Antimicrobial peptides (AMPs) are generally membrane-active compounds that physically disrupt bacterial membranes. Despite extensive research, the precise mode of action of AMPs is still a topic of great debate. This work demonstrates that the initial interaction between the Gram-negative E. coli and AMPs is driven by lipopolysaccharides (LPS) that act as kinetic barriers for the binding of AMPs to the bacterial membrane. A combination of SPR and NMR experiments provide evidence suggesting that cationic AMPs first bind to the negatively charged LPS before reaching a binding place in the lipid bilayer. In the event that the initial LPS-binding is too strong (corresponding to a low dissociation rate), the cationic AMPs cannot effectively get from the LPS to the membrane, and their antimicrobial potency will thus be diminished. On the other hand, the AMPs must also be able to effectively interact with the membrane to exert its activity. The ability of the studied cyclic hexapeptides to bind LPS and to translocate into a lipid membrane is related to the nature of the cationic charge (arginine vs. lysine) and to the distribution of hydrophobicity along the molecule (alternating vs. clumped tryptophan).

One strategy to combat antimicrobial resistance is the discovery of new classes of antibiotics. Most antibiotics will at some point interact with the bacterial membrane to either interfere with its integrity or to cross it. Reliable and efficient tools for determining the dissociation constant for membrane binding (KD) and the partitioning coefficient between the aqueous- and membrane phases (KP) are therefore important tools for discovering and optimizing antimicrobial hits. Here we demonstrate that microscale thermophoresis (MST) can be used for label-free measurement of KD by utilising the intrinsic fluorescence of tryptophan and thereby removing the need for chromophore labelling. As proof of principle, we have used the method to measure the binding of a set of small cyclic AMPs to large unilamellar vesicles (LUVs) and two types of lipid nanodiscs assembled by styrene maleic acid (SMA) and quaternary ammonium SMA (SMA-QA). The measured KD values correlate well with the corresponding measurements using surface plasmon resonance (SPR), also broadly reflecting the tested AMPs’ minimal inhibition concentration (MIC) towards S. aureus and E. coli. We conclude that MST is a promising method for fast and cost-efficient detection of peptide-lipid interactions or mapping of sample conditions in preparation for more advanced studies that rely on expensive sample preparation, labelling and/or instrument time.

Antimicrobial peptides (AMPs) are a promising source of inspiration for new antibiotics discovery, in part because they believed to not trigger rapid resistance development since AMPs have co-existed with bacteria throughout evolution and do not target a single enzyme encoded by a single gene. AMPs is a diverse class of molecules that have diverse and often unspecific modes of action interfering with the membrane potential or the bacterial cell wall integrity, but also intracellular targets have been reported. Regardless of the mode of action(s), the AMPs will first have to interact with-, or penetrate through-, the bacterial cell well, and early characterization of AMP activity relies on the determination of the KD (dissociation constant for binding affinity) and the KP (partitioning constant) of AMP:lipid interactions. Here we demonstrate that microscale thermophoresis (MST) can be used for reliable unlabelled measurement of KD and KP utilising the intrinsic tryptophan fluorescence, thus removing the need for chromophore labelling. The MST results of binding to small unilamellar vesicles (SUVs) and stryrene maleic acid (SMA) based nanodiscs are compared to the corresponding surface plasmon resonance (SPR) measurements. SMA-QA nanodiscs are shown to be best suited for accurate measurements, while vesicles are a viable alternative. Unmodified SMA-nanodiscs proved unsuitable due to interactions between the cationic AMPs and the anionic polymer belt. Significant reduction of KD was observed when 5% anionic lipids were included in the lipid composition of the membrane models. This highlights the preference of the tested AMPs for anionic bacterial membranes, and the measured KD and KP values correlate well with their activity towards S. aureus and E. coli. We conclude that MST is a promising method for fast and efficient detection of peptide-lipid interactions, and the relative strength of the interactions can be reliably ranked within a library of screened compounds.

One strategy to combat antimicrobial resistance is the discovery of new classes of antibiotics. Most antibiotics will at some point interact with the bacterial membrane to either interfere with its integrity or to cross it. Reliable and efficient tools for determining the dissociation constant for membrane binding (KD) and the partitioning coefficient between the aqueous- and membrane phases (KP) are therefore important tools for discovering and optimizing antimicrobial hits. Here we demonstrate that microscale thermophoresis (MST) can be used for label-free measurement of KD by utilising the intrinsic fluorescence of tryptophan and thereby removing the need for chromophore labelling. As proof of principle, we have used the method to measure the binding of a set of small cyclic AMPs to large unilamellar vesicles (LUVs) and two types of lipid nanodiscs assembled by styrene maleic acid (SMA) and quaternary ammonium SMA (SMA-QA). The measured KD values correlate well with the corresponding measurements using surface plasmon resonance (SPR), also broadly reflecting the tested AMPs’ minimal inhibition concentration (MIC) towards S. aureus and E. coli. We conclude that MST is a promising method for fast and cost-efficient detection of peptide-lipid interactions.