# How do antimicrobial peptides disrupt the lipopolysaccharide membrane leaflet of Gram-negative bacteria?

Haoning Gong, Xuzhi Hu, Lin Zhang, Ke Fa, Mingrui Liao, Huayang Liu, Giovanna Fragneto, Mario Campana, Jian Ren Lu

► To cite this version:

Haoning Gong, Xuzhi Hu, Lin Zhang, Ke Fa, Mingrui Liao, et al.. How do antimicrobial peptides disrupt the lipopolysaccharide membrane leaflet of Gram-negative bacteria?. Journal of Colloid and Interface Science, 2023, 637, pp.182-192. 10.1016/j.jcis.2023.01.051. hal-05442012

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# How do antimicrobial peptides disrupt the lipopolysaccharide membrane leaflet of Gram-negative bacteria?

<figure><img src="image_3.png" /></figure>

Haoning Gong<sup>a,b</sup>, Xuzhi Hu<sup>a</sup>, Lin Zhang<sup>a</sup>, Ke Fa<sup>a</sup>, Mingrui Liao<sup>a</sup>, Huayang Liu<sup>a</sup>, Giovanna Fragneto<sup>c</sup>, Mario Campana<sup>d</sup>, Jian Ren Lu<sup>a,*</sup>

## GRAPHICAL ABSTRACT

<figure><img src="image_4.png" /></figure>

## ARTICLE INFO

**Article history:**
Received 3 November 2022
Revised 24 December 2022
Accepted 10 January 2023
Available online 13 January 2023

**Keywords:**
Antimicrobial peptide
Lipopolysaccharide
Neutron reflection
Structural disruption
Anti-infection

## ABSTRACT

**Hypothesis:** It is widely regarded that antimicrobial peptides (AMPs) kill bacteria by physically disrupting microbial membranes and causing cytoplasmic leakage, but it remains unclear how AMPs disrupt the outer membrane (OM) of Gram-negative bacteria (GNB) and then compromise the inner membrane. We hypothesise that different AMPs impose different structural disruptions, with direct implications to their antimicrobial efficacies.

**Experiments:** The antimicrobial activities of three typical AMPs, including the designed short AMP, G₃, and two natural AMPs, melittin and LL37, against E. coli and their haemolytic activities were studied. Lipopolysaccharide (LPS) and anionic di-palmitoyl phosphatidyl glycerol (DPPG) monolayer models were constructed to mimic the outer membrane and inner membrane leaflets of Gram-negative bacteria. The binding and penetration of AMPs to the model lipid monolayers were systematically studied by neutron reflection via multiple H/D contrast variations.

**Finding:** G₃ has relatively high antimicrobial activity, low cytotoxicity, and high proteolytic stability, whilst melittin has significant haemolysis and LL37 has weaker antimicrobial activity. G₃ could rapidly lyse LPS and DPPG monolayers within 10–20 min. In contrast, melittin was highly active against the LPS membrane, but the dynamic process lasted up to 80 min, with excessive stacking in the OM. LL37 caused rather weak destruction to LPS and DPPG monolayers, leading to massive adsorption on the membrane surface without penetrating the lipid tail region. These findings demonstrate that the rationally

<footnote><sup>a</sup>Biological Physics Laboratory, Department of Physics and Astronomy, Faculty of Science and Engineering, The University of Manchester, Manchester M13 9PL, UK</footnote>

<footnote><sup>b</sup>State Key Laboratory of Biochemical Engineering, Institute of Process Engineering, Chinese Academy of Sciences, Beijing 100190, PR China</footnote>

<footnote><sup>c</sup>Institut Laue-Langevin, Grenoble 38042, France</footnote>

<footnote><sup>d</sup>ISIS Pulsed Neutron & Muon Source, STFC Rutherford Appleton Laboratory, Didcot OX11 0QX, UK</footnote>

<footnote>* Corresponding author.
E-mail address: j.lu@manchester.ac.uk (J.R. Lu).</footnote>
designed AMP G3 was well optimised to impose most effective destruction to bacterial membranes, con-
sistent with its highest bactericidal activity. These different interfacial structural features associated with
AMP binding shed light on the future development of active and biocompatible AMPs for infection and
wound treatments.

© 2023 The Authors. Published by Elsevier Inc. This is an open access article under the CC BY license (http://creativecommons.org/licenses/by/4.0/).

**1. Introduction**

Antimicrobial peptides (AMPs) are a promising class of antimicrobial reagents to fight against the ongoing global threat from multi-drug resistant bacteria [1,2]. It is widely accepted that AMPs kill bacteria by disrupting their membranes via a membrane-lytic mechanism, although many other mechanisms have also been reported (e.g. immunoregulation and intracellular targets like nucleic acids, ribosomes, proteins) [3,4]. Compared to the working mechanisms of traditional antibiotics, this membrane-targeted mode of action often leads to rapid killing so that microbes are less likely to become AMP-resistant [5]. Before reaching the cytoplasm of a Gram-negative bacterium (GNB), antimicrobials must penetrate across the entire cell envelop, including the asymmetric outer membrane (OM) consisting of the thick outer lipopolysaccharide (LPS) leaflet and thin inner phospholipid (PL) leaflet, the thin peptidoglycan cell wall, and the PL-based inner membrane (IM) [6]. Since OM is the outermost barrier to mediate the entry of antimicrobials, it has been increasingly recognized as a key component to determine the antimicrobial susceptibility and resistance [7-9]. However, many studies have focused on developing AMPs that are potent against the PL-dominant inner IM, while their specificity against the LPS-rich OM and the peptidoglycan (PDG)-dominated cell wall remains poorly understood. A clear understanding on how AMPs penetrate the bacterial OM will benefit future AMP design in the improvement of antimicrobial efficacy and cytotoxicity to host cells.

LPS, also known as endotoxin, is a distinguished molecule in the
bacterial OM and bears rather different physiochemical properties
to the PL molecules of host mammalian cells and other microbial
strains. Many natural PL molecules are relatively small
(<1000 Da), as they are comprised of 2-4 hydrophobic acyl chains
with a hydrophilic phosphate head group [10,11]. In contrast, a LPS
molecule has a lipid A group composed of 6 C₁₂-C₁₄ tails, a core
oligosaccharide containing some 10 sugar units and many anionic
phosphate groups, and a long O-antigen group covering numerous
repeating oligosaccharide units [12]. LPS molecules with/without
O-antigens are called smooth or rough LPS, respectively. A deep
rough LPS, which is a short version of LPS with about 3-7 sugar
units in the core oligosaccharide, has a molecular weight about
3000 Da. Therefore, a LPS molecule is significantly different to a
PL molecule, and it must interact differently with many membrane
penetrating antimicrobials. Furthermore, with the ever-increasing
interests in understanding how AMPs penetrate both bacterial IM
and OM, it is important to establish more realistic LPS-
incorporated bacterial membrane models to gain reliable insights
of membrane-based structural changes and interactions.

In spite of their large molecular size and strong hydrophobicity,
the physically assembled LPS molecules in the outer OM leaflet can
readily shade off from GNB to cause severe inflammatory
responses, requiring medical treatments to deactivate LPS and con-
trol inflammation [13,14]. Because cationic AMPs can neutralize
LPS and some of them may also recruit immune responses to over-
come inflammatory processes [15], AMPs have been considered as
potential candidates to not only kill antimicrobial resistant GNBs,
but also prevent LPS-induced inflammations. Thus, the mechanistic
processes of AMP-LPS interactions across the membranes have
attracted growing interests.

A simple but useful tool to study the interaction of AMP with
lipid membrane is to constitute a lipid monolayer with varied com-
positions at the air/liquid interface using a Langmuir trough. The
monolayer model has been proven to be advantageous in charac-
terizing how an AMP interacts with a model bacterial membrane
leafletlet as reported in many previous studies [16,17]. We have
developed the experimental capability to control model bacterial
PL membranes so that their interactions with AMPs can be studied
by techniques such as neutron reflection (NR) [18-20].

In this work, we have extracted LPS from an Rc mutant *E. coli* strain and studied its interactions with AMPs using a spread LPS monolayer model from Langmuir trough. Three AMPs including melittin, LL37, and G₃ were chosen to represent AMPs from different origins. Melittin and LL37 are natural AMPs and have been extensively studied due to their strong membrane-lytic capabilities [15,21]. Melittin is isolated from the honeybee venom. LL37 is the human innate defence AMP and belongs to the cathelicidin family. LL37 is released when neutrophils are stimulated. In contrast, G₃ (GIIKKIIKKIIKKI-NH₂) is a *de novo* designed AMP that adopts non-ordered secondary structure in neutral membrane environment but that transforms to α-helical conformation upon exposure to negatively charged bacterial membrane [22]. Such charge induced transition of molecular conformation enhances its membrane-lytic power against bacterial membranes.

NR was used to investigate the dynamic processes and struc-
tural disruptions of AMPs against the planar LPS monolayer model.
By using the H/D contrast variation strategy, NR can detect the
location and the amount of specific materials along surface normal
(perpendicular to the interface) under the resolution of 1–2 Å [23].
Therefore, NR can quantitatively determine the orientation of
lipids, penetration depth and adsorption capacity of AMPs before,
during, and after AMP-LPS interactions. Additionally, the use of
the spread lipid monolayer in NR has the advantages of easy prepa-
ration and higher depth resolution when combined with different
isotopic contrasts [24]. Our study thus provides a useful tool to
examine the membrane-lytic efficiency of AMPs to the bacterial
OM, providing constructive suggestions on future AMP design for
effective resistant bacterial killing and anti-inflammation
treatment.

**2. Experimental section**

**2.1. Materials**

Deuterated and hydrogenous DPPG (dipalmitoyl phosphatidyl-
glycerol) were purchased from Avanti Polar Lipids (purity > 98%).
Acyl chain deuterated DPPG is denoted as d62-DPPG or dDPPG,
while the hydrogenous version is hDPPG. An Rc mutant *Escherichia
coli* strain (*E. coli*, ATCC 43745) was fermented in H2O and D2O
media separately to produce normal and deuterium labelled bacte-
ria [25], followed by phenol/chloroform/petroleum ether extrac-
tion [26] to isolate deuterated and hydrogenous LPS samples
(denoted as dLPS and hLPS). Details of LPS production and charac-
terisation have been previously described [24].

Three peptides were synthesised by Shanghai TopPeptide Com-
pany (China) following an Fmoc solid-phase synthesis method (pu-
rity > 98%). The peptides were characterised by reversed-phase
high performance liquid chromatography (RP-HPLC) and matrix-assisted laser desorption/ionisation mass spectrometry (MALDI MS) [18]. No mixing of other peptide residues with varied amino acid sequences was observed. All other chemicals were obtained from Sigma-Aldrich Co. Ltd with purities > 99% without further purification unless otherwise stated.

## 2.2. Antimicrobial activity

Minimum inhibitory concentration (MIC) was used to determine the antibacterial susceptibility or resistance of AMPs [27]. *E. coli* (ATCC 43745) were cultured using the Mueller Hinton broth (MHB) at 37 °C overnight, followed by dilution in MHB to about $1 \times 10^6$ CFU/mL before adding antimicrobials. AMPs were dissolved in sterilised PBS buffer (10 mM phosphate buffer, 2.7 mM KCl, 137 mM NaCl, pH 7.4), and 2-fold diluted in 96-well plates at a volume of 100 µL solution per well. Aliquots of 100 µL bacterial solution were subsequently mixed with AMP solutions, and incubated at 37 °C overnight. Optical density at 600 nm (OD$_{600}$) was used to measure the turbidity of bacterial solutions after overnight treatments by AMPs. The fractional survival bacteria were calculated by: $(OD_{600, AMP} - OD_{600, 0%})/(OD_{600, 100\%} - OD_{600, 0\%}) \times 100\%$, where 0% and 100% denoted no bacterial growth sample (fresh broth) and 100% bacterial growth sample (without AMP or drug as control), respectively. The lowest peptide concentration to obtain 0% bacterial growth was determined as the MIC. Trilicate experiments were performed to estimate errors.

## 2.3. Haemolytic activity

Fractional haemoglobin release from human red blood cells (hRBCs, sourced from Rockland Immunochemical Inc) as treated by AMPs was used to determine AMP cytotoxicity [22]. Every 100 µL hRBCs (4% v/v diluted in PBS) was mixed with 100 µL AMP (2-fold diluted in PBS) in 96 well-plates, and incubated at 37 °C for 1 h. The unbroken hRBCs and haemoglobin contained solutions were separated by centrifugation of 96 well-plates at $1000 \times g$ for 5 min. Every 100 µL of the supernatant in each well was transferred into a new 96-well plate, followed by measuring optical absorbance at 450 nm ($A_{450}$). The fractional haemolysis of each sample was calculated by: $(A_{450, AMP} - A_{450, 0\%})/(A_{450, 100\%} - A_{450, 0\%}) \times 100\%$, where 0% and 100% denoted 0% hRBC lysis (without AMP) and 100% hRBC disruptions (treated by 0.1% Triton X-100), respectively. Triplicate experiments were performed to estimate errors.

## 2.4. Circular dichroism (CD)

Secondary structures of AMPs were analysed by a Chirascan Series Spectrometer (Applied Photophysics, UK). AMPs were first dissolved in PBS buffer at a concentration of 0.1 mM. CD measurements were then taken on AMPs mixed with/without LPS small unilamellar vesicles (SUVs, 1 mM). LPS SUVs were produced via a syringe extrusion strategy as previously described [28]. The sample path length was 0.2 mm and the scanned wavelength range was 190–260 nm. Each experiment was performed three times.

## 2.5. Proteolytic stability experiments

Matrix metalloproteinase-2 (MMP-2) and matrix metalloproteinase-9 (MMP-9) were selected as typical collagenases to assess the proteolytic stability of AMPs. RP-HPLC (reverse phase – high pressure liquid chromatography) analysis was used to detect the peptides. MMP-2 and MMP-9 were dissolved in PBS buffer at 20 µg/mL to mimic the high protease environment in the

chronic wounds. G$_{3}$, melittin, and LL37 were prepared in PBS buffer at 2 mg/mL, followed by 1/1 (v/v) mixing with MMPs. After interacting for 0.5, 1, 2, 4, 8, and 24 h, samples were 10-fold diluted in 0.1% trifluoroacetic acid (TFA) containing water, followed by injections into a C$_{18}$ reversed-phase column for HPLC experiments. Gradient exchange of eluent A (H$_{2}$O + 0.1% TFA) and eluent B (acetonitrile + 0.1% TFA) were used to flush the HPLC system by following 95% A + 5% B to 5% A + 95% B from 1 min to 40 min. The UV absorbance at 214 nm was used to identify peptides. The percentages of remaining peptides were determined by the remaining peptide (%) = $I_t/I_0$, where $I_t$ and $I_0$ are the integrated peak areas at different time points and 0 h, respectively. The error of each measurement was estimated by 10% due to variations from peak integration.

## 2.6. Lipid monolayers and neutron reflection

Spread RcLPS and DPPG monolayers were constructed using a Langmuir trough as described previously [29]. Some 70 mL of PBS buffer were poured into the trough with the surface area of $12.5 \ cm \times 15 \ cm$ (Nima Technology, UK). RcLPS powders were first suspended in a chloroform/methanol/water solvent solution (60:35:5, v/v/v), and DPPG powders were dissolved in the chloroform/methanol mixture (90:10, v/v). The lipid solutions were carefully placed onto the water surface, followed by 5 min evaporation of the organic solvent. The surface pressure was then controlled at 28 mN/m for at least 15 min. The Langmuir trough was switched to an area control mode, followed by AMP injection in the liquid subphase using a bent needle under the barrier. The injection must be spread uniformly underneath the spread lipid monolayer and care was taken to hold the needle tip close to the trough bottom so that the disruption to the lipid monolayer was kept minimal. The final concentrations of G$_{3}$, melittin, LL37 were at 3 µM, 1 µM, and 1 µM, respectively. The lipid monolayers before and after exposure to the AMPs were characterised by NR.

The SURF reflectometer at the ISIS Pulsed Neutron Source (Didcot, UK) and the FIGARO reflectometer at the ILL Continuous Neutron Source (Grenoble, France) were used to perform NR measurements, with the same setup as described previously [19]. Deuterated or hydrogenous lipids spread on D$_{2}$O or on null reflecting water (NRW, D$_{2}$O/H$_{2}$O = 8/92, v/v, SLD = 0) were the four isotopic contrasts of a typical dataset. Dynamic measurements involving AMP binding to the spread lipid monolayer were performed under the NRW contrasts by the high flux FIGARO reflectometer [30] at the incident angle of 0.62. Each dynamic run was scanned by NR for 4 min. After reaching equilibrium, each monolayer was scanned by either the SURF reflectometer at 3 incident angles (0.35°, 0.65°, and 1.5°), or the FIGARO reflectometer at 2 incident angles (0.62°, 3.8°) and fixed wavelength resolution of 7%. The measured neutron reflectivity $R(Q)$ was plotted as a function of momentum transfer (Q). Both instruments cover the Q-range of 0.01–0.20 Å$^{-1}$, the upper value being determined by the sample background, as analysed in model fittings. The environmental temperature was 20–21 °C.

A dataset with four isotopic contrasts were simultaneously fitted by a Motofit package based on the optical matrix formula [31]. A 2-layer model (lipid tail and head layers) or a 3-layer model (lipid tail, head, and AMP adsorption layers) were used to analyse the monolayer system in the absence and presence of AMPs, respectively. Fitted parameters include layer thickness, $\tau$, and scattering length density (SLD or $\rho$). Roughness of 1 Å was used to smooth interfacial transition. The best-fit SLDs and sample volume fractions in each layer follow: $\rho_{fit} = \sum_n \rho_n \varphi_n$, where $\varphi_n$ is the volume fraction of different species, e.g. lipid tail, lipid head, peptide, and solution, and $\sum_n \varphi_n = 1$. The surface concentration (in the unit
of mg/m²) of a component was calculated by: $\frac{\tau\phi M}{VN_A}$, where M and V are the theoretical mass and volume of a molecule, and $N_A$ is the Avogadro's number.

# 3. Results and discussion

## 3.1. Antibacterial activity, cell toxicity and stability of AMPs

The three amphiphilic peptides possess good water solubility and do not self-assemble into supramolecular structures over the antimicrobial concentration ranges studied [18,32,33]. Thus, they share similar physiochemical properties and work as water-soluble antimicrobials. Their relative amphiphilic behaviour was evaluated by assessing their hydrophobicity and cationicity per amino acid (Table 1). The mean hydrophobicity was found to be 0.476, 0.511 and 0.201 for G₃, melittin and LL37, respectively. G₃ has an intermediate hydrophobicity but is the most cationic AMP with +7 charges out of the 14 residues; melittin is the most hydrophobic AMP with intermediate positive charge density (+6 out of the 26 residues), and LL37 is the most hydrophilic one with the least cationic charge density (+7 out of the 37 residues).

RcLPS used in this study for biophysical characterisations was extracted from a gene modified *E. coli* strain (ATCC 43745). Thus, MIC measurements against this strain were performed to test its antimicrobial susceptibility to the three AMPs. As shown in Fig. 1A, G3 and melittin had similar MICs (10–15 µg/mL), whilst LL37 had a higher MIC of 35 µg/mL. Because the three AMPs have significantly different molecular weights, the use of the mass concentration rather than molarity benefits the evaluation of the amount of peptides used to disrupt bacterial membranes and kill bacteria. The MICs measured against the Rc mutant *E. coli* strain are well consistent with the previous MICs to the standard *E. coli* strains [18], indicating that reducing the uncharged polysaccharide groups of LPS does not significantly influence the AMPs' bactericidal activity. Meanwhile, G₃ and melittin are broad-spectrum antimicrobials, but LL37 shows weaker antimicrobial activity to kill some Gram-positive bacteria (e.g. S. aureus) [18].

Biocompatibility is vital to the development of AMPs with clinical potential, which is commonly tested by measuring their haemolytic activity. As shown in Fig. 1B, melittin is significantly more toxic than G₃ and LL37, as it causes up to 80% of hRBC's rupture at just 10 µg/mL. This is equal to its MIC value against *E. coli*, indicating that melittin is not selective against bacterial membranes. In contrast, LL37 and G₃ are rather biocompatible, with EC₅₀ (effective concentration to cause 50% haemoglobin leakage) of 2300 µg/mL and 1200 µg/mL, respectively. The selectivity indices (SI = EC₅₀/MIC) of LL37 and G₃ were about 65 and 92, respectively. A higher selectivity index indicates that the candidate antimicrobial is safer and more effective. High antimicrobial efficacy and high selectivity index are essential when considering further AMP development toward clinical application.

The antibacterial activity of an AMP is determined by its membrane binding and penetrating capability that is often associated with its secondary structural changes. AMPs are soluble in water as monomers or micelles, but can readily bind to negatively

charged bacterial membranes [19]. To reveal how AMPs initiate membrane binding, their secondary structures in buffer and LPS vesicles were firstly examined by CD. As shown in Fig. 1C &1D, melittin and G₃ are in random coils in the aqueous buffer, featured by a single minimum around 200 nm. In contrast, LL37 adopts α-helical conformation in the physiological buffer, characterised by the CD spectrum with a positive peak at 193 nm and dual negative peaks at 208 nm and 222 nm. When contacting *E. coli*, cationic AMP molecules are attracted to the polyanionic LPS surface of the OM via electrostatic interaction. This process is mimicked by exposing the AMPs to the LPS vesicles. As shown in Fig. 1D, all three AMPs adopt α-helical structures when exposed to the LPS surface. The adoption of α-helical conformation has also been reported from other AMPs after interacting with anionic lipids or surfactants [10]. As shown in Figure S1, an α-helical G₃ molecule has two distinct faces, with one face containing lysine residues and the other containing isoleucine residues, making it better amphiphilically balanced than in the random coil state. The CD spectra have thus revealed that melittin and G₃ under the tested conditions can transform from random coils in aqueous buffer solution to α-helical structures once bound onto the LPS membrane. In contrast, LL37 already adopts α-helical conformation in water and its binding to the LPS surface leads to the strengthening of its α-helical state.

The proteolytic degradation of AMPs is a key challenge that limits their applications. MMP-2 and MMP-9 are common endopeptidases, which can degrade extracellular matrix components and AMPs in wounds [34]. Their expressions in chronic wounds are commonly higher than acute wounds, with the typical concentrations ranging from ng/mL to µg/mL [35,36]. Here, a high concentration of 20 µg/mL of either MMP-2 or MMP-9 was interacted with AMPs to evaluate their proteolytic stability mimicking the harsh chronic wound conditions. After interacting with MMPs for different times, the amounts of remaining peptides were assessed by RP-HPLC via integration of the UV absorbance peaks (Figure S2). As shown in Fig. 1E and 1F, G₃ was relatively stable as exposed to either MMP-2 or MMP-9 proteases. More than 80% of G₃ remained after interacting with MMPs for 8 h. In contrast, melittin and LL37 lost more than 50% of components within 0.5 h of interactions. After 24 h of interactions, MMPs degraded most AMP molecules, whilst G₃ had the most remaining molecules among the three AMPs studied. The use of glycine, isoleucine, and lysine amino acids to produce the simple G₃ peptide makes it less likely to be degraded by enzymes, compared to the naturally occurring melittin and LL37 with relatively complex sequences.

## 3.2. Dynamic binding of AMP to LPS/PL monolayer

To further evaluate the extent of membrane disruptions, AMPs, G₃, melittin and LL37 were brought into contact with the RcLPS and DPPG monolayers, mimicking the OM and IM membrane leaflets, respectively. These lipid models bear the key characteristic features of the membranes with different charges. Although natural bacterial membranes are comprised of multiple components rather than a single negatively charged one, mixed lipid models also bring in other complications [37–39]. Features such as membrane

<table><caption>**Table 1**
Parameters of AMPs. The mean hydrophobicity was calculated using a HeliQuest helical property calculator (https://heliquest.ipmc.cnrs.fr/).</caption><thead><tr><th>Peptide</th><th>Sequence</th><th>Molecular weight</th><th>Charge</th><th>Mean hydrophobicity</th></tr></thead><tbody><tr><td>G<sub>3</sub></td><td>G(IIKK)<sub>3</sub>I-NH<sub>2</sub></td><td>1635.2</td><td>+7</td><td>0.476</td></tr><tr><td>Melittin</td><td>GIGAVLKVLTTGLPALISWI<br>KRKQQ-NH<sub>2</sub></td><td>2846.5</td><td>+6</td><td>0.511</td></tr><tr><td>LL37</td><td>LLGDFFRRKSKEKIGKEFKRI<br>VQRIKDFRLNLPVRTES-NH<sub>2</sub></td><td>4492.3</td><td>+7</td><td>0.201</td></tr></tbody></table>
<figure><img src="image_5.png" /><figcaption>**Fig. 1.** Bioactivities, secondary structures, and proteolytic stabilities of AMPs. (A) Survival fractions of the Rc mutant *E. coli* as treated by G₃ (black square), melittin (red circle), and LL37 (blue diamond) over different concentrations. (B) Fractional haemoglobin releases from AMP disrupted red blood cells. Circular dichroism spectra of the three AMPs: (C) in PBS buffer and (D) after interactions with LPS vesicles. CD spectra were plotted by mean residue ellipticity as a function of optical wavelength. (E-F) Proteolytic stability changes of three AMPs after interacting with (E) MMP-2 and (F) MMP-9 at different time points. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>

instability, partial mixing or formation of lipid rafts are interesting as they often exist in real membranes, but they impose further challenges in measurements and data analysis. The use of the fully charged LPS and DPPG membranes enable us to investigate the extreme charge-driven AMP-lipid membrane interaction and determine the subsequent membrane damage under AMP binding. In addition, evaluating the bioactivity of AMP from the lipid monolayer instead of the bilayer model helps simplify data analysis and achieve high in-depth structural resolution from NR.

As shown in **Figures S3 & S4**, neutron reflectivity with sufficiently high statistical quality can be measured over the low momentum transfer (Q) range in a few minutes on FIGARO [30]. This technical capability allows us to examine how AMP binding onto LPS and DPPG monolayers occurs with time and causes changes in interfacial composition. Each spread lipid monolayer was formed at the air/liquid interface, with the initial surface pressure controlled at 28 mN/m. Once the surface pressure stabilised, AMP was uniformly injected in the liquid subphase over a period of 1 min. As shown in **Fig. 2A & 2B**, air and the water phase do not contribute to the neutron signal in null reflecting water (NRW). Thus, only the components with non-zero SLD in the lipid film can be detected. Through parallel NR runs of acyl chain deuterated (d-lipid) and hydrogenous (h-lipid) lipids on NRW, the amount of lipid removal and AMP binding can be readily determined. NR measurements on D₂O help highlight the lipid head groups immersed under water, and these parallel runs obtained under the same lipid/AMP combination are useful for structural analysis (**Fig. 2A & 2B**). Here, NR profiles at the low-Q region of

0.01–0.04 Å⁻¹ were collected every 4 min. Binding of the hydrogenous AMP to the d- or h-lipid monolayer on NRW was measured in parallel and these two isotopic contrasts were analysed simultaneously by a single layer model to obtain the thickness and average SLD of the AMP bound membrane. The analysis led to volume fractions of lipid and AMP, with the best fit parameters shown in **Tables S1 & S2**.

As shown in **Figure S3**, reflectivity *R*(*Q*) of the deuterated lipid runs on NRW continuously decreases, indicating the loss of lipid at the interface as a result of AMP binding. In contrast, *R*(*Q*) of the hydrogenous lipid runs increases from AMP binding. All three AMPs achieve most of their binding onto the LPS surface within their first dynamic measurements (8 min); some lipids are clearly removed from the water surface (**Fig. 2C**). G₃ reaches its fast equilibration when binding to the LPS monolayer within the first 20 min. This process leads to the removal of 29% LPS molecules and G₃ adsorption of 0.73 mg/m². The binding of LL37 continues up to about 60 min, leading to 15% of LPS molecular loss and 2.57 mg/m² of peptide binding. In contrast, the dynamic binding of the melittin-LPS system is more complex and the process persists over 60–80 min. 1.82 mg/m² melittin binds to the LPS within the first 8 min of the dynamic run, but only removes 22% of the LPS molecules. After reaching equilibrium around 60 min, further 26% LPS molecules is dissolved from melittin binding, with only 1.44 mg/m² (52%) of LPS and 1.52 mg/m² of AMP remain at the interface. Taken together, G₃ causes effective LPS monolayer damage with the least amount of peptides bound onto the LPS membrane, equivalent to 48% of melittin and 28% of LL37 bound.
<figure><img src="image_6.png" /><figcaption>**Fig. 2.** Contrast variations and dynamic measurements in NR. (A-B) Schematic representations of (A) LPS and (B) DPPG monolayers on the water surface. Colour codes from white to black indicate the neutron scattering length density values from low to high. (C-D) Surface concentration dynamic changes of (C) LPS, (D) DPPG, and AMPs as determined by NR low-Q region using NRW contrasts. Detailed fitting parameters are shown in **Figures S3 & S4**.</figcaption></figure>

To compare the susceptibility difference between the LPS and PL monolayers, the three AMPs are also bound to the DPPG monolayer. Our previous studies have shown that AMPs selectively bind to the anionic PG lipids with little affinity to the uncharged PC lipids [40]. This corroborates well with the observation of low MICs and high EC50s from antimicrobial and cytotoxicity studies. The results from the current work reveal that the dominant interaction occurs mainly over the first NR measurements (8 min). However, peptide binding tends to equilibration over 30–40 min (Fig. 2D). This is again shown as the quick decrease of $R(Q)$ from the dDPPG monolayer and slow increase from the hDPPG monolayer (Figure S4). $G_3$, melittin, and LL37 remove 16%, 20%, 4% of DPPG, and achieve the equilibrium adsorbed amount of 0.96, 2.86 and 2.24 $mg/m^2$, respectively. As indicated already, all three AMPs cause weaker damage to the DPPG monolayer than to the LPS monolayer. These AMPs have multiple cationic charges and are rather amphiphilic. They may drag the bound DPPG molecules into bulk solution in a manner dependent on AMP concentration and peptide-to-lipid ratio [41]. $G_3$ again has the lowest adsorption on the DPPG membrane that is 34% and 43% mass of melittin and LL37, respectively. These differences are however linked to the low molecular weight of $G_3$. The differences are much smaller in the micromolar unit (Figure S5).

### 3.3. Structures of the LPS and DPPG monolayers before and after AMP binding

Structural features of the LPS and DPPG monolayers were first analysed by NR under 4 isotopic contrasts, d-lipid/D₂O, h-lipid/D₂O, d-lipid/NRW, and h-lipid/NRW. As d-lipid has deuterated acyl chains and hydrogenous head, the measurement of d-lipid/NRW defines the thickness of the chain layer and area per molecule. The measurement of d-lipid/D₂O defines the entire lipid layer thickness as the immersion of the hydrogenous head in D₂O helps highlight the head layer. Because the SLD of the hydrogenous chain is very low, the measurement of h-lipid/D₂O helps determine the head layer with little influence from the hydrogenous chain layer. All reflectivity profiles shown in Figure S6 were measured from

lipid monolayers kept at 28 mN/m on either D₂O or NRW, and the continuous lines represent the best 2-layer fits, with the top 1st layer representing the lipid tails in air and the bottom 2nd layer representing the head groups immersed in water. The $R(Q)$ profiles in D₂O are close but much higher than those on NRW. Furthermore, the reflectivity measured from the deuterated LPS is higher than the one from the hydrogenous LPS under each solvent contrast. Simultaneous model fitting of the 4-contrast datasets led to thicknesses of 13 Å and 23 Å for the tail and head layers and volume fractions of 0.88 and 0.51, respectively. In contrast, the DPPG monolayer had a thicker tail layer of 18 Å and a thinner head layer of 10 Å, with the volume fractions of 1.00 and 0.59, respectively. These differences are consistent with the longer $C_{16}$ acyl chain and smaller PG head group. The area per molecule was 198 Å² for LPS and 48 Å² for DPPG in their respective monolayer. These lipid measurements are reproducible and consistent with previous observations [24,29].

After AMP injection and equilibrium as described previously (Fig. 2), NR reflectivity profiles were measured following equilibrated AMP binding to the LPS monolayer under the same 4 isotopic contrasts (Figure S6). AMP binding caused dramatic changes in reflectivity profiles under the same water and lipid contrasts. Model fitting of the 4-contrast datasets (Table 2) from the same 2-layer model revealed that the thicknesses of the LPS tail and head layers remained largely unchanged upon $G_3$ binding. However, both the acyl chain and head layers contained $G_3$, with its volume fractions of 0.29 and 0.10 and surface concentrations of 0.46 and 0.27 $mg/m^2$, respectively.

In contrast, binding of the two natural AMPs significantly thickened the LPS monolayer. This is characterised by the increased separation between the D₂O profiles and the occurrence of the weak interference fringes (Fig. 3C & 3E, black squares and green circles). Simultaneous fitting to the 4 parallel contrasts led to 0.86 and 0.18 $mg/m^2$ of melittin inserted into the LPS tail and head layers, respectively. Melittin binding caused removal of 48% of LPS molecules from the monolayer. Interestingly, about 5% of LPS (0.06 $mg/m^2$) aggregated with 0.48 $mg/m^2$ of melittin that became suspended under the main LPS head layer to form layers of lipid/pep-
<caption>**Table 2**</caption>

Best-fit structural parameters for the LPS monolayer without and with AMPs as obtained from NR. $\tau$ denotes layer thickness, $\varphi$ denotes volume fraction and $\Gamma$ denotes mass per unit area.

<table><thead><tr><th>Layer</th><th>τ (Å)</th><th>φ<sub>lipid</sub></th><th>φ<sub>peptide</sub></th><th>φ<sub>solvent</sub></th><th>Γ<sub>lipid</sub> (mg/m<sup>2</sup>)</th><th>Γ<sub>peptide</sub> (mg/m<sup>2</sup>)</th></tr></thead><tbody><tr><td>LPS</td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Tail</td><td>13 ± 1</td><td>0.88 ± 0.08</td><td>-</td><td>-</td><td>0.92 ± 0.09</td><td>-</td></tr><tr><td>Head</td><td>23 ± 2</td><td>0.51 ± 0.05</td><td>-</td><td>0.49 ± 0.04</td><td>1.82 ± 0.18</td><td>-</td></tr><tr><td>LPS + G<sub>3</sub></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Tail</td><td>13 ± 1</td><td>0.62 ± 0.06</td><td>0.30 ± 0.03</td><td>-</td><td>0.65 ± 0.06</td><td>0.46 ± 0.04</td></tr><tr><td>Head</td><td>23 ± 2</td><td>0.36 ± 0.04</td><td>0.10 ± 0.01</td><td>0.54 ± 0.05</td><td>1.28 ± 0.12</td><td>0.27 ± 0.03</td></tr><tr><td>LPS + MLT</td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Tail</td><td>13 ± 1</td><td>0.46 ± 0.05</td><td>0.50 ± 0.05</td><td>-</td><td>0.48 ± 0.05</td><td>0.86 ± 0.08</td></tr><tr><td>Head</td><td>23 ± 2</td><td>0.27 ± 0.03</td><td>0.05 ± 0.01</td><td>0.68 ± 0.06</td><td>0.94 ± 0.09</td><td>0.18 ± 0.01</td></tr><tr><td>Peptide</td><td>54 ± 5</td><td>0.01 ± 0.01</td><td>0.07 ± 0.01</td><td>0.92 ± 0.08</td><td>0.01 ± 0.01</td><td>0.48 ± 0.04</td></tr><tr><td>LPS + LL37</td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Tail</td><td>13 ± 1</td><td>0.75 ± 0.07</td><td>0.22 ± 0.02</td><td>-</td><td>0.78 ± 0.08</td><td>0.38 ± 0.03</td></tr><tr><td>Head</td><td>23 ± 2</td><td>0.43 ± 0.04</td><td>0.20 ± 0.02</td><td>0.37 ± 0.04</td><td>1.54 ± 0.15</td><td>0.60 ± 0.06</td></tr><tr><td>Peptide</td><td>28 ± 2</td><td>-</td><td>0.44 ± 0.04</td><td>0.56 ± 0.05</td><td>0</td><td>1.62 ± 0.16</td></tr></tbody></table>

<figure><img src="image_7.png" /><figcaption>**Fig. 3.** NR reflectivity data and the best model fits to the LPS monolayers after interacting with AMPs. The 4-contrast NR profiles and fitted volume fraction distributions of LPS monolayers after interacting with (A-B) G<sub>3</sub>, (C-D) melittin, and (E-F) LL37. In A/C/E, experimental reflectivity values are plotted as a function of momentum transfer (Q), and the best fitted values are presented by continuous lines. In B/D/F, volume fraction distributions are plotted across the z-axis, with representative cartoons to demonstrate the extent of membrane disruptions and AMP binding. Black curves and grey circles are acyl chains and core oligosaccharides of LPS, respectively. Red circles are peptide aggregates, comprised of peptides denoted by red helices bound to the membrane surface or inserted into the lipid tails. Peptide orientations are plotted assuming the barrel-stave pore, toroidal pore, and carpet models [43]. White and cyan backgrounds are air and water, respectively (Data from FIGARO at ILL). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>

tide stacks with a thickness of 54 Å. The co-assembly of these layered stacks from anionic LPS and cationic melittin must be driven by strong electrostatic interaction but because of their strong hydrophobicity these aggregates must remain largely insoluble in water and become accumulated outside the lipid membrane. Lipid stacking with AMP suggests strong AMP-lipid interactions, which could be common to other AMPs when bound to lipid membranes, especially with the hydrophobic AMPs under high peptide-to-lipid

ratios [42]. Although melittin had a low volume fraction (0.07) of stacking with LPS, the total surface confined amount in this additional layer was 0.48 mg/m² due to the thick stacking formed.

However, LL37 at the same concentration showed the weakest activity against the LPS monolayer with only 15% of lipid loss, corroborating with its weakest antimicrobial activity. Only 0.38 mg/m² of LL37 inserted into the LPS tail layer, with most AMPs being confined to the LPS head layer (0.60 mg/m²) and the outer
condensed AMP layer (1.62 mg/m²) under the lipid head group layer.

In contrast, the three AMPs caused rather weak structural dis-
ruptions to the DPPG monolayer, evident from low lipid dissolu-
tion, low depth penetration in the lipid tail layer and relatively
high AMP binding outside the lipid head layer. As shown in Fig. 4
A and B, G₃ became inserted into the DPPG membrane and
stretched the lipid tail and head layers to 20 Å and 14 Å, with
the amount of peptide binding of 0.28 mg/m² and 0.27 mg/m² in
the two layers, respectively. A protruded layer with a thickness
of 10 Å was also found under the DPPG head layer. As this addi-
tional peptide layer was not observed when G₃ bound to the LPS
monolayer, the difference might indicate that the G₃ aggregate size
spans between the PG head and LPS core oligosaccharide, i.e., 10–
23 Å. Thus, G₃ must be well embedded in the LPS membrane but
protruded outside the DPPG head layer.

Melittin removes some 20% DPPG, which is significantly less
than 48% of the removal of LPS. There is no evidence of the forma-
tion of the thick AMP-lipid stacks either. A layer of condensed
melittin aggregates with a thickness of 29 Å and volume fraction
of 55% are adsorbed outside the DPPG head layer (Fig. 4 C/D).
LL37 only dissolves 4% of DPPG but expands the DPPG monolayer
from 28 Å to 35 Å. It also forms an additional AMP-lipid layer of
23 Å under the lipid head region with the volume fraction of 40%
(Fig. 4 E/F). The best-fit structural parameters of the DPPG mono-
layer with and without AMPs are listed in Table 3. Detailed SLD

distributions of LPS and DPPG monolayers with and without AMP
binding are shown in **Figures S7 & S8**.

3.4. Link between AMP binding to LPS/DPPG monolayers and antimicrobial efficacy

A potent and efficient AMP is likely to disrupt or dissolve lipids using minimal peptide molecules. Here, the overall peptide adsorption across either the LPS or the DPPG monolayers is illustrated in Fig. 5 for comparison. The total mass of adsorbed amount from melittin and LL37 is significantly higher than G₃. It can also be seen clearly from Fig. 5 that melittin and LL37 are strongly associated with the lipid head groups, but they are less well inserted into the tail region. This is well expected from their long sequences and strong aggregation propensity. In contrast, G₃ displays relatively weaker membrane binding in terms of the total adsorbed amount, but its stronger membrane-lytic activity is well illustrated from its deep insertion into the tail region and less association within and under the head group region of the model lipid membranes. These combined interfacial partitioning actions make the designed short G₃ more effective at killing bacteria with competitively low MICs.

The high level of membrane binding of melittin and LL37 must
be associated with in-membrane aggregation, especially around
the lipid head group region, but these aggregates do not appear

<figure><img src="image_8.png" /><figcaption>**Fig. 4.** NR reflectivity data of DPPG monolayers and best model fits after interacting with AMPs. DPPG monolayers at the initial surface pressure of 28 mN/m were brought to interact with G₃ (A-B), melittin (C-D), and LL37 (E-F). Experimental data and model fits of reflectivity profiles plotted against Q are presented in A/C/E. Volume fraction distributions across the z-axis and schematic diagrams of the best-fit interfacial layers are illustrated in B/D/F. Black curves and purple circles are acyl chains and head groups of DPPG, respectively. Red circles are peptide aggregates, comprised of peptides as represented by red helices bound to the membrane surface or inserted into the lipid tails. White and cyan backgrounds are air and water, respectively. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>
<table><caption>**Table 3**
Structural parameters of the DPPG monolayer with and without AMPs as obtained from NR, with τ denoting layer thickness, φ lipid volume fraction and Γ weight per unit area.</caption><thead><tr><th>Layer</th><th>τ (Å)</th><th>φ<sub>lipid</sub></th><th>φ<sub>peptide</sub></th><th>φ<sub>solvent</sub></th><th>Γ<sub>lipid</sub> (mg/m<sup>2</sup>)</th><th>Γ<sub>peptide</sub> (mg/m<sup>2</sup>)</th></tr></thead><tbody><tr><td>DPPG</td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Tail</td><td>18 ± 2</td><td>1.00 ± 0.05</td><td>–</td><td>–</td><td>1.46 ± 015</td><td>–</td></tr><tr><td>Head</td><td>10 ± 1</td><td>0.59 ± 0.06</td><td>–</td><td>0.41 ± 0.04</td><td>1.04 ± 0.10</td><td>–</td></tr><tr><td>DPPG + G<sub>3</sub></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Tail</td><td>20 ± 2</td><td>0.77 ± 0.08</td><td>0.12 ± 0.01</td><td>0.09 ± 0.04</td><td>1.2 ± 0.1</td><td>0.28 ± 0.02</td></tr><tr><td>Head</td><td>14 ± 1</td><td>0.36 ± 0.05</td><td>0.16 ± 0.02</td><td>0.48 ± 0.05</td><td>0.88 ± 0.08</td><td>0.27 ± 0.02</td></tr><tr><td>Peptide</td><td>10 ± 1</td><td>–</td><td>0.35 ± 0.04</td><td>0.65 ± 0.07</td><td>–</td><td>0.41 ± 0.04</td></tr><tr><td>DPPG + MLT</td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Tail</td><td>18 ± 2</td><td>0.75 ± 0.07</td><td>0.15 ± 0.03</td><td>0.05 ± 0.03</td><td>1.1 ± 0.1</td><td>0.35 ± 0.03</td></tr><tr><td>Head</td><td>13 ± 1</td><td>0.36 ± 0.03</td><td>0.27 ± 0.03</td><td>0.37 ± 0.03</td><td>0.77 ± 0.08</td><td>0.45 ± 0.04</td></tr><tr><td>Peptide</td><td>29 ± 2</td><td>–</td><td>0.55 ± 0.06</td><td>0.45 ± 0.05</td><td>–</td><td>2.1 ± 0.2</td></tr><tr><td>DPPG + LL37</td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Tail</td><td>22 ± 2</td><td>0.73 ± 0.07</td><td>0.18 ± 0.01</td><td>0.02 ± 0.01</td><td>1.3 ± 0.1</td><td>0.52 ± 0.05</td></tr><tr><td>Head</td><td>13 ± 1</td><td>0.41 ± 0.04</td><td>0.30 ± 0.02</td><td>0.29 ± 0.04</td><td>0.93 ± 0.09</td><td>0.51 ± 0.05</td></tr><tr><td>Peptide</td><td>23 ± 1</td><td>–</td><td>0.41 ± 0.05</td><td>0.59 ± 0.05</td><td>–</td><td>1.2 ± 0.1</td></tr></tbody></table>

<figure><img src="image_9.png" /><figcaption>**Fig. 5.** Surface concentrations of the three AMPs embedded in different interfacial regions of the LPS and DPPG layers. G3 (black), melittin (red), and LL37 (blue) adsorbed in lipid tail, lipid head, and outer AMP layers under the head layer of (A) LPS and (B) DPPG monolayer. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>

to contribute to high antimicrobial efficacy. However, the hydrophobic melittin does display strong insertion and aggregation into the lipid tail region as well, resulting in high antimicrobial potency. The downside of melittin is its lack of selectivity as the strong hydrophobicity makes it also capable of attacking host cell membranes, eliciting cytotoxicity. High hydrophobicity may also cause other issues such as tendency to aggregate in solution and lack of solubility, both of which reduce bactericidal efficiency through clustering or aggregating in buffer solution and broth

and with bacteria. Nevertheless, these hydrophobic AMPs can be modified to form supramolecular self-assembly based hydrogels that can be applied in controlled release applications, e.g. as antibacterial creams [44–46].

Strong bacterial membrane binding without effective penetration may explain the high antibacterial MICs of many natural AMPs. Thus, it is worth to compare their adsorption behaviour before further biological screening in their development pathway to clinical application. In the NR experiments, the final peptide concentrations as injected in the solution were 3 µM (4.9 µg/mL) of G3, 1 µM (2.8 µg/mL) of melittin, and 1 µM (4.5 µg/mL) of LL37. Considering the liquid volume deposited in the Langmuir trough was 70 mL, the masses of injected peptides in the solution were 0.34, 0.20, and 0.32 mg of G3, melittin, and LL37, respectively. The adsorbed peptides that aggregated with the lipid monolayers were determined by NR (Table S3). Thus, the fraction of total peptides in the solution that interacted with surface lipids was calculated by the amount of surface adsorbed AMP over the total AMP in the solution. The injected G3, melittin, and LL37 had proportions of 0.044, 0.14 and 0.16 that bound to the LPS membrane, and 0.05, 0.26 and 0.13 that bound to the DPPG membrane, respectively. Peptide molecules that aggregate with the negatively charged bacterial membranes may not be available to contribute to further bactericidal effects. Thus, membrane association must lead to the decrease of the AMP solution concentrations available for further antimicrobial actions. The surface mass ratios of AMP and lipid (denoted in mg/mg) can indicate the affinity of an AMP and its consumption by bacterial membranes. Mass ratios of G3/LPS, MLT/LPS and LL37/LPS were found to be 0.38, 1.01, and 1.12, respectively, and those of the G3/DPPG, MLT/DPPG and LL37/DPPG to be 0.46, 1.41, and 0.94, respectively. These data indicate that each unit of anionic lipids consumes significantly higher naturally occurring melittin (MLT) and LL37 but far less G3, showing that in addition to its superior membrane-lytic capability G3 is also efficient and effective for further antimicrobial actions.

It must be noted that these AMP consumptions as outlined above may be underestimated as peptides can also aggregate with ingredients in the broth and buffer solutions, or proteins and nucleic acids of bacteria, or adsorb on the plastic surface of 96-well plates. Under the simplest estimate of the peptide-lipid binding, the rationally designed G3 can kill a given type of bacteria with a small amount of molecules. This ensures its concentration in the solution to remain higher than its MIC, instead of quick decline due to consumptions by bacterial membranes and environments.

These analyses also demonstrate the importance of selecting broth and bacterial density during the MIC measurements. The current method used about $1 \times 10^6$ CFU/mL of the starting bacterial concentration before mixing with the antimicrobials
[27]. It can be expected that the measured MICs would decrease with $1 \times 10^5$ CFU/mL of bacteria, and increase with $1 \times 10^7$ CFU/mL of bacteria.

The interaction intensities of the three AMPs against the LPS leaflet determine their ability to kill *E. coli*. If the bacterial IM is directly disrupted to trigger the irreversible cytoplasmic leakage, the bactericidal effect may well occur within a short exposure time [47]. However, bacterial death is inevitable, as the leakage is driven by the osmotic pressure once the OM structure as transmembrane protection is broken [48]. The lipid monolayer models used in this study consist of full anionic LPS and DPPG, which must be susceptible to the cationic AMPs.

In the absence of other uncharged lipids, this work investigated the extreme interactions of AMP and lipid pairs to determine the most intensive membrane ruptures following each AMP binding, but real bacterial membranes carry large proportions of zwitterionic lipids with PC and PE head groups. A standard *E. coli* bacterium may contain up to 80% PE lipids, making the charge density significantly lower than the DPPG membrane model [10,49]. It is thus more appropriate for the bacterial IM to be comprised of 30% PG and 70% PC or PE [50,51]. In contrast, the bacterial OM model may be covered by 75% LPS [25,52]. Given that the three AMPs disrupted the LPS monolayer more drastically than the DPPG monolayer, they must be more destructive to the OM of the real *E. coli* than their IM. Interestingly, many previous AMP designs have focused on evaluating the ability of them to penetrate and disrupt the phospholipid bilayers of the bacterial IM, but ignored their interactions with the bacterial OM and cell wall components. These studies have however been limited by lack of reliable LPS and peptidoglycan samples and representative lipid models. Meanwhile, if the amphiphilic AMPs can disrupt bacterial IMs, they must be highly potent at attacking the OMs. This OM targeting feature may well help develop AMPs in controlling susceptibility as resistance is often associated with modified OM charges or enzymatic functions such as membrane-fused hydrolytic enzymes or efflux pumps.

In addition to the considerations of other uncharged components in IM and OM models in future work, it is also useful to understand the mode of action of AMPs against different LPS types. Such studies could lead to the development of strain-specific AMPs. Therefore, the development of realistic OM and cell wall models of different microbial strains will also help us understand the dynamic processes and structural disruptions to different microbial membranes and advance AMP research into further stages of their development as therapeutics.

## 4. Conclusion

The rational designed AMP $G_3$ has significantly enhanced antimicrobial activity and proteolytic stability against the natural AMPs melittin and LL37. In addition, $G_3$ has relatively weak cytotoxicity. Following the previously studies on the effects of the charge and saturation of lipids on their affinity to AMPs [29,39,40], this work has revealed that the LPS monolayer is even more susceptible to the $G_3$ destruction than the anionic phospholipid DPPG monolayer. Rapid and effective disruptions of $G_3$ against both LPS and DPPG monolayers with relatively lower surface adsorption than melittin and LL37 corroborate its strong and efficient killing to *E. coli*. In contrast, natural melittin is effective at binding to both LPS and DPPG monolayers due to its strong hydrophobicity. LL37 displayed the highest adsorption onto LPS and DPPG monolayers. However, most LL37 molecules were associated with the lipid head layer, with a low level of insertion into the acyl chain layer of the model membrane. The detailed NR studies have revealed the three AMPs are more potent against the OM

of *E. coli* than their IM. The detailed insights into the interactions between AMPs and bacterial membranes will help the further design and development of small peptide drugs that are more potent and selective against Gram-negative bacteria. Further studies are under way to investigate the design of Gram-positive bacteria specific AMPs by investigating AMPs interact with the thick peptidoglycan.

### CRediT authorship contribution statement

**Haoning Gong:** Conceptualization, Methodology, Data curation, Writing – original draft. **Xuzhi Hu:** Methodology, Software, Investigation, Writing – review & editing. **Lin Zhang:** Methodology, Validation. **Ke Fa:** Methodology, Validation. **Mingrui Liao:** Methodology, Validation. **Huayang Liu:** Methodology, Validation. **Giovanna Fragneto:** Methodology, Validation. **Mario Campana:** Methodology, Validation. **Jian Ren Lu:** Supervision, Project administration, Funding acquisition, Writing – review & editing.

### Data availability

Data will be made available on request.

### Declaration of Competing Interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

### Acknowledgements

We thank the studentship support from the University of Manchester, the China Scholarship Council, Syngenta, Lonza, and STFC. We acknowledge funding support from the National Natural Science Foundation of China (Project No. 22102187). We also thank Dr. Luke Clifton for his assistance to NR experiments. The authors acknowledge the neutron beam time awarded by the ISIS and ILL Neutron Sources (DOI: 10.5286/ISIS.E.RB1820088, DOI: 10.5291/ILL-DATA.9-13-802). We thank a European grant from a Marie Curie Fellowship ITN (grant number 608184) under SNAL (small nano-objects for alteration of lipid bilayers), a BBSRC LINK grant with AstraZeneca (BB/S018492/1) and an Innovate UK Knowledge Transfer Partnership project with Lonza under KTP 10809.

## Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.jcis.2023.01.051.

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