Reconstitution of a Transmembrane Protein, the Voltage-gated Ion Channel, KvAP, into Giant Unilamellar Vesicles for Microscopy and Patch Clamp Studies

Summary

The reconstitution of the transmembrane protein, KvAP, into giant unilamellar vesicles (GUVs) is demonstrated for two dehydration-rehydration methods — electroformation, and gel-assisted swelling. In both methods, small unilamellar vesicles containing the protein are fused together to form GUVs that can then be studied by fluorescence microscopy and patch-clamp electrophysiology.

Abstract

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow GUVs must be modified in order to form GUVs containing functional transmembrane proteins. This article describes two dehydration-rehydration methods — electroformation and gel-assisted swelling — to form GUVs containing the voltage-gated potassium channel, KvAP. In both methods, a solution of protein-containing small unilamellar vesicles is partially dehydrated to form a stack of membranes, which is then allowed to swell in a rehydration buffer. For the electroformation method, the film is deposited on platinum electrodes so that an AC field can be applied during film rehydration. In contrast, the gel-assisted swelling method uses an agarose gel substrate to enhance film rehydration. Both methods can produce GUVs in low (e.g., 5 mM) and physiological (e.g., 100 mM) salt concentrations. The resulting GUVs are characterized via fluorescence microscopy, and the function of reconstituted channels measured using the inside-out patch-clamp configuration. While swelling in the presence of an alternating electric field (electroformation) gives a high yield of defect-free GUVs, the gel-assisted swelling method produces a more homogeneous protein distribution and requires no special equipment.

Introduction

When studying the physical principles that govern living systems, bottom-up approaches allow an experimentalist to control system composition and other parameters that are not easily manipulated in cell-based systems¹. For membrane-based processes, Giant Unilamellar Vesicles (GUVs, diameter ~ 1–100 µm) have proven to be a very useful biomimetic system^2–7 as they are well suited for microscopy studies and micromanipulation^8–10. While there are many different protocols to produce GUVs, most fall into two categories — emulsion based approaches^11,12 and techniques based on rehydrating a lipid film^13–16. In emulsion-based methods, the inner and outer leaflets of the GUV membranes are assembled sequentially from lipid monolayers at water/oil interfaces. This approach is ideal for encapsulating soluble proteins within the GUVs, and for forming GUVs with asymmetric leaflet lipid composition. However, GUVs formed from emulsions can retain traces of solvent that change the membrane’s mechanical properties¹⁷, and the approach is not especially well-suited to trans-membrane protein reconstitution.

Film rehydration methods rely on the fact that drying (dehydration) causes many lipid mixtures to form a multi-lamellar stack of membranes. If this stack is then placed in contact with an aqueous buffer, membranes in the stack will move apart as solvent flows between them and at the surface of the stack, individual membranes can detach to form GUVs^13,18 (as well a veritable zoo of other lipidic objects). However, even for optimal buffer and lipid compositions, this classical “spontaneous swelling” method has a relatively low yield of defect-free GUVs. One widely used method to boost the yield of defect-free GUVs is “electroformation”, in which an alternating current (AC) field is applied during film rehydration. While the mechanism remains poorly understood, “electroformation” can give spectacular GUV yields (> 90% in favorable circumstances) for low salt concentration buffers (< 5 mM)^14,19, and can even work in physiological buffers (~100 mM) using a higher frequency (500 Hz versus 10 Hz) AC field and platinum electrodes¹⁵. An alternative approach to boost the yield of defect-free GUVs is “gel-assisted swelling”, in which the lipid solution is deposited onto a polymeric gel substrate rather than the passive (e.g., glass, PTFE) substrates used in classical “spontaneous swelling”. When the resultant lipid/gel film is rehydrated, GUVs can rapidly form even for physiological buffers^16,20.

All these methods can produce lipid-only GUVs which can be used to study membrane associated phenomena such as the interaction between soluble proteins and membranes. However, to incorporate a trans-membrane protein into GUVs, significant modifications are needed to ensure that the protein remains in a functional state throughout the reconstitution procedure. While solutions of lipids in organic solvents (e.g., chloroform, cyclohexane) are ideal for producing lipid films, trans-membrane proteins are typically only stable when their hydrophobic trans-membrane domain is embedded in a lipid bilayer, or surrounded by a detergent micelle (e.g., during protein purification). Thus, the starting material for a reconstitution is typically native membranes, purified protein in a detergent solution, or small unilamellar protein-containing vesicles (proteo-SUVs) and/or multi-lamellar vesicles (proteo-MLVs) formed by detergent removal in the presence of lipids. Most methods to incorporate these membrane proteins into GUVs fall into three categories.

Direct Insertion: Trans-membrane protein suspended in detergent is mixed with pre-formed, lipid-only, mildly detergent solubilized GUVs, and the detergent then removed using biobeads²¹. While conceptually simple, this method requires precise control of the detergent concentration, as too high a detergent concentration can dissolve the GUVs while too low a concentration can cause the protein to unfold or aggregate.

GUV/Proteo-SUV Fusion: Protein in proteo-SUVs is combined with pre-formed, lipid-only GUVs and fusion is facilitated with special fusogenic peptides²² or detergent²¹. Typically the extent of fusion is limited leading to GUVs with low protein density.

Dehydration/Rehydration: A protein-containing lipid film is formed by partial dehydration of a proteo-SUV (or proteo-MLV) solution and GUVs are then grown as for a pure lipid film. The obvious challenge is to protect the protein during the partial dehydration step²³, but the method has been successfully used to reconstitute trans-membrane proteins such as Bacteriorhodopsin, Calcium-ATPase, Integrin and VDAC into GUVs^7,23–25.

This article describes dehydration/rehydration protocols to make GUVs containing the voltage-gated potassium channel, KvAP, from the hyper-thermophilic Archaea, Aeropyrum pernix. KvAP has a high degree of homology to eukaryotic voltage dependent potassium channels²⁶ and a known crystal structure²⁷, making it a good model for studying the mechanism of voltage gating. Production of the proteo-SUVs has been described in detail previously and is not part of this tutorial^26,28,29. Importantly, KvAP proteo-SUVs do not have to be produced for each GUV preparation, as they can be stored in small (e.g., 10 µl) aliquots at -80 °C for extended periods of time (> 1 year). Electroformation or gel-assisted swelling can then be used to grow GUVs from the KvAP proteo-SUVs (or proteo-MLVs).

The key steps for the electroformation protocol are illustrated in Figure 1. Droplets of a solution of SUVs containing the protein are deposited on platinum wires (shown in Figure 2). Partial dehydration of the SUV suspension leads to the formation of a lipid protein film through the fusion of SUVs. During rehydration, an AC field is applied to the electrodes to assist the lipid layers to delaminate and form GUVs. A 10 Hz field works well when using “low-salt” (< 5 mM) rehydration buffer²⁸ and GUVs take several hours to grow. In contrast, physiological buffers (containing ~100 mM salt) work well with a lower voltage, 500 Hz AC field but require a prolonged (~12 hr) swelling period¹⁵. This method is based upon an earlier protocol using ITO slides²⁴, but uses a custom chamber containing two platinum wires as shown in Figure 2 (see the discussion for design details and suggestions for simpler, improvised chambers).

Figure 3 illustrates the gel-assisted swelling method. The protocol works well with buffers with physiological salt concentrations, is rapid, and produces GUVs with a more homogeneous protein distribution. However, the yield of isolated, apparently defect-free GUVs (i.e., the GUV membrane is uniform at optical length-scales and does not enclose any objects) is lower, although it provides a sufficient number for patch-clamp and micro-manipulation experiments. This method was based on a protocol using agarose gel to produce lipid-only GUVs¹⁶ and requires less specialized equipment than the electroformation method.

The characterization of GUVs with fluorescence microscopy is described, as well as procedures using a standard patch-clamp set-up to measure KvAP activity in “inside-out” excised membrane patches.

Growing protein-containing GUVs can be more difficult than lipid-only GUVs. In particular, the final GUV yield can depend sensitively on exactly how the SUV solution is deposited and dehydrated to form the membrane stack. For someone without any previous experience with GUVs, it may be helpful to first grow lipid-only GUVs following a conventional protocol^15,16 in which the membrane film is formed by depositing lipids from an organic solvent. Once the conventional protocol works well, SUV deposition and partial dehydration can then be mastered using lipid-only SUVs, which are also very helpful when adjusting the protocol for a new lipid composition. When GUVs grow reliably from lipid-only SUVs, it is then only a small step to produce protein-containing GUVs from proteo-SUVs.

Protocol

1. Solution Preparation

1. Prepare 5 ml of ‘SUV buffer’ containing 5 mM KCl, 1 mM HEPES (pH 7.4) or TRIS (pH 7.5), and 2 mM trehalose. Filter the buffer with a 0.2 µm syringe filter and divide into 1 ml aliquots which can be stored at -20 °C.
   NOTE: Additional details for reagents and instruments are given in the materials list.
2. Prepare 40 ml of GUV ‘Growth Buffer’ that will fill the GUV interior during film rehydration. For a ‘low salt’ growth, combine 5 mM KCl, 1 mM HEPES (pH 7.4) or 1 mM TRIS (pH 7.5), and ~400 mM sucrose. For a ‘physiological salt’ growth, combine 100 mM KCl, 5 mM HEPES (pH 7.4) or 5 mM TRIS (pH 7.5), and ~200 mM sucrose.
3. Prepare 40 ml of ‘Observation Buffer’ for the external solution in the experimental chamber by combining 100 mM KCl, 5 mM HEPES (pH 7.4) or 5 mM TRIS (pH 7.5), and ~200 mM glucose.
   NOTE: These buffers are only examples. See the discussion to adapt the buffers for other experiments.
4. Measure the osmolarities of the growth and observation buffers with an osmometer. Add granules of sucrose or glucose to match them to within 1% so that GUVs will not lyse or collapse when transferred from the growth chamber to the observation chamber.
   NOTE: In this concentration range, adding 1 mM of sucrose (13.7 mg per 40 ml) or glucose (7.2 mg per 40 ml) increases osmolarity by ~ 1 mOsm.
5. Filter the growth and observation buffers with a 0.2 µm filter and store them at 4 °C to inhibit bacterial growth.
6. Dissolve 50 mg of beta-casein in 10 ml of 20 mM TRIS (pH 7.5) buffer to form a 5 mg/ml beta-casein solution needed to passivate the surfaces of experimental chambers so that GUVs do not stick, spread and burst. Once the beta-casein is completely dissolved (up to several hours at 4 °C), filter (0.2 µm) it into 0.5 ml aliquots which can be flash frozen and stored at -20 °C for later use (thawed aliquots stored at 4 °C can typically be used for up to a 1 week).

2. SUV Preparation

1. Prepare and freeze aliquots of proteo-SUVs following the previously published detailed protocol²⁸. Use KvAP fluorescently labeled with Alexa-488 maleimide, reconstituted into DPhPC SUVs (10 mg/ml) at a protein to lipid ratio of 1:10 (by mass).
   NOTE: Wild-type KvAP contains one cysteine per monomer located near the intra-cellular C-terminus (amino acid 247).
2. Fluorescent, Lipid-only SUVs:
   NOTE: Handle chloroform under a fume hood wearing nitrile gloves and safety glasses. Avoid the use of any plastic as chloroform can dissolve them. Chloroform solutions can be stored in amber glass vials with Teflon caps and transferred using glass syringes. Take care to rinse all glassware at least 5 to 10 times with chloroform before and after pipetting lipids.
   1. Prepare 100 µl of 10 mg/ml DPhPC SUVs containing 0.5 mol% of the red fluorescent lipid, Texas Red-DHPE, by mixing 100 µl of DPhPC solution (10 mg/ml in chloroform) with 8.2 µl of Texas Red-DHPE solution (1 mg/ml in methanol) in a 1.5 ml amber glass vial.
   2. Dry the lipids down under a stream of nitrogen in a chemical hood while rotating the vial. When the film appears to be dry, place the lipids under a vacuum for 3 hr to remove any residual solvent.
   3. Add 100 µl of SUV buffer to the lipids, and vortex vigorously until no lipid remains stuck to the walls of the vial and the solution is uniformly milky.
   4. Sonicate the lipid solution to form SUVs. Adjust the vial position until the ultrasound causes the most movement and flow inside the vial, and take care not to heat the solution unnecessarily. Continue sonication until the solution becomes translucent, or when possible, transparent (2–5 min for tip sonication, ~20 min for bath sonication).
   5. Aliquot SUVs (e.g., 10 µl or 20 µl) and freeze (-20 °C) for later use.
      NOTE: Lipids, especially unsaturated lipids, can easily breakdown. Store lipid solutions at 20 °C (or 80 °C) under argon and use within 6 months. Lipid breakdown products can be detected with Thin Layer Chromatography.

3. GUV Growth by Electroformation

1. Prepare the electroformation chamber.
   1. If the chamber has not been cleaned, remove the windows, wipe off all sealant and grease, extract the wires, and rinse and scrub the chamber with a tissue using water and ethanol (≥70%) alternately.
   2. Rub the wires well, submerge the wires and chamber in acetone, and sonicate for 5 min. Wipe everything with a tissue again using acetone. Put the chamber in ethanol and sonicate for 5 min.
   3. Assemble the chamber by inserting the wires through the holes, and rotate and wipe the wires to make sure they are clean. Put the chamber in distilled water, sonicate for 5 min and dry the chamber with a stream of nitrogen or air.
2. Prepare 30 µl of 3 mg/ml SUV suspension in SUV buffer. To form protein-containing GUVs, combine 8 µl of proteo-SUVs (DPhPC 10 mg/ml KvAP 1:10), 2 µl of fluorescent SUVs (10 mg/ml DPhPC, 0.5 mol% TexasRed-DHPE) and 20 µl of SUV buffer in a 1.5 ml microcentrifuge tube for a final protein to lipid (mass) ratio of 1:12.5 and 0.1 mol% TexasRed-DHPE. Mix the solution vigorously.
   1. Alternatively, to practice the protocol with lipid-only SUVs, simply combine 10 µl of fluorescent SUVs (10 mg/ml DPhPC and 0.5 mol% TexasRed-DHPE) with 20 µl SUV buffer.
3. Deposit the SUV Solution.
   1. Use a 2 µl pipette or 5 µl glass syringe to deposit small (<0.2 µl) droplets of the SUV solution on the wires. Approximately 1 µl of solution is needed to form a series of drops along 1 cm of wire. Make sure the drops are small enough and spaced far enough apart that they do not touch or fuse.
   2. Let the deposited SUVs dry for ~30 min in open air. When all the drops have settled, rotate the wire so the lipid deposits are easier to observe with the microscope.
      NOTE: If the SUVs do not dry sufficiently, they can just wash off the wires when the growth buffer is added, while drying too much can damage the protein. Because air humidity influences the rate of drying, the drying time and/or air humidity can be adjusted for optimal results³⁰. The lipid film on the wires should be visible under a microscope.
4. Assemble the chamber.
   1. Seal the Chamber Bottom: Use a syringe to apply vacuum grease to the bottom of the chamber around the three wells and press a 40 mm x 22 mm coverslip gently against it to seal the chamber bottom so that it adheres without a gap. Seal the sides of the chamber (where the wires exit) with sealing paste. Apply vacuum grease on top of the chamber outlining the three wells.
   2. Slowly add growth buffer until each well is filled to the top. Avoid any rapid movement of the solution in the wells as this can strip the lipid film off the electrodes.
   3. Close the chamber by pressing the top cover slide gently onto the grease, taking care not to dislodge the bottom coverslip. Use a tissue to remove any drops of buffer at the edges of the top cover slip.
      NOTE: This is a good time to examine the chamber under the microscope to confirm that the lipid film has remained on the wires.
5. Connect the signal generator to the wires using two alligator clips. Set the frequency (10 Hz/500 Hz sine wave for low/high salt buffer) and use a multimeter to measure and adjust the voltage across the wires to 0.7/0.35 V root mean square (Vrms) for the low/high salt buffer. Cover the chamber with aluminum foil to protect the fluorophores from light. Leave the GUVs to grow for 2 to 3 hr for the low salt buffer, and 12 hr or O/N for the high salt buffer.
6. Disconnect the chamber from the generator and carefully place it on an inverted microscope to evaluate GUV growth. Use slow, steady movements or fluid flow in the wells may prematurely detach GUVs from the wires.
   NOTE: GUVs on the wire edges are usually visible in phase contrast (40X long working distance objective), while GUVs anywhere on the lower half of the wires can be seen with epifluorescence. If no GUVs are visible, try rotating the wires to look at the upper surface. GUVs can be stored at 4 °C in a growth chamber for several days.

4. GUV Growth by Gel-assisted Swelling

1. Prepare 10 ml of a 1% agarose solution by mixing 100 mg of agarose with 10 ml of pure water. Heat it until it boils by placing it in a microwave at 480 W for ~20 sec. Stir to make sure the agarose is completely dissolved.
   NOTE: The solution can be stored at 4 °C and reheated when needed.
2. Plasma-clean (air plasma) a cover-slide for 1 min so that the agarose solution will spread nicely on it. Use the cover-slides within the next 15 min as the effect of plasma cleaning wears off quickly.
3. Apply 200 µl of warm agarose solution to each 22 x 22 mm² slide so the solution wets the entire surface. Tilt the slide vertically and touch the lower edge to a tissue to remove excess liquid and leave just a thin smooth layer of agarose on the slide.
4. Place the slide on a hot plate or oven at 60 °C and leave it to dry for at least 30 min. The agarose film is hardly visible by eye. After the slides cool to RT, use them immediately, or store them for up to one week in a closed container at 4 °C.
5. Place the agarose-coated coverslip in a standard 3.5 cm Petri dish.
6. Prepare the SUV solution as in Section 3.2 and apply ~15 µl of the SUV solution (3 mg/ml lipid) in ~30 very small drops gently onto the agarose surface. Take care not to distort the agarose layer too much.
7. Place the slide under a gentle stream of nitrogen for about 10–15 min and follow the evaporation of the buffer by eye as the droplets dry.
8. As soon as the SUVs have dried, add growth buffer to cover the slide surface. For a small 3.5 cm Petri dish use ~1 ml of buffer.
9. Allow the swelling to proceed for ~30 min, and then examine the growth of GUVs in the chamber using an inverted microscope with phase-contrast or Differential Interference Contrast (DIC).
   NOTE: Epifluorescence observation is difficult due to the strong background of fluorophores in the gel and the auto-fluorescence of the agarose.

5. Harvesting and Observing GUVs

1. Passivate the observation chamber (e.g., small Petri dish or glass coverslip) so that GUVs do not stick, spread and burst on the chamber bottom. Cover the chamber bottom with beta-casein solution, incubate for 5 min, rinse out the casein solution with pure water, dry with a stream of air or nitrogen, and finally add observation buffer (e.g., ~5 mm depth for a small Petri dish).
2. Harvest the GUVs. Cut the end of a 100 µl pipette tips so the opening is larger (~2 mm diameter), and aspire slowly as the shear stress of pipetting can easily destroy GUVs.
   1. For electro-formed GUVs, open the growth chamber by gently removing the top coverslip. Place the pipette tip directly above each wire and aspirate ~50 µl while moving the pipette tip along the wire to detach the GUVs.
      NOTE: It may help to rotate the wire to collect GUVs on the “other side” of the wire.
   2. For “gel-assisted swelling” GUVs, first tap the side of the petri dish a few times to help the GUVs detach from the coverslip surface. Position the pipette tip just above the coverslip and aspirate 50 µl while pulling the tip back over the surface. Directly transfer harvested GUVs to an observation chamber, or store in a 1.5 ml microcentrifuge tube at 4 °C for up to 1 week.
3. Place the observation chamber on an inverted microscope, add the GUVs to the observation chamber, and wait a few minutes for the GUVs to settle at the chamber bottom.
4. Survey the chamber with phase contrast or DIC to quickly locate smooth, spherical (‘defect free’) “GUV candidates”. Examine each “GUV candidate” in epifluorescence to exclude any containing smaller liposomes nested inside. Finally, check that the lipid fluorescence intensity is uniform and compatible with a single membrane (i.e., unilamellar).
   NOTE: In some bilamellar (or multi-lamellar) vesicles, the membranes are too close together to be resolved so they appear unilamellar in phase contrast or DIC images. However, these objects can be distinguished from actual unilamellar GUVs by their lipid fluorescence, which is two times (or more) brighter.

6. Patch-clamping GUVs

1. Make patch pipettes with a 1–2 µm tip diameter from standard borosilicate capillary glass using the program recommended for the pipette puller.
   NOTE: Special treatments such as fire polishing are not necessary, and pipettes can be used for several days after they have been pulled if they are kept in a closed box.
2. Passivate the chamber by incubating with a beta-casein solution (5 mg/ml) to ensure that GUVs do not adhere, spread and rupture on chamber surfaces. Rinse the casein off after 5 min.
3. Insert the ground electrode, fill the chamber with observation buffer, transfer 10 µl of the GUV suspension as described in step 5.2 and 5.3, and wait a few minutes for the GUVs to settle at the bottom.
4. Fill a fresh patch pipette with solution (observation buffer or another iso-osmotic solution) and mount it on the patch-clamp amplifier headstage.
5. Search through the chamber to locate a “defect-free” GUV as described in section 5.4, and check that it contains fluorescent protein.
6. Apply a constant positive pressure (> 100 Pa, or roughly 1 cm H₂O in a manometer) to keep the patch pipette interior clean, and insert the patch pipette into the chamber. Bring the patch pipette into the field of view, apply test pulses to measure/compensate the pipette voltage offset and resistance, and examine the pipette under fluorescent illumination to confirm that the tip is clean.
7. Bring the patch pipette towards the GUV, and if necessary, simultaneously reduce the positive pressure so the outward flow from the patch pipette does not make the GUV “run away”. When the patch pipette is close to the GUV, apply a negative pressure (up to 5 cm H₂O) to pull the GUV against the patch pipette. Monitor the resistance as the “tongue” of GUV membrane enters the patch pipette and the gigaseal forms.
8. If a gigaseal did not form, remove the patch pipette from the chamber and return to step 6.4. If the membrane patch formed a gigaseal, but the GUV remains attached to the pipette, excise the patch by pulling away from the GUV, bursting the GUV against the chamber bottom, or briefly moving the pipette out of solution.
9. When the inside-out membrane patch has been excised from the GUV and the gigaseal is stable, switch off the test pulses and apply a voltage protocol such as the one shown in Figure 13.
   NOTE: Figure 13 follows the standard electrophysiological convention for an inside-out patch in which current flowing into the patch-electrode is “positive”, and V = V_bath– V_pipette. Holding the patch at a negative potential (e.g., V = -100 mV) for ~30 sec places KvAP in the resting state, while steps (100 msec to 5 sec) to more positive potentials (e.g., V = 100 mV) can then drive it into conducting (i.e., open) active states.
10. After measurements on a membrane patch are finished, break the patch with a zap or pressure pulse and check that the voltage offset of the patch electrode has not drifted. Remove the patch pipette from the chamber, and return to step 6.4.

Results

The growth of GUVs can be quickly evaluated by examining the growth chamber under the microscope. For electroformation, the GUVs tend to grow in bunches along the platinum wires, as shown in Figure 4. During gel-assisted swelling, GUVs appear as spherical structures that rapidly grow and fuse together (Figure 5).

Defect-free GUVs are more easily identified and evaluated after transferring to an observation chamber. Calibration measurements are needed to rigoro...

Discussion

Biomimetic model systems are an important tool for studying the properties and interactions of proteins and membranes. Compared to other reconstituted systems like BLMs or supported lipid membranes, GUV based systems present several opportunities including considerable control of membrane composition, tension and geometry, as well as being truly oil-free. However, incorporating transmembrane proteins, such as KvAP, into GUVs requires significant adaptations of conventional protocols for lipid-only GUVs. The electroformat...

Materials

Name	Company	Catalog Number	Comments
Name of the Material/Equipment	Company	Catalog Number	Comments/ Description
DPhPC (1,2-diphytanoyl-sn-glycero-3-phosphocholine)	Avanti Polar Lipids	850356P
Egg PC L-α-phosphatidylcholine (Egg, Chicken)	Avanti Polar Lipids	840051P
Egg PA L-α-phosphatidic acid (Egg, Chicken)	Avanti Polar Lipids	840101P
18:1 (Δ9-Cis) PC (DOPC) 1,2-dioleoyl-sn-glycero-3-phosphocholine	Avanti Polar Lipids	850375P
cholesterol (ovine wool, >98%)	Avanti Polar Lipids	700000P
TRed-DHPE	Invirtogen	T-1395MP	labeled lipid
BPTR-Ceramide	Invirtogen	D-7540	labeled lipid
Choloroform	VWR	22711.290	AnalaR Normapur
Acetone	VWR	20066.296	AnalaR Normapur
Ethanol	VWR	20821.330	AnalaR Normapur
Kimwipe	Kimtech	7552
Hamilton syringes	Hamilton Bonaduz AG		diverse
Amber vials and teflon cups	Sigma	SU860083 and SU860076
Parafilm	VWR	291-1214
microcentrifuge tube	eppendorf		diverse
Agarose	Euromex	LM3 (1670-B, Tg 25.7C, Tm 64C)
Sucrose	Sigma	84097-1kg
Glucose	Sigma	G8270-1kg
Trehalose	Sigma	T9531-25G
KCl	Sigma	P9333-1kg
Hepes	Sigma	H3375-100G
TRIS	Euromex	EU0011-A
Osmometer	Wescor	Vapro
pH meter	Schott instruments	Lab850
Sonicator	Elmasonic	S 180 H
Syringe filter 0.2µm	Sartorius steim	Minisart 16532
Function generator	TTi	TG315
Platinum wires	Goodfellow	LS413074	99.99+%, d=0.5mm
Polytetrafluoroethylene	Goodfellow
Dow Corning 'high vacuum grease'	VWR	1597418
sealing paste	Vitrex medical A/S, Denmark	REF 140014	Sigillum Wax
Cover slides 22x40 mm No1,5	VWR	631-0136
Cover slides 22x22 mm No1,5	VWR	631-0125
plasma cleaner	Harrick	PDC-32G	airplasma, setting 'high'
petri dishes	Falcon BD	REF 353001	3.5 cmx1cm
patch clamp amplifier	Axon instruments	Multiclamp700B
DAQ Card	National Instruments	PCI-6221
Labview	National Instruments	version 8.6
Glass pipettes boro silicate OD 1mm ID 0.58mm	Harvard Apparatus	GC100-15
Electrode holder	Warner Instruments	Q45W-T10P
Micromanipulator	Sutter	MP-285
pipette puller	Sutter	P-2000
Camera	ProSilica	GC1380
Zeiss microscope	Zeiss	Axiovert 135
Objective	Zeiss		40x long working distance, Phase contrast
Objective	Zeiss		100x Plan-Apochromat NA 1.3
Filterset GFP (for Alexa-488)	Horiba	XF100-3
Filterset Cy3 (for TexasRed)	Horiba	XF101-2
beta-casein	Sigma	C6905-1G
confocal microscope	Nikon Eclipse TE 2000-E		D-Eclipse C1 confocal head
Objective	Nikon		Plan Fluor 100× NA1.3
matlab	Mathworks		for image processing and analysis of the current traces