Unveiling the potent activity of a synthetic ion transporter against multidrug-resistant Gram-positive bacteria and biofilms

Abstract

The increasing prevalence of drug-resistant infections caused by Gram-positive bacteria poses a significant threat to public healthcare. These pathogens exhibit not only smart resistance mechanisms but also form impenetrable biofilms on various surfaces, rendering them resilient to conventional therapies. In this study, we present the potent antibacterial activity of a synthetic ion transporter T against multi-drug resistant (MDR) Gram-positive pathogens, with minimum inhibitory concentration (MIC) values ranging from 0.5 to 2 μg mL⁻¹. The compound demonstrates high selectivity with negligible toxicity towards mammalian cells (HC₅₀ = 810 μg mL⁻¹). It exhibits fast killing kinetics, completely eliminating >5 log bacterial cells within 12 h. Moreover, the compound displays efficacy against both planktonic bacteria and preformed biofilms of methicillin-resistant S. aureus (MRSA), reducing the bacterial burden within the biofilm by 2 log. Mechanistic investigations reveal that the ion transporter depolarizes the bacterial membrane potential and enhances membrane permeability. Additionally, it generates reactive oxygen species, contributing to its bactericidal activity. Notably, MRSA did not exhibit detectable resistance to the ion transporter even after serial passaging for 10 days. Collectively, this novel class of ion transporter holds promise as a therapeutic candidate for combating infections caused by multi-drug resistant Gram-positive bacteria.

Synthetic ion-transporter kills multidrug-resistant Gram-positive bacteria and disrupts their biofilms.

1. Introduction

The increasing prevalence of infectious diseases and the emergence of microbial resistance pose significant challenges to clinicians worldwide.^1–10 A recent global study reported a staggering 4.5 million deaths in 2019 attributable to antibacterial resistance, with 1.27 million lives lost directly due to resistance to conventional antibiotics.¹¹ In addition to the emergence of Gram-negative bacteria, infections caused by MDR Gram-positive bacteria high-priority pathogens such as MRSA, vancomycin-resistant S. aureus (VRSA), and vancomycin-resistant E. faecium (VRE) identified by the World Health Organization (WHO),¹² endowed one of the major hurdles to tackle with conventional antibiotic therapy.^7,9,10 Furthermore, the ability of these drug-resistant bacteria to form biofilms further complicates treatment strategies.^9,10 Conventional antibiotics struggle to eradicate biofilm-associated infections due to limited penetration of the biofilm extracellular matrix (ECM) and the presence of phenotypically diverse bacterial cells within the biofilm.⁹ This situation has created a significant void in the antibiotic arsenal, demanding the development of new antibacterial agents to combat antimicrobial resistance.⁹

The bacterial membrane, a crucial component of bacteria, offers an attractive target for novel antibacterial agents.^13–21 Small molecular designs that disrupt ionic homeostasis and permeabilize the bacterial cell membrane has garnered considerable interest among researchers.^22–31 These synthetic analogues exhibit enhanced selectivity, stability, and ease of synthesis compared to natural AMPs. Synthetic amphiphilic transporters, inspired by natural antimicrobial peptides (AMPs) known for their activity against various pathogens, including bacteria, viruses, fungi, and cancer cells, have emerged as promising candidates.^22–37 In the recent past, few of these ion-transporters exhibited antibacterial efficacy alone or in combination with other antibiotics.^23–31 Particularly, Gokel and co-workers demonstrated that ion transporters can hamper the bacterial efflux machinery and rejuvenate resistant antibiotics.^24,25 Schmitzer and co-workers recently reported anion transporters based on benzimidazolium salts with good antibacterial activity.^23,27 However, the efficacy of these synthetic ion transporters in disrupting bacterial biofilms and the detailed mechanisms underlying bacterial killing are not well understood. Herein, we report the potent antibacterial activity of a synthetic ion transporter T, designed based on a 3,7-diazabicyclo[3.3.1]nonane (bispidine) backbone substituted with melamine (Fig. 1A). Each melamine ring was further connected with two butyl amino groups at C-2 and C-4 positions, creating hydrogen bonding sites and imparting amphiphilicity. Earlier X-ray crystallographic studies for the HCl salt of T indicated a zig-zag network.³⁸ However, in the lipid membrane, the compound forms a dimeric supramolecule, bridged through four HCl molecules, and facilitates efficient symport of H⁺ and Cl⁻ ions, with maximal activity observed at pH 7.

Fig. 1. (A) Chemical structure of the synthetic ion transporter, T in free form and upon binding with HCl; (B) antibacterial activity (MIC) of T against S. aureus MTCC 737 and E. coli MTCC 443. Vancomycin and colistin was used as control antibiotics; N.D. stands for not determined, HC₅₀ stands for concentration to lyse 50% of the red blood cells. (C) Hemolytic activity of T. (D) Antibacterial activity of T against different Gram-positive bacteria and their clinical isolates. Vancomycin and linezolid were used as control antibiotics. (E) Selectivity index (HC₅₀/MIC) of T against the tested Gram-positive bacteria and their clinical isolates. Each experiment was performed in triplicate and the entire experiment was repeated twice. The average values are provided with standard deviations wherever applicable.

We hypothesized that altering HCl homeostasis using this ion transporter could affect the membrane potential crucial for bacterial cell metabolism, including cell division, efflux machinery operation, and energy generation. Based on this hypothesis, we evaluated the antibacterial activity of the synthetic ion transporter against a wide range of multi-drug resistant Gram-positive bacteria. We assessed its toxicity towards human red blood cells and determined its selectivity towards bacteria. Time-kill kinetics experiments were performed against two superbugs, MRSA and VRSA. Moreover, we investigated the biofilm disruption ability of the ion transporter against mature MRSA biofilms. We conducted membrane depolarisation and permeabilization assays to elucidate its bactericidal mechanism of action. Additionally, we investigated the production of reactive oxygen species (ROS) in bacterial cells and studied the propensity of resistance development against MRSA.

2. Results and discussion

2.1. Antibacterial activity

The ion transporter T was synthesized by reacting 3,7-diazabicyclo[3.3.1]nonane 1 with N²,N⁴-dibutyl-6-chloro-1,3,5-triazine-2,4-diamine 2, facilitated by DIPEA base in THF solvent under reflux conditions (Scheme S1†), following the established protocol.³⁸ The antibacterial activity of the ion transporter was initially assessed against two model bacteria, S. aureus MTCC 737 and E. coli MTCC 443, using the broth dilution method. The compound demonstrated excellent activity against the Gram-positive bacterium S. aureus, with a MIC value of 0.5 μg mL⁻¹, and moderate activity against the Gram-negative bacterium E. coli, with a MIC value of 64 μg mL⁻¹ (Fig. 1B). The potent activity against Gram-positive S. aureus prompted further investigation into its activity against other Gram-positive bacteria and their clinical isolates. When tested against two commonly occurring Gram-positive pathogens, Staphylococcus epidermidis and Enterococcus faecium, the ion transporter exhibited very good antibacterial activity, inhibiting the growth of these bacteria at concentrations of 4 and 0.5 μg mL⁻¹, respectively (Table 1 and Fig. 1D). Moreover, when tested against the superbug MRSA (methicillin resistant Staphylococcus aureus) and its clinical isolates, the ion transporter displayed similar activity (MIC: 0.5–2 μg mL⁻¹) to that of vancomycin, which is considered the last resort antibiotic for Gram-positive bacteria (Table 1 and Fig. 1D). Notably, the ion transporter, T, exhibited significantly superior activity against vancomycin-resistant bacteria, including vancomycin-resistant S. aureus (VRSA) and vancomycin-resistant E. faecium (VRE), with MIC values ranging from 0.5 to 1 μg mL⁻¹, while vancomycin itself exhibited MIC in the range of 512–1024 μg mL⁻¹ (Table 1 and Fig. 1D).

Antibacterial activity and toxicity of T. Cytotoxicity was tested against Madin Darby canine kidney (MDCK) cell-line. N.D. stands for not determined.

Antibacterial agents	MIC (μg mL−1)											HC50 (μg mL−1)	EC50 (μg mL−1) against MDCK cell line
	S. aureus MTCC 737	S. epidermidis MTCC 3615	E. faecium ATCC 19634	MRSA ATCC 33591	MRSA R3545	MRSA R3890	MRSA R3889	VRSA-1	VRSA-4	VRE-903	VRE-909
T	0.5	4	0.25	0.5	0.5	0.5	0.5	0.5	0.5	1	1	810	21
Vancomycin	1	1	1	1	1	1	1	512	512	1024	512	N.D.	N.D.
Linezolid	2	2	4	2	2	2	2	1	1	32	2	N.D.	N.D.

Overall, the compound demonstrated broad-spectrum activity against a wide range of Gram-positive bacteria and their clinical isolates. These findings highlight the potential of the synthetic ion transporter as a promising candidate for combating infections caused by multidrug-resistant Gram-positive bacteria, including those resistant to vancomycin.

2.2. Hemolytic activity and selectivity

In the development of antibacterial agents, potent antibacterial activity with minimal toxicity is a crucial criterion. To evaluate the toxicity of the ion transporter, its effect on human red blood cells (hRBCs) was assessed, and the concentration corresponding to 50% lysis of hRBCs (HC₅₀) was determined. Remarkably, the compound exhibited an HC₅₀ value of 810 μg mL⁻¹, which is approximately 800–1600 times higher than its MIC values against different pathogens (Fig. 1C). Importantly, at its active concentration range of 0.5–2 μg mL⁻¹, the ion transporter caused minimal lysis of red blood cells from 0–3% (Fig. 1C). Furthermore, the selectivity index (SI) of the molecule was calculated by considering the ratio between HC₅₀ and MIC values. For the drug sensitive pathogens (S. aureus, S. epidermidis and E. faecium), the SI was determined to be 400–800 (Fig. 1E). Notably, when tested against the multidrug-resistant clinical isolates, the compound exhibited a selectivity index of 800–1600, validating its selective antibacterial efficacy (Fig. 1E). These findings underscore the promising selectivity and low toxicity profile of the ion transporter.

2.3. Cytotoxicity against MDCK cell line

To further validate the selectivity of the ion transporter, its cytotoxicity was tested against the Madin Darby canine kidney cell line through Alamar blue assay. The molecule showed negligible toxicity (>95% cell viability) against mammalian cells at its active concentration (MIC = 0.5–4 μg mL⁻¹) (Table 1). However, at higher concentrations, it showed a certain level of cell toxicity, underscoring the necessity of further optimisation of the structural parameters for enhanced selectivity.

2.4. Time kill kinetics

Time-kill kinetics experiments were performed to assess the bactericidal properties of the compound. The study focused on two superbugs, MRSA and VRSA, which have been identified as top-priority pathogens by the World Health Organization (WHO). The compound, T, exhibited dose-dependent killing kinetics, effectively eliminating 5.5 log bacteria within a 12 h period (Fig. 2). Specifically, when tested against MRSA ATCC 33591, at 8 μg mL⁻¹, the compound completely eradicated all the bacterial cells (>5 log reduction) within 12 h (Fig. 2A). Similarly, against VRSA-1 at the same concentration, a 3.5 log reduction in bacterial titer was demonstrated after 12 h (Fig. 2B). However, at a higher concentration of 16 μg mL⁻¹, the compound exhibited faster killing kinetics, completely clearing bacterial count within the same time frame. These findings underscore the potent bactericidal activity of the compound.

Fig. 2. Time-kill kinetics of the ion transporter, T, against (A) MRSA ATCC 33591 and (B) VRSA 1; MRSA biofilm disruption ability of T. (C) Reduction in the biofilm biomass upon treatment with T and vancomycin; (D) viability of bacterial cells embedded within biofilm matrix; (E) CLSM images of the MRSA biofilm upon simultaneous staining with SYTO-9 and propidium iodide. Vancomycin was used as an antibiotic control. Each experiment was performed in duplicate and the entire experiment was repeated twice. The average values are provided with standard deviations wherever applicable. Scale bar = 10 μm. Asterisks indicate <50 CFU mL⁻¹.

2.5. Biofilm disruption efficacy

Biofilm disruption is a crucial aspect to address in the battle against chronic and persistent infections, as ∼80% of drug-resistant infections are associated with bacterial biofilm.⁵ Biofilms are complex microbial communities enclosed within a self-generated extracellular matrix (ECM), which creates diffusion barriers for antimicrobial agents and harbors predominantly metabolically inactive or dormant bacterial cells.⁵ Thus, evaluating the ability of the compound to disrupt biofilms was a key focus of this study.

Against a preformed mature biofilm of MRSA, the compound exhibited significant biofilm disruption efficacy. At 32 μg mL⁻¹, it achieved a 50% reduction in biofilm biomass, which further reduced to 40% at 64 μg mL⁻¹ (Fig. 2C). Notably, the compound was not only disrupted by the biofilm matrix but also effectively killed the bacterial cells embedded within the biofilm. At 32 μg mL⁻¹, it caused a 1.7 log reduction in bacterial burden, while a higher concentration of 64 μg mL⁻¹ led to a ∼2 log reduction upon single treatment (Fig. 2D). In comparison, the last resort antibiotic vancomycin, at 32 μg mL⁻¹ exhibited minimal reduction in biofilm biomass and only caused a 0.5 log reduction in bacterial population viability. Although at a higher concentration (64 μg mL⁻¹), vancomycin displayed slightly improved anti-biofilm activity, reducing 30% of biofilm biomass and resulting in a 1.5 log reduction in cell viability (Fig. 2D). However, it is worth noting that at the same concentration, the compound demonstrated superior anti-biofilm activity compared to vancomycin. The cationic charge of the compound and the presence of hydrophobic groups likely facilitated its interaction with the negatively charged ECM, leading to its effective eradication. These findings underscore the potential of the compound as a promising biofilm-disrupting agent, surpassing the limited efficacy of conventional antibiotics like vancomycin.

The dissemination of bacterial biofilm was further visualized using confocal laser scanning microscopy (CLSM) coupled with the LIVE/DEAD assay, providing valuable insights into the effectiveness of the compound. For this purpose, simultaneous staining with Syto-9 (green fluorescence, indicating both live and dead cells) and propidium iodide (red fluorescence, indicating only dead cells) was conducted. CLSM images of the untreated control revealed a dense biofilm measuring 12 μm in thickness, predominantly consisting of live bacteria (Fig. 2E). Treatment with vancomycin at 64 μg ml⁻¹ resulted in a marginal reduction in biofilm thickness with a thickness of 10 μm, but failed to eradicate the enclosed bacteria as indicated by the predominantly green fluorescence (Fig. 2E). In contrast, treatment with the compound led to a remarkable reduction in biofilm thickness to 3 μm, accompanied by the colocalization of green and red cells, which confirmed the successful eradication of MRSA (Fig. 2E). These findings collectively provide strong evidence supporting the ability of the compounds to disrupt biofilms. The CLSM images vividly illustrate the significant reduction in biofilm thickness and the concurrent killing of MRSA cells upon treatment with this cationic lipophilic compound.

2.6. Preliminary mechanism of action

Within the lipid membrane, the receptor forms a dimeric self-assembly, creating a molecular arrangement that facilitates the efficient transport of H⁺ and Cl⁻ ions. This transport process plays a crucial role in the mechanism of action of the compound. The observed pH-dependent Cl⁻ efflux exhibited by the compound can be explained by a gradual protonation model, wherein the compound acts as an active transmembrane carrier at physiological pH. Previous studies have already demonstrated the transport of HCl in a model liposomal membrane using this compound, supporting its potential as a proton carrier. To gain deeper insights into the mechanism of action of the compound, further investigation was carried out using a range of spectroscopic and microscopic techniques.

2.7. Depolarisation of cytoplasmic membrane potential

Previous studies have established that the ion transporter has the ability to modulate membrane permeability, leading to the efflux of ions, as observed in model liposomes.³⁸ This alteration in ion balance has the potential to disrupt the normal functioning of bacterial cells by changing their membrane potential.³⁹ To investigate the impact of the compound on bacterial membrane potential, we utilized a membrane-potential sensitive dye called 3,3′-dipropylthiadicarbocyanine iodide [DiSC₃(5)]. This dye is distributed both inside and outside of the lipid membrane, and its fluorescence is self-quenched in polarised and intact cells. However, when the membrane is depolarised and damaged, the dye is released into the solution, resulting in an increase in fluorescence intensity over time. In this study, MRSA and VRSA were treated with different concentrations of the compound, and the fluorescence intensity of DiSC₃(5) dye was measured. The results showed a concentration-dependent increment in fluorescence intensity, indicating depolarization of the bacterial transmembrane potential upon treatment with the ion transporter (Fig. 3A and B). Notably, a stronger effect was observed at higher compound concentrations, demonstrating a dose-dependent relationship. These findings provide compelling evidence that the ion transporter possesses the capability to depolarise the transmembrane potential of bacterial cells.

Fig. 3. Membrane depolarising ability of T against (A) MRSA ATCC 33591 and (B) VRSA 1 at different concentrations. Arrow indicates the time of compound addition. Quantification of depolarised bacterial cells by calculating R/G ratio upon staining with DiOC₂(3) dye. (C) R/G ratio for control, T (8 μg mL⁻¹) and CCCP (10 μg mL⁻¹); determination of R and G population through flow cytometry (D) control, (E) T (8 μg mL⁻¹) and (F) CCCP (15 μg mL⁻¹).

2.8. Quantification of membrane depolarisation

To quantify the number of depolarised cells following treatment with the compound, we employed the use of DiOC₂(3) (3,3′-diethyloxacarbocyanine iodide) dye. This dye exhibits green fluorescence in all bacterial cells. However, at the higher cytosolic concentrations, it self-aggregates to show red fluorescence caused by high membrane potential in healthy and intact cells. Again, when the membrane potential is compromised or abolished, it returns to a green fluorescent state. Therefore, carbonyl cyanide 3-chlorophenylhydrazone (CCCP), an ionophore that disrupts the membrane potential by eliminating the proton gradient, induces a significant shift in the ratio of red and green fluorescence (R/G ratio). To determine the extent of depolarized cells upon treatment with the compound, we quantified R/G ratio using flow cytometry (Fig. 3C). In the control group, the majority of cells exhibited red fluorescence (red population 63.8% and green population 27.3%; R/G ratio = 2.34), confirming their healthy and polarised state (Fig. 3C and D). In contrast, incubation with the compound (at 8 μg mL⁻¹) resulted in a substantial shift from red to green fluorescence (red population 47.77% and green population 43.18%; R/G ratio = 1.10), indicating the depolarised state of the bacterial cells (Fig. 3C and E). The positive control, CCCP, similarly disrupted the membrane potential and induced depolarization in bacterial cells, leading to a significant presence of green fluorescence (red population 49.13% and green population 47.82%; R/G ratio = 1.10) (Fig. 3C and F). These observations indicate that the compound acts as an ionophore, akin to CCCP, and disrupts the proton gradient to depolarize the transmembrane potential of bacterial cells.

2.9. Inner membrane permeabilisation through LIVE/DEAD assay

The membrane activity of the ion transporter, T was validated using live/dead staining and fluorescence microscopy (Fig. 4A). MRSA cells were incubated with the compound for 4 h, followed by staining the bacterial suspension with Syto-9 (green emission) and PI (red emission) simultaneously, and imaging under a microscope. Syto-9 is a membrane permeable dye that stains both live and compromised bacteria, while PI is cell-impermeable and selectively stains dead or membrane compromised bacteria. In the untreated control group of MRSA, no red cells were observed in the microscopic image, indicating the presence of live bacteria with intact membrane integrity. However, upon treatment with the compound, MRSA cells were stained by both Syto-9 and PI, indicating the presence of dead bacteria with compromised membrane integrity (Fig. 4A). This finding provides further confirmation of the mechanism of action of the compound – as an agent that targets and disrupts bacterial membranes. The live/dead staining results, in conjunction with previous findings, support the notion that the ion transporter exerts its antibacterial activity by acting on bacterial membranes.

Fig. 4. (A) Inner membrane permeabilization of T in MRSA ATCC 33591 determined through LIVE/dead assay. (B) Generation of reactive oxygen species measured upon increment in fluorescence intensity of DCF dye. (C) Propensity of resistance development by MRSA ATCC 33591 against T and fusidic acid upon serial passaging for 10 days. Scale bar = 10 μm.

2.10. Reactive oxygen species (ROS) generation

The bactericidal mechanism of action of the ion transporter was further investigated in MRSA cells by examining its ability to induce the generation of reactive oxygen species (ROS). ROS generation beyond a certain threshold level is a crucial factor in bacterial cell death, as ROS, including superoxide, hydrogen peroxide, and hydroxyl radicals, can cause various intracellular damage. Hydroxyl radicals, for example, can break DNA, induce lipid peroxidation, and participate in the carbonylation of proteins. Agents that target bacterial membranes, such as daptomycin, chlorohexidine, etc., have been shown to trigger a substantial production of ROS inside bacterial cells, which in turn has a detrimental effect on their viability. To evaluate the production of ROS in MRSA cells upon treatment with the compound, 2′,7′-dichlorofluorescin diacetate (DCFH-DA) dye was used. DCFH-DA is a non-fluorescent, cell-permeable molecule that is converted to DCF (a fluorescent compound) after reacting with intracellularly generated ROS. A significant increase in fluorescence intensity observed in the compound-treated cells compared to the untreated cells clearly indicated the generation of intracellular ROS (Fig. 4B). The production of ROS showed a dose-dependent pattern, with a higher concentration of compound (e.g., 16 μg mL⁻¹) leading to greater fluorescence intensity. These results collectively establish that the compound induces the production of intracellular ROS, which subsequently damages the bacterial cell membrane.

The generation of ROS by the ion transporter represents an important aspect of its bactericidal mechanism, as it contributes to the overall antibacterial activity by causing significant intracellular damage.

2.11. Development of resistance propensity

The development of bacterial resistance against conventional antibiotics has emerged as a serious global concern. Therefore, the need for antibacterial agents with sustained activity is paramount in clinical settings. In order to assess the potential of the compound as a long-term treatment option, the propensity for resistance development in MRSA (ATCC 33591) was studied and compared to that of fusidic acid over a period of 10 days (Fig. 4C). Fusidic acid is a widely used antibiotic for MRSA infections. The results revealed that the bacteria did not exhibit any detectable resistance against the compound even after continuous exposure for 10 days. In contrast, fusidic acid, the established antibiotic for MRSA, demonstrated a significant 32-fold increase in MIC after just 3 days of exposure. With further passaging, the MIC of fusidic acid continued to rise, reaching a 128-fold increase on the 5th day, and an astonishing 4000-fold increase after consecutive passaging for 10 days. These findings strongly indicate that the compound possesses a remarkable ability to impede the development of resistance in MRSA over an extended duration. In contrast, fusidic acid, a commonly used antibiotic, exhibited rapid resistance development within a short frame. The resistance profile of the compound suggests its suitability for prolonged treatment against MRSA infections, making it a promising candidate for combating antibiotic-resistant bacterial strains.

3. Conclusions

In conclusion, the manuscript investigated the antibacterial activity of a synthetic ion transporter and demonstrated its effectiveness against Gram-positive bacteria. When tested against drug-resistant bacteria MRSA, VRSA and VRE, the compound exhibited excellent antibacterial activity with fast killing kinetics. Notably, the ion transporter demonstrated very good biocompatibility and exhibited high selectivity towards the tested pathogens. Of particular significance is the ability of the compound to disrupt preformed biofilms of MRSA and reduce the bacterial burden within the biofilm structure. The anti-biofilm activity is noteworthy as very few synthetic ion transporters have been reported to possess this property. Furthermore, the amphiphilic nature of the molecule contributed to its multimodal bactericidal mechanism of action, including depolarisation of bacterial membrane potential, permeabilization of the cytoplasmic membrane, and generation of reactive oxygen species. Importantly, the membrane-active nature of the compound hindered the development of detectable resistance in bacteria for up to 10 days of serial passaging. This characteristic holds great promise for its use as a long-term therapeutic agent against bacterial infections. Overall, the excellent activity of the compound against planktonic bacterial cells, along with its ability to disrupt biofilms through a multimodal bactericidal action, positions it as a promising candidate for further development.

4. Experimental section

4.1. Materials and instrumentation

The following materials were purchased from Sigma-Aldrich: 3,3′-dipropylthiadicarbocyanine iodide [DiSC₃(5)], Measurement of optical density (O.D.) was done using the Tecan Infinite M200 PRO Microplate Reader. An instrument used for UV-visible investigations was a Jasco V-770 spectrophotometer. To quantify fluorescence for the biophysical studies, a Jasco FP-8500 spectrofluorometer was used. The MTCC (Chandigarh, India) provided the bacterial strains S. aureus MTCC 737, E. coli MTCC443, and. The sources of VRE 909 and VRSA 1, VRSA 4 were Anthem Bioscience, located in Bangalore, India. ATCC (Rockville, MD, USA) provided MRSA ATCC33591, E. faecium ATCC19634, and for our purchase. Mueller Hinton Broth was used to cultivate S. aureus, E. coli, MRSA, (MHB-HIMEDIA-M391). E. faecium and VRE were inoculated using brain heart infusion broth (BHI). MacConkey Agar and nutrient agar were utilized as solid medium for both Gram-positive and Gram-negative bacteria. 96 well plates, 6 well plates and transparent black 96 well plates were obtained from Vasa Scientific (Bangalore, India).

4.2. Antibacterial assay⁴⁰

Initially, bacteria from the frozen stock (at −80 °C) were streaked on a MacConkey agar plate for Gram-negative bacteria or nutrient broth for Gram-positive bacteria. After that, the streaked plates were incubated for bacterial growth for the entire night at 37 °C. After that, a single bacterial colony was inoculated for 6 h (mid-log phase) in 3 milliliters of nutrient broth, yielding, depending on the type of bacteria, 10⁸–10⁹ CFU mL⁻¹ cells. The culture that had grown for six h was diluted to ∼10⁵ CFU mL⁻¹ and used for the antibacterial assay. Using sterile Millipore water, compounds were serially diluted two times from their starting concentration in a 96-well plate. Following that, 20 μL of the aqueous solution of the test compound in each well were supplemented with 180 μL of a bacterial solution containing 10⁵ CFU mL⁻¹. After that, the plates were shaken and incubated for 16–18 h at 37 °C. The O.D. at 600 nm was obtained with a TECAN plate reader (Infinite series, M200 PRO). The experiment was run twice, with triplicates of each concentration, and the antibacterial activity (MIC) was assessed using visual turbidity.

4.3. Hemolytic activity⁴¹

The aqueous solution of the compound was diluted sequentially three times by a factor of two in a 96-well plate. After freshly drawn, heparinized human blood was centrifuged, and the supernatant was discarded in order to separate the red blood cells (hRBCs). Afterwards, 5 vol% of collected hRBCs were slowly suspended in 1× PBS (pH = 7.4). Subsequently, 150 μL of this suspension was put into each well of 96 well plates that contained 50 μL of the compound solution. The plate was then incubated for 1 h at 37 °C. After 5 min of centrifugation at 3500 rpm, the supernatant (100 μL) was transferred to a second 96-well plate so that a Tecan Infinite M200 PRO microplate reader could record the absorbance at 540 nm. In this investigation, a positive control was an identical volume of Triton X-100 (1 vol% solution in 1× PBS) and a negative control was an identical volume of 1× PBS without compound. The formula (A_treat − A_nontreat)/(A_{TX treat} − A_nontreat) × 100 was utilized to calculate the percentage of hemolysis. In this formula, A_treat denotes the absorbance of the compound-treated well, A_nontreat denotes the absorbance of the negative controls (without compound), and A_{TX treat} represents the absorbance of the Triton-X-100 treated well. Every concentration was measured in triplicate, and the average of the triplicate O.D. was used to calculate the HC₅₀.

4.4. Cytotoxicity through Alamar blue assay⁴⁰

Briefly 2 × 10⁴ MDCK cells per well were seeded in a 96 well plate for 24 h in complete DMEM media (DMEM + 10% FBS + 1% anti–anti). After 24 h, the cells were washed with 1× PBS and compound solution at different concentrations in complete media were added and incubated for 24 h. After 24 h, Alamar blue solution was added in 1× PBS and incubated for 6 h. After 6 h, absorbance was measured at 570 nm, with 600 nm as the reference wavelength. Here also, 1× PBS and Triton X-100 (1 vol% solution in 1× PBS) was taken as positive and negative control. The cell viability was determined following this formula: {1 − (A_treat − A_nontreat)/(A_{TX treat} − A_nontreat)} × 100. In this formula, A_treat denotes the absorbance of the compound-treated well, A_nontreat denotes the absorbance of the negative controls (without compound), and A_{TX treat} represents the absorbance of the Triton-X-100 treated well. Every concentration was measured in triplicate, and the average of the triplicate O.D. was used to calculate the EC₅₀.

4.5. Bactericidal kinetics⁴¹

To generate 10⁸–10⁹ CFU mL⁻¹ cells, a single colony of MRSA and VRSA was inoculated independently in nutrient broth for 6 h at 37 °C. Subsequently, 180 μL of the diluted bacterial solution in Mueller Hinton broth was added to the 20 μL aqueous solution of test compound T at varying concentrations of 2, 4, and 8 μg mL⁻¹. This mid-log phase bacterial solution was then further diluted to 5 × 10⁵ CFU mL⁻¹. As a control, the same volume of autoclaved water was used, but without the test compound. Next, at various time intervals (0 h, 1 h, 3 h, 6 h, and 12 h), 20 μL of aliquots from the individual mixture of bacteria and compound were serially diluted by a factor of ten in sterile saline. Afterwards, 20 μL of solution from each dilution was utilized for spot plating on agar plates, followed by a 24 h incubation period at 37 °C. After counting the number of bacterial colonies, the results were plotted against time on a logarithmic scale, or log (CFU mL⁻¹).

4.6. Biofilm disruption assay⁴⁰

Crystal violet staining

A study on biofilm disruption was carried out on 18 mm-diameter coverslips. Initially, a mid-log phase MRSA ATCC33591 culture grown for 6 h was diluted to 10⁵ CFU mL⁻¹ in nutritional broth supplemented with 1% NaCl and 1% glucose. Following the addition of this diluted bacterial suspension (2 mL per well) to the wells holding the sterilized coverslips, the plate was left to incubate at 37 °C under static conditions for a full day. Following the media removal, coverslips containing biofilm were carefully cleaned with 1× PBS (pH = 7.4) to get rid of any planktonic bacteria. The coverslips were then put into the well of a fresh 6-well plate. Following that, two solutions (2 mL) of T were added to the wells containing biofilm-coated coverslips and incubated for 24 h at two different concentrations [32 μg mL⁻¹ and 64 μg mL⁻¹]. 2 mL of fresh media devoid of compound was introduced to the well as an untreated control. Vancomycin at 32 and 64 μg mL⁻¹ served as the antibiotic control in this investigation. Following a 24 h period, the planktonic cells were removed from the coverslips using 1× PBS. Subsequently, every coverslip—both treated and untreated—was meticulously placed into an additional 6-well plate. These coverslips were incubated with 1 mL of 0.1% crystal violet (CV) dye in a 6-well plate for 10 min in order to observe biofilm disruption. Following a 1× PBS wash, the coverslip-associated crystal violet was dissolved in 95% ethanol, and the absorbance was measured at 520 nm. The absorbance of CV dye revealed how much biomass was still present on the coverslips.

Cell viability of bacteria embedded in biofilms

In this instance, T and antibiotics were used in accordance with the previously mentioned protocol to treat biofilm growth. Following the washing and plating of the biofilm-containing coverslips in a new well plate, 2 mL of a diluted trypsin–EDTA solution in saline (1 : 4 ratios) was added, and the mixture was shaken for 10 to 15 min. After serially diluting the bacterial suspension ten times, 20 μL of each dilution was spot-plated on nutrient agar plates. Next, the spotted plates were incubated for 18 h. The number of viable bacterial colonies was determined at the conclusion of the incubation period, and the results were reported as log (CFU mL⁻¹).

Confocal laser-scanning microscope (CLSM) of biofilms

For this experiment, T had been used, just like in the earlier research. After being cleaned with 1× PBS, the treated and untreated coverslips (which were previously explained in the biofilm disruption assay section) were put on glass slides. Using a 5 μL solution of Syto-9 (60 μM) and PI (15 μM) dye mixture, biofilm staining was carried out, and images were obtained using a Zeiss 510 Meta confocal laser-scanning microscope. The images were processed using Image J.

4.7. Cytoplasmic membrane depolarization assay⁴¹

MRSA cells in the mid-log phase (working concentration: 10⁸ CFU mL⁻¹) were isolated, centrifuged for 5 min at 3500 rpm, and then rinsed with a 1 : 1 mixture of 5 mM glucose and HEPES buffer (pH = 7.4). Next, a 1 : 1 : 1 mixture of 5 mM HEPES buffer, 100 mM KCl solution, 0.2 mM EDTA, and 5 mM glucose was added to the bacterial plate. In a Corning 96 black well plate with a clear bottom, 2 μM of 3,3′-dipropylthiadicarbocyanine iodide [DiSC₃(5)] and 190 μL of bacterial suspension were used for this investigation. The fluorescence intensity was measured at 622 nm excitation wavelength and 670 nm emission wavelength for 4 min following 60 min incubation of the plates. Next, 10 μL of T was combined with the dye and bacterial suspension at working concentrations of 4, 8, and 16 μg mL⁻¹ in each well. For this experiment, the control was the same volume of water without a compound. A Tecan Infinite M200 PRO microplate reader was used to measure the increase in fluorescence intensity for an additional 25 min.

4.8. Extent of membrane depolarisation through DiOC₂(3) dye

1 mL of ∼10⁸ CFU mL⁻¹ mid-log phase bacterial cells in 1× PBS were incubated in the presence of T (at 8 μg mL⁻¹) and CCCP at 15 μg mL⁻¹. After 4 h of incubation, compounds were removed through centrifugation, and bacterial cells were resuspended in saline. Then, the cells were incubated with the dye at a final concentration of 30 μM for 30 min and subjected to flow cytometry. Two controls were taken in this experiment. Bacteria without any dye and test compound was taken to check the healthy state of bacteria. Bacterial cells with dye was taken to confirm their polarised state. The dye was excited at 488 nm and emission was collected at both green and red channels FACSymphonyA5 (BD Biosciences). The data was analysed at FCS Express 7 software.

4.9. Bacterial live/dead assay⁴²

1 mL of 10⁸ CFU mL⁻¹ midlog phase MRSA suspension in 1× PBS was mixed with the compound T, and the mixture was incubated at 37 °C. The bacterial suspension was centrifuged to extract the compound entirely after 6 h of incubation. Subsequently, the bacterial palate was again suspended in regular saline, and 5 μL of a mixture containing 3 μM Syto-9 and 15 μM PI was added. The mixture was then incubated for 30 min. In addition, the bacterial suspension containing dye was centrifuged to eliminate any excess unbound dye, and the palate was reconstituted using 50 μL of regular saline. Lastly, a glass slide containing 5 μL of the bacterial suspension was prepared for 63× resolution confocal microscopy.

4.10. ROS generation⁴²

The assay was performed using 2′,7′-dichlorofluorescin diacetate (DCFH-DA) through flow cytometry. In summary, 8 and 16 μg mL⁻¹ of the compound were incubated with 1000 μL of 10⁸ CFU mL⁻¹ MRSA cells. After that, the bacterial suspension from each replicate was collected in a 1.5 mL microtube, and the biocide that had been released into the suspension was removed by centrifuging the mixture for 5 min at 3500 RPM. Following this, 1 mL of 1× PBS was used to resuspend the bacterial pellet, and 10 μM of DCFH-DA dye was added to the mixture. Following a 30 min dye incubation period, fluorescence was measured with excitation and emission at 485 nm and 530 nm, respectively.

4.11. Development of resistance propensity⁴³

The experiment was conducted using our previously published protocol.⁴⁰ The MIC of T and fusidic acid was assessed using the previously mentioned antibacterial assay protocol, to put it briefly. The bacterial solution from the sub-MIC concentration was used for another MIC determination in the days that followed, after being diluted to 10⁵ CFU mL⁻¹. The increase in MIC was plotted against the number of days following 10 days of serial passages. Here, each concentration was examined three times, and the difference between the MIC value of the first day and that of each subsequent day was used to calculate the fold increase in MIC.

Conflicts of interest

The authors declare no conflict of interest.