Novel cationic peptide OB1111 is a dual anti-planktonic and anti-biofilm agent against P. aeruginosa strains PA14 and PAO1

Abstract

Background

Pseudomonas aeruginosa is a Gram-negative pathogen frequently responsible for nosocomial infections and a significant problem in intensive care units. P. aeruginosa, as an opportunistic pathogen, increases mortality risks for severely wounded and immunocompromised individuals. The inherent drug-resistance of P. aeruginosa now requires novel therapeutics with multiple mechanisms that will offer lasting potency in the post-antibiotic era. Synthetic antimicrobial peptides (AMPs) are ideal, as their multiple modes of action slow resistance development. In this study, we investigated the potential of novel proprietary AMP OB1111 to effectively treat P. aeruginosa under standard antimicrobial susceptibility testing (AST) and host-mimicking conditions, in planktonic and biofilm states, and at sublethal and lethal concentrations.

Results

The highly virulent PA14 and moderately virulent PAO1 reference strains were used in these studies. OB1111 effectively displayed inhibitory and bactericidal activity against both strains under AST conditions in planktonic and biofilm states. OB1111 demonstrated anti-virulence activity under host-mimicking conditions by reducing pyoverdine production and early biofilm attachment at sublethal concentrations. Under AST conditions, sublethal doses of OB1111 gradually reduced planktonic PA14 and PAO1 growth but showed less efficacy against biofilms. Additionally, PAO1 biofilms showed reduced susceptibility to OB1111 in comparison to PA14 biofilms at sublethal concentrations. Of significance, scanning electron microscopy revealed that OB1111 effectively deformed and disintegrated PA14 and PAO1 membranes in both the planktonic and biofilm states.

Conclusions

OB1111 successfully demonstrated the capacity to combat P. aeruginosa as an anti-planktonic, anti-biofilm, and anti-virulence agent. Future studies should further examine specific mechanisms of action against PA14 and PAO1, along with testing against clinical isolates in AST and host-mimicking conditions.

Background

Pseudomonas aeruginosa is a Gram-negative, opportunistic pathogen whose widespread presence in nosocomial settings has made it a leading cause of hospital-acquired infections, accounting for 7.1–7.3% in the United States alone [1, 2]. P. aeruginosa causes an even higher percentage of infections in the intensive care unit (ICU), with a point-prevalence international study (involving 88 countries), finding that it is responsible for 24% of intensive care units (ICU)-acquired infections [3]. The pathogen’s affinity for moisture makes hospital sinks and medical devices, including respiratory equipment, prime surfaces for colonization [4, 5]. These conditions lead to a range of high-mortality infections, such as hospital-acquired pneumonia (HAP), ventilator-associated pneumonia (VAP), bloodstream infections (BSI), urinary-tract infections (UTI), surgical site and burn wound infections [2]. P. aeruginosa is also a persistent challenge to cystic fibrosis patients, causing up to 70% of chronic airway infections, which are difficult to treat, and lead to high mortality and morbidity [6, 7]. Overall, P. aeruginosa infections contribute to an annual 300,000 deaths worldwide [8, 9].

These trends are largely driven by the bacteria’s expansive intrinsic and acquired resistomes [10]. Intrinsically, P. aeruginosa has reduced outer membrane permeability, AmpC β-lacatamase expression, and constant (MexAB-OprM) and inducible (MexXY) efflux pumps [10]. It also has an impressively large and plastic genome that readily accepts resistance cassettes through horizontal gene transfer. This mechanism has facilitated an increase in carbapenamase-resistant P. aeruginosa strains, which have been labeled high priority for therapeutic development by the World Health Organization (WHO) [8].

P. aeruginosa is also highly mutable, allowing overexpression of efflux pumps to refuse therapeutics [10, 11]. When comparing pathogenic isolates, 54% of P. aeruginosa isolates were found to be hypermutable as opposed to only 1–12% among E. coli isolates [1–3]. Even more troubling, the success of the pathogen’s adaptability to stressful conditions generally leads to the formation of antibiotic-resistant biofilms [11]. All these factors have led to the emergence of difficult-to-treat resistance (DTR) strains, which render first-line antibiotics, such as β-lactams and fluoroquinolones, ineffective. As such, the need for novel therapeutics is apparent [10, 12, 13].

Antimicrobial peptides (AMPs) are short, cationic peptides and promising candidates to meet this need. In comparison to traditional antibiotics, they target multiple low-affinity sites instead of one high-affinity target in the bacterium [14–17]. The cationic nature of most AMPs facilitates electrostatic interaction with the bacterial membrane to destabilize, depolarize, and deform the membrane, allowing content leakage and, ultimately, cell death [18]. AMPs exhibit multiple mechanisms of action, including membrane permeabilization, membrane disruption, and immune system stimulation, which slow selective pressures that foster the emergence of resistant strains [17]. Moreover, some AMPs have displayed anti-virulence properties, even at sublethal concentrations, potentially raising their therapeutic value (4, 19, 20]). In addition to potency against planktonic bacteria, AMPs have also demonstrated efficacy against bacterial biofilms, which are major hurdles to effective P. aeruginosa treatment [13]. Approximately 45 naturally occurring AMPs with anti-pseudomonas activity have been identified, and approximately 20 AMPs have progressed to clinical trials as of 2023 [1]. While this is encouraging, natural AMPs tend to fall short because of poor absorption, distribution, metabolism, and excretion (ADME) [21]. These pitfalls, along with purity and scalability difficulties, can be overcome with intentionally designed synthetic AMPs.

In this study, we hypothesized that the proprietary synthetic AMP OB1111 would be multifunctional – combating P. aeruginosa through inhibitory, bactericidal, and anti-virulence means. To test this hypothesis, we evaluated the therapeutic effects of OB1111 against commonly used P. aeruginosa reference strains, PA14 and PAO1. As a part of the recently developed P. aeruginosa clade system, Clade I PAO1 and Clade II PA14 collectively represent 98% of all P. aeruginosa strains [5, 6]. OB1111 effects were evaluated against these two strains in both planktonic and biofilm states, at lethal and sublethal doses, and under standard antimicrobial susceptibility testing (AST) and host-mimicking testing conditions. This approach allowed for a thorough investigation of OB 1111’s potential to overcome barriers to clinical application in P. aeruginosa infections.

Methods and materials

Strains, media and antimicrobials

P. aeruginosa reference strains PAO1 (ATCC BAA-47, American Type Culture Collection (ATCC), Manassas, VA, USA) and PA14 (a gift from Dr. George O’Toole, Dartmouth College, Hanover, NH, USA) were used in this study. LB broth, Miller (Lysogeny Broth) (Becton Dickinson, Franklin Lakes, NJ, USA), and BD Bacto Tryptic Soy Broth (Soybean-Casein Digest Medium) were, respectively, used to grow PA14 and PAO1 prior to use in experiments. Cation-adjusted, Mueller-Hinton II Broth (CAMHB) (MOLTOX Molecular Toxicology Inc., Boone, NC, USA) was used in all assays and microscopy studies. Dulbecco’s Modified Eagle’s Medium (DMEM) (ATCC 30–2002) supplemented with 10% Fetal Bovine Serum (FBS) (Gibco, ThermoFisher Scientific, Waltham, MA, USA) or 10% Human Serum (HuS) (Sigma Aldrich, Saint Louis, MO, USA) was used for the AST and host-mimicking conditions. The proprietary AMP OB1111 was donated by Owen Biosciences Inc., Baton Rouge, LA, USA. Polymyxin B sulfate was used as a comparative antibacterial control (Fisher Scientific, Hampton, NH, USA).

AMP screening

Initial screenings were conducted to test proprietary AMPs P215, P577, P360, P380, and OB1111 (Owen Biosciences) against P. aeruginosa strain PA14 to identify compounds with anti-pseudomonas activity. Minimum inhibitory concentrations (MICs) were determined according to Clinical and Laboratory Standards Institute (CLSI) M7-A7 standards [7]. Briefly, PA14 was grown to log phase and diluted to 1 × 10⁶ colony-forming units (CFU)/mL in CAMHB. Serially diluted concentrations (128–8 µg/mL) were tested against a final concentration of 1 × 10⁵ CFU/mL PA14 in a 96-well, untreated, flat-bottom plate in triplicate. Untreated PA14 served as a positive growth control. The microtiter plate was incubated at 37 °C for 24 h and OD₆₀₀ for bacterial growth was recorded using a microplate reader (Sky High, ThermoFisher Scientific). The MIC was recorded as the minimal concentration that inhibits visible bacterial growth.

Bacteriostatic assay

To assess the bacteriostatic activity of OB1111 against PA14 and PAO1, we first determined the MIC according to CLSI M7-A7 standards [7]. Briefly, PA14 and PAO1 were grown to log phase in LB and tryptic soy broth, respectively, and diluted to 1 × 10⁶ CFU/mL in CAMHB. Concentrations of OB1111 were serially diluted in CAMHB starting from 128 µg/mL in triplicate using 96-well, untreated, flat-bottom plates. Diluted strains were added to OB1111-treated wells for a final concentration of 1 × 10⁵ CFU/mL. Untreated bacterial strains served as positive growth controls. The MICs were determined as indicated above.

Bacteriostatic and bactericidal assay under AST and host-mimicking conditions

AST and host-mimicking conditions were used to compare the bactericidal effect of OB1111 against PA14 and PAO1. Specifically, PA14 and PAO1 were each exposed to OB1111 at serially diluted concentrations starting at 128 µg/mL in CAMHB, and from 512 µg/mL in DMEM with 10% FBS (DMEM FBS) and DMEM with 10% HuS (DMEM HuS) run in triplicate. Since PA14 and PAO1 had no visible growth in any of the DMEM HuS treated conditions or controls, the MICs for all samples were recorded as the highest concentration with visible growth on agar in 7/9 replicates from three experiments. The minimum bactericidal concentrations (MBCs) were obtained by plating bacteria concentrations on agar and recording the concentration with a 99.9% reduction in growth in 7/9 replicates from three experiments.

Time-kill analysis

M26-A CLSI guidelines were used to perform time-kill studies of OB1111 against PA14 and PAO1 in a microtiter plate [8]. At concentrations of 1 × 10⁵ CFU/mL, both strains were each exposed to OB1111 at 0.5x, 1x, 2x, 4x and 8x of their determined MICs. The number of CFUs on agar was counted after 2, 4, and 8 h of incubation at 37 °C using the Synbiosis Protos 3 Colony Counter (Synoptics, Inc., Frederick, MD, USA).

OB1111 inhibition of biofilm formation

Total Biofilm Mass: The biofilm inhibition and eradication potential of OB111 and polymyxin B against PA14 and PAO1 were assessed using methods similar to those outlined by Haney et al. [9]. To determine the concentration at which OB1111 was able to inhibit PA14 and PAO1 biofilm formation, both strains were grown statically overnight at 37 °C and diluted to 0.01 OD in CAMHB. In a 96-well plate, PA14 and PAO1 were each treated with OB1111 at serially diluted concentrations of 128 µg/mL to 2 µg/mL and incubated for 20 h at 37 °C. After incubation, total bacterial growth was determined by recording the OD₆₀₀ with a microplate reader. The spent media were discarded from the microtiter plates, which were then washed with sterile distilled water. All treated and control wells were stained with crystal violet (CV) (Fisher Healthcare PROTOCOL, Fisher Scientific) and incubated at RT with gentle shaking. The CV stain was discarded, and plates were washed again before destaining with 70% (vol/vol) ethanol. The absorbance at OD₅₉₅ was recorded using a microplate reader. The same process was followed for polymyxin B in a separate plate for antibiotic comparison using concentrations ranging from 8 µg/mL to 0.25 µg/mL. The dose-response curves showing the percentage of overall bacterial growth and total biofilm mass were calculated in comparison to the untreated controls for each strain. The results from three separate experiments in duplicate or triplicate are shown with the mean ± standard deviation (SD).

Biofilm Viability: To determine the viability of biomasses that formed during the biofilm inhibition experiment, PA14 and PAO1 were grown statically overnight at 37 °C and diluted to 0.01 OD in CAMHB. In a 96-well plate, PA14 and PAO1 were each treated with OB1111 at serially diluted concentrations of 128 µg/mL to 2 µg/mL, Afterward, 1 µL of 5% (wt/vol) 2,3,5-Triphenyltetrazolium chloride (TTC) was added to all treated and control wells. OB1111-treated TTC plates were incubated in a 37 °C incubator for 20 h. Subsequently, methanol was added to TTC plates and incubated at RT with gentle shaking. The absorbance of TTC plates was recorded at A₅₀₀ using a microplate reader.

OB1111 biofilm eradication

To evaluate the ability of OB1111 to eradicate PA14 and PAO1 biofilms, both strains were grown statically overnight at 37 °C and diluted to 0.01 OD in CAMHB. Diluted strains were added to two microtiter plates for OB1111 and two other plates for polymyxin B. To quantify the total biofilm mass, two plates were designated for CV staining, while two plates were reserved for 2,3,5-Triphenyltetrazolium chloride (TTC) staining to quantify biofilm cell viability (biofilm metabolism) using a metabolic indicator [10].

All plates containing PAO1and PA14 were incubated for 20 h at 37 °C to allow biofilm formation. Afterwards, planktonic cells were washed away from the attached biofilms as outlined in the protocol [9]. The concentrations of OB1111 and polymyxin B used for the inhibition of biofilm formation were then added to microtiter plate wells. For the TTC plates only, 2 µL of 5% (wt/vol) TTC was added to all treated and control wells. All OB1111-treated TTC and CV plates were incubated in a 37 °C incubator for an additional 20 h to allow activity against pre-formed biofilms. Subsequently, methanol was added to TTC plates and incubated at RT with gentle shaking. The CV staining procedure was conducted as described above for biofilm inhibition. The absorbance of TTC plates was recorded at A500 using a microplate reader. The percentages of overall bacterial growth, biofilm metabolism (TTC), and total biofilm mass (CV) were calculated as outlined in Nature Protocols [10].

Scanning electron microscopy (SEM) analysis

For planktonic PA14 and PAO1 treatment: Log phase strains were exposed to 4x MIC in CAMHB for 2 h at 37 °C and centrifuged at 2,000 x g to pellet the bacteria. The resulting pellets were washed in 0.1% phosphate-buffered saline (PBS) and fixed overnight in a solution comprised of 2.5% glutaraldehyde and 1% formaldehyde in 0.1% PBS. Samples were stained with 1% osmium tetroxide in 0.1% PBS and then dehydrated with increasing concentrations of ethanol in water. Afterwards, samples were deposited on coverslips adhered to SEM stubs (Electron Microscopy Sciences, Hatfield, PA, USA) with carbon tape. Dried samples were sputter-coated with gold (EMS 550X; Electron Microscopy Sciences, Hatfield, PA) before SEM imaging (JEOL JSM-6010LA, Peabody, MA, USA).

For PA14 and PAO1 biofilm eradication: The SEM procedure for biofilm analysis was adopted as described [11]. Briefly, both strains were grown statically overnight at 37 °C, and after normalization to 0.1 OD, they were diluted 1:100 in CAMHB. A volume of 100 µL was dispensed into one side of an uncoated µ-Slide I Luer (0.4 glass bottom, Ibidi chamber slide). Bacteria were allowed to attach to the bottom of the slide at room temperature (RT) for 2 h and then incubated at 37 °C for an additional 24 h. Samples were washed with PBS, and then replaced with either CAMHB, as a control, or with 4x MIC of OB1111 (Table 2) for an additional 2-hour incubation at 37 °C. Samples were washed with PBS and fixed in 2.5% glutaraldehyde in 1x PBS overnight at 4 °C. All samples were dehydrated with increasing concentrations of ethanol in water, followed by drying in an incubator at 55 °C for 2 h and then completely dried in a desiccator. The coverslips were adhered to an SEM stub and coated with gold before SEM imaging.

Table 2.

Minimum inhibitory and bactericidal concentrations against P. aeruginosa

Strains	OB1111 MIC/MBC (µg/mL)	Polymyxin B MIC/MBC (µg/mL)
PA14	16/32	1/1
PAO1	32/64	2/2

MIC minimum inhibitory concentrations, MBC minimum bactericidal concentration

OB1111 effect on biofilm attachment in AST and host-mimicking media

To determine the concentration at which OB1111 was able to reduce or inhibit PA14 and PAO1 biofilm attachment in AST and host-mimicking media, both strains were grown statically overnight at 37 °C and diluted to 0.01 OD in CAMHB. In a 96-well plate, PA14 and PAO1 were each treated with OB1111 at serially diluted concentrations of 64 µg/mL to 4 µg/mL for CAMHB, and 256 µg/mL to 16 µg/mL for DMEM FBS and DMEM HuS. Plates were incubated for 2–4 h at 37 °C. The spent media was discarded from the microtiter plates, which were then washed with sterile distilled water. All treated and control wells were stained with crystal violet (CV) (Fisher Healthcare PROTOCOL, Fisher Scientific) and incubated at RT with gentle shaking. The CV stain was discarded, and plates were washed again before destaining with 70% (vol/vol) ethanol. The absorbance at OD₅₉₅ was recorded using a microplate reader.

OB1111 effect on pyoverdine production in AST and host-mimicking media

To determine if sublethal concentrations of OB1111 could reduce pyoverdine production, PA14 and PAO1 were grown overnight at 37 °C. They were then diluted to 1 × 10⁵ CFU/mL in CAMHB, DMEM FBS, or DMEM HuS for controls or in CAMHB, DMEM FBS, or DMEM HuS containing 0.25 x MIC OB1111 for each strain. The MIC used to determine the concentration for CAMHB was obtained from Table 1. The MICs from Table 2 were used to determine OB1111 concentration for DMEM FBS and DMEM HuS. All samples were incubated at 37 °C while shaking for 18–20 h. Afterward, ODs were measured using a microplate reader, and ODs for all CAMHB samples were normalized to each other, ODs for all DMEM FBS samples were normalized to each other, and ODs for all DMEM HuS samples were normalized to each other. Samples were then centrifuged at 10,000 g and the supernatants were recorded at A₃₈₀ [12–14].

Table 1.

Antimicrobial susceptibility testing against P. aeruginosa PA14 strain

sAMP	MIC (µg/mL)
215	128
577	32
360	> 128
380	> 128
OB1111	16

sAMP synthetic antimicrobial peptide, MIC minimum inhibitory concentrations

Statistical analysis

Statistical analysis was conducted using two-way ANOVA followed by Tukey’s multiple comparisons test for Bacteriostatic Assay. Biofilm Attachment in AST and Host-mimicking media and Pyoverdine Production data was analyzed using two-way ANOVA followed by Sidak’s multiple comparisons test. OB1111 Inhibition of Biofilm Formation and Biofilm Eradication data was analyzed by two-way ANOVA followed by Dunnett’s multiple comparison test utilizing GraphPad Prism 10. Significance between groups was considered at *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001, ****p ≤ 0.0001.

Results

Identification of OB1111 as effective against P. aeruginosa strain PA14

Multiple synthetic AMPs (sAMPs) (Table 1) were first screened against the highly virulent PA14 strain to identify AMPs with anti-pseudomonas activity. MICs were determined from visualization and confirmed by OD₆₀₀ absorbance readings after a 24-hour incubation at 37 °C. From the initial screening process, OB1111 was selected for further studies against both PA14 and PAO1 because it yielded the lowest MIC at 16 µg/mL.

OB1111 is bacteriostatic and bactericidal against PA14 and PAO1

To evaluate the inhibitory and bactericidal capacity of OB1111 against strains PA14 and PAO1, the MICs and MBCs were obtained by broth microdilution assay using CLSI guidelines for AST [7]. To confirm visualized MICs, a microplate reader was used to record absorbance at OD₆₀₀ after a 24-hour incubation at 37 °C and to produce dose-response curves for both strains (Fig. 1). As shown (Table 2), OB1111 achieved an MIC of 16 µg/mL and an MBC of 32 µg/mL against PA14. For PAO1, the MIC and MBC for OB111 were higher at 32 µg/mL and 64 µg/mL, respectively. As a known and effective anti-pseudomonas AMP, polymyxin B was used as a positive control and exhibited an MIC/MBC of 1 µg/mL against PA14 [15, 16]. Again, PAO1 required higher concentrations for inhibition and elimination with an MIC/MBC of 2 µg/mL.

Fig. 1.

Dose response curves of (A) OB1111 and (B) Polymyxin B against PA14 and PAO1. PA14 and PAO1 (1 × 10⁵) were each treated with OB1111 and polymyxin B at various concentrations in untreated 96-well plates. Samples were incubated for 24 h at 37 °C, and OD₆₀₀ was recorded with a microplate reader. Results show the mean ± SD of triplicate samples from four separate experiments. Significance was analyzed using two-way ANOVA followed by Tukey’s multiple comparisons test. Significance between groups was considered at *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001, ****p ≤ 0.0001

The associated dose-response curves for PA14 and PAO1 exposed to OB1111 revealed statistically significant (p ≤ 0.05) downward trends in absorbance values until the MICs were reached for both strains (Fig. 1) This suggests that OB1111 reduces PA14 and PAO1 growth, potentially through suppression, killing, or metabolic interference at sub-MIC concentrations.

To further evaluate the bactericidal capabilities of OB1111, we performed time-kill analyses to quantify bacterial CFU reduction over an 8h period (Fig. 2). The MBC (32 µg/mL) for PA14 achieved CFU elimination at the 8-hour time point, while the MBC (64 µg/mL), 2x MBC, and 4x MBC all effectively eliminated PAO1 in only 2 h. For PA14, total kill for 2x MBC occurred at 4 h, with 4x MBC being required to achieve complete kill within 2 h. Overall, the OB1111 MBC concentration, which is higher for PAO1, effectively eliminates PAO1 in a fourth of the time needed to achieve the same as PA14. Standard AST inhibitory and bactericidal procedures demonstrate that OB1111 effectively inhibits and eliminates the growth of both highly virulent PA14 and moderately virulent PAO1 in vitro.

Fig. 2.

Time-dependent analysis of OB1111 against (A) PA14 and (B) PAO1. An initial inoculum of PA14 and PAO1 (1 × 10⁵) was each treated with serially diluted concentrations of OB1111. Untreated samples of PA14 and PAO1 were used as controls. Samples were incubated at 37 °C in triplicate in 96-well plates for 2, 4, and 8 h. After the indicated time points, samples were serially diluted and plated overnight at 37 °C on LB agar for PA14 and on tryptic soy agar for PAO1. Resulting CFUs were recorded and log values calculated. Results show the mean ± SD of duplicate to triplicate samples from two separate experiments

OB1111 treatments cause structural membrane damage to PA14 and PAO1

To begin understanding the mechanism by which OB1111 exerts its antimicrobial effects, we evaluated the treated structures of PA14 and PAO1 by SEM (Fig. 3). The effects of OB1111 on both strains, with a starting concentration of 10⁷ CFU/mL, were observed after 2 h of treatment with 4x the MICs (Table 2). These concentrations were increased proportionally for both strains to account for the higher inoculation concentration and reduced treatment period used in this experiment. SEM observation revealed deformed and disintegrated membranes for both strains in comparison to the intact structures of the untreated controls. These results suggest that OB1111 may act by deforming and degrading the cell membrane of treated PA14 and PAO1.

Fig. 3.

Scanning electron microscopy images of OB1111-treated PA14 and PAO1. Exponential phase PA14 and PAO1 were adjusted to a concentration of 10⁷ CFU/mL and treated with 4x MIC of OB1111 for 2 h at 37 °C. Fixed bacteria were deposited onto a coverslip, sputter-coated in gold, and imaged. (A) and (C) show untreated PA14 and PAO1, respectively. B and (D) show the effects of OB1111 on the structures of PA14 and PAO1, respectively. Arrows indicate areas of structural damage to PA14 and PAO1

OB1111 effectively inhibits biofilm formation at bactericidal concentrations

Given the antimicrobial effect of OB1111 against planktonic PA14 and PAO1, we sought to assess whether OB1111 can inhibit biofilm formation using a CV staining assay. We selected the following treatment concentrations (OB1111: 128 µg/mL– 2 µg/mL; polymyxin B: 8 µg/mL– 0.25 µg/mL in CAMHB) to capture the MICs and MBCs for OB1111 against both strains (Table 2) and to allow evaluation of sublethal concentrations. OD₆₀₀ was also measured prior to CV staining to show total bacterial growth (Fig. 4A and C). When comparing the effect of the lowest and highest OB1111 doses on PA14 total growth, bacterial dose response curves show that growth is significantly inhibited by OB1111 at 32 µg/mL (p ≤ 0.0001) and polymyxin B at 2 µg/mL (p ≤ 0.05). For PAO1, total growth curves show significant inhibition by OB1111 and polymyxin B, at 64 µg/mL (p ≤ 0.0001) and 4 µg/mL (p ≤ 0.001), respectively (Fig. 4A). Interestingly, PAO1 bacterial growth increased to approximately 170 (± 48)% at 2 µg/mL of polymyxin B (p ≤ 0.0001) (Fig. 4D). This could indicate that biofilm dispersal increases at one-half the polymyxin B concentration needed to inhibit PAO1 bacterial growth. No statistically significant increase in bacterial cells was noted for PA14 treated with OB1111 or polymyxin B.

Fig. 4.

Biofilm inhibition of OB1111 and Polymyxin B against PA14 and PAO1. Biofilm inhibition was evaluated by incubating OB1111 and polymyxin B with PA14 and PAO1 for 20 h at37 °C in an untreated 96-well plate. Results are expressed as percent (%) growth in comparison to the untreated control for total growth biofilm mass, and biofilm viability. Results show the mean ± SD of triplicate samples from three separate experiments. Significance was analyzed using two-way ANOVA followed by Dunnett’s multiple comparisons test. Significance between groups was considered at * p≤ 0.05, ** p≤ 0.01, *** p≤ 0.001, **** p≤ 0.0001. A and (D) show the total PA14 (blue) and PAO1 (red) bacterial growth after treatment with OB1111 and polymyxin B, respectively. B and (E) show the percent total PA14 and PAO1biofilm formation after OB1111 and polymyxin B treatment, respectively. C and (F) show the percent biofilm viability for PA14 (black) and PAO1 (pink) after OB1111 and polymyxin B treatment. Results are expressed as percent (%) growth in comparison to the untreated control

The biofilm inhibition dose-response curves (Fig. 4B and E) show that the MBC concentrations (Table 2) for PA14 and PAO1 effectively inhibited biofilm formation by at least 70% (p ≤ 0.01), relative to the untreated control. Polymyxin B was used as a positive control and required 2x MBCs for both strains to achieve similar inhibition. Interestingly, the patterns for the dose-response curves for each strain differed at the MIC concentrations for PA14 (16 µg/mL) and PAO1 (32 µg/mL) (Fig. 4B and E). For PA14, sub-MICs and MICs of OB1111 and polymyxin B showed no significant biofilm inhibition in comparison to the untreated control (Fig. 4B and E). However, the MIC for PAO1 showed over 400% increase in biofilm mass for OB1111 and an over 200% increase for polymyxin B, which were significant (p ≤ 0.0001) and higher than the untreated control . However, an assessment of biofilm viability at these concentrations showed that the viability was less than 100% of the untreated control: 47 (± 24)% for OB1111 and 65 (± 20)% for polymyxin B (Fig. 4C and F). Moreover, the viability for PA14 at 32 μg/mL of OB1111 neared full inhibition, measuring 1 (±4)% of the untreated control. PMB at 2 μg/mL yielded similar viability inhibition in PA14, measuring approximately 6 (±6)% of the untreated control. Overall, OB1111 effectively inhibited at least 70% biofilm formation at the MBC concentrations.

OB1111 effectively reduces PA14 and PAO1 biofilm viability

To further explore the potential of OB1111 as an anti-biofilm agent, we treated pre-existing PA14 and PAO1 biofilms with the same concentrations of OB1111 and polymyxin B used in the biofilm inhibition studies (OB1111: 128 µg/mL-2 µg/mL; polymyxin B: 8 µg/mL-0.25 µg/mL in CAMHB). The representative dose-response curves show similar patterns in PA14 and PAO1 total growth, with the highest concentrations of OB1111 (128 µg/mL) and polymyxin B (8 µg/mL) reducing growth by at least 70%, in comparison to the untreated control (Fig. 5A and D).

Fig. 5.

PA14 and PAO1 biofilm eradication by OB1111 and polymyxin B. Biofilm eradication was evaluated after growing untreated PA14 and PAO1 biofilms for 20 h at 37 °C in an untreated 96-well plate. Biofilms were then treated with OB1111 and polymyxin B for an additional 16–20 h at 37 °C. A and (D) show the total PA14 (blue) and PAO1 (red) bacterial growth after treatment with OB1111 and polymyxin B, respectively, in comparison to the untreated controls. B and (E) show the PA14 biofilm viability (black) and total biofilm mass (pink) after OB1111 and polymyxin B treatment, respectively. C and (F) show the PAO1 biofilm viability (black) and total biofilm mass (pink) after OB1111 and polymyxin B treatment, respectively. Results are expressed as percent (%) growth in comparison to the untreated control. Results show the mean ± SD of triplicate samples from three separate experiments. Significance was analyzed using two-way ANOVA followed by Dunnett’s multiple comparisons test. Significance between groups was considered at *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001, ****p ≤ 0.0001

However, dose-response patterns between PA14 and PAO1 differed for both OB1111 (Fig. 5B and C) and polymyxin B (Fig. 5E and F). Total biofilm mass reduction (CV) did not reach 50% for either PA14 or PAO1 treated with OB1111 at 128 µg/mL. However, more importantly, the same concentration of OB1111 effectively reduced biofilm metabolism by at least 65% in both strains, relative to the control. This indicates a strong reduction in live cells within the biofilms.

Concerning sublethal concentrations, we observed a marked increase in biofilm cell viability (approximately 230%) at the 32 µg/mL OB1111 concentration for PAO1 (p ≤ 0.0001) (Fig. 5C). This increase in biofilm metabolism was not observed with any OB1111 concentration for PA14.

The polymyxin B dose-response curve for PAO1 was like that of OB1111, showing increases in biofilm cell viability between 200 and 255% at the 2 and 4 µg/mL polymyxin B concentrations (p ≤ 0.0001) (Fig. 5F). These increases in biofilm cell viability suggest that sublethal doses of both therapeutics may promote biofilm cell viability, specifically in PAO1. However, OB1111 overall was able to effectively reduce PA14 and PAO1 biofilm cell viability to at least 20% when treated with higher doses.

OB1111 disintegrates PA14 and PAO1 biofilm structures

After confirming the anti-biofilm potential of OB1111, we used SEM to visualize the impact of OB1111 on PA14 and PAO1 biofilm structures (Fig. 6). PA14 and PAO1 biofilms were formed on uncoated µ-Slide I Luers, which were incubated for 2 h at RT, followed by 24 h at 37 °C. OB1111 biofilm eradication was observed after 2 h of treatment with 4x MIC (Table 2) for PA14 and PAO1. OB1111 treatment showed extensive biofilm disintegration and deformation with no intact cells easily visible within the raised PA14 and PAO1 biofilms. This morphology of OB1111-treated PA14 and PAO1 biofilms drastically contrasts that of untreated control samples. Notably, the raised biofilm shows more disintegration than the single cells adhered to the coverslips in both OB1111-treated PA14 and PAO1 biofilms. These results suggest that OB1111 can markedly deform and disintegrate pre-existing biofilms in a short treatment period.

Fig. 6.

Scanning electron microscopy images of OB1111-treated PA14 and PAO1 biofilms. After a 24-hour biofilm formation period, PA14 and PAO1 were treated with 4x MIC of OB1111 for 2 h at 37° C. Samples were fixed, dehydrated, and covered with gold for imaging. A and (C) show the untreated biofilm controls for PA14 and PAO1, respectively. B and (D) show the effects of OB1111 on the biofilm structures of PA14 and PAO1, respectively

OB1111 efficacy wans under host-mimicking conditions

Recently, studies have highlighted discordance between therapeutic results obtained with AST methods, which use CAMHB and those that use host-mimicking media. Studies found that screening performed with host-mimicking media more closely aligned with clinical results than with AST conditions [17]. Therefore, we evaluated the bactericidal efficacy of OB1111 in host-mimicking media for comparative purposes. In these experiments, host-mimicking conditions were represented by DMEM FBS and DMEM HuS. DMEM FBS was selected for testing because of its wide use in cell culture experiments, and DMEM HuS was selected as another medium to mimic the human environment [18]. Comparing these two conditions could reveal early differences that might impact future clinical efficacy.

Since serum is known to inhibit AMP function, we tested higher concentration ranges of OB1111 (512 µg/mL – 32 µg/mL) against PA14 and PAO1 during a 24 h incubation at 37 °C [19]. We used a lower concentration range (128 µg/mL– 8 µg/mL) with CAMHB based on the AST results (Table 2). Due to the low growth of both strains in DMEM HuS, the MIC was determined as the highest concentration in which growth could be rescued by overnight agar inoculation. No samples of PA14 or PAO1 visually appeared turbid in DMEM HuS, and optical densities were difficult to detect.

In this comparative study, both the MICs and MBCs for OB1111 increased substantially against PA14 and PAO1 (Table 3). The MIC and MBC for OB1111 against PA14 increased 8-fold in DMEM FBS when compared to results obtained in CAMHB. DMEM- HuS yielded an even higher 16-fold change in MIC compared to the results obtained in CAMHB. This is a surprising result, given that PA14 had the lowest growth in this condition [14]. For OB1111 against PAO1 in DMEM FBS, the MIC/MBC increased by 8- and 4-fold, respectively. The DMEM HuS host-mimicking condition yielded an 8-fold increase in the MIC and MBC. These increases potentially point to reduced susceptibility of PA14 and PAO1 to OB1111 under these particular host-mimicking conditions.

Table 3.

Comparison of OB111 against P. aeruginosa in AST and host-mimicking conditions

Strains	Media	MIC/MBC (µg/mL)	Fold change against CAMHB
PA14	CAMHB	8/32
	DMEM FBS	64/256	+ 8/+8
	DMEM HuS	128/256	+ 16/+8
PAO1	CAMHB	16/32
	DMEM FBS	128/128	+ 8/+4
	DMEM HuS	128/256	+ 8/+8

MIC minimum inhibitory concentrations, MBC minimum bactericidal concentration

Sublethal concentrations of OB1111 reduce early biofilm attachment in host-mimicking conditions

Some antimicrobial peptides have demonstrated anti-virulence activity at sublethal concentrations [4, 20]. Since we observed reduced OB1111 efficacy under host-mimicking conditions, we sought to determine if OB1111 might display anti-virulence activity at lower concentrations. Using a biofilm crystal violet staining assay, we tested the ability of OB1111 to reduce or prevent biofilm attachment at 2 and 4 h time periods in CAMHB, DMEM FBS and DMEM HuS and analyzed biofilm formation at 0.25 x MIC [MICs used for CAMHB: (Table 2); DMEM FBS and DMEM HuS (Table 3)]. For PA14, OB1111 showed significant reduction of biofilm formation only in DMEM FBS and DMEM HuS at 0.25 x MIC after 2 h (Table 3) (Fig. 7). However, biofilm attachment significantly declined in all conditions for PAO1 at both the 2 and 4 h time periods with CAMHB and DMEM FBS and at 2 h with DMEM HuS. The OB1111 concentrations analyzed did not exceed the initial MBCs in CAMHB (Table 2). This demonstrates that OB1111 exhibits anti-virulence activity after short-term interaction with PA14 and PAO1.

Fig. 7.

Biofilm attachment inhibition of OB1111 against PA14 and PAO1. Biofilm attachment was evaluated by incubating OB1111 and polymyxin B with PA14 and PAO1 for 2–4 h at 37 °C in an untreated 96-well plate. Bar graphs show OD₆₀₀ measurements of untreated and OB-1111 treated PA14 in CAMHB (A), DMEM FBS (B), and DMEM HuS (C), and PAO1 in CAMHB (D), DMEM FBS (E), and DMEM HuS (F) after 2 (blue) and 4 (red) hours. Results show the mean ± SD of triplicate samples from two separate experiments. Significance was analyzed using two-way ANOVA followed by Sidak’s multiple comparisons test. Significance between groups was considered at *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001, ****p ≤ 0.0001

Sublethal concentrations of OB1111 reduce pyoverdine in host-mimicking conditions

It has recently been shown that early surface attachment and biofilm development is required for effective pyoverdine production [21]. OB1111’s ability to reduce early P. aeruginosa attachment under host mimicking conditions, then, suggests that it may also reduce pyoverdine production. Using the same concentrations used to assess early biofilm attachment (Fig. 7), we treated both strains with OB1111 overnight and measured pyoverdine production at A₃₈₀ with a microplate reader. Interestingly, we noticed different effects of OB1111 in different media conditions (Fig. 8). Sublethal OB1111 treatment increased pyoverdine production in PA14 in CAMHB and decreased it in DMEM HuS. Sublethal OB1111 did not significantly affect PAO1’s pyoverdine production in CAMHB, but decreased pyoverdine production in both DMEM FBS and DMEM HuS. This highlights the influence that host-mimicking conditions have on virulence expression while demonstrating OB1111’s ability to reduce pyoverdine production at sublethal concentrations.

Fig. 8.

OB1111 effect on pyoverdine production in PA14 and PAO1. Pyoverdine production was evaluated by treating PA14 and PAO1 (10⁵ CFU/mL) with 0.25 x MIC OB1111 overnight. Optical densities were normalized only within each culture condition. Afterward, samples were centrifuged and the absorbance was recorded at A₃₈₀. The bar graphs show the pyoverdine production from PA14 and PAO1 after no treatment (blue) or OB1111 treatment (red) in (A) CAMHB, (B), DMEM FBS, and (C) DMEM HuS. Pyoverdine concentrations were estimated with the following equation: Molar concentration = Absorbance/Extinction coefficient (1.9 × 10⁴ M^− 1 cm^− 1). Results show the mean ± SD of triplicate samples from two separate experiments. Significance was analyzed using two-way ANOVA followed by Sidak’s multiple comparisons test. Significance between groups was considered at *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001, ****p ≤ 0.0001

Discussion

P. aeruginosa is a frequent causative agent of hospital-acquired infections, many of which carry increased mortality and morbidity risks [22]. This pathogen has an extensive arsenal of intrinsic and acquired resistance mechanisms that have led to the emergence and spread of multi-drug resistant (MDR), extensively-drug resistant (XDR), pan-drug resistant (PDR), and the recently defined DTR strains [23, 24]. Further limiting treatment options, P. aeruginosa forms antibiotic-resistant biofilms as a stress response to treatment [25]. With the existence of P. aeruginosa strains that fall into every known category of drug resistance, novel therapeutics are an immediate need. Synthetic AMPs are ideal candidates with multifunctional targeting mechanisms capable of slowing resistance development, which is necessary in today’s therapeutic landscape [26–29].

In this study, we proved the fitness of novel, synthetic AMP OB1111 to overcome constant challenges to effective P. aeruginosa treatment, using the highly virulent PA14 and moderately virulent PAO1 reference strains [30]. Our time-kill studies demonstrated the rapid killing ability of OB1111. Even though different agar compositions were used to quantify CFUs for each strain (LB agar for PA14 and tryptic soy agar for PAO1), both sufficiently support P. aeruginosa growth [31]. One of the current challenges to identifying clinically effective therapeutics is the lack of host-mimicking conditions in the screening process [17]. In addition to identifying OB1111 MICs under standard AST conditions, we recorded higher MICs and MBCs for OB1111 in DMEM FBS and DMEM HuS host-mimicking conditions (Table 3). This result is interesting because DMEM HuS did not support enough PA14 or PAO1 growth to be accurately measured by optical density. However, agar plating revealed less susceptibility of both strains to OB1111 than in any other condition. This could be the result of AMP inhibition by high-density lipoproteins (HDL) in the serum [32]. Reduced OB1111 efficacy could also be the result of environmentally induced stress jointly caused by the media components and OB1111.

We chose to use serum-supplemented DMEM as host-mimicking conditions because of their wide use in in vitro cell studies [33]. However, there is a wide range of host-mimicking media that replicate specific P. aeruginosa infections, including wound, bloodstream, and lung infections, and would certainly impact AMP efficacy differently. For example, Masset et al. compared the activity of multiple synthetic AMPs against P. aeruginosa strains in CAMHB, synthetic wound fluid (SWF), and in synthetic cystic fibrosis sputum media (SCFM). Most of their synthetic AMPs yielded very low MICs in SCFM, medium MICs in CAMHB, and the highest MICs in SWF [34]. Of note, SWF contained serum (FBS), and SCFM did not. Additionally, media alone cannot account for the possible benefits that synergy with host immunity and endogenous AMPs may offer under true clinical conditions [27]. Topical application, moreover, may eliminate the AMP interference that serum tends to cause [35–37].

The inherent and drug-tolerant biofilm-forming ability of P. aeruginosa also presents a therapeutic challenge that OB1111 was able to address. This is an important asset, as biofilms are associated with 60–80% of all microbial infections [38]. Our studies demonstrated that OB1111 effectively inhibited and eradicated PA14 and PAO1 biofilms (Figs. 4 and 5). These promising results agree with other studies that found other synthetic AMPs to have anti-biofilm activity [39–43].

OB1111 also showed evidence that it could deform and disintegrate biofilm structures within a 2 h window as observed by SEM (Fig. 6). These results may hint at known AMP anti-biofilm mechanisms, which include membrane disruption, cell signaling, extracellular polymeric substances (EPS) degradation, and biofilm-associated gene down-regulation [44].

Multiple studies have warned against sublethal antibiotic treatment, which has been identified as a driving force behind bacterial resistance development [45–47]. While this has also been observed with AMPs, some studies suggest that sublethal AMP doses could confer therapeutic benefit in some cases [48–50]. Thus, we investigated the effects of sublethal concentrations of OB1111 against planktonic and biofilm forms of PA14 and PAO1. From these studies, the biofilm inhibition and eradication dose-response curves for PAO1 are notable. We observed at least one sublethal concentration of OB1111 that resulted in substantially increased PAO1 biofilm cell viability in biofilm eradication assays when compared to the untreated control (Figs. 4 and 5). The increase in biofilm cell viability suggests a substantial stress response and reduced susceptibility to OB1111 at sublethal concentrations. Further studies are needed to determine if resistance genes are contributing to this phenomenon.

We must note, however, that using microtiter CV and TTC staining to assess total biofilm mass and biofilm viability, respectively, only provides a limited, high-level screening analysis of biofilm inhibition and formation [9]. The effect of OB1111 on total biofilm mass and biofilm viability were measured in separate microtiter plates, although the same cultures were used for each experiment. Our purpose in using this approach was to determine whether OB1111 had anti-biofilm properties that could later be evaluated by more in-depth, comprehensive means of biofilm analysis, including immunofluorescence and cell membrane permeability assays.

Sublethal concentrations of OB1111 against pre-formed PA14 and PAO1 biofilms under AST conditions potentially pointed to improved viability. However, our assessment of biofilm formation at early time points showed that OB1111 was able to significantly reduce biofilm attachment at sublethal concentrations under host-mimicking conditions. Moreover, OB1111 sublethal concentrations significantly reduced pyoverdine concentrations only when host-mimicking conditions were used. These anti-virulence effects under host-mimicking conditions validate our previous study findings that host-mimicking conditions may be important in evaluating anti-virulence candidates [14]. The initial results obtained here warrant further study of the anti-virulence potential of OB1111. For the early biofilm attachment study, only CV staining was used to deliver results for total biofilm mass. According to the protocol by Haney et al., the biofilm inhibition assay only calls for biofilm viability confirmation when the total biofilm mass is high in wells containing high peptide concentrations [9].

Additionally, we measured pyoverdine production at A₃₈₀ in accordance with results from our previous study [14]. However, the addition of OB1111 to this study required lower PA14 and PAO1 concentrations, resulting in overall lower growth and lower pyoverdine production of both strains, specifically in DMEM HuS. The pyoverdine results from this study should be followed with additional confirmation studies, which could include pyoverdine fluorescence analysis [51]. Overall, the identified trend of reduced early biofilm attachment and pyoverdine production is promising. It also agrees with a recent study showing that biofilm disruption results in lower pyoverdine production [21]. Additionally, AMP anti-virulence potential has been noted in other studies. For example, natural AMP temporin L was able to perturb the E. coli bacterial membrane at sublethal concentrations [52]. The potential for such activity is important to explore because sublethal applications would further lower the potential for resistance development and increase the potential for successful synergy with host immunity and traditional antibiotics against drug-resistant strains. The desired result for novel anti-pseudomonas therapeutics should be to display anti-planktonic, anti-biofilm, and anti-virulence activity, increasing opportunities for use.

Overall, positive results obtained from our studies justify further evaluation of OB1111 against clinical isolates and in animal models to determine the extent of OB1111’s potency. Our results demonstrate that the anti-planktonic, anti-biofilm, and anti-virulence activity of OB1111 observed thus far makes it a promising candidate to treat acute infection, where P. aeruginosa has more motility, and chronic infections, where biofilms likely reside [53].

Conclusion

In conclusion, we have assessed the fitness of OB1111 to combat P. aeruginosa through multiple in vitro studies that provide an initial view of its activity under AST and host-mimicking conditions, against planktonic and biofilm forms, and at lethal and sublethal concentrations. We successfully identified anti-planktonic, anti-biofilm, and anti-virulence potential with one synthetic AMP— OB1111. These are the initial screening analyses needed to address the challenges of today’s post-antibiotic therapeutic landscape. Studies that further clarify the mechanism of action for OB1111 and its potential for topical application should be explored in the future.

Abbreviations

AST

Antimicrobial Susceptibility Testing

ICU

Intensive Care Units

HAP

Hospital-acquired Pneumonia

VAP

Ventilator-associated Pneumonia

BSI

Bloodstream Infections

UTI

Urinary-tract Infections

WHO

World Health Organization

DTR

Difficult-to-treat Resistance

AMP

Antimicrobial Peptide

ADME

Absorption, Distribution, Metabolism, and Excretion

CAMHB

Cation-adjusted, Mueller Hinton II Broth

DMEM

Dulbecco's Modified Eagle's Medium

FBS

Fetal Bovine Serum

HuS

Human Serum

MIC

Minimum inhibitory concentration

MBC

Minimum Bactericidal Concentration

CLSI

Clinical and Laboratory Standards Institute

CFU

Colony-Forming Units

LB

Lysogeny Broth

PBS

Phosphate-buffered Saline

EMS

Electron Microscopy Sciences

SEM

Scanning Electron Microscopy

CV

Crystal Violet

OD

Optical Density

TTC

2,3,5-Triphenyltetrazolium chloride

MDR

Multi-Drug Resistant

XDR

Extensively-Drug Resistant

PDR

Pan-Drug Resistant

HDL

High-Density Lipoproteins

SWF

Synthetic Wound Fluid

SCFM

Synthetic Cystic Fibrosis Sputum Media

Authors’ contributions

A.G., R.S. and V.A.D. designed the experiments. A.G., O.F., A.N. and V.T. performed the experiments. A.G. wrote the manuscript. D.R.O. supplied novel cationic antimicrobial peptide OB1111. V.A.D., R.S., and D.R.O. read, edited, and approved the manuscript.

Funding

This research was funded by NSF-HBCU-UP (HRD-1911660), NIH-NIGMS-RISE (1R25GM106995-01), and the Ph.D. Program in Microbiology at Alabama State University.

Data availability

The datasets generated and/or analyzed during the current study are not publicly available due to the use of a proprietary synthetic antimicrobial peptide but are available from the corresponding author on reasonable request.

Declarations

Ethics approval and consent to participate

Not applicable.

Consent for publication

Not applicable.

Competing interests

D.R.O. is the President and CEO of Owen Biosciences Inc. and is holding, or currently applying for, patents relating to the content of the manuscript.