We thank Barbara Windschiegl for her help in establishing SOPs; Dennis Remme for his work on the NanoCalcFX software and Alina Kollmannsperger, Markus Braner and Milan Gerovac for helpful suggestions on the manuscript. The German-Israeli Project Cooperation (DIP) provided by the DFG and the Federal Ministry of Education and Research to R.T., as well as the Federal Ministry of Economics and Technology (ZIM R&amp;D Project) to R.T. and Nanospot GmbH supported this work.

1. Preparation of Large Unilamellar Vesicles (LUVs)
<ol>
	<li>Clean a round bottom flask (10 ml volume) with nitrogen gas to remove any dust particles. Rinse the round bottom flask with ethanol. 
	Note: Residual ethanol can be tolerated and does not disturb subsequent steps.</li>
	<li>Wash a 1 ml glass syringe 4x with chloroform, to remove any contamination. Add 4 ml SoyPC20 (25 mg/ml in chloroform) to the round bottom flask and dope the lipid solution with an additional DOPEATTO390 to a final concentration of 0.1 mol%. 
	Note: Instead of DOPEATTO390 other fluorophore-labeled lipids or lipophilic dyes can also be used, depending on available excitation sources.</li>
	<li>Connect the round bottom flask to a rotary evaporator. Dry the lipid film for 40 min at 300 mbar, 30 &#176;C water bath temperature and maximal rotation speed, to form a homogenous lipid film on the glass wall of the flask.</li>
	<li>Transfer the flask to a high vacuum pump and dry the lipid film for additional 30 min at room temperature.</li>
	<li>Add 5 ml buffer (20 mM Tris, 150 mM NaCl, pH 8.0; sterile filtered) and 8 glass beads (2.7-mm diameter, ethanol washed and dried) to the flask. Connect the flask to the rotary evaporator and rotate without vacuum at 50 &#176;C at maximum speed. 
	Note: Glass beads mechanically displace the lipid film from the glass wall, leading to (multilamellar) vesicle formation. After 45 min rotation the solution appears milky and no residual lipid film should be visible at the flask walls. Instead of glass beads other methods like sonicating the flask in a water bath sonifier, vortexing the flask or the gentle rehydration method are also applicable.</li>
	<li>Transfer the flask under inert gas (argon) to an ultrasonic bath and sonicate for 4 min at room temperature. 
	Note: The ultrasound shockwaves disrupt the liposomes, making them unilamellar.</li>
	<li>Split the LUV solution into 1 ml aliquots.</li>
	<li>Perform 5 freeze/thaw cycles by freezing the LUV solution in liquid nitrogen, followed by thawing at 40 &#176;C in a water bath. 
	Note: This leads to rupture and spontaneous rearrangement of individual liposomes with each other, leading to size increase.</li>
	<li>At the last freezing step store all samples at -80 &#176;C. 
	Note: The LUVs will be stable for at least 6 months.</li>
</ol>
2. Protein Reconstitution
<ol>
	<li>Thaw 1 ml LUV aliquot on ice. Directly before reconstitution, extrude liposomes 17x through a 400 nm polycarbonate filter membrane using a mini extruder. 
	Note: Lipid aggregates will be removed and the final homogenous size distribution is reached.</li>
	<li>Destabilize LUVs by addition of 4% Triton X-100 (v/v) to a final concentration of 0.25% (v/v) and incubate for 5 min at 50 &#176;C. 
	Note: The amount of liposomes used in this step depends on the mass of the purified MscLG22C used (see step 2.3). Purify MscLG22C (derived from E. coli) as described in detail in 28.</li>
	<li>Mix purified MscLG22C and detergent-saturated liposomes at 1:20 (w/w) ratio and incubate for additional 30 min at 50 &#176;C.</li>
	<li>For detergent removal add bio-beads in Tris buffer (20 mM Tris, 150 mM NaCl, pH 8.0; sterile filtered) to the protein/liposome solution, with 16 mg (wet weight) bio-beads per &#181;l detergent used to destabilize liposomes in step 2.2.</li>
	<li>Perform detergent removal overnight (16 hr) at 4 &#176;C on a rotating plate. 
	Note: The constant mixing of the solution ensures optimal detergent removal.</li>
	<li>After detergent removal store proteoliposomes at 4 &#176;C and use for experiments on the same day.</li>
</ol>
3. Activity Assay
<ol>
	<li>During protein reconstitution take a 100 &#181;l aliquot of detergent-destabilized proteoliposome solution after finishing step 2.3.</li>
	<li>Add 1 volume of calcein (30 mM) to the protein-liposome solution and incubate additional 5 min at 50 &#176;C.</li>
	<li>Remove detergent from the solution as described in step 2.4 - 2.6.</li>
	<li>After detergent removal equilibrate a size-exclusion column (20 ml bed volume or more) with Tris buffer (3x the bed volume). 
	Note: Proteoliposome separation can be performed at room temperature, however keeping the sample constantly at 4 &#176;C is advised to ensure best protein activity after fractionation.</li>
	<li>Separate proteoliposomes from free calcein by applying the entire aliquot (200 &#181;l) to the size exclusion column. Allow the mixture to soak into the column, followed by elution of the proteoliposomes from the column via continuous buffer application on top using gravity flow. Observe the calcein containing proteoliposomes as a discrete orange band in the elution front. Collect fractions of 500 &#181;l respectively. 
	Note: Free calcein dye will elute after the proteoliposome fraction with complete separation.</li>
	<li>Pipette 80 &#181;l of each fraction onto a micro-well plate for fluorescent read-out and identify the fraction containing the highest amount of calcein containing proteoliposomes via fluorescence emission. Detect fluorescence at 520 nm with excitation at 490 nm. Use the proteoliposome fraction with the highest signal for the following activity assay.</li>
	<li>Mix 20 &#181;l of the proteoliposome-containing fraction with 980 &#181;l Tris buffer in a 1 ml fluorescence cuvette.</li>
	<li>Measure fluorescence emission at 520 nm (excitation 490 nm) using a spectrofluorometer. Record for 1 min and then add 25 &#181;l of MTSET ([2-(trimethylammonium)ethyl]methanethiosulfonate) from a stock solution (160 mM) and mix well. Keep on recording during mixing. Always prepare the stock solution fresh directly before use. 
	Note: Once solved in buffer MTSET hydrolyzes within 30 min and will be non-reactive. MTSET can be weighed and kept as dry powder aliquot at -20 &#176;C until use. 
	Note: Once activated the MscLG22C channel opens and releases the entrapped calcein, leading to a local concentration decrease of the calcein when effluxing the proteoliposomes and subsequent dequenching of the dye, resulting in an increase of the fluorescence emission.</li>
	<li>After the fluorescence emission has reached a stable plateau again, solubilize the liposomes by addition of 50 &#181;l Triton X-100 (4% v/v). 
	Note: Ideally no further fluorescence increase can be observed, implying active protein in every liposome.</li>
</ol>
4. Size Distribution Measurement of LUVs Using Nano-particle Tracking
<ol>
	<li>Note that only minimal samples of the liposome or proteoliposome solution are necessary for size distribution analysis using the nanoparticle tracker. Dilute a liposome solution of 5 mg/ml lipid concentration 1:5,000 (v/v) in buffer (20 mM Tris, 150 mM NaCl, pH 8.0; sterile filtered) for analysis to a final volume of 1 ml. Filter all buffers used for dilution and prior LUV preparation and reconstitution using a 0.2 &#181;m membrane to remove any suspended particles like dust that would disturb size distribution measurements.</li>
	<li>Wash the flow-chamber with ultra-pure water.</li>
	<li>Use the surface markers to focus on the light beam. Inject approximately 300 &#181;l of the diluted liposome sample and immediately close the exit valve to stop the flow inside the flow chamber, without introducing any air bubbles.</li>
	<li>Adjust focus and camera shutter/gain settings to ensure adequate scattering signal of the sample for detection. 
	Note: 20 - 80 signals should be clearly visible in the field of view. Prevent overcrowding (&#62; 80 signals) as the software will not be able to track individual signals when overlapping.</li>
	<li>Go to the &#34;Capture&#34; screen. Adjust the temperature control to 25 &#176;C and the capture duration to 90 sec using the software controls. Press &#34;Record&#34; to start recording a time series. 
	Note: Upon completion of the time series a new window opens.</li>
	<li>Save the time series in the given format (*.avi). For practical reasons during batch analysis save all files in a single folder.</li>
	<li>Open the exit valve and inject another 300 &#181;l into the flow chamber. Close the valve and repeat steps 4.3 - 4.6 twice.</li>
	<li>After completing all sample runs wash the flow chamber with 5 ml ultra-pure water. Go to the &#34;Pre-Process&#34; screen. Set the calibration parameters temperature = 25 &#176;C and viscosity = 0.91 for the Tris buffer used in this protocol.</li>
	<li>Open the batch process window and load the recorded time series (Advanced/Batch Process/Set File 1-3). Press &#34;GO&#34; to start the size distribution analysis. 
	Note: the analytical software automatically tracks the single particles and calculates their diameter based on their movement behavior using the parameters set in step 4.8. 
	Note: The resulting size distribution analysis saves automatically into the same folder as the time series files.</li>
	<li>Additionally create a report file to summarize the results (Export/Create Report PDF).</li>
</ol>
5. Chip Holder Preparation
<ol>
	<li>Weigh 1.0 g of MDX4 medical grade elastomer base and 0.1 g MDX4 curing agent into a 50 ml reaction tube. Blend both thoroughly with a glass bar.</li>
	<li>Put the tube into a desiccator for 1 hr to degas the adhesive. Afterwards use it within 30 min for chip gluing, as the adhesive will harden and be too hard to work with.</li>
	<li>During elastomer degassing (step 5.1) clean an objective glass slide thoroughly with chloroform to remove any contaminations before chip bonding. Rinse the slide with 5 ml of chloroform using a glass syringe. Collect the chloroform in an underneath positioned beaker. After washing let the slide dry. 
	Note: Perform this step under a fume hood. Alternative to chloroform, other organic solvents like acetone can also be used.</li>
	<li>Deposit a small amount of the adhesive bonding material onto the cover glass using a pipette tip (crystal tips for 10 &#181;l pipettes). Remember that the chips have to fit into the 8-chamber slide holder later on and have to be placed on the slide accordingly.</li>
	<li>Carefully remove one chip at a time from the wafer board using fine tip tweezers and deposit the chip on the drop of adhesive on the cover glass. Ensure that the coated side of the chip faces upwards.</li>
	<li>Gently push the chip onto the cover glass with the back of a blunt PE tweezers. To ensure that the glue is dispensed evenly underneath the chip, carefully slide the chip forth and back. CRITICAL STEP: Ensure that no air bubbles are trapped underneath the chip, as it will disturb optical imaging of the chip cavities later on. Also ensure that no adhesive material contaminates the chip surface.</li>
	<li>Desiccate the finished cover glass for 2 hr at 65 &#176;C in a cabinet dryer. 
	Note: Chips are still coated with a protective layer originating from their fabrication that needs to be removed.</li>
	<li>To remove the protective layer, perform a sequential solvent washing step. During the curing of chips, preheat a water bath to 54 &#176;C.</li>
	<li>Place 3 glass beakers into the water bath, two of them filled with isopropanol and one filled with acetone.</li>
	<li>Insert the chip cover glass into a slide holder and submerge it into the first isopropanol filled dish for 10 min. Afterwards transfer it to the acetone filled dish, incubate again for 10 min before transferring it for the final washing step to the second isopropanol dish and incubate another 10 min.</li>
	<li>Remove the slide holder from the dish and carefully wash it extensively with ultrapure water, followed by drying the chips with a gentle stream of nitrogen gas.</li>
	<li>Take-off the lamination sheet from the 8-well sticky-slide and put it backwards on the bench. Carefully pick up the cover slide and gently press onto the sticky-slide with the chips facing the wells. Let the finished chip holder rest for a couple of minutes before use.</li>
</ol>
6. Assay Preparation
Note: Nanopore chips glued to a cover glass and separated by an 8-well sticky-slide possess the benefit, that multiple chips can be addressed in a single experimental run, with chips being completely separated from each other.
<ol>
	<li>After chip holder preparation, rinse each chip with pure ethanol and blow dry with nitrogen gas, to remove any contaminations like dust.</li>
	<li>Clean the chip surface with air, oxygen or nitrogen plasma. Depending on the type of plasma cleaning time-periods vary: perform oxygen plasma treatment for 1 min, air and nitrogen plasma for 5 min at 0.3 mbar at 80% power setting (RF power medium or high).</li>
	<li>After plasma cleaning incubate chips in pure ethanol for 10-15 min to ensure cavity wetting.</li>
	<li>Stepwise exchange ethanol for buffer by adding 400 &#181;l of Tris buffer (20 mM Tris, 150 mM NaCl, pH 8.0; sterile filtered), followed by mixing of the solution without introducing air bubbles and subsequent removal of 400 &#181;l. Repeat this procedure 12 times, to ensure ethanol removal. 
	Note: Ethanol would be harmful for liposomes and suspended lipid bilayers.</li>
	<li>Dope Tris buffer (20 mM Tris, 150 mM NaCl, pH 8.0; sterile filtered) with 5 mM CaCl2 and 1 &#181;M Oy647 dye. Completely remove the washing buffer from the chip and immediately add the doped buffer to the chip. Note: Chips will be covered with a thin buffer film after wash buffer removal, so the chip is not in direct contact with air, ensuring a constant wetting of the cavities and the chip surface.</li>
	<li>Add 1 mg/ml LUVs or proteo-LUVs in Tris buffer (20 mM Tris, 150 mM NaCl, pH 8.0; sterile filtered) to each well. 
	Note: The final volume in each well is 300 &#181;l. Consider subsequent additions of the control dye and the chemical modifier MTSET during doped Tris buffer addition.</li>
	<li>For pore-spanning lipid bilayer formation on the chip surface incubate the chip for 1 hr at room temperature with the liposome solution. Afterwards wash each chip with 4x 400 &#181;l of Ca2+-free Tris buffer.</li>
	<li>Add the control dye to each well, with a final concentration of 5 &#181;M. The chips are now ready for the translocation assay.</li>
</ol>
7. Assay Setup
<ol>
	<li>Mount the chip holder to the epifluorescence microscope (20X air objective, long working distance). Focus on the rim of the nanopore chip to more easily find the focal plane of the micro-cavities. Once the cavities are in focus scan the nanopore array for a region that displays highest sealing ratio. 
	Note: Every inverted epifluorescence microscope can be used to perform chip assays. Depending on the objective magnification (20X - 60X) the number of assayed cavities will vary, with up to 9,000 cavities in a single field of view using 20X magnification. Air objectives with a long working distance are preferable. The use of inverted confocal laser scanning microscopes (CLSM) is also possible, but image acquisition might be the limiting factor due to the limited depth of focus.</li>
	<li>Optimize the chip illumination. 
	Note: Ensure that both dye channels do not exceed the maximum grey scale value, as changes cannot be observed. For export experiments adjust illumination of the translocated dye to 90% of maximum signal capacity. Adjust the control dye in open cavities to 80-90% of maximum signal capacity to clearly detect any leaky membrane sealing.</li>
	<li>Once all channels are adjusted start the time series. Take images every 10-20 sec for 90 min. 
	Note: This is adequate to likewise follow specific and unspecific efflux events.</li>
	<li>Initialize efflux of the fluorescent substrate by the addition of cysteine-specific, positively charged compound MTSET to a final concentration of 3 mM. Allow at least 5 min of experiment runtime before MTSET addition to be able to later on establish a stable fluorescent signal baseline prior to channel opening. Always prepare MTSET solution fresh directly before usage.</li>
</ol>
8. Data Analysis
<ol>
	<li>Open the time series image stack with a standard version of ImageJ (File/Import/Image Sequence).</li>
	<li>If fluorescent channels are stacked, split the channels in individual stacks for each channel (Image/Stacks/Stack to Images).</li>
	<li>Duplicate the first image of the translocation substrate channel for ROI (regions of interest) identification (Image/Duplicate).</li>
	<li>Convert the image to a binary image (Image/Adjust/Threshold). Ensure that the algorithm used depicts signals for all sealed cavities without overlapping pixilation.</li>
	<li>Automatically define regions of interest with the particle analyzer tool (Analyze/Analyze Particle). Define a minimum particle size defined, to avoid particle noise. Check the options &#34;exclude on edges&#34; and &#34;include holes&#34;. Ensure that resulting ROIs reflect the microcavity array pattern. Interjacent ROIs are artifacts, therefore remove them. Save the ROI list. 
	Note: Measurement parameter has to be set to &#34;area&#34; before particle analysis (Analyze/Set Measurements/Area)</li>
	<li>Open the Time Series Analyzer plug-in (http://rsb.info.nih.gov/ij/plugins/time-series.html), which resizes all ROIs automatically to the same size. Re-size all existing ROIs to a width/height of 6x6 pixels and oval shape (Timer Series Analyzer/AutoROIProperties). Check the option &#34;Resize existing ROIS&#34;. Save the resulting resized ROI list.</li>
	<li>Open the ROI manager (Analyze/Tools/ROI Manager) and load the resized ROI list for each channel of the time series (ROI Manger/More/Open). ROIs will be superposed. To analyze the signal change for each ROI start the Multi Measure tool (ROI Manager/More/Multi Measure), select &#34;Measure all slices&#34; and &#34;One row per slice&#34; and start the multi measure. 
	Note: Measurement parameter has to be set to &#34;mean gray value&#34; for this step (Analyze/Set Measurements/Mean gray value)</li>
	<li>Save the resulting table, giving the mean grey value for each ROI in *.txt or *.xls format.</li>
	<li>Import the ROI analysis of each imaging channel into NanoCalcFX via the &#34;Import File&#34; option. Set &#34;Use first line for naming series&#34;. Set the step size between images according to the time series image acquisition periods and choose the corresponding column and decimal separator.</li>
	<li>Use the &#34;multiple sheet&#34; option to automatically correlate the graphs of the individual fluorescent channels. 
	Note: The events can now be analyzed and classified using the information given by the 3-color spectral information.</li>
	<li>Fit selected curves using the build-in fit function (fit option). 
	Note: The implemented efflux and influx equation is suitable for all monoexponential diffusion-driven events and fit boundaries can be set. Resulting rate constants can be plotted using the histogram tool or exported for further data refinement.</li>
</ol>

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R., He&#769;lix-Nielsen, C., Malmstadt, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lipid+directed+intrinsic+membrane+protein+segregation.">Lipid directed intrinsic membrane protein segregation.</a> J. Am. Chem. Soc. 135, 17294-17297 (2013).</li><li>Heinemann, F., Schwille, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+Micrometer-Sized+Free-Standing+Membranes.">Preparation of Micrometer-Sized Free-Standing Membranes.</a> ChemPhysChem. 12, 2568-2571 (2011).</li><li>Lazzara, T. D., Carnarius, C., Kocun, M., Janshoff, A., Steinem, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Separating+attoliter-sized+compartments+using+fluid+pore-spanning+lipid+bilayers.">Separating attoliter-sized compartments using fluid pore-spanning lipid bilayers.</a> ACS nano. 5, 6935-6944 (2011).</li><li>Winterhalter, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Black+lipid+membranes.">Black lipid membranes.</a> Curr. Opin. Colloid Interface Sci. 5, 250-255 (2000).</li><li>Ko&#231;er, A., Walko, M., Feringa, B. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synthesis+and+utilization+of+reversible+and+irreversible+light-activated+nanovalves+derived+from+the+channel+protein+MscL.">Synthesis and utilization of reversible and irreversible light-activated nanovalves derived from the channel protein MscL.</a> Nat. Protoc. 2, 1426-1437 (2007).</li></ol>

The authors declare no competing financial interests.

The technique presented here allows a highly parallel analysis of membrane protein transport. Reconstituted membrane protein systems can directly be applied to the biochip, making the adaption of theoretically every membrane transporter or channel possible. Transport analysis is only limited by the establishment of a fluorescent read-out system, either via direct fluorescence change (translocation of fluorophores or fluorescently labeled substrates) or indirect fluorescence change (pH-sensitive dyes, secondary enzymatic ...

The analysis of membrane proteins has become of increasing interest for basic and pharmaceutical research in the past 20 years. The development of novel drugs depends on the identification and detailed characterization of new targets, currently being one of the limiting factors. The fact that about 60% of all drug targets are membrane proteins1, makes the development of techniques to elucidate their function most important.
In the past, techniques for the study of electrogenic channels and transporters have been developed in multitude2-4. Non-electrogenic substrates in contrary present a more challenging task. They are however of special interest as prime drug targets, as they control the flux of solutes and nutrients across the cell membrane and function as key receptors in signaling cascades5.
Considerable effort has been put into the development of techniques to study the function of membrane transport proteins6,7. Systems using solid-supported membranes have emerged as most promising tools in this field8-10, including solid supported lipid bilayers, tethered bilayers11,12, microblack lipid membranes13,14 and native vesicle arrays15,16 to name a few. Some of them are even available as commercial setups17,18. Some examples have been published combining the ability to study single membrane proteins in a highly parallel manner14,19, a prerequisite for screening applications. However, these methods rarely bridge from basic research to the industrial environment. The difficulties often lie in the ability of the system to be automatable, the cost-intensive production and/or laborious preparation. An approach overcoming all the above mentioned obstacles is the final aim.
The technique presented here was developed to study membrane channels and transporters in vitro in a controlled environment on the single protein level20-22. Reconstitution of purified membrane proteins into LUVs is much more established than comparable approaches for GUVs23-26 or black lipid membranes27. They can directly be applied to the chip surface, where bilayer formation is taking place via a self-assembly process. The glass-bottom design of the nanoporous chip (Fig. 1) allows for air microscopy, which permits the straightforward automation of the system. In combination with a motorized stage multiple chips can be measured at the same time, with each field of view containing thousands of sealed cavities for analysis.
<img alt="Figure 1" src="/files/ftp_upload/53373/53373fig1.jpg" /> 
Figure 1. Design of multiplexed nanopore biochips. A) A silicon-on-insulator (SOI) wafer is structured by reactive-ion etching. Approximately 1,150 individual chips are fabricated from each wafer with identical properties and quality. B) Each chip comprises 250,000 individual microcavities with nano apertures. Scale bar: 200 &#181;m. C) Each cavity is addressable via multi-spectral fluorescence read-out. An intransparent top layer blocks the fluorescent signals from the buffer reservoir, making the biochip compatible with inverted fluorescence microscopes. D) Atomic force microscopy (AFM) imaging reveals evenly arranged pore openings and surface roughness of the silicon dioxide layer of 3.6 nm (n = 40) optimal for vesicle fusion. Scale bar: 5 &#181;m. E) Scanning electron microscopy (SEM) image shows a cross-section through the nanopore allowing access to the femtoliter cavities inside the silicon chip. This figure was reused with permission from 21. <a href="https://www.jove.com/files/ftp_upload/53373/53373fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
All data analysis is performed using freeware to guarantee unrestricted access for end users. Time series are analyzed using free image processing software and a custom build curve analysis software enabling batch processing and straightforward correlation of large datasets with multiple fluorescent channels and thousands of curves.
The model protein used in this protocol is the mechanosensitive channel of large conductance (MscL) channel protein derived from E. coli. It functions as a valve to release osmotic shock in nature, but was modified in such a way that rationally designed synthetic functionalities can covalently be attached to the channels constriction side. Via charge-repulsion of the covalently bound activator (MTSET) the channel is triggered to open, creating a nano-valve. Small molecules like ions, water, small proteins, but also small fluorophores can permeate through the channel. Here, the protein is used as a model to demonstrate the ability of the system to detect protein-mediated translocation.

<table border="0" cellpadding="0" cellspacing="0" width="1093">
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			<td height="21" style="height:21px;width:389px;">Reagent</td>
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			<td height="21" style="height:21px;width:389px;">[2-(Trimethylammonium)ethyl] 
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			<td height="21" style="height:21px;width:389px;">NyONE microscope</td>
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			<td height="21" style="height:21px;width:389px;">Plasma Cleaner PDC-37G</td>
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			<td height="21" style="height:21px;width:389px;">ImageJ</td>
			<td style="width:232px;">Open Source</td>
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			<td style="width:305px;">http://sourceforge.net/projects/nanocalc/</td>
			<td>data analysis/evaluation software for massive transport kinetic datasets</td>
		</tr>
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			<td height="21" style="height:21px;width:389px;">NTA 2.3 Analytical Software</td>
			<td style="width:232px;">Nanosight</td>
			<td style="width:305px;"></td>
			<td>data acquisition and analysis software for nanoparticle tracking microscope</td>
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			<td height="21" style="height:21px;width:389px;">NTA 2.3 Temperature Comms</td>
			<td style="width:232px;">Nanosight</td>
			<td style="width:305px;"></td>
			<td>temperature controle software for nanoparticle tracking microscope</td>
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</table>

membrane transport processes analyzed by a highly parallel nanopore chip system at single protein resolution

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

Pore-spanning membranes can easily be created on the nanostructured chip surface in a self-assembled manner. However the underlying process is still delicate and influenced by many parameters like liposome size, monodispersity of the liposome population, lamellarity, lipid- and salt-concentration and chemical surface properties. Most of these parameters have been carefully characterized and standardized in the above protocol. Other parameters however should be checked during every new pre...

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

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

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

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JoVE Journal

Biology

11.6K Views.  Goethe University Frankfurt. The presented protocol describes the analysis of membrane protein mediated transport on the single transporter level using pore-spanning solvent-free lipid bilayers. This is achieved by the creation of bulk produced nanopore array chips, combined with highly parallel data acquisition and analysis, enabling the future establishment of membrane protein effector screenings.

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

Measuring Plasma Membrane Protein Endocytic Rates by Reversible Biotinylation

Plasma membrane proteins are a large, diverse group of proteins comprised of receptors, ion channels, transporters and pumps.  Activity of these proteins is responsible for a variety of key cellular events, including nutrient delivery, cellular excitability, and chemical signaling.  Many plasma membrane proteins are dynamically regulated by endocytic trafficking, which modulates protein function by altering protein surface expression.  The mechanisms that facilitate protein endocytosis are complex and are not fully understood for many membrane proteins.  In order to fully understand the mechanisms that control the endocytic trafficking of a given protein, it is critical that the protein s endocytic rate be precisely measured.  For many receptors, direct endocytic rate measurements are frequently achieved utilizing labeled receptor ligands.  However, for many classes of membrane proteins, such as transporters, pumps and ion channels, there is no convenient ligand that can be used to measure the endocytic rate.  In the present report, we describe a reversible biotinylation method that we employ to measure the dopamine transporter (DAT) endocytic rate.  This method provides a straightforward approach to measuring internalization rates, and can be easily employed for trafficking studies of most membrane proteins.

Regulated endocytosis governs the cell surface expression levels of the majority of membrane proteins. Here we utilize reducible, membrane impermeant biotinylation reagents to measure the endocytic rate of the dopamine transporter (DAT), a polytopic membrane protein. The method facilitates a straightforward approach to measuring the endocytic rate of most plasma membrane proteins.

Plasma membrane proteins are a large, diverse group of proteins comprised of receptors, ion channels, transporters and pumps.  Activity of these proteins ...

Split-Ubiquitin Based Membrane Yeast Two-Hybrid (MYTH) System: A Powerful Tool For Identifying Protein-Protein Interactions

The fundamental biological and clinical importance of integral membrane proteins prompted the development of a yeast-based system for the high-throughput identification of protein-protein interactions (PPI) for full-length transmembrane proteins. To this end, our lab developed the split-ubiquitin based Membrane Yeast Two-Hybrid (MYTH) system. This technology allows for the sensitive detection of transient and stable protein interactions using Saccharomyces cerevisiae as a host organism. MYTH takes advantage of the observation that ubiquitin can be separated into two stable moieties: the C-terminal half of yeast ubiquitin (Cub) and the N-terminal half of the ubiquitin moiety (Nub). In MYTH, this principle is adapted for use as a 'sensor' of protein-protein interactions. Briefly, the integral membrane bait protein is fused to Cub which is linked to an artificial transcription factor. Prey proteins, either in individual or library format, are fused to the Nub moiety. Protein interaction between the bait and prey leads to reconstitution of the ubiquitin moieties, forming a full-length 'pseudo-ubiquitin' molecule. This molecule is in turn recognized by cytosolic deubiquitinating enzymes, resulting in cleavage of the transcription factor, and subsequent induction of reporter gene expression. The system is highly adaptable, and is particularly well-suited to high-throughput screening. It has been successfully employed to investigate interactions using integral membrane proteins from both yeast and other organisms.

Split-Ubiquitin Based Membrane Yeast Two-Hybrid MYTH System: A Powerful Tool For Identifying Protein-Protein Interactions

MYTH allows the sensitive detection of transient and stable interactions between proteins that are expressed in the model organism Saccharomyces cerevisiae. It has been successfully applied to study exogenous and yeast integral membrane proteins in order to identify their interacting partners in a high throughput manner.

The fundamental biological and clinical importance of integral membrane proteins prompted the development of a yeast-based system for the high-throughput ...

Using the GELFREE 8100 Fractionation System for Molecular Weight-Based Fractionation with Liquid Phase Recovery

The GELFREE 8100 Fractionation System is a novel protein fractionation system designed to maximize protein recovery during molecular weight based fractionation. The system is comprised of single-use, 8-sample capacity cartridges and a benchtop GELFREE Fractionation Instrument. During separation, a constant voltage is applied between the anode and cathode reservoirs, and each protein mixture is electrophoretically driven from a loading chamber into a specially designed gel column gel. Proteins are concentrated into a tight band in a stacking gel, and separated based on their respective electrophoretic mobilities in a resolving gel. As proteins elute from the column, they are trapped and concentrated in liquid phase in the collection chamber, free of the gel. The instrument is then paused at specific time intervals, and fractions are collected using a pipette. This process is repeated until all desired fractions have been collected. If fewer than 8 samples are run on a cartridge, any unused chambers can be used in subsequent separations.
This novel technology facilitates the quick and simple separation of up to 8 complex protein mixtures simultaneously, and offers several advantages when compared to previously available fractionation methods. This system is capable of fractionating up to 1mg of total protein per channel, for a total of 8mg per cartridge. Intact proteins over a broad mass range are separated on the basis of molecular weight, retaining important physiochemical properties of the analyte. The liquid phase entrapment provides for high recovery while eliminating the need for band or spot cutting, making the fractionation process highly reproducible1.

The accompanying video describes the use of the GELFREE 8100 Fractionation System, which partitions complex protein samples on the basis of molecular weight and recovers the fractions in liquid phase. The video describes how the technology works, how it is used, and provides resultant data, with polyacrylamide gel electrophoresis analysis of fractionated bovine liver homogenate.

The GELFREE 8100 Fractionation System is a novel protein fractionation system designed to maximize protein recovery during molecular weight based ...

Single-Molecule Imaging of Nuclear Transport

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the last decade. The investigation of molecular interactions with high temporal and spatial resolutions deep within cells has remained challenging due to the inherently weak signals arising from individual molecules. Recent works by Yang et al. demonstrated that narrow-field epifluorescence microscopy allows visualization of nucleocytoplasmic transport at the single molecule level. By the single molecule approach, important kinetics, such as nuclear transport time and efficiency, for signal-dependent and independent cargo molecules have been obtained. Here we described a protocol for the methodological approach with an improved spatiotemporal resolution of 0.4 ms and 12 nm. The improved resolution enabled us to capture transient active transport and passive diffusion events through the nuclear pore complexes (NPC) in semi-intact cells. We expect this method to be used in elucidating other binding and trafficking events within cells.

Single molecule microscopy approch provided novel insights into nuclear transport.

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the ...

Direct Analysis of Single Cells by Mass Spectrometry at Atmospheric Pressure

Analysis of biochemicals in single cells is important for understanding cell metabolism, cell cycle, adaptation, disease states, etc. Even the same cell types exhibit heterogeneous biochemical makeup depending on their physiological conditions and interactions with the environment. Conventional methods of mass spectrometry (MS) used for the analysis of biomolecules in single cells rely on extensive sample preparation. Removing the cells from their natural environment and extensive sample processing could lead to changes in the cellular composition. Ambient ionization methods enable the analysis of samples in their native environment and without extensive sample preparation.1 The techniques based on the mid infrared (mid-IR) laser ablation of biological materials at 2.94 &mu;m wavelength utilize the sudden excitation of water that results in phase explosion.2 Ambient ionization techniques based on mid-IR laser radiation, such as laser ablation electrospray ionization (LAESI) and atmospheric pressure infrared matrix-assisted laser desorption ionization (AP IR-MALDI), have successfully demonstrated the ability to directly analyze water-rich tissues and biofluids at atmospheric pressure.3-11 In LAESI the mid-IR laser ablation plume that mostly consists of neutral particulate matter from the sample coalesces with highly charged electrospray droplets to produce ions. Recently, mid-IR ablation of single cells was performed by delivering the mid-IR radiation through an etched fiber. The plume generated from this ablation was postionized by an electrospray enabling the analysis of diverse metabolites in single cells by LAESI-MS.12 This article describes the detailed protocol for single cell analysis using LAESI-MS. The presented video demonstrates the analysis of a single epidermal cell from the skin of an Allium cepa bulb. The schematic of the system is shown in Figure 1. A representative example of single cell ablation and a LAESI mass spectrum from the cell are provided in Figure 2.

Single cell analysis is performed by mass spectrometry on plant and animal cells at atmospheric pressure by using a sharpened optical fiber to sample the cells for laser ablation electrospray ionization (LAESI) mass spectrometry.

Analysis of biochemicals in single cells is important for understanding cell metabolism, cell cycle, adaptation, disease states, etc. Even the same cell ...

Real-time Analyses of Retinol Transport by the Membrane Receptor of Plasma Retinol Binding Protein

Vitamin A is essential for vision and the growth/differentiation of almost all human organs. Plasma retinol binding protein (RBP) is the principle and specific carrier of vitamin A in the blood. Here we describe an optimized technique to produce and purify holo-RBP and two real-time monitoring techniques to study the transport of vitamin A by the high-affinity RBP receptor STRA6. The first technique makes it possible to produce a large quantity of high quality holo-RBP (100%-loaded with retinol) for vitamin A transport assays. High quality RBP is essential for functional assays because misfolded RBP releases vitamin A readily and bacterial contamination in RBP preparation can cause artifacts. Real-time monitoring techniques like electrophysiology have made critical contributions to the studies of membrane transport. The RBP receptor-mediated retinol transport has not been analyzed in real time until recently. The second technique described here is the real-time analysis of STRA6-catalyzed retinol release or loading. The third technique is real-time analysis of STRA6-catalyzed retinol transport from holo-RBP to cellular retinol binding protein I (CRBP-I). These techniques provide high sensitivity and resolution in revealing RBP receptor's vitamin A uptake mechanism.

Here we describe an optimized technique to produce high-quality vitamin A/RBP complex and two real-time monitoring techniques to study vitamin A transport by STRA6, the RBP receptor.

Vitamin A is essential for vision and the growth/differentiation of almost all human organs.  Plasma retinol binding protein (RBP) is the principle and ...

Visualizing Clathrin-mediated Endocytosis of G Protein-coupled Receptors at Single-event Resolution via TIRF Microscopy

Many important signaling receptors are internalized through the well-studied process of clathrin-mediated endocytosis (CME). Traditional cell biological assays, measuring global changes in endocytosis, have identified over 30 known components participating in CME, and biochemical studies have generated an interaction map of many of these components. It is becoming increasingly clear, however, that CME is a highly dynamic process whose regulation is complex and delicate. In this manuscript, we describe the use of Total Internal Reflection Fluorescence (TIRF) microscopy to directly visualize the dynamics of components of the clathrin-mediated endocytic machinery, in real time in living cells, at the level of individual events that mediate this process. This approach is essential to elucidate the subtle changes that can alter endocytosis without globally blocking it, as is seen with physiological regulation. We will focus on using this technique to analyze an area of emerging interest, the role of cargo composition in modulating the dynamics of distinct clathrin-coated pits (CCPs). This protocol is compatible with a variety of widely available fluorescence probes, and may be applied to visualizing the dynamics of many cargo molecules that are internalized from the cell surface.

Clathrin-mediated endocytosis, a rapid and highly dynamic process internalizes many proteins, including signaling receptors. The protocol described here directly visualizes the kinetics of individual endocytic events. This is essential for understanding how core members of the endocytic machinery coordinate with each other, and how protein cargo influence this process.

Many important signaling receptors are internalized through the well-studied process of clathrin-mediated endocytosis (CME). Traditional cell biological ...

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and the inherent instability of many membrane proteins once solubilized in detergents. A protocol is described that combines ligation independent cloning of membrane proteins as GFP fusions with expression in Escherichia coli detected by GFP fluorescence. This enables the construction and expression screening of multiple membrane protein/variants to identify candidates suitable for further investment of time and effort. The GFP reporter is used in a primary screen of expression by visualizing GFP fluorescence following SDS polyacrylamide gel electrophoresis (SDS-PAGE). Membrane proteins that show both a high expression level with minimum degradation as indicated by the absence of free GFP, are selected for a secondary screen. These constructs are scaled and a total membrane fraction prepared and solubilized in four different detergents. Following ultracentrifugation to remove detergent-insoluble material, lysates are analyzed by fluorescence detection size exclusion chromatography (FSEC). Monitoring the size exclusion profile by GFP fluorescence provides information about the mono-dispersity and integrity of the membrane proteins in different detergents. Protein: detergent combinations that elute with a symmetrical peak with little or no free GFP and minimum aggregation are candidates for subsequent purification. Using the above methodology, the heterologous expression in E. coli of SED (shape, elongation, division, and sporulation) proteins from 47 different species of bacteria was analyzed. These proteins typically have ten transmembrane domains and are essential for cell division. The results show that the production of the SEDs orthologues in E. coli was highly variable with respect to the expression levels and integrity of the GFP fusion proteins. The experiment identified a subset for further investigation.

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

A streamlined approach to screening for the expression of recombinant membrane proteins in Escherichia coli based on fusion to green fluorescent protein is presented.

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and ...

CUBIC Protocol Visualizes Protein Expression at Single Cell Resolution in Whole Mount Skin Preparations

The skin is essential for our survival. The outer epidermal layer consists of the interfollicular epidermis, which is a stratified squamous epithelium covering most of our body, and epidermal appendages such as the hair follicles and sweat glands. The epidermis undergoes regeneration throughout life and in response to injury. This is enabled by K14-expressing basal epidermal stem/progenitor cell populations that are tightly regulated by multiple regulatory mechanisms active within the epidermis and between epidermis and dermis. This article describes a simple method to clarify full thickness mouse skin biopsies, and visualize K14 protein expression patterns, Ki67 labeled proliferating cells, Nile Red labeled sebocytes, and DAPI nuclear labeling at single cell resolution in 3D. This method enables accurate assessment and quantification of skin anatomy and pathology, and of abnormal epidermal phenotypes in genetically modified mouse lines. The CUBIC protocol is the best method available to date to investigate molecular and cellular interactions in full thickness skin biopsies at single cell resolution.

This report describes a CUBIC protocol to clarify full thickness mouse skin biopsies, and visualize protein expression patterns, proliferating cells, and sebocytes at the single cell resolution in 3D. This method enables accurate assessment of skin anatomy and pathology, and of abnormal epidermal phenotypes in genetically modified mouse lines.

The skin is essential for our survival. The outer epidermal layer consists of the interfollicular epidermis, which is a stratified squamous epithelium ...

Methods to Study Epithelial Transport Protein Function and Expression in Native Intestine and Caco-2 Cells Grown in 3D

The intestinal epithelium has important transport and barrier functions that play key roles in normal physiological functions of the body while providing a barrier to foreign particles. Impaired epithelial transport (ion, nutrient, or drugs) has been associated with many diseases and can have consequences that extend beyond the normal physiological functions of the transporters, such as by influencing epithelial integrity and the gut microbiome. Understanding the function and regulation of transport proteins is critical for the development of improved therapeutic interventions. The biggest challenge in the study of epithelial transport is developing a suitable model system that recapitulates important features of the native intestinal epithelial cells. Several in vitro cell culture models, such as Caco-2, T-84, and HT-29-Cl.19A cells are typically used in epithelial transport research. These cell lines represent a reductionist approach to modeling the epithelium and have been used in many mechanistic studies, including their examination of epithelial-microbial interactions. However, cell monolayers do not accurately reflect cell-cell interactions and the in vivo microenvironment. Cells grown in 3D have shown to be promising models for drug permeability studies. We show that Caco-2 cells in 3D can be used to study epithelial transporters. It is also important that studies in Caco-2 cells are complemented with other models to rule out cell specific effects and to take into account the complexity of the native intestine. Several methods have been previously used to assess the functionality of transporters, such as everted sac and uptake in isolated epithelial cells or in isolated plasma membrane vesicles. Taking into consideration the challenges in the field with respect to models and the measurement of transport function, we demonstrate here a protocol to grow Caco-2 cells in 3D and describe the use of an Ussing chamber as an effective approach to measure serotonin transport, such as in intact polarized intestinal epithelia.

We describe simple methods to study the regulation of intestinal serotonin transporter (SERT) function and expression using an in vitro cell culture model of Caco-2 cells grown in 3D&#160;and an ex vivo model of mouse intestines. These methods are applicable to the study of other epithelial transporters.

The intestinal epithelium has important transport and barrier functions that play key roles in normal physiological functions of the body while providing ...