All the confocal microscopy images were obtained using the Nikon A1RHD Multiphoton upright confocal microscope. We acknowledge the guidance and assistance of the support staff, especially Guillermo Marques, at the University Imaging Center at the University of Minnesota. This work was supported by NIH Grant HL51177. SI was supported by a Ruth L. Kirschstein NRSA Institutional Research Training Grant F32 HL151128.

1. Preparation of capillary tubes
<ol>
	<li>Place the capillary into a capillary puller and run the desired pulling program to make two tapered capillaries with an outside diameter (OD) of ~1 &#181;m at the tip. 
	NOTE: The OD of the capillary before pulling must be the OD specified to fit in the capillary holder in the microtensiometer cell. The inner diameter (ID) of the capillary can vary, but will affect the critical radius of the capillary following pulling. A pulling program is chosen so that the resulting taper initially reduces the capillary OD and ID quickly, then reaches a radius near the desired capillary OD and ID, and then reduces in diameter more slowly. This will create a greater capillary length that can be scored to yield a usable capillary of 30-100 &#181;m in ID.</li>
	<li>Score the tip of the capillary at the desired spot to obtain an ID of 30-100 &#181;m and break off the tip. The capillary will now have an OD and ID of the desired radius at the tip (Figure 1A). The capillaries can be stored until step 2. 
	NOTE: The cut edge of the capillary must be a 90&#176; clean break. Any defect in the cut edge will lead to bad pinning of the bubble to the capillary and poor surface property measurements. Tapered capillary tips are very delicate. They will be destroyed if they come into contact with anything other than the solutions (e.g., vial walls, air nozzle).</li>
</ol>
2. Hydrophobization of capillaries
<ol>
	<li>Gather pulled glass capillaries, acid cleaning solution, plastic tweezers, deionized (DI) water, hydrophobization solution (2% silane in ethanol), vacuum pump, and ethanol solution. See Table of Materials for details. 
	CAUTION: Acid cleaning solution is acutly toxic, causes skin and eye corrosion/irritation, is oxidizing. Hydrophobization solution is a skin/eye/respiratory irritant. Wear eye protection, lab coats, and gloves and work with solutions in a fume hood.</li>
	<li>Acid-clean the capillary 
	NOTE: Acid-cleaning the capillary removes any organic residues inside the capillary and prepares the glass surface for the silanization reaction that renders the capillary hydrophobic.
	<ol>
		<li>Grab a capillary firmly near its wide end with the tweezers.</li>
		<li>Dip the tapered tip into the acid cleaning solution while attaching the hose from the vacuum pump to the wide end of the capillary. This will suck the solution into the capillary. 
		NOTE: A pipette tip may be attached to the end of the capillary hose to allow for a better fit with the capillary end.</li>
		<li>Stop when the acid cleaning solution has filled about half of the capillary. 
		NOTE: After removal of the capillary tip from the acid cleaning solution, the solution on the exterior of the capillary often forms a bead near the capillary tip. Gently touch the capillary to the neck of the solution vial to remove excess solution.</li>
		<li>Allow the acid cleaning solution to remain in the capillaries for at least 30 min, ensuring that the plug of the liquid remains at the tapered end of the capillary.</li>
		<li>Remove the acid cleaning solution from the capillary by firmly holding the capillary with the tweezers and using the vacuum hose to pull out the liquid from the large end of the capillary.</li>
	</ol>
	</li>
	<li>Rinse the capillary
	<ol>
		<li>Submerge the tapered end of the capillary into DI water ensuring it is submerged deep enough to cover any exterior that was submerged in the acid cleaning solution. While the tip is submerged, use the vacuum hose to pull DI water through the capillary. Remove the capillary from the water and remove the remaining water with the vacuum hose.</li>
		<li>Repeat the above step at least 4x.</li>
	</ol>
	</li>
	<li>Perform step 2.3 again substituting ethanol for DI water.</li>
	<li>Apply suction continuously until the ethanol completely evaporates from the interior of the capillary. The capillary will become cloudy and cool to touch as the ethanol begins to evaporate but will clear after 30 to 45 s.</li>
	<li>Coat the capillary with the hydrophobization solution
	<ol>
		<li>Briefly dip the wide end of the capillary into the ~2% silane in ethanol solution. Capillary action will cause the coating solution to rise within the capillary. Remove the capillary from the solution once a ~1 cm size plug has risen within the capillary.</li>
		<li>Orient the capillary so that the tapered tip faces downward, allowing the coating solution to fall with gravity toward the tapered tip.</li>
		<li>Allow the coating solution to remain in the capillary for at least 3 min. 
		NOTE: There must be no air bubbles in the plug of the coating solution that is in contact with the interior of the tapered tip. If there is an air bubble, then the capillary interior was likely not sufficiently dried in step 2.5. To remedy this, repeat steps 2.4-2.6 as needed.</li>
	</ol>
	</li>
	<li>Rinse the capillaries with ethanol 1x in the same manner as step 2.3.</li>
	<li>Set the hydrophobic coating on the capillary
	<ol>
		<li>Place clean and dry scintillation vials into a vacuum oven set to 120 &#176;C. Place coated capillaries into the vials (ideally one capillary per vial) with wide ends resting on the base of the vial. Allow capillaries to remain in the oven for at least 6 h (overnight preferred) to achieve permanent bonding of the hydrophobic silane layer to the capillaries. The capillaries can be stored until step 4.</li>
	</ol>
	</li>
</ol>
3. Sample preparation and storage
<ol>
	<li>Mix and store surfactant and fluorophore solutions in clean acid-washed vials to avoid contamination. 
	NOTE: Commercially available lipids must be of the highest purity and stored between use at - 20 &#176;C. Old or contaminated lipids often cause results to be difficult to reproduce.</li>
</ol>
4. Setting up the microtensiometer
<ol>
	<li>Assemble the CPM cell as described in Figure 2.
	<ol>
		<li>Place the large side of the capillary into the top of the CPM cell until it pushes through to the underside of the cell.</li>
		<li>Gently tighten the PEEK plug to secure the capillary, and then attach the tube from the microfluidic pump to the large side of the capillary. Be careful not to touch the tapered capillary tip.</li>
	</ol>
	</li>
	<li>As necessary, attach the reservoir exchange and/or temperature control hoses to the respective inlets and outlets on the CPM cell (Figure 2); otherwise, plug the unused inlets and outlets.</li>
	<li>Attach the CPM cell to the confocal microscope stage, roughly aligning it with the CFM objective, CPM camera, and CPM light source (Figure 3).</li>
	<li>Open the gas flow to the microfluidic pump at the recommended operating pressure of the pump (150 mbar for the microfluidic pump used here) and ensure the flow to the capillary is open.</li>
	<li>Start running the CPM virtual interface (Supplemental Coding File 1: Microtensiometer Virtual Interface.vi) in Pressure Control mode with the capillary pressure oscillation frequency and amplitude set to zero (Figure 4-7). Figure 4 shows a screenshot of the virtual interface. For DI water and a capillary radius of ~35 &#181;m, a pressure of ~20 mbar ensures that no water enters the capillary.</li>
	<li>Fill the CPM cell with water using a pipette.</li>
	<li>Focus on the capillary tip using the microtensiometer camera.</li>
	<li>Focus on the capillary tip with the CFM. If there is difficulty finding the capillary, use the CPM camera to find the CFM objective. This will help approximate the distance between the CFM objective and the bubble, achieving the correct working distance.</li>
	<li>After the annulus (green sector projection) is centered on the bubble, manually adjust the focus so that the bubble edge can be seen clearly (Figure 4-3). 
	NOTE: The position, start and end angle, and inner and outer radii of the annulus can be adjusted via the menu below the view window.</li>
	<li>Click on Reset Bubble, and make sure that a new bubble is formed (one will be able to hear the old bubble pop, and the new bubble will be observable from the control panel viewing window; Figure 4-3). If the bubble does not pop, increase the Reset Pressure or increase the Reset Delay Time in the Bubble Reset tab below the viewing window. Check whether the surface tension is around 73 mN/m (for saline or water/air bubbles) (Figure 4-9).</li>
	<li>Take out the water via the direct-to-cell syringe (Figure 3-13), empty it, and reattach it. The sample is ready for loading to run the experiment.</li>
</ol>
5. Adsorption study
<ol>
	<li>Fill the cell with the desired sample using an autoclaved pipette keeping the CPM software in Pressure Control mode. Make sure the initial surface tension is around 73 mN/m when a new bubble interface is created.</li>
	<li>Determine the radius of the newly formed bubble and input that value into the centerline area control (Figure 4-7) and change the control type to area control by clicking on the Area Control tab (Figure 4-8). 
	NOTE: Constant pressure control can also be used, but this causes the bubble radius to change continuously as the surface tension of the interface changes. This changing area can complicate the analysis of surfactant adsorption rates and cause the bubble to pop during the study.</li>
	<li>Start recording the confocal video.</li>
	<li>Click on Reset Bubble (Figure 4-5), and immediately click on Collect Data&#160;(Figure 4-6). The signaling light on the button will turn green.</li>
	<li>Adjust the data recording rate according to the concentration of the sample by sliding the bar shown in Figure 4-6. For slower adsorptions, use a slower recording rate. This can be adjusted in the middle of a run if a higher recording rate is desired early on, but a slower rate is preferable for long studies in order to reduce file size.</li>
	<li>After the end of the experiment (when a final surface tension plateau has been reached), save the file by choosing the correct file path (Figure 4-1) and clicking on the Save button (Figure 4-2).</li>
	<li>Stop and save the recording on the CFM as well.</li>
</ol>
6. Oscillation/relaxation study
<ol>
	<li>Fill the cell with the sample using an autoclaved pipette keeping the CPM software in Pressure Control mode. Make sure the surface tension is around 73 mN/m when a new bubble interface is created.</li>
	<li>Wait until the sample is fully adsorbed to the interface. This can be performed directly after an adsorption study instead of starting over with a new bubble interface.</li>
	<li>Decide whether oscillation will be a pressure oscillation, area oscillation, or curvature oscillation by selecting the appropriate tab (Figure 4-8) and entering the desired baseline value, oscillation% and oscillation frequency (Figure 4-7). 
	NOTE: Sawtooth, square, and triangular wave area oscillations are also accessible from the drop-down menu in the Other Area Oscillation tab.</li>
	<li>Start the recording of the confocal video and click on Collect Data&#160;(Figure 4-6) on the CPM software.</li>
	<li>Start the oscillation. Be sure to record at least seven cycles for best results. Choose a data acquisition rate (Figure 4-6) to give an adequate number of data points for each oscillation cycle.</li>
	<li>If other oscillation amplitudes or frequencies are desired, change the values during the experiment.</li>
	<li>Save the results as in steps 5.6 and 5.7.</li>
</ol>
7. Solvent exchange study
<ol>
	<li>Fill the cell with the sample using an autoclaved pipette keeping the CPM software in pressure control mode. Make sure the surface tension is around 73 mN/m, when a new bubble interface is created. 
	NOTE: Adsorption and/or oscillation studies can be performed prior to the solvent exchange study.</li>
	<li>Connect the inlet tube with the bottle of desired exchange solution (Figure 3-11) to the peristaltic pump (Figure 3-10).</li>
	<li>Start the recording of the video in confocal software and click on Collect Data (Figure 4-6) on the CPM software.</li>
	<li>Set the peristaltic pump speed. This will control the rate of fluid exchange and must be chosen based on the requirements for the experiment, i.e., how fast the solvent needs to be exchanged.</li>
	<li>If multiple fluids need to be exchanged, stop the peristaltic pump, and connect the inlet to another exchange solution.</li>
	<li>After the exchange has finished (~20 min), save the results as in step 5.6 and 5.7.</li>
</ol>
8. Insoluble surfactant adsorption
NOTE: If the surfactant to be adsorbed is not soluble in the reservoir liquid, this method can be used to transfer a monolayer from the air/water interface of the cell to the bubble surface. Many bilayer forming lipids are almost insoluble in saline solution and do not spontaneously absorb to the bubble when suspended in the reservoir solution.
<ol>
	<li>Fill the cell with the sample using an autoclaved pipette keeping the CPM software in Pressure Control mode. Make sure the surface tension is around 73 mN/m, when a new bubble interface is created.</li>
	<li>Deposit a monolayer of insoluble surfactant on air-water interface of the cell from a solution in a volatile organic solution. Using a syringe, deposit small droplets at the interface and allow the solvent to evaporate leaving the lipid behind as a thin film. 
	CAUTION: Chloroform is used as a solvent for phospholipids such as phosphatidylcholines and fatty acids. Spreading solutions are usually 0.01-0.02 mg of lipid per mL of the solvent. Chloroform is acutely toxic, can cause skin and eye irritation, and is carcinogenic. Wear appropriate eye protection, lab coat, and gloves and make the solution in a fume hood.</li>
	<li>Decrease the surface area via the centerline pressure control (Figure 4-7) of the bubble until it is nearly flat. This prevents the bubble from popping after the surfactant has adsorbed.</li>
	<li>Remove the reservoir liquid from the cell via the direct-to-cell syringe until the air/water interface moves past the tip of the capillary. While a syringe pump can be used, this step can be achieved by manually using the syringe.</li>
	<li>Increase the reservoir liquid height to its initial level. 
	NOTE: After the tip is resubmerged, the bubble will be larger due to the surfactant that is now adsorbed on the interface. The monolayer will now be ready for oscillation or solvent exchange experiments.</li>
</ol>
9. Clean up
<ol>
	<li>Turn off the CFM.</li>
	<li>Change to the Pressure Control mode.</li>
	<li>Remove the sample from the cell using a pipette. Load the cell with DI water and turn up the pressure to ~50 mbar to cause bubbles to constantly escape the capillary and clean the capillary tip. Repeat this process 2x.</li>
	<li>Close the safety valve and turn off the CPM by clicking on the red button in the upper-left corner, turn off the light and blue pressure control panel, and close off the pressure source.</li>
	<li>Remove the cell from the confocal microscope stage. Rinse the cell out with ethanol and DI water. Remove the capillary tube from the CPM cell.</li>
</ol>
10. Cleaning the cell
<ol>
	<li>Disassemble the cell. Brush the inside wall with a toothbrush while rinsing under DI water. Submerge the parts in ethanol and sonicate it for ~30 min.</li>
	<li>Rinse all the parts with DI water a few times. Dry the parts by either blowing them with nitrogen gas or drying them inside a vacuum oven.</li>
</ol>
11. Oscillation analysis
<ol>
	<li>Run the Dilatational_Rheology_Analysis.m code (Supplemental Coding File 2), choosing the desired file saved from the CPM virtual interface. Sample data is included in the supplemental files.</li>
	<li>The pressure vs. time plot will appear as shown in Supplementary Figure 1. Left-click the point where the oscillation starts and left-click again where the oscillation ends. If the data contains multiple oscillations, repeat this process for all oscillations.
	<ol>
		<li>When all start and end points have been left-clicked, right-click the mouse anywhere. For example, as shown in Supplementary Figure 1, one can left click at points 1, 2, 3, and 4, followed by a right-click. 
		NOTE: The code will calculate the dilatational modulus and phase angle and the results will be written to a new .csv file in the original file location. The results for the sample data can be seen in the code results given in the Supplemental Coding File 2. MATLAB will also generate several graphical representations of the data as shown in Supplementary Figure 2.</li>
	</ol>
	</li>
</ol>

The authors have no conflicts of interest to disclose.

The combined CPM/CFM is a powerful tool for examining interfacial dynamics, equilibria, and morphology. This protocol describes the steps necessary for obtaining data with CPM/CFM.
Figure 2 shows the cell design with channels for the capillary, solvent, and heat exchange indicated. The inlet for solvent exchange should be at the bottom of the cell while the outlet should be at the top, allowing for the cell to not overflow during the exchange. In practice, the inl...

Air-water interfaces covered by surfactant films are ubiquitous in daily life. Surfactant-water injections are used to enhance oil recovery from depleted fields and are used as hydraulic fracturing solutions for shale gas and oil. Gas-liquid foams and liquid-liquid emulsions are common to many industrial and scientific processes as lubricants and cleaning agents and are common in food. Surfactants and proteins at interfaces stabilize antibody conformations during packaging, storage, and administration1,2,3,4,5, tear film stability in the eye6,7,8, and pulmonary mechanics9,10,11,12,13,14,15.
The study of surface-active agents or surfactants adsorbing to interfaces and their properties has a long history with many different experimental techniques16,17,18,19,20,21,22,23,24,25,26,27. A recent development is the capillary pressure microtensiometer (CPM), which allows the examination of interfacial properties on highly curved interfaces, at much smaller length scales, while using significantly fewer materials than other common methods9,23,24,25. Confocal fluorescence microscopy (CFM) can be used to study the morphology of lipids and proteins at the air-water interfaces in the CPM22 or on Langmuir troughs20,26,27,28,29. Here a CPM and CFM have been combined to connect morphological phenomena to dynamic and equilibrium interfacial properties to develop structure-function relationships for biological and technological interfaces.
There are numerous parameters of importance in interfacial surfactant systems accessible to the CPM-CFM. In the CPM, a 30-200 &#181;m diameter air bubble is pinned to the tip of a glass capillary tube. In earlier versions of the CPM, the capillary pressure difference between the inside and outside of the bubble was controlled via a water column and oscillatory syringe pump9,30 ; the new version described here replaces these with a higher precision, computer-controlled microfluidic pump. The surface tension (&#947;) is determined via the Laplace equation, &#916;P = 2&#947;/R, from the pressure drop across the interface set by the pump, &#916;P, and optical analysis of the radius of curvature of the bubble, R. The dynamic surface tension of the interface can be determined with 10 ms time resolution following the generation of a new bubble in contact with a bulk liquid containing a soluble surfactant. The surfactant adsorption dynamics can be described by the classic Ward-Tordai equation10,31 to determine essential properties of the surfactant, including the diffusivity, surface coverage, and the relationship between bulk concentration and equilibrium surface tension. Once an equilibrium surface tension is achieved, the interfacial area can be oscillated to measure the dilatational modulus, <img alt="Equation 1" src="/files/ftp_upload/64110/64110eq01.jpg" style="margin: auto;" />, by recording the changes in surface tension, induced by small changes in the bubble surface area, A32. For more complex interfaces that develop their own internal structures such as entangled polymers or proteins, the surface tension, , is replaced by a more general surface stress4,33, <img alt="Equation 2" src="/files/ftp_upload/64110/64110eq02.jpg" style="margin: auto;" />.
Lung stability during breathing may be directly tied to maintaining both a low surface tension and a high dilatational modulus at the alveolar air-liquid interface9,10. All internal lung surfaces are lined with a continuous, microns-thick film of epithelial lining fluid to maintain tissue hydration34. This epithelial lining fluid is primarily water, with salts and various other proteins, enzymes, sugars, and lung surfactant. As is the case for any curved liquid-vapor interface, a capillary pressure is induced with the pressure higher on the inside of the alveolus (or bubble). However, if the surface tension was constant everywhere within the lungs, the Laplace equation, &#916;P = 2&#947;/R, shows that smaller alveoli would have a higher internal pressure relative to larger alveoli, forcing the gas contents of the smaller alveoli to flow to larger, lower pressure alveoli. This is known as &#34;Laplace Instability&#34;9,35. The net result is that the smallest alveoli would collapse and be filled with liquid and become difficult to re-inflate causing part of the lung to collapse, and other parts would over-inflate, both of which are typical symptoms of acute respiratory distress syndrome (ARDS). However, in a properly functioning lung, the surface tension changes dynamically as the air-epithelial fluid interface in the alveolus interfacial area expands and contracts during breathing. If <img alt="Equation 3" src="/files/ftp_upload/64110/64110eq03.jpg" style="margin: auto;" />, or <img alt="Equation 4" src="/files/ftp_upload/64110/64110eq04.jpg" style="margin: auto;" />, the Laplace pressure decreases with decreasing radius and increases with increasing radius so as to eliminate the Laplace instability, thereby stabilizing the lung9. Hence, <img alt="Equation 5" src="/files/ftp_upload/64110/64110eq05.jpg" style="margin: auto;" />, and how it depends on frequency, monolayer morphology and composition, and alveolar fluid composition may be essential for lung stability. The CPM-CFM has also provided the first demonstrations of the effects of interfacial curvature on surfactant adsorption25, monolayer morphology22 and dilatational modulus9. The small volume (~1 mL) of the reservoir in the CPM allows for the quick introduction, removal, or exchange of the liquid phase and minimizes the required quantity of expensive proteins or surfactants10.
Contrast in a CPM-CFM image is due to the distribution of small fractions of fluorescently tagged lipids or proteins at the interface16,27. Two-dimensional surfactant monolayers often exhibit lateral phase separation as a function of surface tension or surface pressure, <img alt="Equation 6" src="/files/ftp_upload/64110/64110eq06.jpg" style="margin: auto;" /> &#960; is the difference between the surface tension of a clean fluid-fluid interface,&#160;&#947;0, and a surfactant-covered interface, &#947;. &#960; can be thought as the 2-D &#34;pressure&#34; caused by the interactions of surfactant molecules at the interface that acts to lower the pure fluid surface tension. At low surface pressures, lipid monolayers are in a liquid-like disorganized state; this is known as the liquid expanded (LE) phase. As the surface pressure increases and the area per lipid molecule decreases, the lipids orient with each other and can undergo a first order phase transition to the long-range ordered liquid condensed (LC) phase16,20,27. The LE and LC phases can coexist at various surface pressures and can be visualized as fluorescently tagged lipids are excluded from the LC phase and segregate to the LE phase. Thus, the LE phase is bright and the LC phase is dark when imaged with CFM16.
The goal of this manuscript is to describe the steps necessary to build and operate the combined confocal microscope microtensiometer. This will allow the reader to perform adsorption studies, measure surface tension, rheological behavior, and examine interfacial morphology simultaneously on a micron-scale air/water or oil/water interface. This includes a discussion of how to pull, cut and hydrophobize the required capillaries, instructions for using pressure, curvature, and surface area control modes, and interfacial transfer of insoluble surfactant to the microtensiometer curved interface.

<table><tbody><tr><td>1.5 O.D. Tygon tubing</td><td>Fischer Scientific</td><td></td><td>Tubing</td></tr><tr><td>A1RHD Multiphoton upright confocal microscope</td><td>Nikon</td><td></td><td>Confocal Microscope</td></tr><tr><td>Acid Cleaning Solution</td><td></td><td></td><td>Sulfuric acid and Alnochromix diluted with water 50% by volume, wait until clear befor diluting</td></tr><tr><td>Alnochromix</td><td>Alconox</td><td>2510</td><td>Mixed with sulfuric acid to package instructionand diluted to make acid cleaning solution</td></tr><tr><td>Ceramic glass cutter</td><td>Sutter Instruments</td><td></td><td></td></tr><tr><td>Chloroform</td><td>Sigma-Aldrich</td><td>650471</td><td>HPLC Plus</td></tr><tr><td>Curosurf</td><td>Chiesi</td><td></td><td>&amp;nbsp;Lung Surfactant</td></tr><tr><td>Di Water</td><td></td><td></td><td>18.5 M&amp;Omega; - cm</td></tr><tr><td>Ethanol</td><td>any</td><td></td><td>200 proof used for hydrophobization, denatured used for cleaning</td></tr><tr><td>Fiber-Lite Model 190 fiber optic illuminator</td><td>Dolan-Jenner Industries Inc.</td><td>281900100</td><td>Light source; other light sources should work as well</td></tr><tr><td>Flow EZ F69 mbar w/Link Module</td><td>Fluigent</td><td>LU-FEZ-0069</td><td>Microfluidic Pump</td></tr><tr><td>Fluigent SDK VIs</td><td>Fluigent</td><td></td><td>Required for CPM virtual Interface</td></tr><tr><td>Fluoroelastomer gaskets</td><td></td><td></td><td>Machined from 1 mm thick Viton sheet, See figure 3</td></tr><tr><td>Gas filter</td><td>Norgren</td><td>F07-100-A3TG</td><td>Put between microfluidic pump and pressure regulator</td></tr><tr><td>Gas regulator</td><td>Norgren</td><td>10R0400R</td><td>Steps down pressure from sorce to range of pump, connected to gas filter range 2-120 psi</td></tr><tr><td>Glass Capilary</td><td>Sutter Instruments</td><td>B150-86-10</td><td>Borosilicate glass O.D. 1.5 mm I.D. 0.86 mm</td></tr><tr><td>Glass Slide</td><td>any</td><td></td><td>75 mm x 25 mm</td></tr><tr><td>Glass Syringe</td><td>Hamilton</td><td>84878</td><td>25 &amp;mu;L glass syringe</td></tr><tr><td>Hydrophobizing Agent</td><td>Sigma-Aldrich</td><td>667420</td><td>1H,1H,2H,2H-Perfluoro-octyltriethoxysilane 98%, other hydrophobic triethoxysilane can be substituted</td></tr><tr><td>Insoluble surfactant</td><td>Avanti</td><td>850355C-200mg</td><td>16:0 DPPC in chloroform</td></tr><tr><td>LabVIEW Software</td><td>National Instruments</td><td></td><td>2017</td></tr><tr><td>Longpass Filter</td><td>ThorLabs</td><td>FEL0650</td><td>650 nm Longpass filter, wavelength must remove excitation lazer frequence</td></tr><tr><td>Lyso-PC</td><td>Avanti</td><td>855675P</td><td>16:0 Lyso PC 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphocholine</td></tr><tr><td>Masterflex L/S variable speed analog consol pump system w/&amp;nbsp; Easy-Load II pump head</td><td>Masterflex</td><td>HV-77916-20</td><td>Peristaltic Pump</td></tr><tr><td>MATLAB</td><td>Mathworks</td><td></td><td>R2019</td></tr><tr><td>Micropipette Puller P-1000</td><td>Sutter Instruments</td><td></td><td>Capillary Puller</td></tr><tr><td>Microtensiometer Cell and Holder</td><td></td><td></td><td>Cell machined from PEEK, holder machined from aluminum, See Figure 3 and 4</td></tr><tr><td>Microtensiometer Objective</td><td>Nikon</td><td></td><td>Fluor 20x/0.50W DIC M/N2 &amp;infin;/0 WD 2.0 mm</td></tr><tr><td>NI Vision Development Module</td><td>National Instruments</td><td></td><td>Required for CPM virtual Interface</td></tr><tr><td>PEEK finger tight fittings</td><td>IDEX</td><td>F-120x</td><td>10-32 Coned Ports</td></tr><tr><td>PEEK plug</td><td>IDEX</td><td>P-551</td><td>10-31 Coned Ports</td></tr><tr><td>pippette tips</td><td>Eppendorf</td><td>22492225</td><td>100 &amp;mu;L - 1000 &amp;mu;L, Autoclaved</td></tr><tr><td>Plastic Forceps</td><td>Thermo Scientific</td><td>6320-0010</td><td></td></tr><tr><td>Plastic Syringe</td><td>Fischer Scientific</td><td>14-955-459</td><td>10 mL</td></tr><tr><td>Plumbing parts</td><td>Fischer Scientific</td><td></td><td>3-way valves and other plumbing parts to connect tubing.</td></tr><tr><td>Research Plus 1-channel 100 &amp;mu;L&amp;ndash;1000 &amp;mu;L</td><td>Eppendorf</td><td>3123000063</td><td>Micro pipetter</td></tr><tr><td>Sulfuric Acid</td><td>any</td><td></td><td>Used for acid cleaning solution</td></tr><tr><td>T Plan SLWD 20x/0.30 OFN25 WD 30 mm</td><td>Nikon</td><td></td><td>Confocal Microscope Objective</td></tr><tr><td>Texas Red DHPE triethylammonim salt</td><td>Thermo Fischer Scientific</td><td>1395MP</td><td>Fluorophore</td></tr><tr><td>Vaccum Pump</td><td>Gast</td><td>DOA-P704-AA</td><td></td></tr></tbody></table>

microtensiometer for confocal microscopy visualization of dynamic interfaces

Adsorption of surface-active molecules to fluid-fluid interfaces is ubiquitous in nature. Characterizing these interfaces requires measuring surfactant adsorption rates, evaluating equilibrium surface tensions as a function of bulk surfactant concentration, and relating how surface tension changes with changes in the interfacial area following equilibration. Simultaneous visualization of the interface using fluorescence imaging with a high-speed confocal microscope allows the direct evaluation of structure-function relationships. In the capillary pressure microtensiometer (CPM), a hemispherical air bubble is pinned at the end of the capillary in a 1 mL volume liquid reservoir. The capillary pressure across the bubble interface is controlled via a commercial microfluidic flow controller that allows for model-based pressure, bubble curvature, or bubble area control based on the Laplace equation. Compared to previous techniques such as the Langmuir trough and pendant drop, the measurement and control precision and response time are greatly enhanced; capillary pressure variations can be applied and controlled in milliseconds. The dynamic response of the bubble interface is visualized via a second optical lens as the bubble expands and contracts. The bubble contour is fit to a circular profile to determine the bubble curvature radius, R, as well as any deviations from circularity that would invalidate the results. The Laplace equation is used to determine the dynamic surface tension of the interface. Following equilibration, small pressure oscillations can be imposed by the computer-controlled microfluidic pump to oscillate the bubble radius (frequencies of 0.001-100 cycles/min) to determine the dilatational modulus The overall dimensions of the system are sufficiently small that the microtensiometer fits under the lens of a high-speed confocal microscope allowing fluorescently tagged chemical species to be quantitatively tracked with submicron lateral resolution.

A major source of measurement error arises from the capillaries that have defects either from the cutting process (Figure 5A,B) or the coating process (Figure 5D). Both types of defects lead to errors in determining the bubble shape and size by the optical image analysis system, leading to inaccurate surface tension values. It is important to carefully examine each new capillary after it is pulled and coated under the optical microscope before i...

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Microtensiometer for Confocal Microscopy Visualization of Dynamic Interfaces

This manuscript describes the design and operation of a microtensiometer/confocal microscope to do simultaneous measurements of interfacial tension and surface dilatational rheology while visualizing the interfacial morphology. This provides the real-time construction of structure-property relationships of interfaces important in technology and physiology.

Adsorption of surface-active molecules to fluid-fluid interfaces is ubiquitous in nature. Characterizing these interfaces requires measuring surfactant ...

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2.3K Views.  University of Minnesota. De methode combineert een confocale microscoop met een capillaire druk micro-tensiometer en creëert een krachtig hulpmiddel dat kan worden gebruikt om curvevloeistofinterfaces met een hoge ruimtelijke en temporele resolutie te bestuderen. Deze techniek kan structuur- en functierelaties voor oppervlakteactieve materialen onderzoeken door gelijktijdige metingen van oppervlakte-eigenschappen en confocale beelden van oppervlaktemorfologieën op sterk gekromde interfaces uit te voeren. We veronde...

Video: Microtensiometer for Confocal Microscopy Visualization of Dynamic Interfaces

Visualization of Cortex Organization and Dynamics in Microorganisms, using Total Internal Reflection Fluorescence Microscopy

TIRF microscopy has emerged as a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules in vitro and in living cells1. The optical phenomenon of total internal reflection occurs when light passes from a medium with high refractive index into a medium with low refractive index at an angle larger than a characteristic critical angle (i.e. closer to being parallel with the boundary). Although all light is reflected back under such conditions, an evanescent wave is created that propagates across and along the boundary, which decays exponentially with distance, and only penetrates sample areas that are 100-200 nm near the interface2. In addition to providing superior axial resolution, the reduced excitation of out of focus fluorophores creates a very high signal to noise ratios and minimizes damaging effects of photobleaching2,3. Being a widefield technique, TIRF also allows faster image acquisition than most scanning based confocal setups.
At first glance, the low penetration depth of TIRF seems to be incompatible with imaging of bacterial and fungal cells, which are often surrounded by thick cell walls. On the contrary, we have found that the cell walls of yeast and bacterial cells actually improve the usability of TIRF and increase the range of observable structures4-8. Many cellular processes can therefore be directly accessed by TIRF in small, single-cell microorganisms, which often offer powerful genetic manipulation techniques. This allows us to perform in vivo biochemistry experiments, where kinetics of protein interactions and activities can be directly assessed in living cells.
We describe here the individual steps required to obtain high quality TIRF images for Saccharomyces cerevisiae or Bacillus subtilis cells. We point out various problems that can affect TIRF visualization of fluorescent probes in cells and illustrate the procedure with several application examples. Finally, we demonstrate how TIRF images can be further improved using established image restoration techniques.

Total Internal Reflection Fluorescence (TIRF) microscopy is a powerful approach to observe structures close to the cell surface at high contrast and temporal resolution. We demonstrate how TIRF can be employed to study protein dynamics at the cortex of cell wall-enclosed bacterial and fungal cells.

TIRF microscopy has emerged as  a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules  in vitro and in living cells1. ...

Bioengineering

A TIRF Microscopy Technique for Real-time, Simultaneous Imaging of the TCR and its Associated Signaling Proteins

Signaling is initiated through the T Cell Receptor (TCR) when it is engaged by antigenic peptide fragments bound by Major Histocompatibility Complex (pMHC) proteins expressed on the surface of antigen presenting cells (APCs). The TCR complex is composed of the ligand binding TCR&alpha;&beta; heterodimer that associates non-covalently with CD3 dimers (the &#949;&delta; and &#949;&gamma; heterodimers and the &zeta;&zeta; homodimer)1. Upon engagement of the receptor, the CD3 &zeta; chains are phosphorylated by the Src family kinase, Lck. This leads to the recruitment of the Syk family kinase, Zap70, which is then phosphorylated and activated by Lck. After that, Zap70 phosphorylates the adapter proteins LAT and SLP76, initiating the formation of the proximal signaling complex containing a large number of different signaling molecules2. 
The formation of this complex eventually results in calcium and Ras-dependent transcription factor activation and the consequent initiation of a complex series of gene expression programs that give rise to T cell differentiation2. TCR signals (and the resulting state of differentiation) are modulated by many other factors, including antigen potency and crosstalk with co-stimulatory/co-inhibitory, chemokine, and cytokine receptors 3-4. Studying the spatial and temporal organization of the proximal signaling complex under various stimulation conditions is, therefore, key to understanding the TCR signaling pathway as well as its regulation by other signaling pathways.
One very useful model system to study signaling initiated by the TCR at the plasma membrane in T cells is glass-supported lipid bilayers, as described previously5-6. They can be utilized to present antigenic pMHC complexes, adhesion, and co-stimulatory molecules to T cells-serving as artificial APCs. By imaging the T cells interacting with the lipid bilayer using total internal reflection fluorescence microscopy (TIRFM), we can restrict the excitation to within 100 nm of the space between the glass and the cell surface 7-8. This allows us to image primarily the signaling events occurring at the plasma membrane. As we are interested in imaging the recruitment of signaling proteins to the TCR complex, we describe a two-camera TIRF imaging system wherein the TCR, labeled with fluorescent Fab (fragment antigen binding) fragments of the H57 antibody (purified from hybridoma H57-597, ATCC, ATCC Number:HB-218) which is specific for TCR&beta;, and signaling proteins, tagged with GFP, may be imaged simultaneously and in real time. This strategy is necessary due to the highly dynamic nature of both the T cells and of the signaling events that are occurring at the TCR. This imaging modality has allowed researchers to image single ligands 9-11 as well as recruitment of signaling molecules to activated receptors and is an excellent system to study biochemistry in-situ12-16.

The compartmentalization of proteins either within the plasma membrane or into intracellular locations is one regulatory mechanism that can greatly influence signaling outcomes; hence, to understand signaling it is important to study the spatial and temporal behavior of the proteins involved. We describe here a TIRF microscopy based system to study signal transduction in T cells, but is broadly applicable.

Signaling is initiated through the T Cell Receptor (TCR) when it is engaged by antigenic peptide fragments bound by Major Histocompatibility Complex ...

Immunology and Infection

Simultaneous Visualization of the Dynamics of Crosslinked and Single Microtubules In Vitro by TIRF Microscopy

Microtubules are polymers of &#945;&#946;-tubulin heterodimers that organize into distinct structures in cells. Microtubule-based architectures and networks often contain subsets of microtubule arrays that differ in their dynamic properties. For example, in dividing cells, stable bundles of crosslinked microtubules coexist in close proximity to dynamic non-crosslinked microtubules. TIRF-microscopy-based in vitro reconstitution studies enable the simultaneous visualization of the dynamics of these different microtubule arrays. In this assay, an imaging chamber is assembled with surface-immobilized microtubules, which are either present as single filaments or organized into crosslinked&#160;bundles. Introduction of tubulin, nucleotides, and protein regulators allows direct visualization of associated proteins and of dynamic properties of single and crosslinked microtubules. Furthermore, changes that occur as dynamic single microtubules organize into bundles can be monitored in real-time. The method described here allows for a systematic evaluation of the activity and localization of individual proteins, as well as synergistic effects of protein regulators on two different microtubule subsets under identical experimental conditions, thereby providing mechanistic insights that are inaccessible by other methods.

Simultaneous Visualization of the Dynamics of Crosslinked and Single Microtubules In Vitro by TIRF Microscopy

Here, a TIRF microscopy-based in vitro reconstitution assay is presented to simultaneously quantify and compare the dynamics of two microtubule populations. A method is described to simultaneously view the collective activity of multiple microtubule-associated proteins on crosslinked microtubule bundles and single microtubules.

Microtubules are polymers of &#945;&#946;-tubulin heterodimers that organize into distinct structures in cells. Microtubule-based architectures and ...

Biochemistry

Simultaneous Interference Reflection and Total Internal Reflection Fluorescence Microscopy for Imaging Dynamic Microtubules and Associated Proteins

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. Total-internal-reflection-fluorescence (TIRF) microscopy has a high signal-to-background ratio, but it suffers from photobleaching and photodamage of the fluorescent proteins. Label-free techniques such as interference reflection microscopy (IRM) and interferometric scattering microscopy (iSCAT) circumvent the problem of photobleaching but cannot readily visualize single molecules. This paper presents a protocol for combining IRM with a commercial TIRF microscope for the simultaneous imaging of microtubule-associated proteins (MAPs) and dynamic microtubules in vitro. This protocol allows for high-speed observation of MAPs interacting with dynamic microtubules. This improves on existing two-color TIRF setups by eliminating both the need for microtubule labeling and the need for several additional optical components, such as a second excitation laser. Both channels are imaged on the same camera chip to avoid image registration and frame synchronization problems. This setup is demonstrated by visualizing single kinesin molecules walking on dynamic microtubules.

We present a protocol for implementing interference-reflection microscopy and total-internal-reflection-fluorescence microscopy for the simultaneous imaging of dynamic microtubules and fluorescently labeled microtubule-associated proteins.

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. ...

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence (TIRF) Microscopy

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and regulated by different suites of binding proteins unique for each polymer. Many studies now demonstrate that the dynamics of both cytoskeletal polymers are intertwined and that this crosstalk is required for most cellular behaviors. A number of proteins involved in actin-microtubule interactions have already been identified (i.e., Tau, MACF, GAS, formins, and more) and are well characterized with regard to either actin or microtubules alone. However, relatively few studies showed assays of actin-microtubule coordination with dynamic versions of both polymers. This may occlude emergent linking mechanisms between actin and microtubules. Here, a total internal reflection fluorescence (TIRF) microscopy-based in vitro reconstitution technique permits the visualization of both actin and microtubule dynamics from the one biochemical reaction. This technique preserves the polymerization dynamics of either actin filament or microtubules individually or in the presence of the other polymer. Commercially available Tau protein is used to demonstrate how actin-microtubule behaviors change in the presence of a classic cytoskeletal crosslinking protein. This method can provide reliable functional and mechanistic insights into how individual regulatory proteins coordinate actin-microtubule dynamics at a resolution of single filaments or higher-order complexes.

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence TIRF Microscopy

This protocol is a guide for visualizing dynamic actin and microtubules using an in vitro total internal fluorescence (TIRF) microscopy assay.

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and ...

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue morphogenesis goes in hand with molecular and structural changes at the single cell level that result in variations of subcellular mechanical properties. As a consequence, even within the same cell, different organelles and compartments resist differently to mechanical stresses; and mechanotransduction pathways can actively regulate their mechanical properties. The ability of a cell to adapt to the microenvironment of the tissue niche thus is in part due to the ability to sense and respond to mechanical stresses. We recently proposed a new mechanosensation paradigm in which nuclear deformation and positioning enables a cell to gauge the physical 3D environment and endows the cell with a sense of proprioception to decode changes in cell shape. In this article, we describe a new method to measure the forces and material properties that shape the cell nucleus inside living cells, exemplified on adherent cells and mechanically confined cells. The measurements can be performed non-invasively with optical traps inside cells, and the forces are directly accessible through calibration-free detection of light momentum. This allows measuring the mechanics of the nucleus independently from cell surface deformations and allowing dissection of exteroceptive and interoceptive mechanotransduction pathways. Importantly, the trapping experiment can be combined with optical microscopy to investigate the cellular response and subcellular dynamics using fluorescence imaging of the cytoskeleton, calcium ions, or nuclear morphology. The presented method is straightforward to apply, compatible with commercial solutions for force measurements, and can easily be extended to investigate the mechanics of other subcellular compartments, e.g., mitochondria, stress-fibers, and endosomes.

Here, we present a protocol to investigate the intracellular mechanical properties of isolated embryonic zebrafish cells in three-dimensional confinement with direct force measurement by an optical trap.

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue ...

Developmental Biology

Imaging Integrin Tension and Cellular Force at Submicron Resolution with an Integrative Tension Sensor

Molecular tension transmitted by integrin-ligand bonds is the fundamental mechanical signal in the integrin pathway that plays significant roles in many cell functions and behaviors. To calibrate and image integrin tension with high force sensitivity and spatial resolution, we developed an integrative tension sensor (ITS), a DNA-based fluorescent tension sensor. The ITS is activated to fluoresce if sustaining a molecular tension, thus converting force to fluorescent signal at the molecular level. The tension threshold for ITS activation is tunable in the range of 10&#8211;60 pN that well covers the dynamic range of integrin tension in cells. On a substrate grafted with an ITS, the integrin tension of adherent cells is visualized by fluorescence and imaged at submicron resolution. The ITS is also compatible with cell structural imaging in both live cells and fixed cells. The ITS has been successfully applied to the study of platelet contraction and cell migration. This paper details the procedure for the synthesis and application of the ITS in the study of integrin-transmitted cellular force.

Integrin tension plays important roles in various cell functions. With an integrative tension sensor, integrin tension is calibrated with picoNewton (pN) sensitivity and imaged at submicron resolution.

Molecular tension transmitted by integrin-ligand bonds is the fundamental mechanical signal in the integrin pathway that plays significant roles in many ...

Control of Cell Adhesion using Hydrogel Patterning Techniques for Applications in Traction Force Microscopy

Traction force microscopy (TFM) is the main method used in mechanobiology to measure cell forces. Commonly this is being used for cells adhering to flat soft substrates that deform under cell traction (2D-TFM). TFM relies on the use of linear elastic materials, such as polydimethylsiloxane (PDMS) or polyacrylamide (PA). For 2D-TFM on PA, the difficulty in achieving high throughput results mainly from the large variability of cell shapes and tractions, calling for standardization. We present a protocol to rapidly and efficiently fabricate micropatterned PA hydrogels for 2D-TFM studies. The micropatterns are first created by maskless photolithography using near-UV light where extracellular matrix proteins bind only to the micropatterned regions, while the rest of the surface remains non-adhesive for cells. The micropatterning of extracellular matrix proteins is due to the presence of active aldehyde groups, resulting in adhesive regions of different shapes to accommodate either single cells or groups of cells. For TFM measurements, we use PA hydrogels of different elasticity by varying the amounts of acrylamide and bis-acrylamide and tracking the displacement of embedded fluorescent beads to reconstruct cell traction fields with regularized Fourier Transform Traction Cytometry (FTTC).
To further achieve precise recording of cell forces, we describe the use of a controlled dose of patterned light to release cell tractions in defined regions for single cells or groups of cells. We call this method local UV illumination traction force microscopy (LUVI-TFM). With enzymatic treatment, all cells are detached from the sample simultaneously, whereas with LUVI-TFM traction forces of cells in different regions of the sample can be recorded in sequence. We demonstrate the applicability of this protocol (i) to study cell traction forces as a function of controlled adhesion to the substrate, and (ii) to achieve a greater number of experimental observations from the same sample.

Near-UV lithography and traction force microscopy are combined to measure cellular forces on micropatterned hydrogels. The targeted light-induced release of single cells enables a high number of measurements on the same sample.

Traction force microscopy (TFM) is the main method used in mechanobiology to measure cell forces. Commonly this is being used for cells adhering to flat ...

Tension Gauge Tether Probes for Quantifying Growth Factor Mediated Integrin Mechanics and Adhesion

Multicellular organisms rely on interactions between membrane receptors and cognate ligands in the surrounding extracellular matrix (ECM) to orchestrate multiple functions, including adhesion, proliferation, migration, and differentiation. Mechanical forces can be transmitted from the cell via the adhesion receptor integrin to ligands in the ECM. The amount and spatial organization of these cell-generated forces can be modulated by growth factor receptors, including epidermal growth factor receptor (EGFR). The tools currently available to quantify crosstalk-mediated changes in cell mechanics and relate them to&#160;focal adhesions, cellular morphology, and signaling are limited. DNA-based molecular force sensors known as tension gauge tethers (TGTs) have been employed to quantify these changes. TGT probes are unique in their ability to both modulate the underlying force threshold and report piconewton scale receptor forces across the entire adherent cell surface at diffraction-limited spatial resolution. The TGT probes used here rely on the irreversible dissociation of a DNA duplex by receptor-ligand forces that generate a fluorescent signal. This allows quantification of the cumulative integrin tension (force history) of the cell. This article describes a protocol employing TGTs to study the impact of EGFR on integrin mechanics and adhesion formation. The assembly of&#160;the TGT mechanical sensing platform is systematically detailed and the procedure to image forces, focal adhesions, and cell spreading is outlined. Overall, the ability to modulate the underlying force threshold of the probe, the adhesion ligand, and the type and concentration of growth factor employed for stimulation make this a robust platform for studying the interplay of diverse membrane receptors in regulating integrin-mediated forces.

TGT surface is an innovative platform to study growth factor-integrin crosstalk. The flexible probe design, specificity of the adhesion ligand, and precise modulation of stimulation conditions allow robust quantitative assessments of EGFR-integrin interplay. The results highlight EGFR as a 'mechano-organizer' tuning integrin mechanics, influencing focal adhesion assembly and cell spreading.

Multicellular organisms rely on interactions between membrane receptors and cognate ligands in the surrounding extracellular matrix (ECM) to orchestrate ...

Biology

Interference Reflection Microscopy for Label-Free Visualization of Microtubule Dynamics

Source: Mahamdeh, M., et al. <a href="https://app.jove.com/v/59520">Implementation of Interference Reflection Microscopy for Label-free, High-speed Imaging of Microtubules.</a> J. Vis. Exp. (2019).
In this video, we describe the interference reflection microscopy (IRM) technique to visualize microtubules growing on a coverslip surface in the presence of a suitable buffer. Upon illumination with incident light, the coverslip-buffer interface and buffer-microtubule interface reflect light, which combines to create an interference pattern, enabling the visualization of microtubules as high-contrast images against a bright background.

Source: Mahamdeh, M., et al. Implementation of Interference Reflection Microscopy for Label-free, High-speed Imaging of Microtubules. J. Vis. Exp. 
...

Microtensiometer for Confocal Microscopy Visualization of Dynamic Interfaces

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