Synergistic Ionic Liquid-Antibiotic Combination as a Strategy against Methicillin-Resistant Staphylococcus aureus (MRSA)

Abstract

The accelerating spread of antimicrobial resistance, particularly in methicillin-resistant Staphylococcus aureus (MRSA), demands innovative strategies to restore antibiotic efficacy. Repurposing small molecules as antibiotic adjuvants offers a rapid and translationally feasible solution. Here, we investigated chloroquine-derived ionic liquids (CQILs) as potentiators of conventional antibiotics against MRSA. Although the ILs displayed limited intrinsic antibacterial activity, the top candidate markedly reduced minimum inhibitory concentrations (MICs) in combination, demonstrating strong synergistic interactions with doxycycline and kanamycin (FICI = 0.29 and 0.41, respectively) and an additive interaction with chloramphenicol (FICI = 0.80). Time-kill analyses confirmed accelerated and sustained bactericidal activity relative to monotherapy. Mechanistic studies revealed enhanced intracellular reactive oxygen species accumulation and pronounced disruption of bacterial membrane integrity, corroborated by scanning electron microscopy, which showed severe morphological damage. Importantly, combination treatments achieved effective bacterial eradication at reduced antibiotic doses while maintaining low cytotoxicity toward mammalian cells, thereby improving the therapeutic window. These findings establish chloroquine-derived ionic liquids as promising antibiotic adjuvants that potentiate bactericidal activity through oxidative stress amplification and membrane destabilization, offering a practical strategy to combat multidrug-resistant bacterial infections.

1. Introduction

Antimicrobial resistance (AMR) remains one of the most pressing global health challenges, contributing to millions of deaths annually as once-treatable bacterial infections become increasingly refractory to existing antibiotics. Among resistant pathogens, methicillin-resistant Staphylococcus aureus (MRSA) represents a particularly severe and persistent threat, accounting for a substantial proportion of hospital- and community-acquired infections worldwide. − In the United States alone, MRSA causes more than 70,000 severe infections and approximately 9000 deaths each year, while globally it remains a major contributor to the broader AMR burden, which is estimated to exceed 1.2 million deaths annually. The misuse and overprescription of antibiotics in agricultural settings significantly contribute to the emergence and spread of antimicrobial resistance (AMR). In the cattle industry, antibiotics are routinely used for disease prevention and treatment, creating sustained selective pressure for resistant bacterial populations. Because a substantial fraction of administered antibiotics is excreted unmetabolized, inadequate management of animal waste and agricultural runoff leads to the contamination of soil and water systems with active antimicrobial compounds. These environments function as reservoirs for resistance genes, facilitating their persistence and transmission to humans through contaminated water sources, food chains, and environmental exposure, ultimately undermining the effectiveness of clinically important antibiotics. ,, Despite this situation, the development of new antibacterial agents has progressed at an alarmingly slow pace. To address this growing therapeutic gap, drug repurposing and the identification of novel therapeutic applications for existing compounds have emerged as promising, cost-effective, and time-efficient strategies to accelerate the availability of effective antibacterial interventions. ,

A critical contributor to antibiotic failure in MRSA and other clinically significant pathogens is the structural and functional complexity of the bacterial cell envelope, which restricts drug penetration and promotes resistance. In Gram-negative bacteria, the multilayered cell envelope presents formidable permeability barriers, including an outer membrane enriched in lipopolysaccharides, restrictive porins, and active efflux systems, while periplasmic β-lactamases degrade antibiotics before they reach their intracellular targets. − Although MRSA, a Gram-positive organism that lacks an outer membrane, has a thick peptidoglycan cell wall, altered membrane composition, and resistance-associated modifications, such as changes in penicillin-binding proteins and increased peptidoglycan cross-linking, which significantly impair antibiotic efficacy. − Furthermore, the production of membrane vesicles carrying enzymes and resistance determinants can sequester or inactivate antimicrobial agents, further reducing therapeutic effectiveness. , These persistent and multifaceted resistance mechanisms underscore the need for alternative strategies to enhance antibiotic performance.

Recent therapeutic approaches to overcome MRSA resistance employ membrane-active adjuvants designed to enhance antibiotic intracellular accumulation. Venturicidin A, an ATP synthase inhibitor, rescues gentamicin activity against gentamicin-resistant MRSA clinical isolates by dissipating the proton-motive force (PMF), thereby facilitating aminoglycoside uptake despite enzymatic inactivation. The amphiphilic cyclic peptide [R₄W₄] synergizes with levofloxacin against multidrug-resistant S. aureus (ATCC BAA-1556, FICI = 0.498) through complementary membrane disruption and DNA-targeting mechanisms; however, this synergy weakens against the susceptible reference strain ATCC 29213 (FICI = 0.748, partial synergy only), suggesting that adjuvant efficacy is resistance-dependent. Small-molecule approaches are equally promising: quinoxaline-based compounds reduce fluoroquinolone MICs against MRSA biofilms and prevent resistance emergence over extended passaging. These precedents establish depolarization and membrane permeability enhancement as rational strategies for restoring conventional antibiotic efficacy against multidrug-resistant S. aureus. We hypothesized that a pharmaceutical salt formulation, chloroquine-undecanoate (CQUA), employing similar membrane-depolarizing principles could synergize with clinically relevant antibiotics against MRSA.

Ionic liquids (ILs) are a distinctive class of organic salts with melting points below 100 °C that have attracted increasing interest for their multifunctional roles in biological and pharmaceutical research. Their highly tunable physicochemical properties, arising from the modular combination of diverse cations and anions, enable broad applicability in enzyme stabilization, biopolymer processing, and enhanced drug solubility and delivery. ILs are further valued for their negligible vapor pressure, high thermal stability, and customizable solvation ability, allowing them to function both as solvents and as active pharmaceutical ingredients in the form of API-ionic liquids (API-ILs). By integrating the intrinsic bioactivity of conventional antibiotics with the adaptable properties of ILs, API-ILs have been shown to improve drug solubility, stability, and bioavailability, thereby enhancing antimicrobial efficacy, particularly against resistant bacterial strains. , Originally, the biological activity and toxicity of ILs are strongly governed by structure–activity relationships, notably the length and composition of the cationic alkyl chain. Short-chain ILs tend to form stable nanoaggregates and are generally associated with reduced cytotoxicity and improved oral bioavailability, whereas long-chain ILs preferentially localize to mitochondria, inducing mitochondrial dysfunction, mitophagy, and apoptosis, thereby contributing to increased toxicity. These insights have guided the rational design of IL-based systems with optimized safety and efficacy profiles, positioning ILs as a promising platform for enhancing antimicrobial performance and improving drug delivery strategies against resistant pathogens.

Chloroquine, originally developed as an antimalarial agent, exemplifies the potential of drug repurposing as a strategy to expand therapeutic utility beyond initial indications. Although its clinical use has been limited by well-documented toxicities, including gastrointestinal disturbances, retinopathy, and cardiotoxicity, structural modification led to the development of hydroxychloroquine, which exhibits an improved safety and tolerability profile. As a result, hydroxychloroquine has been successfully repurposed for the long-term management of autoimmune disorders such as systemic lupus erythematosus and rheumatoid arthritis. The improved tolerability profile of hydroxychloroquine relative to chloroquine underscores the importance of structural optimization in expanding therapeutic applicability while mitigating adverse effects.

Considering the persistent clinical burden of MRSA infections and the limitations of existing antibiotic therapies, there is a critical need for antimicrobial strategies that address both resistance and drug delivery challenges. Ionic liquids (ILs) offer a versatile and rationally designable platform capable of overcoming these barriers through tunable ion composition, enhanced solubility, and membrane-active mechanisms. By facilitating improved bacterial membrane permeability and intracellular antimicrobial uptake, IL-based systems have the potential to circumvent conventional resistance pathways and enhance therapeutic efficacy against MRSA. Our group has previously demonstrated that anion manipulation within choline carboxylic acid–based ionic liquid frameworks modulates antimicrobial potency and biocompatibility, establishing key structure–activity relationships that informed the rational design of the present compound series. Building on this foundation, the present study reports the synthesis and antimicrobial screening of a chloroquine-carboxylic acid ionic liquid library against MRSA, identification of chloroquine-undecanoate (CQUA) as the lead candidate, evaluation of synergistic combinations with mechanistically distinct conventional antibiotics, mechanistic characterization through membrane-targeted assays including depolarization, permeabilization, and fluidity measurements, and assessment of mammalian cell biocompatibility to establish a therapeutic window for combination use.

2. Results and Discussion

2.1. Preparation of Ionic Liquids

A large library of chloroquine-carboxylic acid–based ionic liquids (CQILs) was synthesized by combining the cation and anion in a 1:1 molar ratio through an acid–base neutralization reaction (Figure ). In this process, carboxylic acid was added dropwise to the anion while stirring. The mixture was stirred for 3 h, after which it was transferred to a rotary evaporator to remove any excess water. NMR spectroscopy confirmed the expected proton peaks and integrations, indicating the successful synthesis of the ILs, with the mole ratios and identities matching the anticipated structure. Detailed characterizations and anion structures are provided in the Supporting Information.

1.

General structure of chloroquine-carboxylate ionic liquid. The series was designed to investigate the effects of alkyl chain length and degree of saturation on anti-MRSA activity.

2.2. Minimum Inhibitory Concentration (MIC)

The antimicrobial properties of the synthesized ionic liquids were assessed against methicillin-resistant S. aureus (MRSA) using the standard broth microdilution assay in accordance with CLSI guidelines. It should be noted that standard CLSI susceptibility breakpoints (susceptible, intermediate, resistant) are defined for conventional antibiotics and do not apply to ionic liquid compounds, for which no clinical breakpoints have been established; MIC values for the CQILs are therefore reported as absolute inhibitory concentrations. The reference antibiotic MIC values reported here (CHR: 0.039 mM; DOX: 0.0077 mM; KAN: 0.3125 mM) are consistent with established susceptibility ranges, confirming assay validity. Clinically relevant antibiotics, including chloramphenicol (CHR), doxycycline (DOX), and kanamycin (KAN), served as reference controls. As summarized in Table , the parent chloroquine molecule exhibited minimal inhibitory activity against MRSA, with MIC values exceeding 1 mM. In contrast, incorporation of carboxylic acid moieties into the ionic liquid framework markedly enhanced antimicrobial potency. A clear trend emerged showing that longer alkyl chain substituents corresponded with improved antibacterial activity. Notably, chloroquine-undecanoate (CQUA) demonstrated the strongest effect against MRSA (MIC = 0.8 mM). Studies have shown that longer alkyl chains in ionic liquids result in lower MICs against both Escherichia coli and MRSA, with significant reductions observed as the chain length increases in both choline-carboxylic acid and cationic ionic liquids. The increased antibacterial activity observed with longer alkyl-chain ionic liquids arises from enhanced hydrophobic interactions with bacterial membranes, which promote membrane insertion and destabilization. The incorporation of fatty acid-derived anions further increases amphiphilicity, facilitating efficient partitioning into lipid-rich bacterial envelopes. − Importantly, the ionic-liquid form is essential for activity, as neither individual components (CQ MIC = 5 mM, UDA, MIC = 1.56 mM) nor choline alone, nor a simple physical mixture (Table S2, Figure S3), exhibit comparable antibacterial effects, indicating that ion pairing is required to achieve effective membrane disruption. These findings emphasize the potential of this class of ionic liquids as effective agents for targeting both Gram-negative and Gram-positive bacterial pathogens.

1. Antimicrobial Susceptibility of CQILS against MRSA.

treatment	MIC (mM)
chloroquine (CQ)	5
chloroquine phosphate (CQPO4)	>5
chloroquine-butanoate (CQBA)	5
chloroquine-pentanoate (CQPA)	5
chloroquine-hexanoate (CQHA)	5
chloroquine-heptanoate (CQHP)	2.5
chloroquine-octanoate (CQOA)	5
chloroquine-nonanoate (CQNA)	2.5
chloroquine-decanoate (CQDA)	2.5
chloroquine-undecanoate (CQUA)	0.8
chloroquine-dodecanoate (CQDDA)	0.9
chloroquine-2butenoate (CQ2BE)	2.5
chloroquine-3butanoate (CQ3BE)	5
chloroquine-2pentenoate (CQ2PE)	5
chloroquine-2hexenoate (CQ2HE)	2.5
chloroquine-3hexenoate (CQ3HE)	2.5
chloroquine-2octenoate (CQ2OE)	2.5
chloroquine-2nonanoate (CQ2NE)	2.5
chloroquine-3nonenoate (CQ3NE)	2.5
chloroquine-2decenoate (CQ2DE)	1.25
chloramphenicol (CHR)	0.039
doxycycline hyclate (DOX)	0.0077
kanamycin sulfate (KAN)	0.3125

2.3. Checkerboard Synergy Assay

Checkerboard assays revealed distinct combination-dependent interactions between CQUA and conventional antibiotics against MRSA (Table S1). The combination of CQUA with chloramphenicol showed an additive interaction (FICI = 0.80), whereas strong synergism was observed with doxycycline (FICI = 0.41) and kanamycin (FICI = 0.23). The exact CQUA and antibiotic concentrations at the combination MIC were as follows: CQUA-CHR (0.0625 mM/0.029 mM), CQUA-DOX (0.0312 mM/0.0019 mM), and CQUA-KAN (0.125 mM/0.078 mM). These synergistic combinations were associated with pronounced reductions in antibiotic MICs relative to monotherapy (Figure ), indicating substantial enhancement of antibacterial efficacy. Overall, CQUA consistently potentiated antibiotic activity against MRSA.

2.

Interactions between CQUA and conventional antibiotics against MRSA. Checkerboard heatmap showing bacterial growth optical density (OD) following treatment with CQUA in combination with CHR, DOX, and KAN (A–C). The color scale represents OD values: lighter colors indicate lower OD (greater growth inhibition) and darker colors indicate higher OD (bacterial growth). Synergy was quantitatively determined by calculating the fractional inhibitory concentration index (FICI) from the MIC of each agent in combination versus alone; the heatmap provides visual context for growth patterns across the concentration matrix. Calculated FICI values were 0.80 (additive interaction), 0.29, and 0.23 (synergy) for CQUA-CHR, CQUA-DOX, and CQUA-KAN, respectively.

These synergistic interactions align with documented precedent for conventional antibiotic combinations against MRSA. Yu et al. evaluated 55 pairwise antibiotic combinations against 113 MRSA clinical isolates and model strains (COL) using a checkerboard assay and validated synergistic interactions in a murine neutropenic thigh infection model. The study identified vancomycin + oxacillin and vancomycin + rifampicin as synergistic (FICI ≤0.5), achieving in vitro MIC reductions of 50- to 160-fold and demonstrating clinical efficacy in vivo. Our CQUA + kanamycin (FICI 0.41) and CQUA + doxycycline (FICI 0.29) results are consistent with this aminoglycoside and fluoroquinolone synergy pattern. The additive interaction observed with chloramphenicol (FICI 0.80) reflects known reduced synergy between membrane-active agents and β-lactam-class inhibitors, as documented with polymeric adjuvants (HBPL) and peptide adjuvants (T3/T4) paired with ampicillin or oxacillin against MRSA. This antibiotic-class-dependent pattern suggests that depolarization-based enhancement is optimized for uptake-dependent drugs (aminoglycosides, tetracyclines) rather than those relying primarily on transpeptidase inhibition.

The antibiotic selection in this study was guided by a drug repurposing framework: CHR, DOX, and KAN represent mechanistically distinct classes-a 50S ribosomal inhibitor, a 30S ribosomal inhibitor, and an aminoglycoside, respectively-whose clinical utility against MRSA has been significantly curtailed by resistance mechanisms including reduced intracellular accumulation and efflux. Reviving these deprioritized antibiotics through adjuvant combination strategies is a scientifically important goal complementary to new antibiotic discovery, which is consistent with our group’s prior work demonstrating that anion manipulation in ionic liquid frameworks modulates antimicrobial activity. The mechanistic basis for these interactions became apparent from the membrane-targeted assays conducted in this study, where CQUA-mediated membrane depolarization and permeabilization are expected to directly enhance intracellular accumulation of KAN, whose uptake is driven by the proton motive force, and to facilitate access of protein synthesis inhibitors. We acknowledge that this mechanistic interpretation emerged from the experimental data rather than constituting a prospective hypothesis. Future work will evaluate CQUA in combination with current first-line MRSA agents, including vancomycin and daptomycin, to assess broader translational potential. As a weak base, CQ can disrupt pH gradients and interfere with bacterial stress-response and efflux mechanisms, thereby sensitizing cells to ribosomal inhibitors (DOX, CHR) and aminoglycosides (KAN), whose activity depends on efficient intracellular accumulation. −

2.4. Growth Curve

Bacterial growth kinetics over 24 h revealed concentration-dependent inhibition by CQUA, and the reference antibiotics CHR, DOX, and KAN (Figure ). CQUA effectively suppressed exponential growth at ≥ 1× MIC, indicating inhibition of proliferation rather than rapid cell lysis. CHR and DOX produced pronounced growth suppression at comparable concentrations, while KAN exhibited partial growth recovery at sub-MIC levels, consistent with its concentration-dependent activity. The flat, low OD profiles observed in Figure C (CQUA-DOX combination) and Figure G (DOX monotherapy) across all tested concentrations reflect the exceptionally potent and rapid bacteriostatic activity of doxycycline against this MRSA strain, which exhibits an MIC of 0.0077 mM -among the lowest in this data set. Doxycycline binds the 30S ribosomal subunit with high affinity, halting bacterial protein synthesis rapidly upon exposure. At concentrations as low as 0.5× MIC, sufficient intracellular doxycycline accumulation appears to occur to achieve near-complete bacteriostasis from the earliest time points, with no measurable regrowth over 24 h. The low but nonzero OD values confirm bacterial presence without proliferation, consistent with bacteriostasis rather than sterilization. Critically, this pattern was observed in both DOX monotherapy and CQUA-DOX combination, confirming that the effect is driven primarily by doxycycline’s intrinsic potency. These results were reproducible across all three independent replicates, and background subtraction was applied to exclude OD contributions from compounds or media. Across all treatments, growth curves plateaued rather than declined, suggesting predominantly bacteriostatic or nonlytic mechanisms of action. These findings highlight CQUA as a potent growth-inhibitory agent with activity comparable to established antibiotics and support its potential utility against MRSA.

3.

CQUA and its antibiotic combinations enhance the growth inhibition of MRSA compared to parent antibiotics. Growth kinetics monitored over 24 h following treatment with (A) CQUA, (B) CQUA-CHR, (C) CQUA-DOX, (D) CQUA-KAN, (E) representative resazurin microdilution assay following 24 h exposure. Blue wells indicate growth inhibition, and pink wells indicate metabolically active cells (F) CHR, (G) DOX, and (H) KAN. Panel A was evaluated at 0.5×, 1×, 2×, and 4× MIC; in panel A, the 4× MIC curve overlaps substantially with the 2× MIC curve, reflecting a growth inhibitory ceiling effect at those concentrations. Panels B–D and F–H were evaluated at 0.5×, 1×, and 2× MIC only. In panels B–D, MIC multiples refer to both components simultaneously. Actual combination concentrations: CQUA-CHR (0.0625 mM CQUA/0.029 mM CHR); CQUA-DOX (0.0312 mM CQUA/0.0019 mM DOX); CQUA-KAN (0.125 mM CQUA/0.078 mM KAN), while monotherapy concentration of CQUA­(MIC-0.8 mM), CHR (MIC-0.039 mM), DOX (0.0077 mM), KAN (MIC-0.3125 mM). Bacterial growth was measured as optical density at 630 nm (OD₆₃₀). Ctrl denotes untreated cultures grown under identical conditions. Data represent mean ± SD of (3) independent experiments.

2.5. Time-Kill Kinetics

Time-kill assays were conducted to quantitatively compare the antibacterial activity of the test compound administered as monotherapy and in combination regimens over a 24 h period. Bacterial viability was expressed as log₁₀ CFU mL^–1, with untreated cultures serving as growth controls. As a single agent, CQUA exhibited a clear concentration-dependent response but demonstrated limited bactericidal activity at clinically relevant exposures. Sub-MIC and MIC concentrations resulted in minimal suppression of bacterial growth relative to controls (Figure A). In contrast, sustained bactericidal activity was observed only at 2× and 4 × MIC, indicating that elevated concentrations are required to achieve meaningful antibacterial efficacy.

4.

CQUA and antibiotic formulations enhance bactericidal activity. (A–G) Time-kill kinetics of MRSA over 24 h following treatment with (A) CQUA, (B) CQUA-CHR, (C) CQUA-DOX, (D) CQUA-KAN, (E) MBC, (F) CHR, (G) DOX, and (H) KAN at 0.5×, 1×, 2×, and 4 × MIC. Bacterial burden was quantified as log₁₀(CFU mL^–1). Ctrl denotes untreated cultures. Data represent mean ± SEM (n = 3).

For chloramphenicol and doxycycline (Figure F,G), the enhanced activity observed in time-kill experiments was consistent with checkerboard analyses, which revealed additive interaction (FICI = 0.80) and strong (FICI = 0.29) synergy, respectively. In both cases, combination therapy converted predominantly bacteriostatic monotherapy profiles into sustained, time-dependent reductions in bacterial burden (Figure B,C), supporting the functional relevance of the FICI-based interaction metrics. In contrast, the CQUA-KAN (Figure D) demonstrated a clear divergence between static and dynamic interaction assessments. Although checkerboard analysis indicated strong synergy (FICI = 0.41), time-kill studies revealed attenuation of kanamycin-mediated bactericidal activity in the presence of CQUA, with a loss of ≥3 log₁₀ CFU mL^–1 killing at both 1× and 2× MIC. This discrepancy highlights fundamental differences between checkerboard assays, which measure end point growth inhibition, and time-kill analyses, which capture concentration- and time-dependent killing kinetics as well as regrowth behavior. Such discordance between FICI-based synergy and time-kill outcomes has been widely reported , and reflects the inability of checkerboard methods to account for temporal pharmacodynamic effects, inoculum-dependent activity, and mechanistic drug–drug interactions. The biphasic kill profile observed in KAN monotherapy (Figure G), characterized by initial growth suppression between 0 and 12 h followed by regrowth after 12 h, is consistent with adaptive aminoglycoside resistance. , At suboptimal concentrations, a surviving bacterial subpopulation can downregulate electron transport chain activity, thereby reducing the membrane potential that drives electrogenic KAN uptake, resulting in diminished intracellular accumulation and consequent regrowth. This adaptive mechanism provides a mechanistic basis for the CQUA-KAN synergy observed by FICI: CQUA-mediated membrane depolarization would be expected to counteract this adaptive resistance by sustaining membrane potential-dependent KAN uptake, sensitizing bacteria to kanamycin even under conditions that would otherwise permit adaptive resistance. The variability observed in the kanamycin time-kill profile highlights the limitations of monotherapy and further supports the rationale for combination strategies to achieve more consistent bactericidal activity.

2.6. Scanning Electron Microscopy

Scanning electron microscopy (SEM) revealed treatment-dependent alterations in bacterial surface organization across treatment with CQUA. The control sample (Figure A) exhibited a dense bacterial population characterized by associated cells with minimal intercellular spacing, forming compact clusters that extensively covered the surface. Upon partial treatment (Figure B), a noticeable reduction in clustering was observed, with increased intercellular spacing and partial exposure of the underlying substrate. At MIC (Figure C), bacterial distribution appeared further dispersed, with fewer compact aggregates and decreased overall surface coverage relative to the control. At 4 × MIC (Figure D), no intact bacterial cells were observed within the imaged field, and the surface appeared largely devoid of cellular structures. These findings demonstrate a concentration-dependent reduction in bacterial clustering and surface coverage with increasing treatment intensity. These SEM results support the time-kill data, indicating a concentration-dependent action in which ILs initially interact with and weaken the bacterial membrane, then cause irreversible damage and cell lysis at higher doses.

5.

Concentration-dependent morphological disruption of Staphylococcus aureus following 24 h treatment with CQUA. SEM images of S. aureus treated with CQUA at (a) untreated control, (b) 0.5 × MIC, (c) 1× MIC, and (d) 4× MIC. Progressive structural damage is observed with increasing concentration, and no intact cells are visible at 4× MIC.

2.7. Membrane Permeabilization Assay

The membrane-permeabilizing activity of the ionic liquids (ILs) was evaluated using Sytox green uptake in MRSA. Across all tested IL concentrations (0.5× MIC to 4× MIC), fluorescence intensity remained low and comparable to that observed for the melittin-treated control (Figure S4), indicating limited Sytox Green penetration. In Gram-positive bacteria, the presence of a thick peptidoglycan layer restricts dye entry; therefore, low fluorescence intensity does not necessarily reflect an absence of antimicrobial activity. − Instead, these results suggest that IL-mediated bacterial killing may occur predominantly through nonlytic mechanisms rather than extensive membrane disruption.

In contrast to the untreated control, all CQUA-antibiotic combination treatments exhibited greater temporal changes in Sytox Green fluorescence intensity (Figure S4). The untreated control maintained low, stable fluorescence throughout the assay, consistent with intact membrane integrity and restricted dye uptake. However, combinations with chloramphenicol, doxycycline, or kanamycin showed a pronounced fluorescence response over time, characterized by an initial increase followed by a progressive decline in signal intensity. This dynamic fluorescence pattern is indicative of membrane perturbation, transient dye entry, and subsequent leakage or loss of intracellular nucleic acid content. , Mechanistically, these findings suggest that CQUA treatment may act by compromising bacterial membrane integrity, thereby facilitating increased permeability and enhancing intracellular access of antibiotics, particularly those targeting intracellular processes. The observed membrane destabilization provides a plausible mechanistic basis for the synergistic antibacterial activity seen with synergy combinations, exceeding the effects observed with either treatment alone.

2.8. Membrane Depolarization Assay

To investigate whether CQUA and its synergistic combinations exert antibacterial activity through membrane perturbation, membrane depolarization was assessed using the voltage-sensitive dye DiSC₃(5). Changes in fluorescence intensity reflect disruption of the bacterial cytoplasmic membrane potential. Treatment with CQUA alone induced a concentration-dependent increase in DiSC₃(5) fluorescence (Figure A), indicating progressive membrane depolarization. At subinhibitory concentrations (0.5× and 1× MIC), CQUA caused modest but detectable increases in fluorescence relative to the untreated control, suggesting early perturbation of membrane potential without complete membrane collapse. In contrast, higher concentrations (2× and 4 × MIC) resulted in a more pronounced and sustained increase in fluorescence, consistent with significant disruption of membrane integrity. The membrane-active peptide melittin (32 μg/mL), used as a positive control, produced rapid and extensive depolarization. Treatment with the antibiotics alone (CHR, DOX, and KAN) did not induce appreciable membrane depolarization over the time course examined (Figure C), consistent with their established intracellular mechanisms of action and lack of direct membrane targeting. , Importantly, CQUA-antibiotic combinations exhibited minimal additional depolarization compared to CQUA alone at synergistic concentrations (Figure B). This observation suggests that synergy is not driven by additive membrane disruption but rather by CQUA-mediated membrane perturbation that facilitates enhanced antibiotic uptake and intracellular activity. Such a mechanism aligns with the observed reductions in MIC values and enhanced bactericidal activity in time-kill assays. ,,

6.

CQUA induces membrane depolarization in Staphylococcus aureus. DiSC₃(5) fluorescence assay evaluating membrane potential. (A) Fluorescence response following treatment with the CQUA at 0.5×–4× minimum inhibitory concentration (MIC). (B) Combination treatments at synergistic concentrations: CQUA-CHR (0.0625 mM CQUA + 0.029 mM CHR); CQUA-DOX (0.0312 mM CQUA + 0.0019 mM DOX); CQUA-KAN (0.125 mM CQUA + 0.078 mM KAN). (C) Individual antibiotics at their respective MIC values: CHR (0.039 mM), DOX (0.0077 mM), KAN (0.3125 mM). Melittin (32 μg mL^–1) served as the positive control. Increased fluorescence indicates membrane depolarization.

Depolarization-mediated enhancement of antibiotic efficacy is increasingly recognized as a conserved principle in MRSA adjuvant design. Venturicidin A, an ATP synthase inhibitor, achieves rescue of gentamicin in resistant MRSA by directly dissipating the PMF, validating proton-motive force disruption as a sufficient mechanism for aminoglycoside potentiation. Synthetic amphiphile scaffolds, including honokiol/magnolol derivatives, fluoro-lipopeptides (R6F), and cationic polymers (HBPL), similarly achieve synergistic enhancement of aminoglycosides or fluoroquinolones through membrane permeabilization and depolarization. The mechanism of piceatannol, a natural product identified in recent mechanistic surveys, further corroborates this principle: working via PMF dissipation and inhibition of ATP synthesis to enhance antimicrobial effects. Our pharmaceutical salt formulation strategy, in which the undecanoate anion and chloroquine cation jointly contribute to membrane disruption and depolarization, implements these validated design principles within a biocompatible formulation. This positioning suggests that CQUA represents a rational translation of depolarization-based synergy into a therapeutic-grade adjuvant for MRSA infections. Collectively, these results demonstrate that CQUA may function as a membrane-active potentiator rather than a nonspecific membrane-lytic agent. Partial membrane depolarization at sub-MIC levels appears sufficient to sensitize bacteria to antibiotics without inducing extensive membrane damage, providing a mechanistic basis for the observed synergy and improved therapeutic window. This dual behavior, -membrane targeting, coupled with enhanced intracellular antibiotic efficacy, distinguishes the IL from classical membrane-disrupting antimicrobials and supports its potential utility in combination antimicrobial therapy.

2.9. ROS Detection

In MRSA (Figure A), monotherapy induces a significant increase in total intracellular ROS, as indicated by the CQUA signal. However, signals corresponding to individual ROS species (superoxide, hydrogen peroxide, and hydroxyl radicals) were insignificant, preventing definitive identification of the dominant reactive species (Figure B–D). These data indicate that oxidative stress is present under treatment conditions, but that ROS generation is heterogeneous and not attributable to a single, specific radical.

7.

Intracellular reactive oxygen species (ROS) were generated in S. aureus after treatment with CQUA for 30 min. (A) Total ROS, (B) superoxide (O₂•^–), (C) hydroxyl radical (•OH), and (D) hydrogen peroxide (H₂O₂) levels following treatment with the test compound at 1/2×–4× MIC, compared with control and H₂O₂ (500 μM, positive control). (E) ROS levels for combination (synergistic) treatments with antibiotics. (F) ROS levels induced by individual antibiotics. Fluorescence intensity is reported in arbitrary units (AU). Statistical significance is indicated as ns (not significant), p < 0.05, **p < 0.001, and ***p < 0.0001.

Chloramphenicol, doxycycline, and kanamycin primarily target bacterial ribosomes and do not directly induce ROS-mediated cytotoxicity (Figure F). The slightly higher ROS signal observed for CQUA-kanamycin combinations (Figure E) may be due to enhanced metabolic and respiratory stress induced by treatment, rather than by ROS-dependent killing. Nevertheless, antibiotic exposure, particularly with resistant organisms such as MRSA, can disrupt cellular homeostasis, including membrane integrity, central metabolism, and electron transport processes, resulting in nonspecific ROS accumulation. The elevated total ROS observed likely reflects a generalized stress response rather than a defined oxidative killing pathway. ,

The lack of concordance among individual ROS species likely reflects the robust antioxidant and redox-buffering capacity of MRSA. S. aureus expresses catalase, superoxide dismutase, and the carotenoid pigment staphyloxanthin, which collectively neutralize and redistribute oxidative stress, dampening signals from individual ROS species while allowing an overall increase in oxidative burden. −

2.10. Membrane Fluidity

Laurdan GP measurements were used to assess changes in membrane order and fluidity in MRSA following treatment with CQUA, antibiotics, and their combinations. Higher GP values correspond to increased membrane order (reduced hydration), whereas lower GP values indicate increased membrane fluidity and disorder. Laurdan GP measurements (Figure A) show significantly increased membrane order in MRSA. GP values rose sharply from the low levels observed in untreated controls to approximately 0.75–0.80 at concentrations ranging from 1/2 × MIC to 4 × MIC, indicating pronounced membrane rigidification. Melittin, used as a positive control, produced only a moderate increase in GP, consistent with membrane disorder and pore formation rather than rigidification. The ability of CQUA to strongly perturb the packing of bacterial membranes, causing rigidity, likely contributes to its antibacterial activity.

8.

CQUA and synergy combinations induce membrane rigidity in S. aureus. Membrane fluidity assessed by Laurdan generalized polarization (GP). (A) GP values following treatment with the test compound at 0.5×–4× minimum inhibitory concentration (MIC) compared with untreated control and melittin (32 μg mL^–1). (B) Individual antibiotics induce GP changes. (C) GP values for combination treatments. Increased GP values indicate reduced membrane fluidity (membrane rigidity). Data are presented as mean ± SEM. Statistical significance is indicated as ns (not significant), p < 0.05, *p < 0.01, **p < 0.001, and ***p < 0.0001.

Treatment with antibiotics alone resulted in modest and variable changes in GP compared to the control (Figure B). Chloramphenicol and doxycycline slightly reduced GP values, consistent with minimal direct effects on membrane structure. Kanamycin showed no significant difference from the control. These results indicate that the tested antibiotics do not substantially perturb membrane order when used as monotherapy. Synergy combinations produced significantly higher GP values than control and antibiotic-only treatments (Figure C). The increased GP observed for CQUA combined with chloramphenicol, doxycycline, or kanamycin indicates enhanced membrane ordering and reduced lipid mobility. Mechanistically, this suggests that CQUA induces membrane rigidification and alters lipid packing, which may compromise membrane-associated processes and increase susceptibility to antibiotics. Such membrane biophysical changes are consistent with a nonlytic mechanism in which altered membrane organization facilitates antibiotic uptake or sensitizes intracellular targets, thereby contributing to the observed synergistic antibacterial activity. −

2.11. Biocompatibility Studies

The biocompatibility of CQUA was assessed through hemolysis and cytotoxicity assays to evaluate its safety relative to its antimicrobial potency. As shown in the hemolysis profile (Figure A), the IL caused minimal membrane disruption at lower concentrations (0.1–0.3 mM), with hemolysis remaining below 10% at MIC values and increasing in a concentration-dependent manner, becoming more pronounced above 1.3 mM. However, the IL remained largely nonhemolytic within their antimicrobial concentration ranges. A similar trend was observed in the HEK293 cytotoxicity assay, where cell viability exceeded 85–95% at 0.1 mM, then declined progressively with increasing concentration, falling to 40–60% at 0.3 mM and dropping sharply at concentrations ≥0.6 mM, where viability approached baseline levels. Studies have shown that longer-chain fatty acids are more effective at disrupting membrane integrity and causing cell lysis compared to short-chain fatty acids. ,

9.

Hemolytic activity and mammalian cell viability of CQUA and antibiotic treatments. (A) Concentration-dependent hemolysis induced by CQUA, with the minimum inhibitory concentration (MIC) indicated. (B) Hemolysis caused by individual antibiotics. (C) Hemolytic effects of synergistic combinations. (D) Concentration-dependent effects of CQUA on mammalian cell viability. (E) Cell viability following synergistic combination treatments. Data are presented as mean ± SEM.

Despite this concentration-dependent cytotoxicity, the IL maintained low cell viability and low hemolysis near their MIC values, indicating an unfavorable balance between antibacterial activity and mammalian cell tolerance. In comparison, the synergy combination exhibited even lower hemolytic activity and reduced cytotoxicity, further supporting its potential for safer therapeutic applications. Also, the combined approach minimized disruption to healthy cells, highlighting its reduced adverse effects. Collectively, these results demonstrate that CQUA exhibits selective antibacterial activity at a concentration mildly tolerated by mammalian cells and synergistic combinations, retain their efficacy while minimizing damage to mammalian cells, establishing a promising option for MRSA treatment.

3. Conclusion

The rising clinical burden of MRSA infections, set against a backdrop of declining antibiotic discovery, highlights the need for strategies that extend the useful life of existing drugs rather than replace them entirely. This study demonstrates that chloroquine-undecanoate (CQUA), a chloroquine-derived ionic liquid, can function as a membrane-active adjuvant that restores the antibacterial efficacy of conventional antibiotics rendered ineffective against MRSA by resistance. The key finding is mechanistic: CQUA disrupts the bacterial membrane through depolarization, permeabilization, and rigidification. These effects undermine the membrane integrity that MRSA depends on to exclude or expel antibiotics, thereby sensitizing the bacterium to ribosomal inhibitors and aminoglycosides that would otherwise fail to accumulate intracellularly at therapeutic levels. Critically, this synergistic activity is achieved at CQUA concentrations of 0.0312–0.125 mM, well within the range tolerated by mammalian cells, which resolves the cytotoxicity limitation observed with CQUA monotherapy and establishes a viable therapeutic window for combination use. Beyond the specific activity of CQUA, these findings offer a broader proof-of-concept: that the ionic liquid framework, through rational ion-pair design, can confer membrane-active adjuvant properties onto an existing pharmacophore. This represents a structurally tunable and translationally practical platform for antibiotic potentiation. Future studies will extend this approach to first-line MRSA agents and relevant infection models to evaluate its full clinical potential.

4. Methods

4.1. Materials and Reagents

Chloroquine free base (1PlusChem), dimethyl sulfoxide (DMSO), and all commercially available carboxylic acids were purchased from Sigma-Aldrich (St. Louis, MO). Cation-adjusted Mueller–Hinton broth and Dey-Engley neutralizing broth were also obtained from Sigma-Aldrich. Tryptone, yeast extract, agar, and sodium chloride were obtained from Fisher Scientific (Hampton, NH). Fluorescent probes, including H2DCFDA, HKSOX-1, HKOH-1r, and HKPerox-2, were purchased from MedChemExpress (Monmouth Junction, NJ). Melittin was acquired from Selleck Chemicals. Laurdan dye was obtained from TargetMol. SYTOX Green and DISC₃(5) were purchased from Thermo Fisher Scientific. Phosphate-buffered saline (PBS) was obtained from Gibco (Thermo Fisher Scientific), and HEPES buffer was purchased from Cytiva (Marlborough, MA). CellTiter-Glo Luminescent Cell Viability Assay Kit was purchased from Promega Corporation. Bacterial cell viability was determined using the Resazurin Cell Viability Kit (Cell Signaling Technology, Danvers, MA, USA).

4.2. Synthesis and Characterization of Ionic Liquids

Chloroquine free base and the selected carboxylic acids were combined in a 1:1 molar ratio to generate the corresponding ionic liquids via acid–base neutralization reaction with minor modifications. Briefly, chloroquine free base and carboxylic acid were added to a single reaction vessel and subjected to one-pot synthesis at 45 °C with continuous stirring for 3 h in an oil bath. After completion of the reaction, excess water was removed by rotary evaporation at 45 °C under 15 mbar for 2 h. Residual water content in the resulting ionic liquids was quantified using Karl Fischer coulometric titration. Structural confirmation and purity assessment were performed by ¹H nuclear magnetic resonance (NMR) spectroscopy. Complete NMR characterization for all ionic liquids used in this study is provided in the Supporting Information.

4.3. Bacterial Strains and Culture Conditions

Methicillin-resistant S. aureus (MRSA [BAA-1708]) cells were obtained from ATCC (Manassas, VA, USA). The cells were retrieved from frozen stock, streaked onto LB-methicillin agar to isolate resistant colonies with a sterile loop, and then incubated overnight at 37 °C in a static incubator. Confirmed resistant colonies were subsequently cultured in LB-methicillin broth at 37 °C for 18 h with shaking at 150 rpm. For all minimum inhibitory concentration (MIC) assays, bacterial strains were grown in cation-adjusted Muller–Hinton (CAMHB) broth according to standard susceptibility testing guidelines.

4.4. Minimum Inhibitory Concentration (MIC) Determination

Following the Clinical and Laboratory Standards Institute (CLSI) guidelines, the minimum inhibitory concentrations (MICs) of the tested ionic liquids (ILs) were determined using the standard broth microdilution method. Stock solutions of each IL were prepared at 30 and 100 mg/mL in dimethyl sulfoxide (DMSO); molar stock concentrations for each compound, calculated from their respective molecular weights, are provided in Table S3. MIC determination was performed through several iterative rounds of screening with progressively refined concentration ranges, using the appropriate stock as required, until the MIC was reproducibly determined for each compound. Stocks were diluted in CAMHB and subjected to 2-fold serial dilution (10 mM–0.0098 mM) in 96-well microtiter plates. Reference antibiotics (prepared in aqueous solution) were subjected to 2-fold serial dilution from a starting concentration of 5 mM to generate a range of 2.5 mM to 0.00015 mM. Reported MIC values were consistent across screening rounds, regardless of the stock concentration used, indicating that DMSO did not confound the observed antibacterial activity. This was further confirmed by solvent-only controls evaluated at 1–4% (v/v) DMSO, which showed no detectable effect on bacterial growth. Overnight bacterial cultures were grown in CAMHB, adjusted to a turbidity equivalent to 0.5 McFarland (∼1.5 × 10⁸ CFU mL^–1), and subsequently diluted in the same medium to a final inoculum of approximately 1.0 × 10⁶ CFU mL^–1. Serial dilutions of the IL stock solutions were prepared directly in 96-well microtiter plates to achieve the desired test concentrations. Subsequently, 100 μL of the bacterial suspension was added to each well, resulting in a final inoculum concentration of 5 × 10⁵ CFU mL^–1. The cells were incubated for 18–24 h in a static incubator at 37 °C. The MIC was defined as the lowest concentration that inhibited visible bacterial growth. After the initial incubation period, 20 μL of Alamar Blue reagent was added to each well. The plates were then incubated for an additional 2 h to allow for the reduction of resazurin by metabolically active cells. Following incubation, absorbance was measured at an excitation wavelength of 530–570 nm and an emission wavelength of 590–620 nm to assess bacterial viability.

4.5. Checkerboard Synergy Assay

To assess potential interactions between the synthesized ionic liquids (ILs) and other antimicrobial agents, we evaluated them using a checkerboard microdilution assay. Overnight bacterial cultures were washed and adjusted to 1 × 10⁶ CFU mL^–1 in CAMHB. Equal volumes of the bacterial suspension and serially diluted antimicrobials were combined in a 7 × 7 checkerboard layout and incubated at 37 °C for 24 h. Bacterial growth was assessed using the resazurin reduction assay. The fractional inhibitory concentration index (FICI) was determined to characterize the interaction between paired agents, using equation

$$\mathrm{F}\mathrm{I}\mathrm{C}\mathrm{I}=\frac{{\mathrm{M}\mathrm{I}\mathrm{C}}_{\mathrm{a}\mathrm{i}\mathrm{n}\mathrm{c}\mathrm{o}\mathrm{m}\mathrm{b}\mathrm{i}\mathrm{n}\mathrm{a}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n}}}{{\mathrm{M}\mathrm{I}\mathrm{C}}_{\mathrm{a}\mathrm{a}\mathrm{l}\mathrm{o}\mathrm{n}\mathrm{e}}}+\frac{{\mathrm{M}\mathrm{I}\mathrm{C}}_{\mathrm{b}\mathrm{i}\mathrm{n}\mathrm{c}\mathrm{o}\mathrm{m}\mathrm{b}\mathrm{i}\mathrm{n}\mathrm{a}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n}}}{{\mathrm{M}\mathrm{I}\mathrm{C}}_{\mathrm{b}\mathrm{a}\mathrm{l}\mathrm{o}\mathrm{n}\mathrm{e}}}$$

Interpretation of FICI values followed standard criteria: ≤0.5: synergistic interaction; >0.5–1.0: additive effect; >1.0–4.0: indifference; >4.0: antagonism.

4.6. Growth Curves

Growth kinetics and time-kill assays were performed in a complementary manner to evaluate the inhibitory and bactericidal activities of CQUA alone and in combination with conventional antibiotics. Growth kinetics assays were first conducted as a screening approach to assess the effects of treatments on bacterial growth dynamics over time, while time-kill assays were subsequently used to confirm bactericidal activity and validate synergistic interactions. For growth kinetics experiments, bacterial cultures were prepared in CAMHB and adjusted to a final inoculum of approximately 5 × 10⁵ CFU mL^–1. Four experimental groups were evaluated: untreated control (1% DMSO vehicle), CQUA monotherapy, antibiotic monotherapy, and CQUA; antibiotic combination treatment. For monotherapy experiments, CQUA was assessed across a concentration range of 0.5× to 4× MIC. Antibiotics were evaluated at 0.5× to 2× MIC. For combination treatments, CQUA and the respective antibiotic were administered simultaneously, each at concentrations of 0.5× to 2× MIC. Optical density at 630 nm (OD₆₃₀) was measured at 2 h intervals over a 24 h period to monitor growth inhibition and regrowth patterns.

4.7. Time-Kill Kinetics

Time-kill assays were then performed to evaluate the bactericidal and synergistic effects observed in the growth kinetics studies. A bacterial suspension was prepared, adjusted to a final assay inoculum of approximately 5 × 10⁵ CFU mL^–1 in CAMHB. Four experimental groups were evaluated: untreated control (1% DMSO), CQUA monotherapy, antibiotic monotherapy, and combination treatment. CQUA was assessed at final concentrations ranging from 0.5× to 4× MIC, while antibiotics were evaluated at 0.5× to 2 × MIC. For combination treatments, CQUA and antibiotics were administered simultaneously at 0.5× to 2 × MIC. Cultures were incubated at 37 °C with shaking at 150 rpm. At 0, 2, 6, 12, and 24 h, aliquots were collected, serially diluted in sterile saline, and 100 μL was plated onto Mueller–Hinton agar for colony enumeration. Viable counts were expressed as CFU mL^–1. Bactericidal activity was defined as a ≥3 log₁₀ reduction in CFU mL^–1 at 24 h relative to the initial inoculum, while synergistic activity was defined as a ≥2 log₁₀ reduction in CFUmL^–1 for the combination treatment compared with the most active monotherapy at 24 h. All experiments were performed in triplicate.

4.7.1. Scanning Electron Microscopy

SEM was used to study the inhibitory effect of treatment with CQUA. MRSA cultures were incubated in liquid medium containing the test compound at 37 °C with shaking at 150 rpm for 24 h. After incubation, bacterial cells were harvested by centrifugation at 3000g for 10 min at 4 °C. The resulting pellet was resuspended in 1 mL of 1× phosphate-buffered saline (PBS) and washed three times by centrifugation at 1500g for 5 min at 4 °C, discarding the supernatant after each wash. Cells were fixed by resuspension in 2.5% (w/v) glutaraldehyde prepared in 1× PBS and incubated for 2 h at 4 °C. Fixed cells were collected by centrifugation at 4000g for 5 min, washed three additional times with 1× PBS to remove excess fixative, and subjected to graded ethanol dehydration. Briefly, cells were incubated sequentially in 30 and 70% (v/v) ethanol for 5 min each at room temperature, followed by incubation in 100% ethanol for 15 min. After the final wash, the pellet was resuspended in 100% ethanol. A 10 μL aliquot of the ethanol-suspended cells was deposited onto aluminum SEM stubs and air-dried overnight in a closed environment to prevent contamination. Dried samples were sputter-coated with a gold–palladium alloy to enhance conductivity and contrast. SEM images were obtained using a JSM-7200 FLV field-emission scanning electron microscope (JEOL) operated at an accelerating voltage of 10 kV.

4.8. Membrane Permeabilization Assay

Membrane permeabilization of MRSA cells following IL treatment was assessed using Sytox Green, a nucleic acid-binding fluorescent dye that penetrates only cells with compromised membranes. Bacterial suspensions were prepared at a concentration of 1 × 10⁶ CFU mL^–1 in HEPES-glucose buffer (5 mM HEPES, 20 mM glucose). The suspensions were then incubated with the dye (final concentration: 2 μM) in the dark for 30 min at 37 °C, with gentle shaking at 50 rpm. After staining, 100 μL of the cell-dye mixture was transferred into wells of a black 96-well microplate containing the indicated concentration of various treatments. Fluorescence measurements were collected at 2 min intervals over 30 min using excitation and emission wavelengths of 485 and 525 nm, respectively. Melittin (32 μg/mL) served as the positive control for membrane disruption, whereas 1% (v/v) DMSO was used as the negative control. ,

4.9. Membrane Depolarization Assay

Log-phase cultures of MRSA were harvested, washed three times, and resuspended to a concentration of 1 × 10⁶ CFU mL^–1 in HEPES buffer (5 mM) supplemented with 20 mM glucose and 100 mM KCl. The membrane potential-sensitive dye 3,3′-dipropylthiadicarbocyanine iodide (DiSC₃ (5)) was added to the bacterial suspension at a final concentration of 1 μM, followed by incubation in the dark for 30 min to allow the dye to equilibrate across the membrane. After dye loading, cells were exposed to the indicated concentrations of CQUA. Fluorescence intensity was monitored at 2 min intervals for 30 min using excitation and emission wavelengths of 622 and 670 nm, respectively. Melittin served as the positive control for membrane depolarization, whereas 1% (v/v) DMSO served as the negative control.

4.10. Determination of Reactive Oxygen Species (ROS)

Log-phase bacterial cultures were collected, washed, and adjusted to 1 × 10⁶ CFU mL^–1 in PBS supplemented with glucose. To assess overall ROS production, cells were first incubated with 10 μM H₂DCFDA for 30 min at 37 °C in the dark. After staining, the cells were washed twice and resuspended in fresh PBS-glucose buffer. A 100 μL portion of this suspension was transferred into a 96-well plate containing ILs at 0.5× MIC to 4 × MIC. Following a 30 min exposure, fluorescence was recorded at an excitation wavelength of 488 nm and an emission wavelength of 525 nm. Hydrogen peroxide (500 μM) and 1% (v/v) DMSO served as the positive and negative controls, respectively. To monitor specific reactive oxygen species, the probes HKSOX-1, HKPerox-2, and HKOH-1r were employed to detect superoxide anions (O₂•^–), hydrogen peroxide (H₂O₂), and hydroxyl radicals (•OH), respectively. Fluorescence was measured using the appropriate excitation/emission pairs: 500/520 nm for HKSOX-1, 520/543 nm for HKPerox-2, and 500/520 nm for HKOH-1r.

4.11. Membrane Fluidity Assay

Log-phase bacterial cultures were harvested, washed twice with PBS, and adjusted to a final density of 1 × 10⁶ CFU mL^–1. Cells were incubated with Laurdan dye (10 μM) for 30 min at 37 °C in the dark to allow incorporation into the membrane. Following staining, cells were washed twice with PBS and resuspended in fresh PBS before exposure to the indicated concentrations of ILs. Samples were incubated for 30 min, after which fluorescence was recorded at excitation 345 nm and emission wavelengths of 440 and 490 nm, corresponding to ordered (gel-phase) and disordered (liquid-crystalline) membrane states, respectively.

Membrane fluidity was quantified using the Laurdan generalized polarization (GP) parameter, calculated as

$${\mathrm{GP}}_{\mathrm{exc}}=\frac{(A_{440}-A_{490})}{(A_{440}+A_{490})}$$

where A ₄₄₀ and A ₄₉₀ represents fluorescence intensities at 440 and 490 nm.

4.12. Hemolysis Assay

Freshly collected, gender-pooled human whole blood (2 mL) was centrifuged at 1000g for 10 min at 4 °C. The supernatant was discarded, and the red blood cell (RBC) pellet was resuspended in 1× PBS to the original volume. The suspension was centrifuged again at 500g for 5 min at 4 °C. This washing was repeated three times to remove plasma and residual proteins. A 2% hematocrit suspension was prepared by diluting 10 μL of the washed RBC pellet in 490 μL of PBS (1:50, v/v). For each assay, 20 μL of test compound was added to wells of a 96-well microplate, followed by 180 μL of the 2% RBC suspension. Plates were incubated at 37 °C for 1 h and then centrifuged at 1000g for 10 min. A 100 μL aliquot of the supernatant from each well was transferred to a fresh 96-well plate, and absorbance was measured at 410 nm to quantify hemoglobin release. Triton X-100 (5%) and PBS served as positive and negative controls, respectively.

4.13. Cytotoxicity Assay

Human embryonic kidney (HEK293) cells were used to evaluate the effects of the CQ-based ionic liquids (CQILs). Cells were cultured in high-glucose Dulbecco’s modified Eagle medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 5% penicillin–streptomycin and maintained at 37 °C in a humidified incubator with 5% CO₂. Cytotoxicity was assessed using the CellTiter-Glo Luminescent Cell Viability Assay kit. HEK293 cells were seeded into 96-well plates and treated with CQILs at final concentrations ranging from 0.1 to 2.0 mM for 24 h. DMSO-treated cells served as solvent controls. Following incubation, 100 μL of CellTiter-Glo reagent was added to each well, and plates were incubated for an additional 10 min at 37 °C to stabilize the luminescent signal. Luminescence was measured using a Synergy HT multimode microplate reader (BioTek, Winooski, USA). Relative cell viability (%) was calculated by normalizing the luminescence of treated wells to that of untreated control wells, which were defined as 100% viability.

4.14. Statistical Analysis

All statistical analyses were conducted using GraphPad Prism. One-way analysis of variance (ANOVA) was applied to determine differences among groups, followed by Tukey’s post hoc test for pairwise comparisons. Data are presented as mean ± standard error of mean (SEM). Statistical significance was defined as p ≤ 0.05.

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsomega.6c03899.

• ¹H NMR characterization data for all synthesized chloroquine-carboxylic acid ionic liquids (CQ, CQBU, CQ2BE, CQ3BE, CQPE, CQ2PE, CQHE, CQ2HE, CQ3HE, CQHP, CQOC, CQ2OE, CQ3OE, CQNA, CQ2NE, CQ3NE, CQDA, CQ2DE, CQUA, CQDDA); Figure S1, chemical structures of chloroquine and corresponding anions; Figure S2, analytical HPLC of CQUA confirming >95% purity; Table S1, combinatorial antimicrobial effects between CQUA and conventional antibiotics against MRSA; Table S2, MIC and FICI values for the physical mixture of CQ and UA against MRSA; Figure S3, checkerboard heatmap of independently administered CQ and UA; Figure S4, CQUA-induced membrane permeabilization assessed by SYTOX Green fluorescence; Table S3, ionic liquids stock concentrations (PDF)

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Department of Chemistry, The University of Tennessee at Chattanooga, Chattanooga, TN, United States

The authors declare the following competing financial interest(s): E.E.L.T. and M.S. are currently inventors on pending patent applications on the technology disclosed herein.