescence was read for 24 min (3-min time interval) in a microplate reader (BioTek, USA) at the excitation and emission wavelengths of 490 nm and 635 nm, respectively. Paenibacillin was used as a positive control because of its strong membrane-permeabilizing ability (28).

Membrane depolarization assay.

The DiSC₃(5) assay was performed to determine the effect of compounds on the bacterial membrane depolarization, as described previously (65). DiSC₃(5) is a membrane potential-sensitive probe. The bacterial culture S. aureus XU212 was grown to log phase and washed twice in 5 mM HEPES buffer (containing 100 mM KCl, pH 7.2). The cells were incubated with KCl to equilibrate the cytoplasmic and external K⁺ ion concentrations. The cells were prepared as described for the PI assay and then treated with DiSC₃(5) (0.4 μM) for 60 min at 37°C. Afterward, the cultures were incubated with azoles, CCCP, or valinomycin at different concentrations for 30 min in a 96-well black flat-bottom plate. The alteration in fluorescence was measured at an excitation and emission of 622 nm and 670 nm, respectively.

Determination of intracellular ATP levels.

To determine the effect of azoles on the intracellular level of ATP, we used an ATP determination kit (Invitrogen, Life Technologies, USA). The assay was performed as described earlier (66), with some modifications. Briefly, the bacterial strain S. aureus XU212 was grown to the mid-log phase, and OD₆₀₀ was adjusted to 0.3. The cells were incubated with subinhibitory concentrations of azoles for 4 h at 37°C, lysed using an ultrasonic water bath, inactivated by heating, and immediately deep-frozen. The cell lysate was used for the measurement of persistent ATP and plotted as relative luminescence units. CCCP and valinomycin were included for comparison.

Biofilm eradication assay.

The biofilm eradication assay was performed, as mentioned earlier (67), with some modifications. Briefly, the overnight grown culture of S. aureus XU212 was diluted 1:200 times in tryptic soy broth (TSB) supplemented with 2% glucose and dispensed into wells of a 96-well flat-bottom plate. The plate was incubated under static conditions for 48 h. After incubation, the wells with preformed biofilm were washed with PBS to eliminate planktonic bacteria and treated with azoles (at subinhibitory concentration) and tetracycline (128 μg/ml) alone as well as in combination. The plates were incubated for 24 h, again washed with PBS, and then fixed with 99% methanol for 15 min. The plate was allowed to dry under laminar airflow, and the adherent bacteria in the biofilm were stained with filtered CV (0.5%) for 5 min at room temperature. The excess stain was removed by washing with water until the negative-control wells (without biofilms) appeared colorless. Then, the stained cells were solubilized with 33% acetic acid, and the released stain was measured at 570 nm. The results are presented as the percent biofilm formation with respect to the control (without treatment) group.

Determination of viable cell population in biofilm.

Furthermore, we performed the MTT assay for the quantification of viable cells in the biofilm (32). The assay was performed as described above, but MTT (1 mg/ml) was used instead of CV. After the fixation of cells, MTT was added, and plates were incubated for 4 h at 37°C. The cells were washed once to release excess dye and solubilized using a solution (40% [vol/vol] dimethylformamide in 2% [vol/vol] glacial acetic acid added with 16% sodium dodecyl sulfate), and the released stain was measured at 570 nm. The results are presented as the percentage of viable adherent bacteria in the biofilm with respect to the control (without treatment) group.

Confocal microscopy for biofilm eradication.

The bacterial cell suspension was prepared in a similar manner as the CV assay (50). For biofilm attachment, the coverslips were treated with poly-l-lysine for 2 to 3 h at 70°C and kept overnight at room temperature. The coverslips (with coated side upward) were placed in 12-well polystyrene tissue culture plates (Corning, USA) and dispensed with 1 ml of culture suspension. The plates were incubated for 48 h at 37°C, then washed twice with PBS (PBS 1×; Gibco), and treated with tetracycline (128 μg/ml) alone and in combination with butoconazole (12.5 μM) in fresh CA-MHB. The wells with no treatment served as a control, and the plates were again incubated for 24 h. After treatment, the medium was discarded, and the wells were washed thrice with 0.85% saline and then stained with SYTO9 (Invitrogen, Life Technologies, USA) dye for 35 to 40 min. The wells were washed once to remove the excess dye and allowed to dry a little under laminar airflow. The coverslips were mounted inverted on the slides using mounting oil. The stained biofilms were analyzed using inverse confocal laser scanning microscopy (CLSM) [Nikon AI(R)] at 488-nm excitation. The experiment was performed in duplicates, and images were visualized and processed using NIS-Elements Viewer 4.50 software.

Statistical analysis.

All experiments were carried out in biological replicates, and similar results were obtained on all occasions. The data are represented as mean ± standard deviation (SD). Multiple mean t tests (two-tailed) were used to determine the differences among groups by using the GraphPad Prism 8.0.2 software package. A P value of <0.05 (*) was considered statistically significant, and P values of <0.01 (**) and <0.001 (***) were considered highly significant.