This work was supported by Florida Atlantic University Office of Undergraduate Research and Inquiry grant (M.J.S.), Florida Department of Health Ed and Ethel Moore Pilot Grant 20A17 (Q.Z.), Alzheimer&#39;s Association grant AARG-NTF-19-618710 (Q.Z.), and NIA R21 grant AG061656-01A1 (Q.Z.).

The following protocol includes (1) a simplified procedure for establishing mouse hippocampal and cortical cultures based on a well-established protocol22, (2) a brief introduction to an epifluorescence microscope setup for live neurons, (3) a detailed description of loading and imaging ND6 in mouse neurons, (4) a discussion about the quantification of membrane trafficking by ND6 signal. All procedures follow the biosafety and IACUC guidelines at the Florida Atlantic University. The synthesis of ND6 has been described previously20.
1. Preparation of mouse hippocampal and cortical cultures
NOTE: If not specified otherwise, all steps must be performed in a biosafety level 2 laminar flow hood. Sterilize all tools and materials.
<ol>
	<li>Use size 5 forceps to place glass coverslips in multiwell culture plates. 
	NOTE: For example, a 24-well plate is ideal for 12 mm <img alt="Equation 1" src="/files/ftp_upload/62717/62717eq01v2.jpg" /> coverslips.</li>
	<li>Add the appropriate volume of extracellular matrices to coat the cell culture surface (e.g., 75 &#956;L of basement membrane matrix solution per 12 mm <img alt="Equation 1" src="/files/ftp_upload/62717/62717eq01v2.jpg" /> coverslip; see the Table of Materials)</li>
	<li>Place the prepared plates in the tissue culture incubator (5% CO2, 100% humidity, and 37 &#176;C) for 1-4 h to allow basement membrane matrix crosslinking.</li>
	<li>Sacrifice the animals, open the skull, and transfer the whole brain to a 35 mm Petri dish with 3 mL of Hank&#39;s Balanced Salt Solution (HBSS) containing 20% fetal bovine serum (H+20). Cut the brain along the midline, and separate the cortices and hippocampi in a laminar flow dissection hood to reduce contamination.</li>
	<li>Use microscissors to cut the tissues into small pieces (~1 mm3). Transfer those tissues to a 15 mL conical tube containing 5 mL of H+20.</li>
	<li>Wash the tissues three times with 5 mL of H+20 and three times with 5 mL of HBSS.</li>
	<li>Add 1.5 mL of 1% trypsin with ethylenediamine tetraacetic acid and incubate at 37 &#176;C for 10 min for enzyme digestion.</li>
	<li>Repeat step 1.6 and use vacuum to aspirate HBSS in the end.</li>
	<li>Dissociate the tissues mechanically in 1 mL of HBSS containing 2.95 g/L MgSO4&#8226;7H2O using a fire-polished glass pipette.</li>
	<li>Centrifuge the cell suspension at 400 &#215; g, 4 &#176;C for 5 min.</li>
	<li>Aspirate the supernatant and resuspend the cells in an appropriate volume of culture media&#160;to obtain a concentration of ~10,000,000 cells/ml.</li>
	<li>Plate the cells at ~1,000,000 cells/cm2 and place the culture in the incubator (5% CO2, 100% humidity, and 37 &#176;C) for 1-4 h to facilitate cell attachment to the glass coverslips.</li>
	<li>Add an appropriate volume of culture medium (e.g., 1 mL per well for a 24-well plate). Add the same volume of culture medium after 1 week. After 2 weeks, replace half of the existing culture medium with fresh media&#160;every week.</li>
</ol>
2. Microscope setup for live-cell fluorescence imaging
NOTE: An exemplary imaging setup (Figure 5) includes at least an inverted fluorescence microscope (see the Table of Materials), fluorescence light source with automatic shutter, fluorescence filter sets (e.g., for imaging ND6, use 405/10 BP for excitation, 495 LP for dichroic, and 510/20 BP for emission), and a high-sensitivity camera (Table of Materials), all of which are controlled by image acquisition software (Table of Materials).
<ol>
	<li>Prepare an imaging chamber that allows temperature control and solution input/output for live-cell fluorescence imaging. 
	NOTE: For example, a modified open-bath imaging chamber fixed on a heating platform was used in this protocol (Figure 6).</li>
	<li>Set up a programable device to switch the perfusion solutions and deliver the electric stimulus at defined time points during imaging. 
	NOTE: Hardware and software synchronization is necessary for quantitative analyses (Figure 7). For example, in this protocol, a trigger output from the imaging camera was used to start a computer program that controls an automatic switch device, which controls a multichannel perfusion system and a square-pulse stimulator.</li>
	<li>To co-image two or more fluorescent reporters, perform multichannel fluorescence imaging either by sequentially switching filter sets or by simultaneously splitting different fluorescence emissions and projecting to the same camera.</li>
</ol>
3. Loading and imaging ND6 in neuronal cultures
<ol>
	<li>Weigh out an appropriate amount of ND6, dissolve it in dimethylsulfoxide (DMSO), and allow it to solubilize at room temperature; sonicate briefly (e.g., 3 min). Filter the crude stock solution using a 0.22 &#956;m filter to remove large dye aggregates. After filtration, determine the dye concentration by absorption spectroscopy at 405 nm using a conventional or microvolume spectrophotometer using &#949; = 10,700 M-1 cm-1.</li>
	<li>Before application, dilute the stock solution to a concentration of 1 &#956;M&#160;using the bath solution. Keep the stock solution at room temperature in the dark.</li>
	<li>Labeling of endosomes and synaptic vesicles
	<ol>
		<li>To ubiquitously label endocytosed membrane compartments such as endosomes, add ND6 stock solution (e.g., 1 mM in DMSO) to the culture medium at the final concentration of 1 &#181;M, and incubate at 5% CO2, 100% humidity, and 37 &#176;C for 30 min. To reduce the extent of SV labeling, suppress the spontaneous neuronal activity pharmacologically. For example, use tetrodotoxin to block action potentials or 2,3-dioxo-6-nitro-1,2,3,4-tetrahydrobenzo[f]quinoxaline-7-sulfonamide (NBQX) and D-(-)-2-amino-5-phosphonopentanoic acid (D-AP5) to inhibit excitatory transmission23.</li>
		<li>To selectively label SVs, use brief but strong stimulation to evoke presynaptic exo-/endocytosis. First, transfer the culture coverslip(s) to a 35 mm <img alt="Equation 1" src="/files/ftp_upload/62717/62717eq01v2.jpg" /> Petri dish containing 2 mL of normal Tyrode&#39;s solution at room temperature. Second, dilute the ND6 stock solution in high-K+ Tyrode&#39;s solution (90 mM KCl) at the final concentration of 1 &#956;M. Third, replace the normal Tyrode&#39;s solution in the Petri dish with ND6-containing high-K+ Tyrode&#39;s solution and incubate at room temperature for 2 min.</li>
	</ol>
	</li>
	<li>Transfer the ND6-labeled cell culture to the imaging chamber filled with prewarmed normal Tyrode&#39;s solution containing 10 &#956;M NBQX and 10 &#956;M D-AP5.</li>
	<li>Adjust the perfusion speed to ~0.2 mL/s (i.e., 1 drop per second) and start the perfusion of prewarmed normal Tyrode&#39;s solution containing NBQX and D-AP5 to remove excessive ND6 in the culture.</li>
	<li>Adjust the focus and locate the appropriate field of view containing healthy and well-spread neurons bearing connected neurites. Avoid areas containing unresolved dye colloids.</li>
	<li>Try imaging ND6-loaded cells with different exposure times to identify the best imaging settings. 
	NOTE: For the best exposure time, the highest pixel fluorescence intensity in the resulting image is about half of the bit range (e.g., for a 16-bit image, the bit range is from 0 to 65,535), which will allow a further increase in fluorescence when the acidic bath solution is applied. The selected exposure time should be used for all ND6 imaging.</li>
	<li>Set up the stimulation and perfusion protocol, frame interval, and total duration. For example, in the following protocol, use a 30 s baseline, a 10 s 30 Hz electric field stimulation, 50 s recovery, 120 s 90 mM K+, 60 s recovery, 60 s Tyrode&#39;s solution with 50-mM NH4Cl, 60 s Tyrode&#39;s solution at pH 5.5, and a frame interval of 3 s.</li>
	<li>Start the image acquisition accompanied by the synchronized stimulation and perfusion. Monitor the simulation and solution exchange during imaging.</li>
	<li>Stop the perfusion after the imaging ends, remove the coverslip, and clean the imaging chamber for the next trial.</li>
</ol>
4. Quantification of membrane trafficking by a change in ND6 fluorescence
<ol>
	<li>Back up and/or make an electronic copy of all image files.</li>
	<li>Choose an analysis program for data extraction, e.g., an open-source image analysis software such as ImageJ24 and/or FIJI25.</li>
	<li>Open or import an image stack to the analysis program.</li>
	<li>Set the first image as the reference and align the rest to it using functions/plugins such as Rigid Registration26, which will mitigate artifacts due to xy-drifting. Refer to Figure 8 for example images.</li>
	<li>Average all images acquired during the 30-s pre-baseline to produce a reference image. Save a copy of this image for future reference.</li>
	<li>Set the intensity threshold to generate a binary image from the baseline-averaged image.</li>
	<li>Use Watershed in ImageJ or similar functions in another program to separate connecting neurites or cells.</li>
	<li>Use the Analyze Particle function with appropriate area size and circularity to solicit regions of interest (ROIs) corresponding to cell membranes, endosomes, lysosomes, or synaptic boutons. Save all selected ROIs.</li>
	<li>Select four background ROIs in cell-free regions within the field of view.</li>
	<li>Measure the average pixel intensities for each ROI in every frame of the image stack, and export the results for statistical analysis.</li>
	<li>Calculate the mean intensity of the background ROIs as the baseline noise, which will be subtracted from the average pixel intensities for each ROI.</li>
	<li>Average the three highest average intensities for every selected ROI during the application of pH 5.5 Tyrode&#39;s solution to obtain the maximal fluorescence intensity, which is defined as 100% for normalization.</li>
	<li>Average the three lowest average intensities for every selected ROI during the application of 50 mM NH4Cl to set the minimal fluorescence intensity, which is defined as 0% for normalization.</li>
	<li>Calculate the relative fluorescence changes for every ROI based on its own 0% and 100% intensities. Derive mean fluorescence changes, change kinetics, and other values/plots from individual ROI data.</li>
</ol>

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The authors declare no competing financial interests.

Lipid-based dyes, such as 1,1&#8242;-dioctadecyl-3,3,3&#8242;,3&#8242;-tetramethylindocarbocyanine (DiI) and 3,3&#8242;-Dioctadecyloxacarbocyanine perchlorate (DiO), have long been used to illustrate cell morphology and track cellular processes such as the axon projections of neurons. Styryl dyes, such as FM1-43, have been invented and used successfully for the study of exocytosis34. Due to their low membrane affinity, they selectively label endocytosed vesicles where they are trapped while dyes r...

The lipid bilayer is one of the most common biomolecule assemblies and is indispensable for all cells. Outside cells, it forms the plasma membrane interfacing cells and their environment; inside cells, it compartmentalizes various organelles specialized for designated functionalities. Rather dynamic than still, lipid membranes constantly experience fusion and fission, which mediates biomaterial transport, organelle reform, morphology change, and cellular communication. Undoubtedly, the lipid membrane is the physical foundation for almost all cellular processes, and its dysfunction plays a crucial role in various disorders ranging from cancer1 to Alzheimer&#39;s disease2. Although lipid molecules are far less diverse than proteins, membrane research so far has mainly been protein-centric. For example, a lot more is known about protein machinery than about lipids in exocytosis3,4,5. Moreover, the organization, distribution, dynamics, and homeostasis of lipids across surface and intracellular membranes largely remain unexplored in comparison to membrane proteins6.
This is not surprising as modern molecular biology techniques, such as mutagenesis, provide a methodological advantage for studying proteins rather than lipids. For example, transgenic tagging of pH-sensitive green fluorescent protein (a.k.a., pHluorin) to vesicular proteins facilitates the quantitative measurement of the amount and rate of vesicular protein turnover during exo-/endocytosis7,8,9. However, it is almost impossible to genetically modify membrane lipids in vivo. Moreover, qualitative and even quantitative manipulations of protein amounts and distributions are much more feasible than those of lipids10. Nevertheless, native and synthetic fluorescent lipids have been isolated and developed to simulate endogenous membrane lipids in vitro and in vivo11. One group of widely used fluorescent lipids are styryl dyes, e.g., FM1-43, which exhibit membrane-enhanced fluorescence and are a powerful tool in studying synaptic vesicle (SV) release in neurons12. Lately, environment-sensitive lipid dyes have been invented and widely used as a new class of reporters to study various cell membrane properties, including membrane potential11, phase order13, and secretion14.
A new class of lipid mimetics whose fluorescence is both pH-sensitive (e.g., pHluorin) and membrane-sensitive (e.g., FM1-43) was developed to directly measure the lipid distribution in the plasma membrane and endosomes/lysosomes and the lipid traffic during exo-/endocytosis. The well-known solvatochromic fluorophores exhibiting push-pull characteristics due to intramolecular charge transfer were selected for this purpose. Among existing solvatochromic fluorophores, the 1,8-naphthalimide (ND) scaffold is relatively easy to modify, versatile for tagging, and is unique in photo-physics15 and has therefore been used in DNA intercalators, organic light-emitting diodes, and biomolecule sensors16,17,18.
Attaching an electron-donating group to the C4 position of the ND scaffold generates a push-pull structure, which leads to an increased dipole moment by redistributing the electron density in the excited state19,20. Such an intramolecular charge transfer produces large quantum yields and Stokes shifts, resulting in bright and stable fluorescence21. This group has recently developed a series of solvatochromic lipid analogs based on the ND scaffold and obtained them with good synthetic yields20.
Spectroscopic characterization shows that among those products, ND6 possesses the best fluorescence properties (Figure 1)20. First, it has well-separated excitation and emission peaks (i.e., ~400 nm and ~520 nm, respectively, in Figure 2A,B) compared to popular fluorophores such as fluorescein isothiocyanate, rhodamine, or GFP, making it spectrally separatable from them and thus useful for multicolor imaging. Second, ND6 fluorescence exhibits a more than eight-fold increase in its fluorescence in the presence of micelles (Figure 2C), suggesting a strong membrane-dependency. Prior live-cell fluorescence imaging studies with different types of cells showed excellent membrane staining by ND620. Third, when the solution&#39;s pH is decreased from 7.5 (commonly found in extracellular or cytosolic environments) to 5.5 (commonly found in endosomes and lysosomes), ND6 shows an approximately two-fold increase in fluorescence (Figure 2D), showing its pH-sensitivity. Moreover, molecular dynamics simulation indicates that ND6 readily integrates into the lipid bilayer with its ND scaffold facing out of the membrane and piperazine residue showing strong interactions with phospholipid head groups (Figure 3). Altogether, these features make ND6 an ideal fluorescent lipid analog to visualize and measure membrane lipid turnover during exo-/endocytosis.
This paper presents a method to study the turnover rate and dynamics of SV lipids using cultured mouse hippocampal neurons. By stimulating neurons with high K+ Tyrode&#39;s solutions, SVs and the plasma membrane were loaded with ND6 (Figure 4A,B). Subsequently, neurons were re-stimulated with different stimuli followed by NH4Cl-containing and pH 5.5 Tyrode&#39;s solutions (Figure 4D). This protocol facilitates the quantitative measurement of the assembled exocytosis and endocytosis rates under different circumstances (Figure 4C).

<table><tbody><tr><td>Digidata 1440A Data Acquistion System</td><td>Molecular Devices</td><td>Digidata 1440A</td><td>For synchronized stimulation and solution exchange</td></tr><tr><td>Dual Channel Temperature Controller</td><td>Warner Instruments</td><td>TC-344B</td><td>For live-cell imaging</td></tr><tr><td>Fetal Bovine Serum</td><td>OMEGA Scientific</td><td>FB-01</td><td>For making H+20 solution used in dissection and tissue culture</td></tr><tr><td>Hamamatsu Flash4.0 sCOMS camera</td><td>Hamamatsu Inc.</td><td>C13440-20CU</td><td>high-sensitivity camera</td></tr><tr><td>Hank&#039;s Balanced Salt Solution</td><td>Sigma</td><td>H6648</td><td>For making H+20 solution used in dissection and tissue culture</td></tr><tr><td>Heated Platform</td><td>Warner Instruments</td><td>PH-1</td><td>For live-cell imaging</td></tr><tr><td>Matrigel</td><td>BD Biosciences</td><td>354234</td><td>For tissue culture</td></tr><tr><td>Micro-G Vibration Isolation Table</td><td>TMC</td><td>63-564</td><td>For live-cell imaging</td></tr><tr><td>Micro-manager</td><td>https://micro-manager.org/</td><td>NA</td><td>For image acquisition control</td></tr><tr><td>Multi-Line In-Line Solution Heater</td><td>Warner Instruments</td><td>SHM-6</td><td>For live-cell imaging</td></tr><tr><td>Neurobasal Plus Medium</td><td>THermoFisher Scientific</td><td>A3582901</td><td>For tissue culture</td></tr><tr><td>Nikon Ti-E Inverted Microscope</td><td>Nikon</td><td>Ti-E/B</td><td>For live-cell imaging</td></tr><tr><td>ORCA-Flash4.0 Digital CMOS camera</td><td>Hamamatsu</td><td>C1340-20CU</td><td>For live-cell imaging</td></tr><tr><td>Perfusion Chamber</td><td>Warner Instruments</td><td>RC-26G</td><td>For live-cell imaging</td></tr><tr><td>Six-Channel Valve Control Perfusion System</td><td>Warner Instruments</td><td>VC-6</td><td>For solution exchange</td></tr><tr><td>Square Pulse Stimulator</td><td>Grass Instrument</td><td>SD9</td><td>For electric field stimulation</td></tr></tbody></table>

measuring membrane lipid turnover with the ph-sensitive fluorescent lipid analog nd6

Exo-/endocytosis is a common process mediating the exchange of biomolecules between cells and their environment and among different cells. Specialized cells use this process to execute vital body functions such as insulin secretion from &#946; cells and neurotransmitter release from chemical synapses. Owing to its physiological significance, exo-/endocytosis has been one of the most studied topics in cell biology. Many tools have been developed to study this process at the gene and protein level, because of which much is known about the protein machinery participating in this process. However, very few methods have been developed to measure membrane lipid turnover, which is the physical basis of exo-/endocytosis.
This paper introduces a class of new fluorescent lipid analogs exhibiting pH-dependent fluorescence and demonstrates their use to trace lipid recycling between the plasma membrane and the secretory vesicles. Aided by simple pH manipulations, those analogs also allow the quantification of lipid distribution across the surface and the intracellular membrane compartments, as well as the measurement of lipid turnover rate during exo-/endocytosis. These novel lipid reporters will be of great interest to various biological research fields such as cell biology and neuroscience.

SVs are specialized for neurotransmitter release via evoked exo-/endocytosis27. SVs have highly acidic lumen (i.e., pH 5.5), which is ideal for ND6. We used high K+ stimulation to evoke SV exo-/endocytosis in order to allow ND6 to access SV. Expectedly, bright green fluorescent puncta along neuronal processes showed up after loading (Figure 9A). The line profile shown in Figure 4B demonstrated a strong overlap between ND6 (green curve) and ...

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Measuring Membrane Lipid Turnover with the pH-sensitive Fluorescent Lipid Analog ND6

This protocol presents a fluorescence imaging method that uses a class of pH-sensitive lipid fluorophores to monitor lipid membrane trafficking during cell exocytosis and the endocytosis cycle.

Exo-/endocytosis is a common process mediating the exchange of biomolecules between cells and their environment and among different cells. Specialized ...

measuring-membrane-lipid-turnover-with-ph-sensitive-fluorescent-lipid

Research

JoVE Journal

Neuroscience

1.5K Views.  The Brain Institute at Florida Atlantic University. Dit protocol beschrijft de karakterisering en toepassing van een nieuwe pH-gevoelige fluorescerende lipidensonde, specifiek voor celmembranen, waarmee onderzoekers exo- en endocytose in levende cellen kunnen volgen. De meeste eerdere verslaggevers over membraaneiwit op basis van pH-gevoelig groen fluorescentie-eiwit, dat indirect is. Als lipidenmimeticum integreert ND6 in het celmembraan en rapporteert zo getrouw het verkeer van lipidemembranen.ND6-sonde wordt alleen fluorescerend als ...

Video: Measuring Membrane Lipid Turnover with the pH-sensitive Fluorescent Lipid Analog ND6

A Fluorescence-based Assay of Phospholipid Scramblase Activity

Scramblases translocate phospholipids across the membrane bilayer bidirectionally in an ATP-independent manner. The first scramblase to be identified and biochemically verified was opsin, the apoprotein of the photoreceptor rhodopsin. Rhodopsin is a G protein-coupled receptor localized in rod photoreceptor disc membranes of the retina where it is responsible for the perception of light. Rhodopsin&#39;s scramblase activity does not depend on its ligand 11-cis-retinal, i.e., the apoprotein opsin is also active as a scramblase. Although constitutive and regulated phospholipid scrambling play an important role in cell physiology, only a few phospholipid scramblases have been identified so far besides opsin. Here we describe a fluorescence-based assay of opsin&#39;s scramblase activity. Opsin is reconstituted into large unilamellar liposomes composed of phosphatidylcholine, phosphatidylglycerol and a trace quantity of fluorescent NBD-labeled PC (1-palmitoyl-2-{6-[7-nitro-2-1,3-benzoxadiazole-4-yl)amino]hexanoyl}-sn-glycero-3-phosphocholine). Scramblase activity is determined by measuring the extent to which NBD-PC molecules located in the inner leaflet of the vesicle are able to access the outer leaflet where their fluorescence is chemically eliminated by a reducing agent that cannot cross the membrane. The methods we describe have general applicability and can be used to identify and characterize scramblase activities of other membrane proteins.

We describe a fluorescence-based assay to measure phospholipid scrambling in large unilamellar liposomes reconstituted with opsin.

Scramblases translocate phospholipids across the membrane bilayer bidirectionally in an ATP-independent manner. The first scramblase to be identified and ...

Biochemistry

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous number of imaging techniques currently available and the progress in instrumentation, there is still a need for highly sensitive probes that are suitable for in vivo imaging. One typical problem of available preclinical fluorescent probes is their rapid clearance in vivo, which reduces their imaging sensitivity. To circumvent rapid clearance, increase number of dye molecules at the target site, and thereby reduce background autofluorescence, encapsulation of the near-infrared fluorescent dye, DY-676-COOH in liposomes and verification of its potential for in vivo imaging of inflammation was done. DY-676 is known for its ability to self-quench at high concentrations. We first determined the concentration suitable for self-quenching, and then encapsulated this quenching concentration into the aqueous interior of PEGylated liposomes. To substantiate the quenching and activation potential of the liposomes we use a harsh freezing method which leads to damage of liposomal membranes without affecting the encapsulated dye. The liposomes characterized by a high level of fluorescence quenching were termed Lip-Q. We show by experiments with different cell lines that uptake of Lip-Q is predominantly by phagocytosis which in turn enabled the characterization of its potential as a tool for in vivo imaging of inflammation in mice models. Furthermore, we use a zymosan-induced edema model in mice to substantiate the potential of Lip-Q in optical imaging of inflammation in vivo. Considering possible uptake due to inflammation-induced enhanced permeability and retention (EPR) effect, an always-on liposome formulation with low, non-quenched concentration of DY-676-COOH (termed Lip-dQ) and the free DY-676-COOH were compared with Lip-Q in animal trials.

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

The use of fluorophores for in vivo imaging can be greatly limited by opsonization, rapid clearance, low detection sensitivity and cytotoxic effects on the host. Encapsulation of fluorophores in liposomes by film hydration and extrusion leads to fluorescence quenching and protection which enables in vivo imaging with high detection sensitivity.

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous ...

Bioengineering

Measuring Nucleotide Binding to Intact, Functional Membrane Proteins in Real Time

We have developed a method to measure binding of adenine nucleotides to intact, functional transmembrane receptors in a cellular or membrane environment. This method combines expression of proteins tagged with the fluorescent non-canonical amino acid ANAP, and FRET between ANAP and fluorescent (trinitrophenyl) nucleotide derivatives. We present examples of nucleotide binding to ANAP-tagged KATP ion channels measured in unroofed plasma membranes and excised, inside-out membrane patches under voltage clamp. The latter allows for simultaneous measurements of ligand binding and channel current, a direct readout of protein function. Data treatment and analysis are discussed extensively, along with potential pitfalls and artefacts. This method provides rich mechanistic insights into the ligand-dependent gating of KATP channels and can readily be adapted to the study of other nucleotide-regulated proteins or any receptor for which a suitable fluorescent ligand can be identified.

This protocol presents a method for measuring adenine nucleotide binding to receptors in real time in a cellular environment. Binding is measured as F&#246;rster resonance energy transfer (FRET) between trinitrophenyl nucleotide derivatives and protein labeled with a non-canonical, fluorescent amino acid.

We have developed a method to measure binding of adenine nucleotides to intact, functional transmembrane receptors in a cellular or membrane environment. ...

Fluorescence Recovery after Merging a Droplet to Measure the Two-dimensional Diffusion of a Phospholipid Monolayer

We introduce a new method to measure the lateral diffusivity of a surfactant monolayer at the fluid-fluid interface, called fluorescence recovery after merging (FRAM). FRAM adopts the same principles as the fluorescence recovery after photobleaching (FRAP) technique, especially for measuring fluorescence recovery after bleaching a specific area, but FRAM uses a drop coalescence instead of photobleaching dye molecules to induce a chemical potential gradient of dye molecules. Our technique has several advantages over FRAP: it only requires a fluorescence microscope rather than a confocal microscope equipped with high power lasers; it is essentially free from the selection of fluorescence dyes; and it has far more freedom to define the measured diffusion area. Furthermore, FRAM potentially provides a route for studying the mixing or inter-diffusion of two different surfactants, when the monolayers at a surface of droplet and at a flat air/water interface are prepared with different species, independently.

We present a new technique to measure the lateral diffusion of a surface active species at the fluid-fluid interface by merging a droplet monolayer onto a flat monolayer.

We introduce a new method to measure the lateral diffusivity of a surfactant monolayer at the fluid-fluid interface, called fluorescence recovery after ...

Fluorescent Leakage Assay to Investigate Membrane Destabilization by Cell-Penetrating Peptide

Cell-penetrating peptides (CPPs) are defined as carriers that are able to cross the plasma membrane and to transfer a cargo into cells. One of the main common features required for this activity resulted from the interactions of CPPs with the plasma membrane (lipids) and more particularly with components of the extracellular matrix of the membrane itself (heparan sulphate). Indeed, independent of the direct translocation or the endocytosis-dependent internalization, lipid bilayers are involved in the internalization process both at the level of the plasma membrane and at the level of intracellular traffic (endosomal vesicles). In this article, we present a detailed protocol describing the different steps of a large unilamellar vesicles (LUVs) formulation, purification, characterization, and application in fluorescence leakage assay in order to detect possible CPP-membrane destabilization/interaction and to address their role in the internalization mechanism. LUVs with a lipid composition reflecting the plasma membrane content are generated in order to encapsulate both a fluorescent dye and a quencher. The addition of peptides in the extravesicular medium and the induction of peptide-membrane interactions on the LUVs might thus induce in a dose-dependent manner a significant increase in fluorescence revealing a leakage. Examples are provided here with the recently developed tryptophan (W)- and arginine (R)-rich Amphipathic Peptides (WRAPs), which showed a rapid and efficient siRNA delivery in various cell lines. Finally, the nature of these interactions and the affinity for lipids are discussed to understand and to improve the membrane translocation and/or the endosomal escape.

The fluorescence leakage assay is a simple method that enables the investigation of peptide/membrane interactions in order to understand their involvement in several biological processes and especially the ability of cell-penetrating peptides to disturb phospholipids bilayers during a direct cellular translocation process.

Cell-penetrating peptides (CPPs) are defined as carriers that are able to cross the plasma membrane and to transfer a cargo into cells. One of the main ...

Fluorescence-Based Measurements of Phosphatidylserine/Phosphatidylinositol 4-Phosphate Exchange Between Membranes

Several members of the evolutionarily conserved oxysterol-binding protein (OSBP)-related proteins(ORP)/OSBP homologs (Osh) family have recently been found to represent a novel lipid transfer protein (LTP) group in yeast and human cells. They transfer phosphatidylserine (PS) from the endoplasmic reticulum (ER) to the plasma membrane (PM) via PS/phosphatidylinositol 4-phosphate (PI(4)P) exchange cycles. This finding allows a better understanding of how PS, which is critical for signaling processes, is distributed throughout the cell and the investigation of the link between this process and phosphoinositide (PIP) metabolism. The development of new fluorescence-based protocols has been instrumental in the discovery and characterization of this new cellular mechanism in vitro at the molecular level. This paper describes the production and the use of two fluorescently labelled lipid sensors, NBD-C2Lact and NBD-PHFAPP, to measure the ability of a protein to extract PS or PI(4)P and to transfer these lipids between artificial membranes. First, the protocol describes how to produce, label, and obtain high-purity samples of these two constructs. Secondly, this paper explains how to use these sensors with a fluorescence microplate reader to determine whether a protein can extract PS or PI(4)P from liposomes, using Osh6p as a case study. Finally, this protocol shows how to accurately measure the kinetics of PS/PI(4)P exchange between liposomes of defined lipid composition and to determine lipid transfer rates by fluorescence resonance energy transfer (FRET) using a standard fluorometer.

Here, we describe protocols using fluorescent lipid sensors and liposomes to determine whether a protein extracts and transports phosphatidylserine or phosphatidylinositol 4-phosphate in vitro.

Several members of the evolutionarily conserved oxysterol-binding protein (OSBP)-related proteins(ORP)/OSBP homologs (Osh) family have recently been found ...

Measuring the Time-Evolution of Nanoscale Materials with Stopped-Flow and Small-Angle Neutron Scattering

This paper presents the use of a stopped-flow small-angle neutron-scattering (SANS) sample environment to quickly mix liquid samples and study nanoscale kinetic processes on time scales of seconds to minutes. The stopped-flow sample environment uses commercially available syringe pumps to mix the desired volumes of liquid samples that are then injected through a dynamic mixer into a quartz glass cell in approximately 1 s. Time-resolved SANS data acquisition is synced with the sample mixing to follow the evolution of the nanostructure in solution after mixing. 
To make the most efficient use of neutron beam time, we use a series of flow selector valves to automatically load, rinse, and dry the cell between measurements, allowing for repeated data collection throughout multiple sample injections. Sample injections are repeated until sufficient neutron scattering statistics are collected. The mixing setup can be programmed to systematically vary conditions to measure the kinetics at different mixing ratios, sample concentrations, additive concentrations, and temperatures. The minimum sample volume required per injection is approximately 150 &#181;L depending on the path length of the quartz cell. 
Representative results using this stopped-flow sample environment are presented for rapid lipid exchange kinetics in the presence of an additive, cyclodextrin. The vesicles exchange outer-leaflet (exterior) lipids on the order of seconds and fully exchange both interior and exterior lipids within hours. Measuring lipid exchange kinetics requires in situ mixing to capture the faster (seconds) and slower (minutes) processes and extract the kinetic rate constants. The same sample environment can also be used to probe molecular exchange in other types of liquid samples such as lipid nanoparticles, proteins, surfactants, polymers, emulsions, or inorganic nanoparticles. Measuring the nanoscale structural transformations and kinetics of exchanging or reacting systems will provide new insights into processes that evolve at the nanoscale.

This protocol presents the use of a stopped-flow sample environment to quickly mix multiple liquid solutions in situ during a small-angle neutron scattering measurement and to study kinetic processes on nanometer length scales and second time scales.

This paper presents the use of a stopped-flow small-angle neutron-scattering (SANS) sample environment to quickly mix liquid samples and study nanoscale ...

Engineering

Gramicidin-based Fluorescence Assay; for Determining Small Molecules Potential for Modifying Lipid Bilayer Properties

Many drugs and other small molecules used to modulate biological function are amphiphiles that adsorb at the bilayer/solution interface and thereby alter lipid bilayer properties. This is important because membrane proteins are energetically coupled to their host bilayer by hydrophobic interactions. Changes in bilayer properties thus alter membrane protein function, which provides an indirect way for amphiphiles to modulate protein function and a possible mechanism for "off-target" drug effects. We have previously developed an electrophysiological assay for detecting changes in lipid bilayer properties using linear gramicidin channels as probes 3,12. Gramicidin channels are mini-proteins formed by the transbilayer dimerization of two non-conducting subunits. They are sensitive to changes in their membrane environment, which makes them powerful probes for monitoring changes in lipid bilayer properties as sensed by bilayer spanning proteins. We now demonstrate a fluorescence assay for detecting changes in bilayer properties using the same channels as probes. The assay is based on measuring the time-course of fluorescence quenching from fluorophore-loaded large unilamellar vesicles due to the entry of a quencher through the gramicidin channels. We use the fluorescence indicator/quencher pair 8-aminonaphthalene-1,3,6-trisulfonate (ANTS)/Tl+ that has been successfully used in other fluorescence quenching assays 5,13. Tl+ permeates the lipid bilayer slowly 8 but passes readily through conducting gramicidin channels 1,14. The method is scalable and suitable for both mechanistic studies and high-throughput screening of small molecules for bilayer-perturbing, and potential "off-target", effects. We find that results using this method are in good agreement with previous electrophysiological results 12.

We introduce a fast fluorescence-based assay that monitors the rate of fluorescence quenching as a measure of gramicidin channel activity. The gramicidin channels are used as molecular force transducers to monitor changes in lipid bilayer properties as sensed by bilayer spanning proteins.

Many drugs and other small molecules used to modulate biological function are amphiphiles that adsorb at the bilayer/solution interface and thereby alter ...

Biology

Assessing Hepatic Steatosis with 3D Fluorescence Lipid Droplet Reconstruction

Analysis of Fluorescent-Stained Lipid Droplets with 3D Reconstruction for Hepatic Steatosis Assessment

Lipid droplets (LDs) are specialized organelles that mediate lipid storage and play a very important role in suppressing lipotoxicity and preventing dysfunction caused by free fatty acids (FAs). The liver, given its critical role in the body's fat metabolism, is persistently threatened by the intracellular accumulation of LDs in the form of both microvesicular and macrovesicular hepatic steatosis. The histologic characterization of LDs is typically based on lipid-soluble diazo dyes, such as Oil Red O (ORO) staining, but a number of disadvantages consistently hamper the use of this analysis with liver specimens. More recently, lipophilic fluorophores 493/503 have become popular for visualizing and locating LDs due to their rapid uptake and accumulation into the neutral lipid droplet core. Even though most applications are well-described in cell cultures, there is less evidence demonstrating the reliable use of lipophilic fluorophore probes as an LD imaging tool in tissue samples. Herein, we propose an optimized boron dipyrromethene (BODIPY) 493/503-based protocol for the evaluation of LDs in liver specimens from an animal model of high-fat diet (HFD)-induced hepatic steatosis. This protocol covers liver sample preparation, tissue sectioning, BODIPY 493/503 staining, image acquisition, and data analysis. We demonstrate an increased number, intensity, area ratio, and diameter of hepatic LDs upon HFD feeding. Using orthogonal projections and 3D reconstructions, it was possible to observe the full content of neutral lipids in the LD core, which appeared as nearly spherical droplets. Moreover, with the fluorophore BODIPY 493/503, we were able to distinguish microvesicles (1 &#181;m &lt; d &#8804; 3 &#181;m), intermediate vesicles (3 &#181;m &lt; d &#8804; 9 &#181;m), and macrovesicles (d &gt; 9 &#181;m), allowing the successful discrimination of microvesicular and macrovesicular steatosis. Overall, this BODIPY 493/503 fluorescence-based protocol is a reliable and simple tool for hepatic LD characterization and may represent a complementary approach to the classical histological protocols.

Author Spotlight: Analysis of Fluorescent-Stained Lipid Droplets with 3D Reconstruction for Hepatic Steatosis Assessment  

Herein, we demonstrate an optimized BODIPY 493/503 fluorescence-based protocol for lipid droplet characterization in liver tissue. Through the use of orthogonal projections and 3D reconstructions, the fluorophore allows for successful discrimination between microvesicular and macrovesicular steatosis and may represent a complementary approach to the classical histological protocols for hepatic steatosis assessment.

Lipid droplets (LDs) are specialized organelles that mediate lipid storage and play a very important role in suppressing lipotoxicity and preventing ...

Exploring the Regulation of Lipid Droplet Catabolism through Lipophagy

Macroautophagy, commonly referred to as autophagy, is a highly conserved cellular process responsible for the degradation of cellular components. This process is particularly prominent under conditions such as fasting, cellular stress, organelle damage, cellular damage, or aging of cellular components. During autophagy, a segment of the cytoplasm is enclosed within double-membrane vesicles known as autophagosomes, which then fuse with lysosomes. Following this fusion, the contents of autophagosomes undergo non-selective bulk degradation facilitated by lysosomes. However, autophagy also exhibits selective functionality, targeting specific organelles, including mitochondria, peroxisomes, lysosomes, nuclei, and lipid droplets (LDs). Lipid droplets are enclosed by a phospholipid monolayer that isolates neutral lipids from the cytoplasm, protecting cells from the harmful effects of excess sterols and free fatty acids (FFAs). Autophagy is implicated in various conditions, including neurodegenerative diseases, metabolic disorders, and cancer. Specifically, lipophagy -- the autophagy-dependent degradation of lipid droplets -- plays a crucial role in regulating intracellular FFA levels across different metabolic states. This regulation supports essential processes such as membrane synthesis, signaling molecule formation, and energy balance. Consequently, impaired lipophagy increases cellular vulnerability to death stimuli and contributes to the development of diseases such as cancer. Despite its significance, the precise mechanisms governing lipid droplet metabolism regulated by lipophagy in cancer cells remain poorly understood. This article aims to describe confocal imaging acquisition and quantitative imaging analysis protocols that enable the investigation of lipophagy associated with metabolic changes in cancer cells. The results obtained through these protocols may shed light on the intricate interplay between autophagy, lipid metabolism, and cancer progression. By elucidating these mechanisms, novel therapeutic targets may emerge for combating cancer and other metabolic-related diseases.

Lipophagy is a selective form of autophagy that involves the degradation of lipid droplets. Dysfunctions in this process are associated with cancer development. However, the precise mechanisms are not yet fully understood. This protocol describes quantitative imaging approaches to better understand the interplay between autophagy, lipid metabolism, and cancer progression.

Macroautophagy, commonly referred to as autophagy, is a highly conserved cellular process responsible for the degradation of cellular components. This ...

Measuring Membrane Lipid Turnover with the pH-sensitive Fluorescent Lipid Analog ND6

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