U.S. patent application number 12/733275 was filed with the patent office on 2011-02-24 for bilayers.
Invention is credited to Mark Ian Wallace.
Application Number | 20110041978 12/733275 |
Document ID | / |
Family ID | 38566709 |
Filed Date | 2011-02-24 |
United States Patent
Application |
20110041978 |
Kind Code |
A1 |
Wallace; Mark Ian |
February 24, 2011 |
BILAYERS
Abstract
A method for producing a bilayer of amphipathic molecules
comprising providing a hydrated support and providing a hydrophilic
body, and bringing the hydrated support and hydrophilic body into
contact to form a bilayer of amphipathic molecules. A bilayer
produced by the method of the invention, and uses of the
bilayer.
Inventors: |
Wallace; Mark Ian; (Oxford,
GB) |
Correspondence
Address: |
Jeffrey J. King, Esq.;Patent Networks Law Group PLLC
5000 Carillon Point, Suite 400
Kirkland
WA
98033
US
|
Family ID: |
38566709 |
Appl. No.: |
12/733275 |
Filed: |
August 19, 2008 |
PCT Filed: |
August 19, 2008 |
PCT NO: |
PCT/GB2008/002805 |
371 Date: |
November 4, 2010 |
Current U.S.
Class: |
156/60 |
Current CPC
Class: |
G01N 33/5432 20130101;
G01N 33/6872 20130101; Y10T 156/10 20150115; G01N 33/68
20130101 |
Class at
Publication: |
156/60 |
International
Class: |
B32B 37/14 20060101
B32B037/14 |
Foreign Application Data
Date |
Code |
Application Number |
Aug 21, 2007 |
GB |
0716264.7 |
Claims
1. A method for producing a bilayer of amphipathic molecules
comprising the steps of: (i) providing a hydrated support in a
hydrophobic medium, wherein a first monolayer of amphipathic
molecules is present on the surface of the hydrated support; (ii)
providing a hydrophilic body in a hydrophobic medium, wherein a
second monolayer of amphipathic molecules is present on the surface
of the hydrophilic body; and (iii) bringing the first monolayer and
the second monolayer into contact to form a bilayer of amphipathic,
wherein either: (a) the hydrophobic medium in which the hydrated
support is provided contains amphipathic molecules and the
hydrophobic medium in which the hydrophilic body is provided
contains amphipathic molecules; or (b) the hydrated support
contains amphipathic molecules and the hydrophilic body contains
amphipathic molecules.
2-7. (canceled)
8. The method of claim 1 wherein the amphipathic molecules are
lipid molecules.
9. The method of claim 8 wherein the lipid molecules are selected
from the group comprising fatty acyls, glycerolipids,
glycerophospholipids, sphingolipids, sterol lipids, prenol lipids,
saccharolipids, polyketides, phospholipids, glycolipids and
cholesterol.
10. The method of claim 8 wherein the lipid molecules are selected
from the group comprising monoolein;
1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC);
1,2-diphytanoyl-sn-glycero-3-phosphatidylcholine (DPhPC); palmitoyl
oleoyl phosphatidylcholine (POPC);
1-palmitoyl-2-oleoyl-phosphatidylethanolamine (POPE);
1-palmitoyl-2-oleoyl-phosphatidylethanolamine; and
1-palmitoyl-2-oleoylphosphatidylglycerol (POPE/POPG) mixtures; or
mixtures thereof.
11. (canceled)
12. The method of claim 1 wherein the hydrated support comprises a
solid or a semi-solid substrate.
13. The method of claim 1 wherein the hydrated support is
hydrophilic.
14. (canceled)
15. The method of claim 1 wherein the hydrated support is selected
from the group comprising hydrogels, agarose, polyacrylamide,
[cross-linked] polyethylene glycol, nitro-cellulose, polycarbonate,
anodisc material, polyethersulphone, cellulose acetate, nylon,
Naphion materials, mesoporous silica, water and glass.
16. The method of claim 1 wherein the hydrated support is a protein
or analyte separation gel.
17. (canceled)
18. The method of claim 1 wherein the hydrophilic body comprises a
droplet of aqueous solution.
19. The method of claim 18 wherein the droplet is from 5 nm to 10
cm in diameter.
20. The method of claim 1 wherein the hydrophilic body comprises a
hydrated solid or semi-solid support/substrate.
21. The method of claim 1 wherein the hydrophobic medium is an
oil.
22. The method of claim 21 wherein the oil is a hydrocarbon.
23. The method or bilayer of claim 21 wherein the oil is selected
from the group comprising alkanes, alkenes, fluorinated oils,
silicone based oils and carbon tetrachloride.
24. The method of claim 1 wherein the bilayer has a diameter from
about 1 .mu.m to greater than about 1 cm.
25. (canceled)
26. The method of claim 1 wherein a membrane-associated protein is
present in at least one of the hydrated support, the hydrophilic
body and the hydrophobic medium, said membrane-associated protein
being capable of insertion into the bilayer.
27. (canceled)
28. The method of claim 26 wherein the membrane-associated protein
is selected from the group comprising a selective or non-selective
membrane transport protein, an ion channel, a pore forming protein
and a membrane-resident receptor.
29. The method of claim 26 wherein the one or more protein is
inserted into the bilayer after the bilayer has formed.
30. The method of claim 1 wherein the area of the bilayer can be
varied by varying the relative positions of the hydrophilic body
and the hydrated substrate.
31. The method of claim 1 wherein the bilayer can be disassembled
by removing contact between the hydrated support and the
hydrophilic body.
32. The method of claim 31 wherein the bilayer can be reformed by
restoring contact between a lipid monolayer on the hydrated support
and a lipid monolayer on the hydrophilic body.
33-56. (canceled)
Description
[0001] The present invention relates to a bilayer, such as a lipid
bilayer, to a method of producing a bilayer, to the use of a
bilayer and to apparatus to produce and/or use a bilayer.
[0002] Artificial planar lipid bilayers serve as simplified models
of biological membranes and are widely used for the electrical
characterisation of ion-channels and protein pores. Ion-channels
are a diverse group of membrane proteins that selectively control
the movement of specific ions across cell membranes, establishing
voltage and electrochemical gradients that are fundamental to a
wide variety of biological processes. In humans, ion-channels
regulate everything from heartbeat and muscle contraction to
hormone secretion and the release of neurotransmitters. Defective
ion channel function is implicated in a growing list of disorders,
including cardiac arrhythmia, periodic paralysis, epilepsy and
diabetes (Ashcroft, F. M. 2000, Academic Press, San Diego;
Ashcroft, F. M. 2006, Nature 440, 440-447; Kass, R. S. 2005,
Journal of Clinical Investigation 115, 1986-1989). Protein pores
are non-specific channels that allow molecules to pass across cell
membranes. Protein pores can be exploited for many applications
such as molecular sensing (Bayley, H. et al., 2000, Advanced
Materials 12, 139-142; Bayley, H. & Cremer, P. S. 2001. Nature
413, 226-230) and DNA sequencing (Kasianowicz, J. J. et al., 1996.
Proceedings of the National Academy of Sciences of the United
States of America 93, 13770-13773; Howorka, S. et al., 2001. Nature
Biotechnology 19, 636-639; Astier, Y. 2006. Journal of the American
Chemical Society 128, 1705-1710).
[0003] Single-channel recording (SCR) of individual proteins is a
powerful means of studying channel protein function (Sakmann, B.
& Neher, E. 1995. Plenum Press, New York; London).
Single-channel recording measures changes in ion-current through
single protein channels, and can examine voltage dependence, gating
behaviour, ligand binding affinity, and ion selectivity at the
single-molecule level. Consequently, single-channel recording can
help determine the molecular basis of an ion-channel disease. It is
also an important technique for the development of new drugs
specifically targeting channelopathies, and for screening other
medicines for unwanted side-effects (Ashcroft, F. M. 2006. Nature
440, 440-447; Roden, D. M. 2004. New England Journal of Medicine
350, 1013-1022). Advances in these areas require much higher
throughput assays of ion-channel behaviour than are currently
available.
[0004] Single-channel recording typically uses either
patch-clamping (Sakmann, B. & Neher, E. 1984. Annual Review of
Physiology 46, 455-472) or artificial planar lipid bilayers
(Mueller, P. et al., 1962. Nature 194, 979-980; White, S. H. 1986.
ed. Miller, C. Plenum Press: New York). Although other methods may
also be used, including excised-patch, tip-dip and on-chip
methods.
[0005] Patch-clamping of whole cells is a versatile and sensitive
means of examining channels, but is time-consuming and often
complicated by the heterogeneous nature of cell membranes. In
contrast, artificial planar lipid bilayers control the constituents
of the system and can be used to study purified proteins. Planar
lipid bilayers are usually formed either by painting, where a
solution of lipid in an organic solvent is directly applied to an
aperture separating two aqueous compartments (Mueller, P. et al.,
1962. Nature 194, 979-980; White, S. H. 1986. ed. Miller, C. Plenum
Press: New York), or variants of the Langmuir-Blodgett technique,
where two air/water monolayers are raised past an aperture (Montal,
M. & Mueller, P. 1972. Proceedings of the National Academy of
Sciences of the United States of America 69, 3561-3566). Although
widely used, planar lipid bilayers are difficult to prepare, and
their short lifetime prohibits their use in many situations.
[0006] Alternative emulsion-based approaches to forming bilayers
have also been proposed (Tsofina, L. M. et al., 1966. Nature 212,
681-683), where bilayers are created between aqueous surfaces
immersed in a solution of lipid in oil. When immersed in an
immiscible lipid/oil solution, aqueous surfaces spontaneously
self-assemble a lipid monolayer (Cevc, G. 1993. Phospholipids
handbook, ed. Cevc, G., Marcel Dekker, New York); Seddon, J. M.
& Templer, R. H. 1995. eds. Lipowsky, R. & Sackmann, E.,
Elsevier, Amsterdam, Oxford), and when monolayers from two aqueous
components are brought into contact they can `zip` together to form
a lipid bilayer (Tien, H. T. 1974. M. Dekker, New York; Fujiwara,
H. et al., 2003. Journal of Chemical Physics 119, 6768-6775).
Recent studies have shown that microfluidic flows (Malmstadt, N. et
al., 2006. Nano Letters 6, 1961-1965; Funakoshi, K. et al., 2006.
Analytical Chemistry 78, 8169-8174) and droplets (Funakoshi, K. et
al., 2006. Analytical Chemistry 78, 8169-8174; Holden, M. A. et
al., 2007. Journal of the American Chemical Society p8650-5) can be
contacted in a lipid/oil solution to create bilayers suitable for
single-channel recording experiments.
[0007] According to a first aspect of the invention there is
provided a method for producing a bilayer of amphipathic molecules
comprising the steps of: [0008] (i) providing a hydrated support in
a hydrophobic medium, wherein the hydrophobic medium contains
amphipathic molecules and a first monolayer of amphipathic
molecules is present on the surface of the hydrated support; [0009]
(ii) providing a hydrophilic body in a hydrophobic medium, wherein
the hydrophobic medium contains amphipathic molecules and a second
monolayer of amphipathic molecules is present on the surface of the
hydrophilic body; and [0010] (iii) bringing the first monolayer and
the second monolayer into contact to form a bilayer of amphipathic
molecules.
[0011] Preferably step (i) of the first method of the invention
comprises providing a hydrated support in a hydrophobic medium,
wherein the hydrophobic medium contains amphipathic molecules, and
then forming a first monolayer of amphipathic molecules on the
surface of the hydrated support.
[0012] Preferably step (ii) of the first method of the invention
comprises providing a hydrophilic body in a hydrophobic medium,
wherein the hydrophobic medium contains amphipathic molecules, and
then forming a second monolayer of amphipathic molecules on the
surface of the hydrophilic body.
[0013] According to a second aspect of the invention there is
provided a method for producing a bilayer of amphipathic molecules
comprising the steps of: [0014] (i) providing a hydrated support
containing amphipathic molecules in a hydrophobic medium, wherein a
first monolayer of amphipathic molecules is present on the surface
of the hydrated support; [0015] (ii) providing a hydrophilic body
containing amphipathic molecules in a hydrophobic medium, wherein a
second monolayer of amphipathic molecules is present on the surface
of the hydrophilic body; and [0016] (iii) bringing the first
monolayer and the second monolayer into contact to form a bilayer
of amphipathic molecules.
[0017] Preferably step (i) of the second method of the invention
comprises providing a hydrated support containing amphipathic
molecules in a hydrophobic medium, and then forming a first
monolayer of amphipathic molecules on the surface of the hydrated
support.
[0018] Preferably step (ii) of the second method of the invention
comprises providing a hydrophilic, body containing amphipathic
molecules in a hydrophobic medium, and then forming a second
monolayer of amphipathic molecules on the surface of the
hydrophilic body.
[0019] It is found that the method of both the first and the second
aspect of the invention spontaneously forms a bilayer of
amphipathic molecules which has the added advantage that it is
stable over long periods of time and when subjected to
environmental and/or physical stress.
[0020] The life-time of the bilayer of amphipathic molecules made
according to any method of the invention may be greater than about
1 hour, 5 hours, 10 hours, 24 hours, 2 days, 1 week, 1 month, 2
months, 3 months or more.
[0021] Preferably the method of the invention forms bilayers with
at least about 90% efficiency, more preferably with about 95%, 98%
or 99% efficiency.
[0022] Preferably the bilayers form within 1 minute of contact
between the monolayer on the hydrated support and the monolayer on
hydrophilic body.
[0023] The bilayer of amphipathic molecules may be capable of
withstanding physical shock, for example, the bilayer may be stable
after being dropped one or more times from heights greater than 0.5
metres or 1 metre. This bilayer stability makes bilayers produced
by any method of the invention easier to work with than bilayers
produced by known conventional methods.
[0024] The surprisingly long life-time and stability of the bilayer
of amphipathic molecules produced by any method of the invention
has the benefit that a user is able to set up the bilayer and use
it in long term and/or multiple experiments. It also has the
benefit of being capable of being more readily used in a portable
device, for example outside of a controlled laboratory environment,
where the physical environment is less controllable or predictable,
than conventional bilayers.
[0025] Preferably a monolayer of amphipathic molecules self
assembles on the hydrated support and the hydrophilic body when
each is placed in a hydrophobic medium containing amphipathic
molecules.
[0026] The orientation of the amphipathic molecules in the
monolayers means that a bilayer forms when the monolayers are
brought into contact.
[0027] Amphipathic molecules have both a hydrophilic group and a
hydrophobic group. In the monolayers, formed in any method of the
invention, the amphipathic molecules are aligned on the surface of
the hydrophilic body and the hydrated substrate with the
hydrophilic groups (or "heads") towards the water interface and the
hydrophobic groups (or "tails") away from the water interface.
[0028] The amphipathic molecules used in any method of the
invention may be lipid molecules, in particular, surfactant
molecules may be used. The lipid molecules may be selected from the
group comprising fatty acyls, glycerolipids, glycerophospholipids,
sphingolipids, sterol lipids, prenol lipids, saccharolipids,
polyketides, phospholipids, glycolipids and cholesterol.
[0029] The lipid may include any of the group comprising monoolein;
1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC);
1,2-diphytanoyl-sn-glycero-3-phosphatidylcholine (DPhPC); palmitoyl
oleoyl phosphatidylcholine (POPC);
1-palmitoyl-2-oleoyl-phosphatidylethanolamine (POPE);
1-palmitoyl-2-oleoyl-phosphatidylethanolamine; and
1-palmitoyl-2-oleoylphosphatidylglycerol (POPE/POPG) mixtures; or
mixtures thereof.
[0030] The amphipathic molecules in the monolayers or bilayers may
be of the same or different types. For example, each monolayer of
the bilayer may comprise a different type of amphipathic molecule
such that the bilayer produced is asymmetric. An asymmetric bilayer
may be produced using the method of the first aspect of the
invention wherein the first monolayer is formed on the hydrated
support in a first hydrophobic medium containing a first type of
amphipathic molecule. The second monolayer is formed on the
hydrophilic body in a second hydrophobic medium containing a second
type of amphipathic molecule. The first and second amphipathic
molecules may be different. The two monolayers, one on the
hydrophilic body and one on the hydrated support, are then brought
together in a third hydrophobic medium which does not contain an
amphipathic molecule to form an asymmetric bilayer of amphipathic
molecules. Alternatively, an asymmetric bilayer may be produced by
the second method of the invention by using a different type of
amphipathic molecule in the hydrated support to that used in the
hydrophilic body.
[0031] Alternatively, each monolayer may comprise the same type or
mixtures of types of amphipathic molecules.
[0032] The hydrated support may comprise a solid or a semi-solid
substrate. The terms "solid" and "semi-solid" as used herein are
understood to have their ordinary meaning to a person skilled in
the art. Essentially the term "solid" refers to a substrate that is
rigid and resistant to deformation, and "semi-solid" refers to a
substrate that has properties between those of a solid and a
liquid. Preferably a semi-solid substrate has some degree of
flexibility but is rigid enough to maintain its shape when placed
in a container, and will not immediately conform to the shape of
the container. An example of a semi-solid substrate is a gel.
[0033] Preferably the hydrated support is hydrophilic. Preferably a
monolayer of amphipathic molecules will self assemble on the
surface of the hydrated support if it is placed in the presence of
amphipathic molecules in a hydrophobic medium.
[0034] Alternatively, the hydrated support may not be hydrophilic
and the lipid monolayer may be formed by the attachment of lipid
molecules to the surface, for example, the surface of the support
may be such that modified lipids will react with the surface and
attach to form a monolayer.
[0035] The hydrated support may be porous or non-porous. Preferably
the hydrated support is porous.
[0036] The hydrated support may be a hydrogel. The hydrogel may be
photocrosslinked.
[0037] The hydrated support may comprise agarose, polyacrylamide,
[cross-linked] polyethylene glycol, nitro-cellulose, polycarbonate,
anodisc material, polyethersulphone, cellulose acetate, nylon,
Naphion materials, mesoporous silica, water and/or glass.
[0038] The hydrated support may be a protein or analyte separation
gel, for example, an electrophoresis gel. The separation gel may
contain proteins, DNA or other samples separated, for example, on
the basis of their size, molecular weight or ionic properties.
[0039] The surface of the hydrated support which carries the
amphipathic molecule monolayer may be any suitable configuration.
The surface may be substantially flat, or the surface may be
substantially uneven. The surface may be curved. The surface may be
patterned.
[0040] The hydrated support may be partially or substantially
transparent. Alternatively, the hydrated support may be largely
opaque.
[0041] The hydrated support may comprise a substrate of any
thickness, preferably from about 1 nm to about 10 cm, more
preferably from about 1 .mu.m to about 1 cm, most preferably from
about 100 .mu.m to about 1 cm.
[0042] The hydrophilic body may be a liquid, solid, or semi-solid
or a mixture thereof. Preferably the hydrophilic body comprises a
droplet of aqueous solution, such as water.
[0043] Where the hydrophilic body is a droplet of an aqueous
solution it preferably has a diameter of from about 5 nm to 10 cm
or more, preferably from 1 .mu.m to 1 mm. In one embodiment,
droplets are around 100 .mu.m in diameter.
[0044] The hydrophilic body may comprise a hydrated solid or
semi-solid support/substrate. The hydrophilic body may comprise a
hydrogel, such as hydrated agarose.
[0045] The composition of the hydrophilic body is preferably
controlled to contain the correct salts to allow an electrical
current to be carried, for example, NaCl, KCl, MgCl.sub.2 and/or
other salts may be included.
[0046] The hydrophilic body may also comprise common buffering
agents to control pH, for example, Bis-tris, Tris, Hepes, sodium
phosphate and/or potassium phosphate.
[0047] Salts may also be included for other reasons, for example,
to stabilise proteins, to control binding components, to control
the osmotic gradient across the bilayer and/or to activate
fluorescent probes.
[0048] The hydrophilic body may also contain varying amounts of
other components, such as, sucrose or PEG which may be used to
stabilise osmotic stresses, fluorescent probes, microspheres or
beads. The hydrophilic bodies may also comprise denaturants such as
urea or guanidine HCl.
[0049] Where more than one hydrophilic body is used each may be of
the same or a different composition.
[0050] The hydrophobic medium may be an oil. The oil may be a
hydrocarbon, which may be branched or unbranched, and may be
substituted or unsubstituted. For example, the hydrocarbon may have
from 5 to 20 carbon atoms, more preferably from 10 to 17 carbon
atoms. Suitable oils include alkanes or alkenes, such as
hexadecane, decane, pentane or squalene, or fluorinated oils, or
silicone based oils, or carbon tetrachloride.
[0051] Preferably, the method of the invention uses a lipid
(amphipathic molecules) in oil (hydrophobic medium) solution.
Preferably the lipid in oil solution contains from about 1 mg/ml to
about 30 mg/ml of lipid in the oil. Preferably, the lipid in oil
solution contains about 5 mg/ml of lipid. Preferably the lipid/oil
solution comprises 1,2-diphytanoyl-sn-glycero-3-phophocholine
(DPhPC) in n-hexadecane (C.sub.16).
[0052] The terms "contacting" or "contact" used herein with
reference to the contacting of monolayers to form a bilayer are
understood to mean actual physical contact, and/or close enough
proximity, to allow the assembly of an amphipathic molecule bilayer
from separate amphipathic molecule monolayers. Preferably the
contacting process is a form of Gibbs-plateau border action.
[0053] The bilayer of amphipathic molecules may be from less than
about 1 .mu.m to greater than about 1 cm in diameter, preferably
from about 5 .mu.M to about 5000 .mu.m in diameter, more preferably
from about 30 .mu.m to about 3000 .mu.m in diameter. The bilayer
may be from about 5 .mu.m to about 500 .mu.m in diameter, more
preferably from about 30 .mu.m to about 300 .mu.m in diameter. The
skilled man will appreciate that the bilayer does not need to be
circular. Where a non-circular bilayer is formed preferably the
bilayer includes a portion which has the aforementioned preferred
diameter. For example, a non-circular bilayer according to the
invention may include a portion which has diameter from about 1
.mu.m to about 1 cm or more. Preferably, a non-circular bilayer
according to the invention has one dimension which is from about 1
.mu.m to about 1 cm or more.
[0054] The area of the bilayer of amphipathic molecules may be
adjustable. Preferably the area of the bilayer layer is adjustable
before, during, or in intervals between uses of the bilayer. The
area of the bilayer may be adjustable by increasing or decreasing
the contact area between the hydrophilic body and the hydrated
support. The contact area between the hydrophilic body and the
hydrated support may be adjusted by moving the centre of the
hydrophilic body towards or away from the hydrated support, or by
moving the hydrated support towards or away from the centre of the
hydrophilic body. Preferably to increase the area of the bilayer,
the centre of the hydrophilic body is moved towards the hydrated
support. Preferably to decrease the area of the bilayer, the centre
of the hydrophilic body is moved away from the hydrated support.
Where an electrode is used to hold the hydrophilic body, the area
of the bilayer may be increased or decreased by moving the
electrode towards or away from the hydrated support respectively.
In an alternative embodiment the position of the hydrophilic body
may be controlled by using an applied electric or magnetic field,
by using light beams (such as optical traps) or pressure. The area
of the bilayer may be adjusted by increasing or decreasing the
volume, size and/or shape of the hydrophilic body. Preferably the
area of the bilayer can be changed without breaking the
bilayer.
[0055] Preferably the area of the bilayer can be moved to any
diameter between 5 .mu.m and about 500 .mu.m, or an up to 100 fold
change in bilayer size can be achieved, in less than about 10
seconds, less than about 5 seconds, less than about 3 second, less
than about 2 seconds, less than about 1 second or less than about
0.5 seconds.
[0056] Controlling the area of the bilayer is surprisingly easy and
fast. Controlling the area of the bilayer has the advantage of
being able to control the number/amount of membrane proteins that
associate with the bilayer. Surprisingly, altering the area of the
bilayer does not denature or remove membrane-associated proteins
from the bilayer. Indeed, if trans-membrane proteins are already
inserted into the bilayer they will become concentrated if the area
of the bilayer is decreased. If contact between the hydrophilic
body and the hydrated support is removed, the bilayer will
disassemble and any trans-membrane protein located therein will be
no longer present in the bilayer or either monolayer.
[0057] The method may include a stabilisation period to allow the
monolayer and/or bilayer of amphipathic molecules to form. The
stabilisation period may be to allow the system to reach
equilibrium. The stabilisation period may be from about 0 seconds
to about 5 hours, preferably the stabilisation period is from about
10 seconds to about 1 hour. Preferably the stabilisation period is
about 15 minutes. In some embodiments if one component of the
method, for example either the first or second monolayer, has been
left to stabilise after formation, then there may be no need to
allow the other monolayer to stabilise before formation of the
bilayer.
[0058] The stabilisation period has the advantage of reducing the
chances of the hydrophilic body and the hydrated support coalescing
without bilayer formation.
[0059] The bilayer may be visualised through the hydrated support
with an inverted microscope or in some circumstances even by the
naked eye. The visualisation of the lipid bilayer may be used to
track the formation, position, size, or other property of the
bilayer. Visualisation of the bilayer allows labelled
analytes/proteins/compounds at or in the bilayer to be seen and
studied.
[0060] The bilayer of amphipathic molecules may be used to study
processes occurring at, in or through the bilayer. The bilayer can
be used as an artificial/model system in which to study cell
membrane behaviour.
[0061] Proteins may be inserted into the bilayer of amphipathic
molecules.
[0062] Proteins in the environment of the bilayer, for example in
the hydrophobic medium and/or in the hydrophilic body and/or in the
hydrated support, may insert spontaneously into the bilayer.
Alternatively proteins may be driven into the bilayer by the
application of a voltage and/or by fusion of protein loaded
vesicles with the bilayer. The vesicles may be contained within or
introduced to the hydrophilic body. Proteins may be introduced into
the membrane by using the probe method disclosed in GB0614835.7.
Proteins may insert into the bilayer in the same manner as if the
bilayer was formed by known techniques.
[0063] The inserted protein may be a known membrane-associated
protein.
[0064] The protein may be a membrane-associated protein which is
anchored directly or indirectly to the bilayer. The protein may be
a selective or non-selective membrane transport protein, an ion
channel, a pore forming protein or a membrane-resident
receptor.
[0065] Membrane-associated proteins which may associate with and/or
insert into the bilayer include any of the group comprising
peptides, e.g. gramicidin; .alpha.-helix bundles, e.g.
bacteriorhodopsin or K.sup.+ channels; and .beta.-barrels, e.g.
.alpha.-hemolysin, leukocidin or E. coli porins; or combinations
thereof.
[0066] The bilayer of amphipathic molecules may be used to detect
compounds/analytes which are capable of interaction with
amphipathic molecules in the bilayer or with a membrane-associated
protein in the bilayer. The interaction with the
membrane-associated protein or the amphipathic molecules may be by
the specific or non-specific translocation of the
analytes/compounds across the bilayer, this may be mediated by the
membrane-associated protein or by the amphipathic molecules.
Alternatively compounds/analytes may interact with a trans-membrane
protein or with the lipid bilayer to cause physical, optical,
electrical, or biochemical changes. Such interaction may be
detected in many different ways, including, but limited to, by
visual changes, changes in specific capacitance, or by the
activation of fluorescently labelled lipids or proteins in the
bilayer.
[0067] The bilayer may be used to detect membrane-associated
proteins. Preferably the membrane-associated proteins are ion
channel proteins and/or pore forming proteins. Preferably the
membrane-associated proteins diffuse into and/or associate with the
bilayer causing a detectable change in the properties at the
bilayer. The properties changed may be physical, optical,
electrical or biochemical.
[0068] Bilayers of amphipathic molecules made by any method of the
invention may be used to investigate and/or screen
membrane-associated proteins; to investigate and/or screen for
analytes that interact with membrane-associated proteins; and to
investigate and/or screen for compounds that interact with bilayers
made of different amphipathic molecules. Bilayers of different
amphipathic molecules may be screened to study the role of and/or
the interaction of different amphipathic molecules with various
analytes/compounds.
[0069] The bilayer may be used to study the voltage dependence
properties of a membrane-associated protein inserted in the
bilayer. For example, a bilayer composed of DPhPC may have a
specific capacitance between about 0.3 and about 0.9 .mu.F
cm.sup.-2 at 22.degree. C., preferably about 0.65 .mu.F cm.sup.-2
at 22.degree. C. By studying changes in the specific capacitance
properties the changes in the properties of the bilayer caused by
different conditions can be studied. For example, by studying the
ionic current crossing a bilayer the properties of an ion permeable
transmembrane-associated protein may be studied, as may the
interaction of the transmembrane-associated protein with different
analytes/compounds.
[0070] The bilayer may be used to study the ability of a
membrane-associated protein inserted in the bilayer to transport
molecules across the membrane. For example, the amount of a
molecule being transported across a membrane, either through a
protein or simply non-mediated transport across the bilayer, may be
determined by using voltage studies, mass spectrometry, enzymatic
techniques, such as ELISA, or by using a fluorescently or
radioactively labelled substrate.
[0071] A bilayer produced by any method of the invention may also
be used to study the effects of mechanical changes of the bilayer
on proteins in the bilayer or on the bilayer itself. Mechanical
changes which can be studied include, for example, changes in
membrane curvature, lateral forces, surface tension etc.
[0072] A compound/analyte to be tested, studied and/or used in a
screen may be introduced into the system by placing it in the
hydrophilic body and/or in the hydrated support and/or in the
hydrophobic medium. If included in the hydrophilic body the
compound/analyte may be incorporated when the body is formed or it
may be added later, for example, by injection into the formed body.
Similarly, if the compound/analyte is in the hydrated support it
may be incorporated when the support is formed or added later.
[0073] In one embodiment the hydrated support may be a protein
separation gel or membrane containing proteins (analytes/compounds)
for analysis. For example the sample may be a polyacrylamide gel
containing proteins which have been separated on the basis of size.
In this embodiment the analytes/compounds are introduced to the
bilayer via the hydrated support.
[0074] The analyte/compound may be a purified protein or a crude
protein extract.
[0075] The analytes/compounds to be tested may be in a sample. The
sample may be an environmental sample, for example from a body of
water such as a river or reservoir, or the sample may be a
biological sample, for example a sample of blood, urine, serum,
saliva, cells or tissue. The sample may be a liquid from a cell
growth medium.
[0076] One or more detection means may be used to detect chemical,
biochemical, electrical, optical, physical and/or environmental
properties of the bilayer of amphipathic molecules or of
membrane-associated proteins inserted into the bilayer. In
particular the one or more detection means may be used to detect
changes in or at the bilayer induced by the compounds/analytes.
[0077] The detection means may comprise electrodes which may be
used to detect changes in ionic current passing through a protein
channel inserted into a bilayer or the electrochemical properties
of molecules in the hydrated support, hydrophilic body or
bilayer.
[0078] Chemical or biochemical changes may be detected using
enzymatic assays or immunoassays. Alternatively, the use of
labelled, for example radio or fluorescently labelled, proteins
which are activated under certain conditions can be used to monitor
changes at the bilayer. Colormetric methods that respond to changes
in light absorption upon reaction may also be used to detect
changes in the bilayer, in particular this method may be used to
detect a change in the size of the bilayer.
[0079] The detection means may be capable of constantly or
intermittently detecting properties of, or changes at, the
bilayer.
[0080] Detection reagents, like membrane-associated proteins,
analytes and other compounds may be delivered to the bilayer by
incorporation into the hydrophilic body and/or the hydrated
support, injection directly into the hydrophilic body and/or the
hydrated support, and/or incorporation in, or addition to, the
hydrophobic medium. Injection into the hydrophilic body and/or the
hydrated support may be achieved using a micropipette.
[0081] In an embodiment where changes in membrane capacitance are
being studied electrodes may be used as the detection means. The
electrodes may be Ag/AgCl, such electrodes may be from
approximately 10 microns to 1 mm in diameter. A first electrode may
be electrically contacted with the hydrophilic body and a second
electrode may be electrically contacted with the hydrated support.
Electrical properties of the bilayer, such as the specific
capacitance of the bilayer, may be determined using the
electrodes.
[0082] A micromanipulator may be used to insert electrodes into the
hydrophilic body and/or the hydrated support.
[0083] The physical, chemical or electrical environment of the
bilayer may be controlled by the introduction, removal, or
sequestering of reagents, analytes, compounds and/or proteins to or
from the bilayer and/or hydrophilic body and/or hydrated support,
e.g. the pH of the environment surrounding the bilayer may be
controlled by the addition of a buffer to the hydrophilic aqueous
body and/or the hydrated support.
[0084] The bilayer may be repeatedly reformed, by removing contact
between the hydrophilic body and the hydrated support and then
re-establishing contact to re-form the bilayer. The bilayer may be
disassembled by increasing the distance between the centre of the
hydrophilic body and the hydrated support to a point where the
bilayer would become unstable and spontaneously disassemble.
[0085] Membrane-associated proteins in the bilayer may be removed
from association with the bilayer by disassembling the bilayer for
example by removing contact between the hydrophilic body and the
hydrated support. Once the bilayer has been disassembled to remove
the membrane-associated proteins, it may be reformed such that no
membrane-associated proteins, or different membrane-associated
proteins, are associated with the bilayer.
[0086] The concentration of membrane-associated proteins associated
with the bilayer may be increased by decreasing the area of the
bilayer. Conversely, the concentration of membrane-associated
proteins associated with the bilayer may be decreased by increasing
the area of the bilayer.
[0087] The ability to increase the concentration of
membrane-associated proteins in the bilayer may be used to produce
2D crystals of membrane-associated proteins by decreasing the area
of the bilayer comprising the membrane-associated proteins and
hence restricting the area within which a membrane protein can
diffuse.
[0088] The ability to increase or decrease the concentration of
membrane-associated proteins in the bilayer may be used to modulate
protein-protein interactions, for example between sub-units of a
protein or between components of a protein complex.
[0089] Once formed, a bilayer made according to any method of the
invention may be translocated or moved across the surface of the
hydrated support. Preferably this is achieved by moving the
hydrophilic body across the surface of the hydrated support. The
lipid bilayer may be translocated across the surface of the
hydrated support at speeds of about 1 or 2 mm s.sup.-1 or more.
Preferably membrane-associated proteins within the bilayer do not
disassociate from the bilayer when the bilayer is translocated
across the surface of the hydrated support.
[0090] The translocation of the bilayer across the surface of the
hydrated support may be achieved by moving a member contacted with
the hydrophilic body or the hydrated support. Preferably the member
is an electrode. Preferably a micromanipulator is used to move the
member in order to translocate the bilayer by moving either the
hydrophilic body across the hydrated support or the hydrated
support across the hydrophilic body. Alternatively both the
hydrated support and the hydrophilic body may move.
[0091] The translocation of the bilayer may be used to apply forces
to proteins in the bilayer, for example to study mechano-sensitive
protein channels or to study the effect of such force on the
properties of the bilayer itself.
[0092] The translocation of the bilayer may be used to scan across
the surface of the hydrated support to identify analytes/compounds
in the hydrated support which alter properties of the bilayer
and/or membrane-associated proteins in the bilayer.
[0093] The surprising capability of the bilayer to translocate
across the surface of the hydrated support provides the benefit of
being able to scan across the hydrated support with the bilayer to
detect analytes/compounds, such as membrane-associated proteins
and/or reagents or substrates located at different regions of the
same hydrated support. This advantageously can be done without
having to disassemble the bilayer between each sampling region.
[0094] The bilayer may be translocated across a hydrated support
comprising an array or library of different compounds. The
different compounds may be spotted onto the support in
predetermined positions. Alternatively the hydrated support may
comprise a separation gel or membrane, containing compounds such as
proteins or DNA, that have been separated on the basis of their
size or ionic properties.
[0095] The translocation of one or more bilayers may allow a
bilayer comprising one or more particular membrane-associated
proteins to be rapidly screened against a library of compounds in
the hydrated support. The screen may allow compounds in a library
which interact with a membrane-associated protein and cause a
detectable change in properties at the bilayer to be detected. The
compounds in the library may be proteins, DNA or other small
molecules. The detectable change may be, for example, a change in
conductance or a change in fluorescence or other marker
pattern.
[0096] Alternatively, or additionally, translocation of the bilayer
may allow a library of compounds in a hydrated support to be
screened for potential membrane-associated proteins. Again, the
membrane-associated proteins may be detected by a change in
properties at the bilayer, for example, a change in conductance or
capacitance across the membrane. The library may comprise protein
extracted from a cell or a population of cells.
[0097] The bilayer may be formed on a porous hydrated support which
may then be scanned across the surface of a cell to detect local
concentration differences in excreted compounds, such as ATP. The
cell may be prokaryotic or eukaryotic. The bilayer may be used to
detect analytes/compounds, such as cell excretions, on a cell
surface in vitro or in vivo.
[0098] The bilayer may be formed on a suspended hydrated support in
a mobile device, such as a pipette tip, which may then be scanned
across the surface of a cell. Alternatively the mobile device may
be used to probe different solutions, for example, different
biological samples.
[0099] A plurality of separate bilayers may be formed between a
plurality of separate hydrophilic bodies and one or more hydrated
supports. A hydrophilic body may be contacted with one or more
other hydrophilic bodies on the same hydrated support to form a
plurality of separate lipid bilayers between each of the
hydrophilic bodies as well as between the hydrophilic bodies and
the hydrated support. The hydrophilic bodies may be arranged in a
two, or potentially three, dimensional array.
[0100] Two or more separate hydrophilic bodies on the same hydrated
support may comprise the same or different detection means and/or
different reagents, and/or the same or different
membrane-associated proteins relative to each other.
[0101] An array of aqueous droplets may be deposited over a
hydrated support surface to detect the location of
analytes/compounds in the hydrated support, for example by the
fluorescence of a hydrophilic body or a change in recorded
conductance when an analyte/compound is detected.
[0102] According to another aspect the invention provides a bilayer
product comprising a hydrophilic body and a hydrated support with a
bilayer of amphipathic molecules therebetween. Preferably the
bilayer is formed by interaction of a monolayer of amphipathic
molecules on the hydrophilic body and a monolayer of amphipathic
molecules on the hydrated support.
[0103] The bilayer may be formed by any method of the
invention.
[0104] The skilled man will appreciate that all the preferred
features discussed with reference to the first or second aspect of
the invention, and in particular those relating to the bilayer
itself and not its production, can be applied to a bilayer product
according to the invention and to all aspects of the invention
which use a bilayer.
[0105] According to another aspect of the invention, there is
provided the use of a bilayer product according to the invention in
conjunction with fluorescence microscopy.
[0106] The nature of the hydrated support preferably allows the
bilayer to be viewed using a microscope. Thus in this aspect of the
invention the hydrated support preferably is a layer no more than
about 2 mm thick. Preferably the hydrated support is from about 1
nm to about 2 mm thick, preferably from about 100 nm to about 1 mm
thick, more preferably the hydrated support is from about 100 nm to
about 400 nm thick.
[0107] Preferably when a thin hydrated support of less than about
400 nm thick is used it is kept in contact with a re-hydrating
medium, such as a larger bulk of hydrating liquid or hydrated
support material, to prevent drying out of the thin support
material. The rehydrating medium may be the same or different in
material/composition to the thin hydrated support, and is present
to prevent the thin layer from dehydrating. The rehydrating support
may be agarose gel, water or polyacrylamide gel. The rehydrating
support may be porous or solid.
[0108] Molecules to be observed may be fluorescently labelled with
fluorophores.
[0109] Preferably fluorophores in the hydrophilic droplet and/or
hydrated support are observed using total internal reflection
fluorescence.
[0110] Preferably observations using total internal reflection
fluorescence are used in combination with electrical
measurements.
[0111] Total internal reflection fluorescence microscopy may be
used to observe fluorescence from entities present in the bilayer
either as a bulk property of the bilayer, or with suitable
detection down to the level of individual molecules.
[0112] The advantage of using total internal reflection
fluorescence to observe the fluorophores is that only fluorophores
within about 200 nm of the lipid bilayer are illuminated and thus
observed, whilst other fluorophores not close to the lipid bilayer
are not illuminated and not observed. Using total internal
reflection fluorescence measurements in combination with electrical
measurements has the advantage that protein-protein interactions
can be studied, for example, the assembly of channel proteins such
as .alpha.-hemolysin from labelled subunits or the electrical
response of ion channels to the binding of a fluorescent substrate
can be studied.
[0113] According to a yet further aspect of the invention there is
provided a method of screening for an interaction between a bilayer
of amphipathic molecules and one or more compounds in a library
comprising: [0114] i) providing a bilayer product comprising a
hydrophilic body and a hydrated support with a bilayer of
amphipathic molecules therebetween; [0115] ii) translocating the
hydrophilic body and thus the bilayer across the surface of the
hydrated support; and [0116] iii) detecting any interaction between
the bilayer and a compound in the hydrated support.
[0117] Preferably the bilayer product is made by the method of the
first or second aspect of the invention.
[0118] Preferably the hydrated support comprises the library of
compounds to be tested.
[0119] In one embodiment a membrane-associated protein may be
inserted into the bilayer before or as the bilayer is translocated
across the hydrated support.
[0120] Translocation of the bilayer may be achieved by the direct
or indirect contact of the hydrophilic body with a micromanipulator
arranged to move the hydrophilic body. The hydrophilic body may be
in contact with an electrode which may be moved to translocate the
bilayer across the hydrated support.
[0121] The method of screening may be automated to allow high
throughput screening to be undertaken.
[0122] The skilled man will appreciate that the preferred features
of any aspect of the invention relating the method of producing a
bilayer, to a bilayer per se and to the translocation of a bilayer
can be applied to this aspect of the invention.
[0123] According to a further aspect of the invention, there is
provided the use a bilayer product according to the invention to
identify one or more membrane-associated protein present in and/or
on the hydrated support or the hydrophilic body.
[0124] According to another aspect of the invention, there is
provided the use of a bilayer product according to the invention to
identify substances capable of interaction with a
membrane-associated protein located in the bilayer.
[0125] According to another aspect of the invention, there is
provided a method for detecting one or more analytes present in an
aqueous solution comprising the steps of: [0126] (a) providing a
lipid-in-oil solution in a walled vessel, wherein at least part of
a wall of the vessel comprises a porous hydrated support; [0127]
(b) providing a hydrophilic body in the lipid-in-oil solution;
[0128] (c) forming a first monolayer of lipid molecules on the
surface of the porous hydrated support and a second monolayer of
lipid molecules on the surface of the hydrophilic body; [0129] (d)
contacting the porous hydrated support with the hydrophilic body
such that a lipid bilayer forms between the lipid monolayer on the
hydrophilic body and the lipid monolayer on the porous hydrated
support; [0130] (e) contacting the porous hydrated support with the
aqueous solution such that the analytes present in the aqueous
solution are available to the lipid bilayer; [0131] (f) detecting
the insertion of analytes into the lipid bilayer and/or the
translocation of the analytes across the lipid membrane and/or the
interaction of analytes with the bilayer, using detection
means.
[0132] An apparatus for use with a lipid bilayer comprising a
walled vessel for retaining a lipid/oil solution, wherein at least
one portion of a wall of the vessel comprises a porous support
arranged to be hydrated in use.
[0133] The apparatus may comprise a detection means. Preferably the
detection means is an electrode or a photodetector.
[0134] Preferably the apparatus is portable. Preferably the
apparatus is handheld.
[0135] The walled vessel may be a pipette tip.
[0136] It will be appreciated that all the optional and/or
preferred features of the invention discussed in relation to only
some aspects of the invention can be applied to all aspects of the
invention.
[0137] Preferred embodiments/aspects of the invention will now be
described by way of example only with reference to the accompanying
figures, in which:
[0138] FIG. 1--illustrates an aqueous droplet (hydrophilic body) on
a hydrated support bilayer, referred to herein in as a
droplet-on-hydrated-support bilayer (DHB); FIG. 1A is a diagram of
a droplet-on-hydrated-support bilayer. A lipid monolayer
spontaneously forms on the aqueous surface of the aqueous (water)
droplet and the hydrated support (hydrogel) when each is immersed
in a solution of lipid in hydrophobic oil. When the monolayers of
the two components are brought into contact they form a lipid
bilayer; FIG. 1B shows a droplet bilayer (DHB) visualised from
below with an inverted microscope--the image shows a droplet,
without an electrode, supported on a hydrogel surface. The single
continuous bilayer area in the centre of the droplet is easily seen
due to the large change in refractive index at the interface; FIG.
1C demonstrates that the lipid bilayer between the droplet and
hydrated support can be electrically accessed via electrodes
inserted into both the droplet and the hydrated support. The
electrical capacitance trace shows the formation of a lipid
bilayer. Bilayer capacitance is determined by applying a triangular
potential waveform to the lipid bilayer and measuring the square
wave peak-to-peak current response;
[0139] FIG. 2--illustrates the scanning of proteins in hydrogels
using a droplet-on-hydrated-support bilayer; FIGS. 2A and B show
composite images of polyacrylamide gels after
droplet-on-hydrated-support bilayer scanning, created by overlaying
an image of the dried gel (containing visible pre-stained maker
lane M) with autoradiographs to visualise the radio-labelled
protein bands. Marker lane (M) bands correspond to molecular
weights of approximately 210, 111, and 71 kDa; FIG. 2A is an
SDS-PAGE gel containing the potassium channel Kcv; FIG. 2B is an
SDS-PAGE gel containing the pore forming protein alpha hemolysin
(aHL) aHL-WT (WT) and aHLM113F-D8 (113F) which forms heptameric
protein pores. After immersion in DPhPC/C16 (lipid/oil) solution
the gels were scanned with 200 mL aqueous droplets which had formed
a bilayer on contact with the gel (hydrated support). Protein
insertion and binding at the bilayer was monitored via patch-clamp
amplified electrical recordings as a function of droplet-bilayer
position on the gel surfaces; FIG. 2C shows typical electrical
recordings from the regions of the gels containing Kcv (+20 mV, 500
mM KCl, 10 mM Hepes, pH 7.0), .alpha.HL-WT and .alpha.HL-M113F-D8
(+10 mV, 1 M KCl, 10 mM Na.sub.1PO.sub.4, pH 7.0). The aHL channels
were scanned with droplets containing 10 .mu.M .beta.-cyclodextrin
(.beta.CD) to differentiate between the two mutants; FIG. 2D shows
that channel-protein insertion was only observed in localised
regions about the separated protein bands. This is illustrated by a
12 mm linear scan across the aHL-WT band (dotted line marked on gel
in FIG. 2B), which shows the rate of channel-protein insertion as a
function of bilayer position;
[0140] FIG. 3--illustrates the results of scanning cell extracts in
gels using droplet-on-hydrated-support bilayers; FIG. 3A shows a
coomassie stained polyacrylamide gel after
droplet-on-hydrated-support bilayer scanning, showing SDS-PAGE
purified E. coli cell extracts. The E. coli cell lines were
separately transformed to produce .alpha.HL-WT (lane 1) and
.alpha.HL-M113F-D8 (lane 2) through leaky expression. After
immersion in DPhPC/C, solution the gel was scanned with 200 mL
droplets containing 10 .mu.M .beta.-cyclodextrin. Protein insertion
and binding in the bilayer was monitored via patch clamp amplified
electrical recordings as a function of droplet-bilayer position on
the gel surface; FIG. 3B shows typical electrical recordings from
scans of the regions circled on the gel. Large numbers of
.alpha.HL-WT (top) and .alpha.HL-M113F-D8 (middle) channels
inserted from the regions indicated. Small numbers of unidentified
porin-like channels (bottom) were found in the lower region of the
gel;
[0141] FIG. 4--illustrates scanning of cyclodextrins in gels using
droplet-on-hydrated-support bilayers; FIG. 4A shows a schematic of
the experimental approach employed to scan molecules doped into
polyacrylamide gels by absorption. .gamma.-cyclodextrin (.gamma.CD)
and heptakis (2,3,6-tri-O-methyl)-.beta.-cyclodextrin (h.beta.CD)
were introduced to the bottom of the gel approximately 10 mm apart.
After immersion and stabilisation in the lipid/oil solution the gel
was scanned with droplets containing .alpha.HL-WT channels.
Cyclodextrin binding to .alpha.HL-WT channels in the droplet
bilayer was monitored via patch clamp amplified electrical
recordings as a function of droplet-bilayer position on the gel
surface; FIG. 4B shows the binding characteristics of .gamma.CD
(top, 68% block) and h.beta.CD (bottom, 95% block) to aHL-WT are
clearly distinguishable in electrical recordings (-50 mV, 1 M KCl,
10 mM Na.sub.1PO.sub.4, pH 7.0); FIG. 4C shows the binding
frequency of the two cyclodextrins plotted as a function of
distance in a scan between the two doped locations. Lines indicate
Gaussian fits to the measured binding frequency;
[0142] FIG. 5--shows a diagram of a droplet (hydrophilic body)
ejected from a pipette tip, whilst immersed in a lipid/oil
solution. The droplet is contacted with a hydrogel (hydrated
support) to form a lipid bilayer;
[0143] FIG. 6--illustrates a droplet-on-hydrated-support bilayer
produced using the method of FIG. 5 visualised from below using an
inverted microscope;
[0144] FIG. 7--shows a diagram of a lipid bilayer formed on a
hydrated support fixed to a tip of a pipette which can be used to
scan or probe an aqueous system;
[0145] FIG. 8--shows a diagram of how the lipid bilayer can be
increased and decreased in area by moving the centre of the aqueous
droplet towards or away from the hydrated support using an
electrode inserted in the droplet or in electrical contact with the
droplet;
[0146] FIG. 9--illustrates how a second aqueous droplet carrying a
cargo of reagents can be burst together with an aqueous droplet
without rupturing the bilayer;
[0147] FIG. 10--shows a diagram of how reagents can be introduced
into an aqueous droplet by injection from (A) a micro-pipette, or
(B) a micro-pipette with a multi bore capillary;
[0148] FIG. 11--illustrates multiple droplets-on-hydrated-support
bilayers on a single hydrated support. FIG. 11A--shows
independent/separate droplets; FIG. 11B--shows connected droplets
forming multiple lipid bilayers between the droplets and with the
underlying hydrated support;
[0149] FIG. 12--illustrates total internal reflection fluorescence
microscopy on a droplet-on-hydrated-support bilayer;
[0150] FIG. 13A--shows an alternative portable device for use in
producing a bilayer according to the invention;
[0151] FIG. 13B--illustrates a bilayer produced using the device of
FIG. 13A visualised from below using an inverted microscope;
[0152] FIG. 14A--shows a device for producing an array of
bilayers;
[0153] FIG. 14B--illustrates a bilayer produced using the device of
FIG. 14A;
[0154] FIG. 15A--shows a device for producing a bilayer in which
the aqueous phase is capable of perfusion;
[0155] FIG. 15B--illustrates electrical traces produced using the
device of FIG. 15A, in the top trace .alpha.-hemolysin channels are
shown inserting into the bilayer, in the bottom trace cyclodextrins
are shown binding;
[0156] FIG. 15C--illustrates a bilayer produced by the device of
FIG. 15A;
[0157] FIG. 16--shows an alternative device to that of FIG. 15A in
which the bulk aqueous volume is a microfluidic channel;
[0158] FIG. 17--illustrates a large bilayer produced by the method
of the invention;
[0159] FIG. 18--illustrates a further large bilayer produced by the
method of the invention; and
[0160] FIG. 19--illustrates the concentration of membrane bound
proteins using a bilayer according to the invention.
[0161] "A droplet-on-hydrated-support bilayer" as referred to in
the specific examples is the same as "a bilayer of amphipathic
molecules" as previously discussed.
METHODS
[0162] 1,2-Diphytanoyl-sn-glycero-3-phosphocholine (Avanti Polar
Lipids), hexadecane (Sigma-Aldrich), .beta.-cyclodextrin
(Sigma-Aldrich), and .gamma.-cyclodextrin (Sigma-Aldrich), heptakis
(2,3,6-tri-o-methyl)-.beta.-cyclodextrin (Cyclolab) were used
without further purification.
In Vitro Transcription/Translation of Proteins
[0163] .alpha.HL-WT, .alpha.HL-M113F-D8 (Gu, L. Q. et al., 2001.
Journal of General Physiology 118, 481-493) (RL2 background
(Cheley, S. et al., 1999. Protein Science 8, 1257-1267) and a
C-terminal D8 extension to produce a gel shift relative to
.alpha.H-WT) and Kcv were prepared from genes cloned in the pT7.SC1
vector (Cheley, S. et al., 1997. Protein Engineering 10, 1433-1443)
using a coupled in vitro transcription/translation (IVTT) kit
(Promega Corporation) as previously described (Walker, R. et al.,
1992. Journal of Biological Chemistry 267, 10902-10909).
.sup.35S-methionine was incorporated into the proteins for
visualisation by autoradiography. 50 .mu.L in vitro
transcription/translation reactions of the aHL proteins were
oligomerised as described previously (Walker, B. et al., 1992.
Journal of Biological Chemistry 267, 10902-10909), then pelleted
and resuspended (20 .mu.L, 10 mM MOPS buffer, pH 7.4, 150 mM NaCl,
1 mg mL.sup.-1 BSA). Prior to electrophoresis, the 20 .mu.L
resuspended oligomer samples were mixed with 5 .mu.L of
5.times.SDS-containing Laemmli buffer (final concentration: 10%
(v:v) glycerol, 5% 2-mercaptoethanol, 2.3% SDS, 0.0625 M Tris, pH
7.5).
[0164] 100 .mu.L in vitro transcription/translation reactions were
performed for Kcv, and the products were subsequently separated in
a 10% Tris-HCl gel by electrophoresis. The gel was dried onto paper
under vacuum at room temperature and then imaged by
autoradiography. The band corresponding to the Kcv tetramer was cut
from the gel and rehydrated (300 .mu.l, 10 mM Hepes, pH 7.4). The
rehydrated gel was crushed and transferred to a 0.2 .mu.m cellulose
acetate microfiltration tube (Rainin) and centrifuged at 25000 g
for 30 minutes to recover the solubilised protein.
Electrophoresis of In Vitro Transcription/Translation Proteins
[0165] 5 .mu.L aliquots of the gel-purified Kcv tetramers were
loaded into 8.5% Tris-acetate polyacrylamide gels and subjected to
electrophoresis (200 V, 20 minutes) in TBE buffer (8.9 mM Tris, pH
8.3, 8.9 mM boric acid, 200 .mu.M EDTA, 0.1% SDS) to separate the
protein bands. The gel tank was then refilled with SDS-free TBE
buffer, and electrophoresis was continued (50 V, 2 hours) to remove
SDS in the gel.
[0166] 5 .mu.L aliquots of the in vitro transcription/translation
.alpha.-hemolysin oligomers were loaded into 7% Tris-acetate
polyacrylamide gels (XT Criterion; Bio-Rad Laboratories Inc.) and
subjected to electrophoresis (200 V, 1 hour) in XT Tricine buffer
(Bio-Rad Laboratories Inc.) to separate the protein bands. The gel
tank was then refilled with SDS-free Laemmli buffer, and
electrophoresis was continued (100 V, 2 hours) to remove SDS.
[0167] All gels were run with a pre-stained marker lane (SeeBlue
Plus2, Invitrogen). Following droplet-on-hydrated-support bilayer
gel scanning the gels were imaged by autoradiography.
E. coli Crude Extraction and Electrophoresis
[0168] Competent E. coli JM109(DE3) cells (Promega Corporation)
were transformed by heat-shock with pT7-plasmids encoding either
.alpha.HL-WT or .alpha.HL-M113F-D8. Single colony transformants
were picked and cultured for 16 hours in 2 mL LB-medium containing
50 .mu.g mL-1 ampicillin. The cells were then centrifuged at 2500 g
for 20 minutes and resuspended (200 .mu.L, 25 mM MOPS, pH 7.4, 150
mM NaCl, 0.5% (w:v) SDS, 500 ng DNase 1). Following 30 minutes of
incubation on ice the 200 .mu.L samples were mixed with 50 .mu.L of
5.times.SDS-containing Laemmli buffer (final concentration: 10%
(v:v) glycerol, 5% 2-mercaptoethanol, 2.3% SDS, 0.0625 M Tris, pH
7.5). 45 .mu.L of this solution was then loaded into 10% Bis-Tris
polyacrylamide gels (XT Criterion; Bio-Rad Laboratories Inc.) and
subjected to electrophoresis (200 V, 30 minutes) in XT MOPS buffer
(Bio-Rad Laboratories Inc.). The gel tank was then refilled with
SDS-free buffer (50 mM MOPS, 50 mM Bis-Tris, pH 7.0), and
electrophoresis was continued (100 V, 2 hours) to remove SDS.
[0169] Following droplet-on-hydrated-support bilayer gel scanning
the gels were stained with Coomassie Brilliant Blue
(Sigma-Aldrich).
Droplet-on-Hydrated-Support Bilayer Gel Scanning
[0170] After electrophores the Kcv gels were immersed in 10 mM
Hepes buffer (pH 7.0) containing 500 mM KCl for at least 30
minutes. The aHL and E. coli gels were immersed in 10 mM
Na.sub.2PO.sub.4 (pH 7.0) buffer containing 1M KCl. After dialysis
the gels were left in 10 mM DPhPC/C.sub.16 solution for 15 minutes,
then scanned with approximately 200 nL droplets of the same buffer
as that in the gel. Droplets were moved about the surface of the
hydrogels with the inserted Ag/AgCl electrode attached to a dxdydz
micromanipulator (NMN-21; Narishige).
Electrical Measurements and Bilayer Imaging
[0171] 100 .mu.m diameter Ag/AgCl wire electrodes were used to
electrically access the droplets and gels. Currents were recorded
with a patch clamp amplifier (Axopatch 200B; Axon Instruments), and
digitized at 1 kHz (MiniDigi-1A; Axon Instruments). Electrical
traces were filtered post-acquisition (100 Hz lowpass Gaussian
filter) and analyzed using pClamp 9.0 software (Axon Instruments).
The gel scanning apparatus and amplifying headstage were enclosed
in a Faraday cage attached to an inverted microscope (TE-2000;
Nikon Instruments UK) equipped with a camera (DS-1QM; Nikon) for
imaging and positional tracking of the droplet-on-hydrated-support
bilayers.
Results
Creating Droplet-on-Hydrated-Support Bilayers
[0172] 10 mM 1,2-diphytanoyl-sn-glycero-3-phosphocholine (DPhPC) in
hexadecane (C.sub.16) was used as the lipid/oil solution in all
experiments. Aqueous volumes immersed in this solution
spontaneously self-assemble a DPhPC monolayer, and when the
monolayers of two components are brought into contact they
spontaneously form a lipid bilayer (Tsofina, L. M. et al., 1966.
Nature 212, 681-683; Malmstadt, N. et al., 2006. Nano Letters 6,
1961-1965; Funakoshi, K. et al., 2006. Analytical Chemistry 78,
8169-8174; Holden, M. A. et al., 2007. Journal of the American
Chemical Society--in press). A droplet-on-hydrated-support bilayer
is formed by contacting aqueous droplets with porous hydrated
supports such as hydrogels (FIG. 1A). A stabilisation period of at
least 15 minutes was required before contacting monolayers to
prevent fusion. After this period, bilayer formation was observed
with almost 100% efficiency within a few seconds to a minute of
contact of an aqueous droplet with the hydrated support. The
droplet-on-hydrated-support bilayers were visualised on an inverted
microscope (FIG. 1B), this technique was used to track the position
of a lipid bilayer during experiments.
[0173] The lipid bilayers were electrically accessed by inserting a
100 .mu.M diameter Ag/AgCl electrode into the droplets (FIG. 1A)
using a micromanipulator. With a corresponding Ag/AgCl ground
electrode in the hydrated support, electrical measurements across
the lipid bilayer were carried out. The lipid bilayers were
typically able to withstand voltages up to approximately 300 mV
while retaining seals in excess of 100 G.OMEGA.. Electrical noise
levels were typically of the order of .+-.0.5 pA rms with a 1 kHz
recording bandwidth. This reflects the limitations of this
apparatus and not the inherent noise in droplet-on-hydrated-support
bilayers.
[0174] Synchronous optical measurements of bilayer area (FIG. 1B)
in conjunction with capacitance measurements (FIG. 1C) yielded a
specific capacitance of 0.65 .mu.F cm.sup.-2 at 22.degree. C. for
the DPhPC bilayers in this system. This agrees well with other
reported values (0.4 to 0.8 .mu.F cm.sup.-2) (Montal, M. &
Mueller, P. 1972. Proceedings of the National Academy of Sciences
of the United States of America 69, 3561-3566; Fujiwara, H. et al.
2003. Journal of Chemical Physics 119, 6768-6775; Funakoshi, K. et
al., 2006. Analytical Chemistry 78, 8169-8174), indicating the
lipid bilayers are similar in thickness to their planar bilayer
counterparts.
Gel Scanning with Droplet-on-Hydrated-Support Bilayers
[0175] By scanning the position of a bilayer across a gel while
making single-channel recording measurements it is possible to
resolve the location of isolated membrane channels as they insert
into the lipid bilayer. When applied to electrophoretically
separated protein bands in hydrogels, scanning allows the
determination of whether a particular band contained a
channel-forming protein. To validate this technique, polyacrylamide
gels containing the viral potassium channel Kcv (Plugge, B. et al.,
2000. Science 287, 1641-1644; Gazzarrini, S. et al., 2003. Febs
Letters 552, 12-16), and two mutants of the staphylococcal pore
forming toxin .alpha.-hemolysin (.alpha.HL) (Song, L. Z. et al.,
1996. Science 274, 1859-1866) were scanned.
[0176] SDS-PAGE was used to separate in vitro
transcription/translation (IVTT) expressed Kcv, wild-type
.alpha.-hemolysin (.alpha.HL-WT) and a M113F .alpha.-hemolysin
mutant (.alpha.HL-M113F-D8) (Gu, L. Q. et al., 2001. Journal of
General Physiology 118, 481-493) in polyacrylamide gels (FIG. 2A
and FIG. 2B). SDS-PAGE was followed by electrophoretic cleaning to
remove free SDS from the gels (which would otherwise destabilise
the lipid bilayer). This step can be omitted when running gels
under native detergent-free conditions. After electrophoresis, the
gels were dialysed in appropriate buffers containing KCl to
introduce the electrolyte necessary for electrical recordings.
Following dialysis the gels were immersed in the DPhPC/C.sub.16
lipid/oil solution for 30 minutes, and then scanned with 200 nL
droplets (producing bilayers of approximately 200 .mu.m in
diameter). The droplets were moved about the surface of the
hydrogels by translating an inserted Ag/AgCl electrode, and the
lipid bilayer position was tracked visually on an inverted
microscope.
[0177] When droplet-on-hydrated-support bilayers were positioned
over regions of the gels containing channel proteins stepwise
changes in ion-current resulting from the spontaneous insertion of
channels could be detected. FIG. 2C shows typical examples of
electrical traces acquired when scanning the respective regions of
the gels containing Kcv, .alpha.HL-WT and .alpha.HLM113F-D8. Kcv
behaviour is characterised by stepwise bursts of current as the
channels transiently open and close (FIG. 2C top). .alpha.HL-WT
pores remain open, resulting in stepwise increases in current for
each insertion event (FIG. 2C middle).
[0178] To demonstrate the ability to differentiate between the two
.alpha.-hemolysin mutants the .alpha.HL gels were scanned with
droplets containing .beta.-cyclodextrin (.beta.CD).
.beta.-cyclodextrin acts as a non-covalent blocker that lodges
inside the .beta.-barrel of .alpha.HL, which is observed as a
reversible stepwise change in current in an electrical recording
(Gu, L. Q. et al., 1999. Nature 398, 686-690; Gu, L. Q. &
Bayley, H. 2000. Biophysical Journal 79, 1967-1975). .alpha.HL-WT
does not bind .beta.-cyclodextrin strongly (Gu, L. Q. et al., 1999.
Nature 398, 686-690; Gu, L. Q. & Bayley, H. 2000. Biophysical
Journal 79, 1967-1975), whereas the aHL-M113F-D8 mutant binds
.beta.-cyclodextrin strongly with a voltage-dependent mean dwell
time of approximately 10 seconds (Gu, L. Q. et al., 2001. Journal
of General Physiology 118, 481-493). Without .beta.-cyclodextrin
the electrical characteristics of the two aHL variants are
essentially identical. With .beta.-cyclodextrin the
.alpha.HL-M113F-D8 channels are easily distinguishable by the
.beta.-cyclodextrin binding events overlaying the stepwise
increases in conductance (FIG. 2C bottom).
[0179] It was found that during gel scanning the proteins did not
appear to diffuse, and insertion events were only observed in
highly localised regions about the focused bands in the gel. This
is illustrated quantitatively in FIG. 2D, which shows protein
insertion rate in a linear scan across the wild-type .alpha.HL band
using a droplet with a lipid bilayer of approximately 200 .mu.m in
diameter.
[0180] The gel scanning experiments where extended to SDS-PAGE
purified cell extracts. FIG. 3 shows the results of scanning a
SDS-PAGE gel (FIG. 3A) containing crude extracts from E. coli,
transformed to produce .alpha.HL-WT (lane 1) and .alpha.HL-M113F-D8
(lane 2) through leaky expression. As with the previous gel example
in FIG. 2, these proteins could be electrically characterised (FIG.
3B) from the expected regions of the gel as shown by subsequent
Coomassie staining. Channel insertion rates were higher than
observed from the in vitro transcription/translation gels,
reflecting the substantially higher concentrations of protein
produced from expression in E. coli. Surprisingly, in addition to
aHL channels a number of other channel proteins were detected with
markedly different behaviour (e.g. FIG. 3B bottom), again localised
to specific areas of the gel. The channels typically insert in
multiples of three and show substantial voltage-dependent gating
behaviour.
[0181] It was found that extended immersion in the lipid/oil
solution during gel scanning does not result in any substantial
loss of protein from the gel matrix. Similarly, extended immersion
in electrolyte buffer during the dialysis step does not noticeably
deplete the proteins in the gel. As a result, individual gels can
be re-used in many, at least six, consecutive gel scanning
experiments, and the gel buffer conditions can be varied as
required. Furthermore, the scanning procedure does not affect the
ability to subsequently stain or image the gel, or to recover
specific proteins from the gel for further analysis.
Analyte Detection with Droplet-on-Hydrated-Support Bilayers
[0182] Essentially reversing the gel scanning experiment, protein
channels in droplets on hydrated support bilayers can be used as
molecular sensors to scan different analytes within hydrogels.
[0183] 2% polyacrylamide gels were doped via absorption from
blotted protein solution with approximately 10 .mu.M of
.gamma.-cyclodextrin (.gamma.CD) and
heptakis(2,3,6-tri-O-methyl)-.beta.-cyclodextrin (h.beta.CD) in two
regions spaced approximately 10 mm apart (FIG. 4A). Following a 30
minute immersion in a DPhPC/C.sub.16 solution, the gel was scanned
between the two cyclodextrin regions with a 200 mL droplet
containing .alpha.HL-WT. Under the experimental conditions
.gamma.CD binding to .alpha.HL-WT produces a current block of 68%,
and h.beta.CD binding to aHL-WT produces a current block of 95%.
These different binding amplitudes permit positive identification
of both analytes with aHL-WT (FIG. 4B).
[0184] In this experiment the droplet was translated across the
hydrogel without removing the lipid bilayer from the surface,
retaining the .alpha.HL-WT channels in the lipid bilayer throughout
the scan. The position of the lipid bilayer was recorded by imaging
its position on an inverted microscope, and cyclodextrin binding
events were observed electrically. Cyclodextrin binding frequency
was determined by dividing the total number of events by the number
of aHL-WT channels in the lipid bilayer. FIG. 4C shows the
diffusion limited localised binding of the two cyclodextrin
analytes, plotting .gamma.CD and h.beta.CD binding frequency in a
scan between the two regions.
Bilayer Production Using a Pipette
[0185] With reference to FIG. 5, a hydrogel 11 is layered over a
substrate 13 to form a hydrated support which is then immersed in a
lipid/oil solution 7, such that a lipid monolayer forms on the
upper surface of the hydrogel. An aqueous droplet 9 is partially
ejected from a pipette tip 3 whilst the pipette tip 3 is immersed
in a lipid/oil solution 7. A lipid monolayer forms on the surface
of the droplet. The size of the droplet is controlled by how much
of the aqueous solution is pushed out of the pipette tip on the
application of pressure in the pipette. The size of the area of
contact between the droplet and hydrogel will determine the length
of the bilayer formed. The area of contact can be controlled by
controlling the size of the droplet and the proximity of the
pipette tip to the hydrogel.
[0186] The aqueous droplet 9 is contacted with the hydrogel 11 to
form a lipid bilayer 1. A first electrode 5 in contact with the
aqueous droplet 9 is used in conjunction with a second electrode
(not shown) in contact with the hydrogel 11 to measure electrical
activity across the lipid bilayer 1. The electrodes are
Ag/AgCl.
[0187] By moving the pipette in the x/y plane the bilayer can be
translocated across the surface of the hydrogel.
[0188] FIG. 6 uses inverted microscopy to visualise the formation
of a lipid bilayer 801 between an aqueous droplet 809 and a
transparent hydrated support (not shown) as the droplet and support
are brought into contact.
Droplets in a Scanning `Pipette`
[0189] FIG. 7 illustrates that droplet-on-hydrated-support bilayers
can be formed on a hydrated supported mounted in a translatable
scanning pipette tip. More specifically, a hydrated support 111 is
fixed to the opening of a pipette tip 103, and the pipette tip is
filled with a lipid/oil solution 107. An aqueous droplet 109 is
immersed in the lipid/oil solution and contacted with the hydrated
support 111 to form a lipid bilayer 101. The lipid bilayer 101 can
be used to scan or probe an aqueous system 113 by immersing the
pipette tip 103 into an aqueous system 113. A pair of electrodes
105 is provided to measure electrical activity across the lipid
bilayer 101. One electrode 105 is in contact with the aqueous
droplet 109 and the other electrode 105 is in contact with the
aqueous system 113.
[0190] As long as the hydrated support is porous the bilayer can
sense molecules in the aqueous system which can pass though the
pores. The size of the pores in the support can be controlled to
filter what material actually reaches the bilayer.
[0191] Alternative Device for the Production of a Bilayer
[0192] FIG. 13A illustrates an alternative portable device for
producing a bilayer according to the invention. The device 801 is
made of PMMA and comprises a first screw 811, a second screw 812, a
chamber 816 and a hole 820.
[0193] In use, the chamber 816 is filled with oil and a droplet or
agarose ball 822 is placed on the end of the second screw 812. The
first screw 811 is adjustable to adjust the oil volume in the
chamber 816. The second screw 812 is adjustable to adjust the
height of the droplet or agarose ball 822.
[0194] To form a bilayer 830, the device 801, containing oil and
including a droplet or agarose ball 822, is placed in a solution
825, the solution may be water. The first screw 811 is then raised
to increase the volume in the chamber 816 which draws solution 825
into the chamber. When the solution 825 contacts the droplet or
agarose ball 822 a bilayer 830 spontaneously forms. By adjusting
the height of the second screw 812 the size of the bilayer 830 can
be adjusted. By connecting the second screw 812 to an electrode,
and placing a separate electrode in the solution 825, electrical
access to the bilayer 830 is allowed. Furthermore, when the device
801 is removed from the solution 825 the device holds onto enough
solution 825 to maintain the bilayer 830. This allows the bilayer
to be removed from one solution and returned to a different
solution.
[0195] FIG. 13B illustrates a bilayer 830 formed using the
apparatus of FIG. 13A. The edge 829 of the bilayer 830 is clearly
visible.
Device for Producing an Array of Bilayers
[0196] FIG. 14A depicts a device which can be used to produce an
array of bilayers. The device comprises a base 842 and a lid 841.
The base 840 is filled with agarose in the lower channel, then
filled with a lipid/oil solution and left to equilibrate so that a
monolayer forms on the agarose substrate. The lid 841 is dipped in
a bulk aqueous solution, and through hydrophilic interaction with
the plastic used to make the lid, small droplets remain on the lid.
The lid is then lowered into the base, such that the droplets
remain in oil for a time to equilibrate, before the lid is then
lowered further to bring the droplets into contact with the
underlying agarose. An array of bilayers then forms spontaneously
where the droplets contact the agarose surface. By using electrodes
connected to each of the droplets through the lid, and with a
common electrode in the agarose, each bilayer in the array is
individually accessible for electrical measurements. The nature of
the device allows individual bilayers 844 to be imaged from below
with a microscope, as can be seen in FIG. 14B, in which an agarose
wall 845, a plastic support 846 and the suspended droplet 847 are
also visible.
Device Capable of Perfusion of the Aqueous Phase
[0197] FIG. 15A illustrates how a basic device to produce
droplet-on-hydrated-support bilayers according to the invention can
be extended to provide control of the volume of the oil phase (in
this case with an adjustment screw) to manipulate the bulk liquid
phase being drawn into the device.
[0198] In this embodiment a PMMA device 870 is attached to an
underlying agarose layer 872 mounted on glass 873. An internal
cavity 875, channel, or network of channels is filled with a
lipid/oil solution. The internal oil-filled cavity 875 is open to
the environment at one end, and terminates with a mechanism to
control the oil volume at the other end, in this case a screw. When
the screw is wound in or unwound, the oil is pushed into or pulled
out of the cavity. The means of adjusting volume need not be a
screw, and can extend to any means of controlling the volume of the
oil, such as, it may be solid object such as a pin or needle pushed
into the sample, which could be actuated by a stepper motor or
syringe driver, for example. Alternatively, the volume in the
cavity may be controlled by a syringe upstream, or by a
microfluidic pump. Alternatively, the cavity itself, or parts of
the cavity, could be made of a deformable material or include a
membrane section, that when compressed adjusts the volume of the
oil. When the oil is pulled into the cavity 875, bulk water or bulk
aqueous volume 878 (added to the outside of the device) is also
pulled into the cavity. Where the water/aqeuos volume contacts the
underlying agarose a bilayer will spontaneously form.
[0199] With an electrode 879 in the bulk water or aqueous volume
878, and another electrode 872 in the agarose 872 the bilayers are
accessible for electrical measurements. FIG. 15B illustrates an
electrical trace produced using the device of FIG. 15A showing an
example of .alpha.-hemolysin channels inserting into the bilayers
(top), and binding cyclodextrins (bottom).
[0200] The bilayers can also be imaged using a microscope, see FIG.
15C, which show the bilayer 880 and the bilayer edge 882.
[0201] The bulk aqueous volume in the device can be a microfluidic
channel 885 as depicted in FIG. 16.
[0202] A device according to this embodiment, and illustrated in
FIGS. 15A and 16, has the advantage that the bulk aqueous volume is
open to the external environment which means that there is easier
access for adding components to the system, which is difficult in a
closed droplet system, and also that the volume can be perfused to
fully exchange and alter the contents of the aqueous volume.
Control on Bilayer Area and Capacitance Measurements
[0203] The droplet-on-hydrated-support bilayers subject of this
application can be very large, greater than 1 cm, or very small,
less than 1 micrometre. The area of the bilayer can be adjusted
very quickly by adjusting the height of the droplet relative to the
hydrated support. As is illustrated in FIG. 8, the size/area 215,
315, 415 of the lipid bilayer 201 can be increased or decreased by
moving the centre of the aqueous droplet 109 towards or away from
the hydrated support 111. In this example the droplet is moved by
moving the electrode 205 in the droplet towards or away from the
hydrated support 111.
[0204] By monitoring capacitance across the bilayer the area of the
bilayer can be monitored. Bilayers formed by the method of the
invention shows a linear response of bilayer to area to
capacitance. This relationship is regardless of whether the bilayer
has been reformed and what area changes the bilayer has been
through.
The Production of Large Bilayers
[0205] By using devices such as those described herein, and the
method of the invention, larger bilayers, of 1 mm, 1 cm or more,
can be formed. For example, by using the oil withdrawing techniques
discussed herein, bulk water can be drawn further and further down
channels into the cavity of a device to produce a large
bilayer.
[0206] FIG. 17 shows a composite of images (the bilayer is too big
to be observed in its entirety) taken on a microscope using the
10.times. objective in which the bilayer 900 is approximately 1 mm
by 2 mm. The bilayer edge 902 is visible as are small oil
inclusions 903 and the bulk water inlet 905.
[0207] FIG. 18 shows a slightly larger bilayer as the bilayer of
FIG. 17 is pulled further into the channel, this bilayer is
approximately 1 mm by 5 mm in size. Bilayers of 1 mm by 1 cm in
size were also produced, and larger bilayers of many cms in size
could be created. These larger bilayers may have a capacitance of
.about.100,000 pFarads or more for a DiPhytanoylPC bilayer. This is
in comparison to artificial bilayers produced using previously
known methods which are a few hundred micrometers in diameter, and
which have capacitances of a few hundred pFarads for DiphytanoylPC
bilayers.
[0208] Bursting, Injection and Perfusion of
Droplet-on-Hydrated-Support Bilayers
[0209] The droplet part of a droplet-on-hydrated-support bilayer
can be burst without rupturing the bilayer. This is illustrated in
FIG. 9 where the bursting of a cargo-carrying droplet into an
existing droplet does not disrupt the bilayer. More specifically, a
second aqueous droplet 517 carrying a cargo of reagents is shown
bursting/fusing together with the existing aqueous droplet 509
without rupturing the bilayer 501.
[0210] FIG. 10 illustrates that reagents can be introduced into an
aqueous droplet 609 by injection from a micro-pipette 619 or a
micro-pipette with a multi bore capillary 621. This offers a simple
way of introducing compounds into an existing droplet.
Droplet-on-Hydrated-Support Bilayers in Large Connected or
Unconnected Networks
[0211] Multiple droplet-on-hydrated-support bilayers may be formed
between individual unconnected droplets on a hydrated support or
between multiple connected droplets on a hydrated support.
[0212] With reference to FIG. 11, multiple
droplet-on-hydrated-support bilayers 709 are dispersed on a single
hydrated support 111. FIG. 11A shows a plurality of
independent/separate aqueous droplets 709 on the hydrated support
111 all of which are forming a bilayer with the support.
[0213] FIG. 11B shows a plurality of aqueous droplets 709 connected
in an array on the hydrated support 111, forming multiple bilayers
between the droplets 709 and with the hydrated support 111.
[0214] Such large arrays are suitable for fluorescent experiments.
These arrays may also be used for larger statistical studies. For
example, an array of tiny droplets, from about 10s of nanolitres to
10s of picolitres, can be made with the probability tuned that each
contains only one molecule of a given protein/reagent. Experiments
can then be performed that track the turnover of an enzyme, for
example, at the single molecule level.
[0215] Large arrays could also be used to fluorescently scan
hydrogels containing isolated protein bands. For example if an
array of droplets containing a Ca-sensitive fluorophore containing
polymer is deposited over a gel containing calcium and isolated
alpha-hemolysin channels in an electrophoretically focussed band,
then droplets positioned over the protein band would insert the aHL
and become fluorescent as calcium entered the droplets.
Droplet-on-Hydrated-Support Bilayers for Fluorescence
[0216] Droplet-on-hydrated-support bilayers laid down on thin
hydrated supports can be fluorescently examined using total
internal reflection fluorescence (TIRF) microscopy.
[0217] FIG. 12, which illustrates total internal reflection
fluorescence microscopy on a droplet-on-hydrated-support bilayer, a
supporting substrate comprised of a thin layer of agarose is formed
on a glass coverslip. This thin substrate is rehydrated by filling
a polymethyl methacrylate (PMMA) micro-channel device with aqueous
agarose. The device wells are filled with a solution of lipid in
oil. An aqueous droplet is placed on top of the hydrogel underneath
the oil. A lipid bilayer forms at the interface between the two
aqueous phases. The evanescent field propagates into the
droplet-on-hydrated-support bilayer illuminating the lipid bilayer
and fluorophore-tagged biomolecules in the droplet.
[0218] TIRF techniques may also be used in combination with other
analysis techniques, for example, in combination with acquiring
electrical data. The combination of data may provide improved
information on protein function.
Efficiency of Bilayer Formation
[0219] Experiments in which the hydrophilic body was a water
droplet, and a planar 1% agarose gel made with ultrapure water was
the hydrated support, demonstrated that 20 water droplets all
formed bilayers within 1 minute of contact between the water
droplet and the agarose gel. The bilayers were observed to be
intact after 2 weeks.
[0220] Concentration and Crystallisation of Membrane Proteins
[0221] FIG. 19 shows how bilayers according to the invention can be
used to concentrate membrane proteins. This could be a means to
produce 2-dimensional crystals or semi-ordered lattices of membrane
proteins for further study. By inserting membrane proteins into a
droplet-on-hydrogel bilayer, then shrinking the bilayer size by
pulling the droplet perpendicularly from the surface, it is
possible to drag membrane proteins inwards along the bilayer edge,
without removing them from the bilayer. This results in a
concentration of the inserted membrane proteins towards the centre
of the bilayer.
[0222] FIG. 19 demonstrates the concentration of .alpha.-hemolysin
pores embedded into a droplet-on-hydrogel bilayer. A 1% (w:v) thin
agarose gel was deposited onto a coverslip and dehydrated, it was
then rehydrated using a 1.5% (w:v) agarose containing buffer and
750 mM CaCl.sub.2. A droplet containing the calcium indicator dye
Fluo-4 (25 .mu.M), .alpha.-hemolysin heptamers, 1.5M KCl and a
buffer, was used to create a droplet-on-hydrogel bilayer on this
thin agarose gel.
[0223] Bilayer fluorescence (afforded by the non-chelating Fluo-4)
was imaged as a circular disc through TIRF illumination. Upon
insertion of .alpha.-hemolysin pores into the bilayer, calcium flux
into the droplet is possible. In this example the flux was enhanced
by the application of an externally applied negative potential from
electrodes inserted into the gel (ground) and droplet. This
resulted in an enhanced fluorescence emanating from the vicinity of
the pores (imaged as a spot). This was due to the fluorescence of
the Fluo-4 near the pore greatly increasing in intensity upon its
immediate chelation of calcium upon entry into the droplet.
[0224] FIG. 19 shows diffuse .alpha.-hemolysin pores (as
fluorescent spots) 940 diffusing in the bilayer using the method
described above. The bilayer is then gradually shrunk 941,
resulting in the pores being condensed as they were dragged inwards
by the encroaching bilayer edge 942. When the bilayer area was
enlarged again 943, it was possible to see that the pores had been
concentrated to where the bilayer edge was shrunk. Further
concentration of the pores ensued 944 by repeating this process,
leading to further condensation of the pores towards the centre of
the bilayer area 945.
Discussion
[0225] Although droplet-on-hydrated-support bilayers represent a
significant departure from conventional planar lipid bilayers, they
are easier to prepare and work with, and are similarly amenable to
single-channel recording examinations of both major classes of
membrane protein. Droplet-on-hydrated-support bilayers are more
stable than planar bilayers, and in contrast to the typical planar
bilayer lifetime of a few hours (Miller, C. 1986. Plenum Press: New
York), droplet-on-hydrated-support bilayers are often still
functional several weeks after formation. This opens up avenues for
long timescale experiments that have not been previously
possible.
[0226] Droplet-on-hydrated-support bilayers also possess a number
of other unique properties: (i) the lipid bilayers can be moved
across the surface of a hydrated support. This has been exploited
in hydrogel scanning experiments; (ii) droplet-on-hydrated-support
bilayers provide reliable access to stable bilayers over a much
great size range (<1 .mu.m to >1 mm) than possible using
alternative techniques; (iii) the area of the lipid bilayer can be
adjusted during an experiment. For example, this may be used to
control the number of inserted proteins in single-channel recording
experiments; where the lipid bilayer is initially enlarged to
increase the probability of protein insertion, then rapidly reduced
once a single protein has inserted to minimise the chances of
further insertions. Reducing the lipid bilayer area may also be
used to concentrate transmembrane proteins inserted in a lipid
bilayer. This may provide an alternative means to crystallise
membrane-proteins, and for improving the probability of observing
protein-protein interactions; and (iv) the lipid bilayer can be
removed and reformed many times without mixing the droplet and
hydrogel solutions. This may be used to reset single-channel
recording experiments, as removing the lipid bilayer also appears
to remove inserted transmembrane proteins. The fact that no
contents mixing occurs is also important for experiments where
cross-contamination is an issue.
[0227] The sensitivity of droplet-on-hydrated-support bilayer gel
scanning allows direct study of low levels of endogenous protein
from cell extracts without the need for over-expression. In
contrast, examining proteins without over-expression using either
traditional patchclamp techniques or planar lipid bilayers is
difficult. Although whole-cell patch clamping can examine low
levels of endogenous protein, it is often necessary to compensate
for other constituents of the system due to the heterogeneous
nature of cell membranes (Hamill, O. P. et al., 1981. Pflugers
Archly-European Journal of Physiology 391, 85-100; Ashley, R. H.
1995. IRL). It is possible to circumvent this problem using
artificial lipid bilayers, however, it is difficult to extract and
concentrate protein in sufficient quantities to successfully
reconstitute in these bilayers (Miller, C. 1986. Plenum Press: New
York; Ashley, R. H. 1995. IRL).
[0228] Droplet-on-hydrated-support bilayer gel scanning may be
incorporated in existing proteomic methods that rely upon 2D gel
electrophoresis to separate complex mixtures of cellular components
for the discovery and characterisation of new proteins (Palzkill,
T. 2002 Proteomics, Kluwer Academic Publishers, Boston, London;
Simpson, R. J. 2003. Proteins and proteomics: a laboratory manual,
Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.).
Droplet-on-hydrated-support bilayer gel scanning provides means to
identify channel proteins in 2D gels. The observation that
droplet-on-hydrated-support bilayer gel scanning does not appear to
affect the proteins within the gel matrix means that repeated
droplet-on-hydrated-support bilayer scanning of an individual gel
under varying conditions, and subsequent analysis with conventional
proteomic techniques (e.g. mass spectrometry) can be performed.
[0229] This invention provides a new platform for high-throughput
studies of ion-channels. In particular, the requirement for only
nanolitre volumes permits the application of many established
emulsion-based technologies (Joanicot, M. & Ajdari, A. 2005.
Science 309, 887-888; Ahn, K. et al., 2006. Applied Physics Letters
88; Link, D. R. et al., 2006. Angewandte Chemie-International
Edition 45, 2556-2560; Hung, L. H. et al., 2006. Lab on a Chip 6,
174-178) for scaling-up and automating droplet-on-hydrated-support
bilayers. For example, by combining flows of lipid/oil and water
(Thorsen, T. et al., 2001. Physical Review Letters 86, 4163-4166)
thousands of droplets with a controlled size can be created. Large
numbers of droplet-on-hydrated-support bilayers may also be
manipulated in an automated fashion with existing microfluidic
techniques that can create and sort nanolitre droplets in oil (Ahn,
K. et al., 2006. Applied Physics Letters 88; Link, D. R. et al.,
2006. Angewandte Chemie-International Edition 45, 2556-2560).
[0230] The ability to image droplet-on-hydrated-support bilayers
also allows the incorporation of fluorescence techniques.
Single-channel recording experiments have provided a wealth of
functional detail on many ion-channels, but it is difficult to link
this to dynamic changes in protein structure. Single-molecule
fluorescence of labelled proteins is one possible method of
providing additional structural and dynamic information. Moreover,
droplet-on-hydrated-support bilayers allows simultaneous optical
and electrical measurements, which have the potential to uncover
new aspects of channel function that cannot be elucidated with the
individual techniques alone (Borisenko, V. et al., 2003.
Biophysical Journal 84, 612-622; Ide, T. & Yanagida, T. 1999.
Biochemical and Biophysical Research Communications 265, 595-599;
Ide, T. et al., 2002. Single Molecules 3, 33-42; Macdonald, A. G.
& Wraight, P. C. 1995. Progress in Biophysics & Molecular
Biology 63, 1-29; Suzuki, H. et al., 2007. Biosensors &
Bioelectronics 22, 1111-1115; Suzuki, H. et al., 2006. Langmuir 22,
1937-1942); for example, the dynamics of the folding and insertion
of ion-channels.
[0231] Overall, the combination of enhanced stability, the ability
to manipulate the lipid bilayer, electrical access, and imaging
demonstrate that droplet-on-hydrated-support bilayers according to
the invention provide a versatile platform for examining many
aspects of membrane protein function.
* * * * *