Quantitative live-cell imaging of lipidated peptide transport through an epithelial cell layer.

Oral drug delivery increases patient compliance and is thus the preferred administration route for most drugs. However, for biologics the intestinal barrier greatly limits the absorption and reduces their bioavailability. One strategy employed to improve on this is chemical modification of the biologic through the addition of lipid side chains. While it has been established that lipidation of peptides can increase transport, a mechanistic understanding of this effect remains largely unexplored. To pursue this mechanistic understanding, end-point detection of biopharmaceuticals transported through a monolayer of fully polarized epithelial cells is typically used. However, these methods are time-consuming and tedious. Furthermore, most established methods cannot be combined easily with high-resolution live-cell fluorescence imaging that could provide a mechanistic insight into cellular uptake and transport. Here we address this challenge by developing an axial PSF deconvolution scheme to quantify the transport of peptides through a monolayer of Caco-2 cells using single-cell analysis with live-cell confocal fluorescence microscopy. We then measure the known cross-barrier transport of several compounds in our model and compare the results with results obtained in an established microfluidic model finding similar transport phenotypes. This verifies that already after two days the Caco-2 cells in our model form a tight monolayer and constitute a functional barrier model. We then apply this assay to investigate the effects of side chain lipidation of the model peptide drug salmon calcitonin (sCT) modified with 4­carbon and 8­carbon-long fatty acid chains. Furthermore, we compare that with experiments performed at lower temperature and using inhibitors for some endocytotic pathways to pinpoint how lipidation length modifies the main avenues for the transport. We thus show that increasing the length of the lipid chain increases the transport of the drug significantly but also makes endocytosis the primary transport mechanism in a short-term cell culture model.

Imaging therapeutic peptide transport across intestinal barriers.

Oral delivery is a highly preferred method for drug administration due to high patient compliance. However, oral administration is intrinsically challenging for pharmacologically interesting drug classes, in particular pharmaceutical peptides, due to the biological barriers associated with the gastrointestinal tract. In this review, we start by summarizing the pharmacological performance of several clinically relevant orally administrated therapeutic peptides, highlighting their low bioavailabilities. Thus, there is a strong need to increase the transport of peptide drugs across the intestinal barrier to realize future treatment needs and further development in the field. Currently, progress is hampered by a lack of understanding of transport mechanisms that govern intestinal absorption and transport of peptide drugs, including the effects of the permeability enhancers commonly used to mediate uptake. We describe how, for the past decades, mechanistic insights have predominantly been gained using functional assays with end-point read-out capabilities, which only allow indirect study of peptide transport mechanisms. We then focus on fluorescence imaging that, on the other hand, provides opportunities to directly visualize and thus follow peptide transport at high spatiotemporal resolution. Consequently, it may provide new and detailed mechanistic understanding of the interplay between the physicochemical properties of peptides and cellular processes; an interplay that determines the efficiency of transport. We review current methodology and state of the art in the field of fluorescence imaging to study intestinal barrier transport of peptides, and provide a comprehensive overview of the imaging-compatible in vitro, ex vivo, and in vivo platforms that currently are being developed to accelerate this emerging field of research.

Compositional inhomogeneity of drug delivery liposomes quantified at the single liposome level.

Liposomes are the most used drug delivery vehicle and their therapeutic function is closely linked to their lipid composition. Since most liposome characterization is done using bulk techniques, providing only ensemble averages, the lipid composition of all liposomes within the same formulation are typically assumed to be identical. Here we image individual liposomes using confocal microscopy to quantify that liposomal drug delivery formulations, including multiple component mixtures mimicking Doxil, display more than 10-fold variation in their relative lipid composition. Since liposome function is tightly regulated by the physicochemical properties bestowed by the lipid composition, such significant variations could render only a fraction of liposomes therapeutically active. Additionally, we quantified how this degree of compositional inhomogeneity was modulated by liposome preparation method, the saturation state of the membrane lipid, and whether anti-fouling polyethylene glycol (PEG) conjugated lipids were added to the initial lipid mix or inserted after liposome formation. We believe the insights into the factors governing the degree of inhomogeneity offers the possibility for producing more uniform liposomal drug delivery systems, potentially increasing their therapeutic efficacy.

A Quantitative Fluorescence Microscopy-based Single Liposome Assay for Detecting the Compositional Inhomogeneity Between Individual Liposomes.

Most research employing liposomes as membrane model systems or drug delivery carriers relies on bulk read-out techniques and thus intrinsically assumes all liposomes of the ensemble to be identical. However, new experimental platforms able to observe liposomes at the single-particle level have made it possible to perform highly sophisticated and quantitative studies on protein-membrane interactions or drug carrier properties on individual liposomes, thus avoiding errors from ensemble averaging. Here we present a protocol for preparing, detecting, and analyzing single liposomes using a fluorescence-based microscopy assay, facilitating such single-particle measurements. The setup allows for imaging individual liposomes in a massive parallel manner and is employed to reveal intra-sample size and compositional inhomogeneities. Additionally, the protocol describes the advantages of studying liposomes at the single liposome level, the limitations of the assay, and the important features to be considered when modifying it to study other research questions.

Dissociation of fluorescently labeled lipids from liposomes in biological environments challenges the interpretation of uptake studies.

Within nanomedicine, liposomes are investigated for their ability to deliver drug cargoes specifically into subcellular compartments of target cells. Such studies are often based on flow cytometry or microscopy, where researchers rely on fluorescently labeled lipids (FLLs) incorporated into the liposomal membrane to determine the localization of the liposomes within cells. These studies assume that the FLLs stay embedded in the liposomal membrane throughout the duration of the experiment. Here, we used size exclusion chromatography (SEC) to investigate the validity of this assumption by quantitatively determining the propensity of various widely used FLLs to dissociate from liposomes during incubation in human plasma. For certain commonly used off-the-shelf FLLs, up to 75% of the dye dissociated from the liposomes, while others dissociated less than 10%. To investigate the implications of this finding, we measured the peripheral blood leukocyte uptake of liposomes formulated with different FLLs using flow cytometry, and observed a significant difference in uptake correlating with the FLL's dissociation tendencies. Consequently, the choice of FLL can dramatically influence the conclusions drawn from liposome uptake and localization studies due to uptake of dissociated FLLs. The varying dissociation propensities for the FLLs were not reflected when incubating in buffer, showing that non-biological environments are unsuitable to mimic liposomal stability in a drug delivery context. Overall, our findings suggest that it is crucial for researchers to evaluate the stability of their FLL-labeled liposomes in biological environments, and the simplicity of the SEC assay put forward here makes it very applicable for the purpose.

Membrane curvature enables N-Ras lipid anchor sorting to liquid-ordered membrane phases.

Trafficking and sorting of membrane-anchored Ras GTPases are regulated by partitioning between distinct membrane domains. Here, in vitro experiments and microscopic molecular theory reveal membrane curvature as a new modulator of N-Ras lipid anchor and palmitoyl chain partitioning. Membrane curvature was essential for enrichment in raft-like liquid-ordered phases; enrichment was driven by relief of lateral pressure upon anchor insertion and most likely affects the localization of lipidated proteins in general.