WO2005069004A2 - Solid-supported membranes inside porous substrates and their use in biosensors - Google Patents

Solid-supported membranes inside porous substrates and their use in biosensors Download PDF

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Publication number
WO2005069004A2
WO2005069004A2 PCT/US2005/000069 US2005000069W WO2005069004A2 WO 2005069004 A2 WO2005069004 A2 WO 2005069004A2 US 2005000069 W US2005000069 W US 2005000069W WO 2005069004 A2 WO2005069004 A2 WO 2005069004A2
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lipid
receptor
membrane
protein
molecule
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PCT/US2005/000069
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French (fr)
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WO2005069004A3 (en
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Klaus Gawrisch
Holly C. Gaede
Keith M. Luckett
Ivan V. Polozov
Alexei Yeliseev
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The United States Of America As Represented By The Secretary Department Of Health And Human Services
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    • G—PHYSICS
    • G01—MEASURING; TESTING
    • G01N—INVESTIGATING OR ANALYSING MATERIALS BY DETERMINING THEIR CHEMICAL OR PHYSICAL PROPERTIES
    • G01N33/00—Investigating or analysing materials by specific methods not covered by groups G01N1/00 - G01N31/00
    • G01N33/48—Biological material, e.g. blood, urine; Haemocytometers
    • G01N33/50—Chemical analysis of biological material, e.g. blood, urine; Testing involving biospecific ligand binding methods; Immunological testing
    • G01N33/53—Immunoassay; Biospecific binding assay; Materials therefor
    • G01N33/543—Immunoassay; Biospecific binding assay; Materials therefor with an insoluble carrier for immobilising immunochemicals
    • G—PHYSICS
    • G01—MEASURING; TESTING
    • G01N—INVESTIGATING OR ANALYSING MATERIALS BY DETERMINING THEIR CHEMICAL OR PHYSICAL PROPERTIES
    • G01N33/00—Investigating or analysing materials by specific methods not covered by groups G01N1/00 - G01N31/00
    • G01N33/48—Biological material, e.g. blood, urine; Haemocytometers
    • G01N33/50—Chemical analysis of biological material, e.g. blood, urine; Testing involving biospecific ligand binding methods; Immunological testing
    • G01N33/53—Immunoassay; Biospecific binding assay; Materials therefor
    • G01N33/543—Immunoassay; Biospecific binding assay; Materials therefor with an insoluble carrier for immobilising immunochemicals
    • G01N33/54366—Apparatus specially adapted for solid-phase testing

Definitions

  • This invention relates to reagents and methods for forming membranes that may be deposited into porous solid supports for use in biosensors.
  • Immobilized Membranes and Biosensors Immobilized membranes may be employed in a wide range of applications, including enabling biofunctionalization of inorganic solids (semiconductors, gold- covered surfaces, and optoelectronic devices) and polymeric materials; providing a natural, non-denaturing, and defined environment for the immobilization of biomolecules; and allowing the preparation of ultrathin, high-electric-resistance layers on conductors and the incorporation of receptors into insulating layers for the design of biosensors based on electrical and optical detection of ligand binding (Ho, W.S.
  • Biosensors are analytical devices that are used to measure the presence and/or concentration of desired biological molecules in a sample.
  • biomolecules such as antigens, hormone receptors, enzymes, cytokine receptors, antibodies, etc.
  • biosensors can be analyzed using biosensors (see, e.g., Keusgen M. (2002) “BIOSENSORS: NEW APPROACHES IN DRUG DISCOVERY,” Naturwissenschaften. 89(10):433-44. Epub 2002 Sep 11; Albers, J. et al. (2003) "ELECTRICAL BIOCHIP TECHNOLOGY ⁇ A TOOL FOR MICROARRAYS AND
  • Biosensors function by generating a detectable physical signal from the sensor's physical transducer component in response to the binding of a target biological molecule to the sensor's biological component.
  • the physical transducer component is typically an optical or electrical signal, e.g. a fluorescence signal elicited by the ligand binding event, a modulation of a current in response to binding, a pH change in response to binding, a modulation of electrical resistance in response to binding, etc.
  • Care must be taken to ensure that the target biological molecule of the biological component of the biosensor is immobilized, and that the immobilization procedure forms a stable layer of biomolecules.
  • the immobilization procedure must also not undesirably diminish the activity or structure of the target biological molecule of the biological component, or change its substrate reactivity (i.e., perturbations of the biological component is optimally minimized).
  • Integrated bilayers are characterized by an inner monolayer that is either covalently or ionically bonded to the support surface.
  • Freely supported bilayers are characterized as having an inner monolayer that is separated from the support by an ultrathin water layer ( ⁇ 10 A).
  • the third type of supported membrane consists of a bilayer membrane that rests on an ultrathin polymer film (see, Sackmann, E. (1996) "SUPPORTED MEMBRANES: SCIENTIFIC AND PRACTICAL APPLICATIONS,” Science 271 :43-48).
  • Such supported membranes can be associated with silicon microchips, or with beads (as in a column, etc.).
  • Immobilization of the biological molecule can be achieved by adsorbing the biological molecule onto the surface of a solid support, integrating the molecule within a gel or other matrix, or covalently coupling it to a solid support.
  • Some approaches achieve immobilization by adsorbing the biological molecule onto the surface of a solid support, integrating the molecule within a gel or other matrix, or covalently coupling it to a solid support.
  • Other approaches interpose a polymer cushion between the lipid bilayer and the polymer support. Supported lipid-protein bilayers separated from the solid surface by nanometer-thick water layers or ultrathin soft polymer cushions maintain the thermodynamic and structural properties of free bilayers.
  • Flat membrane supports have the advantage of being able to physically separate membranes (and thus provide protection against their disruption). They suffer from the disadvantage of low surface area. Beaded surfaces provide much greater surface area than flat membranes, however, the membranes can touch one another and are not protected. Porous supports provide very large surface area, but their application has suffered from issues related to uniformity and homogeneity of membrane preparation as well as from limited membrane accessibility for ligands that are delivered via the water phase.
  • AAO filters have a 60 ⁇ m thick support layer with pore diameters of 0.2 ⁇ m that is capped by a thin, ⁇ 1 ⁇ m layer with nominal pore diameters of 0.02, 0.1, or 0.2 ⁇ m.
  • AAO filters have a variety of other applications, including support for cell cultures and microscopy, sample preparation for HPLC, IC, and electrophoresis, and liposome extrusion.
  • Lipids have previously been added to AAO as monolayers to change liquid crystal director orientation (Crawford, G.P. et al. (1991) "SURFACE-INDUCED ORIENTATIONAL ORDER IN THE ISOTROPIC PHASE OF A LIQUID-CRYSTAL MATERIAL,” Phys.Rev.A 44:2558-2569), as pore-spanning bilayers (Hennesthal, C. et al. (2000) “PORE-SPANNING LIPID BILAYERS VISUALIZED BY SCANNING FORCE MICROSCOPY,” J.Am.Chem.Soc. 122:8085-8086), as the upper leaflet of hybrid bilayers (Marchal, D. et al.
  • Lipid monolayers have been used to make inner pore surfaces hydrophobic so that they may be used for thermotropic liquid crystal display applications (Marchal, D. et al. (1998) "ELECTROCHEMICAL MEASUREMENT OF LATERAL DIFFUSION COEFFICIENTS OF UBIQUINONES AND PLASTOQUINONES OF VARIOUS ISOPRENOID CHAIN LENGTHS INCORPORATED IN MODEL BILAYERS," Biophys.J. 74: 1937-1948). Multilamellar lipid bilayers have been deposited in AnoporeTM filters by the diffusion of small liposomes (Smirnov, A.I. et al. (2003) “SUBSTRATE-SUPPORTED LIPID NANOTUBE ARRAYS,” JAm.Chem.Soc. 125:8434-8435).
  • G-Protein Coupled Receptors G-Protein Coupled Receptors
  • G-Protein Coupled Receptors convey signals from extracellular hormones and neurotransmitters to intracellular effectors and linked signaling pathways.
  • G-Protein Coupled Receptors are a class of integral membrane proteins belonging to the "7TM" superfamily of transmembrane receptors. The GPCRs are characterized by the possession of seven intramembrane helices, and by domains that extend both into the extracellular environments and the cytosol (Wojcikiewicz, R.J. (2004) "REGULATED UBIQUITINATION OF PROTEINS IN GPCR-INITIATED SIGNALING PATHWAYS," Trends Pharmacol Sci.
  • the GPCRs use an amazing number of different domains both to bind their ligand and to activate G proteins. More than 150 GPCRs have been identified, in at least six families of proteins that show no sequence similarity. The fine-tuning of their coupling to G proteins is regulated by splicing, RNA editing and phosphorylation.
  • the G-Protein Coupled Receptor trigger cellular processes through a conformational shift that is caused by the binding of the GPCR to a ligand molecule (Gether, U. et al. (2002) "STRUCTURAL BASIS FOR ACTIVATION OF G-PROTEIN-COUPLED RECEPTORS," Pharmacol Toxicol. 91(6):304-12). As a consequence of the conformational shift, the GPCR acquires the capacity to bind and cleave intracellular proteins ("G-proteins"). The release of the cleavage products into the cytosol triggers cellular processes to occur.
  • GPCRs include the receptors of the olfactory sensory epithelium that bind odorants and neurotransmitter receptors (e.g., the serotonin receptor), the cannabinoid receptor (Picone, R.P. et al. (2002) “LigAND BASED STRUCTURAL STUDIES OF THE CB1 CANNABINOID RECEPTOR,” J Pept Res. 60(6):348-56; Onaivi, E.S. et al. (2002) “ENDOCANNABINOIDS AND CANNABINOID RECEPTOR GENETICS,” Prog Neurobiol. 66(5):307-44), and the rhodopsin receptor (Ballesteros, J. et al.
  • G protein-coupled receptors mediate the perception of smell, light, taste, and pain (Ahmad, S. et al. (2004) “NOVEL G PROTEIN-COUPLED RECEPTORS AS PAIN TARGETS,” Curr Opin Investig Drugs.
  • the microstructure of the membrane significantly impacts upon the properties of the biosensor.
  • Single lipid bilayers that are directly immobilized to a solid support, or immobilized via a polymer or "hairy rod” cushion see,
  • Figure 1 shows a schematic representation of the lipid bilayer adsorbed to the inner surface of an AAO pore.
  • the z-axis defines the direction of the pore, and the x-y axes define the plane of the filter.
  • the bilayer normal of the membrane is given by D
  • the vector of the external magnetic field is B
  • the angle between these vectors is given by ⁇ .
  • the angle between the pore axis and the magnetic field is defined by ⁇ .
  • the orientation of the bilayer normal in the x-y plane is defined by the angle ⁇ .
  • Panel A Unilamellar samples prepared by flushing the pores after lipid loading.
  • the angular distribution function used in the fit, displaying the mosaic spread of the lipid, is shown below the spectra.
  • the order parameters used in the simulation were obtained from the POPC-d 3 ⁇ MLV 2 H NMR spectrum and the profile is shown in Figure 2, Panel C, Multilamellar samples prepared with no flushing step after lipid loading.
  • the extra signal in the center of the experimental spectrum of lipid adsorbed in AAO pores appears to be mostly from a residual 2 H resonance of AAO hydroxyl groups.
  • the small deviation in signal intensity between measured and calculated spectra in the frequency range of ⁇ 5 kHz could indicate existence of a few percent of lipid with lower headgroup order.
  • Figure 5 shows 300 MHz ⁇ NMR MAS spectra at a rotor spinning frequency of 5 kHz and a temperature of 30°C.
  • Trace A POPC MLVs
  • Trace B POPC in AAO pores
  • Trace C POPC in AAO pores exposed to 5 mM Pr 3+ .
  • Inset Expansion of ⁇ -choline signal in POPC in AAO samples. The POPC resonance assignments are given in the lower spectrum.
  • Figure 6 shows 31 P MAS NMR spectra at a rotor spinning frequency of 5 kHz and a temperature of 30°C.
  • Trace A POPC MLVs
  • Trace B POPC single bilayers adsorbed on AAO pores
  • Trace C POPC adsorbed onto AAO pores and exposed to Pr 3+ .
  • the integral intensity of the superimposed ⁇ POPC +PEG methylene resonance at 3.57 ppm relative to the ⁇ POPC resonance at 3.15 ppm is 8.6:9.
  • Figure 8 show the normalized methylene intensity plotted versus temperature for DMPC supported on AAO and DMPC MLVs. The lines are included as a guide to the eye.
  • Figure 9 shows 500 MHz ⁇ PFG-MAS NMR diffusion measurements on crushed POPC/AAO with trapped PEG8000 at a spinning frequency of 10 kHz, temperature of 30.0 °C, and a diffusion time of 200 ms.
  • Trace A Water resonance and Trace B: choline resonances of spectra acquired at 16 different gradient strengths from 0.01 - 0.37 T/m.
  • Trace C The signal intensity decay of the choline resonance as a function of k, fit to Equation 1.
  • Figure 10A shows a model for lipid adsorption consistent with the NMR data. A single bilayer forms a good seal with the AAO surface by the interaction of a small percentage of the lipids.
  • the lipids adsorb as wavy tubules with an average length of 0.4 ⁇ m. These tubules posses undulation with a radius of curvature of 100 - 400 nm. Trapped between the tubules and the AAO surface are pockets of water with an average thickness of 3 nm.
  • Figure 10B shows an illustration of shape of the lipid bilayer tubules inside the AAO pore.
  • Figure 11 shows the solid state 2 H NMR spectrum of POPC -d 3 ⁇ lipid tubules in anopore filters containing bovine rhodopsin at a lipid/protein molar ratio of 100/1.
  • the anopore disks were aligned with their normal parallel to the B 0 field of the NMR instrument. The spectrum indicates that bilayers adhere to the surface of the cylindrical pores as lipid tubules.
  • Figures 12A-H show 2 H NMR spectra of 18:0(d35)-22:6 PC membranes containing bovine rhodopsin at protein/lipid molar ratios from zero to 1/100. On the left the experimental spectra and on the right the simulated spectra are shown. The simulation yields the chain order parameters as a function of rhodopsin concentration, the mosaic spread of bilayer orientations, and the resonance linewidth. Results confirm formation of lipid tubules containing reconstituted bovine rhodopsin.
  • Figure 13 shows chain order parameter profile of the sn-1 hydrocarbon chain in 18:0(d35)-22:6 PC as a function of rhodopsin concentration reported as a function of molar protein/lipid ratio. In the presence of rhodopsin a small reduction of sn-1 chain order parameters for carbon atoms 2-8 is observed.
  • Figure 14 shows ! H MAS NMR spectrum of a cytoplasmic membrane preparation of E. coli BL21 cells recorded at a MAS spinning frequency of 5 kHz at ambient temperature. Trace A: membrane pellet, Trace B. membranes deposited inside the pores of an anopore filter.
  • Figure 15 shows a comparison of the conventional competitive filter- binding assay (Whatman GF/B filters) with ligand-binding performed on Anopore membranes. Results indicate that depositioning of the cannabinoid receptor in single tubular lipid bilayers at the inner surface of pores did not alter the ligand binding properties of the receptor. Furthermore, nonspecific binding of ligands was reduced significantly, allowing much more accurate ligand binding measurements.
  • Figure 16 shows the use of Anopore filters in a ligand binding study. Shown are scintillation count rates for binding of CP55,940 to the cannabinoid receptor CB2. Depositioning of membranes as tubular lipid bilayers in Anopore filters reduced nonspectific binding to less than 15% of total radioactivity, therefore greatly increasing the accuracy and reproducibility of ligand binding assays. Each data point represents the average of three filters.
  • Figure 17 shows the reconstitution of the CB2 fusion protein into a SOPC lipid bilayer.
  • Panel A Excitation: 532 nm (green laser); Emission: 580 nm.
  • PMT 320; Sensitivity: normal;
  • Panel B Excitation: 532 nm (green laser); Emission: 610 nm.
  • PMT 320; Sensitivity: normal.
  • Alexa 532 Exc conflicton:532 nm; Emission: 561 nm; T Red: Excitation: 583 nm; emission: 602 nm.
  • Figure 18 shows the deposition of SOPC into Anodisk filters. Presented is the lipid fluorescence count as a function of SOPC concentration in the reconstitution mixture. Reconstitution from lipid dispersions in octylglucoside is significantly more efficient than reconstitution from the triple detergent mix that is used for protein solubilization.
  • Figure 19 shows the deposition of CB2/SOPC into Anodisk filters.
  • cannabinoid receptor CB2 fluorescence count as a function of CB2 concentration in the dispersion.
  • CB2 binding per filter saturated at a concentration of 10 micrograms.
  • Figure 20 shows the deposition of SOPC/CB2 into Anodisk filters.
  • lipid fluorescence count as a function of cannabinoid receptor CB2 concentration in the reconstituted membranes. In the concentration range from 0 - 10 ⁇ g of CB2 a reduction of the amount of deposited lipid with increasing protein content was observed. The count rate remained constant at a protein content of 10 ⁇ g or higher.
  • Figure 21 illustrates the depositioning of CB2 into Anodisk filters using a Western blot with anti-MBP antibody.
  • 1 CB2-POPC mixloaded onto Anopore Filter; 2, 2a: flowthrough solution/wash with Tris buffer; 3: CB2 eluted from the Anopore filter with 2% SDS
  • This invention relates to compositions and methods for forming membranes containing membrane proteins that may be deposited into solid supports for use in biosensors.
  • the present invention particularly concerns the development of a procedure capable of forming a high surface area, supported single-lipid bilayer in which the membrane is separated from the support by a closed and stable aqueous cushion.
  • the present invention particularly concerns the development of a procedure for depositing single lipid membranes onto a solid support so as to provide, for example, a total exposed surface area of 500 cm 2 in a filter with a diameter of only 13 mm and thickness of 60 ⁇ m, orienting approximately 2.4 x 10 "7 moles of lipid.
  • 2 H NMR spectra on chain deuterated phosphatidylcholine it was established that lipids adsorb as wavy, tubular bilayers to the inner pore surface.
  • H magic angle spinning NMR it is found that the sample preparation procedure resulted in formation of a single lipid bilayer inside every pore.
  • lipid tubule length may vary from a fraction of a micrometer to several micrometers depending on membrane composition and preparation procedures.
  • the membranes are separated from the solid surface by a thick water layer with an average thickness of 30 A such that lipids are completely unperturbed by the solid support.
  • This makes the membranes suitable for incorporation of biological components, such as sensitive integral and peripheral membrane proteins, e.g. G- Protein Coupled Membrane Receptors (GPCR), receptor tyrosine kinases, ion channels, etc.
  • GPCR G- Protein Coupled Membrane Receptors
  • Adhesion between membrane and solid support is, nevertheless, quite strong. Membranes are not removed from the surface by passing water through the pores at high rate. Furthermore, the invention evidences that the supporting water layer is well sealed by the membrane from the bulk water phase. This permits entrapment of solutes, like soluble components of signal transduction pathways between the solid support and the bilayers. It is demonstrated that water soluble polymers with a molecular weight of 8,000 can be trapped in this water layer. Also, the substrate used for sample preparation, anodized aluminum oxide, is compatible with magnetic resonance spectroscopy, optical spectroscopy, radiotracer studies, neutron- and x-ray experiments, calorimetry, etc., permitting easy detection of receptor binding events as well as structural studies.
  • the large accessible membrane surface raises sensitivity of assays based on our technology by up to three orders of magnitude per unit of chip area.
  • An additional advantage is that substrates are applied by slowly flowing solutions through pores. The flow rate of liquid through the pores is easily controlled, permitting substrate binding studies to be conducted under very reproducible conditions. Because of the unique features of those solid-supported membranes, including lack of perturbation from the substrate, large surface area, strong adhesion to the support, stability to water flow, and the ease of sample preparation, the invention provides considerable promise for preparation of biosensors.
  • the invention concerns a composition
  • a composition comprising a single bilayer lipid membrane supported on a solid support, wherein the membrane comprises a lipid tubule having a wavy tubular geometry and rims, wherein the tubule has rims that are in contact with the solid support, and a center region between the rims that is spaced apart from the support by an aqueous cushion, the aqueous cushion being located between the membrane and the support.
  • the invention further concerns the embodiment of such a composition wherein the solid support is a porous aluminum oxide support.
  • the invention further concerns the embodiment of such compositions wherein the lipid comprises a phospholipid.
  • the invention further concerns the embodiment of such compositions wherein the membrane comprises an incorporated biological molecule.
  • the invention further concerns the embodiment of such compositions wherein the incorporated biological molecule is an integral membrane protein or a peripheral membrane protein.
  • the invention further concerns the embodiment of such compositions wherein the integral membrane protein or the peripheral membrane protein is a receptor molecule, an enzyme, or an antigen.
  • the invention further concerns the embodiment of such compositions wherein the incorporated biological molecule is a receptor molecule capable of binding an agonist or an antagonist of a signal transduction pathway.
  • the invention further concerns the embodiment of such compositions wherein the receptor molecule is a G-protein coupled membrane receptor.
  • the invention further concerns the embodiment of such compositions wherein the solid support is porous, and wherein an agonist or antagonist of the receptor, a G- protein, or an analog of such molecules is provided to the composition by flowing a solution through the pores of the porous support so as to permit the receptor agonist or antagonist to bind to the receptor from inside the lipid tubule from the side of the lipid monolayer opposite from the AAO surface.
  • the invention further concerns the embodiment of such compositions wherein the agonist or antagonist of the receptor, the G-protein, or the analog of such molecules receptor agonist or antagonist is contained in the aqueous cushion.
  • the invention further concerns a biosensor device comprising a solid support, a biological component, and a physical transducer; wherein: the biological component comprises a single bilayer lipid membrane supported on the solid support, wherein the membrane comprises a lipid tubule having a wavy tubular geometry and rims, wherein the tubule has rims that are in contact with the solid support, and a center region between the rims that is spaced apart from the support by an aqueous cushion, the aqueous cushion being located between the membrane and the support; the membrane comprises an incorporated biological molecule selected from the group consisting of an enzyme, a receptor molecule, and an antigen; and the physical transducer serves to generate a signal in response to the binding of a target molecule to the incorporated biological molecule.
  • the biological component comprises a single bilayer lipid membrane supported on the solid support, wherein the membrane comprises a lipid tubule having a wavy tubular geometry and rims, wherein the tubule has rims that
  • the invention further concerns the embodiment of such a biosensor device wherein the solid support is a porous aluminum oxide support.
  • the invention further concerns the embodiment of such biosensor devices wherein the lipid comprises a phospholipid.
  • the invention further concerns the embodiment of such biosensor devices wherein the incorporated biological molecule is a receptor molecule.
  • the invention further concerns the embodiment of such biosensor devices wherein the receptor molecule is a G-protein coupled membrane receptor.
  • the invention further concerns the embodiment of such biosensor devices wherein the solid support is porous, and wherein an agonist or antagonist of the receptor, a G-protein, or an analog of such molecules is provided to the composition by flowing a solution through the pores of the porous support so as to permit the receptor agonist or antagonist to bind to the receptor from inside the lipid tubule from the side of the lipid monolayer opposite from the AAO surface.
  • the invention further concerns the embodiment of such biosensor devices wherein the aqueous cushion contains a biological molecule that interacts with the receptor molecule.
  • the invention further concerns the embodiment of such biosensor devices wherein the aqueous cushion contains a G-Protein, and the incorporated biological molecule is a G-protein coupled membrane receptor.
  • the invention further concerns the embodiment of such biosensor devices wherein the physical transducer comprises an optical sensor, an electrochemical sensor, a potentiometric sensor, a conductometric sensor, or a piezoelectrical sensor; wherein the sensor generates a detectable physical signal in response to the binding of a target biological molecule to the biosensor's incorporated biological molecule.
  • the physical transducer comprises an optical sensor, an electrochemical sensor, a potentiometric sensor, a conductometric sensor, or a piezoelectrical sensor; wherein the sensor generates a detectable physical signal in response to the binding of a target biological molecule to the biosensor's incorporated biological molecule.
  • the invention further concerns the embodiment of such biosensor devices wherein the physical transducer is an optical sensor selected from the group consisting of a colorimetric optical sensor operating in the visible or non-visible spectral range.
  • the optical sensor is an optical sensor that detects fluorescent light.
  • the invention further concerns a method of detecting a pharmacological agent that binds to a biological molecule, wherein the biological molecule is selected from the group consisting of an enzyme, a receptor molecule, an antigen, and an antibody, wherein the method comprises the steps: (A) incubating the pharmacological agent in the presence of a biosensor device, the biosensor device comprising a solid support, a biological component, and a physical transducer; wherein the biological component comprises a single bilayer lipid membrane supported on the solid support, wherein: the membrane comprises a lipid tubule having a wavy tubular geometry and rims, wherein the tubule has rims that are in contact with the solid support, and a center region between the rims that is spaced apart from the support by an aqueous cushion, the aqueous cushion being located between the membrane and the support; the membrane comprises the biological molecule incorporated therein; and the physical transducer serves to generate a signal in response to the binding of a target
  • the present invention relates to compositions and methods for forming membranes that may be deposited into solid supports e.g., for use in biosensors.
  • the invention relates to compositions and methods for forming a high surface area, supported single-lipid bilayer matrix in which the membrane is separated from the support by a closed and stable aqueous cushion.
  • Such compositions comprise a lipid bilayer, a solid support, and where used for a biosensor, a biological molecule (such as a GPCR).
  • lipids may be employed in accordance with the methods of the present invention.
  • such lipids will be glycerol based lipids (e.g., phosphocholine, phosphatidyl-DL-glycerol, phosphatidylethanolamine, phosphatidylinositol, phosphatidylserine, etc.) and their derivatives and salts.
  • lipids will have fatty acid side chains of 10-20 carbon atoms, more preferably of 12-16 carbon atoms.
  • the membrane bilayer compositions of the biosensor devices of the present invention will comprise l-palmitoyl-2-oleoyl-sn-glycero-3 phosphocholine (POPC).
  • POPC l-palmitoyl-2-oleoyl-sn-glycero-3 phosphocholine
  • Any of a variety of solid supports may be employed in the biosensor devices of the present invention, however, the most preferred solid support is a porous aluminum oxide support. In particular, the porous aluminum oxide AnoporeTM support (Structure Probe, Inc., West Chester, PA, USA) is preferred. Although any of a variety of solid supports may be employed in accordance with the principles of the present invention, the most preferred solid support is a porous aluminum oxide support. In particular, the porous aluminum oxide AnoporeTM support (Whatman, Inc.) is preferred.
  • the AnoporeTM support is fabricated from a unique form of aluminum oxide with a highly controlled, uniform capillary pore structure that is tightly controlled at 0.2 ⁇ m.
  • the starting purity of the aluminum metal use in the first step for the manufactured of the membranes is quite high.
  • the support provides the advantages of high flow rates, efficient particle retention, rigid, uniform surface, transparency (when wet), ability to retain virtually no background stain, low levels of extractable materials, promotes sieving of particles at the surface, is temperature resilient (stable to 400° C) and electron beam radiation resistant.
  • the proteins of the biological component of the biosensor can be prepared either by reconstituting a lipid bilayer using, for example, purified membrane protein(s), or by forming bilayers from the cellular membranes of spheroplasts.
  • the present invention provides a facile process for reconstituting single lipid bilayers that contain functional membrane proteins into AnoporeTM filters.
  • a desired membrane protein is preferably recombinantly expressed as a fusion protein containing a C-terminal "tail" (e.g., a Hisio tail) (Grisshammer, R. et al. (1997) "QUANTITATIVE
  • membrane proteins and lipids are solubilized in detergent micelles preferably using either a mixture of 3-[(3- cholamidopropyl)dimethylammonio]-l-propanesulfonate (Chaps), cholesteryl hemisuccinate (CHS), and dodecyl-,B-D-maltoside (LM )] (see, Tucker, J. et al.
  • Liposomes are formed from the mixed detergent-lipid-protein components by lowering the detergent concentration below the critical micelle concentration (cmc) (see, U.S. Patent No. 6,143,321; Urbaneja, M.A. et al. (1990) "DETERGENT SOLUBILIZATION OF PHOSPHOLIPID VESICLE. EFFECT OF ELECTRIC CHARGE,” Biochem J. 270(2):305-308; Marsh, D. et al. (1986) "PREDICTION OF THE CRITICAL MICELLE CONCENTRATIONS OF MONO- AND DI-ACYL PHOSPHOLIPIDS,” Chem Phys Lipids 42(4):271-277; Lasch, J. et al.
  • cmc critical micelle concentration
  • liposomes with lipid bilayers are formed spontaneously when the added buffer causes the detergent concentration to fall below the critical micelle concentration (about 20 mM) (see e.g. Mitchell, D.C. et al. (2001) "OPTIMIZATION OF RECEPTOR-G PROTEIN COUPLING BY BILAYER LIPID COMPOSITION I - KINETICS OF RHODOPSIN- TRANSDUCIN BINDING," J. Biol. Chem. 276(46):42801-42806; Litman, B. J. (1982) "PURIFICATION OF RHODOPSIN BY CONCANAVALIN A AFFINITY
  • the liposomes formed after dilution form bilayers that cover the cylindrical surface on pores in the Anopore® support.
  • the residual detergent is easily flushed out the lipid bilayers by passing detergent- free buffer through the support.
  • the present invention greatly shortens and simplifies the procedure for detergent removal.
  • bilayers can be prepared by extruding spheroplasts through stacked membranes (especially stacked AnoporeTM porous aluminum oxide membranes).
  • the spheroplasts will be bacterial spheroplasts that have been genetically engineered to express membrane proteins. Spheroplasts from genetically engineered E. coli are particularly preferred.
  • Multilayer components may be removed by controlled flushing with buffer.
  • the supported membranes of the present invention will be at least partially spaced apart from the support by a water (or aqueous buffer) layer or cushion.
  • Water-soluble biopolymers may be advantageously entrapped in this water layer or cushion.
  • the bilayer membranes of the present invention have the advantages that the incorporated protein is functional, and the effective membrane area is up to three orders of magnitude larger than the surface area of a flat chip.
  • the formed membranes are stable at moderate flow rates through pores; the flow rate can be controlled by, for example, an infusion pump.
  • the bilayer membranes produced in accordance with the methods of the present invention are wavy tubules as shown in Figures 10A and 10B. The upper and lower ends of tubules are in contact with the solid support, and seal off a layer of trapped water.
  • Membrane protein that is incorporated into such bilayers can be in an orientation that "faces" the support, or in an orientation that "faces" away from the support. Where desired, buffer, charge, pH, and membrane curvature parameters may be altered in order to cause more membrane proteins to adopt a particular orientation relative to the support.
  • the supported membranes of the present invention can be employed in biosensor devices.
  • biosensor device is intended to denote an analytical device that is capable of generating a signal in response to the presence of, and preferably, the binding of, a target molecule.
  • the biosensor devices of the present invention possess a biological component, which comprises an immobilized, or substantially immobilized, single bilayer membrane, and one or more membrane proteins incorporated therein; and a physical transducer, which serves to initiate the generated signal of the device.
  • the biosensor devices of the present invention may be augmented to additionally contain signal enhancers and signal multipliers. Likewise, co-factors, accessory proteins, and other relevant biomolecules may be incorporated into the biosensor.
  • the biological component of the biosensors of the present invention will typically comprise one, two, three, four or more membrane-bound proteins.
  • a single species of such protein may be incorporated into the membranes of the present invention, or, in alternative embodiments, two, three, four or more different species of receptor proteins.
  • Such additional species may have related functions, such as, for example, all being cytokine receptors, etc., or may have diverse functions, or may have subsets of receptors with related functions.
  • Different proteins (or sets of proteins) can be incorporated into membrane located at different regions of the support, so as to produce a solid support having a pattern of pores each containing different types of sensor protein.
  • membrane associated proteins can be incorporated in the membranes of the present invention.
  • biological components may be enzymes, or other proteins, whose expression is characteristic of disease (e.g., bone specific alkaline phosphatase, aldose reductase, myoglobin, troponin I, etc.). More preferably, however, they will be receptor molecules.
  • Suitable receptor molecules include cell-surface receptors (including protein-tyrosine kinase receptors (e.g., EGFR, PDGFR, MCSFR, SCFR, insulin-R, VEGFR, Trk, Met, Ron, Axl, Eph), ion channel receptors; or receptors for TNF and related factors (e.g., Trk, Met, Ron, Axl, Eph, Fas, TNFRI, TNFRII, CD40, CD30, CD27, 4-1BB, LNGFR, OX40), serine-threonine kinase receptors (e.g., TGF ⁇ R), transmembrane 7 or G protein-coupled receptors (e.g., receptors for CCR1, CCR2 ⁇ , ⁇ , CCR3, CCR4, CCR5, CXCR1, CXCR2, CXCR3, CXCR4, BLR1, BLR2, V28, and class I and class II cytokines), CD4 + receptors, class I (hematopoi
  • pylori or M. tuberculosi, hepatitis virus, rubella, CMV or immunodeficiency virus (HIV, FIV), prostate specific antigen, etc.); or membrane- associated antibodies to such antigens, or autoimmune immunoglobulins, thyroglobulin, anti-thyroglobulin, IgE, IgG, or IgM immunoglobulins, tumor markers (e.g., prostate specific antigen, AFP CEA, etc.). It is, however particularly preferred that such biological molecule be a
  • GPCR so as to form a biosensor capable of detecting ligands that affect GPCR activity.
  • the G protein peptides simulating elements of G-protein function will be included in the reconstituted membrane preparations so as to be present in the aqueous cushion separating the bilayer from the support.
  • the external side of the bilayer will exhibit the attributes of the external side of a biological membrane.
  • the G protein will not be included in the reconstituted membrane preparation, but will be provided in the buffer being flushed through the support.
  • the external side of the bilayer will exhibit the attributes of the internal side of a biological membrane.
  • the biological component(s) can be recombinantly produced (i.e., produced in a heterologous host cell), genetically produced (i.e., produced in a homologous host cell), or synthetically produced (i.e., synthesized using in vitro chemical synthetic procedures).
  • Such biological component(s) can be incorporated into the bilayer membranes of the present invention by any of a variety of means, such as by reconstitution or by extrusion of spheroplasts membranes, as discussed above.
  • the physical transducer component of the biosensors of the present invention is preferably configured to exploit the open pore nature of the preferred solid support (i.e., Anopore®), so as to permit detection of signal by flowing a fluid through the pores of the support.
  • the physical transducer component of the biosensors of the present invention preferably comprises an optical sensor, such as a colorimetric (i.e., a sensor that modulates the production of a visible spectrum optical signal in response to target molecule binding), a non- visible spectrum optical sensor (i.e., a sensor that modulates the production of a non-visible spectrum optical (e.g., infra-red, UV, fluorescent, etc.) signal in response to target molecule binding).
  • a colorimetric i.e., a sensor that modulates the production of a visible spectrum optical signal in response to target molecule binding
  • a non-visible spectrum optical sensor i.e., a sensor that modulates the production of a non-visible spectrum optical (e.g., in
  • the use of the translucent AnoporeTM support facilitates the use of colorimetric and non-visible spectrum optical sensors.
  • An optical sensor that detects fluorescent light is particularly preferred.
  • Receptor activation may be sensed by retention/release of radioactive substances such as tritium labeled receptor agonists or antagonists.
  • the physical transducer component of the biosensor of the present invention may be configured as an electrochemical sensor (e.g., an amperometric sensor (i.e., a sensor that modulates a current in response to target molecule binding), a potentiometric sensor (i.e., a sensor that modulates a pH change in response to target molecule binding), a conductometric sensor (i.e., a sensor that modulates resistance in response to target molecule binding), or a piezoelectrical sensor (i.e., a sensor that modulates a piezoelectric response to target molecule binding).
  • an electrochemical sensor e.g., an amperometric sensor (i.e., a sensor that modulates a current in response to target molecule binding), a potentiometric sensor (i.e., a sensor that modulates a pH change in response to target molecule binding), a conductometric sensor (i.e., a sensor that modulates resistance in response to target molecule binding), or a pie
  • the signal generated by the physical transducer may be quantitative (i.e., varying in intensity, duration, etc., with the concentration of the target molecule in a sample), qualitative (i.e., producing a signal in response to a threshold concentration of the target molecule), or partially qualitative and partially quantitative.
  • the biosensor devices of the present invention may be employed in ligand binding studies to detect ligand concentrations in a sample.
  • ⁇ -, 2 H, 13 C, and 31 P NMR with and without "magic angle” spinning can be used to verify that the facile sample preparation method of the present invention results in the attachment of single lipid bilayers to the solid support.
  • MAS NMR spectra are simpler, better resolved, and have larger signal amplitudes, as only the isotropic chemical shift and / couplings contribute to the resonance frequency.
  • Chemical shift anisotropy and dipole-dipole couplings are averaged by fast tumbling of the molecules in solution. The physical reason for this averaging is that the orientation of the molecule with respect to the static field changes on a timescale that is faster than the inverse anisotropy of the interactions. Such an irregular fast tumbling of the sample inside the spectrometer cannot be realized technically. However, as was shown by Lowe, I.J. (1959) ("FREE
  • the technique is called “magic angle spinning” (MAS) (see, STRUCTURE DETERMINATION II (NMR, EPR), G. Jeschke (WS 2003/04 9) "High-Resolution Solid-State NMR: Magic Angle Sample Spinning and Cross Polarization," web site: mpip-mainz.mpg.de/ ⁇ jeschke/lect9.pdf).
  • the preferred solid-supported membranes of the present invention are separated from the solid surface by a thick water cushion over most of their area. Therefore it can be expected that proteins in reconstituted membranes, prepared according to our procedure, will function without any interference from the solid support. Furthermore, the present sample preparation procedure will raise sensitivity of assays by up to three orders of magnitude per unit of biochip area. It is therefore envisioned that the invention has utility in the production of biosensors for pharmaceutical applications, environmental screening, detection of warfare agents, etc.
  • the solid supported membranes can be used for rapid screening of binding affinity of drugs to membrane receptors, e.g. G-protein coupled membrane receptors (GPCR).
  • GPCR G-protein coupled membrane receptors
  • the solid substrate is compatible with optical studies, radiotracer binding studies, NMR, EPR, small angle neutron scattering, x-ray diffraction, calorimetry, and other methods, permitting the development of binding assays as well as structural studies on proteins and bound ligands.
  • l-Palmitoyl-2-oleoyl-sn-glycero-3 phosphocholine (POPC), 1- palmitoyl(d 3 ⁇ )-2-oleoyl-sn-glycero-3 phosphocholine (POPC-d 3 ⁇ ), l-palmitoyl-2- oleoyl-sn-glycero-3 phosphocholine-d (POPC-d 4 ), and 1,2-dimyristoyl-sn- glycero-3 -phosphocholine (DMPC) were purchased from Avanti Polar Lipids (Alabaster, AL).
  • D 2 O (99.8% D) and deuterium-depleted H 2 O (2-3 ppm D) were purchased from Cambridge Isotope Labs (Andover, MA).
  • Anopore aluminum oxide filters (13 mm diameter) with nominal pore sizes of 0.02, 0.1, and 0.02 ⁇ m were purchased from SPI Supplies (West Chester, PA).
  • Praseodymium (III) nitrate hexahydrate (99.9%) was purchased from Aldrich (Milwaukee, WI).
  • Polyethylene glycol 8000 (PEG8000) with an average molecular weight of 8455 g/mol was purchased from Spectrum.
  • Piperazine-N,N'-bis(ethanesulfonic acid) sesquisodium salt (Calbiochem; San Diego, CA) was used to prepare 10 mM PIPES buffer with 100 mM NaCl (Sigma; St. Louis, MO) in deuterium-depleted water for use in 2 H NMR measurements on POPC-d 4 .
  • lipids were loaded using one of two different extruders.
  • the majority of the experiments utilized a mini-extruder (Avanti; Alabaster, AL) equipped with 1-mL syringes.
  • One to five 13 -mm diameter anopore filters were placed into the extruder and flushed with 1 mL of water 11 times before bringing them into contact with the lipid.
  • the water was removed and 5 mg lipid dispersed in 1 mL of water, buffer, or aqueous PEG solution was passed through the filters 15 times.
  • the lipid dispersion was removed and one or two 1 mL aliquots of water, buffer, or aqueous PEG solution was passed through the filters at an rates of 3 - 12 mL/min, either by hand or using an infusion pump, to remove any remaining liposomes and to remove additional bilayers that are loosely attached to the first bilayer at the AAO surface.
  • the filters were then flushed with 1 mL of an isotonic solution (100 mOsmole) of NaCl in D 2 O at a rate of 0.04 mL/min using an infusion pump to remove any PEG not trapped beneath the bilayers.
  • the extrusion was performed at room temperature, but for DMPC, the lipid and extruder were maintained at 35 °C during loading using a dry heat incubator (Fisher Scientific; Pittsburgh, PA).
  • H spectra are acquired at a resonance frequency of 300.14 MHz with a 3.6 ⁇ s 90°pulse and a 4 s delay between scans.
  • the spectral width is 5 kHz and the number of acquisitions is 32.
  • P spectra are acquired at a resonance frequency of 121.4 MHz with a 2.5 ⁇ s 90°pulse and a repetition rate of one acquisition per second.
  • the spectral width is 50 kHz, and the number of acquisitions varied from 25,000 to 90,000.
  • the data are transferred to a personal computer and processed as described by Holte, L.L. et al. ((1995) " 2 H NUCLEAR MAGNETIC RESONANCE ORDER PARAMETER PROFILES SUGGEST A CHANGE OF MOLECULAR SHAPE FOR PHOSPHATIDYLCHOLINES CONTAINING A POLYUNSATURATED ACYL CHAIN," Biophys. J. 68:2396-2403) and Huster, D. et al. ((1998) "INFLUENCE OF
  • Diffusion measurements are conducted at sixteen different values of gradient strength varying from 0.01 - 0.37 T/m with a stimulated echo sequence using sine-shaped bipolar gradient pulses (Cotts, R.M. et al. (1989) J. Magn.Reson. 83:252-266) of 5 ms duration. A longitudinal eddy current delay of 5 ms is used. Diffusion times are varied from 20 - 200 ms. At every gradient strength, 64 scans are acquired with a recycle delay of 4 s.
  • Mathcad is used to fit the diffusion data to the equation that relates signal intensity to the diffusion constant for powder samples with filter pores oriented at random to the orientation of the magnetic field gradient (Gaede, H.C. et al. (2003) "LATERAL DIFFUSION RATES OF LIPID, WATER, AND A HYDROPHOBIC DRUG IN A MULTILAMELLAR LIPOSOME,” Biophys J 85:1734-1740),
  • Equation (3) a formula for the calculation of the FID for a 2 H NMR experiment on a lipid bilayer is obtained. Resonances corresponding to all possible values of ⁇ and ⁇ are superimposed by integration:
  • y realn ⁇ J Jli cos(2 ⁇ v q • s( ⁇ , ⁇ ) • S(mo ⁇ )i • n-DW)-exp(-n-DW/T 2 ⁇ )p( ⁇ )d ⁇ d ⁇ (4) i 0 0 0
  • the molecular order parameters S(mol)i were determined by dePakeing of spectra obtained by 2 H NMR experiments on randomly oriented bilayers (multilamellar liposomes) (Sternin, E. et al. (1983) J. Magn.Reson. 55:274-282; Mccabe, MA. et al. (1995) J. Magn. Reson. Ser.B 106:80-82).
  • the angular distribution function p( ⁇ ) and the line broadening, T 2 ⁇ > were adjusted to match experimental signal intensities.
  • the value of ⁇ measured in units of degrees, represents the width of the Gaussian distribution function.
  • p( ⁇ ) sin ⁇ .
  • the simulation included an angular distribution function to describe the spread in bilayer orientation.
  • the bilayer normals have a circular distribution relative to the external magnetic field which was the starting point for the simulation.
  • a mosaic spread of cylinder axis orientation was considered.
  • T 2 ⁇ a second adjustable parameter
  • the spectra in both the MLVs and the single bilayers at AAO are a superposition of two quadrupolar splittings with values of 6.3 and 5.2 kHz, corresponding to the quadrupolar splittings of POPC choline ⁇ and ⁇ resonances, respectively (Koenig, B.W. et al. (1996) Langmuir 12:1343-1350).
  • the extra signal in the center of the experimental spectrum of lipid adsorbed to AAO appears to be mostly from a residual 2 H resonance of AAO hydroxyl groups.
  • there was an additional small but systematic deviation in signal intensity between measured and calculated spectra in the frequency range of +5 kHz that could indicate existence of a few percent of lipid with lower, but not well defined headgroup order.
  • Lipids 6:343-350 This commonly used assay was employed here to determine whether the membranes formed were well-sealed and if they formed single or multiple bilayers. Addition of the shift reagent Pr 3+ at a concentration of 5 mM after adsorption of bilayers to AAO pores, either by dropwise addition or slow extrusion at rates of 0.02 mL/minute, shifted a fraction of the ⁇ choline resonance downfield by 0.12 ppm (Bystrov, V.F. et al. (1971) Chem.Phys. Lipids 6:343-350).
  • the seal was also perturbed by higher flow rates of water.
  • Pr 3+ shift reagent was applied by extrusion of a Pr 3+ solution through the AAO pores at rates of 12 mL/minute, then more than 50% of the ⁇ -choline resonances were shifted. Often two or more ⁇ choline peaks were detected, indicating that different regions of the monolayers were exposed to different concentrations of the shift reagent. However, extrusion at rates of 0.02 mL/minute resulted again in 1:1 peak ratios.
  • Fluid-gel phase transitions of lipids To probe the influence of the AAO support on the phase transition temperature of supported lipids, ⁇ MAS NMR spectra of DMPC supported on AAO pores were acquired as a function of temperature. At a MAS frequency of 5 kHz the ! H resonances of lipid hydrocarbon chains are well resolved in the fluid phase but broadened beyond detection in the gel state. To follow the main phase transition of DMPC, the normalized intensity of the 100 Hz line-broadened methylene resonance at 1.3 ppm was plotted versus temperature in Figure 8 for both DMPC in multilamellar liposomes and for single DMPC bilayers adsorbed on AAO pores.
  • the water resonance is a superposition of signals of at least two water pools. At our experimental conditions two thirds of water resides outside pores while one third is inside.
  • the signal decay of the water resonance as a function of gradients strength is complex due to the random orientation of filter pieces as well as water- filled pores in the spinning rotor, water interaction with lipid and AAO surfaces, chemical exchange of protons with AAO hydroxyl groups, and permeation of water through lipid bilayers. From the signal decay at low gradient strength it was estimated that within very generous error limits most of the water moves at the rate of free water, D ⁇ 2 TO "9 m 2 /s .
  • the width of the angular distribution of 20° must result predominantly from a distribution in bilayer orientations rather than the AAO surface, though it is likely to contain some contributions from the pores, such as deviations from parallelism, roughness along the long axis, and distortion from cylindrical shape.
  • the lipid bilayers must maintain curvature in excess to curvature from the cylindrical symmetry, suggesting that bilayers do not adhere flatly to the inner surface of the pores.
  • the T 2 value for the proton ⁇ resonance of AAO-adsorbed POPC of 8.8 ms may be compared to 97 ms, the value for the ⁇ -resonance of DMPC MLVs at 30°C (Huster, D. et al. (1999) J. Phys. Chem. B 103:243-251).
  • the reduced ! H NMR T 2 for POPC on AAO suggests an additional reorientation of the lipids with correlation times in the millisecond range.
  • a POPC lateral diffusion constant of 9.5 X 10 "12 m 2 /s see Fillipov, A. et al. (2002) Biophys. J.
  • the order parameters of the lipid choline resonances of the outer monolayers at the AAO surface are identical to order parameters in the inner monolayer. Therefore the headgroup order and motions of the majority of lipids in the outer monolayer adjacent to the AAO surface are not influenced by the association, in good agreement with existence of a thick water layer between AAO and lipid underneath most of the bilayer. The data are consistent with only a small percentage of the lipids acting as points of attachment to the AAO surface.
  • Lipid order parameters and phase transitions The deuterium spectra of the perdeuterated palmitic acid chain in POPC-d 3 ⁇ were used to assess order and dynamics of lipids in adsorbed single bilayers.
  • the molecular chain order parameters S(mol)i are indistinguishable from order parameters in multilamellar liposomes, which is remarkable considering the high sensitivity of order parameters to changes in lipid area per molecule as demonstrated in experiments conducted as a function of temperature or hydration.
  • E m J/mol equivalent to a lowering of the main lipid phase transition temperature AT - — ⁇ T j of about 0.9
  • Multilamellar liposomes become cylindrical after being forced into the pores.
  • the cylinders are stable until their length reaches a critical value that depends on the surface bending energy.
  • Cylinders with variable diameter have lowest energy if they belong to the family of surfaces with constant total curvature J, called Delaunay surfaces (Delauney, C.
  • lipid cylinders have a tendency to form constrictions and to break up at a certain length, forming short, capped tubules or spherical vesicles.
  • Rhodopsin purification and reconstitution into POPC membranes Rod outer segment discs from bovine retinas were solubilized in the detergent octylglucoside (OG), and the rhodopsin purified by affinity chromatography according to the procedure of Litman et al. (Litman BJ. (1982) "PURIFICATION OF RHODOPSIN BY CONCANAVALIN A AFFINITY CHROMATOGRAPHY,” Methods Enzymology, 81, 150- 153) using a Pharmacia concanavaline A columnm (Pharmacia Biotech, Piscataway, NJ).
  • Rhodopsin concentration after purification was determined by measuring light adsorption at 500 nm using a diode array UV/Vis spectrophotometer Agilent 8453 (Agilent Waldbronn, Germany). Care was taken to minimize exposure of samples to light during the experiment.
  • the lipid dispersion was added to the purified rhodopsin dispersed in OG to yield a rhodopsin/lipid molar ratio in the range from 1/100 to 1/1000.
  • the OG concentration was kept at a concentration in the range from 40-100 mM OG and the OG/lipid molar ratio was 10/1.
  • Small liposomes with membrane incorporated rhodopsin were formed according to the dilution/reconstitution method of Jackson and Litman (Jackson M.L. and Litman, BJ. (1985) "RHODOPSIN-EGG PHOSPHATIDYLCHOLINE RECONSTITUTION BY AN OCTYL GLUCOSIDE DILUTION PROCEDURE," Biochim.
  • a mini-extruder (Avanti Polar Lipids, Alabaster, AL) or stainless steel thermobarrel extruder (Lipex Biomembranes, Inc; Vancouver, BC Canada) were loaded with up to five 13 mm diameter Whatman Anopore filters (SPI Supplies, West Chester, PA) with a nominal pore diameter of 0.2 ⁇ m.
  • Anopore filters were packed between two filter supports.
  • a total of 5 ml of rhodopsin reconstituted into POPC-d 3 ⁇ liposomes were sent through the filters in 1 ml increments at a rate of 0.06 ml/s using a gas-tight Hamilton syringe.
  • the rhodopsin from the 4 th milliliter was partially adsorbed (rhodopsin concentration 0.19 mg/ml), and the rhodopsin from the 5 th milliliter passed the filters essentially without binding within anopore (rhodopsin concentration 0.35 mg/ml, compared to 0.37 mg/ml in the incoming solution).
  • rhodopsin concentration 0.35 mg/ml 0.35 mg/ml, compared to 0.37 mg/ml in the incoming solution.
  • a total of 1.3 mg of rhodopsin did bind within the 5 anopore disks, corresponding to 0.26 mg of rhodopsin in 0.52 mg of POPC-d 3 ⁇ bilayers per disk. Again, extrusion was performed in complete darkness to prevent the bleaching of rhodopsin.
  • the Anopore membranes were placed on a stainless steel grid and sealed against the upper barrel by an O-ring. In this approach, water was flushed through the filters several times before the lipid dispersion was passed through ten times using compressed argon at pressures in the range from 2 - 8 bar.
  • the anopore filter disks with rhodopsin in lipid membranes were loaded into a flat glass cell that was filled with PIPES/NaCl buffer prepared in deuterium depleted water (Cambridge Isotopes, Cambridge MA) and sealed with a silicon stopper. The cell was inserted in darkness into a Doty flat cell probe for a DMX500 solid-state NMR.
  • the carrier frequency of the spectrometer was adjusted to be exactly at the center of the symmetric spectra.
  • FID free induction decay
  • the location of the maximum of the quadrupolar echo was determined with a resolution of l/10 th of a dwell time unit, and the time base of the spectra was corrected such that the FID began exactly at the echo maximum using a spline interpolation function to calculate new digital data points.
  • the FID was multiplied with an exponential decaying window function corresponding to a line broadening of 100 Hz. After the Fourier transformation of the FID the spectra shown in Figure 11 and Figures 12A-H were obtained.
  • the spectra are characteristic for lipid bilayers oriented preferentially with their bilayer normal perpendicular to the magnetic field. This orientation of the bilayer normal is consistent with membranes adhering to the inner surface of pores as lipid tubules. With increasing amounts of protein in the samples we observed an increase of the linewidth of resonances and some increase of mosaic spread. The mosaic spread of preferred bilayer orientations was reduced after freezing the sample in a deep freezer before investigating it at ambient temperature. Exact values of sn-1 chain order parameters and of mosaic spread were determined by simulating the experimental spectra using a program written for Mathcad 200 li Professional (MathSoft Engineering & Education, Inc., Cambridge, MA).
  • Order parameter analysis revealed a small reduction of hydrocarbon chain order in the order parameter plateau region (carbon atoms 2-8) due to the presence of the protein (see Figure 13).
  • Mosaic spread could be reasonably well modeled as Gaussian distribution with a half width of 8 degrees (pure 18:0(d35)-22:6 PC bilayers) to about 20 degrees (18:0(d35)-22:6 PC bilayers containing rhodopsin at a lip id/protein molar ratio of 100/1.
  • the presence of the protein in the reconstituted membranes is also detectable as a substantial reduction of spin-spin relaxation times (broadening of resonance lines) and small reductions of spin-lattice relaxation times of rapidly moving lipid segments.
  • the anopore filters containing reconstituted rhodpsin had a bright pink color that turned to yellow within one minute after exposure to light. This transition is consistent with conversion of dark-adapoted rhodopsin to a meta- I/meta-II rhodopsin equilibrium, demonstrating that rhodopsin was successfully adsorbed and functional.
  • Example 3 Adsorption of E. coli Protoplast Membranes with Incorporated Human Peripheral Cannabinoid Receptor (CB2) into AAO Pores Human peripheral cannabinoid receptor (CB2) was expressed in E. coli BL21 cells as a fusion protein, containing E. coli maltose-binding protein (MBP) attached at the N-terminal end of CB2, and thioredoxin followed by a stretch often histidine residues attached at the C-terminal end of CB2. Cell density was of the order of 10 9 cells per ml. Every cell expressed about 1,000 copies of CB2.
  • MBP E. coli maltose-binding protein
  • E-coli cytoplasmic membrane preparation By Western Blot analysis using specific antibodies we established that CB2 is preferentially located in the cytoplasmic membranes of E. coli. Formation of spheroplasts and fractionation to obtain cytoplasmic membranes were conducted according to the protocols by R.L. Weiss (Weiss, R.L. (1976) "PROTOPLAST FORMATION IN ESCHERICHIA COLI," J. Bacteriol. 128:668-670) and Thai and Kaplan (Tai, S.-P. et al.
  • E. coli cells were collected by centrifugation, washed twice with 0.1 M Tris-HCl pH 8.0 buffer and re-suspended in 0.1 M Tris-HCl buffer pH 8.0 containing 20% (w/v) sucrose, such that the resulting cell suspension had an optical density of 10 at 600 nm.
  • a cocktail of protease inhibitors F. Hoffmann-La Roche Ltd, Basel, Switzerland was added to prevent enzymatic digestion of CB2.
  • the temperature was adjusted to 37° C, and a solution of lysozyme (2 mg/ml) was added slowly, under constant stirring, until a final lysozyme concentration of 0.1 mg/ml was reached.
  • the cell suspension was incubated at 37° C for additional 15 minutes.
  • a solution of 0.1 M EDTA pH 7.0 was added slowly under continuous stirring, until a final EDTA concentration of 10 mM was reached. Incubation continued for another 10 minutes.
  • the spheroplasts were centrifuged at 12,000 g for 20 minutes, the pellet collected and washed once with 0.1 M Tris-HCl buffer, pH 8.0, containing 20% sucrose. Spheroplasts were centrifuged again at 12,000 for 20 minutes. The pellet of spheroplasts was re-suspended in ice-cold water, resulting in osmotic lysis of sphoroplasts.
  • MAS NMR experiments one milliliter of the cytoplasmic membrane preparation was suspended in Tris buffer prepared in 99.9% D 2 O (Cambridge Isotopes, Cambridge MA) and pelleted at 150,000 g for thirty minutes. The pellet with a volume of approximately 15 ⁇ L was transferred by centrifugation to a 4 mm outer diameter MAS rotor outfitted with a Kel-F insert to generate a spherical sample volume of 11 ⁇ L (Bruker Biospin Inc., Billerica MA).
  • the sample was spun at a MAS frequency of 5 kHz using a Bruker H-X resonance MAS probe for a solid state DMX300 NMR spectrometer (Bruker Biospin, Billerica MA) equipped with a Bruker widebore 300 MHz magnet.
  • Proton resonance spectra were acquired at ambient temperature using a ⁇ /2 pulse length of 4 ⁇ s and Cyclops phase cycling. About 1,000 free induction decays (FID) with a relaxation delay of 4 s were acquired. Before Fourier transformation the FID was multiplied with an exponential window function equivalent to a signal broadening of 1Hz.
  • Radioligand [ 3 H]CP55,940 bound to the CB2 receptor deposited into the Anopore filters could be competed off by increased concentrations of unlabeled ligand CP 55,940.
  • Ligand-binding parameters determined for the CB2 receptor deposited into Anopore filters were identical to the ligand-binding characteristics of CB2 receptor measured by a conventional filter-binding assay ( Figure 15).
  • the conventional competitive filter-binding assay was performed by incubating a suspension of E. coli membranes with radioligand [3H]CP55,940 and variable concentrations of the unlabeled competing ligand. Upon incubation, the reaction mixture was rapidly filtered through Whatman GF/B paper filters, and retained activity on the filters was counted with a scintillation counter.
  • This assay typically results in significant nonspecific binding of the hydrophobic ligand (CP55,940) in the multilamellar deposits on the filter surface which requires to work with the active receptor at much higher concentrations and reduces accuracy and reproducibility of the binding parameters K and B max . In the convential assay nonspecific binding constitutes 40-50% of the total radioactivity count.
  • Example 5 Reconstitution of Purified Recombinant Cannabinoid Receptor, Adsorption into AAO Pores, and Detection of Membrane Adsorption by Fluorescence Spectroscopy Human peripheral cannabinoid receptor (CB2) was expressed in E. coli BL21 cells as a fusion protein, containing E. coli maltose-binding protein (MBP) attached at the N-terminal end of CB2, and thioredoxin followed by a stretch often histidine residues attached at the C-terminal end of CB2.
  • MBP E. coli maltose-binding protein
  • Recombinant protein was solubilized from the bacterial membranes in a mixture of 0.5% CHAPS, 0.1 % cholesteryl hemisuccinate and 1% of dodecylmaltoside, and purified to approximately 90% purity by affinity chromatography on Ni-agarose (Qiagen) and ion-exchange chromatography on HiTrap-Q Sepharose (GE-Amersham Biosciences).
  • fusion protein was covalently labeled with AlexaFluoro 532 carboxylic acid, succinimidyl ester (Molecular Probes) according to the protocol recommended by the manufacturer.
  • Non-reacted fluorescent dye was (partially) removed by sequential gel-filtration (PD-10 desalting columns, Amersham), centrifugation in the Centricon-30 filter device (TVIillipore) and dialysis (Slide-A-Lyzer, Pierce). About 200 ⁇ g of labeled protein was obtained.
  • lipid 150 ⁇ g of l-stearoyl-2-oleoyl-sn-glycero-3 -phosphocholine (SOPC, Avanti) were dissolved in methanol and mixed with a methanol solution of 1.5 ⁇ g of Texas Red l,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine, triethylammonium salt (Texas Red DHPE, Molecular Probes). The methanol was removed under the stream of nitrogen, and lipids were dispersed in a solution of 3% octylglucoside (OG) in 50 mM Tris-HCl buffer pH 7.5.
  • OG octylglucoside
  • Filters containing a mixture of fluorescence labeled CB2 and SOPC produced bright fluorescence at 580 nm (excitation at 532 nm), characteristic for AlexaFluoro 532 fluorophore. Intensity of fluorescence of sample 3 containing labeled CB2 as well as labeled lipid was lower than the fluorescence of sample 1 which contained labeled CB2 and non-labeled lipid, the effect that can possibly be attributed to quenching of fluorescence of the AlexaFluoro532 fluorophore by the Texas Red.
  • Quantification of the amounts of lipid and CB2 protein deposited onto Anopore filters was performed as follows.
  • SOPC was mixed with the Dil-Cl 8 DilC i s ( 1 , 1 ' -dioctadecyl-3 , 3 ,3 ' ,3 ' -tetramethy lindodicarbocyanine, 4- chlorobenzenesulfonate salt) fluorescently labeled lipid at a ratio of 1000: 1 (w/w, SOPC to Dil-Cl 8) in either 3% OG or mixture of triple detergents (0.5% CHAPS, 0.1% CHS, 0.1% DM).
  • SOPC/ Dil C18 mix was rapidly diluted (20-fold) into 50 mM Tris-HCl buffer pH 7.5, and solution was slowly filtered through the Anopore filter.
  • the fluorescence of the lipid retained on the filter was measured by scanning of the wet filter on Typhoon 8600 fluorescence scanner at following settings: excitation: 633 nm, emission: 670 nm, PMT: 450, sensitivity: normal.
  • FIG. 21 demonstrates that CB2 receptor can be eluted from the Anopore filter in the presence of strong ionic detergent SDS, and that no degradation of the receptor occurred during deposition/ elution procedures.

Abstract

This invention relates to reagents and methods for forming peripheral and integral membrane protein containing membranes inside a porous solid support for use in biosensors. The present invention particularly concerns the development of a procedure for depositing single layered membranes inside those pores so as to provide a total exposed pore surface area that is orders of magnitude larger than the corresponding area of the filter.

Description

Title of the Invention:
Solid-Supported Membranes Inside Porous Substrates and Their Use in Biosensors
Field of the Invention: This invention relates to reagents and methods for forming membranes that may be deposited into porous solid supports for use in biosensors.
Cross-Reference to Related Applications:
This application claims priority to U.S. Patent Applications Serial Nos. 60/534,380 (filed January 6, 2004) and 60/578,067 (filed June 9, 2004) both herein incorporated by reference in their entirety.
Statement of Governmental Interest:
This invention was funded by the National Institute of Alcohol Abuse and Alcoholism at the National Institutes of Health. The United States Government has certain rights to this invention.
Background of the Invention:
Immobilized Membranes and Biosensors Immobilized membranes may be employed in a wide range of applications, including enabling biofunctionalization of inorganic solids (semiconductors, gold- covered surfaces, and optoelectronic devices) and polymeric materials; providing a natural, non-denaturing, and defined environment for the immobilization of biomolecules; and allowing the preparation of ultrathin, high-electric-resistance layers on conductors and the incorporation of receptors into insulating layers for the design of biosensors based on electrical and optical detection of ligand binding (Ho, W.S. (2003) "REMOVAL AND RECOVERY OF METALS AND OTHER MATERIALS BY SUPPORTED LIQUID MEMBRANES WITH STRIP DISPERSION," Ann N Y Acad Sci. 984:97-122; Groves, J.T. (2002) "MEMBRANE ARRAY TECHNOLOGY FOR DRUG DISCOVERY," Curr Opin Drug Discov Devel. 5(4):606-612; Sinner, E.K., (2001) "FUNCTIONAL TETHERED MEMBRANES," Curr Opin Chem Biol. 5(6):705-711 ; Jonsson, J.A. et al. (2000) "MEMBRANE-BASED TECHNIQUES FOR SAMPLE ENRICHMENT," J Chromatogr A. 902(l):205-225; Schuster, B. et al. (2000) "S- LAYER-SUPPORTED LIPID MEMBRANES," J Biotechnol. 74(3): 233-254; Knoll, W. et al. (2000) "FUNCTIONAL TETHERED LIPID BILAYERS," J Biotechnol. 74(3) : 137- 158; Boxer, S.G. (2000) "MOLECULAR TRANSPORT AND ORGANIZATION IN SUPPORTED LIPID MEMBRANES," Curr Opin Chem Biol. 4(6): 704-709; Sackmann, E. et al. (2000) "SUPPORTED MEMBRANES ON SOFT POLYMER CUSHIONS: FABRICATION, CHARACTERIZATION AND APPLICATIONS," Trends Biotechnol. 18(2):58-64; Sackmann, E. (1996) "SUPPORTED MEMBRANES: SCIENTIFIC AND PRACTICAL APPLICATIONS," Science 271:43-48; Tien, H.T. etal. (1990) "SELF- ASSEMBLING BILAYER LIPID MEMBRANES ON SOLID SUPPORT," Biotechnol Appl Biochem. 12(5):478-84).
Immobilized membranes are particularly suitable in the production of "Biosensors." Biosensors are analytical devices that are used to measure the presence and/or concentration of desired biological molecules in a sample. Depending upon the biosensor format employed, any of a wide variety of biological molecules non-denaturing, and defined environment for the immobilization of biomolecules (such as antigens, hormone receptors, enzymes, cytokine receptors, antibodies, etc.); can be analyzed using biosensors (see, e.g., Keusgen M. (2002) "BIOSENSORS: NEW APPROACHES IN DRUG DISCOVERY," Naturwissenschaften. 89(10):433-44. Epub 2002 Sep 11; Albers, J. et al. (2003) "ELECTRICAL BIOCHIP TECHNOLOGY~A TOOL FOR MICROARRAYS AND
CONTINUOUS MONITORING," Anal Bioanal Chem. 377(3):521-527. Epub 2003 Aug 30; Brenner- Weiss, G. et al. (2003) "APPROACHES TO BIORESPONSE-LINKED INSTRUMENTAL ANALYSIS IN WATER ANALYSIS," Anal Bioanal Chem. 377(3):408- 416. Epub 2003 Aug 16; Stett, A. et al. (2003) "BIOLOGICAL APPLICATION OF MICROELECTRODE ARRAYS IN DRUG DISCOVERY AND BASIC RESEARCH," Anal Bioanal Chem. 377(3):486-495. Epub 2003 Aug 16; Baeumner, A.J. (2003) "BIOSENSORS FOR ENVIRONMENTAL POLLUTANTS AND FOOD CONTAMINANTS," Anal Bioanal Chem. 377(3):434-445. Epub 2003 Aug 12; Cosnier, S. (2003) "BIOSENSORS BASED ON ELECTROPOLYMERIZED FILMS: NEW TRENDS," Anal Bioanal Chem. 377(3): 507-520. Epub 2003 Aug 12; Lindholm-Sethson, B. et al. (2003) "ARE BIOSENSOR ARRAYS IN ONE MEMBRANE POSSIBLE? A COMBINATION OF MULTIFREQUENCY IMPEDANCE MEASUREMENTS AND CHEMOMETRICS," Anal Bioanal Chem. 377(3):478-485. Epub 2003 Aug 12; Dickert, F.L. et al. (2003) "SENSOR STRATEGIES FOR MICROORGANISM DETECTION-FROM PHYSICAL PRINCIPLES TO IMPRINTING PROCEDURES," Anal Bioanal Chem. 377(3):540-549. Epub 2003 Aug 12; D'Orazio, P. (2003) "BIOSENSORS IN CLINICAL CHEMISTRY," Clin Chim Acta. 334(l-2):41-69; Ligler, F.S. et al. (2003) "ARRAY BIOSENSOR FOR DETECTION OF TOXINS," Anal Bioanal Chem. 2003 Oct; 377(3): 469-477. Epub 2003 Jun 13; Nakamura, H. et al. (2003) "CURRENT RESEARCH ACTIVITY IN BIOSENSORS," Anal Bioanal Chem. 377(3):446-468. Epub 2003 Jun 13; Jain, K.K. (2003) "CURRENT STATUS OF MOLECULAR BIOSENSORS," Med Device Technol. 14(4):10-15; Kasianowicz, J.J. (2002) "NANOMETER-SCALE PORES: POTENTIAL APPLICATIONS FOR ANALYTE DETECTION AND DNA CHARACTERIZATION," Dis Markers. 18(4):185-191; Vercoutere, W. et al. (2002) "BIOSENSORS FOR DNA SEQUENCE DETECTION," Curr Opin Chem Biol. 6(6):816-822; Rishpon, J. (2002) "ELECTROCHEMICAL BIOSENSORS FOR ENVIRONMENTAL MONITORING," Rev
Environ Health 17(3):219-247; Kroger, S. et al. (2002) "BIOSENSORS FOR MARINE POLLUTION RESEARCH, MONITORING AND CONTROL," Mar Pollut Bull. 45(1- 12):24-34.
Biosensors function by generating a detectable physical signal from the sensor's physical transducer component in response to the binding of a target biological molecule to the sensor's biological component. The physical transducer component is typically an optical or electrical signal, e.g. a fluorescence signal elicited by the ligand binding event, a modulation of a current in response to binding, a pH change in response to binding, a modulation of electrical resistance in response to binding, etc. Care must be taken to ensure that the target biological molecule of the biological component of the biosensor is immobilized, and that the immobilization procedure forms a stable layer of biomolecules. The immobilization procedure must also not undesirably diminish the activity or structure of the target biological molecule of the biological component, or change its substrate reactivity (i.e., perturbations of the biological component is optimally minimized).
Several approaches have been developed in order to address these constraints. In the context of biosensors that employ immobilized lipid bilayer membranes, this is typically accomplished by supporting the membrane on a solid support, and introducing the target biological molecule into the membrane. Three types of supported membranes have been described. Integrated bilayers are characterized by an inner monolayer that is either covalently or ionically bonded to the support surface. Freely supported bilayers are characterized as having an inner monolayer that is separated from the support by an ultrathin water layer (~10 A). The third type of supported membrane consists of a bilayer membrane that rests on an ultrathin polymer film (see, Sackmann, E. (1996) "SUPPORTED MEMBRANES: SCIENTIFIC AND PRACTICAL APPLICATIONS," Science 271 :43-48). Such supported membranes can be associated with silicon microchips, or with beads (as in a column, etc.).
Immobilization of the biological molecule can be achieved by adsorbing the biological molecule onto the surface of a solid support, integrating the molecule within a gel or other matrix, or covalently coupling it to a solid support. Some approaches achieve immobilization by adsorbing the biological molecule onto the surface of a solid support, integrating the molecule within a gel or other matrix, or covalently coupling it to a solid support. Other approaches interpose a polymer cushion between the lipid bilayer and the polymer support. Supported lipid-protein bilayers separated from the solid surface by nanometer-thick water layers or ultrathin soft polymer cushions maintain the thermodynamic and structural properties of free bilayers. This enables the application of several surface-sensitive techniques (micro interferometry, ellipsometry, surface plasmon spectroscopy, Fourier transform infrared (FTIR) spectroscopy, nuclear magnetic resonance (NMR), and neutron and x-ray surface reflectivity).
Three basis support formats have been employed: flat membrane supports, bead supports and porous supports. Flat membrane supports have the advantage of being able to physically separate membranes (and thus provide protection against their disruption). They suffer from the disadvantage of low surface area. Beaded surfaces provide much greater surface area than flat membranes, however, the membranes can touch one another and are not protected. Porous supports provide very large surface area, but their application has suffered from issues related to uniformity and homogeneity of membrane preparation as well as from limited membrane accessibility for ligands that are delivered via the water phase.
Anodic oxidation of aluminum has been found to produce γ-alumina with a reproducible pore size, high pore density, and a narrow pore size distribution (Furneaux, R. C. et al. (1989) "GLUCOFURANOSYLATION WITH PENTA-O- PROPANOYL-BETA-D-GLUCOFURANOSE," Nature 337:147-149). The commercially available aluminum oxide (AAO) filters have a 60 μm thick support layer with pore diameters of 0.2 μm that is capped by a thin, ~1 μm layer with nominal pore diameters of 0.02, 0.1, or 0.2 μm. AAO filters have a variety of other applications, including support for cell cultures and microscopy, sample preparation for HPLC, IC, and electrophoresis, and liposome extrusion.
Lipids have previously been added to AAO as monolayers to change liquid crystal director orientation (Crawford, G.P. et al. (1991) "SURFACE-INDUCED ORIENTATIONAL ORDER IN THE ISOTROPIC PHASE OF A LIQUID-CRYSTAL MATERIAL," Phys.Rev.A 44:2558-2569), as pore-spanning bilayers (Hennesthal, C. et al. (2000) "PORE-SPANNING LIPID BILAYERS VISUALIZED BY SCANNING FORCE MICROSCOPY," J.Am.Chem.Soc. 122:8085-8086), as the upper leaflet of hybrid bilayers (Marchal, D. et al. (1998) "ELECTROCHEMICAL MEASUREMENT OF LATERAL DIFFUSION COEFFICIENTS OF UBIQUINONES AND PLASTOQUINONES OF VARIOUS ISOPRENOID CHAIN LENGTHS INCORPORATED IN MODEL BILAYERS," Biophys.J. 74:1937-1948, and as streptavidin-supported lipid bilayers (Proux- Delrouyre, et al. "FORMATION OF STREPTAVIDIN-SUPPORTED LIPID BILAYERS ON POROUS ANODIC ALUMINA: ELECTROCHEMICAL MONITORING OF TRIGGERED VESICLE FUSION," (2001) J.Am.Chem.Soc. 123:9176-9177). The EPR signal of bilayers doped with spin labels has been interpreted as proof for adhesion of lipid membranes to AAO (Smirnov, A.I. et al (2003) "SUBSTRATE-SUPPORTED LIPID NANOTUBE ARRAYS," J.Am.Chem.Soc. 125:8434-8435). Multilamellar lipid bilayers have been formed on AAO supports. Smirnov, A.I. et al (2003) "SUBSTRATE-SUPPORTED LIPID NANOTUBE ARRAYS," J Am Chem Soc. 125(28):8434-8435.
A variety of different solid supports have been described that possess the desired attributes of a membrane support. Lipid monolayers have been used to make inner pore surfaces hydrophobic so that they may be used for thermotropic liquid crystal display applications (Marchal, D. et al. (1998) "ELECTROCHEMICAL MEASUREMENT OF LATERAL DIFFUSION COEFFICIENTS OF UBIQUINONES AND PLASTOQUINONES OF VARIOUS ISOPRENOID CHAIN LENGTHS INCORPORATED IN MODEL BILAYERS," Biophys.J. 74: 1937-1948). Multilamellar lipid bilayers have been deposited in Anopore™ filters by the diffusion of small liposomes (Smirnov, A.I. et al. (2003) "SUBSTRATE-SUPPORTED LIPID NANOTUBE ARRAYS," JAm.Chem.Soc. 125:8434-8435).
G-Protein Coupled Receptors (GPCR) G-Protein Coupled Receptors (GPCR) convey signals from extracellular hormones and neurotransmitters to intracellular effectors and linked signaling pathways. G-Protein Coupled Receptors are a class of integral membrane proteins belonging to the "7TM" superfamily of transmembrane receptors. The GPCRs are characterized by the possession of seven intramembrane helices, and by domains that extend both into the extracellular environments and the cytosol (Wojcikiewicz, R.J. (2004) "REGULATED UBIQUITINATION OF PROTEINS IN GPCR-INITIATED SIGNALING PATHWAYS," Trends Pharmacol Sci. 25(1):35-41; Becker, O.M. et al. (2003) "MODELING THE 3D STRUCTURE OF GPCRS: ADVANCES AND APPLICATION To DRUG DISCOVERY," Curr Opin Drug Discov Devel. 6(3):353-61; Bockaert, J. et al. (2003) "THE 'MAGIC TAIL' OF G PROTEIN-COUPLED RECEPTORS: AN ANCHORAGE FOR FUNCTIONAL PROTEIN NETWORKS," FEBS Lett. 546(l):65-72; Hermans, E. (2003) "BIOCHEMICAL AND PHARMACOLOGICAL CONTROL OF THE MULTIPLICITY OF COUPLING AT G-PROTEIN-COUPLED RECEPTORS," Pharmacol Ther. 99(l):25-44).
The GPCRs use an amazing number of different domains both to bind their ligand and to activate G proteins. More than 150 GPCRs have been identified, in at least six families of proteins that show no sequence similarity. The fine-tuning of their coupling to G proteins is regulated by splicing, RNA editing and phosphorylation. The G-Protein Coupled Receptor trigger cellular processes through a conformational shift that is caused by the binding of the GPCR to a ligand molecule (Gether, U. et al. (2002) "STRUCTURAL BASIS FOR ACTIVATION OF G-PROTEIN-COUPLED RECEPTORS," Pharmacol Toxicol. 91(6):304-12). As a consequence of the conformational shift, the GPCR acquires the capacity to bind and cleave intracellular proteins ("G-proteins"). The release of the cleavage products into the cytosol triggers cellular processes to occur.
Examples of GPCRs include the receptors of the olfactory sensory epithelium that bind odorants and neurotransmitter receptors (e.g., the serotonin receptor), the cannabinoid receptor (Picone, R.P. et al. (2002) "LigAND BASED STRUCTURAL STUDIES OF THE CB1 CANNABINOID RECEPTOR," J Pept Res. 60(6):348-56; Onaivi, E.S. et al. (2002) "ENDOCANNABINOIDS AND CANNABINOID RECEPTOR GENETICS," Prog Neurobiol. 66(5):307-44), and the rhodopsin receptor (Ballesteros, J. et al. (2001) "G PROTEIN-COUPLED RECEPTOR DRUG DISCOVERY: IMPLICATIONS FROM THE CRYSTAL STRUCTURE OF RHODOPSIN," Curr Opin Drug Discov Devel. 4(5):561-74; Filipek, S. et al. (2003) "G PROTEIN-COUPLED RECEPTOR RHODOPSIN: APROSPECTUS," Annu Rev Physiol. 65:851-79. Epub 2002 May 01. G protein-coupled receptors (GPCRs) mediate the perception of smell, light, taste, and pain (Ahmad, S. et al. (2004) "NOVEL G PROTEIN-COUPLED RECEPTORS AS PAIN TARGETS," Curr Opin Investig Drugs. 5(l):67-70; Luca, S. et al. (2003) "THE CONFORMATION OF NEUROTENSIN BOUND TO ITS G PROTEIN- COUPLED RECEPTOr," Proc Natl Acad Sci U S A. 100(19):10706-11. Epub 2003 Sep 05). They are involved in signal recognition and cell communication and are some of the most important targets for drug development (Neubig, R.R. (2002) "REGULATORS OF G PROTEIN SIGNALING (RGS PROTEINS): NOVEL CENTRAL NERVOUS SYSTEM DRUG TARGETS," J Pept Res. 2002 Dec;60(6):312-6; Neubig, R.R. et al (2002) "REGULATORS OF G-PROTEIN SIGNALLING AS NEW CENTRAL NERVOUS SYSTEM DRUG TARGETS," Nat Rev Drug Discov. 1 (3): 187-97; Fang, Y. et al (2003) "G PROTEIN-COUPLED RECEPTOR MICROARRAYS FOR DRUG DISCOVERY," Drug Discov Today. 8(16):755-61; Conway, B.R. et al. (2002) "THE USE OF BIOSENSORS TO STUDY GPCR FUNCTION: APPLICATIONS FOR HIGH- CONTENT SCREENING," Receptors Channels. 8(5-6):331-41; Rees, S. et al. (2002) "GPCR DRUG DISCOVERY THROUGH THE EXPLOITATION OF ALLOSTERIC DRUG BINDING SITES," Receptors Channels. 8(5-6):261-8).
The microstructure of the membrane significantly impacts upon the properties of the biosensor. Single lipid bilayers that are directly immobilized to a solid support, or immobilized via a polymer or "hairy rod" cushion (see,
Sackmann, E. (1996) "SUPPORTED MEMBRANES: SCIENTIFIC AND PRACTICAL APPLICATIONS," Science 271:43-48) can be used in a wide array of different biosensor applications, but are difficult to prepare, and may exhibit undesired perturbation of the biological component that adversely affect the integrity of such biosensors. By immobilizing membranes onto beads, one can dramatically increase surface area (up to 1000 fold); however, such membranes are difficult to prepare and store, and may undesirably perturb the function of the biological component. Detection of ligand binding is also encumbered.
Unfortunately, the limitations of existing procedures for immobilizing membranes has encumbered the use of biosensors to analyze proteins, such as the GPCRs, that possess domains which extend beyond the cellular membrane. Biosensors involving GRCRs thus advantageously need to possess two "compartments" (one cushioning the receptors from contact with the solid support, and the other protecting the receptors from the milieu beyond the membrane. Immobilization methods that immobilize a GPCRs-containing membrane directly to a solid support cause conformational perturbations in the receptor's structure. Although such perturbation is in some respects ameliorated through the use of immobilization techniques that create polymer cushions between the immobilized bilayer and the solid support, the non-aqueous nature of such cushions creates its own set of perturbation concerns.
Thus, despite the efforts that have been expended to prepare supported membranes, a need remains for a procedure that will permit the immobilization of receptors and other proteins that would yield stable membranes that would not impair the function of membrane proteins, in particular those with large extracellular domains, and that would be useful to form biosensors having large surface area, easier manufacture, storage and use and that would be less affected by issues of perturbations. The present invention is directed to this and other goals.
Brief Description of the Figures:
Figure 1 shows a schematic representation of the lipid bilayer adsorbed to the inner surface of an AAO pore. The z-axis defines the direction of the pore, and the x-y axes define the plane of the filter. The bilayer normal of the membrane is given by D , the vector of the external magnetic field is B , and the angle between these vectors is given by θ. The angle between the pore axis and the magnetic field is defined by β. The orientation of the bilayer normal in the x-y plane is defined by the angle α.
Figure 2 shows a 2H NMR spectrum of POPC-d3ι of multilamellar liposomes (T=24.2°C) (Panel A); the dePaked spectrum (Panel B), and smoothed order profile of POPC-d3ι (Panel C). Figure 3 shows 2H NMR spectra (T=24.2°C) of POPC-d3ι adsorbed in AAO pores for β0=0 and 90°. Panel A: Unilamellar samples prepared by flushing the pores after lipid loading. Panel B: The simulated spectra produced with equation 4 and a Gaussian angular distribution function with σ=20°. The angular distribution function used in the fit, displaying the mosaic spread of the lipid, is shown below the spectra. For comparison, the angular distribution of powder, p(β)=sin(β), is shown with a dashed line. The order parameters used in the simulation were obtained from the POPC-d3ι MLV 2H NMR spectrum and the profile is shown in Figure 2, Panel C, Multilamellar samples prepared with no flushing step after lipid loading.
Figure 4 shows 2H NMR spectra of POPC-d4 adsorbed on AAO pores at βo=0° (left) and POPC-d4 MLVs (right) at T=24.2 °C, together with simulated spectra (below). Both samples show two quadrupolar splittings of 6.3 and 5.2 kHz, corresponding to POPC choline α and β resonances, respectively. The extra signal in the center of the experimental spectrum of lipid adsorbed in AAO pores appears to be mostly from a residual 2H resonance of AAO hydroxyl groups. The small deviation in signal intensity between measured and calculated spectra in the frequency range of ±5 kHz could indicate existence of a few percent of lipid with lower headgroup order. Figure 5 shows 300 MHz Η NMR MAS spectra at a rotor spinning frequency of 5 kHz and a temperature of 30°C. Trace A: POPC MLVs, Trace B: POPC in AAO pores, and Trace C: POPC in AAO pores exposed to 5 mM Pr3+. Inset: Expansion of γ-choline signal in POPC in AAO samples. The POPC resonance assignments are given in the lower spectrum. Figure 6 shows 31P MAS NMR spectra at a rotor spinning frequency of 5 kHz and a temperature of 30°C. Trace A: POPC MLVs, Trace B: POPC single bilayers adsorbed on AAO pores, and Trace C: POPC adsorbed onto AAO pores and exposed to Pr3+. Figure 7 shows a 500 MHz Η MAS spectrum at T = 30.0 °C and vr = 10 kHz of 11 wt% PEG8000 in D2O trapped beneath a single POPC bilayer adsorbed in aluminum oxide pores. The integral intensity of the superimposed β POPC +PEG methylene resonance at 3.57 ppm relative to the γ POPC resonance at 3.15 ppm is 8.6:9.
Figure 8 show the normalized methylene intensity plotted versus temperature for DMPC supported on AAO and DMPC MLVs. The lines are included as a guide to the eye.
Figure 9 shows 500 MHz Η PFG-MAS NMR diffusion measurements on crushed POPC/AAO with trapped PEG8000 at a spinning frequency of 10 kHz, temperature of 30.0 °C, and a diffusion time of 200 ms. Trace A: Water resonance and Trace B: choline resonances of spectra acquired at 16 different gradient strengths from 0.01 - 0.37 T/m. Trace C: The signal intensity decay of the choline resonance as a function of k, fit to Equation 1. Figure 10A shows a model for lipid adsorption consistent with the NMR data. A single bilayer forms a good seal with the AAO surface by the interaction of a small percentage of the lipids. The lipids adsorb as wavy tubules with an average length of 0.4 μm. These tubules posses undulation with a radius of curvature of 100 - 400 nm. Trapped between the tubules and the AAO surface are pockets of water with an average thickness of 3 nm. Figure 10B shows an illustration of shape of the lipid bilayer tubules inside the AAO pore.
Figure 11 shows the solid state 2H NMR spectrum of POPC -d3ι lipid tubules in anopore filters containing bovine rhodopsin at a lipid/protein molar ratio of 100/1. The anopore disks were aligned with their normal parallel to the B0 field of the NMR instrument. The spectrum indicates that bilayers adhere to the surface of the cylindrical pores as lipid tubules.
Figures 12A-H show 2H NMR spectra of 18:0(d35)-22:6 PC membranes containing bovine rhodopsin at protein/lipid molar ratios from zero to 1/100. On the left the experimental spectra and on the right the simulated spectra are shown. The simulation yields the chain order parameters as a function of rhodopsin concentration, the mosaic spread of bilayer orientations, and the resonance linewidth. Results confirm formation of lipid tubules containing reconstituted bovine rhodopsin.
Figure 13 shows chain order parameter profile of the sn-1 hydrocarbon chain in 18:0(d35)-22:6 PC as a function of rhodopsin concentration reported as a function of molar protein/lipid ratio. In the presence of rhodopsin a small reduction of sn-1 chain order parameters for carbon atoms 2-8 is observed. Figure 14 shows !H MAS NMR spectrum of a cytoplasmic membrane preparation of E. coli BL21 cells recorded at a MAS spinning frequency of 5 kHz at ambient temperature. Trace A: membrane pellet, Trace B. membranes deposited inside the pores of an anopore filter. Signal assignments: 1- terminal methyl groups of lipid hydrocarbon chains, 2- methylene groups of lipid hydrocarbon chains, 3- methylene groups next to double bonds in lipid hydrocarbon chains, 4- DMSO (minor component in one of the chemicals added for cytoplasmic membrane preparation; the DMSO was flushed out of the membranes deposited into anopore filters with buffer). 5- superimposed resonances of phosphatidylethanolamine- and phosphatidylglycerol headgroups. 6- residual H-OD resonance of water, 7- superimposed amid- and aromatic resonances of proteins).
Figure 15 shows a comparison of the conventional competitive filter- binding assay (Whatman GF/B filters) with ligand-binding performed on Anopore membranes. Results indicate that depositioning of the cannabinoid receptor in single tubular lipid bilayers at the inner surface of pores did not alter the ligand binding properties of the receptor. Furthermore, nonspecific binding of ligands was reduced significantly, allowing much more accurate ligand binding measurements. Figure 16 shows the use of Anopore filters in a ligand binding study. Shown are scintillation count rates for binding of CP55,940 to the cannabinoid receptor CB2. Depositioning of membranes as tubular lipid bilayers in Anopore filters reduced nonspectific binding to less than 15% of total radioactivity, therefore greatly increasing the accuracy and reproducibility of ligand binding assays. Each data point represents the average of three filters.
Figure 17 shows the reconstitution of the CB2 fusion protein into a SOPC lipid bilayer. In Panel A, Excitation: 532 nm (green laser); Emission: 580 nm. PMT: 320; Sensitivity: normal; In Panel B, Excitation: 532 nm (green laser); Emission: 610 nm. PMT: 320; Sensitivity: normal. Alexa 532: Excitatiion:532 nm; Emission: 561 nm; T Red: Excitation: 583 nm; emission: 602 nm. Expperimental samples: 150 ug of SOPC; 7 ug of CB2 dissolved in 3% OG; Dilution: 1:10 in 50 mM Tris-HCl pH 7.5. Legend: 1-Labeled (Alexa Fluor 532) CB2 + non-labeled SOPC; 2-Dye (Alexa532) + non-labeled SOPC; 3-Labeled (Alexa532) CB2 + labeled (Tx Red) SOPC; 4-Dye (Alexa532) + labeled (TxRed) SOPC; 5-Labeled (Alexa532) CB2; 6-Dye (Alexa532); 7-Labeled (Tx Red) SOPC.
Figure 18 shows the deposition of SOPC into Anodisk filters. Presented is the lipid fluorescence count as a function of SOPC concentration in the reconstitution mixture. Reconstitution from lipid dispersions in octylglucoside is significantly more efficient than reconstitution from the triple detergent mix that is used for protein solubilization.
Figure 19 shows the deposition of CB2/SOPC into Anodisk filters. Presented is the cannabinoid receptor CB2 fluorescence count as a function of CB2 concentration in the dispersion. CB2 binding per filter saturated at a concentration of 10 micrograms.
Figure 20 shows the deposition of SOPC/CB2 into Anodisk filters. Presented is the lipid fluorescence count as a function of cannabinoid receptor CB2 concentration in the reconstituted membranes. In the concentration range from 0 - 10 μg of CB2 a reduction of the amount of deposited lipid with increasing protein content was observed. The count rate remained constant at a protein content of 10 μg or higher.
Figure 21 illustrates the depositioning of CB2 into Anodisk filters using a Western blot with anti-MBP antibody. Presented is a Western Blot of recombinant cannabinoid receptor CB2 reconstituted into SOPC lipid bilayers before and after depositioning into Anopore porous filters. The results confirm that the membrane protein is retained inside the pores. There is no indication for damage to the protein from reconstitution and depositioning. 1 : CB2-POPC mixloaded onto Anopore Filter; 2, 2a: flowthrough solution/wash with Tris buffer; 3: CB2 eluted from the Anopore filter with 2% SDS
Summary of the Invention:
This invention relates to compositions and methods for forming membranes containing membrane proteins that may be deposited into solid supports for use in biosensors. The present invention particularly concerns the development of a procedure capable of forming a high surface area, supported single-lipid bilayer in which the membrane is separated from the support by a closed and stable aqueous cushion.
The present invention particularly concerns the development of a procedure for depositing single lipid membranes onto a solid support so as to provide, for example, a total exposed surface area of 500 cm2 in a filter with a diameter of only 13 mm and thickness of 60 μm, orienting approximately 2.4 x 10"7 moles of lipid. By 2H NMR spectra on chain deuterated phosphatidylcholine it was established that lipids adsorb as wavy, tubular bilayers to the inner pore surface. By !H magic angle spinning NMR it is found that the sample preparation procedure resulted in formation of a single lipid bilayer inside every pore. Using a novel pulsed field gradient NMR technique, an average length of lipid tubules of 0.4μm was measured. Tubule length may vary from a fraction of a micrometer to several micrometers depending on membrane composition and preparation procedures. The membranes are separated from the solid surface by a thick water layer with an average thickness of 30 A such that lipids are completely unperturbed by the solid support. This makes the membranes suitable for incorporation of biological components, such as sensitive integral and peripheral membrane proteins, e.g. G- Protein Coupled Membrane Receptors (GPCR), receptor tyrosine kinases, ion channels, etc.
Adhesion between membrane and solid support is, nevertheless, quite strong. Membranes are not removed from the surface by passing water through the pores at high rate. Furthermore, the invention evidences that the supporting water layer is well sealed by the membrane from the bulk water phase. This permits entrapment of solutes, like soluble components of signal transduction pathways between the solid support and the bilayers. It is demonstrated that water soluble polymers with a molecular weight of 8,000 can be trapped in this water layer. Also, the substrate used for sample preparation, anodized aluminum oxide, is compatible with magnetic resonance spectroscopy, optical spectroscopy, radiotracer studies, neutron- and x-ray experiments, calorimetry, etc., permitting easy detection of receptor binding events as well as structural studies. The large accessible membrane surface raises sensitivity of assays based on our technology by up to three orders of magnitude per unit of chip area. An additional advantage is that substrates are applied by slowly flowing solutions through pores. The flow rate of liquid through the pores is easily controlled, permitting substrate binding studies to be conducted under very reproducible conditions. Because of the unique features of those solid-supported membranes, including lack of perturbation from the substrate, large surface area, strong adhesion to the support, stability to water flow, and the ease of sample preparation, the invention provides considerable promise for preparation of biosensors.
The feasibility to deposit membranes containing GPCR was demonstrated using the GPCR bovine rhodopsin. Phosphatidylcholine (PC) membranes containing rhodopsin at a protein-to-lipid ratio of 100:1 adsorbed to the inner surface of AAO pores in a similar fashion as pure PC membranes as demonstrated by 2H NMR on the lipid. One AAO filter with a diameter of 13 mm oriented about 5 x 10"9 moles of rhodopsin, corresponding to 3 x 1015 copies of the GPCR. By 2H NMR measurements it was established that the detergent octylglucoside that was used for rhodopsin reconstitution is washed away within minutes by flowing water through the pores. This eliminates detergent perturbation of GPCR function and grossly simplifies reconstitution procedures.
The invention concerns a composition comprising a single bilayer lipid membrane supported on a solid support, wherein the membrane comprises a lipid tubule having a wavy tubular geometry and rims, wherein the tubule has rims that are in contact with the solid support, and a center region between the rims that is spaced apart from the support by an aqueous cushion, the aqueous cushion being located between the membrane and the support.
The invention further concerns the embodiment of such a composition wherein the solid support is a porous aluminum oxide support.
The invention further concerns the embodiment of such compositions wherein the lipid comprises a phospholipid.
The invention further concerns the embodiment of such compositions wherein the membrane comprises an incorporated biological molecule.
The invention further concerns the embodiment of such compositions wherein the incorporated biological molecule is an integral membrane protein or a peripheral membrane protein. The invention further concerns the embodiment of such compositions wherein the integral membrane protein or the peripheral membrane protein is a receptor molecule, an enzyme, or an antigen.
The invention further concerns the embodiment of such compositions wherein the incorporated biological molecule is a receptor molecule capable of binding an agonist or an antagonist of a signal transduction pathway.
The invention further concerns the embodiment of such compositions wherein the receptor molecule is a G-protein coupled membrane receptor. The invention further concerns the embodiment of such compositions wherein the solid support is porous, and wherein an agonist or antagonist of the receptor, a G- protein, or an analog of such molecules is provided to the composition by flowing a solution through the pores of the porous support so as to permit the receptor agonist or antagonist to bind to the receptor from inside the lipid tubule from the side of the lipid monolayer opposite from the AAO surface. The invention further concerns the embodiment of such compositions wherein the agonist or antagonist of the receptor, the G-protein, or the analog of such molecules receptor agonist or antagonist is contained in the aqueous cushion. The invention further concerns a biosensor device comprising a solid support, a biological component, and a physical transducer; wherein: the biological component comprises a single bilayer lipid membrane supported on the solid support, wherein the membrane comprises a lipid tubule having a wavy tubular geometry and rims, wherein the tubule has rims that are in contact with the solid support, and a center region between the rims that is spaced apart from the support by an aqueous cushion, the aqueous cushion being located between the membrane and the support; the membrane comprises an incorporated biological molecule selected from the group consisting of an enzyme, a receptor molecule, and an antigen; and the physical transducer serves to generate a signal in response to the binding of a target molecule to the incorporated biological molecule.
The invention further concerns the embodiment of such a biosensor device wherein the solid support is a porous aluminum oxide support.
The invention further concerns the embodiment of such biosensor devices wherein the lipid comprises a phospholipid.
The invention further concerns the embodiment of such biosensor devices wherein the incorporated biological molecule is a receptor molecule. The invention further concerns the embodiment of such biosensor devices wherein the receptor molecule is a G-protein coupled membrane receptor. The invention further concerns the embodiment of such biosensor devices wherein the solid support is porous, and wherein an agonist or antagonist of the receptor, a G-protein, or an analog of such molecules is provided to the composition by flowing a solution through the pores of the porous support so as to permit the receptor agonist or antagonist to bind to the receptor from inside the lipid tubule from the side of the lipid monolayer opposite from the AAO surface.
The invention further concerns the embodiment of such biosensor devices wherein the aqueous cushion contains a biological molecule that interacts with the receptor molecule. The invention further concerns the embodiment of such biosensor devices wherein the aqueous cushion contains a G-Protein, and the incorporated biological molecule is a G-protein coupled membrane receptor.
The invention further concerns the embodiment of such biosensor devices wherein the physical transducer comprises an optical sensor, an electrochemical sensor, a potentiometric sensor, a conductometric sensor, or a piezoelectrical sensor; wherein the sensor generates a detectable physical signal in response to the binding of a target biological molecule to the biosensor's incorporated biological molecule.
The invention further concerns the embodiment of such biosensor devices wherein the physical transducer is an optical sensor selected from the group consisting of a colorimetric optical sensor operating in the visible or non-visible spectral range. The invention further concerns the embodiment of such biosensor devices wherein the optical sensor is an optical sensor that detects fluorescent light.
The invention further concerns a method of detecting a pharmacological agent that binds to a biological molecule, wherein the biological molecule is selected from the group consisting of an enzyme, a receptor molecule, an antigen, and an antibody, wherein the method comprises the steps: (A) incubating the pharmacological agent in the presence of a biosensor device, the biosensor device comprising a solid support, a biological component, and a physical transducer; wherein the biological component comprises a single bilayer lipid membrane supported on the solid support, wherein: the membrane comprises a lipid tubule having a wavy tubular geometry and rims, wherein the tubule has rims that are in contact with the solid support, and a center region between the rims that is spaced apart from the support by an aqueous cushion, the aqueous cushion being located between the membrane and the support; the membrane comprises the biological molecule incorporated therein; and the physical transducer serves to generate a signal in response to the binding of a target molecule to the incorporated biological molecule; and (B) determining whether the physical transducer generates a signal, the generated signal indicating that the pharmacological agent binds to the biological molecule. The invention further concerns the embodiment of such method wherein the biological agent is a G-protein coupled membrane receptor, and the method detects a pharmacological agent that binds to a G-protein coupled membrane receptor.
Description of the Preferred Embodiments: The present invention relates to compositions and methods for forming membranes that may be deposited into solid supports e.g., for use in biosensors. In a preferred embodiment, the invention relates to compositions and methods for forming a high surface area, supported single-lipid bilayer matrix in which the membrane is separated from the support by a closed and stable aqueous cushion. Such compositions comprise a lipid bilayer, a solid support, and where used for a biosensor, a biological molecule (such as a GPCR).
Any of a variety of lipids may be employed in accordance with the methods of the present invention. Preferably, such lipids will be glycerol based lipids (e.g., phosphocholine, phosphatidyl-DL-glycerol, phosphatidylethanolamine, phosphatidylinositol, phosphatidylserine, etc.) and their derivatives and salts. Preferably, such lipids will have fatty acid side chains of 10-20 carbon atoms, more preferably of 12-16 carbon atoms. In preferred embodiments, the membrane bilayer compositions of the biosensor devices of the present invention will comprise l-palmitoyl-2-oleoyl-sn-glycero-3 phosphocholine (POPC). Any of a variety of solid supports may be employed in the biosensor devices of the present invention, however, the most preferred solid support is a porous aluminum oxide support. In particular, the porous aluminum oxide Anopore™ support (Structure Probe, Inc., West Chester, PA, USA) is preferred. Although any of a variety of solid supports may be employed in accordance with the principles of the present invention, the most preferred solid support is a porous aluminum oxide support. In particular, the porous aluminum oxide Anopore™ support (Whatman, Inc.) is preferred. The Anopore™ support is fabricated from a unique form of aluminum oxide with a highly controlled, uniform capillary pore structure that is tightly controlled at 0.2 μm. The starting purity of the aluminum metal use in the first step for the manufactured of the membranes is quite high. The support provides the advantages of high flow rates, efficient particle retention, rigid, uniform surface, transparency (when wet), ability to retain virtually no background stain, low levels of extractable materials, promotes sieving of particles at the surface, is temperature resilient (stable to 400° C) and electron beam radiation resistant.
The proteins of the biological component of the biosensor can be prepared either by reconstituting a lipid bilayer using, for example, purified membrane protein(s), or by forming bilayers from the cellular membranes of spheroplasts. In a preferred embodiment, the present invention provides a facile process for reconstituting single lipid bilayers that contain functional membrane proteins into Anopore™ filters. In a first preferred procedure, a desired membrane protein is preferably recombinantly expressed as a fusion protein containing a C-terminal "tail" (e.g., a Hisio tail) (Grisshammer, R. et al. (1997) "QUANTITATIVE
EVALUATION OF NEUROTENSIN RECEPTOR PURIFICATION BY IMMOBILIZED METAL AFFINITY CHROMATOGRAPHY," Protein Expression Purification 11:53-60; Tucker, J. et al. (1996) "PURIFICATION OF A RAT NEUROTENSIN RECEPTOR EXPRESSED IN ESCHERICHIA COLI," Biochem. J. 317:891-899. The presence of the C-terminal tail permits the membrane protein to be recovered using immobilized metal affinity chromatography (IMAC) with a Ni2+ - nickel nitrilotriacetic acid (NTA) column, preferably as described by Tucker, J. et al. (1996) "PURIFICATION OF A RAT NEUROTENSIN RECEPTOR EXPRESSED IN ESCHERICHIA COLI " Biochem. J. 317:891- 899. For producing membranes via reconstitution, membrane proteins and lipids are solubilized in detergent micelles preferably using either a mixture of 3-[(3- cholamidopropyl)dimethylammonio]-l-propanesulfonate (Chaps), cholesteryl hemisuccinate (CHS), and dodecyl-,B-D-maltoside (LM )] (see, Tucker, J. et al. (1996) "PURIFICATION OF A RAT NEUROTENSIN RECEPTOR EXPRESSED IN ESCHERICHIA COLI," Biochem. J. 317:891-899; Grisshammer R. et al. (2002) "CHARACTERIZATION OF AN ANTIBODY Fv FRAGMENT THAT BINDS TO THE HUMAN, BUT NOT TO THE RAT NEUROTENSIN RECEPTOR NTS-1 ," Protein Expression Purif. 24:505-512; Luca, S. et al. (2003) "THE CONFORMATION OF NEUROTENSIN BOUND TO ITS G PROTEIN-COUPLED RECEPTOR," Proc. Natl. Acad., Sci. (U.S.A.) 10:10706-10711), or, more preferably, for incorporation within AAO supports, using octylglycoside (in the range from 40 to 80 mM).
Liposomes are formed from the mixed detergent-lipid-protein components by lowering the detergent concentration below the critical micelle concentration (cmc) (see, U.S. Patent No. 6,143,321; Urbaneja, M.A. et al. (1990) "DETERGENT SOLUBILIZATION OF PHOSPHOLIPID VESICLE. EFFECT OF ELECTRIC CHARGE," Biochem J. 270(2):305-308; Marsh, D. et al. (1986) "PREDICTION OF THE CRITICAL MICELLE CONCENTRATIONS OF MONO- AND DI-ACYL PHOSPHOLIPIDS," Chem Phys Lipids 42(4):271-277; Lasch, J. et al. (1983) "A METHOD TO MEASURE CRITICAL DETERGENT PARAMETERS. PREPARATION OF LIPOSOMES," Anal Biochem. 133(2):486-491; Allen, T.M. et al. (1980) "DETERGENT REMOVAL DURING MEMBRANE RECONSTITUTION," Biochim Biophys Acta 601(2):328-342), extruding the liposomes through stacked Anopore™ membranes, and removing the detergent and multilayer components by controlled flushing with buffer.
When employing the detergent octylglucoside, liposomes with lipid bilayers are formed spontaneously when the added buffer causes the detergent concentration to fall below the critical micelle concentration (about 20 mM) ( see e.g. Mitchell, D.C. et al. (2001) "OPTIMIZATION OF RECEPTOR-G PROTEIN COUPLING BY BILAYER LIPID COMPOSITION I - KINETICS OF RHODOPSIN- TRANSDUCIN BINDING," J. Biol. Chem. 276(46):42801-42806; Litman, B. J. (1982) "PURIFICATION OF RHODOPSIN BY CONCANAVALIN A AFFINITY
CHROMATOGRAPHY," Methods Enzymol. 81:150-153; Jackson, M.L. et al. (1985) "RHODOPSIN-EGG PHOSPHATIDYLCHOLINE RECONSTITUTION BY AN OCTYL GLUCOSIDE DILUTION PROCEDURE," Biochim. Biophys Acta 812:369-376; Miller, J.L. et al (1987) "BINDING AND ACTIVATION OF ROD OUTER SEGMENT PHOSPHODIESTERASE AND GUANOSINE TRIPHOSPHATE BINDING PROTEIN BY DISC MEMBRANES: INFLUENCE OF REASSOCIATION METHOD AND DIVALENT CATIONS," Biochim. Biophys Acta 898:81-89).
When such procedures are conducted, the liposomes formed after dilution form bilayers that cover the cylindrical surface on pores in the Anopore® support. The residual detergent is easily flushed out the lipid bilayers by passing detergent- free buffer through the support. Prior to the present invention, it might take as long as 12-24 hours of dialysis detergent from a lipid bilayer preparation; The present invention greatly shortens and simplifies the procedure for detergent removal. To form membranes from spheroplasts, bilayers can be prepared by extruding spheroplasts through stacked membranes (especially stacked Anopore™ porous aluminum oxide membranes). Preferably, the spheroplasts will be bacterial spheroplasts that have been genetically engineered to express membrane proteins. Spheroplasts from genetically engineered E. coli are particularly preferred.
Multilayer components may be removed by controlled flushing with buffer. Most preferably, the supported membranes of the present invention will be at least partially spaced apart from the support by a water (or aqueous buffer) layer or cushion. Water-soluble biopolymers may be advantageously entrapped in this water layer or cushion.
The bilayer membranes of the present invention have the advantages that the incorporated protein is functional, and the effective membrane area is up to three orders of magnitude larger than the surface area of a flat chip. The formed membranes are stable at moderate flow rates through pores; the flow rate can be controlled by, for example, an infusion pump. The bilayer membranes produced in accordance with the methods of the present invention are wavy tubules as shown in Figures 10A and 10B. The upper and lower ends of tubules are in contact with the solid support, and seal off a layer of trapped water. Membrane protein that is incorporated into such bilayers can be in an orientation that "faces" the support, or in an orientation that "faces" away from the support. Where desired, buffer, charge, pH, and membrane curvature parameters may be altered in order to cause more membrane proteins to adopt a particular orientation relative to the support.
As indicated above, the supported membranes of the present invention can be employed in biosensor devices. As used herein, the term "biosensor device" is intended to denote an analytical device that is capable of generating a signal in response to the presence of, and preferably, the binding of, a target molecule. The biosensor devices of the present invention possess a biological component, which comprises an immobilized, or substantially immobilized, single bilayer membrane, and one or more membrane proteins incorporated therein; and a physical transducer, which serves to initiate the generated signal of the device. As desired, the biosensor devices of the present invention may be augmented to additionally contain signal enhancers and signal multipliers. Likewise, co-factors, accessory proteins, and other relevant biomolecules may be incorporated into the biosensor.
The biological component of the biosensors of the present invention will typically comprise one, two, three, four or more membrane-bound proteins. A single species of such protein may be incorporated into the membranes of the present invention, or, in alternative embodiments, two, three, four or more different species of receptor proteins. Such additional species may have related functions, such as, for example, all being cytokine receptors, etc., or may have diverse functions, or may have subsets of receptors with related functions. Different proteins (or sets of proteins) can be incorporated into membrane located at different regions of the support, so as to produce a solid support having a pattern of pores each containing different types of sensor protein.
Any of a wide variety of membrane associated proteins can be incorporated in the membranes of the present invention. Such biological components may be enzymes, or other proteins, whose expression is characteristic of disease (e.g., bone specific alkaline phosphatase, aldose reductase, myoglobin, troponin I, etc.). More preferably, however, they will be receptor molecules. Suitable receptor molecules include cell-surface receptors (including protein-tyrosine kinase receptors (e.g., EGFR, PDGFR, MCSFR, SCFR, insulin-R, VEGFR, Trk, Met, Ron, Axl, Eph), ion channel receptors; or receptors for TNF and related factors (e.g., Trk, Met, Ron, Axl, Eph, Fas, TNFRI, TNFRII, CD40, CD30, CD27, 4-1BB, LNGFR, OX40), serine-threonine kinase receptors (e.g., TGFβR), transmembrane 7 or G protein-coupled receptors (e.g., receptors for CCR1, CCR2α, β, CCR3, CCR4, CCR5, CXCR1, CXCR2, CXCR3, CXCR4, BLR1, BLR2, V28, and class I and class II cytokines), CD4+ receptors, class I (hematopoietic cytokine) receptors (e.g., IL-lβ, IL-2R β and γ chains, IL-3Rα, IL-5Rα, GMCSFRα, the IL-3/IL- 5/GM-CSF receptor common β-chain, IL-4Rα, IL-7Rα, IL-9Rα, IL-10R, IL-l lRα, IL-13Rα, LIFR β, TPOR, OBR, IL-6Rα, gpl30, OSMRβ , GCSFR, IL-1 lRα, IL- 12RM and IL-12Rb2, GHR, PRL, and EPO), EGFR, PDGFR, MCSFR, SCFR, insulin-R, VEGFR, and class II receptors (e.g., IFNgRα, IFNgRβ, IL-10R, tissue factor receptor (TFR), and IFNαRl), etc.), opiate receptors, cannabinoid receptors, ***e receptors; or hormone receptors (such as the receptors for adrenaline (epinephrine), adrenocorticotropic hormone (ACTH), androgens (e.g., testosterone), angiotensinogen, antidiuretic hormone (ADH) (vasopressin), atrial- natriuretic peptide (ANP), calciferol (vitamin D3), calcitonin, calcitriol, cholecystokinin, chorionic gonadotropin (CG), dopamine, erythropoietin, estrogens (e.g., estradiol), follicle-stimulating hormone (FSH), gastrin, glucagon, glucocorticoids (e.g., cortisol and urinary cortisol), gonadotropin-releasing hormone (GnRH), gorticotropin-releasing hormone (CRH), growth hormone (GH), growth hormone-releasing hormone (GHRH), insulin, insulin-like growth factor- 1 (IGF-1), leptin, luteinizing hormone (LH), melatonin, mineralocorticoids (e.g., aldosterone), neuropeptide Y, noradrenaline (norepinephrine), oxytocin, parathyroid hormone (PTH), progesterone, prolactin (PRL), renin, secretin, somatostatin, theophylline, thiiodothyronine T3, thrombopoietin, thyroid- stimulating hormone (TSH) , thyrotropin-releasing hormone (TRH), thyroxine (T4); or cytokine receptors (such as the receptors for the interleukins (e.g., IL-2, IL-3, IL-4, IL-5, IL-6, IL-7, IL-8, IL-10, IL-12, IL-13) or TNFα, VEGF, GMCSF, IL-lβ, FGFβ, LNFγ, EGF, PDGF, MCSF, SCF, insulin, VEGF, Trk, Met, Ron, Axl, Eph, Fas, CD40, CD30, CD27, 4-lBB, LNGFR, OX40, TGFβR, or a ligand of CCR1, CCR2α, β, CCR3, CCR4, CCR5, CXCR1, CXCR2, CXCR3, CXCR4, BLR1, BLR2, V28 receptor , or a receptor of IL-2, IL-3, IL-4, IL-5, IL-6, IL-7, IL- 8, IL-10, IL-12, or IL-13; or antigen recptors or binding ligands (such as those that bind antigens characteristic of Chlamydia, Streptococcus pyogenes Group A bacteria, H. pylori, or M. tuberculosi, hepatitis virus, rubella, CMV or immunodeficiency virus (HIV, FIV), prostate specific antigen, etc.); or membrane- associated antibodies to such antigens, or autoimmune immunoglobulins, thyroglobulin, anti-thyroglobulin, IgE, IgG, or IgM immunoglobulins, tumor markers (e.g., prostate specific antigen, AFP CEA, etc.). It is, however particularly preferred that such biological molecule be a
GPCR, so as to form a biosensor capable of detecting ligands that affect GPCR activity. In designing such a biosensor, it may be desired to include G protein in the biosensor. In one embodiment, the G protein peptides simulating elements of G-protein function will be included in the reconstituted membrane preparations so as to be present in the aqueous cushion separating the bilayer from the support. In such an embodiment, the external side of the bilayer will exhibit the attributes of the external side of a biological membrane. In an alternate embodiment, the G protein will not be included in the reconstituted membrane preparation, but will be provided in the buffer being flushed through the support. In such an embodiment, the external side of the bilayer will exhibit the attributes of the internal side of a biological membrane.
The biological component(s) can be recombinantly produced (i.e., produced in a heterologous host cell), genetically produced (i.e., produced in a homologous host cell), or synthetically produced (i.e., synthesized using in vitro chemical synthetic procedures). Such biological component(s) can be incorporated into the bilayer membranes of the present invention by any of a variety of means, such as by reconstitution or by extrusion of spheroplasts membranes, as discussed above.
The physical transducer component of the biosensors of the present invention is preferably configured to exploit the open pore nature of the preferred solid support (i.e., Anopore®), so as to permit detection of signal by flowing a fluid through the pores of the support. For such purposes, the physical transducer component of the biosensors of the present invention preferably comprises an optical sensor, such as a colorimetric (i.e., a sensor that modulates the production of a visible spectrum optical signal in response to target molecule binding), a non- visible spectrum optical sensor (i.e., a sensor that modulates the production of a non-visible spectrum optical (e.g., infra-red, UV, fluorescent, etc.) signal in response to target molecule binding). The use of the translucent Anopore™ support facilitates the use of colorimetric and non-visible spectrum optical sensors. An optical sensor that detects fluorescent light is particularly preferred. Receptor activation may be sensed by retention/release of radioactive substances such as tritium labeled receptor agonists or antagonists.
Notwithstanding the above, the physical transducer component of the biosensor of the present invention may be configured as an electrochemical sensor (e.g., an amperometric sensor (i.e., a sensor that modulates a current in response to target molecule binding), a potentiometric sensor (i.e., a sensor that modulates a pH change in response to target molecule binding), a conductometric sensor (i.e., a sensor that modulates resistance in response to target molecule binding), or a piezoelectrical sensor (i.e., a sensor that modulates a piezoelectric response to target molecule binding). Detection of an electric signal can be enhanced by coating the pores of the solid support with a conductive layer. For purposes of employing such physical transducer components, it is desirable to employ a support that has a closed end capable of serving to impede or block the flow of electrons or electrolytes (see, Marchal, D. et al. (1998) "ELECTROCHEMICAL MEASUREMENT OF LATERAL DIFFUSION COEFFICIENTS OF UBIQUINONES AND PLASTOQUINONES OF VARIOUS ISOPRENOID CHAIN LENGTHS INCORPORATED IN MODEL BILAYERS," Biophys.J. 74:1937-1948).
The signal generated by the physical transducer may be quantitative (i.e., varying in intensity, duration, etc., with the concentration of the target molecule in a sample), qualitative (i.e., producing a signal in response to a threshold concentration of the target molecule), or partially qualitative and partially quantitative. The biosensor devices of the present invention may be employed in ligand binding studies to detect ligand concentrations in a sample.
Η-, 2H, 13C, and 31P NMR with and without "magic angle" spinning (MAS) can be used to verify that the facile sample preparation method of the present invention results in the attachment of single lipid bilayers to the solid support. MAS NMR spectra are simpler, better resolved, and have larger signal amplitudes, as only the isotropic chemical shift and / couplings contribute to the resonance frequency. Chemical shift anisotropy and dipole-dipole couplings are averaged by fast tumbling of the molecules in solution. The physical reason for this averaging is that the orientation of the molecule with respect to the static field changes on a timescale that is faster than the inverse anisotropy of the interactions. Such an irregular fast tumbling of the sample inside the spectrometer cannot be realized technically. However, as was shown by Lowe, I.J. (1959) ("FREE
INDUCTION DECAYS OF ROTATING SOLIDS," Phys. Rev. Lett. 2:285) and Andrew, E.R. et al (1959) ("REMOVAL OF DIPOLAR BROADENING OF NUCLEAR MAGNETIC
RESONANCE SPECTRA OF SOLIDS BY SPECIMEN ROTATION," Nature 183:1802), it is sufficient to rotate the sample about one axis, if this axis is inclined by the "magic angle" of 54.74° with respect to the static magnetic field. At this magic angle,
B0 = a cos vl/3 , the second order Legendre polynomial, P2 = (3 cos2 θ - 1) / 2 , vanishes. The technique is called "magic angle spinning" (MAS) (see, STRUCTURE DETERMINATION II (NMR, EPR), G. Jeschke (WS 2003/04 9) "High-Resolution Solid-State NMR: Magic Angle Sample Spinning and Cross Polarization," web site: mpip-mainz.mpg.de/~jeschke/lect9.pdf).
As opposed to previous preparation procedures, the preferred solid- supported membranes of the present invention are separated from the solid surface by a thick water cushion over most of their area. Therefore it can be expected that proteins in reconstituted membranes, prepared according to our procedure, will function without any interference from the solid support. Furthermore, the present sample preparation procedure will raise sensitivity of assays by up to three orders of magnitude per unit of biochip area. It is therefore envisioned that the invention has utility in the production of biosensors for pharmaceutical applications, environmental screening, detection of warfare agents, etc. The solid supported membranes can be used for rapid screening of binding affinity of drugs to membrane receptors, e.g. G-protein coupled membrane receptors (GPCR). The solid substrate is compatible with optical studies, radiotracer binding studies, NMR, EPR, small angle neutron scattering, x-ray diffraction, calorimetry, and other methods, permitting the development of binding assays as well as structural studies on proteins and bound ligands. Having now generally described the invention, the same will be more readily understood through reference to the following Examples, which are provided by way of illustration, and not intended to be limiting of the present invention. Example 1
Materials. l-Palmitoyl-2-oleoyl-sn-glycero-3 phosphocholine (POPC), 1- palmitoyl(d3ι)-2-oleoyl-sn-glycero-3 phosphocholine (POPC-d3ι), l-palmitoyl-2- oleoyl-sn-glycero-3 phosphocholine-d (POPC-d4), and 1,2-dimyristoyl-sn- glycero-3 -phosphocholine (DMPC) were purchased from Avanti Polar Lipids (Alabaster, AL). D2O (99.8% D) and deuterium-depleted H2O (2-3 ppm D) were purchased from Cambridge Isotope Labs (Andover, MA). Anopore aluminum oxide filters (13 mm diameter) with nominal pore sizes of 0.02, 0.1, and 0.02 μm were purchased from SPI Supplies (West Chester, PA). Praseodymium (III) nitrate hexahydrate (99.9%) was purchased from Aldrich (Milwaukee, WI). Polyethylene glycol 8000 (PEG8000) with an average molecular weight of 8455 g/mol was purchased from Spectrum. Piperazine-N,N'-bis(ethanesulfonic acid) sesquisodium salt (Calbiochem; San Diego, CA) was used to prepare 10 mM PIPES buffer with 100 mM NaCl (Sigma; St. Louis, MO) in deuterium-depleted water for use in 2H NMR measurements on POPC-d4.
Sample Preparation. Lipid Loading. The lipids were loaded using one of two different extruders. The majority of the experiments utilized a mini-extruder (Avanti; Alabaster, AL) equipped with 1-mL syringes. One to five 13 -mm diameter anopore filters were placed into the extruder and flushed with 1 mL of water 11 times before bringing them into contact with the lipid. The water was removed and 5 mg lipid dispersed in 1 mL of water, buffer, or aqueous PEG solution was passed through the filters 15 times. The lipid dispersion was removed and one or two 1 mL aliquots of water, buffer, or aqueous PEG solution was passed through the filters at an rates of 3 - 12 mL/min, either by hand or using an infusion pump, to remove any remaining liposomes and to remove additional bilayers that are loosely attached to the first bilayer at the AAO surface. In the PEG entrapment experiments, the filters were then flushed with 1 mL of an isotonic solution (100 mOsmole) of NaCl in D2O at a rate of 0.04 mL/min using an infusion pump to remove any PEG not trapped beneath the bilayers. In the case of POPC, the extrusion was performed at room temperature, but for DMPC, the lipid and extruder were maintained at 35 °C during loading using a dry heat incubator (Fisher Scientific; Pittsburgh, PA).
*H and 31P MAS NMR Samples. A filter was removed from the extruder and a drop of water was added to its surface to keep the lipids fully hydrated. In the case of the shift-reagent studies, a drop of 5 mM Pr(NO3)3 was added and allowed to sit for ten minutes. In later experiments, which gave identical spectra, an infusion pump was used to push 1 mL of the shift reagent through the filter at a rate of 0.02 ml/min. The filter was then cut into small pieces using a blade. The wet filter pieces were transferred into an 11 -μL spherical insert made of Kel-F that was placed into a 4-mm NMR rotor (Bruker Biospin Corp. Billerica, MA). 2H NMR Samples. The filters were removed from the extruder and placed between glass slides (12 x 9 mm) with a drop of deuterium -depleted water to keep the lipids fully hydrated. The protruding edges of the filters were removed with a blade. The entire stack was wrapped in Teflon tape and placed in a flat NMR tube along with deuterium-depleted water. The tube was sealed with a silicone stopper. NMR Experiments 1H- and 31P MAS experiments are performed on a Bruker (Billerica, MA) DMX300 spectrometer equipped with a 4-mm broadband Bruker MAS probe. Experiments are conducted at a rotor spinning frequency of 5 kHz and a temperature of 30.0 ± 0.1 °C, unless otherwise specified. !H spectra are acquired at a resonance frequency of 300.14 MHz with a 3.6 μs 90°pulse and a 4 s delay between scans. The spectral width is 5 kHz and the number of acquisitions is 32. P spectra are acquired at a resonance frequency of 121.4 MHz with a 2.5 μs 90°pulse and a repetition rate of one acquisition per second. The spectral width is 50 kHz, and the number of acquisitions varied from 25,000 to 90,000. PFG-MAS diffusion, 1H MAS T2, and 2H NMR experiments are performed on a Bruker DMX500 spectrometer running XWINNMR v3.1. The 2H NMR spectra are recorded using a Doty flat coil probe with variable angle housing (Doty Scientific; Columbia, SC) whose coil could be rotated relative to the external magnetic field. Spectra are acquired at an operating frequency of 76.8 MHz using a quadrupolar echo sequence (Davis, J.H. et al. (1976) Chem.Phys.Lett. 42:390-394) with a 3.4 μs 90°pulse, a 75 μs interpulse delay, and a 250 ms delay between scans. Spectra are acquired at a temperature of 24.2 ± 0.1 °C with the filter pore axis both parallel (βo=0°) and perpendicular (β0=90°) to the external magnetic field, and the number of acquisitions varied from approximately 200,000 to 800,000. For data analysis, the data are transferred to a personal computer and processed as described by Holte, L.L. et al. ((1995) "2H NUCLEAR MAGNETIC RESONANCE ORDER PARAMETER PROFILES SUGGEST A CHANGE OF MOLECULAR SHAPE FOR PHOSPHATIDYLCHOLINES CONTAINING A POLYUNSATURATED ACYL CHAIN," Biophys. J. 68:2396-2403) and Huster, D. et al. ((1998) "INFLUENCE OF
DOCOSAHEXAENOIC ACID AND CHOLESTEROL ON LATERAL LlPID ORGANIZATION
IN PHOSPHOLIPID MIXTURES," Biochemistry 37:17299-17308) using software written for Mathcad version 200 li (MathSoft Engineering & Education, Inc., Cambridge, MA). The diffusion experiments and spin-spin relaxation measurements are performed at 27.6 ± 0.1°C at a proton frequency of 500.17 MHz with a Bruker 4- mm PFG-MAS probe with z-axis gradients operated at a spinning frequency of 5 kHz. Spectra were obtained at a sweep width of 5,000 Hz. The relaxation measurements are performed by the Carr-Purcell-Meiboom-Gill pulse sequence (Meiboom, S. et al. (1958) Rev.Sci.Instrum. 29:6881; Can, H.Y. et al. (1954)
Phys.Rev. 94:630-638). Diffusion measurements are conducted at sixteen different values of gradient strength varying from 0.01 - 0.37 T/m with a stimulated echo sequence using sine-shaped bipolar gradient pulses (Cotts, R.M. et al. (1989) J. Magn.Reson. 83:252-266) of 5 ms duration. A longitudinal eddy current delay of 5 ms is used. Diffusion times are varied from 20 - 200 ms. At every gradient strength, 64 scans are acquired with a recycle delay of 4 s. Mathcad is used to fit the diffusion data to the equation that relates signal intensity to the diffusion constant for powder samples with filter pores oriented at random to the orientation of the magnetic field gradient (Gaede, H.C. et al. (2003) "LATERAL DIFFUSION RATES OF LIPID, WATER, AND A HYDROPHOBIC DRUG IN A MULTILAMELLAR LIPOSOME," Biophys J 85:1734-1740),
Figure imgf000034_0001
where D is the diffusion constant, and k is a factor whose exact nature depends on the pulse sequence and on instrumental settings. For the stimulated echo sequence, 9 9 91 T O l k = 4γ g δ Δ , where γ is the gyromagnetic ratio of protons, g is the V 2 8) gradient strength, δ is the gradient pulse length, and T is the time between the gradient pulses sandwiching the 180° pulses (Fordham, E.J. et al (1996) J. Magn.
Reson. Ser.A 121:187-192.
2H NMR Lineshape Simulation. The 2H NMR spectra of deuterated lipids were simulated using Mathcad on a personal computer. Since 2H spectra with quadrupole splittings are symmetric about the origin, the free induction decay (FID) for a lipid containing / deuterated carbons may be represented as:
y(t)rea, = ∑I, ∞s{ω,t)e)φ(- t/T2) (2)
where Ii is an intensity factor representing the number of deuterons attached to the tth carbon atom andω; = 2πVj . The time is defined as t = n • DW , where n is the number of the data point in the digitized FID and DW is the dwell time between points. The frequency of each resonance of the deuterium doublet for the tth carbon is given by (Abragam, A "FINE STRUCTURE OF RESONANCE LINES - QUADRUPOLE EFFECTS," In: Principles of Nuclear Magnetism; Oxford University Press: 1961; pp 216-263): v(l,2)i = v(iso)i ± Δvq • S(θ) • S(mol)i (3)
where the isotropic frequency, v(iso)i , is 0 for spectra acquired on resonance, the constant Δvq=(3/4)e2qQ/h=125 kHz (Burnett, L.J. et al (1971) J Chem.Phys. 55:5829-5831) characterizes the strength of quadrupolar interactions in saturated hydrocarbon chains, S(θ) = — (3 cos2 θ - 1) is an order parameter reflecting the orientation of lipid bilayer normals with respect to the outer magnetic field, and S(mol)i is a molecular order parameter that depends on orientation and fast motions (>10"5 sec) of lipid C-2H bonds with respect to the bilayer normal. The angle θ is formed between the bilayer normal whose unit vector is d = D D and the unit
vector of the external magnetic field b = BQ / B0 (see Figure 1). The orientation of the external magnetic field with respect to the bilayer normals is defined by the angles α and β of the spherical coordinate system shown in Figure 1. The angle θ is related to α and β via the scalar vector product: cos θ = b • d . By substitution of Equation (3) into Equation (2), a formula for the calculation of the FID for a 2H NMR experiment on a lipid bilayer is obtained. Resonances corresponding to all possible values of α and β are superimposed by integration:
yrealn = Σ J Jli cos(2πΔvq • s(α, β) • S(moι)i •n-DW)-exp(-n-DW/T2α)p(β)dαdβ (4) i 0 0
To reproduce the experimental lineshapes, the molecular order parameters S(mol)i were determined by dePakeing of spectra obtained by 2H NMR experiments on randomly oriented bilayers (multilamellar liposomes) (Sternin, E. et al. (1983) J. Magn.Reson. 55:274-282; Mccabe, MA. et al. (1995) J. Magn. Reson. Ser.B 106:80-82). The angular distribution function p(β) and the line broadening, T2α> were adjusted to match experimental signal intensities. For lipid adsorbed onto the AAO pores, p(β) = exp[- (β — β0 ) / 2σ , where the angle β0= 0° for AAO pores oriented parallel to the magnetic field and β0= 90° for pores oriented perpendicular. The value of σ, measured in units of degrees, represents the width of the Gaussian distribution function. For a powder distribution of lipid bilayer normals, as in unoriented multilamellar liposomes, p(β) = sinβ . Fourier transformation of Equation (4) yields the simulated spectrum.
Orientational distribution function of lipid bilayer normals. The order parameters, S(mol)l5 of POPC-d3ι necessary to simulate the 2H NMR spectra of POPC-d3ι adsorbed onto AAO pores (Equation (4)) were obtained from the dePaked spectrum of multilamellar liposomes composed of POPC-d3ι at 24.2 °C (Figure 2). The effective quadrupolar splittings of C-2H bonds in multilamellar liposomes and in single bilayers at AAO surfaces are indistinguishable within the very narrow error margins of ±200 Hz. The 2H NMR spectra of POPC-d31 adsorbed on AAO filters with the filter pore axis aligned parallel (βo=0°) and perpendicular (β0=90°) to the external magnetic field were acquired to determine the orientation and mosaic spread of the adsorbed lipids. The experimental and simulated spectra are shown in Figure 3. Since all bilayer normals in the βo=0° orientation have the same orientation with respect to the magnetic field, namely 0=90°, the starting point for the simulation was narrow resonance lines with quadrupolar splittings related to the order parameters by equation 4. However, the experimental spectra are broadened by mosaic spread in the bilayer orientation and intrinsic linebroadening due to relaxation. To account for the first factor, the simulation included an angular distribution function to describe the spread in bilayer orientation. The calculated spectrum of POPC-d3ι in AAO pores matches the experimental spectrum almost perfectly with a Gaussian angular distribution function p(β) using a width of σ=20°. In the βo=90° orientation, the bilayer normals have a circular distribution relative to the external magnetic field which was the starting point for the simulation. In addition, a mosaic spread of cylinder axis orientation was considered. In fact, exactly the same width of the Gaussian distribution, σ=20° used for βo=0°, results in the best match of experimental and simulated spectra at βo=90c as well. Note that this distribution is quite distinct from what is expected from a random distribution of orientations, p(β)=sin(β), as would be seen in an unoriented sample, and the spectra with such a distribution would appear quite differently. For comparison, both the angular distribution functions used in these simulations and those of a powder sample are shown in Figure 3.
To account for the homogenous linebroadening of the lipid resonances from relaxation, a second adjustable parameter, T2α, was included in the spectral simulations. A T value of 0.5 ms, corresponding to a linebroadening of 600 Hz was used. While this value was a perfect match for the orientation βo=0, spectral resolution in the experimental spectra acquired at βo=90° was lower, and required linebroadening of ~800 Hz. The intense resonance line at the center of the experimental spectra in both βo=0° and βo=90° orientations arises mostly from natural abundance deuterated hydroxyl groups at the AAO surface that are not completely removed by exchange with deuterium-depleted water, as a large hydroxyl signal was observed in AAO samples even in the absence of lipid. However, a small contribution from liposomes is also possible as freezing the AAO/lipid sample reduced the intensity of this central resonance.
The spectrum of POPC-d3ι shown in Figure 3 (Panel C) was recorded without flushing the membranes with water to remove extra layers of lipid. Intensity in the central region of this spectrum acquired at βo=90° is somewhat higher, indicating the presence of more randomly oriented lipid bilayers. The appearance of the spectra was independent of the nominal pore size of the 1 μm- thin active layer that caps the filters, consistent with the fact that pores in the supporting layer composing -98% of the filter length always have a diameter of 0.2 μm. Headgroup conformation and flexibility. To determine whether the lipid headgroups interact strongly with the AAO surface, 2H NMR spectra of POPC-d4 with α and β methylene groups of choline deuterated were acquired. Experiments were conducted in PIPES buffer at pH 7 with 100 mM NaCl. The filter pore axis was aligned parallel (βo=0°) to the external magnetic field. The spectrum is shown in Figure 4, together with the spectrum of POPC-d4 MLVs prepared in the same buffer. Experimental results were compared with simulated spectra. The simulation was conducted using the same width of the Gaussian distribution (σ=20°) and the linebroadening of 600 Hz as for POPC-d3ι at AAO. The spectra in both the MLVs and the single bilayers at AAO are a superposition of two quadrupolar splittings with values of 6.3 and 5.2 kHz, corresponding to the quadrupolar splittings of POPC choline α and β resonances, respectively (Koenig, B.W. et al. (1996) Langmuir 12:1343-1350). The extra signal in the center of the experimental spectrum of lipid adsorbed to AAO appears to be mostly from a residual 2H resonance of AAO hydroxyl groups. However, there was an additional small but systematic deviation in signal intensity between measured and calculated spectra in the frequency range of +5 kHz that could indicate existence of a few percent of lipid with lower, but not well defined headgroup order.
Both samples show two quadrupolar splittings of 6.3 and 5.2 kHz, corresponding to POPC choline α and β resonances, respectively. The extra signal in the center of the experimental spectrum of lipid adsorbed in AAO pores appears to be mostly from a residual 2H resonance of AAO hydroxyl groups. The small deviation in signal intensity between measured and calculated spectra in the frequency range of ±5 kHz could indicate existence of a few percent of lipid with lower headgroup order.
Adsorption of single lipid bilayers. The Η MAS spectra of POPC in crushed AAO filters is shown in Figure 5, Trace B, along with the assigned spectrum of POPC multilamellar vesicles (MLVs) for comparison in Figure 5, Trace A broad water/hydroxyl group resonance partially obscures the lipid glycerol resonances of bilayers in AAO pores, but headgroup β and γ as well as hydrocarbon chain resonances are visible, albeit broadened. Rinsing or boiling of the AAO in D2O reduced the intensity of the large water/hydroxyl signal but did not entirely remove it. Spin-spin relaxation time, T2, measurements were conducted at 27.6 °C of the 1H MAS NMR signals in bilayers adsorbed to AAO to determine if resonances are broadened homogeneously. The T2 value for the γ resonance of POPC adsorbed onto AAO was found to be 8.8 ms, corresponding to a linewidth of 36 Hz which is close to the experimentally measured value of 34 Hz. The introduction of a paramagnetic ion as a chemical shift reagent is often employed to distinguish between the external and internal surface of the membranes of phosphatidylcholine lipid vesicles (Bystrov, V.F. et al. (1971) Chem.Phys. Lipids 6:343-350). This commonly used assay was employed here to determine whether the membranes formed were well-sealed and if they formed single or multiple bilayers. Addition of the shift reagent Pr3+ at a concentration of 5 mM after adsorption of bilayers to AAO pores, either by dropwise addition or slow extrusion at rates of 0.02 mL/minute, shifted a fraction of the γ choline resonance downfield by 0.12 ppm (Bystrov, V.F. et al. (1971) Chem.Phys. Lipids 6:343-350). If Pr3+ was added immediately after extrusion, before flushing the pores with plain water, the shifted signal had low intensity, indicating that only a small fraction of bilayers was exposed to the ions. However, if Pr3+ was added after flushing the pores with water at rates of 3 - 12 mL/minute, the intensity of the shifted signal corresponded to 50% of total intensity as determined by the curve fitting routine in XWINNMR (Bruker Biospin Inc., Billerica, MA), indicating the formation of a single bilayer with one monolayer in contact with the excess water phase and the other monolayer at the AAO surface, shielded from exposure. (Figure 5, Trace C and insert to Figure 5).
31P MAS spectra of POPC MLVs and POPC in crushed AAO filters with and without addition of Pr3+ are shown in Figure 6. POPC in multilamellar liposomes has a single, well-resolved 31P resonance. POPC adsorbed to the surface of AAO pores and exposed to Pr3+ ions has two phosphorous resonances, one with the same chemical shift as in MLVs and the other one shifted downfϊeld from exposure to the shift reagent. Due to the short delay time between scans, the unshifted phosphorous signal is attenuated relative to the shifted signal, whose relaxation time is decreased by complexation with Pr3+. The 20 ppm-broad resonance, 10 ppm upfield is visible even in the absence of lipid and arises from the preparation of the AAO in phosphoric acid electrolyte. It has been reported that these filters contain 7.4 wt% phosphate (Furneaux, R.C. et al (1989) Nature 337:147-149).
Durability of Bilayer Seal. To test the sucsceptibility of the bilayer seal to shear forces, Pr3+ was applied by extrusion through the lipid-loaded pores at a variety of flow rates. Without shear, the appearance of the !H and 31P MAS spectra after addition of Pr3+ ions did not change over a period of 24 hours, suggesting that bilayers are well-sealed. To test the durability of the seal to the main lipid phase transition, the temperature of the filters was lowered after Pr3+ was added. Lowering the temperature resulted in shifting both the resonances of inner and outer monolayers, indicating that the seal was broken.
The seal was also perturbed by higher flow rates of water. When the Pr3+ shift reagent was applied by extrusion of a Pr3+ solution through the AAO pores at rates of 12 mL/minute, then more than 50% of the γ-choline resonances were shifted. Often two or more γ choline peaks were detected, indicating that different regions of the monolayers were exposed to different concentrations of the shift reagent. However, extrusion at rates of 0.02 mL/minute resulted again in 1:1 peak ratios.
Existence of Water Pockets. To determine the average thickness of water trapped beneath the bilayer, 1H MAS spectra at 30 °C were obtained on a sample in which an 11 wt% PEG8000 solution in D2O was trapped beneath the bilayer surface. This was achieved by extruding a dispersion of multilamellar liposomes in a PEG8000 solution. Multiple bilayers were flushed out rapidly with a PEG8000 solution without lipid, followed by slow flushing with an isotonic NaCl solution to remove untrapped PEG. AAO filters loaded with a single bilayer showed the H PEG methylene resonance, which completely superimposed the β headgroup resonance of POPC at 3.57 ppm (Figure 7). The ratio of this superimposed PEG resonance to the γ-choline resonance at 3.15 ppm was found to be 8.6:9 by integration. This resonance was absent in samples extruded with PEG in the absence of lipid.
Fluid-gel phase transitions of lipids. To probe the influence of the AAO support on the phase transition temperature of supported lipids, Η MAS NMR spectra of DMPC supported on AAO pores were acquired as a function of temperature. At a MAS frequency of 5 kHz the !H resonances of lipid hydrocarbon chains are well resolved in the fluid phase but broadened beyond detection in the gel state. To follow the main phase transition of DMPC, the normalized intensity of the 100 Hz line-broadened methylene resonance at 1.3 ppm was plotted versus temperature in Figure 8 for both DMPC in multilamellar liposomes and for single DMPC bilayers adsorbed on AAO pores.
The gel-fluid phase transition in MLV at 23°C, which is seen as a rapid drop of signal intensity to zero, agrees with reported values (Dixon, G.S. et al. (1982) Anal. Biochem. 121:55-61; Boggs, J.M. et al. (1985) Biochim. Biophys. Acta 816:221-233. The phase transition of DMPC single bilayers at AAO surfaces was more complex. The DMPC at AAO shows a rapid drop in signal intensity at 22°C, but the overall transition is broader, with tailing intensity toward lower temperatures. This behavior is consistent with the bilayer area reduction upon the transition from fluid to gel state. Experiments in the presence of Pr3+ ions indicated that the shift reagent started penetrating into the water layer between the AAO surface and the bilayer at temperatures corresponding to the onset of the main phase transition.
Physisorption of lipids to the AAO surface. The mode of lipid attachment to the AAO surface was probed by removing the lipid with methanol. By Η MAS NMR signal intensities of crushed membranes before and after solvent extraction, it was determined that only ~1% of the lipid remained on the AAO surface after extraction, indicating the absence of a covalent bond between adsorbed lipid and AAO. This is supported by the measurement of lipid 31P chemical shifts of the outer monolayer that is exposed to AAO, as the values are identical to the shift of lipids in multilamellar liposomes. It is known that lipid hydrolysis may be triggered at pH-values that deviate from neutral, and that alumina surfaces may be acidic or basic. However, chemical stability of lipid bilayers at AAO surfaces was satisfactory. Most samples did not show any signs of lipid hydrolysis after one week of storage or longer.
Water and POPC lateral diffusion within AAO pore. To determine the mobility of the lipid bilayers and the size of lipid domains, diffusion of lipid and water was measured on crushed POPC/AAO samples containing trapped PEG. Experiments were conducted by Η PFG-MAS NMR at a spinning frequency of 10 kHz. Spectra acquired at 16 different gradient strengths at a temperature of 30.0 °C and a diffusion time of 200 ms are superimposed in Figures 10A and 10B.
The water resonance is a superposition of signals of at least two water pools. At our experimental conditions two thirds of water resides outside pores while one third is inside. The signal decay of the water resonance as a function of gradients strength is complex due to the random orientation of filter pieces as well as water- filled pores in the spinning rotor, water interaction with lipid and AAO surfaces, chemical exchange of protons with AAO hydroxyl groups, and permeation of water through lipid bilayers. From the signal decay at low gradient strength it was estimated that within very generous error limits most of the water moves at the rate of free water, D∞2 TO"9 m2/s .
In contrast to the water resonance that decayed to almost undetectable values, there is a barely measurable decay of lipid resonance intensities of 5% recorded as a function of k. This decay may be fit to equation 1 to obtain an apparent diffusion constant of (4.6 ± 1.3) x 10"13 m2/s for the lipid. The apparent diffusion rate was diffusion time dependent, with higher rates at the shorter diffusion time of 100 ms, indicating that diffusion distances are restricted by the size of lipid patches in the pores. Though the small change in signal intensity causes the numerical value of the diffusion constant to be of questionable reliability, it is the small change in signal intensity itself that reveals that diffusion is restricted. The signal intensity decay of the lipid would be at least ten times greater if the lipid tubules would have infinite length.
Evidence for bilayers inside AAO pores. The distribution function of bilayer normals (Figure 3) obtained from the 2H NMR experiments on deuterated lipids deposited onto aluminum oxide pores proved unambiguously that lipid layers have cylindrical symmetry and that cylinder orientation coincides with the orientation of pores in the filters. In particular, when the filters are oriented with their pores parallel to the magnetic field (β=0°), the lipid bilayer normals in the AAO pores are preferentially oriented perpendicular to the field (0=90°), resulting in a well- resolved 2H NMR spectrum with quadrupolar splittings of half the maximum value (equation 3). Turning the filters by 90° results in a circular distribution of bilayer normals with respect to the magnetic field and a more complex 2H NMR lineshape with significant intensity in the spectral wings (see Figure 3).
The fit of the experimental spectra yields an angular distribution function, p(β), of the pore axis relative to the external magnetic field. A Gaussian distribution with a width of 20+2° resulted in a perfect match between experimental and calculated spectra for both orientations. This value is very reproducible for independently prepared samples. It applies to the fit of quadrupole splittings of chain resonances in POPC-d3ι (Figure 3) as well as to splittings of deuterated headgroups in POPC-d4 (Figure 4).
The question arises if this distribution function reflects a disorder in pore orientations or a disorder that only applies to bilayer normals. Though no numerical values are available for an angular distribution for AAO pores, their parallelism is often touted. Examination of several published scanning electron micrographs indeed show that the pores are smooth and parallel to within -5° (Furneaux, R.C. et al. (1989) Nature 337:147-149; Crawford, G.P. et al. (1992) J. Chem. Phys. 96:7788-7796; Platt, M. et al. (2003) Langmuir 19:8019-8025). Therefore, the width of the angular distribution of 20° must result predominantly from a distribution in bilayer orientations rather than the AAO surface, though it is likely to contain some contributions from the pores, such as deviations from parallelism, roughness along the long axis, and distortion from cylindrical shape. The lipid bilayers must maintain curvature in excess to curvature from the cylindrical symmetry, suggesting that bilayers do not adhere flatly to the inner surface of the pores.
Radius of bilayer curvature. Because of membrane curvature in the submicrometer range it can be expected that individual NMR resonances are broadened homogeneously by lipid reorientation in the magnetic field from lateral diffusion over curved surfaces. Indeed, the 600-Hz linewidth required in the 2H NMR simulations to match experimental 2H NMR spectra at β=0° is much larger than the 350-Hz linewidth used to simulate the spectra of large liposomes with radii of curvature in the μm-range.
If pores are oriented perpendicular to the field (β0=90°), then lateral diffusion of lipids over the circumference of the pore (r=0.1 μm) may also contribute to linebroadening. Indeed, spectral resolution for this orientation is even lower (linewidth -800 Hz). The underlying cause for this broadening, orientational exchange from lateral diffusion over surfaces with small radius of curvature, was confirmed by conducting 2H NMR experiments at higher temperature that raised rates diffusion rates and resulted in further broadening. The broadening of MAS proton resonances of lipids in aluminum oxide pores compared to lipids in large liposomes appears to be the result of curvature- driven orientational exchange as well. Indeed, the T2 value for the proton γ resonance of AAO-adsorbed POPC of 8.8 ms may be compared to 97 ms, the value for the γ-resonance of DMPC MLVs at 30°C (Huster, D. et al. (1999) J. Phys. Chem. B 103:243-251). The reduced !H NMR T2 for POPC on AAO suggests an additional reorientation of the lipids with correlation times in the millisecond range. Using a POPC lateral diffusion constant of 9.5 X 10"12 m2/s (see Fillipov, A. et al. (2002) Biophys. J. 82:754) it is estimated that the radius of curvature for the reorientation must be 100 nm < r < 400 nm, as smaller radii would average out the 2H NMR quadrupolar splittings of headgroups and hydrocarbon chains and larger radii would not influence the homogeneous linewidth of lipid resonances from orientational exchange due to lateral diffusion (Gawrisch, K. et al. (1986) Biochim. Biophys. Acta 856:443-447; Bloom, M. et al. (1987) Biochemistry 26:2101-2105). The data indicated that bilayers adhere to the cylindrical pores with additional curvature, most likely from some waviness of lipid tubules in the pores.
Single lipid bilayers. Pr3+ ions interact with the phosphate groups of phospholipids and shift accessible lipid headgroup resonances to lower field (Bystrov, V.F. et al. (1971) Chem.Phys. Lipids 6:343-350). Extrusion of multilamellar liposomes through AAO pores deposited multilamellar layers of lipid at the AAO surface, as demonstrated by the low intensity of shifted choline γ- resonances after gentle flushing of pores with a solution containing Pr3+ ions (water velocity of 0.01 mm/s in the pores). Multiple bilayers were washed away by flushing the pores with water moving through the pores at a velocity of 1.7 mm/s or higher, leaving only the last bilayer that is nearest the AAO surface. This was confirmed by the area ratio of the shifted and unshifted LH MAS NMR γ-choline resonances of POPC. After very slow flushing with Pr3+ shift reagent the ratio of shifted to unshifted signal was 1 : 1 (Figure 5), as expected for an inner tubular leaflet exposed to the shift reagent and an outer leaflet not exposed to the ions. Flushing of surfaces with water was used by us previously to remove multiple bilayers from the surface of polished Si-single crystals (Koenig, B.W. et al. (1996) Langmuir 12:1343-1350). As in the case of lipids on Si-oxide, the last bilayer adheres to the aluminum oxide surface with greater force. Even at water velocities of 6.7 mm/s in the pores, no detachment was observed; however, those high flow rates were capable of breaking the seal between the lipid bilayer and the AAO surface that prevented Pr3+ ions from entering this water-filled space, seen as a Pr3+-induced shift for almost all lipids. The data clearly point at the existence of a rather well sealed water volume between the AAO surface and attached lipid bilayers.
Trapped water volume. To probe this water volume 11 wt% PEG8000 in D2O was entrapped between the AAO surface and the single bilayer. From the intensity ratio of NMR signals of the lipid γ-choline resonance and the PEG8000 signal it is estimated that there are 64 molecules of trapped water per lipid in the outer leaflet of the bilayer apposing the AAO surface. Assuming an area per lipid molecule for POPC of 0.6 nm2, this corresponds to an average water layer thickness of 3 nm. However, at least a thin layer of lipids around the boundary of adsorbed lipid bilayers must have close contact to AAO such that PEG8000 remained trapped.
Nature of Lipid Attachment. The chemical shift of the 31P NMR resonance attributable to the AAO filters themselves suggests that the aluminum oxide surface is partially covered with phosphate groups that are connected via ester bonds to one or two aluminate units (Mcnatt, J.S. et al. (2003) Langmuir 19: 1148- 1153). In contrast, the absence of severe broadening of the lipid 31P NMR resonance, its chemical shift, as well as the 99% removal of lipid by flushing of pores with organic solvent indicate that AAO does not form covalent bonds with phospholipids. Instead, it is proposed that lipids are attached via hydrogen bonds between AAO hydroxyl- and phosphate groups and the lipid. Ionic interactions between charges in the AAO surface and the zwitterionic lipid headgroup may contribute to the attachment as well.
The order parameters of the lipid choline resonances of the outer monolayers at the AAO surface are identical to order parameters in the inner monolayer. Therefore the headgroup order and motions of the majority of lipids in the outer monolayer adjacent to the AAO surface are not influenced by the association, in good agreement with existence of a thick water layer between AAO and lipid underneath most of the bilayer. The data are consistent with only a small percentage of the lipids acting as points of attachment to the AAO surface.
Bilayer Patches. The apparent POPC diffusion rate of 4.6 x 10"13 m2/s, measured at a diffusion time of 200 ms, is much less than the 9.5 x 10"12 m2/s measured for large POPC liposomes at 30°C (Fillipov, A. et al. (2002) Biophys.J. 82:754).
Since there is no physical reason for reducing the lateral diffusion rates by almost an order of magnitude, this diffusion value indicates that lipid displacement along pores is confined to cylindrical bilayer patches. This is confirmed by the diffusion time dependence of apparent diffusion rates. The average length of those patches corresponds to the distance, 1, traveled over the diffusion time / = 2DappA ~ 0.4 μm, where Dapp is the apparent diffusion constant and Δ the diffusion time (Figures 10A and 10B). The uncertainty of the value of the POPC diffusion rate does not greatly impact the patch length estimate. Changing the value of the diffusion constant by 50% changes the patch length by only 0.1 μm.
Lipid order parameters and phase transitions. The deuterium spectra of the perdeuterated palmitic acid chain in POPC-d3ι were used to assess order and dynamics of lipids in adsorbed single bilayers. The molecular chain order parameters S(mol)i are indistinguishable from order parameters in multilamellar liposomes, which is remarkable considering the high sensitivity of order parameters to changes in lipid area per molecule as demonstrated in experiments conducted as a function of temperature or hydration. Our spectral resolution would have permitted us to resolve changes in order parameters smaller than ΔS = ±0.002, equivalent to changes in temperature (Holte, L.L. et al. (1995) Biophys. J. 68:2396-2403) of less than 1 K or changes in area per molecule (Gawrisch, K. et al. (2002) Chem. Phys. Lipids 116:135-151) of less than 0.2 A2. To maintain the same order parameters as in multilamellar liposomes the lipids in the outer monolayer must be fully hydrated, equivalent to a water concentration in excess of 20 water molecules per lipid (Koenig, B.W. et al. (1997) Biophys. J. 73:1954- 1966). Indirect evidence for the adsorption of a single bilayer along fixed lines that coincide with the boundaries of patches is provided by the phase behavior of DMPC adsorbed to the surface of the AAO pores. The reduction in DMPC area per molecule upon the fluid-gel phase transition at 23 °C from ao = 59.6 A2 in the fluid phase at 30 °C (Koenig, B.W. et al. (1997) Biophys. J. 73:1954-1966; Nagle, J.F. et al. (2000) Biochim. Biophys. Acta 1469:159-195) to 47.2 A2 in the gel phase at 10 °C interferes with AAO-lipid bilayer interaction. Assuming an elastic area compressibility modulus of K= 240 dyn/cm (Olbrich, K. et al. (2000) Biophys. J. 79:321-327) this area change generates a bilayer tension which exceeds by far the typical lysis tension of bilayers, τ= 10 mN/m (Olbrich, K. et al. (2000) Biophys. J. 79:321-327). Because of attachment at the boundaries, the bilayers break open at the phase transition. This explains the observed penetration of Pr3+ ions into the water layer between the solid support and the membrane when samples are cooled below the phase transition of the lipid. Furthermore, attachment of lipid bilayers to the AAO surface at the boundaries of patches is in agreement with the observed shift of the main phase transition to lower temperatures. A lysis tension of τ= 10 mN/m is equivalent to an area change, Δα = τ ■ a0 IK , of 2.5 A2. The corresponding change in membrane
energy from elastic stretching is Em J/mol, equivalent to a
Figure imgf000048_0001
lowering of the main lipid phase transition temperature AT - —^Tj of about 0.9
K, assuming an enthalpy of the main phase transition of DMPC of zLH=24.7 kJ/mol (Boggs, J.M. et al. (1985) Biochim. Biophys. Acta 816:221-233) and Tfg=296 K. This calculated shift agrees well with the observed shift of the transition midpoint by 1 K (see Figure 8). The tailing of the transition to lower temperatures suggests that there is a wide distribution of stresses present in the adsorbed bilayers.
Model of lipid adsorption. The experimental observations support the following model of bilayer adsorption to the cylindrical surface of pores in the anopore filters (Figures 10A and 10B): bilayers adsorb as wavy lipid tubules with an average length, /, of 0.4 μm. The seal between the AAO surface and the bilayer is formed by a thin line of lipids at the boundary of the adsorbed tubule. Between the AAO and the bilayer exists a trapped water volume that varies from close apposition to several nanometers in the middle of the patches, resulting in an average water layer thickness of 3 nm. Using a crude geometrical model of the wavy lipid tubules it is estimated that, on average, they have four inward constrictions before breaking up inside the pore.
How are such lipid tubules formed? Multilamellar liposomes become cylindrical after being forced into the pores. The cylinders are stable until their length reaches a critical value that depends on the surface bending energy.
Cylinders with variable diameter have lowest energy if they belong to the family of surfaces with constant total curvature J, called Delaunay surfaces (Delauney, C.
(1841) J. Math. Pures Appl. 6:309-315). The curvature energy of such cylindrical bilayers has been analyzed recently (Shemesh, T. et al. (2003) Biophys. J.
85:3813-3827). It was reported that the lipid cylinders have a tendency to form constrictions and to break up at a certain length, forming short, capped tubules or spherical vesicles.
The process of breaking up those cylinders inside pores appears to be somewhat different than in non-supported vesicles. It is proposed that the edges of the broken-up cylinders widen and adhere to the AAO surface, driven by attractive interactions between lipid and AAO. The bilayer edges form a good seal to the wall of AvAO pores that prevents penetration of Pr3+ ions into the trapped water layer. Taken together, these experiments point to possible applications for lipids loaded into AAO pores. The thick water layer should allow for both integral and peripheral membrane proteins to be included without perturbation. Moreover, solutes may be stored in the trapped water layer beneath the lipid. Considering the ease of preparation of such systems, lipids supported on AAO filters have considerable promise for use in biosensors.
Example 2 Reconstitution of the G-Protein Coupled Membrane Receptor Rhodopsin into Lipid Bilayers and Adsorption into AAO pores
Rhodopsin purification and reconstitution into POPC membranes. Rod outer segment discs from bovine retinas were solubilized in the detergent octylglucoside (OG), and the rhodopsin purified by affinity chromatography according to the procedure of Litman et al. (Litman BJ. (1982) "PURIFICATION OF RHODOPSIN BY CONCANAVALIN A AFFINITY CHROMATOGRAPHY," Methods Enzymology, 81, 150- 153) using a Pharmacia concanavaline A columnm (Pharmacia Biotech, Piscataway, NJ). Rhodopsin concentration after purification was determined by measuring light adsorption at 500 nm using a diode array UV/Vis spectrophotometer Agilent 8453 (Agilent Waldbronn, Germany). Care was taken to minimize exposure of samples to light during the experiment.
A proper amount of the monounsaturated chain-deuterated phospholipid 1- palmitoyl(d3ι)-2-oleoly-sn-glycero-3-phosphocholine (POPC-d3ι) or the polyunsaturated lipids 1-stearoyl (d35)-2-docosahexaenoyl-sn-glycero-3- phosphocholine (18:0(d35)-22:6 PC) (Avanti Polar Lipids) were dispersed in a micellar solution of OG in 10 mM PIPES buffer, pH 7.0, 100 mM NaCl, with 50 μm of the chelator DTPA added. The lipid dispersion was added to the purified rhodopsin dispersed in OG to yield a rhodopsin/lipid molar ratio in the range from 1/100 to 1/1000. The OG concentration was kept at a concentration in the range from 40-100 mM OG and the OG/lipid molar ratio was 10/1. Small liposomes with membrane incorporated rhodopsin were formed according to the dilution/reconstitution method of Jackson and Litman (Jackson M.L. and Litman, BJ. (1985) "RHODOPSIN-EGG PHOSPHATIDYLCHOLINE RECONSTITUTION BY AN OCTYL GLUCOSIDE DILUTION PROCEDURE," Biochim. Biophys. Acta 812, 369- 376). The OG-solubilized POPC-d3] -rhodopsin mixture was added drop-wise into detergent free PIPES buffer/NaCl solution at constant stirring. Upon dilution, the final concentration of OG in the sample was reduced to less than 10 mM OG which is well below the critical micelle concentration of OG. The procedure yielded 5 ml of a liposome dispersion with membrane incorporated rhodopsin. The final rhodopsin concentration was 0.37 mg/ml. Rhodopsin purification and reconstitution were conducted at ambient temperature in complete darkness using night vision goggles to avoid rhodopsin bleaching.
Adsorption into AAO nanopores. A mini-extruder (Avanti Polar Lipids, Alabaster, AL) or stainless steel thermobarrel extruder (Lipex Biomembranes, Inc; Vancouver, BC Canada) were loaded with up to five 13 mm diameter Whatman Anopore filters (SPI Supplies, West Chester, PA) with a nominal pore diameter of 0.2 μm. For sample preparation with the mini-extruder, Anopore filters were packed between two filter supports. A total of 5 ml of rhodopsin reconstituted into POPC-d3ι liposomes were sent through the filters in 1 ml increments at a rate of 0.06 ml/s using a gas-tight Hamilton syringe. Solutions that had passed through the filter were collected using a second syringe and rhodopsin concentration was determined by a measurement of light adsorption at 500 nm. The rhodopsin from the first three milliliters was completely adsorbed within the anopore filters (residual rhodopsin concentration in the solution that had passed through the filters was 0.01-0.02 mg/ml). The rhodopsin from the 4th milliliter was partially adsorbed (rhodopsin concentration 0.19 mg/ml), and the rhodopsin from the 5th milliliter passed the filters essentially without binding within anopore (rhodopsin concentration 0.35 mg/ml, compared to 0.37 mg/ml in the incoming solution). A total of 1.3 mg of rhodopsin did bind within the 5 anopore disks, corresponding to 0.26 mg of rhodopsin in 0.52 mg of POPC-d3ι bilayers per disk. Again, extrusion was performed in complete darkness to prevent the bleaching of rhodopsin.
For sample preparation using the stainless steel thermobarrel extruder the Anopore membranes were placed on a stainless steel grid and sealed against the upper barrel by an O-ring. In this approach, water was flushed through the filters several times before the lipid dispersion was passed through ten times using compressed argon at pressures in the range from 2 - 8 bar. The anopore filter disks with rhodopsin in lipid membranes were loaded into a flat glass cell that was filled with PIPES/NaCl buffer prepared in deuterium depleted water (Cambridge Isotopes, Cambridge MA) and sealed with a silicon stopper. The cell was inserted in darkness into a Doty flat cell probe for a DMX500 solid-state NMR. spectrometer (Bruker Biospin Inc., Billerica MA) equipped with a wide-bore 500 MHz NMR magnet (Oxford Instruments, Oxford U.K.). The flat cell was oriented such that the normal to the anopore filters was oriented parallel to the B0 field of the superconducting magnet. 2H NMR spectra of the chain deuterated POPC-d3ιwere acquired in quadrature detection mode at a resonance frequency of 76.8 MHz using the quadrupolar echo sequence π/2-τ- π/290-τ-acquire, with a π/2 pulse length of 3.4 μs and a delay time τ=75 μs. A total of 200,000 scans with a relaxation delay time, dj, of 250 ms, were acquired. Every scan consisted of 16 K data points recorded at a dwell time of 2.5 μs corresponding to a spectral width of 200 kHz. The carrier frequency of the spectrometer was adjusted to be exactly at the center of the symmetric spectra. For processing the real and imaginary parts of the free induction decay (FID) were transferred from the spectrometer computer to a Windows workstation. Further processing was conducted using a program written for Mathcad 200 li Professional (MathSoft Engineering & Education, Inc., Cambridge, MA). The phase of the FID was adjusted to minimize the signal in the imaginary channel. The location of the maximum of the quadrupolar echo was determined with a resolution of l/10th of a dwell time unit, and the time base of the spectra was corrected such that the FID began exactly at the echo maximum using a spline interpolation function to calculate new digital data points. The FID was multiplied with an exponential decaying window function corresponding to a line broadening of 100 Hz. After the Fourier transformation of the FID the spectra shown in Figure 11 and Figures 12A-H were obtained.
The spectra are characteristic for lipid bilayers oriented preferentially with their bilayer normal perpendicular to the magnetic field. This orientation of the bilayer normal is consistent with membranes adhering to the inner surface of pores as lipid tubules. With increasing amounts of protein in the samples we observed an increase of the linewidth of resonances and some increase of mosaic spread. The mosaic spread of preferred bilayer orientations was reduced after freezing the sample in a deep freezer before investigating it at ambient temperature. Exact values of sn-1 chain order parameters and of mosaic spread were determined by simulating the experimental spectra using a program written for Mathcad 200 li Professional (MathSoft Engineering & Education, Inc., Cambridge, MA). Order parameter analysis revealed a small reduction of hydrocarbon chain order in the order parameter plateau region (carbon atoms 2-8) due to the presence of the protein (see Figure 13). Mosaic spread could be reasonably well modeled as Gaussian distribution with a half width of 8 degrees (pure 18:0(d35)-22:6 PC bilayers) to about 20 degrees (18:0(d35)-22:6 PC bilayers containing rhodopsin at a lip id/protein molar ratio of 100/1.
The presence of the protein in the reconstituted membranes is also detectable as a substantial reduction of spin-spin relaxation times (broadening of resonance lines) and small reductions of spin-lattice relaxation times of rapidly moving lipid segments.
It was established that the residual OG in the preparation was easily removed by flushing the loaded anopore filters with several ml of PIPES/NaCl buffer. Removal of OG resulted in a slight increase of lipid order. After OG removal lipid order was identical to lipid order in rhodopsin-containing liposomes after OG removal by dialysis over 14-24 hours against at least 50-fold excess of PIPES/NaCl buffer containing 3 g/1 of detergent-adsorbing SM-2 Bio-Beads (Bio- Rad Laboratories, Hercules, CA). For dialysis the rhodopsin-containing liposomes were kept in a slide-A-Lyser dialysis cassette (Pierce Chemical Company,
Rockford, IL). It is remarkable that in anopore membranes a similar degree of removal of OG was achieved in less than one minute by simply flushing the filters with buffer. The reconstituted membranes within anopore filters could be stored in a deep freezer for one month and possibly much longer, without compromising membrane integrity.
The anopore filters containing reconstituted rhodpsin had a bright pink color that turned to yellow within one minute after exposure to light. This transition is consistent with conversion of dark-adapoted rhodopsin to a meta- I/meta-II rhodopsin equilibrium, demonstrating that rhodopsin was successfully adsorbed and functional.
Example 3 Adsorption of E. coli Protoplast Membranes with Incorporated Human Peripheral Cannabinoid Receptor (CB2) into AAO Pores Human peripheral cannabinoid receptor (CB2) was expressed in E. coli BL21 cells as a fusion protein, containing E. coli maltose-binding protein (MBP) attached at the N-terminal end of CB2, and thioredoxin followed by a stretch often histidine residues attached at the C-terminal end of CB2. Cell density was of the order of 109 cells per ml. Every cell expressed about 1,000 copies of CB2.
E-coli cytoplasmic membrane preparation. By Western Blot analysis using specific antibodies we established that CB2 is preferentially located in the cytoplasmic membranes of E. coli. Formation of spheroplasts and fractionation to obtain cytoplasmic membranes were conducted according to the protocols by R.L. Weiss (Weiss, R.L. (1976) "PROTOPLAST FORMATION IN ESCHERICHIA COLI," J. Bacteriol. 128:668-670) and Thai and Kaplan (Tai, S.-P. et al. (1985) "INTRACELLULAR LOCALIZATION OF PHOSPHOLIPIDS TRANSFER ACTIVITY IN RHODOPSEUDOMONAS SPHAEROIDES AND A POSSIBLE ROLE IN MEMBRANE BIOGENESIS," J. Bacteriol. 164: 181-186) with some modifications as outlined below.
Briefly, E. coli cells were collected by centrifugation, washed twice with 0.1 M Tris-HCl pH 8.0 buffer and re-suspended in 0.1 M Tris-HCl buffer pH 8.0 containing 20% (w/v) sucrose, such that the resulting cell suspension had an optical density of 10 at 600 nm. A cocktail of protease inhibitors (F. Hoffmann-La Roche Ltd, Basel, Switzerland) was added to prevent enzymatic digestion of CB2.
The temperature was adjusted to 37° C, and a solution of lysozyme (2 mg/ml) was added slowly, under constant stirring, until a final lysozyme concentration of 0.1 mg/ml was reached. The cell suspension was incubated at 37° C for additional 15 minutes.
To disrupt the outer membrane lipopolysaccharides, a solution of 0.1 M EDTA (pH 7.0) was added slowly under continuous stirring, until a final EDTA concentration of 10 mM was reached. Incubation continued for another 10 minutes. The spheroplasts were centrifuged at 12,000 g for 20 minutes, the pellet collected and washed once with 0.1 M Tris-HCl buffer, pH 8.0, containing 20% sucrose. Spheroplasts were centrifuged again at 12,000 for 20 minutes. The pellet of spheroplasts was re-suspended in ice-cold water, resulting in osmotic lysis of sphoroplasts. Immediately, 1 M Tris buffer pH 8.0 containing protease inhibitors was added, to produce the final Tris concentration of 50 mM. A 1M solution of MgCl2 was added to yield a final concentration 1 mM. A solution of DNAse was added, the cell-free extract briefly sonicated, and incubated on ice for 1 hour. The extract was centrifuged at 150,000 g for 1 hour, the membrane pellet washed with Tris buffer, and centrifuged again at 150,000 for 1 hour. The membrane pellet was re-suspended in a small volume of Tris buffer, frozen in liquid nitrogen, and stored at -80° C before use.
For 1H magic angle spinning (MAS) NMR experiments one milliliter of the cytoplasmic membrane preparation was suspended in Tris buffer prepared in 99.9% D2O (Cambridge Isotopes, Cambridge MA) and pelleted at 150,000 g for thirty minutes. The pellet with a volume of approximately 15 μL was transferred by centrifugation to a 4 mm outer diameter MAS rotor outfitted with a Kel-F insert to generate a spherical sample volume of 11 μL (Bruker Biospin Inc., Billerica MA). The sample was spun at a MAS frequency of 5 kHz using a Bruker H-X resonance MAS probe for a solid state DMX300 NMR spectrometer (Bruker Biospin, Billerica MA) equipped with a Bruker widebore 300 MHz magnet. Proton resonance spectra were acquired at ambient temperature using a π/2 pulse length of 4 μs and Cyclops phase cycling. About 1,000 free induction decays (FID) with a relaxation delay of 4 s were acquired. Before Fourier transformation the FID was multiplied with an exponential window function equivalent to a signal broadening of 1Hz.
In a second experiment 1 ml of the cytoplasmic membrane preparation was extruded through two 13 mm diameter anopore filters with a pore size of 0.2 μm using a mini-extruder (Avanti Polar Lipids, Alabaster, AL). The top filter was discarded and the second filter carefully cut into fine pieces using a razor blade. The pieces were transferred to a 4 mm OD MAS rotor with a spherical Kel-F insert. The anopore filter pieces were collected in the spherical volume by centrifugation before inserting the upper piece of the insert. For !H MAS NMR experiments the sample was spun at a MAS frequency of 5 kHz and the Η NMR spectra acquired as described above.
The [H MAS NMR spectra of the protoplast pellet and of protoplast membranes adsorbed into anopore filters (Figure 14, Trace A and Trace B) indicate that the cytoplasmic membranes containing the CB2 receptor was successfully deposited inside anopore filters, ready to be used for ligand binding studies. Preliminary ligand- and G-protein binding studies using standard protocols for membrane preparations confirmed the presence of functional CB2 in those preparations. The present invention enables the development of a protocol to conduct quantitative binding studies using cytoplasmic membranes deposited into anopore disks. Example 4 Porous Substrates Reduce Nonspecific Hydrophobic Ligand Binding Anopore filters were used to perform the ligand-binding assay on the E. coli membranes expressing recombinant cannabinoid receptor. Briefly, the procedure used to perform the binding assay was as follows: Anopore filters (25 mm diameter, 0.2 μm pore size) were pre-loaded with the E. coli BL21-107 protoblast membranes expressing CB2 by slowly passing suspension of the membranes in 50 mM Tris-HCl buffer pH 7. 5, on a Millipore vacuum-filtration manifold. Filters were washed by passing 10 ml of binding buffer containing 50 mM Tris buffer pH 7.5, 3 mM MgCl2, and 0.1% bovine serum albumine. A 1 ml solution of radiolabeled ligand and variable concentrations of unlabeled ligand (CP55,940) in binding buffer were slowly passed through the Anopore filters with application of vacuum. Filters were washed three times by passing 2 ml of binding buffer. The filters were then immersed into the scintillation liquid and the activity counted.
Radioligand [3H]CP55,940 bound to the CB2 receptor deposited into the Anopore filters could be competed off by increased concentrations of unlabeled ligand CP 55,940. Ligand-binding parameters determined for the CB2 receptor deposited into Anopore filters were identical to the ligand-binding characteristics of CB2 receptor measured by a conventional filter-binding assay (Figure 15).
These results indicate that deposition on the Anopore surface preserves functional activity of CB2 receptor.
The conventional competitive filter-binding assay was performed by incubating a suspension of E. coli membranes with radioligand [3H]CP55,940 and variable concentrations of the unlabeled competing ligand. Upon incubation, the reaction mixture was rapidly filtered through Whatman GF/B paper filters, and retained activity on the filters was counted with a scintillation counter. This assay typically results in significant nonspecific binding of the hydrophobic ligand (CP55,940) in the multilamellar deposits on the filter surface which requires to work with the active receptor at much higher concentrations and reduces accuracy and reproducibility of the binding parameters K and Bmax. In the convential assay nonspecific binding constitutes 40-50% of the total radioactivity count.
In contrast, the assay performed on E. coli membranes deposited as single tubular lipid layers into Anopore filters resulted in significant reduction of background binding to values of less than 15% (Figure 16). Thus, the use of single tubular membrane layers in porous substrates offers significant promise as solid support in receptor-ligand interaction studies. Careful inspection of the filter material revealed that the use of O-rings in the filter devices was the most important source of nonspecific background radioactivity when using Anopore porous filters. We expect a further reduction in background radioactivity from using a device without O-rings.
Two important conclusions can be drawn from the above experiments: 1. Deposition of the recombinant receptor into the porous material preserves functional activity. 2. The use of single tubular lipid bilayers in porous substrates reduces nonspecific binding to very low levels, thus permitting a more accurate determination of ligand binding parameters.
Example 5 Reconstitution of Purified Recombinant Cannabinoid Receptor, Adsorption into AAO Pores, and Detection of Membrane Adsorption by Fluorescence Spectroscopy Human peripheral cannabinoid receptor (CB2) was expressed in E. coli BL21 cells as a fusion protein, containing E. coli maltose-binding protein (MBP) attached at the N-terminal end of CB2, and thioredoxin followed by a stretch often histidine residues attached at the C-terminal end of CB2. Recombinant protein was solubilized from the bacterial membranes in a mixture of 0.5% CHAPS, 0.1 % cholesteryl hemisuccinate and 1% of dodecylmaltoside, and purified to approximately 90% purity by affinity chromatography on Ni-agarose (Qiagen) and ion-exchange chromatography on HiTrap-Q Sepharose (GE-Amersham Biosciences).
Labeling of protein. Purified fusion protein was covalently labeled with AlexaFluoro 532 carboxylic acid, succinimidyl ester (Molecular Probes) according to the protocol recommended by the manufacturer. Non-reacted fluorescent dye was (partially) removed by sequential gel-filtration (PD-10 desalting columns, Amersham), centrifugation in the Centricon-30 filter device (TVIillipore) and dialysis (Slide-A-Lyzer, Pierce). About 200 μg of labeled protein was obtained.
Labeling of lipid. 150 μg of l-stearoyl-2-oleoyl-sn-glycero-3 -phosphocholine (SOPC, Avanti) were dissolved in methanol and mixed with a methanol solution of 1.5 μg of Texas Red l,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine, triethylammonium salt (Texas Red DHPE, Molecular Probes). The methanol was removed under the stream of nitrogen, and lipids were dispersed in a solution of 3% octylglucoside (OG) in 50 mM Tris-HCl buffer pH 7.5.
Reconstitution of recombinant CB2 into lipid bilayer and deposition into Anopore membranes. A mixture of 150 μg of SOPC and 7.5 μg of fluorescently labeled CB2 dissolved in 1 ml of 3% OG in Tris-HCl buffer, pH 7.5 was diluted with 9 ml of Tris-HCl buffer, pH 7.5. The diluted solution was kept at room temperature for 30 min, and then passed through 25 mm Anodiscs (Whatman, 0.2 um thickness) at a flow rate of approximately 5 ml/min. Discs were washed twice with 5 ml of Tris-HCl buffer, and kept wet until ready for scanning.
Scanning of Anopore membranes. Wet Anopore discs were scanned using a Typhoon 8600 fluorescent scanner (Molecular Dynamics) at the following instrument settings: for AlexaFluoro 532: excitation 532 nm (green laser); emission 580 nm, PMT 320 V, normal sensitivity; for Texas Red: excitation 532 nm (green laser), emission 610 nm, PMT 320 V, normal sensitivity. The results of experiments are presented in Figure 17, Panel A and Panel B.
Filters containing a mixture of fluorescence labeled CB2 and SOPC (samples 1 and 3) produced bright fluorescence at 580 nm (excitation at 532 nm), characteristic for AlexaFluoro 532 fluorophore. Intensity of fluorescence of sample 3 containing labeled CB2 as well as labeled lipid was lower than the fluorescence of sample 1 which contained labeled CB2 and non-labeled lipid, the effect that can possibly be attributed to quenching of fluorescence of the AlexaFluoro532 fluorophore by the Texas Red. If labeled CB2 was passed through anopore filters without prior addition of lipid (sample 5) a significantly lower fluorescence of the respective Anodisc was obtained (compare with samples 1 and 3). Filtration of the solution of the non- reacted dye AlexaFluoro532 through Anodiscs resulted in much lower levels of fluorescence of the corresponding Anodisc (sample 6), suggesting that the remaining intensity in the previous experiment could be a weak background signal from unreacted dye. Addition of SOPC to the solution of free AlexaFluoro532 dye (samples 2 and 4) increased background fluorescence somewhat. However, the resulting intensity of fluorescence in these samples was still significantly lower than the intensity of fluorescence derived from the labeled CB2 samples deposited into Anodiscs in the presence of lipid (samples 1 and 3). The presence of lipid in the reconstituted samples was confirmed by a scan at 610 nm that confirmed the presence of the lipid label Texas Red (samples 3, 4, and 7).
The results presented above confirm unambiguously that CB2 adheres to the inner surface of anopore filters only after incorporation into SOPC bilayers. The binding of the lipids to the surface of Anodiscs was confirmed by using fluorescently labeled (Texas Red DHPE) lipids. Thus, it is possible to use dual fluorescent labeling of both protein and lipids to follow the reconstitution of receptor into the lipid bilayer, and their depositioning onto the inner pore surface of Anopore membranes, and to quantify the amounts of lipid and protein deposited onto Anodiscs using fluorescence spectroscopy.
Quantification of the amounts of lipid and CB2 protein deposited onto Anopore filters was performed as follows. SOPC was mixed with the Dil-Cl 8 DilC i s ( 1 , 1 ' -dioctadecyl-3 , 3 ,3 ' ,3 ' -tetramethy lindodicarbocyanine, 4- chlorobenzenesulfonate salt) fluorescently labeled lipid at a ratio of 1000: 1 (w/w, SOPC to Dil-Cl 8) in either 3% OG or mixture of triple detergents (0.5% CHAPS, 0.1% CHS, 0.1% DM). SOPC/ Dil C18 mix was rapidly diluted (20-fold) into 50 mM Tris-HCl buffer pH 7.5, and solution was slowly filtered through the Anopore filter. The fluorescence of the lipid retained on the filter was measured by scanning of the wet filter on Typhoon 8600 fluorescence scanner at following settings: excitation: 633 nm, emission: 670 nm, PMT: 450, sensitivity: normal.
A typical lipid depositioning curve obtained in these experiments is presented in Figure 18. Up to 250 μg of SOPC can be deposited per sing Anopore filter (25 mm diameter, 0.2 μm pore size).
In order to determine the amounts of CB2 protein deposited onto anopore filter, the following experiments were conducted. Variable amounts of the purified CB2 labeled with Alexa Fluor 532 in triple detergent mix (0.5% CHAPS, 0.1% CHS, 0.1% DM) were mixed with 250 μg of SOPC/DHC18 dissolved in 3% OG. Upon 20-fold dilution into 50 mM Tris-HCl buffer pH 7.5 the mixture was slowly filtered through the Anopore filters, and filters were further subjected to extensive wash with Tris-HCl buffer. Wet filters were scanned at the following settings of the Typhoon 8600 fluorescence scanner:
excitation: 532 nm, emission: 555 nm, PMT: 450, sensitivity: normal. Results are summarized in Figure 19. Up to 10 μg of CB2 can be deposited per single Anopore filter.
The amounts of lipid retained in the presence of increasing concentrations of protein was determined by scanning of the Anopore filters at the following settings of the Typhoon 8600 fluorescence scanner: excitation: 633 nm, emission: 670 nm, PMT: 450, sensitivity: normal. Depositioning of SOPC into Anopore filters decreased with the increasing CB2 concentration, reaching saturation at a CB2 concentration of 10 μg per sample (Figure 20).
It was demonstrated that simultaneous depositioning of CB2 and SOPC into Anopore filters (25 mm diameter, 0.2 μm pore size) permits retaining of 10 μg of the CB2 protein and 100-120 μg of SOPC per filter. A further increase in the protein/SOPC ratio did not result in depositioning of larger amounts of lip id or protein. The achieved CB2 concentration is well within range of concentrations required for highly accurate ligand binding studies that are currently underway. A further increase of deposited protein concentration is expected after cleaving the strongly negatively charged maltose binding protein from the fusion taking advantage of the Tev-protease cleavage site.
The integrity of the CB2 receptor before and after depositioning into the pores of the Anopore filters, in the presence of lipid matrix (SOPC), was confirmed by SDS-page and Western-blot analysis. Anopore filters were loaded with purified CB2 reconstituted into POPC matrix. A mixture of 10 μg of purified recombinant CB2 and 200 μg of POPC in 3% octyl glucoside was rapidly diluted 20-fold in 50 mM Tris-HCl buffer pH 7.5. The resulting liposome dispersion was passed through the Anopore filters at a low flow rate of 2 ml/ min, and filters were further subjected to extensive washing with Tris-HCl buffer. A solution of 2% SDS was than applied to the filter, and the eluted material collected and subjected to the SDS-PAGE and Western-blot analysis. Figure 21 demonstrates that CB2 receptor can be eluted from the Anopore filter in the presence of strong ionic detergent SDS, and that no degradation of the receptor occurred during deposition/ elution procedures.
All publications and patents mentioned in this specification are herein incorporated by reference to the same extent as if each individual publication or patent application was specifically and individually indicated to be incorporated by reference.
While the invention has been described in connection with specific embodiments thereof, it will be understood that it is capable of further modifications and this application is intended to cover any variations, uses, or adaptations of the invention following, in general, the principles of the invention and including such departures from the present disclosure as come within lαiown or customary practice within the art to which the invention pertains and as may be applied to the essential features hereinbefore set forth.

Claims

What is Claimed Is:
Claim 1. A composition comprising a single bilayer lipid membrane supported on a solid support, wherein said membrane comprises a lipid tubule having a wavy tubular geometry and rims, wherein said tubule has rims that are in contact with the solid support, and a center region between said rims that is spaced apart from said support by an aqueous cushion, said aqueous cushion being located between said membrane and said support.
Claim 2. The composition of claim 1, wherein said solid support is a porous aluminum oxide support.
Claim 3. The composition of claim 1, wherein said lipid comprises a phospholipid.
Claim 4. The composition of claim 1, wherein said membrane comprises an incorporated biological molecule.
Claim 5. The composition of claim 4, wherein said incorporated biological molecule is an integral membrane protein or a peripheral membrane protein.
Claim 6. The composition of claim 5, wherein said integral membrane protein or said peripheral membrane protein is a receptor molecule, an enzyme, or an antigen.
Claim 7. The composition of claim 6, wherein said incorporated biological molecule is a receptor molecule capable of binding an agonist or an antagonist of a signal transduction pathway.
Claim 8. The composition of claim 7, wherein said receptor molecule is a G- protein coupled membrane receptor.
Claim 9. The composition of claim 8, wherein said solid support is porous, and wherein an agonist or antagonist of said receptor, a G-protein, or an analog of such molecules is provided to said composition by flowing a solution through the pores of said porous support so as to permit said receptor agonist or antagonist to bind to the receptor from inside said lipid tubule from the side of the lipid monolayer opposite from the AAO surface.
Claim 10. The composition of claim 9, wherein said agonist or antagonist of said receptor, said G-protein, or said analog of such molecules receptor agonist or antagonist is contained in said aqueous cushion.
Claim 11. A biosensor device comprising a solid support, a biological component, and a physical transducer; wherein: said biological component comprises a single bilayer lipid membrane supported on said solid support, wherein said membrane comprises a lipid tubule having a wavy tubular geometry and rims, wherein said tubule has rims that are in contact with the solid support, and a center region between sai rims that is spaced apart from said support by an aqueous cushion, said aqueous cushion being located between said membrane and said support; said membrane comprises an incorporated biological molecule selected from the group consisting of an enzyme, a receptor molecule, and an antigen; and said physical transducer serves to generate a signal in response to the binding of a target molecule to said incorporated biological molecule.
Claim 12. The biosensor device of claim 11, wherein said solid support is a porous aluminum oxide support.
Claim 13. The biosensor device of claim 11, wherein said lipid comprises a phospholipid.
Claim 14. The biosensor device of claim 11, wherein said incorporated biological molecule is a receptor molecule.
Claim 15. The biosensor device of claim 14, wherein said receptor molecule is a G-protein coupled membrane receptor.
Claim 16. The biosensor device of claim 15, wherein said solid support is porous, and wherein an agonist or antagonist of said receptor, a G- protein, or an analog of such molecules is provided to said composition by flowing a solution through the pores of said porous support so as to permit said receptor agonist or antagonist to bind to the receptor from inside said lipid tubule from the side of the lipid monolayer opposite from the AAO surface.
Claim 17. The biosensor device of claim 14, wherein said aqueous cushion contains a biological molecule that interacts with said receptor molecule.
Claim 18. The biosensor device of claim 16, wherein said aqueous cushion contains a G-Protein, and said incorporated biological molecule is a G-protein coupled membrane receptor.
Claim 19. The biosensor device of claim 11, wherein said physical transducer comprises an optical sensor, an electrochemical sensor, a potentiometric sensor, a conductometric sensor, or a piezoelectrical sensor; wherein said sensor generates a detectable physical signal in response to the binding of a target biological molecule to said biosensor's incorporated biological molecule.
Claim 20. The biosensor device of claim 19, wherein said physical transducer is an optical sensor selected from the group consisting of a colorimetric optical sensor operating in the visible or non-visible spectral range.
Claim 21. The biosensor device of claim 19, wherein said optical sensor is an optical sensor that detects fluorescent light.
Claim 22. A method of detecting a pharmacological agent that binds to a biological molecule, wherein said biological molecule is selected from the group consisting of an enzyme, a receptor molecule, an antigen, and an antibody, wherein said method comprises the steps: (A) incubating said pharmacological agent in the presence of a biosensor device, said biosensor device comprising a solid support, a biological component, and a physical transducer; wherein said biological component comprises a single bilayer lipid membrane supported on said solid support, wherein: said membrane comprises a lipid tubule having a wa^vy tubular geometry and rims, wherein said tubule has rims that are in contact with the solid support, and a center region between said rims that is spaced apart from said support by an aqueous cushion, said aqueous cushion being located between said membrane and said support; said membrane comprises said biological molecule incorporated therein; and said physical transducer serves to generate a signal in response to the binding of a target molecule to said incorporated biological molecule; and (B) determining whether said physical transducer generates a signal, said generated signal indicating that said pharmacological agent binds to said biological molecule.
Claim 23. The method of claim 22, wherein said biological agent is a G- protein coupled membrane receptor, and said method detects a pharmacological agent that binds to a G-protein coupled membrane receptor.
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