This work was funded by the Danish Council for Independent Research [grant number 5054-00165B].

1. Liposome Preparation
NOTE: Briefly, preparation of liposomes usually includes three crucial steps: 1) preparation of dry lipid films of the desired lipid composition; 2) rehydration of the lipids for formation of liposomes; and 3) controlling the size and lamellarity of the liposome population.
<ol>
	<li>Weigh out the lipids and dissolve them in tert-butanol:water (9:1) in glass vials.
	<ol>
		<li>Dissolve POPC (1-palmitoyl-2-oleoyl-glycero-3-phosphocholine; MW = 760 g/mol) to 50 mM.</li>
		<li>Dissolve cholesterol (MW = 387 g/mol) to 25 mM.</li>
		<li>Dissolve DOPE-Atto488 (1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-Atto488; MW = 1,316 g/mol) to 0.1 mM.</li>
		<li>Dissolve DOPE-Atto655 (1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-Atto655; MW = 1,368 g/mol) to 0.1 mM.</li>
		<li>Dissolve DSPE-PEG2000-biotin (1,2-distearyl-sn-glycero-3-phosphatethanolamine-N-[biotinyl(polyethylene glycol)-2000]; MW = 3,017 g/mol) to 0.1 mM. 
		NOTE: Heat lipids to 55 &#176;C and use magnetic stirring in order to ensure complete dissolution of the lipids. Alternatively, use a sonication bath. Unused lipid stocks can be stored at -20 &#176;C for several months.</li>
	</ol>
	</li>
	<li>Mix the lipid stocks prepared in step 1.1 to a molar ratio of POPC:cholesterol:DOPE-Atto488:DOPE-Atto655:DOPE-PEG-biotin 68.95:30:0.5:0.5:0.05, by adding 138 &#181;L of POPC, 120 &#181;L of cholesterol, 500 &#181;L of each fluorescently labeled lipid, and 50 &#181;L of DSPE-PEG-biotin to a fresh glass vial. 
	NOTE: The exact liposome composition can easily be modified in order to address the specific question of interest. See discussion for more detail.</li>
	<li>Loosen the lid of the glass vial, and snap-freeze the vial in liquid nitrogen.</li>
	<li>Lyophilize the frozen lipid mixture overnight.</li>
	<li>Add 1 mL of 200 mM D-sorbitol buffer (sorbitol buffer) to the dry lipids.</li>
	<li>Heat the mixture to 45 &#176;C and expose to magnetic stirring for at least 1 h. 
	NOTE: The buffer should reflect the specific question that is being addressed (e.g., physiological conditions for studying membrane-protein interactions, or a specific clinically approved buffer for studying drug delivery liposomes). However, if a specific buffer is not required for the study, a buffer without ions can be applied for rehydration in order to reduce the multilamellarity of the liposomes.</li>
	<li>Freeze the lipid suspension by dipping the vial in liquid nitrogen, and wait until the suspension is completely frozen.</li>
	<li>Dip the frozen suspension in a heating bath at 55 &#176;C until the mixture is completely thawed.</li>
	<li>Repeat steps 1.7 and 1.8 until the liposome suspension has been exposed to a total of 11 freeze/thaw cycles. 
	NOTE: Repeated freeze/thaw cycles have shown to reduce liposome multilamellarity23, which is paramount for the accuracy of the single liposome assay, as multilamellar liposomes will skew the fluorescence intensity versus liposome size ratio of the liposomes. The multilamellarity is usually inherently low when including more than 0.5% PEGylated lipid in the formulation (such as commonly done in liposomes for drug delivery)24.</li>
	<li>Extrude the liposome suspension once through an 800 nm polycarbonate filter using a mini extrusion kit. Follow the manufacturer&#39;s instructions for assembly of the extrusion kit (see Table of Materials).</li>
	<li>Store the liposomes at 4 &#176;C overnight.</li>
</ol>
2. Surface Preparation of Imaging Chamber
<ol>
	<li>Prepare bovine serum albumin (BSA; 1 mg/mL), BSA-biotin (1 mg/mL) and streptavidin (0.025 mg/mL) in 10 mM HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid) 95 mM NaCl buffer (HEPES buffer). 
	NOTE: To prevent liposomal burst, the sorbitol buffer used for rehydration and the HEPES buffer used for surface preparation should be isotonic. It is thus recommended to check the osmolarity of the buffers before the experiment.</li>
	<li>Mix 1,200 &#181;L of BSA and 120 &#181;L of BSA-biotin and add 300 &#181;L of the mixture to each well in an 8 well slide for microscopy. 
	NOTE: Here we use a commercially available 8 well microscope slide with a glass bottom (see Table of Materials), but the protocol can easily be adapted to customized microscope chambers.</li>
	<li>Incubate the slide for 20 min at room temperature (RT).</li>
	<li>Wash the slide 8x with 300 &#181;L of HEPES buffer. 
	NOTE: Be careful not to leave the wells without buffer for more than a few seconds, as drying out the surface will damage it. Furthermore, be careful not to scratch the surface with the pipette tip as this will also damage it. Thus, when aspirating buffer from a well, do it from the edge or corner.</li>
	<li>Add 250 &#181;L of streptavidin and incubate for 10 min at RT.</li>
	<li>Repeat the 8 washes described in step 2.4.</li>
	<li>Store the microscope slide with 300 &#181;L of sorbitol buffer in each well at 4 &#176;C. Evaporation of the solvent will damage the surface, so put the microscope slide in a Petri dish and seal it with parafilm unless the prepared slide is used immediately.</li>
</ol>
3. Liposome Immobilization
<ol>
	<li>Dilute the liposome suspension to about 20 &#181;M total lipid in sorbitol buffer. The procedure in section 1 will yield a liposome suspension of approximately 10 mM total lipid.</li>
	<li>Place the slide on the microscope and focus on the surface of the chamber using the increased laser reflection signal from the glass/buffer interface as a guide.</li>
	<li>Wash the chamber 4x with 300 &#181;L of fresh HEPES buffer.</li>
	<li>Add 10 &#181;L of diluted liposome stock (20 &#181;M total lipid) to a microcentrifuge tube.</li>
	<li>Take out 100 &#181;L of buffer from the chamber, add to the microcentrifuge tube prepared in step 3.4, mix properly, and put the 110 &#181;L back into the chamber.</li>
	<li>Put the specimen under the microscope, and use a rudimental microscope setting capable of detecting signal from the liposome fluorophore(s) to observe the liposomes being immobilized on the surface.</li>
	<li>Aim for a liposome surface density that is simultaneously sparse enough to indentify individual liposomes and dense enough that it facilitates high-throughput measurements. Typically using a 50 &#181;m x 50 &#181;m field of view, 300&#8722;400 liposomes per frame is optimal (Figure 2). This can usually be achieved within 5&#8722;10 min. 
	NOTE: If only rapidly moving fluorescent particles are detected in the field of view, an absence or too low concentration of any of the components critical for the immobilization process (BSA-biotin, streptavidin, DOPE-PEG-biotin) might be the cause. If no liposomes are detected it might either be related to a low concentration of liposomes, improper settings for the fluorescence detection, or potentially imaging with a focal plane not at the glass surface.</li>
	<li>Once an appropriate liposome surface density is reached, wash the chamber 3x with 200 &#181;L of HEPES buffer. 
	NOTE: Be aware that the immobilization kinetics also depend on liposome properties such as size and charge and should thus always be optimized for each liposome formulation.</li>
</ol>
4. Image Acquisition
NOTE: This section will depend a lot on the microscope system available to the researcher performing the experiment. Thus, overall guidelines on how to perform the imaging will be described. However, the exact settings and how to apply them will vary between the different microscope setups. For example, some systems allow choosing any emission filter combination desired, while other microscopes are equipped with specific, preset filters.
<ol>
	<li>Set up the microscope for imaging single liposomes. To ensure optimal image quality and subsequent data analysis use a high greyscale resolution and a pixel scheme that allows for oversampling individual liposomes. A bit depth of 16 and at least 1,024 x 1,024 pixels for a 50 &#181;m x 50 &#181;m area is recommended. If available, line-averaging can beneficially be applied to reduce noise (e.g., 3 scans per line).</li>
	<li>Select an excitation laser power that both ensures nonsignificant frame-to-frame bleaching and strong enough signal to clearly discriminate individual liposomes from the background. The optimal setting will depend on the specific microscope used as well as the fluorophore combination. Make sure that detectors are not saturated, as this will bias intensity quantification.</li>
	<li>Imaging both the DOPE-Atto488 and DOPE-Atto655 fluorophores in the liposomes requires imaging multiple channels. Thus, make sure each channel is imaged sequentially to avoid cross-excitation. For example, first take one image by exciting at 488 nm and reading emission at 495&#8722;560 nm. Thereafter, take another image by exciting at 633 nm but reading emission only at 660&#8722;710 nm. The specifics will depend on the microscope system (e.g., which lasers and filters) available.</li>
	<li>Make sure to cover different areas of the surface, acquiring at least 10 images of the sample, thus imaging at least 3,000 individual liposomes. If the surface density is lower than 300 liposomes/frame, acquire more images. Make sure to refocus the microscope for every new image. 
	NOTE: For two-channel imaging, make sure to name the image files so that pairs of images with the same liposomes in different fluorescence channels can easily be identified during data analysis.</li>
	<li>In order to quantify the experimental uncertainty relating to the measurement of compositional inhomogeneity, image the same area of liposomes before and after refocusing (see Figure 3 and description in the text).</li>
	<li>Export the images from the microscope software as .tiff files. Export the two channels of the same liposomes individually.</li>
</ol>
5. Data Analysis
NOTE: Specially developed automated 2D Gaussian fitting routines have previously been employed6,11,12. However, to increase the applicability of the method a data analysis process that can be easily implemented in all laboratories is described.
<ol>
	<li>Load the corresponding pair of .tiff images of two different fluorescence channels in the same imaging field into the FIJI (FIJI Is Just ImageJ) software.</li>
	<li>In the Image menu, choose Color, and use the Merge Channel function to create a composite of the two channels.</li>
	<li>Observe if the liposomes imaged in two different channels display good colocalization or whether visible drift occurred. 
	NOTE: In case of drift between the two fluorescence channels (recognized as a systematic and equal X-Y offset between the signal in the two channels), one of the frames can be translated using the Transform &#62; Translate function in the Image menu of FIJI. However, care should be taken with such image manipulation. It is thus recommended to instead avoid drift as much as possible when imaging the liposomes.</li>
	<li>Make sure the ComDet plugin (v.0.3.6.1 or newer) is installed, or do this in the Plugins menu.</li>
	<li>Open the ComDet plugin to detect particles by going to the Plugins menu and choosing ComDet v.0.3.6.1 &#62; Detect Particles.</li>
	<li>In ComDet, make sure to choose the Detect Particles on Both Channels Individually function. Set the Max Distance Between Co-localized Spots similar to the Approximate Particles Size. Usually, 4 pixels is appropriate for the settings described here.</li>
	<li>The signal-to-noise ratio should allow for detection of even dim particles with a low amount of fluorescence. In ComDet, set this ratio to 3 by setting the Sensitivity of Detection in both channels to Very Dim Particles (SNR = 3). 
	NOTE: While this SNR value is usually appropriate, it might be necessary to set it higher (e.g., if there is a lot of image noise that may be identified as liposomes and lead to false positives).</li>
	<li>Make sure the boxes with Calculate Colocalization and Plot Detected Particles in Both Channels are checked, before pressing OK.</li>
	<li>After running the analysis, two pop-up windows with Results and Summary will show. Export the data table Results containing the colocalization data (particle coordinates and integrated intensity of each particle detected) to a data handling software of choice by saving the table as a .txt file and importing it into the software.</li>
	<li>Make sure that each liposome is only included once in the data set and not both the channel1/channel2 as well as channel2/channel1 ratio. Thus, filter the data so only Abs_frame = 1 is plotted in step 5.11. 
	NOTE: By choosing Colocalized = 1, false positives from noise in the images can be removed from the analysis. However, any liposomes with only one of the two fluorescent components present will also be excluded from the analysis, thus potentially removing important data points from the analysis.</li>
	<li>Plot a histogram of the column with data containing the Intensity Ratio for each detected liposome.</li>
	<li>The degree of compositional inhomogeneity for the studied liposomal system is represented by the width of the intensity ratio distribution. To quantify the inhomogeneity, fit the intensity ratio histogram with a Gaussian function and extract the mean (&#181;) and standard deviation (sigma). See the Representative Results section.</li>
	<li>A value for the degree of inhomogeneity (DI) can now be calculated using the coefficient of variation defined as DI = sigma / &#181;.</li>
</ol>
6. Liposome Size Calibration
<ol>
	<li>Take out part of the liposome stock, and extrude 21x through a 50 nm polycarbonate filter as described in step 1.10.</li>
	<li>Dilute the liposomes by adding 10 &#181;L of the liposome suspension to 800 &#181;L of sorbitol buffer in a microcentrifuge tube.</li>
	<li>Transfer the diluted liposome sample to a polypropylene single-use cuvette.</li>
	<li>Measure the size using dynamic light scattering (DLS). Perform at least three independent runs to measure the size and polydispersity of the liposome suspension. 
	NOTE: If necessary, use a more concentrated liposome suspension for the measurement. Alternatives to DLS (e.g., nanoparticle tracking analysis) can also be applied for measuring the size of the liposomes. A description of how to execute such particle-size determination is beyond the scope of this protocol.</li>
	<li>Image the calibration liposomes on the microscope using exactly the same experimental settings as defined in steps 4.1 and 4.2.</li>
	<li>Extract the integrated intensity for each calibration liposome in the calibration images: Extract the filtered results sheet containing liposome fluorescence intensities as described in steps 5.1&#8722;5.10. From the results table, extract the &#34;IntegratedInt&#34; column.</li>
	<li>Because the total integrated intensity of a liposome labeled in its membrane is proportional to the surface area of the liposome and is thus proportional to the square of its diameter, plot a square root intensity histogram of the fluorescence intensity of the calibration liposomes.</li>
	<li>Fit the integrated intensity histogram produced in step 6.7 with a log normal distribution and extract the average fluorescence intensity of the calibration liposomes.</li>
	<li>To determine the relation between the square root intensity (IntSqrt) and liposome size, calculate the correction factor (C) using the average liposome diameter (Dia) weighed by the number obtained from the DLS measurements: Dia = C x IntSqrt is equivalent to C = Dia / IntSqrt.</li>
	<li>Calculate IntSqrt values for the liposomes in the compositional inhomogeneity experiment and convert these to diameters by multiplying with the correction factor.</li>
	<li>Plot the intensity ratio value as a function of diameter for the compositional inhomogeneity liposomes, thus achieving the inhomogeneity as a function of liposome size for a population of liposomes spanning from approximately 50 nm&#8722;800 nm.</li>
</ol>

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The authors declare no conflict of interest.

It is important to note that while we describe in detail how the single liposomes assay can be used to study the compositional inhomogeneity between individual liposomes, the platform is very versatile. As previously shown and discussed in the introduction, the protocol can easily be adapted to study aspects of membrane-membrane fusion, protein-membrane interactions, or liposomal drug carrier characterization. For any scientific questions being addressed, the power of the single liposome assay lies in the ability to detect the individual components of the ensemble and thus having a quantitative readout, not biased by ensemble averaging effects.
The single liposome assay is optimally suited for surface imaging modalities such as total internal reflection or confocal microscope, where the Z-direction sectioning can eliminate unwanted background fluorescence from the solution above the liposome layer. However, this aspect is most important for studies where fluorescently labeled compounds, such as peptides, proteins, or other liposomes are added to the solution and remain there during imaging. For example, if the assay is used in the format described in detail here, where the fluorescent dyes are restricted to the immobilized liposomes, the assay could in principle be performed using more broadly available widefield microscopes, providing the detection system is sensitive enough.
A priori there are no limitations with respect to the method used to prepare the liposomes used in the assay. Here we describe in detail the use of a freeze-drying technique. However, previously chloroform-based lipid rehydration or ethanol injection-based techniques have been employed to fabricate liposomes used in the assay5,6. A critical parameter is the inclusion of a minute fraction of biotinylated lipids in the lipid mixture to ensure immobilization (0.05 mol% recommended). Another necessary element is to include a small amount of fluorescently labeled lipid. Overall, the amount of lipid-dye added should be kept as low as possible to both avoid quenching effects and to avoid the lipid-dye from significantly altering the physicochemical properties of the liposome. The lower limit of the amount of lipid dye is set by the concentration, where the stochastic variation in the number of individual lipid dyes per liposome becomes significant compared to the average amount of lipid-dyes (see Larsen et al.6 for an in-depth discussion). Going below this limit will introduce large uncertainties in the extracted intensities. Finally, the amount of lipid dye needs to be high enough for accurate detection of the single liposomes with a good signal-to-noise ratio. While this criterion of course heavily depends on the imaging system, we found that lipid dye concentrations between 0.05 and 0.5 mol% work well in the vast majority of cases.
To choose which lipid dye to include in the liposome, the first prerequisite is that the excitation and emission properties match the illumination and detection capabilities of the imaging system. Second, to increase the sensitivity and accuracy of the method we recommend using dyes with both high quantum yields and photostability, and we have previously successfully used dyes with different excitation wavelengths6,11. Care should be taken in choosing lipid anchors with similar physicochemical properties to ensure that lipid partitioning does not lead to artificially high inhomogeneity. For example, for this protocol, we have anchored both fluorophores using DOPE, while having one fluorophore on DOPE and another on DPPE could lead to fluorophore distribution heterogeneities not related to the inherent inhomogeneity in the liposome population. Especially for dual color imaging it is important to choose dyes with narrow excitation and emission spectra and the smallest spectral overlap as possible to avoid significant bleed-through, cross talk, and fluorescence resonance energy transfer (FRET) between dyes and channels. To avoid artifacts related to variation in the fluorophore environment we prefer to work with dyes, which have been proven experimentally to show extremely low membrane interaction propensity as determined by Hughes et al.26. Finally, if a specific liposome lipid composition is not a hard requirement for the experimental design, it can be beneficial to include up to 10 mol% of negatively charged lipids to ensure that individual liposomes repel each other and are efficiently immobilized as single liposomes12.
An advantage of the single liposome assay is the small experimental volume, which can greatly reduce the amount of expensive or rare compounds used per experiment. Here we describe the use of standard and commercially available chambers with an experimental volume between 150&#8211;300 &#181;L. However, custom-made microscope chambers that can reduce the volume to a range of 50&#8211;80 &#181;L can be used. Also, the liposome consumption is very low, with a final concentration around 2 &#181;M total lipid.
A disadvantage of the single liposome assay is that it is hard to control the lipid concentration in the chamber due to the variations in immobilization tendencies described above. Additionally, with respect to applying the assay for studying membrane-protein interactions, it is challenging to determine the concentration or number of bound proteins directly from the fluorescence intensity.
When using the liposomes in the assay as membrane model systems (e.g., to study the membrane interaction of fluorescent compounds, peptides, or proteins) it is important to ensure low non-specific binding to the protein components that facilitates the immobilization. If this is not the case, high background signal can be detected that could prevent the detection of the specific binding to the liposomes. If high non-specific background is detected we have previously successfully reduced the background by changing from streptavidin to Neutravidin or avidin.
Performing single liposome detection using other techniques like flow cytometry could potentially offer an increased detection throughput compared to the microscope-based assay. However, given that flow cytometers are optimized for studying much larger and brighter samples like cells, the detection system of most is not sensitive enough to detect the relatively weak fluorescence from an individual liposome. So while new and more sensitive flow cytometry is being developed the microscope-based assay offers the best solution for elucidating liposome inhomogeneities when performed efficiently and monitoring thousands of liposomes per experiment.

Liposomes are spherical phospholipid-based vesicles that are heavily used both in basic and applied research. They function as excellent membrane model systems, because their physiochemical properties can be easily manipulated by varying the lipid components making up the liposome1,2. Also, liposomes constitute the most used drug delivery nanocarrier system, offering improved pharmacokinetics and pharmacodynamics as well as high biocompatibility3.
For many years, liposomes have primarily been studied using bulk techniques, giving only access to ensemble average read-out values. This has led the majority of these studies to assume that all liposomes in the ensemble are identical. However, such ensemble-averaged values are only correct if the underlying dataset is uniformly distributed around the mean value, but can represent a false and biased conclusion if the dataset includes multiple independent populations, for example. Additionally, assuming the ensemble mean to represent the whole population can overlook the information harbored within the inhomogeneity between liposomes. Only recently have quantitative assays emerged that are able to probe single liposomes, revealing large inhomogeneities between individual liposomes with respect to important physicochemical properties including liposome size4, lipid composition5,6, and encapsulation efficiency7, highlighting the importance of studying liposomes at the single liposome level.
A research area where ensemble averaging of liposome properties has been shown to bias results is studying liposome size-dependent protein-membrane interactions8,9. Traditionally, researchers studying such processes have been restricted to preparing liposomes with different ensemble average diameters by extrusion through filters with different pore sizes9. However, extracting the diameter of individual liposomes using single liposome assays has revealed large population overlaps, with liposomes extruded using 100 nm and 200 nm filters displaying up to 70% overlap in their size distribution4. This could severely bias bulk measurements of liposome size-dependent protein-membrane interactions10. Performing the membrane-protein interaction studies using the single liposome assay, researchers instead took advantage of the size-polydispersity within the sample, allowing them to study a wide range of liposome diameters within each single experiment, facilitating new discoveries of how membrane curvature and composition can affect protein recruitment to membranes4,11,12. Another field where the application of single liposome assays has proven instrumental is in mechanistic studies of protein-mediated membrane fusion13,14. For such kinetic measurements, the ability to study individual fusion events alleviated the need for the experimental synchronization of the fusion process, allowing new mechanistic insights that would otherwise have been lost in the spatiotemporal averaging done in bulk ensemble measurements. Additionally, single liposomes have been used as a membrane scaffold, allowing the measurement of individual proteins and offering new knowledge on transmembrane protein structural dynamics15,16. Furthermore, such proteoliposome-based setups made it possible to study the function of individual transmembrane transporters17 and pore-forming protein complexes18 as well as the mechanism of bioactive membrane-permeabilizing peptides19. Single liposomes have also been used as soft matter nanofluidics with surface-immobilized single liposomes serving as chambers for enzymatic reactions in volumes of 10-19 L, increasing the throughput and complexity of the screening assays with minimal product consumption20.
Recently, single liposome assays have been used for characterizing drug delivery liposomes at a previously unprecendented level of detail. Researchers were able to quantify significant inhomogeneities in the amount of polymer attached to the surface of individual liposomes21. The single liposome assays also allowed measurements of drug delivery liposomes in complex media, such as blood plasma, revealing how elements anchored to the liposome surface through lipid anchors can be susceptible to dissociation when liposomes are exposed to conditions mimicking those experienced during in vivo circulation22. Overall, the versatility and usefulness of the single liposome assays are substantiated by the great variety of problems these setups have been employed to address, and we envision that the methodology will continue to be developed and find use in new scientific fields.
Here we describe a fluorescence microscopy-based single liposome assay that allows individual liposomes to be studied in a high-throughput manner (Figure 1). To illustrate the method, we use it to quantify the size and compositional inhomogeneity between individual liposomes within an ensemble. The assay employs fluorescence microscope imaging of single liposomes immobilized on a passivated glass surface. We first describe the critical steps in the liposome fabrication process that ensures proper fluorescent liposome labeling and immobilization. Then, we describe the surface preparation needed to facilitate liposome immobilization before outlining the procedure for ensuring appropriate liposome surface densities. We discuss the microscopy parameters important for acquiring high-quality images and delineate how to perform simple data analysis, allowing the extraction of liposome size and compositional inhomogeneity. This generic protocol should provide a good basis for the interested researcher to develop the assay further for his or her specific research interest.

<table><tbody><tr><td>8-well microscopy slides (&amp;micro; slides)</td><td>Ibidi</td><td>80827</td><td>Microscopy slides with glass bottom</td></tr><tr><td>Avanti Mini Extrusion kit</td><td>Avanti Polar Lipids</td><td>610000</td><td>Consumables (Whatman filters) can be aquired from GE Healthcare</td></tr><tr><td>BSA</td><td>Sigma</td><td>A9418</td><td></td></tr><tr><td>BSA-Biotin</td><td>Sigma</td><td>A8549</td><td></td></tr><tr><td>Cholesterol</td><td>Avanti Polar Lipids</td><td>700000</td><td>Traded trough Sigma</td></tr><tr><td>Computer with FIJI (Fiji Is Just ImageJ)</td><td></td><td></td><td>ComDet plugin must be installed. Also, a data handling software (Excel, MatLab, OpenOffice, GraphPad Prism etc.) able to load .txt files will be needed to plot the data</td></tr><tr><td>DOPE-Atto488</td><td>Atto-Tech</td><td>AD488-165</td><td></td></tr><tr><td>DOPE-Atto655</td><td>Atto-Tech</td><td>AD655-165</td><td></td></tr><tr><td>DOPE-PEG-Biotin</td><td>Avanti Polar Lipids</td><td>880129</td><td>Traded trough Sigma</td></tr><tr><td>D-Sorbitol</td><td>Sigma</td><td>S-6021</td><td></td></tr><tr><td>Freeze-dryer</td><td></td><td></td><td>e.g. ScanVac Coolsafe from Labogene</td></tr><tr><td>Glass vials</td><td>Brown Chromatography</td><td>150903</td><td>Glass vials that can resist snap-freezing in liquid nitrogen. The 8 mL version of the vials has a size that also fits with the syringes of the extrusion kit</td></tr><tr><td>HCl</td><td>Honeywell Fluka</td><td>258148</td><td></td></tr><tr><td>Heating bath</td><td></td><td></td><td>Capable of heating to minimum 65 &amp;deg;C</td></tr><tr><td>Heating plate with Magnet stirring</td><td></td><td></td><td>Capable of heating to minimum 65 &amp;deg;C</td></tr><tr><td>HEPES</td><td>Sigma</td><td>H3375</td><td></td></tr><tr><td>Liquid nitrogen</td><td></td><td></td><td>Including container for storage, e.g. Rubber-bath</td></tr><tr><td>Magnetic stirring bars</td><td>VWR</td><td>442-4520 (EU)</td><td></td></tr><tr><td>Microcentrifuge tubes 1.5 mL</td><td>Eppendorf</td><td>0030 120.086 (EU)</td><td></td></tr><tr><td>Microscope</td><td></td><td></td><td>For the images in this protocol a Leica SP5 confocal microscope has been used</td></tr><tr><td>Na HEPES</td><td>Sigma</td><td>H7006</td><td></td></tr><tr><td>NaCl</td><td>Sigma</td><td>S9888</td><td></td></tr><tr><td>NaOH</td><td>Honeywell Fluka</td><td>71686</td><td></td></tr><tr><td>POPC</td><td>Avanti Polar Lipids</td><td>850457</td><td>Traded trough Sigma</td></tr><tr><td>Streptavidin</td><td>Sigma</td><td>S4762</td><td></td></tr><tr><td>tert-Butanol (2-methyl-2-propanol)</td><td>Honeywell Riedel-de Ha&amp;euml;n</td><td>24127</td><td></td></tr><tr><td>Ultrapure water</td><td></td><td></td><td>e.g. MilliQ</td></tr></tbody></table>

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.

Following the protocol described makes it possible to image single liposomes in a massive parallel manner (Figure 1). The successful surface immobilization of liposomes should be immediately apparent upon the addition of the liposome solution to the chamber (step 3.6 in the protocol) as diffraction limited intensity spots should appear in the image (Figure 1B and Figure 1C).
In order to achieve good statistics and exploit the high-throughput abilities of the assay, several thousand liposomes should be imaged. In order to do this in a reasonable number of images, it is recommended to immobilize enough liposomes to achieve a density of 300&#8722;400 liposomes per frame, as this will bring the number of images per sample down to ~10, thus limiting the number of images that has to be acquired and analyzed. A lower density will make the data analysis more time-consuming, while a higher density can make it challenging for the image analysis software to distinguish single liposomes. To get a visual impression of what 300&#8722;400 liposomes look like, see Figure 2A. It should be noted, however, that for some applications (e.g., membrane-protein interactions with a protein that tends to elicit strong background binding to the BSA surface) it is recommended to use a slightly lower density, such as in Figure 2A at the top right.
Occasionally, when acquiring images it is hard to get the whole field of view in proper focus, as illustrated in Figure 2B. Such issues might indicate that the sample plate is tilting, which can arise from the plate not being placed properly on the specimen holder on the microscope. Also, if the liposomes seem large and blurry, the buffer in the chamber might have evaporated and the surface dried out. It is especially important to keep this issue in mind when imaging for long time periods or when performing measurements at temperatures higher than RT. To illustrate the difference between properly stored and dried out liposomes, an image of the same sample before and after drying out the chamber is shown in Figure 2C.
After having ensured optimal image quality, the integrated intensity for each liposome in the two imaging channels can be extracted following the steps in section 5 of this protocol. Doing this will create a list of unique intensity values, allowing us to calculate the intensity ratio for each individual liposome. The compositional inhomogeneity is evaluated from intensity ratio histograms, typically revealing a Gaussian distribution around a mean ratio value (Figure 3A). Notice that if strong deviations from a Gaussian distribution are observed (Figure 3B), it indicates a detection sensitivity issue for at least one of the imaging channels, suggesting that a subset of liposomes showing a weaker signal have been excluded. This could be due to both the detection limit of the imaging system, or excluding liposomes in the data analysis (e.g., when applying a certain minimum threshold; see step 5.7).
Calculating the DI value as described in steps 5.11&#8722;5.13 in the protocol will provide a quantitative measure of the compositional inhomogeneity between the individual liposomes of the preparation. For the liposomal system studied here, DI = 0.23 &#177; 0.01 (Figure 3C). This value can be used to systematically compare how varying liposome properties or preparation methods affect the compositional inhomogeneity. To conceptualize the meaning of the DI value, we refer to the normal distribution displayed by the intensity ratio histogram, meaning they will obey the empirical 68-95-99.7 rule. This rule describes the percentage of a population that falls within one, two, and three standard deviations around the mean value. For the DI value of 0.23 &#177; 0.01 found here, it means that 32% of the liposomes in the population will have an intensity ratio that differs by more than 23% from the mean molar ratio of the ensemble (Figure 3D).
For the control experiment imaging the same liposomes before and after refocusing (section 4.6), the same data analysis and result plotting as for the actual experiment is performed. Doing this allows for the quantification of the experimental uncertainty of the DI value, which was found to be DIuncertainty = 0.10 &#177; 0.01 (Figure 3E). Although the DIuncertainty value will depend on the employed imaging system, it was found that confocal microscope setups consistently give rise to DIuncertainty values of around 0.105,6. The DI value of DI = 0.23 &#177; 0.01 found for the liposomal system here is more than twice the experimental uncertainty, which suggests the presence of significant compositional inhomogeneity between the individual liposomes of the preparation.
Performing the size calibration experiment described in section 6 will allow the arbitrary liposome intensity values to be transformed to physical diameters in nanometers. The method has been validated against other imaging techniques25 such as cryogenic electron microscopy, which has an ability to optically resolve the nanometer-sized liposomes, although with much lower throughput. Imaging the control sample using the exact same microscope settings as used for the actual experiments, it is now possible to correlate the mean liposome intensity to the mean liposome diameter determined by DLS (Figure 4A). The next step is to depict the liposome intensity distribution, which based on the physical constraints against creating extremely small liposomes, typically will display a log normal distribution (Figure 4B). If the distribution is not well described by a log normal distribution (most often due to missing liposomes with low intensity values) it means that a part of the liposome population was excluded either due to low microscope detection sensitivity or overly strict parameters during data analysis (Figure 4C). It is important to catch and address these issues to ensure unbiased data interpretation. Next, the intensity values for each individual liposome in Figure 4B can be converted to actual diameters using the correction factor determined in Figure 4A (Figure 4D). After determining the size of each liposome, the DI as a function of diameter can now be investigated by plotting the intensity ratio versus diameter for each individual liposome (Figure 4E). This funnel-like data structure corroborates earlier findings that the DI value increases when the liposome size decreases6. Importantly, the symmetrical increase in the spread of intensity ratios around a mean value suggests that there is no systematic size-dependent variation in the average composition of liposomes.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60538/60538fig1a.jpg" /> 
Figure 1: The single liposome assay. (A) Liposomes are formulated with an equimolar content of the fluorescently labeled lipids DOPE-Atto488 and DOPE-Atto655 and immobilized on a BSA surface using a biotin/streptavidin linkage. (B) The immobilized liposomes are imaged using a confocal microscope, allowing for detection of the fluorescence intensities from the single liposomes. Scale bars = 8 &#181;m. (C) Zoom of the green area in (B) depicted as surface intensity plot for two adjacent liposomes displaying inverted fluorescent signal between the two channels, illustrating the concept of compositional inhomogeneity. <a href="https://www.jove.com/files/ftp_upload/60538/60538fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60538/60538fig2a.jpg" /> 
Figure 2: Optimization and pitfalls. (A) The top left shows 29 liposomes immobilized in a 50 &#181;m x 50 &#181;m frame. These are too few liposomes per frame to exploit the possibility for high-throughput investigation of liposome inhomogeneity. The top right shows 123 liposomes per frame. This is a suboptimal density but can be used if many images are acquired. The bottom left shows 300&#8722;400 liposomes per frame. This is optimal for the protocol described here. The bottom right shows more than 1,500 liposomes per frame, which is too many to distinguish individual liposomes. There is a high risk that a single spot will actually be two liposomes immobilized within the same diffraction limited spot. (B) If the chamber is not placed properly in the specimen holder, getting the whole field of view in proper focus will be impossible. In the left micrograph the plate is tilting around the traverse axis, giving rise to the lower left corner being out of focus. In the right micrograph the plate is tilting around the longitudinal axis, giving rise to a heavier tilt. Both the lower left and upper right corner are out of focus. (C) If imaging for long time periods or at elevated temperatures the buffer might evaporate, leading to drying out of the chamber. We illustrated this scenario here by imaging liposomes before and after the chamber was left to dry for 3 min, and then rewetted by adding fresh buffer to the chamber. The resulting liposomes are larger, blurry, have lower fluorescence intensity, and appear to be spread out on the plate. <a href="https://www.jove.com/files/ftp_upload/60538/60538fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60538/60538fig3a.jpg" /> 
Figure 3: Intensity ratio histograms for quantifying liposome compositional inhomogeneity. (A) The intensity ratio of DOPE-Atto488 and DOPE-Atto655 fluorescence for each individual liposome plotted as a histogram. (B) A truncated Gaussian distribution illustrating a potential sensitivity issue, either during imaging or in the data treatment step. (C) Fitting the intensity histogram with a Gaussian function makes it possible to extract the mean and standard deviation used to calculate the DI value. (D) A DI value of 0.23 can be translated to 32% of the population differing by more than 23% from the mean molar ratio of the ensemble. (E) Control experiment imaging the same liposomes before and after refocusing and then performing the same data treatment procedure as for the actual experiment. This allows the experimental error for the DI quantification to be determined. <a href="https://www.jove.com/files/ftp_upload/60538/60538fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60538/60538fig4a.jpg" /> 
Figure 4: Converting liposome intensity to physical diameter in nm. (A) Intensity histogram of the 50 nm extruded calibration liposomes compared with the DLS data in order to determine the calibration factor. (B) A histogram of the IntSqrt value from the experimental liposomes typically yields a log-normal distribution due to the physical constraints against creating very small liposomes. (C) A non log-normal IntSqrt value histogram indicates either issues with detection sensitivity or that small liposomes were excluded during data treatment. (D) Liposome intensities in (B) are converted to actual liposome diameters using the correction factor. (E) The intensity ratio of each liposome plotted as a function of liposome diameter can now be displayed allowing the investigation of compositional inhomogeneity as a function of liposome size. <a href="https://www.jove.com/files/ftp_upload/60538/60538fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Quantitative Fluorescence Microscopy-based Single Liposome Assay for Detecting the Compositional Inhomogeneity Between Individual Liposomes at JoVE.com

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

This protocol describes the fabrication of liposomes and how these can be immobilized on a surface and imaged individually in a massive parallel manner using fluorescence microscopy. This allows for the quantification of the size and compositional inhomogeneity between single liposomes of the population.

Most research employing liposomes as membrane model systems or drug delivery carriers relies on bulk read-out techniques and thus intrinsically assumes ...

a-quantitative-fluorescence-microscopy-based-single-liposome-assay

Universidad San Sebastian

Research

JoVE Journal

Bioengineering

7.8K Views.  Technical University of Denmark. Het meeste onderzoek met liposomen vertrouwen op gewaagde technieken intrinsiek veronderstellend alle liposomen om identiek te zijn. Door het bestuderen van enkele liposomen, significante inhomogeniteiten kunnen worden waargenomen. De enkele liposoom techniek die hier wordt gepresenteerd stelt ons in staat om honderden liposomen tegelijkertijd te bestuderen en inhomogeniteiten in hun grootte en samenstelling te detecteren.De methode kan eenvoudig worden aangepast om andere systemen te ...

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

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

Single Liposome Measurements for the Study of Proton-Pumping Membrane Enzymes Using Electrochemistry and Fluorescent Microscopy

Proton-pumping enzymes of electron transfer chains couple redox reactions to proton translocation across the membrane, creating a proton-motive force used for ATP production. The amphiphilic nature of membrane proteins requires particular attention to their handling, and reconstitution into the natural lipid environment is indispensable when studying membrane transport processes like proton translocation. Here, we detail a method that has been used for the investigation of the proton-pumping mechanism of membrane redox enzymes, taking cytochrome bo3 from Escherichia coli as an example. A combination of electrochemistry and fluorescence microscopy is used to control the redox state of the quinone pool and monitor pH changes in the lumen. Due to the spatial resolution of fluorescent microscopy, hundreds of liposomes can be measured simultaneously while the enzyme content can be scaled down to a single enzyme or transporter per liposome. The respective single enzyme analysis can reveal patterns in the enzyme functional dynamics that might be otherwise hidden by the behavior of the whole population. We include a description of a script for automated image analysis.

Here, we present a protocol to study the molecular mechanism of proton translocation across lipid membranes of single liposomes, using cytochrome bo3 as an example. Combining electrochemistry and fluorescence microscopy, pH changes in the lumen of single vesicles, containing single or multiple enzyme, can be detected and analyzed individually.

Proton-pumping enzymes of electron transfer chains couple redox reactions to proton translocation across the membrane, creating a proton-motive force used ...

Proteome-wide Quantification of Labeling Homogeneity at the Single Molecule Level

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a fluorescent dye and are spotted by a photodetector following their separation. Single molecule fluorescence imaging can provide ultrasensitive protein detection with its ability for visualizing individual fluorescent molecules. However, the application of this powerful imaging method to electrophoresis assays is hampered by the lack of ways to characterize the homogeneity of fluorescent labeling of each protein species across the proteome. Here, we developed a method to evaluate the labeling homogeneity across the proteome based on a single molecule fluorescence imaging assay. In our measurement using a HeLa cell sample, the proportion of proteins labeled with at least one dye, which we termed &#8216;labeling occupancy&#8217; (LO), was determined to range from 50% to 90%, supporting the high potential of the application of single molecule imaging to sensitive and precise proteome analysis.

Here, we present a protocol to assess the labeling homogeneity for each protein species in a complex protein sample at the single molecule level.

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a ...

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 ...

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 ...

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

SNARE-mediated Fusion of Single Proteoliposomes with Tethered Supported Bilayers in a Microfluidic Flow Cell Monitored by Polarized TIRF Microscopy

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. During neurotransmitter or hormone release via exocytosis, the fusion pore can transiently open and close repeatedly, regulating cargo release kinetics. Pore dynamics also determine the mode of vesicle recycling; irreversible resealing results in transient, &#34;kiss-and-run&#34; fusion, whereas dilation leads to full fusion. To better understand what factors govern pore dynamics, we developed an assay to monitor membrane fusion using polarized total internal reflection fluorescence (TIRF) microscopy with single molecule sensitivity and ~15 msec time resolution in a biochemically well-defined in vitro system. Fusion of fluorescently labeled small unilamellar vesicles containing v-SNARE proteins (v-SUVs) with a planar bilayer bearing t-SNAREs, supported on a soft polymer cushion (t-SBL, t-supported bilayer), is monitored. The assay uses microfluidic flow channels that ensure minimal sample consumption while supplying a constant density of SUVs. Exploiting the rapid signal enhancement upon transfer of lipid labels from the SUV to the SBL during fusion, kinetics of lipid dye transfer is monitored. The sensitivity of TIRF microscopy allows tracking single fluorescent lipid labels, from which lipid diffusivity and SUV size can be deduced for every fusion event. Lipid dye release times can be much longer than expected for unimpeded passage through permanently open pores. Using a model that assumes retardation of lipid release is due to pore flickering, a pore &#34;openness&#34;, the fraction of time the pore remains open during fusion, can be estimated. A soluble marker can be encapsulated in the SUVs for simultaneous monitoring of lipid and soluble cargo release. Such measurements indicate some pores may reseal after losing a fraction of the soluble cargo.

Here, we present a protocol to detect single, SNARE-mediated fusion events between liposomes and supported bilayers in microfluidic channels using polarized TIRFM, with single molecule sensitivity and ~15 msec time resolution. Lipid and soluble cargo release can be detected simultaneously. Liposome size, lipid diffusivity, and fusion pore properties are measured.

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. ...

Neuroscience

Preparation, Administration, and Assessment of In Vivo Tissue-Specific Cellular Uptake of Fluorescent Dye-Labeled Liposomes

There is a growing interest in using liposomes to deliver compounds in vivo particularly for targeted treatment approaches. Depending on the liposome formulation, liposomes may be preferentially taken up by different cell types in the body. This may influence the efficacy of the therapeutic particle as progression of different diseases is tissue- and cell-type-specific. In this protocol, we present one method for synthesizing and fluorescently labeling liposomes using DSPC, cholesterol, and PEG-2000 DSPE and the lipid dye DiD as a fluorescent label. This protocol also presents an approach for delivering liposomes in vivo and assessing cell-specific uptake of liposomes using flow cytometry. This approach can be used to determine the types of cells that take up liposomes and quantify the distribution and proportion of liposome-uptake across cell types and tissues. While not mentioned in this protocol, additional assays such as immunofluorescence and single-cell fluorescence imaging on a cytometer will strengthen any findings or conclusions made as they permit assessment of intracellular staining. Protocols may also need to be adapted depending on the tissue(s) of interest.

The goal of this protocol is to synthesize fluorescently-labeled liposomes and use flow cytometry to identify in vivo localization of liposomes at a cellular level.

There is a growing interest in using liposomes to deliver compounds in vivo particularly for targeted treatment approaches. Depending on the liposome ...

A Fluorescence-based Assay for Characterization and Quantification of Lipid Droplet Formation in Human Intestinal Organoids

Dietary lipids are taken up as free fatty acids (FAs) by the intestinal epithelium. These FAs are intracellularly converted into triglyceride (TG) molecules, before they are packaged into chylomicrons for transport to the lymph or into cytosolic lipid droplets (LDs) for intracellular storage. A crucial step for the formation of LDs is the catalytic activity of diacylglycerol acyltransferases (DGAT) in the final step of TG synthesis. LDs are important to buffer toxic lipid species and regulate cellular metabolism in different cell types. Since the human intestinal epithelium is regularly confronted with high concentrations of lipids, LD formation is of great importance to regulate homeostasis. Here we describe a simple assay for the characterization and quantification of LD formation (LDF) upon stimulation with the most common unsaturated fatty acid, oleic acid, in human intestinal organoids. The LDF assay is based on the LD-specific fluorescent dye LD540, which allows for quantification of LDs by confocal microscopy, fluorescent plate reader, or flow cytometry. The LDF assay can be used to characterize LD formation in human intestinal epithelial cells, or to study human (genetic) disorders that affect LD metabolism, such as DGAT1 deficiency. Furthermore, this assay can also be used in a high-throughput pipeline to test novel therapeutic compounds, which restore defects in LD formation in intestinal or other types of organoids.

This protocol describes an assay for the characterization of lipid droplet (LD) formation in human intestinal organoids upon stimulation with fatty acids. We discuss how this assay is used for quantification of LD formation, and how it can be used for high throughput screening for drugs that affect LD formation.

Dietary lipids are taken up as free fatty acids (FAs) by the intestinal epithelium. These FAs are intracellularly converted into triglyceride (TG) ...

Medicine

Membrane Transport Processes Analyzed by a Highly Parallel Nanopore Chip System at Single Protein Resolution

Membrane protein transport on the single protein level still evades detailed analysis, if the substrate translocated is non-electrogenic. Considerable efforts have been made in this field, but techniques enabling automated high-throughput transport analysis in combination with solvent-free lipid bilayer techniques required for the analysis of membrane transporters are rare. This class of transporters however is crucial in cell homeostasis and therefore a key target in drug development and methodologies to gain new insights desperately needed.
The here presented manuscript describes the establishment and handling of a novel biochip for the analysis of membrane protein mediated transport processes at single transporter resolution. The biochip is composed of microcavities enclosed by nanopores that is highly parallel in its design and can be produced in industrial grade and quantity. Protein-harboring liposomes can directly be applied to the chip surface forming self-assembled pore-spanning lipid bilayers using SSM-techniques (solid supported lipid membranes). Pore-spanning parts of the membrane are freestanding, providing the interface for substrate translocation into or out of the cavity space, which can be followed by multi-spectral fluorescent readout in real-time. The establishment of standard operating procedures (SOPs) allows the straightforward establishment of protein-harboring lipid bilayers on the chip surface of virtually every membrane protein that can be reconstituted functionally. The sole prerequisite is the establishment of a fluorescent read-out system for non-electrogenic transport substrates.
High-content screening applications are accomplishable by the use of automated inverted fluorescent microscopes recording multiple chips in parallel. Large data sets can be analyzed using the freely available custom-designed analysis software. Three-color multi spectral fluorescent read-out furthermore allows for unbiased data discrimination into different event classes, eliminating false positive results.
The chip technology is currently based on SiO2 surfaces, but further functionalization using gold-coated chip surfaces is also possible.

The presented protocol describes the analysis of membrane protein mediated transport on the single transporter level using pore-spanning solvent-free lipid bilayers. This is achieved by the creation of bulk produced nanopore array chips, combined with highly parallel data acquisition and analysis, enabling the future establishment of membrane protein effector screenings.

Membrane protein transport on the single protein level still evades detailed analysis, if the substrate translocated is non-electrogenic. Considerable ...

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

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