This work is supported by the KAIST-funded K-Valley RED&#38;B Project for 2014 and the Basic Science Research Program through the National Research Foundation of Korea (NRF- 2012R1A6A3A040395, NRF-2013R1A1A2057708, NRF- 2012R1A1A1011023).

Caution: Refer to material safety data sheets (MSDS) before use of acetone and chloroform which are carcinogenic.
1. Preparation of Phospholipid Monolayer at a Flat Air-water Interface
<ol>
	<li>Formation of a phospholipid monolayer
	<ol>
		<li>Preparation of a phospholipid solution
		<ol>
			<li>Clean a 4 ml vial with a polytetrafluoroethylene coated cap using acetone, ethanol, and deionized water at least three times, and blow nitrogen gas thoroughly into the vial to get rid of water.</li>
			<li>Dissolve 1 mg of phospholipid (e.g., dioleoylphosphatidylcholine, DOPC) to 1 ml of chloroform in the vial to obtain 1 mg/ml of concentration. Perform this procedure in a fume hood for safety. Use lower or higher concentrations of phospholipid solution if necessary.</li>
			<li>Add dye tagged phospholipids (e.g., Rhodamine dipalmitoylphosphatidylethanolamine, Rhodamine DPPE) with less than 1 mol % of the phospholipid solution, in order to visualize the phospholipid monolayer with a fluorescence microscope. Perform this procedure in a fume hood for safety.</li>
			<li>Wrap the vial with a polytetrafluoroethylene tape to prevent solvent evaporation, and store the sample in a freezer at -20 &#176;C.</li>
		</ol>
		</li>
		<li>Deposition of phospholipids onto the air-water interface
		<ol>
			<li>Clean a Petri dish (55 mm in diameter and 12 mm in height) with ethanol, and deionized water at least three times.</li>
			<li>Fill the Petri dish with 10 ml of deionized water to create an air-water interface.</li>
			<li>Spread few microliters of the phospholipid solution with a micro-syringe onto a clean interface to achieve desired surface pressure and wait for at least 30 min&#160;to evaporate the solvent completely, prior to doing the experiment. 
			Note: A Langmuir trough can be used instead of a Petri dish if precise control of surface pressure is necessary.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Surface pressure measurement
	<ol>
		<li>Measure the surface pressure of the phospholipid monolayer with a Wilhelmy plate tensiometer. Wait for at least 30 min to get the filter paper wetted enough if filter paper is used as a Wilhelmy plate. The detailed protocol is available in Kuhn et al.20.</li>
		<li>Adjust the deposited amount of the phospholipid to control the surface pressure precisely. Add 4 &#181;l of the DOPC solution (1 mg/ml) onto the 30.25 cm2&#8211;area of the interface if ~ 5 mN/m of surface pressure is required. 
		Note: If the filter paper is not fully wetted, surface pressure changes dramatically during the first 30 min of the experiment due to the weight change of the filter paper.</li>
	</ol>
	</li>
	<li>Minimization of the convective flow of the monolayer
	<ol>
		<li>Use a cone-shaped apparatus that contains a 3 mm reservoir with two thin channels, connected to a large section of the Petri dish, to suppress the convective flow of the monolayer, which can disturb morphology imaging and surface pressure measurement.</li>
		<li>Make sure to match the water levels between the inside and outside of the cone-shape apparatus to ensure that phospholipids move freely over the whole region before depositing phospholipids onto the air-water interface.</li>
	</ol>
	</li>
</ol>
2. Preparation of Phospholipid Monolayer at the Curved Surface of a Droplet
<ol>
	<li>Tapering process of a glass capillary using micropipette puller
	<ol>
		<li>Place a glass capillary (o.d. 1 mm, i.d. 0.78 mm, length 100 mm) on a capillary holder of a micropipette puller.</li>
		<li>Design a program for pulling the capillaries with appropriate parameter values (Heat: Ramp, Pull: 60, Vel: 70, Delay: 70 and Pressure 200) and pull the capillaries with the designed program according to manufacturer&#8217;s protocol. 
		Note: A capillary end with a few micrometers-diameter is necessary to form a 100 &#181;m-droplet by applying a pressure with ~ 10 kPa. If the tip end of the capillary is too small, much higher pressure, &#62; 600 kPa, is required to obtain the droplet, while it is hard to control the size of the droplets with a too large capillary tip end.</li>
	</ol>
	</li>
	<li>Absorption of phospholipids onto the droplet surface 
	Note: To form a phospholipid monolayer at the curved interface of a droplet, a phospholipid solution without dye tagged phospholipids is used to obtain an intensity contrast, against the flat monolayer, prepared by procedure 1. In addition, both procedures 2.2.1 and 2.2.2 are allowable here. If precise control of the surface pressure at the droplet surface is necessary, procedure 2.2.2 is highly recommended. However, if not, procedure 2.2.1, which is a much easier way than procedure 2.2.2, is useful.
	<ol>
		<li>Coating process of phospholipids at a tip end
		<ol>
			<li>Clean a slide glass using acetone, ethanol, and deionized water at least three times.</li>
			<li>Place the tapered capillary on a cleaned slide glass, and tilt the capillary to facilitate touching the slide glass with a tip end.</li>
			<li>Drop a few droplets of phospholipid solution (1 mg/ml) onto the tip end of a capillary that is adhered to the glass slide using a glass syringe, and wait at least 30 min to evaporate the solvent completely. 
			Note: It is highly recommended to adhere the tip end of the capillary to the glass slide during procedure 2.2.1.3 because the perimeter of the tip end is too small to hold the droplet of the phospholipids at the tip end only by capillary force.</li>
		</ol>
		</li>
		<li>Preparation of a vesicle solution
		<ol>
			<li>Clean a vial, and remove water from the vial, as introduced in procedure 1.1.1.1.</li>
			<li>Dry a 2 ml volume of phospholipid solution (1 mg/ml) by applying nitrogen gas gently, and desiccate the vial for 1 hr at RT to eliminate any remaining solvent. Perform this procedure in a fume hood for safety.</li>
			<li>Add 2 ml of deionized water into the vial containing dried lipids, and incubate the vial in an oven at 60 &#176;C for 1 hr.</li>
			<li>Shake the vial several times, and sonicate (HF-frequency: Up to 40 kHz, Power: 370 W) for 30 min to obtain vesicles.</li>
			<li>Perform extrusion and freeze-thaw processes to obtain monodisperse unilamellar vesicles. The detailed protocol for preparing vesicles is described in Mayer et al.21. 
			Note: Surface pressure of the droplet interface is controlled precisely by adjusting the waiting time after formation of the droplet that contains monodisperse unilamellar vesicles. Using a pendant drop method22, it is necessary to measure the change of surface pressure according to time, prior to doing the experiment.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Formation of a droplet that contains a phospholipid monolayer at the curved surface
	<ol>
		<li>Fill in the tapered capillary with 10 &#181;l of deionized water from procedure 2.2.1. Alternatively, use 10 &#181;l of the vesicle solution from procedure 2.2.2.</li>
		<li>Connect the capillary to an automated micro-injector to provide the pressure to form the droplet.</li>
		<li>Mount the capillary that is connected to a micro-injector to a micromanipulator to control the position of the capillary precisely.</li>
		<li>Prepare a bright field microscope (Microscope 1, objective lens: 10X NA 0.3) for imaging the lateral view of the capillary with a CCD camera. Microscope 1 thus enables to observe the precise position of the droplet along the z-axis and estimate the size of the droplet.</li>
		<li>Move the tip end to a position where the lateral view of the tip end is well visualized by Microscope 1 using a micromanipulator, and apply a variable pressure (~ 100 hPa) with the tip end of the capillary, until an appropriate size (~100 &#181;m in diameter) of droplet is formed. 
		Note: It is recommended that automated micro-injector and micromanipulator be used, if fine control of the droplet size and location of the droplet is necessary. It is also possible to use manual ones.</li>
	</ol>
	</li>
</ol>
3. Imaging Fluorescence Recovery after Droplet Merging
Note: The fundamental principles of this protocol are identical to those of the FRAP technique, except for the drop coalescence process. The detailed protocol and related theories of FRAP are available in A. Lopez et al.15 and D. Axelrod et al.16.
<ol>
	<li>Monitoring and controlling the location of the droplet
	<ol>
		<li>Prepare an inverted microscope set (Microscope 2, objective lens: 10X NA 0.3, tube lens: focal length 17 cm) that enables both a fluorescence microscope with a proper filter set for Rhodamine-DPPEs (excitation at 560 nm, emission at 583 nm) and a bright field microscope. Use a CCD camera here for visualizing the top views of the droplet with a fluorescence microscope mode and a bright-field microscope mode.</li>
		<li>Move the droplet coated with a phospholipid to a flat air-water interface along the z-axis, but do not merge the droplet yet, using the micromanipulator. Use Microscope 1 to visualize the lateral view of the droplet.</li>
		<li>Locate the droplet at the center of the top view of the flat monolayer, using the micromanipulator. Use the bright-field microscope mode of Microscope 2 to visualize top view of flat monolayer.</li>
	</ol>
	</li>
	<li>Merging the droplet onto the flat air-water interface
	<ol>
		<li>Move the droplet further toward the flat interface until the droplet merges onto the interface, using the micromanipulator. If the merging process is done successfully, a dark region with a circular shape that is surrounded by a white background, is observed using the fluorescence mode of Microscope 2.</li>
		<li>Record the series of fluorescent images according to time after merging the droplet, using the fluorescence mode of Microscope 2. Use a faster frame rate than the diffusion time scale of the monolayer here. In DOPC monolayer, it takes a few minutes to diffuse into the 200 &#181;m-dark area completely.</li>
	</ol>
	</li>
</ol>
4. Determining the Diffusion Coefficient by Image Analysis
Note: To determine the diffusion coefficient from a series of images, customized program for image analysis is built as described below. A detailed source code of this program is available in JoVE website.
<ol>
	<li>Detection of a circular region of interest
	<ol>
		<li>Detection of the center of the region of interest
		<ol>
			<li>Obtain a series of fluorescent images that have been recorded during a recovery process which contains a circular dark area and a white background, and set the first image of the series as a reference image. Here, RD is the radius of the dark area in a reference image.</li>
			<li>Convolute the reference image with a white circle whose intensity is uniform over a whole region. Use embedded function named &#8216;conv2&#8217; for convolution as shown in the source code of the customized program line 124. Here, the radius of the white circle is slightly smaller than RD.</li>
			<li>Find a position that indicates a minimum value in the convolution calculation, and set this position as the center of the region of interest in the reference image.</li>
		</ol>
		</li>
		<li>Determining a radius of the region of interest
		<ol>
			<li>Convolute the reference image with a white circle that contains a center position, determined by a procedure 4.1 and uniform intensity over a whole region. Here, RS is the radius of the white circle. Use iteration such as &#8216;for&#8217; or &#8216;while&#8217; with equation of a circle to make the white circle as shown in the source code of the customized program line 102 to 109.</li>
			<li>Convolute the reference image with another white circle which has a slightly larger radius (5%&#8211;10%), RL, than RS. Use same method with procedure 4.1.2.1 to make the circle as shown in the source code of the customized program line 113 to 120.</li>
			<li>Obtain the difference in value between the two convolution calculations of 4.1.2.1 and 4.1.2.2.</li>
			<li>Repeat the above procedures from RS = RD/2 to RS = 2RD, by increasing both the RS and RL with a pixel level. Make an iteration that contains whole source codes from procedure 4.1.2.1 to 4.1.2.3 to repeat.</li>
			<li>Find the radius of RS that indicates the maximum difference in value between the two convolution calculations. Use embedded function named &#8216;max&#8217; to find the radius indicates the maximum difference in value as shown in the source code of the customized program line 148 to 149. This RS thus indicates the radius of the dark area in the reference image.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Calculation of a fractional intensity
	<ol>
		<li>Set a circle that contains a center position and a radius, determined by procedure 4.1 as a region of interest.</li>
		<li>Calculate the average intensities of the region of interest in a series of fluorescence images according to time. Convolute each frame with region of interest and divide it by area of region of interest to calculate the average intensity. The details are available in the source code of the customized program which is available in JoVE website.</li>
		<li>Calculate a fractional intensity, defined as the equation below, where F(t) is the average intensity in the circle that is a region of interest according to time, Fi is the initial intensity in the circle, and Fo is the intensity of a white background. 
		f(t) = (F(t) &#8211; Fi)/(Fo &#8211; Fi)&#160;&#160;&#160;&#160;&#160; (Equation 1).</li>
	</ol>
	</li>
	<li>Fitting the fractional intensity to FRAP theory
	<ol>
		<li>Fit the fractional intensity with the equation below, where &#964; is a characteristic diffusion time and I0, I1 are modified Bessel functions, using a fitting program, and obtain &#964; 
		<img alt="Equation 2" src="/files/ftp_upload/53376/53376eq2.jpg" />&#160;&#160;&#160;&#160;&#160; (Equation 2).</li>
		<li>Obtain a diffusion coefficient based on the relation, &#964; = a2/4D, where D is the diffusion coefficient and a is the radius of the dark area. 
		Note: During the fitting process, it is possible to shift the fractional intensity profile along a time axis when the recovery process has already started, before recording the images.</li>
	</ol>
	</li>
</ol>

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The authors declare no competing financial interest.

FRAM shares a lot of fundamental principles with FRAP, especially for measuring fluorescence recovery after bleaching a specific area, but FRAM uses a process of merging a droplet onto the flat interface to form a bleached area, instead of applying an intensive light. The merging process is thus most important step in FRAM, and especially, the shape of the dark stain at the flat monolayer after merging a droplet determines the accuracy of the diffusivity measurement. Specifically, based on a current method, we only set the circle with a radius R as a region of interest to calculate the average intensity of the dark stain, shown in procedure 4, and if there is a too much deviation from the circular shape, this would disturb the precise measurement of a diffusion coefficient. Therefore, it is necessary to obtain a circular shaped dark region, but, noncircular shapes are formed occasionally due to several reasons. First, if there exists a convective flow in the Petri dish, the shape of the dark area becomes elongated along the direction of flow. As mentioned in procedure 1.3, a cone-shape apparatus helps to suppress the convective flow, and this elongation would be minimized. Second, if the tip end of the capillary touches the flat interface during a merging process, the dark area is formed with a variety of shapes since the capillary end interrupts the merging process. By obtaining a droplet that is only hanging from the very end of the capillary, this interruption can be prevented. In particular, a hydrophobic treatment of the capillary helps the droplet to sit at the very end of the capillary. Therefore, above trouble shootings and modifications enable us to measure diffusion properties more precisely.
This well troubleshot process thus allows FRAM to have several significant advantages over FRAP. First, FRAM requires simpler equipment compared to FRAP. For FRAM, it is sufficient to use a simple fluorescence microscope, instead of an expensive confocal microscope with a high power laser whose wavelength must match with the absorption spectrum of the dye. In addition, it is necessary to use additional apparatus to adjust the size to the bleached area in the FRAP, while the FRAM controls the size of the area easily by adjusting the size of the droplet, or the surface pressure at the droplet surface. Second, it is possible to use various species of dyes in FRAM. FRAP only uses dye molecules whose bleaching process is much faster than the diffusion process. If the diffusion of the dyes occurs during the bleaching process, it is hard to estimate the diffusion property accurately. The FRAM, however, does not require a process for bleaching dyes and this thus enables the use of various types of dyes in the fluorescence imaging. In addition, the FRAP requires additional high power lasers and filter sets to change the dye species, while it is only necessary to replace the filter sets in the FRAM. Finally, since the monolayers at the droplet surface and the flat interface can be formed independently, thus enabling the study of inter-diffusion or mixing between two different lipid monolayers by merging A-type monolayer to B-type monolayer, while the FRAP only measures the self-diffusion of a monolayer.
Despite these advantages, the process of merging a droplet onto a flat air-water interface can potentially limit this technique due to several issues that might be of concern, such as a surface pressure mismatch between the two monolayers, the time scale of the merging process, and an increase in the surface pressure of the flat monolayer after merging droplets. These issues, however, do not affect the diffusion measurement significantly for the following reasons. First, even though the surface pressures at the two different monolayers are quite different, an equilibration process is completed at a very early stage during the recovery process. For example, if the surface pressure of the monolayer at the droplet surface is higher than that of the monolayer at the flat air-water interface, the dark area expands until the surface pressure equilibrates. Since this process is completed within 10 msec, the surface pressure will be equilibrated before it affects the diffusion process significantly. Second, if the time scale of the merging process is fast enough, it does not limit the diffusion measurement at all. Fortunately, a typical merging process is also completed within 10 msec. Finally, even multiple merging of the droplets onto the flat interface do not increase the surface pressure since the surface area of the trough is far larger than the surface area of the droplet. According to the sizes of the trough and the droplets, described in the protocol, the area per molecule increases less than 0.015% by merging a droplet. Accordingly, the increase of surface pressure due to the change of area per molecule is less than 0.1%, which is negligible because it is much smaller than the typical error in surface pressure, as measured by Wilhelmy plate tensiometer.
In summary, we introduced a new method to measure the lateral diffusion property of a phospholipid monolayer by merging a droplet monolayer onto the flat interface. This technique requires relatively simple equipment and also enables the use of various sorts of dye species. The FRAM is thus potentially applicable for diffusion measurement of any surface active agents, including phospholipids, block copolymers, proteins and even nanoparticles at a fluid-fluid interface. Furthermore, we expect that the FRAM provides a new way to study the inter-diffusion or mixing between two different surface active agents.

Phospholipids are major components of cell membranes and the membranes of organelles, and the fluidity of these membrane layers often affects cellular functions by altering the activity of membrane proteins1&#8211;3. For instance, membrane&#160;lipid&#160;homeostasis&#160;is achieved by regulating membrane fluidity, which is detected by membrane proteins, and an abnormal level of fluidity causes severe diseases, such as hepatic steatosis and cholestasis4. In addition, fluidity of a phospholipid monolayer at the air-alveoli fluid interface has important implications in its functions. High fluidity of the lung surfactant phospholipid monolayer facilitates spreading of the layer during inhalation, whereas a low fluidity enables the layer to stay within the alveoli during exhalation5,6. It is thus important to estimate the fluidity of the phospholipid layers to understand their roles.
A direct measure of the fluidity is viscosity, and the viscosity of phospholipid layers has been measured by using micron-sized colloids7, magnetic needles8,9, and magnetic micro-buttons6,10,11. However, these techniques are limited to relatively rigid films, and cannot measure less viscous films. For such cases, diffusivity would be an alternative to quantify fluidity of phospholipid layers. For several decades, a variety of techniques to measure the diffusion properties of phospholipid layers have been developed, such as fluorescence quenching method12, pulsed field gradient NMR13, and fluorescence correlation spectroscopy (FCS)14. One of the most representative methods is fluorescence recovery after photobleaching (FRAP)15,16. Due to the simplicity of the measurement procedure and related theories, several studies on the diffusion properties of phospholipid layers have been performed using FRAP17&#8211;19. However, FRAP usually requires an expensive confocal microscope setup with a high-power laser.
Here, we present a new technique to measure the lateral diffusion of phospholipid monolayers, termed as fluorescence recovery after merging (FRAM). The key difference between FRAP and FRAM is that the photobleaching step is replaced by the drop coalescence. Merging a droplet covered by non-fluorescence monolayer onto a fluorescently labeled flat monolayer leaves a circular dark stain on a bright flat monolayer, thus setting up the same initial state with the photobleaching step. We then observe the fluorescence recovery in the dark stain with time to measure the lateral diffusivity. This new &#8220;bleaching&#8221; step by drop coalescence provides significant advantages over FRAP: FRAM only requires a fluorescence microscope rather than an expensive confocal microscope with a high power laser that is necessary for FRAP. In addition, FRAM has a wide selection of fluorescence dyes since it does not involve the photobleaching process. Finally, the two different monolayers, a droplet monolayer and a flat monolayer, can be prepared independently, thus allowing interdiffusion to be measured, while FRAP only measures the self-diffusion of monolayers.
FRAM provides a wide range of diffusion measurements from highly viscous materials to nearly inviscid materials. In principal, FRAM can measure the maximum diffusivity of 106 &#181;m2/sec, which is comparable to existing techniques, when the diffusion occurs across the area 100 &#181;m by 100 &#181;m during the time for the drop coalescence process (~ 10 msec). Furthermore, slow diffusion process can be easily measured unless monolayers are solid. FRAM can be used for any kinds of surface active materials as long as appropriate fluorescently tagged molecules are available.

<table><tbody><tr><td>Acetone</td><td>OCI corporation</td><td>Acetone 3.8L Extra Pure</td><td>Purity: &amp;ge; 99.5%, Please consult material safety data sheets (MSDS) before use.</td></tr><tr><td>Ethanol</td><td>OCI corporation</td><td>Ethanol 3.8L Extra Pure</td><td>Purity: &amp;ge; 94%, Please consult material safety data sheets (MSDS) before use.</td></tr><tr><td>Dioleoylphosphatidylcholine, DOPC</td><td>Avanti Polar Lipids</td><td>850375C</td><td>Please consult material safety data sheets (MSDS) before use. Chloroform is carcinogenic.</td></tr><tr><td>Chloroform</td><td>LiChrosolv</td><td>Chloroform ultrapure (A3633)</td><td>Purity: &amp;ge; 99.8%, Please consult material safety data sheets (MSDS) before use. Carcinogenic</td></tr><tr><td>Rhodamine DPPE</td><td>Avanti Polar Lipids</td><td>810158C, 810158P</td><td>Avoid direct light exposure to prevent photobleaching</td></tr><tr><td>Wilhelmy plate tensiometer</td><td>R&amp;amp;K ultrathin organic film technology</td><td>Wilhelmy tensiometer</td><td>http://www.rieglerkirstein.de/index.htm</td></tr><tr><td>Micropipette puller</td><td>Sutter instrument</td><td>P-1000</td><td></td></tr><tr><td>Micro-injector</td><td>Eppendorf</td><td>Femtojet</td><td></td></tr><tr><td>Micromanipulator</td><td>Eppendorf</td><td>Micromanipulator 5171</td><td></td></tr><tr><td>Microscope 1</td><td>Objective lens: Olympus</td><td>Objective lens: UPlanFl 10x</td><td>Objective lens: 10X NA 0.3</td></tr><tr><td>Microscope 2</td><td>Objective lens: Olympus, Tube lens: Thorlabs</td><td>Objective lens: UPlanFl 10x</td><td>Objective lens: 10X NA 0.3, tube lens: focal length 17 cm</td></tr><tr><td>CCD for Microscope 1</td><td>Jai</td><td>CV 950 camera</td><td></td></tr><tr><td>CCD for Microscope 2</td><td>Andor</td><td>iXon3 EMCCD</td><td></td></tr><tr><td>Filter set</td><td>Chroma technology</td><td>Catalog Set 49004: ET545, T565, ET605</td><td>Prepare suitable for dye molecules</td></tr><tr><td>Sonicator</td><td>DAIHAN Scientific</td><td>Wiseclean WUC-D06H</td><td></td></tr></tbody></table>

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.

A series of fluorescence images were obtained with time during the recovery process after merging a droplet coated with a DOPC monolayer onto a flat DOPC monolayer, as listed in Figure 1. The DOPC monolayer at the flat air-water interface was doped with low amounts of Rhodamine-DPPE, and this thus made it possible to visualize a background with a bright color and a dark region newly added to the flat interface. There, a recovery process was observed at 23 mN/m of fixed surface pressure. A fit of equation 1 to the change of fractional intensity according to time is shown in Figure 2. The R2 value of this fit is 0.999, and this fit still works well even at lower or higher surface pressures. The diffusion coefficient of the DOPC monolayer obtained from this fit was 27.54 &#181;m2/sec at 23 mN/m of surface pressure.
For further validation of the FRAM, diffusion coefficients of the DOPC and dipalmitoylphosphatidylcholine (DPPC) monolayers were measured according to surface pressure. As shown in Figure 3, FRAM captures the rapid decrease of diffusion coefficient of the DPPC monolayer at ~ 9 mN/m of surface pressure, where a LC (liquid condensed) &#8211; LE (liquid expanded) phase transition occurs10, and the values of diffusion coefficients are also agreed well with the previously measurements by Peters et al.19. In addition, an exponential decay of the diffusion coefficient with the surface pressure was observed in the DOPC monolayer, and this tendency is almost identical with the one measured by Caruso et al.12.
<img alt="Figure 1" src="/files/ftp_upload/53376/53376fig1.jpg" /> 
Figure 1. (A) Schematic illustration of the FRAM (fluorescence recovery after merging) technique. As the lipid monolayer coated droplet is merged to flat air-water interface, the lipid monolayer without fluorescence tagged lipid is inserted into flat fluorescent lipid monolayer. (B) Fluorescence microscope images with time during the recovery process of a DOPC monolayer at 23 mN/m of surface pressure (scale bar = 100 &#181;m). The dark region of the DOPC monolayer is fully recovered by the diffusion process within several minutes. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53376/53376fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53376/53376fig2.jpg" /> 
Figure 2. Fractional intensity vs. time. Black circles and the gray dotted line indicate the values of fractional intensities obtained from image analysis (procedures 4.1 and 4.2) and the fit of equation 1 to the fractional intensities with time, shown in procedure 4.3, respectively. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53376/53376fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53376/53376fig3.jpg" /> 
Figure 3. Diffusion coefficients of DOPC (triangle) and DPPC (empty square) monolayers as a function of surface pressure. The lines with the bright gray color and with the dark gray color indicate the values of the diffusion coefficient previously reported by Peters et al. and Caruso et al., respectively. This figure has been modified from Jeong et al.23. Reprinted with permission from Jeong et al. Langmuir. 30(48), 14369-14374. Copyright 2014 American Chemical Society. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53376/53376fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Fluorescence Recovery after Merging a Droplet to Measure the Two-dimensional Diffusion of a Phospholipid Monolayer at JoVE.com

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

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 ...

fluorescence-recovery-after-merging-droplet-to-measure-two

Universidad de Cartagena UDEC

Research

JoVE Journal

Bioengineering

8.0K Views.  KAIST. 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.

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

A Fluorescence Fluctuation Spectroscopy Assay of Protein-Protein Interactions at Cell-Cell Contacts

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring cells. Of interest, only few assays are capable of specifically probing such interactions directly in living cells. Here, we present an assay to measure the binding of proteins expressed at the surfaces of neighboring cells, at cell-cell contacts. This assay consists of two steps: mixing of cells expressing the proteins of interest fused to different fluorescent proteins, followed by fluorescence fluctuation spectroscopy measurements at cell-cell contacts using a confocal laser scanning microscope. We demonstrate the feasibility of this assay in a biologically relevant context by measuring the interactions of the amyloid precursor-like protein 1 (APLP1) across cell-cell junctions. We provide detailed protocols on the data acquisition using fluorescence-based techniques (scanning fluorescence cross-correlation spectroscopy, cross-correlation number and brightness analysis) and the required instrument calibrations. Further, we discuss critical steps in the data analysis and how to identify and correct external, spurious signal variations, such as those due to photobleaching or cell movement.
In general, the presented assay is applicable to any homo- or heterotypic protein-protein interaction at cell-cell contacts, between cells of the same or different types and can be implemented on a commercial confocal laser scanning microscope. An important requirement is the stability of the system, which needs to be sufficient to probe diffusive dynamics of the proteins of interest over several minutes.

This protocol describes a fluorescence fluctuation spectroscopy-based approach to investigate interactions among proteins mediating cell-cell interactions, i.e. proteins localized in cell junctions, directly in living cells. We provide detailed guidelines on instrument calibration, data acquisition and analysis, including corrections to possible artefact sources.

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring ...

Biochemistry

Lateral Diffusion and Exocytosis of Membrane Proteins in Cultured Neurons Assessed using Fluorescence Recovery and Fluorescence-loss Photobleaching

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This directed trafficking is of fundamental importance to deliver, maintain or remove synaptic proteins.
Super-ecliptic pHluorin (SEP) is a pH-sensitive derivative of eGFP that has been extensively used for live cell imaging of plasma membrane proteins1-2. At low pH, protonation of SEP decreases photon absorption and eliminates fluorescence emission. As most intracellular trafficking events occur in compartments with low pH, where SEP fluorescence is eclipsed, the fluorescence signal from SEP-tagged proteins is predominantly from the plasma membrane where the SEP is exposed to a neutral pH extracellular environment. When illuminated at high intensity SEP, like every fluorescent dye, is irreversibly photodamaged (photobleached)3-5. Importantly, because low pH quenches photon absorption, only surface expressed SEP can be photobleached whereas intracellular SEP is unaffected by the high intensity illumination6-10. 
FRAP (fluorescence recovery after photobleaching) of SEP-tagged proteins is a convenient and powerful technique for assessing protein dynamics at the plasma membrane. When fluorescently tagged proteins are photobleached in a region of interest (ROI) the recovery in fluorescence occurs due to the movement of unbleached SEP-tagged proteins into the bleached region. This can occur via lateral diffusion and/or from exocytosis of non-photobleached receptors supplied either by de novo synthesis or recycling (see Fig. 1). The fraction of immobile and mobile protein can be determined and the mobility and kinetics of the diffusible fraction can be interrogated under basal and stimulated conditions such as agonist application or neuronal activation stimuli such as NMDA or KCl application8,10. 
We describe photobleaching techniques designed to selectively visualize the recovery of fluorescence attributable to exocytosis. Briefly, an ROI is photobleached once as with standard FRAP protocols, followed, after a brief recovery, by repetitive bleaching of the flanking regions. This 'FRAP-FLIP' protocol, developed in our lab, has been used to characterize AMPA receptor trafficking at dendritic spines10, and is applicable to a wide range of trafficking studies to evaluate the intracellular trafficking and exocytosis.

This report describes the use of live cell imaging and photobleach techniques to determine the surface expression, transport pathways and trafficking kinetics of exogenously expressed, pH-sensitive GFP-tagged proteins at the plasma membrane of neurons.

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This ...

Neuroscience

From Fast Fluorescence Imaging to Molecular Diffusion Law on Live Cell Membranes in a Commercial Microscope

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the regulation of many cellular functions. However, due to the fast dynamics and the tiny structures involved, a very high spatio-temporal resolution is required to catch the real behavior of molecules. Here we present the experimental protocol for studying the dynamics of fluorescently-labeled plasma-membrane proteins and lipids in live cells with high spatiotemporal resolution. Notably, this approach doesn&#8217;t need to track each molecule, but it calculates population behavior using all molecules in a given region of the membrane. The starting point is a fast imaging of a given region on the membrane. Afterwards, a complete spatio-temporal autocorrelation function is calculated correlating acquired images at increasing time delays, for example each 2, 3, n repetitions. It is possible to demonstrate that the width of the peak of the spatial autocorrelation function increases at increasing time delay as a function of particle movement due to diffusion. Therefore, fitting of the series of autocorrelation functions enables to extract the actual protein mean square displacement from imaging (iMSD), here presented in the form of apparent diffusivity vs average displacement. This yields a quantitative view of the average dynamics of single molecules with nanometer accuracy. By using a GFP-tagged variant of the Transferrin Receptor (TfR) and an ATTO488 labeled 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphoethanolamine (PPE) it is possible to observe the spatiotemporal regulation of protein and lipid diffusion on &#181;m-sized membrane regions in the micro-to-milli-second time range.

Spatial distribution and temporal dynamics of plasma membrane proteins and lipids is a hot topic in biology. Here this issue is addressed by a spatio-temporal image fluctuation analysis that provides conceptually the same physical quantities of single particle tracking, but it uses small molecular labels and standard microscopy setups.

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the ...

Single Molecule Fluorescence Microscopy on Planar Supported Bilayers

In the course of a single decade single molecule microscopy has changed from being a secluded domain shared merely by physicists with a strong background in optics and laser physics to a discipline that is now enjoying vivid attention by life-scientists of all venues 1. This is because single molecule imaging has the unique potential to reveal protein behavior in situ in living cells and uncover cellular organization with unprecedented resolution below the diffraction limit of visible light 2. Glass-supported planar lipid bilayers (SLBs) are a powerful tool to bring cells otherwise growing in suspension in close enough proximity to the glass slide so that they can be readily imaged in noise-reduced Total Internal Reflection illumination mode 3,4. They are very useful to study the protein dynamics in plasma membrane-associated events as diverse as cell-cell contact formation, endocytosis, exocytosis and immune recognition. Simple procedures are presented how to generate highly mobile protein-functionalized SLBs in a reproducible manner, how to determine protein mobility within and how to measure protein densities with the use of single molecule detection. It is shown how to construct a cost-efficient single molecule microscopy system with TIRF illumination capabilities and how to operate it in the experiment.

Preparation of protein-functionalized planar glass-supported lipid bilayers, determination of protein mobility within and measurement of protein densities is shown here. A roadmap to building a noise-reduced Total Internal Reflection microscope is outlined, which allows visualizing single bilayer-resident fluorochromes with high spatiotemporal resolution.

In the course of a single decade single molecule microscopy has changed from being a secluded domain shared merely by physicists with a strong background ...

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 Diffusion Coefficients via Two-photon Fluorescence Recovery After Photobleaching

Multi-fluorescence recovery after photobleaching is a microscopy technique used to measure the diffusion coefficient (or analogous transport parameters) of macromolecules, and can be applied to both in vitro and in vivo biological systems. Multi-fluorescence recovery after photobleaching is performed by photobleaching a region of interest within a fluorescent sample using an intense laser flash, then attenuating the beam and monitoring the fluorescence as still-fluorescent molecules from outside the region of interest diffuse in to replace the photobleached molecules. We will begin our demonstration by aligning the laser beam through the Pockels Cell (laser modulator) and along the optical path through the laser scan box and objective lens to the sample. For simplicity, we will use a sample of aqueous fluorescent dye. We will then determine the proper experimental parameters for our sample including, monitor and bleaching powers, bleach duration, bin widths (for photon counting), and fluorescence recovery time. Next, we will describe the procedure for taking recovery curves, a process that can be largely automated via LabVIEW (National Instruments, Austin, TX) for enhanced throughput. Finally, the diffusion coefficient is determined by fitting the recovery data to the appropriate mathematical model using a least-squares fitting algorithm, readily programmable using software such as MATLAB (The Mathworks, Natick, MA).

In this article we will describe the procedure for measuring diffusion coefficients using multi-photon fluorescence recovery after photobleaching. We will begin by aligning the laser along the optical path to the sample and determining the proper experimental parameters, then continue generating and finally fitting fluorescence recovery curves.

Multi-fluorescence recovery after photobleaching is a microscopy technique used to measure the diffusion coefficient (or analogous transport parameters) ...

Biology

Ligand Nano-cluster Arrays in a Supported Lipid Bilayer

Currently there is considerable interest in creating ordered arrays of adhesive protein islands in a sea of passivated surface for cell biological studies. In the past years, it has become increasingly clear that living cells respond, not only to the biochemical nature of the molecules presented to them but also to the way these molecules are presented. Creating protein micro-patterns is therefore now standard in many biology laboratories; nano-patterns are also more accessible. However, in the context of cell-cell interactions, there is a need to pattern not only proteins but also lipid bilayers. Such dual proteo-lipidic patterning has so far not been easily accessible. We offer a facile technique to create protein nano-dots supported on glass and propose a method to backfill the inter-dot space with a supported lipid bilayer (SLB). From photo-bleaching of tracer fluorescent lipids included in the SLB, we demonstrate that the bilayer exhibits considerable in-plane fluidity. Functionalizing the protein dots with fluorescent groups allows us to image them and to show that they are ordered in a regular hexagonal lattice. The typical dot size is about 800 nm and the spacing demonstrated here is 2 microns. These substrates are expected to serve as useful platforms for cell adhesion, migration and mechano-sensing studies.

We present a protocol to functionalize glass with nanometric protein patches surrounded by a fluid lipid bilayer. These substrates are compatible with advanced optical microscopy and are expected to serve as platform for cell adhesion and migration studies.

Currently there is considerable interest in creating ordered arrays of adhesive protein islands in a sea of passivated surface for cell biological ...

PIP-on-a-chip: A Label-free Study of Protein-phosphoinositide Interactions

Numerous cellular proteins interact with membrane surfaces to affect essential cellular processes. These interactions can be directed towards a specific lipid component within a membrane, as in the case of phosphoinositides (PIPs), to ensure specific subcellular localization and/or activation. PIPs and cellular PIP-binding domains have been studied extensively to better understand their role in cellular physiology. We applied a pH modulation assay on supported lipid bilayers (SLBs) as a tool to study protein-PIP interactions. In these studies, pH sensitive ortho-Sulforhodamine B conjugated phosphatidylethanolamine is used to detect protein-PIP interactions. Upon binding of a protein to a PIP-containing membrane surface, the interfacial potential is modulated (i.e. change in local pH), shifting the protonation state of the probe. A case study of the successful usage of the pH modulation assay is presented by using phospholipase C delta1 Pleckstrin Homology (PLC-&#948;1 PH) domain and phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2) interaction as an example. The apparent dissociation constant (Kd,app) for this interaction was 0.39 &#177; 0.05 &#181;M, similar to Kd,app values obtained by others. As previously observed, the PLC-&#948;1 PH domain is PI(4,5)P2 specific, shows weaker binding towards phosphatidylinositol 4-phosphate, and no binding to pure phosphatidylcholine SLBs. The PIP-on-a-chip assay is advantageous over traditional PIP-binding assays, including but not limited to low sample volume and no ligand/receptor labeling requirements, the ability to test high- and low-affinity membrane interactions with both small and large molecules, and improved signal to noise ratio. Accordingly, the usage of the PIP-on-a-chip approach will facilitate the elucidation of mechanisms of a wide range of membrane interactions. Furthermore, this method could potentially be used in identifying therapeutics that modulate protein&#39;s capacity to interact with membranes.

Here we present a supported lipid bilayer in the context of a microfluidic platform to study protein-phosphoinositide interactions using a label-free method based on pH modulation.

Numerous cellular proteins interact with membrane surfaces to affect essential cellular processes. These interactions can be directed towards a specific ...

Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence (TIRF) Microscopy

High-resolution imaging techniques have shown that many ion channels are not static, but subject to highly dynamic processes, including the transient association of pore-forming and auxiliary subunits, lateral diffusion, and clustering with other proteins. However, the relationship between lateral diffusion and function is poorly understood. To approach this problem, we describe how lateral mobility and activity of individual channels in supported lipid membranes can be monitored and correlated using total internal reflection fluorescence (TIRF) microscopy. Membranes are fabricated on an ultrathin hydrogel substrate using the droplet interface bilayer (DIB) technique. Compared to other types of model membranes, these membranes have the advantage of being mechanically robust and suitable for highly sensitive analytical techniques. This protocol measures Ca2+ ion flux through single channels by observing the fluorescence emission of a Ca2+-sensitive dye in close proximity to the membrane. In contrast to classical single-molecule tracking approaches, no fluorescent fusion proteins or labels, which can interfere with lateral movement and function in the membrane, are required. Possible changes in ion flux associated with conformational changes of the protein are only due to protein lateral motion in the membrane. Representative results are shown using the mitochondrial protein translocation channel TOM-CC and the bacterial channel OmpF. In contrast to OmpF, the gating of TOM-CC is very sensitive to molecular confinement and the nature of lateral diffusion. Hence, supported droplet-interface bilayers are a powerful tool to characterize the link between lateral diffusion and the function of ion channels.

Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence TIRF Microscopy

This protocol describes how to use TIRF microscopy to track individual ion channels and determine their activity in supported lipid membranes, thereby defining the interplay between lateral membrane movement and channel function. It describes how to prepare the membranes, record the data, and analyze the results.

High-resolution imaging techniques have shown that many ion channels are not static, but subject to highly dynamic processes, including the transient ...

Molecular Diffusion in Plasma Membranes of Primary Lymphocytes Measured by Fluorescence Correlation Spectroscopy

Fluorescence correlation spectroscopy (FCS) is a powerful technique for studying the diffusion of molecules within biological membranes with high spatial and temporal resolution. FCS can quantify the molecular concentration and diffusion coefficient of fluorescently labeled molecules in the cell membrane. This technique has the ability to explore the molecular diffusion characteristics of molecules in the plasma membrane of immune cells in steady state (i.e., without processes affecting the result during the actual measurement time). FCS is suitable for studying the diffusion of proteins that are expressed at levels typical for most endogenous proteins. Here, a straightforward and robust method to determine the diffusion rate of cell membrane proteins on primary lymphocytes is demonstrated. An effective way to perform measurements on antibody-stained live cells and commonly occurring observations after acquisition are described. The recent advancements in the development of photo-stable fluorescent dyes can be utilized by conjugating the antibodies of interest to appropriate dyes that do not bleach extensively during the measurements. Additionally, this allows for the detection of slowly diffusing entities, which is a common feature of proteins expressed in cell membranes. The analysis procedure to extract molecular concentration and diffusion parameters from the generated autocorrelation curves is highlighted. In summary, a basic protocol for FCS measurements is provided; it can be followed by immunologists with an understanding of confocal microscopy but with no other previous experience of techniques for measuring dynamic parameters, such as molecular diffusion rates.

A method to measure protein diffusion in membranes of primary immune cells using fluorescence correlation spectroscopy (FCS) is described. In this paper, the use of antibodies for fluorescent labeling is illustrated.

Fluorescence correlation spectroscopy (FCS) is a powerful technique for studying the diffusion of molecules within biological membranes with high spatial ...