Dimeric lipo-α/sulfono-γ-AA hybrid peptides as broad-spectrum antibiotic agents

Abstract

There is an urgent need to develop novel antibiotic agents that can combat emerging drug resistance. Herein, we report the design and investigation of a class of short dimeric antimicrobial lipo-α/sulfono-γ-AA hybrid peptides. Some of these peptides exhibit potent and broad-spectrum antimicrobial activity toward both clinically related Gram-positive and Gram-negative bacteria. The TEM study suggests that these hybrid peptides can compromise bacterial membranes and lead to bacterial death. Membrane depolarization and fluorescence microscopy studies also indicate that the mechanism of action is analogous to host-defense peptides (HDPs). Furthermore, the lead compound shows the ability to effectively inhibit biofilms formed from MRSA and E. coli. Further development of the short dimeric lipo-α/sulfono-γ-AA hybrid peptides may lead to a new generation of antimicrobial biomaterials to combat drug resistance.

Introduction

Conventional antibiotics have been used to treat bacterial infections for many years. However, the misuse and overuse of antibiotics aggravating antibiotic resistance is one of the biggest issues to threaten public health over the world.^1,2 Multidrug-resistant (MDR) bacteria such as methicillin-resistant Staphylococcus aureus (MRSA) and Staphylococcus epidermidis (MRSE) occur frequently in community-acquired severe infections. Pandrug-resistant (PDR) Gram-negative strains such as E. coli and Pseudomonas aeruginosa could not be eradicated by most antibiotics.³ It is urgent to develop a new generation of antibacterial agents for combating drug resistance.^4–7 Host-defense peptides (HDPs), natural cationic amphiphilic peptides, are found in all organisms and act as the first line of nonspecific defense against a variety of pathogens including bacteria, fungi, and viruses.^8–10 Unlike traditional antibiotics targeting and breaking down intracellular components or specific cell wall of bacteria, HDPs interact with bacterial membranes firstly by taking advantage of electrostatic interaction between their cationic groups and negatively charged bacterial membranes.^11–13 After that, the hydrophobic residues of HDPs interact with the fatty acyl tails of membrane phospholipids, causing bacterial membrane penetration, depolarization, and disruption.¹⁴ HDPs interact with bacterial membranes by biophysical force and involve no defined specific membrane targets, endowing HDPs with lower antibiotic resistance compared to traditional antibiotics.^15–17 Consequently, HDPs could be a potentially viable strategy to develop a new antibiotic agent.¹⁸ However, intrinsic drawbacks are limiting the development of HDPs as antibiotics for practical application, such as susceptibility to proteolytic degradation and low-to-moderate activity.^19,20 Non-natural peptidomimetics antibiotic agents that mimic the structure and action of HDPs, such as α-peptides,^21,22 β-peptides,^23–25 peptoids,²⁶ arylamide oligomers,²⁷ cationic polymers,^28,29 and others,³⁰ have been explored over the past several decades. In the past years, our group has reported a class of antimicrobial peptidomimetics “γ-AApeptide” which mimics the amphipathic structure of HDPs (Fig. 1).^31–35 Among them, some short lipid γ-AA hybrid peptides, bearing α-amino acid residues and γ-AA units, exhibited potent and broad activity against Gram-positive and/or Gram-negative bacteria.^33,35 Recently, we found that the strategy of dimerization could lead to even stronger activity with relatively small size.^36,37 Herein, we report the application of the dimerization strategy to design a new series of small molecular mimics of HDPs (Fig. 2).

Fig. 1.

General structures of canonical α-peptides, γ-AApeptides, sulfono-γ-AApeptides, and α/sulfono-γ-AA hybrid peptides.

Fig. 2.

(A) Structure of the monomer lipo-α/sulfono-γ-AA hybrid peptides.³⁸ (B) Structure of the dimeric lipo-α/sulfono-γ-AA hybrid peptides.

Results and Discussion

The dimeric lipo-α/sulfono-γ-AA hybrid peptides were synthesized on the solid phase and purified by HPLC (see the ESI^† for details). The ratios of cationic groups, length of lipid tails, and hydrophobic groups in mimicking the amphipathic structure of HDPs were investigated.

The antimicrobial activity of hybrid peptides was tested and is listed in Table 1. Many compounds show good antibacterial activity against both Gram-positive and Gram-negative bacteria, and the relationship between structure and antibacterial activity could lead us to further develop the class of peptidomimetics in the future. Compound 1, containing dimeric sulfono-AApeptide building blocks bearing a chiral positively charged side chain and a hydrophobic phenyl sulfonamido group as well as a C12 lipid tail, already exhibited good activity toward both Gram-positive and Gram-negative bacteria. The structural difference between compounds 1 and 2 is that 2 has a C16 lipid tail instead of C12, but the activity dropped significantly. It is consistent with our previous findings that the length of the lipid tail plays an important role in influencing the compound to interact with the bacterial membrane.^20,39,40 It seems that the C12 lipid tail is the optimal length as compounds 3–5 possess short lipid tails but did not show any antibacterial activity. We next synthesized compounds 6–8 which do not contain the terminal lysine amino acid residues on the two arms. Interestingly, these compounds were much less potent compared with compound 1, indicating the importance of cationic charge. Compounds 9 and 10, bearing two lipid tails instead of one, however, almost completely abolished their antibacterial activity. It is interesting that compound 11, bearing a sulfono-γ-AApeptide building block in each arm with switched positions for both side chains compared with 1, virtually completely lost its antibacterial activity, even though 11 contains an identical ratio of cationic to hydrophobic groups as compound 1. The result may indicate that the arrangement of hydrophobic and cationic patches in the compound is crucial for its antimicrobial activity. Next, we synthesized compound 12 containing just cationic groups as arms. To our surprise, the compound did not exhibit any antimicrobial activity, suggesting that the ratio of cationic groups and hydrophobic groups is critical. Furthermore, we kept the molecular scaffold of 1 and only changed hydrophobic groups on the sulfono side-chain. Among them, compound 17, containing two chlorophenyl sulfonyl side chains, shows the most potent antibacterial activity toward both Gram-positive and Gram-negative all tested bacterial strains. It is possible that the chlorophenyl groups could have both hydrophobic and polar interactions with bacterial membranes, thus enhancing the ability to compromise the membranes.

Table 1.

Structures and antibacterial activity of compounds 1–20. The bacteria used in the study were methicillin-resistant S. aureus (MRSA) (ATCC 33591), methicillin-resistant S. epidermidis (MRSE) (RP62A), E. coli (ATCC 25922), P. aeruginosa (ATCC 27853).

		MIC (μg mL−1)
		Gram-positive		Gram-negative
#	Structure	MRSA	MRSE	E. coli	P. aeruginosa	Hemolysis (HC50, μg mL−1)
1		3–6	3–6	3–6	6–12.5	>250
2		6–12.5	NA	6–12.5	NA	ND
3		NA	NA	12.5–25	NA	ND
4		NA	NA	NA	NA	ND
5		NA	NA	NA	NA	ND
6		3–6	NA	6–12.5	12.5–25	>250
7		3–6	NA	NA	NA	ND
8		NA	NA	12.5–25	NA	ND
9		NA	6–12.5	NA	NA	ND
10		NA	NA	NA	NA	ND
11		6–12.5	NA	12.5–25	NA	ND
12		NA	NA	NA	NA	ND
13		3–6	3–6	1.5–3	NA	ND
14		3–6	12.5–25	6–12.5	NA	ND
15		6–12.5	NA	6–12.5	NA	ND
16		3–6	NA	6–12.5	NA	ND
17		3–6	3–6	3–6	3–6	>250
18		3–6	1.5–3	3–6	NA	>250
19		3–6	1.5–3	1.5–3	NA	>250
20		3–6	3–6	3–6	NA	>250
	Ciprofloxacin	0.75–1.5	0.75–1.5	0.75–1.5	1.5–3	ND

ND: not determined. Hemolysis activity was not measured for peptidomimetics showing antimicrobial activity less than three tested strains

Based on the HPLC retention time (RT) of all the compounds, we estimated the amphiphilicity of these compounds and tried to correlate it with antimicrobial activity. From the HPLC data (Table S2 in ESI^†), the most potent compound 17 has a RT of 19.70 min. These compounds with RT shorter than 18 min (compounds 3, 4, 5, and 12) did not exhibit any antibacterial activity. The antibacterial activity of compounds in general increased with an increase in the RT value from 18 to 19.7 min; it then started to decrease when the RT value was more than 19.70 min. The analysis seems to be consistent with our previous statement that the balance of hydrophobicity and hydrophilicity plays an important role in influencing antibacterial activity.^20,41

After that, hemolytic assay was conducted to evaluate the selectivity of these compounds.⁴² As shown in Table 1, for breaking down 50% of human red blood cells, compound 17’s concentration must be greater than 250 μg mL⁻¹. The concentration is more than 40-fold of compound 17’s antibacterial activity concentration toward all four tested bacterial strains. It attests that the lead compound 17 has low toxicity and high selectivity toward bacteria.

Then, we conducted the cell viability assay to test the selectivity of the lead compound. In this study, we used DAPI for staining cells’ nucleus and Phalloidin-iFluor 594 for staining cells’ cytoskeleton. The HeLa cell line has been chosen since it is the oldest and most commonly used human cell line.⁴³ After HeLa cells have been incubated with compound 17 at 12 μg mL⁻¹ (2-fold of MIC) for 24 h, both the cells’ cytoskeleton and nucleus are in good shape (Fig. 3B) compared with the control sample (Fig. 3A). The result suggests that the lead compound could eradicate bacteria without affecting human cell viability.

Fig. 3.

Cell viability of HeLa that are treated or not treated with 12 μg mL⁻¹ compound 17 for 24 h: (A) control, untreated; (B) cell treated with 17.

As we hypothesized that the dimeric lipo-α/sulfono-γ-AA hybrid peptides could mimic the mechanism of HDPs, these peptides should be able to disrupt the bacterial membrane and eradicate bacteria rapidly.⁴⁴ The time-kill study was then conducted to explore the bacterial killing kinetics for the most potent compound 17. As shown in Fig. 4, compound 17 exhibited a concentration-dependent effect on both MRSA and E. coli, with faster killing kinetics at higher concentrations. The complete killing of both MRSA and E. coli was observed within 6 h at 2 × MIC. Compound 17 exhibited more rapid bactericidal kinetics eradicating MRSA compared to E. coli at a concentration higher than 2 × MIC. Notably, MRSA could be eradicated in roughly 2 h at 8 × MIC of compound 17. Overall, these results indicate that compound 17 is kinetically favorable as it relates to the killing of bacteria in a similar manner compared to HDPs.

Fig. 4.

Time-kill plots of 17 against MRSA (A) and E. coli (B).

Subsequently, fluorescent microscopy was conducted to further study the compounds’ effect to compromise bacterial membranes. In this assay, DAPI could stain bacterial cells’ membranes with blue fluorescence irrespective of their viability. However, PI can only stain the dead or impaired cells’ membranes with red fluorescence. As shown in Fig. 5, the control, without treatment with compound 17, shows blue fluorescence with DAPI (b1 and b3) but no fluorescence with PI (a1 and a3), proving that both MRSA and E. coli bacteria’s membranes were intact. After incubating MRSA and E. coli with compound 17 for 2 h, both DAPI and PI channels (a2, b2, a4, and b4) were observed with strong fluorescence. It demonstrates that the membranes of both MRSA and E. coli were compromised.

Fig. 5.

Fluorescence micrographs of MRSA and E. coli that are treated or not treated with 12 μg mL⁻¹ of compound 17 for 2 h: (a1) control, no treatment, PI stained; (b1) control, no treatment, DAPI stained; (c1) control, no treatment, PI and DAPI channel merge; (a2) MRSA treatment with 17,PI stained; (b2) MRSA treatment with 17, DAPI stained; (c2) MRSA treatment with 17, PI and DAPI channel merge;(a3) control, no treatment, PI stained; (b3) control, no treatment, DAPI stained; (c3) control, no treatment, PI and DAPI channel merge; (a4) E. coli treatment with 17, PI stained; (b4) E. coli treatment with 17, DAPI stained; (c4) E. coli treatment with 17, PI and DAPI channel merge.

To further test whether compound 17 mimics the mechanism of action of HDPs, we carried out membrane depolarization. DiSC3(5) is a membrane-potential voltage-sensitive dye that could accumulate on hyperpolarized membranes and translocate into the lipid bilayer. The weak fluorescence is observed in living bacterial cells due to self–quenching; however, the fluorescence intensity would increase dramatically when the bacterial membrane potential is lost. As such, bacteria were incubated with DiSC3(5) for 30 min, after that immediately treated bacteria with compound 17 in different concentrations of 1 × MIC, 2 × MIC, and 4 × MIC. As shown in Fig. 6, both MRSA and E. coli show dramatic enhancement of fluorescence intensity after treatment of compound 17, and the increase of fluorescence intensity is concentration-dependent. The results are consistent with that of the fluorescence microscopy that compound 17 could disrupt the bacterial membrane.

Fig. 6.

Membrane depolarization of MRSA (A) and E. coli (B). Untreated control is the culture without antibacterial treatment. The experiment was repeated three times with duplicates each time.

Furthermore, the TEM study was conducted to visualize if our compounds can damage bacteria’s membranes as the TEM micrograph is a straightforward way to show cell membrane morphology. As shown in Fig. 7A and C, the control, MRSA and E. coli had intact membrane structures. After treatment with 12 μg mL⁻¹ concentration of compound 17 for 2 h, both MRSA and E. coli membranes have been damaged, and even some cells became debris (Fig. 7B and D). The result visually proves that compound 17 could destroy bacterial cell membranes.

Fig. 7.

Observation of cell membrane damage by transmission electron micrographs (TEM): (A) MRSA cells without any antibacterial treatment (control); (B) MRSA cells treated with compound 17 at 12 μg mL⁻¹; (C) E. coli cells without any antibacterial treatment (control); (D) E. coli after treatment with compound 17 at 12 μg mL⁻¹.

Next, we explored the ability of the lead compound 17 to permeabilize the outer membrane (OM) of Gram-negative bacteria by using the hydrophobic fluorescent probe N-phenyl-1-naphthylamine (NPN). NPN’s fluorescence emission is increased when it transfers to a glycerophospholipid environment.⁴⁵ Hence, improved fluorescence of NPN reveals disrupting or functional impairment of the OM structure of Gram-negative bacteria.⁴⁶ In the study, melittin was used as a positive control. As shown in Fig. 8, compound 17 improved the permeability of OM in a concentration-dependent manner. Conspicuously, compound 17 induced the permeabilization of the E. coli’s OM more than 50% even at the low concentration (1 × MIC). Compound 17 exhibited a high potency in the permeability of the OM. The result emphasizes that the lead compound 17 can interrupt with the bacterial membrane by mimicking the mechanism of action of HDPs.

Fig. 8.

Outer membrane permeability induced by compound 17 and melittin. The uptake of NPN of E. coli in the presence of different concentrations of compound 17 was determined using the fluorescent dye (NPN) assay. The percent uptake is relative to 16 mM melittin.

It is known that most bacterial infections are followed by biofilm formation, and bacteria in the biofilm are much harder to kill than independent cells since biofilms have potent resistance to antibiotics.⁴⁷ As such, we conducted the biofilm study to evaluate compound 17’s inhibitory effect on biofilm formation of MRSA and E. coli. As shown in Fig. 9, at a concentration of just 0.094 μg mL⁻¹, almost 50% of biofilm formation of MRSA was inhibited by compound 17. With regard to the biofilm formation of E. coli, the suppressed capability of compound 17 was not that strong, even though, at a concentration of 0.188 μg mL⁻¹, compound 17 could inhibit around 40% of biofilm formation of E. coli. The results reinforced the potential of compound 17 as the novel antibiotic agent.

Fig. 9.

Biofilm disruption by compound 17: (A and B) Biological activity of 17 in the inhibition of a biofilm by MRSA(A) and E. coli (B). (C and D) images of the treated and untreated biofilms of MRSA (C) and E. coli (D), respectively after staining with crystal violet.

Low metabolic stability of HDPs, which is an inherent risk of therapeutic peptides in general, is considered a key factor limiting their clinical application.⁴⁸ Proteolytic stability is a key property in evaluating these dimeric compounds’ potential for clinical application. We next conducted the enzymatic stability of the lead compound 17. The assays were performed by incubating 0.1 mg mL⁻¹ compound 17 with pronase (0.1 mg mL⁻¹) in 100 mM ammonium bicarbonate buffer (pH 7.8) at 37 °C for 24 hours. We chose pronase because it is a mixture of several nonspecific endo- and exoproteases that digest proteins down to single amino acids.⁴⁹ As shown in Fig. 10, compound 17 shows almost no degradation after incubating with pronase for 24 hours. The result exhibits good stability of the dimerized γ-AA peptides against enzymatic degradation, augmenting their potential in therapeutic application.

Fig. 10.

Stability study: analytic HPLC traces of 17 before and after incubation with pronase (0.1 mg mL⁻¹).

As more and more conventional antibiotics are rendered ineffective by drug-resistant bacteria,⁵⁰ it would be important to evaluate the lead compound 17 potential in developing drug resistance. Therefore, the study on bacterial resistance by E. coli toward compound 17 was conducted. Ciprofloxacin, a common antibiotic used to treat a variety of bacterial infections,⁵¹ was used as a positive control for E. coli. As shown in Fig. 11, serial 16 passages of E. coli in sub-MIC concentrations of compound 17 generated strains that remained highly sensitive to compound 17. In contrast, an obvious increase in MIC was observed in the case of ciprofloxacin. The results suggest that lead compound 17 is difficult to develop drug resistance.

Fig. 11.

Fold increase in MIC of compound 17 and ciprofloxacin toward E. coli.

Conclusion

In summary, we designed and synthesized a series of short dimeric lipo-α/sulfono-γ-AA hybrid peptides through a dimerization strategy at ease. The lead compounds showed potent bacteria-killing efficacy against a series of Gram-positive and Gram-negative bacteria with low hemolytic toxicity, suggesting the therapeutic potential of these compounds. Fluorescence microscopy, TEM, OM permeabilization and membrane depolarization studies demonstrated that the lead compound 17 could kill bacteria by interrupting with the bacterial membrane, mimicking the mechanism of action of HDPs. Compound 17 also has good ability to inhibit biofilm formation. The further development of the class of dimerized γ-AA peptides could lead to a new generation of antibacterial agents.

Experimental section

General information

Rink-amide resin (0.64 mmol g⁻¹, 200–400 mesh) and Fmoc protected α-amino acids were purchased from Chem-impex Int’l Inc. Solvents, coupling reagents, and other chemicals of reagent grade were purchased from either Fisher Scientific or Sigma-Aldrich. Column chromatography was carried out with silica gel (200–300 mesh). All compounds were synthesized by using solid-phase peptide synthesis. The compound was synthesized in a peptide reaction vessel clamped on a Burrell wrist-action shaker and purified on a High-Performance Liquid Chromatography (HPLC) system with both analytical and preparative functions. The final product’s molecular weight was confirmed by using a Bruker AutoFlex MALDI-TOF mass spectrometer, and the desired final pure products were dried on a Labcono lyophilizer.

Synthesis of sulfono-γ-AApeptide building blocks

The synthesis of the Alloc-γ-AApeptide building blocks ((Fig. S1^†) was followed using the previously reported procedure.^52,53

Synthesis of the desired short dimeric lipo-α/sulfono-γ-AA hybrid peptides

The synthesis of dimeric lipo-α/sulfono-γ-AA hybrid peptides was conducted on the solid phase following our previously reported protocol.^53,54 The alloc protected building blocks were attached to the solid phase, which were subjected to the alloc removal and reaction with various sulfonyl chlorides. The synthetic routes of the desired short dimeric lipo-α/sulfono-γ-AA hybrid peptides attached are shown in the ESI (Fig. S3–S6^†).

Compound 17

¹H NMR (600 MHz, DMSO-d6) δ 10.32 (s, 2H), 8.39 (d. J = 8.4 Hz, 1H), 8.22 (s, 4H), 7.80–7.89 (m, 5H), 7.79 (s, 2H), 7.78 (s, 2H), 7.60 (s, 2H), 7.59(s, 2H), 7.54 (s, 1H), 7.12 (s, 2H), 4.35 (dd. J = 8.4, 13.8 Hz, 1H), 4.22 (d. J = 12 Hz, 2H), 4.10 (d. J = 16.8 Hz, 4H), 3.77–3.82 (m, 2H), 3.3 (s, 4H), 2.98–3.01 (m, 2H), 2.74–2.78 (m, 8H), 2.01(t. J = 7.2 Hz, 2H), 1.70–1.83 (m, 6H), 1.50–1.55(m, 10H), 1.43–1.47 (m, 4H), 1.36–1.39(m, 10H), 1.27–1.33 (m, 8H), 1.22–1.26 (m, 18H), 0.85 (t. J = 6.6 Hz, 3H). ¹³C NMR (150 MHz, DMSO-d6) δ 174.25, 172.54, 169.03, 166.72, 158.72, 139.08, 138.45, 138.27, 136.07, 129.66, 129.43, 118.60, 116.62, 114.30, 53.47, 52.59, 52.41, 49.99, 47.36, 40.83, 39.12, 35.89, 32.04, 31.76, 31.46, 31.18, 30.94, 29.46, 29.18, 27.34, 27.11, 25.80, 22.75, 22.57, 21.60, 14.44. HRMS (ESI) C65H107Cl2N15O11S2 [M + H]⁺ calcd = 1408.71; found = 1408.74.

Minimum inhibitory concentrations (MICs) antimicrobial assays⁴¹

We tested the antimicrobial activity of the compounds against four different bacterial strains including MRSA (ATCC 33591), MRSE (RP62A), P. aeruginosa (ATCC27853), and E. coli (ATCC 25922). The experiment was completed with the following protocol. The bacteria were grown overnight in 5 mL of tryptic soy broth (TSB) buffer at 37 °C to the mid-logarithmic phase and diluted to 1 × 10⁶ CFU mL⁻¹. 50 μL of these bacterial suspension were added to every well of a 96 well plate followed by 50 μL of different concentrations of dimeric lipo-α/γ-AA peptides diluted with the same TSB medium. The mixtures were incubated at 37 °C for 18 h, and the Minimum Inhibitory Concentrations (MICs) were recorded by measuring optical density at the absorption at 600 nm wavelength on a Biotek Synergy HT microtiter plate reader. MICs are the lowest concentrations of the compounds which can completely inhibit the bacterial growth. Results were repeated three times with duplicates each time.

Hemolytic assays³⁶

Fresh human red blood cell (hRBCs) was washed with 1× PBS and centrifuged at 3000 rpm for 10 min twice. The top clear supernatant was removed. The bottom hRBC layer was diluted to 5% v/v suspension in 1× PBS. 50 μL of the diluted hRBC solution was added into a 96-well plate, and to each well was added 50 μL of 2-fold serial dilution lead compounds. The plate was incubated at 37 °C for 1 hour. After that, the mixture solution was centrifugated for 10 minutes at 3000 rpm. 30 μL of the supernatant was transferred to another 96-well plate containing 100 μL of PBS in each vial. The absorbance of the mixture was obtained on a Biotek Synergy HT plate reader at a wavelength of 540 nm. The hemolysis activity was determined by the formula: % hemolysis = (Abs_sample -Abs_PBS)/(Abs_Triton -Abs_PBS) × 100. 1× PBS was used as the negative control and 2% Triton X-100 was used as the positive control.

Cell viability assay⁵⁵

12 μg mL⁻¹ concentration of compound 17 was incubated with HeLa cells on a confocal dish at 37 °C for 24 h. The cells were then washed with PBS (pH = 7.4) twice, the cells were fixed with 4% formaldehyde solution at 37 °C for 15 min. After that, the cells were stained with DAPI (10 μL) and Phalloidin-iFluor 594 (1 μL) in PBS for 10 min, respectively. Subsequently, the cleaned cells were observed and recorded using a biological confocal laser scanning microscope (fv1200, OLYMPUS).

MTT assay³⁸

MTT assay was determined using the cell counting kit-8 (CCK8) assay. HeLa cells were cultured in high glucose DMEM supplemented with 10% FBS, 4 mM L-glutamine, and 1% penicillin/streptomycin, in 5% CO₂ at 37 °C. HeLa cells under good conditions were placed in 96-well plates at a concentration of 5 × 10³ cells per well with six replicates in 100 μL of complete medium, cultured for 24 h. Then, the medium was removed and the cells were washed with phosphate buffered saline (PBS). 100 μL of serum-free DMEM medium containing various concentrations of compound 17 was added to the wells. After culturing for 24 h in 5% CO₂ at 37 °C, 10 μL of CCK8 reagent was added to each well. OD values were read at 450 nm on a BioTek Synergy Hybrid plate reader after 2 h in the incubator. The final cell viability was calculated as: cell viability % = (OD - ODblank)/(ODctrl - ODblank) × 100%. ODctrl is the OD of the well containing cells only. ODblank is the OD of the well containing serum-free DMEM only. Experiments were repeated three times.

Time-kill assays³³

The time-kill assay was performed to determine the efficacy of the best compound 17 against MRSA (Gram-positive) and E. coli (Gram-negative). The bacteria were grown overnight in TSB at 37 °C to the mid-logarithmic phase and diluted to 1 × 10⁶ CFU mL⁻¹. The bacteria were then incubated with different concentrations of compound 17. Then, the suspensions were placed on agar plates at different time points. The suspensions were placed onto agar plates. The plates were incubated 20 h at 37 °C and then colonies were counted. The assays were repeated three times.

Fluorescence microscopy⁵⁶

Two dyes were used in the study: 4',6-diamidino-2-phenylindole dihydrochloride (DAPI) and propidium iodide (PI). DAPI has good ability to dye any bacterial cells; however, PI can only dye dead cells because it can only pass through damaged membranes. Bacteria were cultured to the mid-logarithmic phase at 37 °C and diluted to 1 × 10⁶ CFU mL⁻¹, then, incubated with compound 17 for 2 h at 37 °C (no compound was added to the untreated controls), the cell pellets were collected after centrifugation at 3000 rpm for 15 min at 4 °C, then washed with PBS and incubated with DAPI (10 μg mL⁻¹) for 15 min on ice in the dark. Next, the bacteria were washed twice with PBS to remove excess dye. The procedure was repeated with PI (5 μg mL⁻¹). A Zeiss Axio Imager Z1 optical microscope with an oil-immersion objective was then used to observe the cells.

Bacterial membrane depolarization assay³³

The bacterial cells were collected at the mid-log phase, then washed with 5 mM HEPES and 5 mM glucose. The bacteria were re-suspended in a 1: 1: 1 ratio (10⁶ CFU mL⁻¹) of 5 mM glucose, 5 mM HEPES, and 100 mM KCL solution. 192 μL of bacterial suspension and 8 μL of 10 μM DiSC3(5) were added into a 96-well plate at 37 °C. After that, the fluorescence of the suspension was monitored for a half-hour at 37 °C at the excitation wavelength of 622 nm and the emission wavelength of 670 nm. Compound 17 was added to the wells immediately when the minimum value of fluorescence was reached. An increase in fluorescence was recorded. The experiment was repeated three times.

Transmission electron microscopy (TEM)⁴¹

Bacterial cells grew in the mid log phase and were incubated with compound 17 in TSB at 37 °C for 2 h. Then, the mixture was centrifuged at 3000 rpm for 15 min and the bacterial pellets were collected at the bottom of the centrifuge tube. The bacterial pellets were washed with PBS four times, after that, the bacteria were suspended in deionized water. The samples without compound 17 were used as the control samples. A drop of each sample’s solution was added onto the carboncoated Cu grids and the excess sample was wiped off with filter paper. The grids were set to dry for about 1 h and then used for the TEM study. TEM images were taken under an FEI Morgagni 268D TEM with an Olympus MegaView III camera on the microscope. The microscope uses Analysis software to run the camera. The microscope was operated at 60 kV.

Outer membrane permeability assay⁵⁷

Mid-logarithmic phase E. coli cells were collected and washed twice with washing buffer (5 mM HEPES with 5 mM glucose, pH 7.4). After that, the cells were suspended in 5 mM HEPES buffer solution to an optical density at 600 nm (OD600) = 0.4. 2 mL of 40 μM NPN was added to 2 mL of the bacterial suspension. The desired concentrations of compound 17 were then added to the bacterial suspension. The mixtures were then incubated at 37 °C for 1 h. Fluorescence readings were measured using a Biotek Synergy HT plate reader with an excitation wavelength of 355 nm and an emission wavelength of 405 nm. 16 mM melittin was used to induce maximal NPN uptake. The fluorescence was recorded as a function of time until no further increase in the fluorescence was observed. The percentage of NPN uptake in the presence of peptide was calculated based on the formula below

$$\%\mathrm{N}\mathrm{P}\mathrm{N}\mathrm{u}\mathrm{p}\mathrm{t}\mathrm{a}\mathrm{k}\mathrm{e}\mathrm{=}\frac{\mathit{F}{-\mathit{F}}_{\mathrm{0}}}{{\mathit{F}}_{\max }-{\mathit{F}}_{\mathrm{0}}}\mathrm{\times }\mathrm{100}\%$$

where F₀ is the fluorescence intensity corresponding to NPN uptake in the absence of peptide and melittin. F_max and F are the fluorescence intensities after the addition of 16 mM melittin or peptide, respectively.

Inhibition of biofilms⁴²

Bacteria (MRSA and E. coli) were allowed to grow for 12 h at 37 °C. 50 μL of 2-fold serial dilutions of compound 17 were placed in a 96-well plate, followed by the addition of 50 μL of diluted bacterial medium (1 × 10⁶ CFU mL⁻¹) each vial. Then after incubation for 48 h at 37 °C, the wells were gently washed with TSB three times to remove unattached biofilms. The wells were dried after 24 h, and then 200 μL of 0.1% crystal violet solution was added to stain the attached biofilms for 15 min. The wells were washed with TSB three times to remove excess stained solution and allowed to dry again. After that, 200 μL of 30% acetic acid was added to each vial to dissolve the stain for 15 min. 125 μL of the dissolved solution was transferred to a new 96-well plate and recorded for the OD reading at 595 nm wavelength.

Enzymatic stability study⁵⁸

Lead compound 17 (0.1 mg mL⁻¹) was incubated with protease (0.1 mg mL⁻¹) in 100 mM ammonium bicarbonate buffer (pH 7.8) at 37 °C for 24 hours. Then, the reaction mixtures were concentrated in a speed vacuum at medium temperature to remove water and ammonium bicarbonate. The resulting residues were redissolved in DI H₂O/ACN and analyzed on a High-Performance Liquid Chromatography (HPLC) system with a flow rate of solvent B (0.1% TFA in ACN) in A (0.1% TFA in DI H₂O) over a duration of 50 min. The ultraviolet detector was set to 215 nm.

Drug resistance assay²⁰

Lead compound 17 was chosen for the drug resistance study. The first MIC value was measured following the same method as the MIC assay mentioned before. Then, it was well suspended with bacteria after the last clear well was employed to make the bacterial suspension (approximately 10⁶ CFU mL⁻¹) for the next experiment to test new MIC until it was repeated for 16 passages. Ciprofloxacin was used as the positive control. The assay was conducted independently in triplicate with three biological replicates.