Patent Grant
10576456
The present invention relates to systems and methods of preparing lipid assemblies for enhancing stability, in particular, systems and methods for stabilizing synthetic and natural phospholipid membranes using direct polymerization of lipid monomers or polymer scaffolding of non-lipid monomers.
Section 1
Suspended lipid bilayers, also known as Black lipid membranes (BLMs), are a model system used to study the function and activity of transmembrane proteins, engineered proteins, phospholipid organization, and as a key component of ion channel-based biosensors. Ion channels (IC) possess a number of desirable properties that make them useful for analytical applications, including ion selectivity, chemical or mechanical gating, inherent signal amplification, well-defined open and closed states and simple electrical readout. BLMs provide an important synthetic membrane environment to study the function and activity of ion channels and serve as key components of ion channel-functionalized analytical platforms.
Additionally, BLMs have potential in high-throughput applications, including drug screening, due to the formation of an array of BLM-IC-based biosensors. A major limitation of BLM-based platforms (i.e. ion channel-functionalized sensor platforms) is the ability to form membranes with adequate electrical, mechanical and temporal stability. BLM instability arises from the relatively weak noncovalent forces of interaction between lipid molecules in the membrane, which are insufficient to maintain the structure of BLMs under mechanical, chemical and electrical stresses. Further, the interaction forces between the lipid membrane and the underlying substrate significantly affect the temporal stability of BLMs.
The development of robust BLMs has been a major focus of research efforts. A few of the methods developed to enhance the formation and stability of BLMs include; miniaturization of aperture size, reducing the surface energy of aperture substrates, sandwiching the BLM between hydrogel layers, and chemical cross-linking by photopolymerization of reactive amphiphiles. Benz et al. pioneered the direct polymerization of lipid membranes as a method of stabilizing BLMs, and identified lipid compositions for developing synthetic ion channel-functionalized sensors (Benz, R.; Elbert, R.; Prass, W.; Ringsdorf, H. Eur. Biophys. J. 1986, 14, 83-92). Reactive chemical functionalities can be introduced in the structure of lipid amphiphiles during synthesis to allow cross-linking at the lipid head group, the middle or the distal end of the lipid tail, or via a linker attached to the lipid head group. The degree of cross-linking in polymeric membranes depends on the type of polymerizable lipid and method of polymerization used, and affects the fluidity and stability of the lipid membranes. While polymerization can significantly enhance the stability of BLMs, stiff, viscous polymeric membranes may inhibit the function of some ICs.
A number of approaches have been explored to address the challenge of creating stable membranes that retain fluidity. Schmidt and coworkers created stable long lived BLM platforms for single-channel measurements by encapsulating a pre-existing free-standing membrane within a gel polymerized around it in situ (Jeon, T. J.; Malmstadt, N.; Schmidt, J. J. J. Am. Chem. Soc. 2006, 128, 42-43). Although the lifetime of BLM was greatly enhanced, the method reduces the effective diffusion of IC into BLM by ca. 70%, thus increasing IC reconstitution time and making application of the method for sensor development impractical. BLMs have been prepared from mixtures of polymerizable and nonpolymerizable phospholipids which allowed adequate fluidity to observe normal ion channel activity. Shenoy and co-workers reported improved bilayer lifetime using a mixture of polymerizable and non-polymerizable lipids (Daly, S. M.; Heffernan, L. A.; Barger, W. R.; Shenoy, D. K. Langmuir 2005, 22, 1215-1222). They observed wide fluctuations in the lifetime of UV irradiated BLMs due to variation in the amount of reactive polymerizable lipids that formed the BLM. To address the challenge of membrane fluidity, Heitz et al demonstrated the preparation of highly stable BLMs from a mixture of polymerizable (bis-dienoyl phosphatidylcholine) and nonpolymerizable (1, 2-diphytanoyl-sn-glycero-3-phosphocholine (DPhPC)) phospholipids, a mixture that retained sufficient fluidity for reconstitution and proper function of ion channels and allowed for observation of normal IC activity (Heitz, B. A.; Jones, I. W.; Hall, H. K.; Aspinwall, C. A.; Saavedra, S. S. J. Am. Chem. Soc. 2010, 132, 7086-7093). Meier et al. used a different approach to enhance the electrical stability of free standing lipid membranes, in which BLMs were formed from a mixture of nonpolymerizable lipids and nonlipid hydrophobic monomers consisting of polymerizable styrene and divinylbenzene (Meier, W.; Graff, A.; Diederich, A.; Winterhalter, M. Phys Chem Chem Phys 2000, 2, 4559-4562). Despite the enhanced electrical stability of BLMs after UV-initiated polymerization, the longevity of the BLMs, fluidity and compatibility with ion channel reconstitution were not investigated.
Here, the present invention features a simple and cost effective method of improving the stability of BLMs from a mixture of nonpolymerizable lipids and commercially available, hydrophobic polymerizable monomers (i.e. methacrylate monomers) that partition into the lamella region of the lipid bilayer. BLMs prepared in equimolar mixture with nonlipid, hydrophobic monomers (BMA and EDGMA) were evaluated for their electrical, mechanical and physical properties before and after UV photopolymerization. The present invention shows dramatically enhanced BLM stability and maintenance of incorporated ion channel activity.
Section 2
Phospholipid membranes play key roles in the regulation of biological function by serving as a barrier between the extracellular and intracellular environments, as well as in the evaluation of physiological and pharmaceutical modulators of biological function. Additionally, phospholipid membranes provide a suitable chemical environment for expression and solubilization of transmembrane proteins. Due to the importance of phospholipid membranes in biological function, many pharmaceutical modulators interact with the macromolecular assemblies either via direct partitioning into the membrane or through interactions with transmembrane proteins.
Current drug screening assays commonly utilize intact cells to identify novel small molecule agonists and antagonists that interact directly with membranes or, more specifically, transmembrane proteins. Unfortunately, cell-based assays suffer from irreproducibility due to variability among heterogeneous cell populations, exhibit false positives and false negatives due to non-specific interactions, and are difficult to interpret due to the complexity associated with monitoring downstream effects of signal transduction. In contrast, affinity chromatography platforms that integrate phospholipid membranes present a unique capability for identifying compounds that interact directly with the membrane or with integrated membrane proteins. Additionally, membrane-functionalized affinity platforms tend to minimize non-specific interactions.
Phospholipid membrane-functionalized affinity stationary phases have been utilized in chromatography to study partitioning and binding interactions. In immobilized liposome chromatography (ILC), liposomes are retained on a support matrix through steric, hydrophobic, covalent, avidin-biotin, or other types of specific or non-specific interactions. ILC has been primarily used to study small molecule partitioning through lipid membranes and interactions between peptides and phospholipids. Various membrane proteins have been immobilized in ILC stationary phases and used to study ligand binding; however, the liposomes are formed by non-covalent interactions, which are inherently unstable. Thus, the stationary phases have limited lifetimes, as well as reduced pressure stability, requiring low flow rates that reduce separation efficiency. Furthermore, ILC phases lack the chemical and mechanical stability to withstand variations in solution conditions (e.g. small fractions of organic solvents, varying ionic strength, etc.) and physical and mechanical insults (air bubbles, shear forces, etc.), decreasing the reproducibility of the columns and limiting their utility.
Immobilized artificial membranes (IAMs) provide an alternative chromatographic stationary phase that exhibits greater stability and reproducibility than ILCs. IAMs are prepared by covalent attachment of lipid tails to an underlying support, resulting in formation of a lipid monolayer on the particle surface. IAMs have been used to study partitioning and interactions between small molecules and phospholipids. Additionally, nicotinic acetylcholine receptors, μ and κ opioid receptors, and other membrane proteins were separately immobilized in IAMs and packed into columns. Using frontal chromatography, dissociation constants for the various membrane proteins were calculated against a series of small molecules, revealing similar trends to binding constants calculated using cell-based assays. However, there were quantitative differences, likely due to the altered conformation of membrane proteins upon interaction with the underlying silica support and the truncated lipid membrane.
Affinity chromatographic matrices for analyzing membrane proteins and molecules that interact with them would benefit from the presence of a more stable lipid structure that more accurately represents a lipid bilayer to allow incorporation of a larger number of membrane proteins, while maintaining their native conformations. Bilayer stability can be increased by a number of methods, such as incorporating cholesterol, adsorbing a protective overlayer, membrane tethering, and polymerizing phospholipid monomers. Of these methods, direct polymerization of phospholipid monomers yields the most stable structures.
Polymerizable phospholipids have been used in various analytical platforms to form stable phospholipid bilayers. As a non-limiting example, planar supported lipid bilayers (PSLBs) prepared by polymerizing bis-SorbPC (1,2-bis[10-(2′,4′-hexadieoyloxy)decanoyl]-sn-glycero-2-phosphocholine) have exhibited stability against surfactants, organics, and exposure to high vacuum. Polymerized bis-SorbPC (poly(bis-SorbPC)) membranes have also been utilized in capillary zone electrophoresis as stable surface coatings for reducing the electro-osmotic flow and minimizing non-specific adsorption of proteins. These polymerized surface coatings were stable to surfactant solutions, shear forces, applied electric fields, and dry storage. Additionally, Rhodopsin, a transmembrane protein, was incorporated into PSLBs prepared from poly(bis-SorbPC) and retained its activity in the stabilized bilayer. When combined, these data support further investigation of polymeric lipid bilayers for enhancing stability of phospholipid-based stationary phases and the utility of the resulting polymeric lipid stationary phases for chromatographic separations.
Though polymeric-lipid coatings have been used to minimize non-specific adsorption of proteins in a range of materials, to the knowledge of the inventors, polymeric-lipid membranes have not yet been utilized as a stationary phase material in packed columns. The present invention features a method of preparing poly(bis-SorbPC) coatings onto silica particles that were subsequently packed into capillary LC columns. The chemical, physical, and temporal stability of the polymerized-phospholipid bilayers were assessed and their utility as a lipid-based stationary phase for liquid chromatography is demonstrated.
Polymer lipid membranes can be prepared by photochemical or redox initiated polymerization of synthetic, polymerizable lipid. The resulting polymerized stationary phases exhibited enhanced stability compared to particles coated with non-polymerizable lipid bilayers when exposed to chemical and physical assaults over a period of time. However, a drawback to redox polymerization is that initiation and progression of the polymerization using conventional redox mixtures proceed only under acidic conditions, which poses a problem for proteins incorporated into the lipid bilayer membrane. For example, transmembrane proteins often get denatured, and activities of these proteins can diminish or disappear after redox polymerization with current redox mixtures. Hence there is a need for redox mixtures that provide milder conditions when used in the polymerization of lipids. The present invention features a redox mixture that allows for polymerization methods to proceed under neutral pH conditions while preserving the proteins incorporated into the polymerized lipid membrane.
Any feature or combination of features described herein are included within the scope of the present invention provided that the features included in any such combination are not mutually inconsistent as will be apparent from the context, this specification, and the knowledge of one of ordinary skill in the art. Additional advantages and aspects of the present invention are apparent in the following detailed description and claims.
Section 1
Black lipid membranes (BLMs) provide a synthetic environment that facilitates measurement of ion channel activity in diverse analytical platforms. For example, the development of next-generation transmembrane protein-based biosensors relies heavily on the use of BLMs. BLMs instability resulting in rupture within hours (<4 h) of formation poses a significant challenge to biosensor development. Enhanced temporal, mechanical, and electrical stability of BLMs is needed to support the development of membrane-based biosensors and other highly stable measurement platforms.
Suspended lipid bilayers formed across glass apertures are greatly affected by the surface energy of the underlying substrate. Thus, glass pipette apertures were modified with silanes that define the surface energy of the underlying substrate, which affects the stability of the suspended lipid bilayer. The silanized microapertures typically form a hybrid bilayer membrane (HBM) with the phospholipid by adsorption through the assembly of the lipid monolayer on the hydrophobic substrate. The assembly of a lipid monolayer oriented in a tail-down configuration is due to van der Waals and hydrophobic forces of interaction between the hydrophobic lipid tails and the hydrophobic monolayer of the substrate.
When suspended across a decreased energy surface of a silanized aperture, the enhanced stability of BLMs may be due to the amphiphobic (H2O/oil repellency) character induced by PFDCS modifiers on the surface of aperture substrates. The force of interaction between the lipid membrane and the substrate (FMS) enhanced the electrical and mechanical stability of the BLMs by 5 and >25 fold respectively in comparison to conventional 3-cyanopropyldimethylchlorosilane (CPDCS) pipette apertures.
Despite the substantial increase in the stability of BLMs that were suspended across decreased energy surfaces of apertures, the inherent weak hydrophobic force of interaction between the self-assembled lipid molecules (FH) and electrostatic forces (FE) limits the average lifetime of the BLMs to ca 8±1 h. To overcome the weak hydrophobic forces, small polymerizable hydrophobic monomers (butylmethacrylate (BMA) and ethylene glycol dimethacrylate (EGDMA) were introduced into the BLMs. Diethoxyacetophenone (DEAP) was used to initiate the chemical cross-linking of the monomers by UV activation. The extent of cross-linking in the monomer doped BLMs was monitored by measuring the electrical and physical properties of the membranes before and after UV irradiation.
One embodiment of the subject disclosure features chemically modified aperture surfaces and chemical cross-linking within the lipid membrane to dramatically improve BLM stability. Glass microapertures were modified using PFDCS. The amphiphobic property (H2O/oil repellency) of the perfluorinated surfaces facilitated the rapid formation of highly stable BLMs. Perfluorinated patch pipettes showed decreased background capacitance and ionic conduction by 83% and 77% respectively, compared to unmodified pipettes. The reduced background current led to 48% noise reduction compared to conventional pipettes.
Another embodiment features photopolymerization of EGDMA and BMA partitioned into lipid membranes composed of 1, 2-diphytanoyl-sn-glycero-3-phosphocholine (DPhPC) to create a cross-linked polymer scaffold in the bilayer lamella for further improving BLM stability. The commercially available methacrylate monomers provide a simple, low cost, and broadly accessible approach for preparing highly stabilized BLMs used for ion channel analytical platforms. As a non-limiting example, when prepared on silane-modified glass microapertures, the resulting polymer scaffold-stabilized (PSS)-BLMs exhibited significantly improved lifetimes of 23±9 to 40±14 h and >10-fold increase in mechanical stability, with breakdown potentials >2000 mV attainable, depending on surface modification and polymer cross-link density. Additionally, the polymer scaffold exerted minimal perturbations to membrane electrical integrity as indicated by mean conductance measurements. When gramicidin A and α-hemolysin were reconstituted into PSS-BLMs, the ion channels retained function comparable to conventional BLMs.
Overall, cross-linked BLMs suspended on perfluorinated apertures exhibited significantly improved lifetimes (>24 h), 25-fold increase in mechanical stability, 50% increase in electrical resistance, and 53% increase in electrical stability. The present invention demonstrates key advances in the formation of stabilized BLMs and can be amenable to a wide range of receptor and ion channel functionalized platforms.
Section 2
The capability to rapidly screen complex libraries of pharmacological modulators is paramount to modern drug discovery efforts. This task is particularly challenging for agents that interact with lipid bilayers or membrane proteins due to the limited chemical, physical, and temporal stability of traditional lipid-based stationary phases in affinity chromatography. The present invention features the preparation of liquid chromatography (LC) columns using a novel stationary phase prepared from highly stable, polymeric phospholipid bilayers assembled onto silica micro-particles.
Polymer lipid membranes were prepared by photochemical or redox initiated polymerization of 1,2-bis[10-(2′,4′-hexadieoyloxy)decanoyl]-sn-glycero-2-phosphocholine (bis-SorbPC), a synthetic, polymerizable lipid. The resulting polymerized bis-SorbPC (poly(bis-SorbPC)) stationary phases exhibited enhanced stability compared to particles coated with non-polymerizable lipid 1,2-dioleoyl-sn-glycero-phosphocholine (DOPC) bilayers when exposed to chemical (organic solvents and surfactants) and physical (shear forces) insults over a 30 day period. Further, poly(bis-SorbPC)-coated particles could be packed into the column using slurry packing with no degradation of the lipid bilayer, compared to unpolymerized lipid membranes where the lipid bilayer was removed during packing. Frontal chromatographic analyses of acetylsalicylic acid, benzoic acid, and salicylic acid using poly(bis-SorbPC)-coated microspheres showed increased retention times compared to bare silica microspheres (P<0.0001), supporting that the lipophilic molecules were retained on the polymerized phospholipid bilayer stationary phases. The capability to prepare highly stabilized phospholipid membranes that withstand typical capillary LC conditions should greatly expand the range of applications utilizing lipid membrane-functionalized separations.
The present invention features a mixture of redox initiators, comprising an initiator-buffer component and NaHSO3, that is surprisingly is effective to polymerize lipid bilayer membranes, such as bis-SorbPC vesicles, at slightly acidic or near neutral pH solutions. Without wishing to limit the invention to any particular theory or mechanism, it is believed that one of the features that facilitates for this effectiveness is the use of ammonium persulfate (AP, (NH4)2S2O8) as the initiator-buffer component. Most importantly, the ammonium persulfate and NaHSO3 redox mixture shows the ability to generate radicals at near neutral pH, surprisingly converts 90+% of monomers to polymers in about an hour, and is mild enough to prevent denaturation of transmembrane proteins, such as Rhodopsin. For example, both Rhodopsin incorporated directly and in rod outer segment (ROS) membranes with bis-SorbPC show retention of activity after the polymerization of poly(bis-SorbPC) vesicles. In addition, the poly(bis-SorbPC) vesicles polymerized with the AP/NaHSO3 redox mixture show stability similar to or better than that of cross-linked polymerized lipid bilayers prepared with conventional redox mixtures such as K2S2O8/NaHSO3. Another advantageous property of the ammonium persulfate and NaHSO3 redox mixture is that it does not have undesirable spectroscopic properties that mask the structural information of the protein/lipid assembly.
As far as the inventors are aware, the prior arts and conventional redox mixtures do not teach or suggest a redox mixture for polymerizing lipid monomers and having an initiator-buffer component that functions as both an initiator and buffer, that is capable of generating radical species and maintaining near neutral pH conditions, and that does not interfere with redox chemistry and protein activity. For example, the prior art generally teaches a redox mixture comprising as K2S2O8/NaHSO3, which causes acidic conditions that can denature proteins.
The features and advantages of the present invention will become apparent from consideration of the following detailed description presented in connection with the accompanying drawings in which:
The following is a list of chemical abbreviations as used herein:
As used herein, the term “buffer” is defined as a chemical species that can resist pH change upon the addition of an acidic or basic component in a solution by neutralizing small amounts of the added component, thus maintaining a relatively stable pH of the solution.
As used herein, the term “initiator” is defined as a chemical species that can start a reaction. For example, in a polymerization reaction, an initiator can generate radical species that can subsequently react with monomers to form intermediate compounds capable of linking successively with other monomers to form a polymeric compound.
As known to one of ordinary skill in the art, redox polymerization is defined as the use of an oxidizing and a reducing agent for oxidation-reduction reactions to produce radicals that can be used to initiate polymerization.
As used herein, the term “lipid”, or alternatively, “lipid monomer”, and any of their derivatives, i.e. phospholipid, is defined as an organic molecule that is amphiphilic, or having a polar end and a non-polar end. The general structure of a lipid monomer is a polar head group attached to two hydrocarbon chains, or tails, which are usually fatty acids. The structural and amphiphilic properties of the lipids cause them to spontaneously assemble into supramolecular structures in aqueous solutions, of which a lipid bilayer is one example. As used herein, the term “lipid bilayer” or “lipid bilayer membrane” is defined as two thin molecular sheets, each sheet comprising lipid monomers. As used herein, the term “lipid leaflet” is defined as one of the molecular sheets, or monolayer, of the lipid bilayer. As used herein, the term “vesicle” is defined as a spherical lipid bilayer.
As used herein, there term “membrane protein” is defined as a protein that is inserted into the membrane. In one embodiment, a “transmembrane protein” is a protein that is embedded in the lipid membrane and spans from one side of the membrane through to the other side of the membrane. Alternatively, another embodiment of a membrane protein is a protein that contains a hydrophobic portion that inserts into, but does not span, the membrane.
Section 1
Referring now to
Another embodiment of the present invention features a suspended lipid system comprising a supporting substrate having a substrate surface and an aperture, and a modified lipid membrane. In one embodiment, an energy modifying layer is disposed on the substrate surface at or near the aperture to effect a lowering of a surface energy of the substrate surface. The modified lipid membrane may comprise a plurality of lipid monomers and a plurality of polymerized, hydrophobic non-lipid monomers comprising a methacrylate and a cross-linking agent. The modified lipid membrane may be disposed on the substrate such that the lipid monomers form a lipid bilayer suspended across the aperture and the energy modifying layer is disposed between the lipid monomers and the substrate surface. Preferably, the non-lipid monomers are disposed in or on the lipid bilayer and are polymerized at a near neutral pH by redox polymerization using a redox polymerization mixture comprising an initiator-buffer component and NaHSO3.
The present invention may further feature a method of enhancing stability of a suspended lipid membrane. The method may comprise providing a supporting substrate having a substrate surface and an aperture, depositing non-polymerizable lipid monomers on the supporting substrate such that the lipid monomers form a lipid bilayer suspended across the aperture and the energy modifying layer is disposed between the lipid monomers and the substrate surface, inserting polymerizable, hydrophobic non-lipid monomers in the lipid bilayer to form a modified lipid membrane, and polymerizing the non-lipid monomers to stabilize the modified lipid membrane. In some embodiments, the method may further comprise depositing an energy modifying layer on the substrate surface at or near the aperture prior to the depositing non-polymerizable lipid monomers on the supporting substrate in order lower a surface energy of the substrate surface. In some embodiments, the non-lipid monomers are polymerized by UV irradiation, visible irradiation, gamma irradiation, redox polymerization, or thermal polymerization may be used to polymerize the non-lipid monomers. In other embodiments, the method may further comprise inserting photoinitiators, such as DEAP, prior to polymerizing the non-lipid monomers by UV or visible irradiation. In still other embodiments, a redox polymerizing mixture is used to polymerize the non-lipid monomers by redox polymerization.
Another embodiment of the present invention features a method of enhancing stability of a suspended lipid membrane comprising depositing an energy modifying layer on a substrate surface at or near an aperture of a supporting substrate such that the energy modifying layer lowers a surface energy of the substrate surface, depositing non-polymerizable lipid monomers on the supporting substrate such that the lipid monomers form a lipid bilayer suspended across the aperture and the energy modifying layer is disposed between the lipid monomers and the substrate surface, inserting polymerizable, hydrophobic non-lipid monomers comprising a methacrylate and a cross-linking agent in the lipid bilayer to form a modified lipid membrane, and polymerizing the non-lipid monomers using a redox polymerization mixture to stabilize the modified lipid membrane. Preferably, the redox polymerization mixtures comprise an initiator-buffer component and NaHSO3 that allows for the redox polymerization to occur at a near neutral pH.
In some embodiments, the lipid bilayer can be suspended across the aperture such that a first lipid leaflet and a second lipid leaflet of the lipid bilayer are both disposed above or below the aperture. In other embodiments, the lipid bilayer can be suspended across the aperture such that a first lipid leaflet of the lipid bilayer is disposed above the aperture and a second lipid leaflet of the lipid bilayer is disposed below the aperture. In other embodiments, the non-lipid monomers are disposed in or on (i.e. deposited into or coated onto) a first lipid leaflet of the lipid bilayer, a second lipid leaflet of the lipid bilayer, or both. In some embodiments, the non-lipid monomers are disposed on the polar head groups of the lipids, the hydrocarbon tails of the lipids or both.
In some embodiments, the supporting substrate may be constructed from a material selected from a group consisting of a glass, a polymeric material, an epoxy, a metal oxide, or any other suitable material. Non-limiting examples of the supporting substrates include glass pipettes or solid sheets. The supporting substrate may be planar or curved.
One embodiment of the invention may feature a supporting substrate with an inherently low surface energy. For example, the surface of supporting substrate may have a surface energy of less than about 40 mJ/m2 or less than about 60 mJ/m2.
In other embodiments, an energy modifying layer may be disposed on the substrate surface at or near the aperture such that the energy modifying layer is disposed between the lipid monomers and the substrate surface. The energy modifying layer may be directly deposited on the substrate surface to form a coating. Preferably, the energy modifying layer can lower a surface energy of the substrate surface. In some embodiments, the energy modifying layer lowers the surface energy of the supporting substrate to less than about 40 mJ/m2. In other embodiments, the energy modifying layer lowers the surface energy of the supporting substrate to less than about 60 mJ/m2. In still further embodiments, the energy modifying layer lowers the surface energy of the supporting substrate to less than about 80 mJ/m2.
In some embodiments, the energy layer modifying is a silane-modified layer. For example, the silane-modified layer may comprise an alkylated silane. Non-limiting examples of alkylated silanes includes (tridecafluoro 1, 1, 2, 2-tetrahydrooctyl)dimethylchlorosilane (PFDCS), (heptadecafluoro 1, 1, 2, 2-tetrahydrodecyl)dimethylchlorosilane (PFDDCS), (trideca-fluoro 1, 1, 2, 2-tetrahydrooctyl)trichlorosilane (PFTCS), 3-cyanopropyldimethyl-chlorosilane (CPDCS), aminopropyldimethylethoxyosilane (APDES), 3,3,3-trifluoro-propyldimethylchlorosilane (FPDCS), ethyldimethylchlorosilane (EDCS), or n-octyl-dimethylchlorosilane (ODCS). In some embodiments, the silane-modified layer comprises any chemically related substance capable of lowering the surface energy of the substrate.
In some embodiments, the lipid monomers can be polymerizable lipids or non-plymerizable lipids. For example, the lipid monomers may be polymerizable bis-SorbPC (1,2-bis[10-(2′,4′-hexadieoyloxy)decanoyl]-sn-glycero-2-phosphocholine), non-polymerizable 1,2-diphytanoyl-sn-glycero-3-phosphocholine monomers, cell membrane fragments, naturally occurring lipids, or synthetic lipids.
In other embodiments, the plurality of polymerized, hydrophobic non-lipid monomers may comprise a methacrylate and a cross-linking agent. Exemplary methacrylates include aliphatic methacrylates and aromatic methacrylates. The aliphatic methacrylate may be alkyl substituted. For example, the alkyl substitution can be C4-C10. The aromatic methacrylate may be a benzyl methacrylate, a napthyl methacrylate, or any other aromatic methacrylate. In some embodiments, the cross-linking agent is a dimethacrylate, such as ethylene glycol dimethacrylate and derivatives thereof.
In some embodiments, the non-lipid monomers can be polymerized by UV irradiation, visible irradiation, gamma irradiation, redox polymerization, or thermal polymerization. When the non-lipid monomers are polymerized by UV or visible irradiation, the modified lipid membrane may further comprise photoinitiators. Preferably, the photoinitiators are lipophaic, such as DEAP. In some embodiments, the duration of UV irradiation or visible irradiation is sufficient to photopolymerize the non-lipid monomers. In preferred embodiments, the duration of irradiation is between about 5 to 10 minutes. In some embodiments, the duration of UV irradiation is between about 1 to 5 minutes. In other embodiments, the duration of UV irradiation is between about 5 to 15 minutes.
In some embodiments, the modified lipid membrane has an enhanced electrical stability. For example, a breakdown voltage of the modified lipid membrane is at least 1,000 mV. In other embodiments, a breakdown voltage of the modified lipid membrane is at least 1,250 mV. In still other embodiments, a breakdown voltage of the modified lipid membrane is at least 1,500 mV or at least 1,750 mV.
In other embodiments, the modified lipid membrane has an enhanced mechanical stability. For instance, the membrane is sufficiently flexible to support functional ion channels. In still other embodiments, the modified lipid membrane has an enhanced temporal stability. For example, the modified lipid membrane has a temporal stability of at least 8 hours, at least 10 hours, at least 15 hours, at least 24 hours, at least 36 hours, or at least 48 hours.
In some embodiments, the non-lipid monomers are polymerized by redox polymerization. A redox polymerization mixture comprising an initiator-buffer component and NaHSO3 may be used for redox polymerization. In some embodiments, the redox polymerization mixture comprises any redox mixture that polymerizes the non-lipid monomers, i.e. a mixture having a reductant and an oxidant. In preferred embodiments, the initiator-buffer component comprises ammonium persulfate. Suitable mole ratios of ammonium persulfate, NaHSO3, and lipids can include about 10-500 AP: 10-500 NaHSO3:1 lipid, about 50-400 AP:50-400 NaHSO3: 1 lipid, or about 100-300 AP:100-300 NaHSO3: 1 lipid.
Preferably, the redox polymerization mixtures allows for the reaction to occur at a near neutral pH and to maintain a constant pH. In some embodiments, the redox polymerization occurs at a pH between about 5 to 9. In other embodiments, the redox polymerization occurs at a pH between about 5.5 to 7.5. In still other embodiments, the redox polymerization occurs at a pH between about 6 to 7.
A kit for a preparing a suspended lipid membrane having enhanced stability is provided in the present invention. The kit may comprise a supporting substrate having a substrate surface and an aperture, a plurality of non-polymerizable lipid monomers, a plurality of polymerizable, hydrophobic non-lipid monomers comprising a methacrylate and a cross-linking agent, and a redox polymerization mixture comprising an initiator-buffer component and NaHSO3. To use the kit to prepare the suspended lipid membrane, the non-polymerizable lipid monomers are deposited on the supporting substrate such that the lipid monomers form a lipid bilayer suspended across the aperture, and the polymerizable, hydrophobic non-lipid monomers are disposed in the lipid bilayer. Preferably, the non-lipid monomers are polymerized by redox polymerization at a near neutral pH using the redox polymerization mixture to form a modified lipid membrane.
In some embodiments, the supporting substrate has an inherently low surface energy. In other embodiments the kit may further comprising an energy modifying compound that can be deposited on the substrate surface at or near the aperture to form an energy modifying layer. The energy modifying layer is disposed between the lipid monomers and the substrate surface in order to lowers a surface energy of the substrate surface and to increase stability of the modified lipid membrane. The energy modifying compound may comprise a silane, such as an alkylated silane. Exemplary alkylated silannes include (tridecafluoro 1, 1, 2, 2-tetrahydrooctyl)dimethylchlorosilane (PFDCS), (heptadecafluoro 1, 1, 2, 2-tetrahydrodecyl)dimethylchlorosilane (PFDDCS) (tridecafluoro 1, 1, 2, 2-tetrahydrooctyl)trichlorosilane (PFTCS), ethyldimethylchlorosilane (EDCS), aminopropyldimethylethoxyosilane (APDES), 3,3,3-trifluoropropyl-dimethylchlorosilane (FPDCS), ethyldimethylchlorosilane (EDCS), or n-octyl-dimethylchlorosilane (ODCS).
Reagents and Materials
Gramicidin A, ethylene glycol dimethacrylate (EGDMA), KCl, 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) and α-hemolysin (α-HL) were purchased from Sigma-Aldrich (St. Louis, Mo.). Gramicidin A was diluted to 10 μg/mL in ethanol. Tridecafluoro 1, 1, 2, 2-tetrahydrodimethylchlorosilane (PFDCS) was purchased from Gelest, Inc. (Morrisville, Pa.). 3-cyanopropyldimethylchlorosilane (CPDCS) was purchased from TCI America, Inc. (Portland, Oreg.). Anhydrous acetonitrile (ACN) and NaCl were purchased from EMD Chemical Inc. (Gibbstown, N.J.). Ethanol was purchased from Decon Laboratories (King of Prussia, Pa.). Butyl methacrylate (BMA) was purchased from Alfa Aesar (Ward Hill, Mass.) and diethoxyacetophenone (DEAP) was purchased from Acros Organics (Pittsburgh, Pa.). 1, 2-diphytanoyl-sn-glycero-3-phosphocholine (DPhPC) lipid in chloroform was purchased from Avanti Polar Lipids, Inc. (Alabaster, Ala.). Nanopure water was obtained from a Barnstead EasyPure UV/UF purifier with resistivity of 18.3 MΩ cm.
Pipette Aperture Fabrication and Surface Modification
Borosilicate capillaries (1.5 outer diameter and 1.1 mm inner diameter) were purchased from World Precision Instruments, (Sarasota, Fla.) and were fabricated into pipette apertures with 25-30 μm diameter using a P-97 micropipette puller (Sutter Instruments, Novato, Calif.) and fire polished with a model MF-900 microforge (Narishige, East Meadow, N.Y.) for BLM formation. A schematic of the pipette fabrication process is shown in
Formation and Characterization of BLMs
DPhPC dissolved in n-decane to a final concentration of 20 mg/mL was used to form BLMs by the painting method. Briefly, stock lipids suspended in chloroform were dried using compressed Ar followed by overnight vacuum. Conventional (unpolymerized) BLMs were prepared with this solution. Methacrylate-doped BLMs (MA-BLMs) describe BLMs prepared using a mixture of lipids, methacrylate monomers and photoinitiators in the absence of UV photopolymerization. PSS-BLMs describe MA-BLMs that were subsequently photopolymerized. For MA-BLMs and PSS-BLMs, DPhPC solutions were prepared with BMA, EGDMA and DEAP as follows. Initially, radical inhibitors were removed from BMA and EGDMA using an alumina column (Al2O3, 50-200 μm, 60 Å, Acros). The monomers were then combined in 1:1:1 ratios with DEAP to yield a solution referred to as a monomer mixture, followed by addition of one equivalent of lyophilized DPhPC to yield and overall mixture of 1:1:1:1 composition of BMA:EGDMA:DEAP:DPhPC. The lipid/monomer mixture was vortexed for 30 s prior to the addition of n-decane.
BLMs were formed by addition of 2 μL of lipid or lipid/monomer mixture solution dissolved in n-decane to the pipette tip and dried with N2 gas. Pipettes were back filled with recording buffer (1 M KCl, 5 mM HEPES, pH 7.4) and mounted on the head stage of a patch clamp amplifier (EPC-10, HEKA Electronics, Bellmore, N.Y.). The bath chamber was filled with recording buffer and connected to the reference electrode via a salt bridge. The lipid or methacrylate-doped lipid solution was painted by gently sweeping a plastic micropipette tip across the silanized pipette aperture submerged in the recording buffer.
Formation of BLMs or MA-BLMs across silanized pipette apertures was monitored by the spontaneous increase in electrical resistance from open pipette resistance (50-100 KΩ) to >2 GΩ. Further, the formation of BLM or MA-BLMs was verified by applying an increasing potential from 0 to 2000 mV in 10 mV increments of 50 ms duration. Additionally, the appearance of transient pores in BLMs under applied electrical fields was used to indicate the existence of BLMs prior to UV irradiation, although care was taken not to allow complete rupture of the BLM upon observation of transient pores. Subsequently, MA-BLMs were polymerized, forming PSS-BLMs, by UV irradiation using a pen lamp (UVP, Upland, Calif., Model 90-0012-01) at a distance of 3-5 cm from the BLM.
The biophysical properties of conventional, MA- and PSS-BLMs were characterized by the reconstitution and measurement of gramicidin A or α-HL activity. A 0.5 μL aliquot of stock gramicidin peptide (10 μg/mL) in ethanol was added to 500 μL bath solution to a final concentration of 10 ng/mL and allowed to incubate with the BLMs. The activity of gramicidin was monitored with a potential of 70 mV applied across the BLMs. Quantized changes in current were typically observed within 2 min of adding gramicidin to the bath solution. Two μl of α-HL (0.5 mg/mL in recording buffer) was added to bath solution containing 500 μl of recording buffer and the insertion of IC measured a bias potential of +40 mV across the BLM.
Conductance Measurement
The conductance of conventional BLMs, MA-BLMs and PSS-BLMs was measured by applying a square wave of increasing potential from −100 to +100 mV in 10 mV increments of 50 ms duration. The potential was held for 10 ms at 0 mV before and after applying each pulse. The average of the steady state current between 30 to 50 ms (following capacitive decay) was plotted versus the applied potential. The conductance is reported as the slope of the current versus potential plot and normalized for pipette aperture area, as a first approximation of BLM area. A minimum of three pipettes each was used for evaluating BLMs suspended across CPDCS or PFDCS-modified pipette apertures. For each pipette, a minimum of three BLMs was analyzed to determine the mean conductance.
Assessment of BLM Stability
The stability of conventional, MA and PSS-BLMs was quantified by measuring breakdown voltage (VB), longevity and air-water transfer count (AWT). VB is the potential at which the BLM undergoes irreversible rupture, and is measured by applying an increasing potential from 0 to 2000 mV in 10 mV increments of 50 ms duration and observing the potential at which a large, non-linear increase in current occurs. The mean VB indicates the electrical stability of BLMs. AWT refers to the number of times a BLM survives transport across the air-water interface before it ruptures. To assess AWT, the aperture was removed from aqueous buffer and maintained in air for 1 s, prior to re-submersion in buffer. Each cycle of removal and return to buffer is indicated as one AWT. Longevity was measured as the average time required for the bilayer to undergo rupture under the application of a ±5 mV 20 Hz square-wave.
Statistical Analysis
All data is presented as mean±standard deviation. For each measurement, a minimum of three BLM replicates on at least three different pipettes were collected. For each BLM stability metric analyzed, outlying data was assessed using the Q test at the 90% confidence level. All statistical comparisons were performed using Student's t-test at the 95% confidence interval.
Single-channel recordings with corresponding all-points histograms and mean open times were analyzed using TAC (X4.3.3) and TACfit X4.3.3 (Bruxton). The fit duration histogram for open probability and construction uses the Sigworth and Sine transformations.
Redox Polymerization of Bis-DenPC
The following is a non-limiting example of redox polymerization of bis-DenPC (10 mg/mL)
Bis-DenPC stored in benzene was dried over argon gas and lyophilized overnight. The resultant solid was suspended in n-decane to a final concentration of 10 mg/mL. About 0.5 μL of this solution was then painted across a glass pipette aperture (ca. 30 μm diameter) that had been silanized with a perfluorinated silane, forming a black lipid membrane (BLM).
A redox initiation solution containing NaHSO3 and (NH4)2S2O8 was prepared in buffer solution containing 1M KCL and 5 mM HEPES (pH 7.4). Redox initiation solutions were prepared separately and mixed before perfusing into the bath solution into which the BLM was immersed. BLM polymerization was performed using a mole ratio of lipid (bis-DenPC):NaHSO3:(NH4)252O8=1:300:300)
Redox Polymerization of DPhPC BLMs Doped with Methacrylate Monomers
The following is a non-limiting example of redox polymerization of DPhPC BLMs doped with methacrylate monomers.
DPhPC dissolved in chloroform was dried over argon gas and lyophilized overnight followed by suspension in n-decane to a final concentration of 20 mg/mL.
Doping Polymerizable Monomers into BLMs
Butyl methacrylate (BMA), ethylene glycol dimethacrylate (EGDMA) and the initiator diethoxy acetophenone (DEAP) in a 1:1:1 ratio were doped into the DPhPC BLM. All experiments were performed under yellow light (low pressure sodium lamp). As shown in
Table 1 provides exemplary mole and volume amounts of DPhPC lipid and monomers used for preparation of BLMs containing poly(methacrylate) networks.
Redox Polymerization of Methacrylate Doped BLMs
Using newly fabricated and silanized glass pipettes, methacrylate doped BLMs were formed via a tip-dip or a painting method. The fabricated pipette was filled with buffer solution (1 M KCl and 5 mM HEPES, pH 7.4) and mounted onto the head stage of a patch clamp amplifier.
After formation of methacrylate doped BLMs, redox reagents containing equal moles of NaHSO3:(NH4)2S2O8 were prepared separately. An exemplary mole ratio of lipid (DPhPC):NaHSO3:(NH4)2S2O8 is 1:300:300. To perform redox polymerization, 500 μL each of NaHSO3 and (NH4)2S2O8 were mixed together to give a total volume of 1000 μL, which was sufficient to displace the recording buffer in the bath reaction chamber. The methacrylate doped BLM was allowed to react for 5 to 10 min. After reaction, the bath solution was exchanged 5× with 1M KCl and 5 mM HEPES, pH 7.4 buffer.
Table 2 shows a non-limiting example of concentrations of lipid and redox reagents that may be used.
Table 3 shows an exemplary comparison of breakdown voltages for a DPhPC control and a redox polymerized monomer mixture.
Insertion of Ion Channel
About 0.5 μL of Gramicidin A (0.5 mg/mL) was added close to the tip of pipette containing methacrylate doped BLMs to allow for insertion. Insertion was observed after a few minutes from the addition of gramicidin. Single channel activities with multiple states were observed after application of a holding potential of 70 mV. Next, the bath solution was exchange with redox buffer and allowed to react for 10 min, after which the conductance drastically decreased due to extended cross-linking in the methacrylate monomers, as shown in
Results and Discussion
The lifetime, electrical and physical properties of monomer doped BLMs were investigated to probe the effect of various degrees of cross-linking in BLMs on the properties of IC proteins.
Decreased substrate surface energy enhances BLM stability by improving the force of interaction between the lipid membrane and the substrate (Fms)—However, the weak van der Waals forces (FV) and electrostatic forces (FE) of interactions between adjacent lipid molecules limit the temporal, electrical, and mechanical stability of BLMs. For example, when BLMs are suspended across silane-functionalized glass apertures with decreased surface energy, the inherently weak forces of interaction between the lipid molecules in the self-assembled bilayer (
Table 4 shows physical and electrical properties of conventional BLMs, MA-BLMs, and PSS-BLMs on CPDCS-modified pipette apertures.
Table 5 shows physical and electrical properties of conventional BLMs, MA-BLMs, and PSS-BLMs on PFDCS-modified pipette apertures.
To further improve the stability of BLMs for applications requiring high mechanical and temporal stability, the integration of a polymer network into the lamella region of the BLM was evaluated. A methacrylate polymer scaffold prepared from BMA and EGDMA was utilized. The BMA forms linear polymer chains that are cross-linked by the EGDMA to improve the polymer stability. Without wishing to limit the present invention to a particular theory or mechanism, while the resulting polymer does not covalently link the DPhPC monomers that form the BLM, it is hypothesized that the enhanced structural stability provided by the polymer network would enhance BLM lifetime and mechanical stability. The resulting BLM architecture is referred to herein as a polymer scaffold-stabilized BLM (PSS-BLM).
Physical and Electrical Properties of PSS-BLMs
To evaluate the physical and electrical properties of PSS-BLMs, the air-water transfer (AWT), VB, and longevity were measured as metrics of the physical, electrical, and temporal stability, respectively. A higher value of AWT and VB indicates enhanced mechanical and electrical stability of the specific BLM composition.
The successful formation of a BLM was indicated when VB was observed in the range of 0-1000 mV, prior to polymerization, as opposed to a high resistance blockage in the pipet which cannot be broken down in this range. The mean VB observed for conventional BLMs suspended across CPDCS-modified pipette apertures was 460±21 mV. Furthermore, electrical, physical, and temporal stability were statistically similar before and after 15 min of UV irradiation of conventional BLMs, indicating no deleterious effects of UV exposure. When the monomer mixture was incorporated into the BLM in the absence of UV-irradiation to form MA-BLMs (Table 4), similar longevity and VB were observed compared to conventional BLMs. Though inclusion of the monomer mixture increases membrane conductance, the magnitude of the change is within the normal working range of BLMs on a range of aperture substrate materials. Upon cross-linking of MA-BLMs via UV irradiation for 5 min to yield PSS-BLMs, a >10 fold increase in AWT and longevity and a 30% increase in VB were observed compared to conventional BLMs. Additional improvements in electrical stability were observed upon increasing UV irradiation time to 10 min. Importantly, the membrane conductance, a key measure of membrane integrity, was statistically similar in MA-BLMs and PSS-BLMs irrespective of UV irradiation time.
Stability metrics for conventional BLMs on PFDCS-modified apertures were statistically similar before and after 15 min of UV irradiation (Table 5). Conventional BLMs formed on PFDCS-modified apertures exhibit marked stability increases compared to those formed on CPDCS apertures due to enhanced surface/lipid interactions. Unlike on CPDCS-modified apertures, MA-BLMs formed on PFDCS-modified apertures exhibited decreased electrical and mechanical stability as indicated by VB and AWT (47% and 30% decreases, respectively) compared to conventional BLMs, though the magnitudes of these values are still comparable to BLMs formed on CPDCS-modified apertures. Thus, it is likely that the inclusion of the monomer mixture disrupts the lipid-surface interactions and yields BLM stabilities comparable to those formed on surfaces with weaker surface-lipid interactions.
Upon formation of PSS-BLMs via 5 min of UV irradiation, VB recovered to values equivalent to conventional BLMs; however, >3 fold increase in longevity was observed compared to conventional BLMs. Furthermore, a 45% reduction in membrane conductance was observed for MA-BLMs, and persisted in PSS-BLMs formed via 5 minutes of UV irradiation. When UV irradiated for 10 minutes, an increase in membrane conductance was observed, though the values were still lower than conventional BLMs prepared with this surface modification. Furthermore, VB and longevity were increased by >2- and 5-fold, respectively, compared to conventional BLMs.
Based on the aggregate of the measurements, it appears that formation of the polymer scaffold exhibits no deleterious effects on membrane stability or membrane integrity within the BLM. In fact, PSS-BLMs formed via 5 min or 10 min of UV irradiation of MA-BLMs on CPDCS- and PFDCS-modified apertures surprisingly yielded significantly improved membrane longevity and reduced membrane conductance with enhanced VB and little or no adverse effect on AWT. For membranes formed on both CPDCS and PFDCS-modified apertures, membrane conductance was unchanged when comparing MA-BLMs and PSS-BLMs formed via 5 minutes of UV irradiation. Though the conductance was increased with further irradiation time, the conductance was still lower than that obtained from a conventional BLM.
Electrical Properties of Monomer Doped BLMs
To evaluate the physical and electrical properties of monomer doped BLMs and the extent of cross-linking of monomers introduced into BLMs, the AWTs, VB and longevity were measured as a metric of the mechanical, electrical and temporal stability respectively.
In addition to using pipettes that gave >90% success rate in the formation of BLMs, the presence of BLMs prior to UV irradiation was confirmed by the appearance of the first transient pores under induced electric field. Further, the transient pores were used to indicate an existing BLM with thickness similar to cell membranes. Subsequently, increasing pore size or number preceded the rupture of BLMs, thus the complete rupture of a BLM was prevented by quickly removing the applied electric field before irradiation of monomer doped BLMs. The transient pores were observed to occur between 500-700 mV for BLMs and 400-500 mV for monomer doped BLMs suspended across perfluorinated apertures. Numerous transient pores were often observed in monomer doped BLMs under applied electric field as compared to BLMs without monomers.
Irreversible breakdown of BLMs suspended across conventional pipettes was observed at 418±18 mV. However, BLMs suspended on low energy perflourinated pipette surfaces surprisingly demonstrated reversible pore formation under a progressively increasing electric field ranging from 400-600 mV and much higher irreversible breakdown potentials. Without wishing to limit the present invention to a particular theory or mechanism, the high energy barrier within the pores and the improved interaction between the hydrophobic lipid tails and the aperture substrate may prevent the irreversible rupture of BLMs within the observed potential range (400-600 mV). In other cases, high viscosity in the lipid membrane may inhibit fast growth of the pores, thereby preventing irreversible breakdown of BLMs. At high electric fields (>900 mV), an irreversible rupture of BLMs often occurred.
When BLMs were doped with monomers, the accumulation of transient pores that preceded the irreversible rupture of the membranes suspended across CPDCS and PFDCS was observed at 493±59 and 520±119 mV respectively.
Table 6 shows physical and electrical properties of monomer doped BLMs suspended on low energy (PFDCS) silane-modified pipette apertures.
Table 7 shows physical and electrical properties of monomer doped BLMs suspended across (CPDCS) silane-modified pipette apertures.
The electrical stability of conventional BLMs suspended across PFDCS modified glass pipette apertures before and after 15 min of UV irradiation (984±210 vs 980±254 mV respectively) were statistically the same, suggesting that UV irradiation had no chemical effect on non-polymerizable lipids. Additionally, the conductance, the number of AWTs and longevity were statistically the same before and after 15 minutes of UV irradiation for BLMs without monomers, confirming that UV irradiation had no effect on the structural organization of BLMs. When BLMs were doped with hydrophobic monomers, the electrical stability decreased by ca. 50%, which is possibly due to the interference of free monomers on the packing density of self-assembled lipid molecules.
Interestingly, the conductance decreased by ca. 50%. for monomer doped BLMs, suggesting that the necessary arrangement of the small hydrophobic monomers in the lamella region of the bilayer without the formation of domains thereby decreases the permeability of the membranes to conducting ions. In addition, the mechanical stability or the number of AWTs slightly decreased from >50 for BLMs without monomers to 32±11 upon introduction of monomers into the bilayer. In general, the observed decrease in the electrical and mechanical stability of monomer doped BLMs, and the decrease in temporal stability from 8±1 to 6±1, suggest that further weakening of the hydrophobic interaction between lipid molecules leads to a gradual collapse of the bilayer scaffold with time.
After 5 minutes of UV irradiation of the monomer doped BLMs, the electrical stability was improved by a factor of 3 from 520±119 to 1398±552 mV, while the mechanical stability was statistically the same before and after UV irradiation (32±11 to 36±13 respectively). The large standard deviation reported on the electrical stability of monomer doped BLMs after UV irradiation may be due to the variation in the amount of hydrophobic monomers that undergo cross-linking in the lipid bilayer in each individual experiment. There was no significant change in the conductance before and after 5 minutes of UV irradiation due to insufficient cross-linking in the BLM scaffold. Although the change in the conductance and mechanical stability of monomer doped BLMs before and after UV irradiation was indistinguishable, the observed increase in the electrical stability indicated cross-linking in BLMs through the hydrophobic monomers. Improvement in the temporal stability from 4 to >8 h also confirms cross-linking in BLMs. Further UV irradiation for 10 minutes showed a significant change in the electrical stability as observed by >4 fold increase in the breakdown voltage from 520±119 to >2000 mV before and after cross-linking. The mechanical stability of monomer doped BLMs suspended across PFDCS-modified apertures was improved by a factor of 2 after 10 minutes of UV irradiation.
Although traditional BLMs suspended across perfluorinated glass pipette apertures demonstrated enhanced electrical and mechanical stability, the introduction of small hydrophobic polymerizable monomers in BLMs further improved the electrical stability by overcoming the weak force of interaction within the lipid moieties via chemical cross-linking. The conductance of monomer doped BLMs before and after 10 min of UV irradiation increased from 4.81±0.14 to 6.93±1.08, suggesting a slight leakage in the bilayer, which is probably due to the formation of domains. Previous results demonstrated that polymeric lipid membranes that become leaky after cross-linking is cause by the lateral contraction of the lipids which leads to the formation of holes. During chemical cross-linking via UV irradiation, the bilayer undergoes a structural rearrangement. In the case where cross-linking occurs at only one end of an EGDMA monomer, instead of both ends, the lipid head groups are likely to be drawn together at the surface of the bilayer. Further, UV irradiation may trigger the reorganization of the molecules to result in either improved packing or domain formation.
Reconstitution of Protein Ion Channel into Monomer Doped BLMs
To validate the formation of lipid membranes with a bilayer thickness from a mixture of polymerizable monomers and non-polymerizable lipids, a protein was reconstituted and single-channel recording was monitored. Ion channels in cell membranes or model artificial lipid membranes serve as transducers for label-free chemical measurements of molecules and ions. Additionally, the reconstitution of protein ion channels provide an indication of successful formation of BLMs with thickness similar to cell membranes and the activity of the IC can be used to probe the lipid environment.
Robust ion channel-based biosensors and sequencing platforms necessitate high stability suspended lipid bilayers into which functional ion channels can be reconstituted. While PSS-BLMs showed significant stability improvements, the effects of UV irradiation and the presence of the polymer scaffold on the ion channel function was a major concern for the application of this technology. Gramicidin A, a channel forming peptide that requires membrane fluidity to function, was reconstituted into conventional BLMs, MA-BLMs, and PSS-BLMs to probe the relationship between methacrylate cross-linking and ion channel function.
The structure and function of Gramicidin A is one of the most well studied and described cation-selective channels. Gramicidin A forms an ion-conducting channel by the dimerization of the pentadeca peptide subunits which freely diffuse in each monolayer leaflet of a lipid bilayer. The dynamic process of formation and dissociation of transmembrane dimers of gramicidin A leads to a quantized change in current with conductance ranging from 21-24 pS. Owing to a shorter hydrophobic length of gramicidin (ca. 2.2 nm) than the hydrophobic thickness of BLM, the bilayer is locally deformed to allow dimerization of two gramicidin monomers to form pores (
Kelkar et al. reported the structure and function of gramicidin in a lipid bilayer to be dependent upon the oriented dipole moments of the four C-terminal tryptophan residues of the peptide (Kelkar, D. A.; Chattopadhyay, A. Biochim. Biophys. Acta 2007, 1768, 2011-2025). Thus changes in membrane properties upon PSS-BLM formation or degradation of C-terminal tryptophan residues during UV irradiation may lead to disruption of gramicidin A function.
To monitor IC insertion and activity, Gramicidin A was added to the cis side of a conventional BLM while applying a potential of 70 mV across the bilayer. Successful insertion and dimerization was indicated by quantized changes in ion current with amplitudes of ca. 1.5 pA. (
The formation of BLMs with thickness similar to cell membranes was crucial for monitoring the transport of ions across reconstituted ion channels. In the case where the thickness of BLMs exceeded that of cell membranes, measurement of IC current across the bilayer was impaired. Mueller and Montal reported the adverse effect of trapped hydrocarbon solvent within a bilayer leaflet on the insertion and activity of IC. Rovin and co-workers also demonstrated the dependence of membrane composition on the lifetime of gramicidin ion channels (Rudnev, V. S.; Ermishkin, L. N.; Fonina, L. A.; Rovin, Y. G. Biochim. Biophys. Acta 1981, 642, 196-202). Since the composition of the lipid bilayer greatly influenced the insertion, average lifetime, and gating of an ion channel, the effect of free monomers that partitioned in and out of the lipid bilayer and organic solvent within the annulus region or outside of bilayer was a major concern.
To evaluate the effect of monomer doped BLMs on the activity of ICs, gramicidin was reconstituted in DPhPC BLMs without monomers, and the IC activity was monitored before and after UV irradiation. Results in
Interestingly, channel activity was observed when gramicidin was reconstituted in equimolar monomer/lipid BLMs. The IC conductance in monomer doped BLMs that were not UV irradiated was reduced by 30% compared to gramicidin in DPhPC BLMs (
Table 8 shows conductance of IC under variable lipid bilayer environment.
Owing to the fact that the conductance or current amplitude of gramicidin A in BLMs was not preserved after UV irradiation, the monomer doped BLMs were UV irradiated prior to IC insertion. The monomer doped BLMs were partially polymerized for 5 minutes before IC insertion. Gramicidin activity was successfully maintained in the partially polymerized monomer doped BLMs as shown in
Table 9 shows gramicidin A activity in conventional BLMs, MA-BLMs, and PSS-BLMs.
21 ± 0.4
21 ± 0.6
Upon 5 minutes of UV irradiation of MA-BLMs to form PSS-BLMs (
In an effort to circumvent the deleterious effects of UV irradiation and/or monomer mixture on gramicidin conductance, gramicidin was reconstituted into pre-formed PSS-BLMs (
When PSS-BLMs were formed via 10 min of UV irradiation (
The observed number of functional ion channels provides an additional indication of the degree of fluidity and/or compressibility in PSS-BLMs. When gramicidin activity was observed in PSS-BLMs, no more than two active gramicidin channels were observed concurrently, compared to >6 in both conventional and MA-BLMs. Thus the probability of forming a functional dimer is decreased upon increased photopolymerization.
Table 10 shows gramicidin A activity when reconstituted in pre-formed PSS-BLMs.
Importantly, gramicidin reconstituted into PSS-BLMs maintained function for 7-9 h before permanent loss of peptide function was observed, possibly due to peptide denaturation, as shown in
To broaden the application of PSS-BLM ion channel platforms, α-HL was reconstituted, a pore forming channel with characteristic conductance of ca. 1 nS into the various BLM configurations. α-HL is routinely used to prepare stochastic sensors and nucleic acid sequencing platforms, thus it represents an important application for stabilized BLMs. Table 11 summarizes the results obtained for α-HL reconstituted into differing BLM configurations. In each case, the mean conductance values for α-HL were within the accepted experimentally measured values as shown in
α-HL is a homoheptamer that requires insertion and assembly of the seven monomer units to form the functional channel. Thus, the membrane must retain sufficient fluidity to support diffusion and assembly of the channel subunits. Reconstitution of α-HL into pre-formed PSS-BLMs via 5 min of UV irradiation further confirms the existence of sufficient fluidity required for ion channel reconstitution and function, similar to that observed for gramicidin A, whereas no evidence of functional ion channel assembly was observed after extended cross-linking via 10 min of UV irradiation (Table 12). Overall, reconstitution of ion channels into pre-formed PSS-BLMs prepared via 5 min of photopolymerization show great promise for the construction of ion channel functionalized sensor technologies.
Importantly, UV-photopolymerization might also be useful to limit the number of α-HL insertions into the BLM. In typical α-HL reconstitutions, an excess of ion channel is added to the bath and immediately upon insertion of a functional channel, the bath is diluted. Thus polymerization of the PSS-BLM may provide an easily automated alternative approach for controlling ion channel insertion density, if the excessive electrical noise introduced by the UV lamp can be overcome.
Table 11 shows α-HL activity in conventional BLMs, MA-BLMs, and PSS-BLMs on PFDCS-modified pipette apertures.
Table 12 shows α-HL activity reconstituted in pre-formed PSS-BLMs on PFDCS-modified pipette apertures.
Finally, it should be noted that while direct insertion of ion channels used here was readily achieved, insertion of more hydrophobic channels typically requires either surfactant dialysis or fusion of proteolysosomes.
Conclusion
The stability of lipid bilayers suspended across glass pipette apertures is dependent on the underlying surface energy of the aperture and the interactive force existing between the lipid molecules. The use of low surface energy (PFDCS) modifiers enhanced the mechanical, electrical and temporal stability of BLMs compared to conventional pipettes. The lifetime of BLMs was improved from 8 to >15 h by overcoming the weak noncovalent interactions among lipid molecules via chemical cross-linking using small hydrophobic non-lipid monomers. Monomers doped into BLMs without cross-linking decreased the stability of BLMs by 50% due to the interference of free monomers on the self-assembled lipid interactions. However, UV irradiation to convert the monomers to a polymeric network further improved the electrical stability of BLMs by a factor of 3 and >4 for 5 and 10 minute irradiations respectively.
In addition, UV irradiation of BLMs after the reconstitution of ICs resulted in the photolytic degradation of IC indicated by a decrease in current amplitude. Partial cross-linking in monomer doped BLMs for 5 minutes allowed for the insertion and free diffusion of IC without any deleterious effect on the conductance, which is due to preserved fluidity in BLMs. UV irradiation for 10 minutes resulted in the loss of fluidity in BLMs, which was confirmed by the inability to reconstitute ICs. Thus, partial polymerization was necessary to maintain fluidity and stability of BLMs, while allowing for the reconstitution of ICs. The improved IC scaffold by partial cross-linking using commercially available monomers is promising for the development of the next generation of robust, BLM-based, ion channel sensors.
Furthermore, improved stability of BLMs can be attained by chemically cross-linking methacrylate monomers within the lipid membranes to form PSS-BLMs. This approach is simpler, broadly applicable, less costly and more widely accessible compared to prior efforts utilizing reactive lipid monomers. PSS-BLMs can withstand potentials >2000 mV without experiencing dielectric breakdown and show >10 fold increase in measures of mechanical stability and >5 fold increase in BLM lifetime compared to conventional BLMs, with no deleterious effect on membrane integrity or structure. The average membrane lifetime was improved such that the lifetime of the reconstituted ion channel, gramicidin A, and not the BLM lifetime, was the fundamental limitation on sensor lifetime. PSS-BLMs offer stability advantages similar to those obtainable with polymerizable lipids but with few of the associated limitations. Thus, PSS-BLMs can offer substantial advantages for ion channel-based sensors and other BLM technologies, and may address the limitations of membrane stability on the development of these technologies.
Section 2A
Referring now to
An embodiment of the present invention may feature a stabilized, supported lipid bilayer system for use in a chromatography column comprising the chromatography column, a supporting substrate, and a stable, polymeric lipid bilayer comprising a plurality of polymerized lipid monomers and disposed on the supporting substrate. Preferably, the lipid monomers are polymerized at a near neutral pH using a redox polymerization mixture comprising NaHSO3 and an initiator-buffer component. The initiator-buffer component functions to initiate redox polymerization and to buffer the redox polymerization mixture.
Another embodiment of the present invention may feature a method of producing a stabilized, supported lipid bilayer system. For example, the method may comprise obtaining a plurality of polymerizable lipid monomers, obtaining a supporting substrate, depositing the lipid monomers onto the supporting substrate such that the lipid monomers form a lipid bilayer on the supporting substrate, and polymerizing the lipid monomers to increase a stability of the lipid bilayer. Teh method may further comprise integrating membrane proteins into the lipid bilayer prior to polymerizing the lipid monomers.
In an alternative embodiment, the method may comprise obtaining a plurality of polymerizable lipid monomers, obtaining a supporting substrate, depositing the lipid monomers onto the supporting substrate to form a lipid bilayer on the supporting substrate, and polymerizing the lipid monomers to increase the stability of the lipid bilayer using a redox polymerization mixture comprising an initiator-buffer component and NaHSO3. The initiator-buffer component can initiate polymerization of the lipid monomers and buffer the redox polymerization mixture to maintain a near neutral pH during polymerization of the lipid monomers. In other embodiments, the method may further comprise integrating membrane proteins into the lipid bilayer prior to polymerizing the lipid monomers. Preferably, the activity and native conformations of the proteins are maintained after polymerization of the lipid monomers.
Another embodiment of the present invention features a chromatography column packed with microparticles having a polymeric lipid bilayer disposed on a surface of each microparticle. The lipid bilayer may comprise polymerized lipid monomers.
In an alternative embodiment, the present invention features a chromatography column comprising a polymeric lipid bilayer deposited on an inner surface of the chromatography column. The chromatography column may be an open-tubular capillary column. The lipid bilayer may comprise polymerized lipid monomers disposed on (i.e. covering) at least about 50% of the inner surface of the chromatography column.
The present invention may also feature an embodiment comprising a method of preparing a packed chromatography column. The method may comprise providing a chromatography column, preparing a plurality of vesicle-coated microparticles, and packing the vesicle-coated microparticles inside the chromatography column. In an exemplary embodiment, preparing said vesicle-coated microparticles may comprise obtaining a plurality of polymerizable lipid monomers, obtaining a plurality of microparticles, depositing the lipid monomers onto each microparticle such that the lipid monomers form a lipid bilayer on a surface of each microparticle, and polymerizing the lipid monomers of each lipid bilayer using a redox polymerization mixture to form a stable, polymeric vesicle coating on each microparticle. Preferably, the redox polymerization mixture may comprise an initiator-buffer component and NaHSO3. The initiator-buffer component initiates polymerization of the lipid monomers and buffers the redox polymerization mixture to maintain a near neutral pH during polymerization of the lipid monomers. In other embodiments, the method may further comprise integrating membrane proteins into the lipid bilayer prior to polymerizing the lipid monomers. The activity and native conformations of the proteins are maintained after polymerization of the lipid monomers.
The present invention may further feature a kit for preparing a stabilized, supported lipid bilayer system. The kit may comprise at least one supporting substrate, a plurality of polymerizable lipid monomers, and a redox polymerization mixture comprising an initiator-buffer component and NaHSO3. To prepare the lipid bilayer system, the lipid monomers are deposited onto the supporting substrate such that the lipid monomers form a lipid bilayer on the supporting substrate and the lipid monomers are polymerized using the redox polymerization mixture to increase stability of the lipid bilayer. Preferably, the initiator-buffer component initiates polymerization of the lipid monomers and buffers the redox polymerization mixture to maintain a near neutral pH during polymerization of the lipid monomers. In some embodiments, the kit may further comprise an affinity platform. The affinity platform is a chromatography column, such as an open-tubular capillary chromatography column or a packed-bed chromatography column. In one embodiment, the supporting substrate is an inner wall of the open-tubular capillary column. In another embodiment, the supporting substrate comprises a plurality of microparticles.
In one embodiment, when the chromatography column is an open-tubular capillary chromatography column, the supporting substrate may be an inner wall of the open-tubular capillary column. In another embodiment, when the chromatography column is a packed-bed chromatography column, the supporting substrate may comprise a plurality of microparticles. In some embodiments, the supporting substrate has a planar or curved surface. In other embodiments, the lipid bilayer may be disposed on a surface of each microparticle to form a vesicle-coated microparticle.
In some embodiments, the lipid bilayer covers at least about 50% of the inner surface of the chromatography column. In some embodiments, the lipid bilayer covers at least about 60% of the inner surface of the chromatography column. In some embodiments, the lipid bilayer covers at least about 70% of the inner surface of the chromatography column. In some embodiments, the lipid bilayer covers at least about 80% of the inner surface of the chromatography column. In some embodiments, the lipid bilayer covers at least about 90% of the inner surface of the chromatography column.
In some embodiments, the chromatography column is an open-tubular capillary chromatography column or a packed-bed chromatography column. In some embodiments, the chromatography column can be any dimension.
In some embodiments, the supporting substrate is a micro-particle. In some embodiments, the supporting substrate is an inner wall of the open-tubular capillary column. In some embodiments, the supporting substrate is planar or curved. In some embodiments, the supporting substrate is a vesicle.
In some embodiments, the lipid bilayer covers at least 50% of the surface of each micro-particle. In other embodiments, the lipid bilayer covers at least 60% of the surface of each micro-particle. In still further embodiments, the lipid bilayer covers at least 70% of the surface of each micro-particle. In some embodiments, the lipid bilayer covers at least 80% of the surface of each micro-particle. For example, the lipid bilayer covers at least 90% of the surface of each micro-particle. In some embodiments, the micro-particles are silica micro-particles or metal-oxide particles.
In some embodiments, a diameter of the micro-particles is between about 0.5 μm to 40 μm. In some embodiments, a diameter of the micro-particles is between about 0.5 μm to 30 μm. In some embodiments, a diameter of the micro-particles is between about 10 μm to 30 μm. In some embodiments, a diameter of the micro-particles is between about 20 μm to 30 μm. In some embodiments, a diameter of the micro-particles is between about 30 μm to 40 μm.
In some embodiments, the lipid monomers are sorbyl- or dienoyl-containing lipid molecules, mono-functionalized lipid molecules or lipid molecules containing one or more polymerizable dienoyl groups. In some embodiments, the dienoyl-containing lipid molecule is 1,2-bis[10-(2′,4′-hexadieoyloxy)decanoyl]-sn-glycero-2-phosphocholine (bis-SorbPC) or 1,2-bis(octadeca-2,4-dienoyl)-sn-glycero-3-phosphocholine (bis-DenPC). In other embodiments, the mono-functionalized lipid molecule is mono-dienoylphosphatidylcholine (mono-DenPC) or mono-sorbylphophostidylcholine (mono-SorbPC).
In some embodiments, the lipid monomers can be polymerized by UV irradiation, visible irradiation, gamma irradiation, redox polymerization, or thermal polymerization. When the lipid monomers are polymerized by UV or visible irradiation, the lipid bilayer may further comprise photoinitiators. In some embodiments, the duration of UV irradiation or visible irradiation is sufficient to photopolymerize the lipid monomers and/or the photoinitiators. In preferred embodiments, the duration of irradiation is between about 1 to 30 minutes. In some embodiments, the duration of UV irradiation is between about 5 to 30 minutes. In other embodiments, the duration of UV irradiation is between about 10 to 30 minutes, or between about 15 to 30 minutes, or between about 20 to 30 minutes.
In some embodiments, the lipid monomers are polymerized by redox polymerization. A redox polymerization mixture comprising an initiator-buffer component and NaHSO3 may be used for redox polymerization. In some embodiments, the redox polymerization mixture comprises any redox mixture that polymerizes the lipid monomers, i.e. a mixture having a reductant and an oxidant. In preferred embodiments, the initiator-buffer component comprises ammonium persulfate. Suitable mole ratios of ammonium persulfate, NaHSO3, and lipids can include about 10-500 AP: 10-500 NaHSO3:1 lipid, about 50-400 AP: 50-400 NaHSO3: 1 lipid, or about 100-300 AP: 100-300 NaHSO3: 1 lipid.
Preferably, the redox polymerization mixtures allows for the reaction to occur at a near neutral pH and to maintain a constant pH. In some embodiments, the redox polymerization occurs at a pH between about 5 to 9. In other embodiments, the redox polymerization occurs at a pH between about 5.5 to 7.5. In still other embodiments, the redox polymerization occurs at a pH between about 6 to 7.
1,2-bis[10-(2′,4′-hexadieoyloxy)decanoyl]-sn-glycero-2-phosphocholine (bis-SorbPC, Tm=29° C.) was synthesized according to previous protocols. Before use, a 0.5 mL aliquot of 18 mg mL−1 bis-SorbPC in 7:3 (v/v) MeOH:H2O was purified on a C18 column (Shimadzu Chromegabond WR C18, 5 μm, 250 mm×23 mm) using a 10 mL min−1 gradient (Shimadzu BCM-20A controller and LC-8A pumps) of H2O and MeOH. The MeOH volume in the mobile phase was increased from 50-70% in the first minute, increased from 70-85% over the next 20 min, increased from 85-100% over the next 40 min, and held at 100% for 5 min. The column was then flushed with 5% MeOH and 70% MeOH. All changes in the gradient were linear. The bis-SorbPC fraction was collected (elution time=40 min), dried under vacuum, and washed 3 times with chloroform before dissolving in 500 μL chloroform. The concentration of bis-SorbPC was determined by UV absorbance at 258 nm (∈=47100 M−1 cm−1 in MeOH) (Model 440CCD Array UV-Vis Spectrophotometer; Spectral Instruments, Inc., Tucson, Ariz.). Purified bis-SorbPC was stored at −80° C. Sorbyl functional groups are light sensitive; thus, purification and preparation of bis-SorbPC-coated particles were performed under UV-free, yellow light.
1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC, Tm=−17° C.) was purchased from Avanti Polar Lipids (Alabaster, Ala.). 3 μm silica particles (10% in water) were purchased from Polysciences, Inc. (Warrington, Pa.). FM1-43 was purchased from Invitrogen (Eugene, Oreg.). K2S2O8, NaHSO3, and (NH4)2S2O8 were purchased from Sigma (St. Louis, Mo.). All other chemicals were purchased from Fisher (Pittsburgh, Pa.). H2O was obtained from a Barnstead EasyPure UV/UF purification system. Buffers were filtered using membranes with 0.2 μm pores before use.
The chloroform from lipid stock solutions (DOPC and bis-SorbPC) was evaporated under Ar and lipids were dried by vacuum overnight (FreeZone 6, Labconco, Kansas City, Mo.). Lipid-coated particles were prepared by vesicle fusion. Briefly, dried lipids were suspended in phosphate buffered saline (PBS), pH 7.4, to a concentration of 1 mg mL−1. Lipid solutions were sonicated at 40% output power until clarity using a cup horn sonicator (Model W-380, Heat Systems-Ultrasonics, Inc., Newtown, Conn.) to produce small unilamellar vesicles (SUVs). Silica particles (3.0 μm diameter) were added to the SUV solution at a surface area ratio of 1 silica:6 lipid. The mixture was sonicated for 5 min and then allowed to rest for 30 min to promote vesicle fusion. Vesicle formation and fusion onto silica were performed at 35° C., above the phase transition temperature of both lipids. For some studies, 5 μm diameter silica particles were used and the lipid deposition was performed at room temperature using a UV pen lamp for 30 min at a lipid to surface area ratio of 1 silica:25 lipid.
Bis-SorbPC-coated particles were polymerized by one of three methods. Before all polymerizations, the lipid/silica solutions were degassed by bubbling N2 for 10 min. UV-polymerized particles were irradiated with a 100 W Hg Arc lamp with an IR-water filter and a band pass filter (U-330, Edmund Optics, Barrington, N.Y.) for 30 min while stirring. Redox polymerizations were performed using two different methods. Bis-SorbPC-coated particles were polymerized in the presence of 65 mM K2S2O8 and 22 mM NaHSO3 for 3 h at 35° C. or at a mole ratio of 1 lipid:300 (NH4)2S2O8: 300 NaHSO3 for 1 h at 35° C. After preparation (DOPC-coated particles) or polymerization (bis-SorbPC-coated particles) each set of particles was washed by 6 cycles of centrifugation, removal of supernatant, and suspension in fresh PBS buffer.
Particles were packed in capillaries (50 μm or 75 μm i.d.; 360 μm o.d.; Polymicro Technologies, Phoenix, Ariz.) against Microtight filter assemblies (Upchurch Scientific, Oak Harbor, Wash.). Particles were slurried in PBS and packed at 1000 psi N2 until an 8 cm packed bed was formed (50 μm) or at 560 psi He (75 μm). After packing, the N2 tank was turned off and the pressure was allowed to slowly decrease by bleeding out of the capillary over the next 1 h.
Aliquots of particles were treated on days 1, 2, 15, and 30 with triton X-100 or acetonitrile to monitor the stability of the lipid bilayer. Particles were aliquoted into individual tubes, centrifuged, and suspended in fresh PBS, pH 7.4, at 1.8×107 particles per 100 μL. Particles were treated with 25 μL of 50 mM triton X-100 at room temperature for 15 min or with 50% acetonitrile (v/v) and sonicated for 15 min. After incubation, particles were rinsed by 4 cycles of centrifugation, removal of supernatant, and suspension in fresh PBS to a final concentration of 1.8×108 particles mL−1.
FM1-43 was added to a particle slurry at a final concentration of 57 nM and allowed to intercalate into the lipid bilayer before imaging. Fluorescent images of particles were acquired on a Nikon Eclipse TE300 Quantum inverted microscope using a 40×/1.30 N.A. oil objective. Fluorescent images were obtained using rhodamine filters: λex=540/25 nm; λem=620/60 nm. Images were collected using a Quantix 57 back-illuminated CCD camera (Roper Scientific, Tucson, Ariz.) operated by MetaVue imaging software (Universal Imaging, Downingtown, Pa.). Images were analyzed using Image J. Data is presented as the mean±standard deviation (graphically represented as error bars) for n=3×100, with the intensity of 100 particles quantified for each of three batches of particles that were prepared separately.
Packed capillary columns were imaged with or without FM1-43, which was allowed to intercalate into the bilayers before excess was rinsed away. Images were acquired using the instrument described above, but with a 4×/0.13 N.A. objective. Data is presented as the mean±standard deviation for n=3×3, 3 measurements from 3 capillaries.
Flow cytometric analysis was performed using a FACScan flow cytometer (BD Biosciences, San Jose, Calif.) equipped with an air-cooled 15 mW Ar+ laser tuned to 488 nm. The emission fluorescence of FM1-43 was detected and recorded through a 582/42 nm bandpass filter in the FL2 channel. Data files consisting of approximately 50,000 events gated on forward scatter versus side scatter were acquired and analyzed using CellQuest PRO software (BD Biosciences). Appropriate electronic compensation was adjusted by acquiring particle populations with and without FM1-43. Data is presented as the average±standard deviation for n=3×50,000, with 50,000 gated events quantified for each of three batches of particles that were coated and polymerized independently.
Frontal chromatography was performed using a lab-built instrument. Isocratic elution with PBS, pH 7.4, as the mobile phase utilized pressure driven flow that was applied by a Micropro Syringe Pumping System (Eldex, Napa, Calif.) connected to a Cheminert injection valve (Valco, Houston, Tex.) with a 600 nL sample loop. The eluent was pumped at 1 μL min−1 at room temperature, resulting in 2750±50 psi when a packed column was present in the instrument. The elution profile was monitored by ultraviolet absorbance detection (Model 500 Detector, ChromTech, Apple Valley, Minn.) at 220 nm. Signal from the detector was collected with an A/D converter (NI USB-6221, National Instruments, Austin, Tex.) and software written in LabVIEW (National Instruments). All samples were prepared at a concentration of 100 μM in PBS buffer, pH 7.4. Statistical significance was determined using the two-tailed Student's t-test.
Results and Discussion
Fusing small unilamellar vesicle (SUV) with a hydrophilic, silica surface results in rapid rupture of the vesicles to form a supported bilayer. Bilayer formation via vesicle fusion results from a balance between the increase in adhesion energy and loss of curvature energy during adsorption, deformation, and rupture of the vesicles. The process is dependent on the support material, the type of lipid, the size of the vesicles, and the aqueous environment (e.g., presence of dissolved salts).
Silica (non-coated) particles, DOPC-coated particles, and poly(bis-SorbPC)-coated particles exhibited low fluorescence background prior to addition of FM1-43. After incubation with FM1-43, the fluorescence intensity of DOPC- and poly(bis-SorbPC)-coated particles increased, indicating the presence of lipid membranes on the surfaces of these particles. Additionally, the uniform fluorescence intensity indicates the presence of a bilayer membrane on the particle surface. To test the stability of the lipid bilayer coating, particles were exposed to surfactant (Triton X-100) or a high concentration of organic solvent (50% (v/v) ACN), washed, and then stained with FM1-43. These conditions, while more harsh than would be expected for most analyses involving lipid stationary phases, were chosen to provide a clear indication of the stability enhancements that might be obtained utilizing poly(lipid) membranes. The fluorescence intensity of DOPC-coated particles decreased significantly after exposure to surfactant or organic solvents, suggesting that the lipid bilayers were degraded. However, when poly(bis-SorbPC)-coated particles were exposed to either insult, the fluorescence intensity was retained after staining with FM1-43, indicating the enhanced bilayer stability after polymerization of the lipids.
To increase the throughput of particle characterization, flow cytometry was used to study large groups of particles since the scatter associated with each particle could be correlated with the fluorescence. Table 13 shows the mean fluorescence intensity of silica, DOPC-coated, and poly(bis-SorbPC)-coated particles in the presence and absence of FM1-43 and after treatment with surfactant or organic solvent. This data correlates well with the images presented in
Table 13 shows chemical stability of lipid bilayer coatings analyzed by flow cytometry.
In addition to examining the stability of the bilayer coatings against common chemical insults, the long-term stability was analyzed by imaging. Particles were stained with FM1-43 prior to each experiment. Images were collected on the day of preparation, and again on days 15 and 30. Between experiments, particles were dispersed in PBS, pH 7.4, and stored at 4° C. The data in
The physical stability of the bilayer coatings was analyzed by imaging capillaries packed with UV poly(bis-SorbPC)-coated silica particles, DOPC-coated silica particles, or bare silica particles in the presence or absence of FM1-43 (
The imaging and flow cytometry studies presented above show that poly(bis-SorbPC) membranes deposited on silica particles exhibit enhanced chemical and temporal stability compared to non-polymerized lipid bilayers (DOPC). However, aggregates of particles were observed after UV polymerization and these were exacerbated following membrane destabilizing conditions, e.g. surfactant or organic rinse (
To demonstrate the efficacy of poly(bis-SorbPC)-coated silica following packing into capillary columns, frontal analyses of three lipophilic small molecules that are known to cross cell membranes were performed using capillary LC.
Table 14 shows frontal chromatography retention analyses.
Importantly, column performance was highly reproducible. Frontal chromatograms yielded retention time relative standard deviations less than 4% for acetylsalicylic acid and less than 1% for benzoic acid and salicylic acid (n 4) over the course of 1 week, values that are on par with silica particles lacking a coating. This is particularly impressive when considering the operating pressure for each run exceeded 2700 psi. Thus not only do poly(bis-SorbPC) coated particles withstand slurry packing but also high pressures associated with the separation which would delaminate unpolymerized lipid coatings and lipid architectures. These key results provide the foundation for a new range of high pressure and thus higher throughput evaluation of mixtures for lipid membrane interactions.
Radical polymerization of bis-SorbPC and other lipids with functional groups can be achieved with redox mixtures. One of the most commonly used is the 1:1 mixture of potassium persulfate (K2S2O8, oxidant) and sodium hydrogen sulfite (NaHSO3, reductant). However, initiation and progression of the polymerization with this redox couple proceed only under acidic conditions (pH of the aqueous redox mixture is 1-2 depending on the concentration). This could be problematic for proteins incorporated into lipid vesicles. Unless milder conditions are used in the polymerization of lipids, transmembrane proteins often get denatured. For example, activity of Rhodopsin incorporated into bis-SorbPC vesicles, measured spectroscopically at 500 nm, diminished or disappeared after redox polymerization with the above redox couple. Therefore, other polymerization methods that proceed under neutral pH were investigated.
Both direct UV-induced and photo-initiated radical polymerizations were tried as alternatives. Photo-initiated radical polymerization was performed using DMABN and a UG-1 band pass filter (ca. 300-400 nm). Both UV and radical polymerization methods were applied to Rhodopsin incorporated bis-SorbPC vesicles in pH 5.5 MES buffer. Bis-SorbPC polymerization occurs in both conditions, but Rhodopsin activity also disappears concurrently (
Another alternative is to use another redox mixture that generates radicals under different pH conditions. One of those mixtures is AP and NaHSO3.
The photoactivity of Rhodopsin embedded in vesicles is usually measured in a pH 5.5 MES buffer at 10° C. Therefore, the polymerization of bis-SorbPC in these conditions using the initiators and the lipid ratio mentioned earlier is shown in
The effect of redox mixture on the spectroscopic properties of the vesicles prepared with MES buffer solution is shown in
Data from
A highly crosslinked polymer results when K2S2O8 and NaHSO3 are used as the redox couple to polymerize bis-SorbPC (Xn=40-600). The initiation reactions between S2O82− and HSO3− are shown below (Scheme 1).
S2O82−+HSO3−→HSO3*SO4*−+SO42−
SO4*−+H2O→HSO4−+OH*− (1)
HSO4− (pKa 1.9) is produced as a byproduct of initiation, resulting in an acidic solution. When bis-SorbPC lipids undergo redox-initiated polymerization, the decrease in pH does not affect the lipid structure or the resulting polymer; however, if membrane proteins are to be incorporated into bis-SorbPC membranes before redox polymerization, the resulting decrease in pH may alter the native protein conformation. Thus, the inventors sought to identify conditions that would be more compatible with membrane protein-functionalized matrices.
When (NH4)2S2O8 was substituted for K2S2O8 in the redox reaction, a solution with approximately neutral pH resulted. The mixture could be buffered to pH 7.4 and the solution still maintained a high degree of monomer to polymer conversion, as opposed to the previous redox couple, as indicated by the stability and lack of aggregation (
Stability tests of polymerized vesicles often reveal the type of polymer formed by the polymerization method, such as whether the polymers are linear or cross-linked. One commonly used stability test is Triton X-100 insertion.
The redox mixture was applied to Rhodopsin embedded bis-SorbPC vesicles to check on whether the initiator mixture is mild enough to maintain Rhodopsin activity and also to achieve a high degree of lipid polymerization. At first, crude samples with Rhodopsin (ROS membranes) were incorporated into bis-SorbPC vesicles in MES buffer and the polymerization was performed. The results are summarized in
Conclusions
Poly(bis-SorbPC) coatings on silica particles exhibited increased chemical, temporal, and physical stability compared to unpolymerized phospholipid bilayer coatings (DOPC). Redox-initiated polymerization yielded bilayers with greater stability than UV poly(bis-SorbPC) coatings, which aggregated after exposure to chemical insults. Additionally, chromatographic frontal analyses of small molecule analytes showed reproducible lipophilic retention on poly(bis-SorbPC)-coated particles polymerized with AP/NaHSO3, indicating that the highly stabilized lipid membrane environment of the poly(lipid) stationary phase presents an excellent platform for further development of biomimetic stationary phases. Thus, poly(bis-SorbPC) stationary phases show enhanced stability and high reproducibility compared to ILCs. The presence of a full phospholipid bilayer may also make poly(lipid) stationary phases more compatible with future incorporation of membrane proteins, presenting an excellent opportunity for new affinity chromatography approaches to study physiological and pharmaceutical modulators of phospholipid membranes and/or membrane proteins.
The AP and NaHSO3 redox mixture polymerizes bis-SorbPC vesicles at slightly acidic or near neutral pH solutions at similar conversion rates to the K2S2O8 and NaHSO3 redox mixture. Polymerization at near neutral pH is a critical feature when incorporating transmembrane proteins into bilayers composed of polymerizable lipids and stabilizing these supramolecular assemblies by cross-linking polymerization. The experimental evidence suggests that the AP and NaHSO3 redox mixture has all these desirable properties. In addition, the AP and NaHSO3 redox mixture is mild enough to maintain the photoactivity of the model transmembrane protein, Rhodopsin, during the polymerization.
Section 2B
Referring now to
Affinity capture coupled with matrix-assisted laser desorption/ionization-time of flight mass spectrometry (MALDI-TOF MS) is a label-free method to detect and identify a peptide or protein ligand that binds to a receptor immobilized on a solid support. In this approach, water-soluble receptors (typically antibodies) are used to extract the ligand from a complex mixture. The affinity capture surface is prepared by covalently immobilizing the receptor on a substrate surface that is compatible with MALDI-TOF MS analysis (the “direct” method), or on a solid support from which the captured analytes are subsequently desorbed and deposited on a standard MALDI plate (the “indirect” method).
In principle, affinity capture coupled with MALDI-TOF MS could be a powerful approach for characterizing ligands that target receptors embedded in artificial phospholipid membranes, which are widely used as models of natural cell membranes and recognition processes that occur at these membranes. A seminal paper by Marin et al. demonstrated a strategy for MALDI analysis of a ligand bound to a receptor incorporated into an artificial membrane. Bovine rhodopsin (Rho) was reconstituted into His-tagged lipid nanodiscs that were captured onto a self-assembled monolayer (SAM) on Au. Transducin was incubated with the Rho/nanodisc/SAM assembly and then subsequently detected using MALDI-TOF MS. However, background peaks for Rho, lipids, and the nanodisc scaffold protein were also present in the mass spectrum, which illustrates a major drawback of the nanodisc strategy—matrix deposition and laser ionization causes dissociation of the entire assembly, producing a complex background spectrum. Ideally, the assembly would be less structurally complex, and matrix deposition/laser ionization would cause only the ligand to dissociate from the receptor, leaving the rest of the molecular assembly intact.
A structurally simpler alternative is a PSLB that is deposited directly on a solid substrate. However, conventional PSLBs are typically composed of fluid-phase glycerophospholipids. The relatively weak, noncovalent interactions in the bilayer and between it and the underlying planar substrate are insufficient to maintain the PSLB structure upon exposure to air, organic solvents, and high vacuum; thus conventional PSLBs are not stable to the analysis conditions inherent to affinity capture coupled with direct MALDI-TOF MS.
PSLB stability can be greatly enhanced by polymerization of synthetic lipids bearing cross-linkable moieties, such as bis-Sorbyl phosphatidylcholine (bis-SorbPC), which suggests that a polymerized PSLB could be a functional platform for MALDI-TOF MS detection of ligands captured by incorporated receptors.
Bacterial toxins that target membrane-bound gangliosides are used to assess the suitability of polymerized PSLBs as an affinity capture surface for MALDI-TOF MS detection. Gangliosides are major membrane receptors for toxins such as cholera toxin, heat-labile enterotoxin and pertussis toxin. Poly(bis-SorbPC) PSLBs were doped with GM1, a monosialoganglioside that binds to the B-subunit of cholera toxin and heat-labile enterotoxin, and GD1a, a disialoganglioside that is a receptor for the B-subunit of pertussis toxin. The three individual ganglioside-toxin B pairs were characterized first, followed by simultaneous detection and identification of all three toxins. Cholera toxin B (CTB) and GM1 were used to assess the minimal detectable toxin concentration, as well as detectability in a complex matrix. The results show that a poly(bis-SorbPC) PSLB is stable to MALDI-TOF MS analysis conditions; the mass spectrum of the dissociated toxin is largely free of background peaks due to other components in the molecular assembly. Finally, the feasibility of on-plate trypsin digestion of CTB bound to GM1 in a PSLB, followed by MS/MS analysis of digested peptides, also was demonstrated. Overall, these results show that combining a poly(lipid) affinity capture platform with MALDI-TOF MS detection is a viable approach for identification and proteomic characterization of membrane-associated proteins in a label-free manner.
GM1 and 1,2-diphytanoyl-sn-glycero-3-phosphocholine (DPhPC) were purchased from Avanti Polar Lipids (Alabaster, Ala.). GD1a was purchased from Sigma-Aldrich (St. Louis, Mo.). 1,2-bis[10-(2′,4′-hexadienoloxy)decanoyl]-sn-glycero-3-phosphocholine (bis-SorbPC) was synthesized and purified by preparatory scale high performance liquid chromatography as described in Electronic Supplementary Material (SM). For safety considerations, only the binding domains of toxins were used. CTB and heat-labile enterotoxin B (LTB) were obtained from Sigma-Aldrich. Pertussis toxin B oligomer (PTB) was purchased from List Biological Laboratories, Inc (Campbell, Calif.). Water from a Barnstead Nanopure system with a minimum resistivity of 18 MΩ·cm was used. Stock toxin solutions were made by dissolving each toxin in nanopure water at 0.5 mg/mL. Phosphate buffered saline, pH 7.4 (PBS) contained the following components: 140 mM sodium chloride, 3 mM potassium chloride, 10 mM dibasic sodium phosphate, 2 mM monobasic potassium phosphate, 0.2 mM sodium azide. Silicon wafers were obtained from Wacker Chemie AG. Fetal bovine serum (FBS) was purchased from Invitrogen (Grand Island, N.Y.). Sinapinic acid was purchased from Fluka Analytical (St. Louis, Mo.) and alpha-cyano-4-hydroxycinnamic acid (HCCA) was provided by Bruker Daltonics Inc. (Auburn, Calif.). Peptide calibration standard II, containing Angiotensin II, Angiotensin I, Substance P, Bombesin, ACTH clip 1-17, ACTH clip 18-39, Somatostatin 28, Bradykinin Fragment 1-7, and Renin Substrate Tetradecapeptide porcine, was supplied by Bruker Daltonics Inc. Cytochrome c and myoglobin were purchased from Sigma-Aldrich. Trypsin Gold, mass spectrometry grade, was purchased from Promega (Madison, Wis.).
Preparation of Small Unilamellar Vesicles (SUVs) with Incorporated GM1 and GD1a.
Stock solutions of bis-SorbPC were prepared in pure chloroform. GM1 was dissolved in methanol and GD1a in 2/1 chloroform/methanol (v/v). GM1 and GD1a were mixed with bis-SorbPC at molar ratios of 1/99 and 1/4, respectively (expressed below as 1 mol % and 20 mol %, respectively). Organic solvents were evaporated from the lipid mixtures under a stream of argon, followed by vacuum drying for at least four hours. The lipids were then rehydrated with PBS to a concentration of 0.5 mg/mL, vortexed, and then sonicated in a Branson Sonicator with a cup horn at 35° C. until the solution was visibly clear (usually 30 min).
Preparation of Polymerized PSLBs.
Silicon wafers (cut to 0.8 cm×0.8 cm) were cleaned in piranha solution (7/3 concentrated H2SO4/H2O2) for 30 minutes and rinsed thoroughly in nanopure water. The silicon wafers were dried with a stream of nitrogen and incubated in 200 μL SUV solutions at 35° C. on a hot plate for at least 15 minutes to form PSLBs. Unfused SUVs were rinsed away with copious PBS buffer (at least 10 mL) without exposing the PSLB to air. A low-pressure mercury pen lamp with a rated intensity of 4500 μW/cm2 at 254 nm was directed through a bandpass filter (325 nm, 140 nm FWHM; U330, Edmund Optics) for 60 minutes to polymerize bis-SorbPC. The distance between the lamp and the PSLB was 7.6 cm.
Mass Spectrometric Detection of Toxins.
The toxin solution (0.5 mL of 0.24 μM CTB, 0.24 μM LTB, and/or 1 μM PTB) was incubated with GM1- and/or GD1a-incorporated PSLBs on 0.8 cm×0.8 cm silicon wafers for 1 hour. PSLBs were then rinsed thoroughly with water and dried under a nitrogen stream. Nonspecific binding was assessed by incubating toxins with PSLBs that lacked gangliosides, followed by rinsing and drying. The MS mass calibration standard was prepared by mixing 0.5 μL of myoglobin solution (3.8 mg in 500 μL), 1.0 μL of cytochrome c solution (1.2 mg in 500 μL) and 8.5 μL of saturated sinapinic acid (SA) in 70/30/1 H2O/acetonitrile/trifluoroacetic acid. The dried silicon wafers were mounted onto a MALDI plate (a microtiter plate (MTP) adapter for Prespotted AnchorChip Targets (Bruker)) using double-sided tape. The calibration standard (1 μL) was spotted on each wafer for external calibration. Three or four different spots of 1 μL SA solution were added to the remaining surface of each wafer and the solvent was allowed to evaporate under ambient conditions, crystallizing the SA. The plate was mounted in a Bruker Ultraflex III MALDI TOF/TOF mass spectrometer (Bruker Daltonics) equipped with a Smartbeam laser (Nd:YAG laser, 355 nm; spot diameter at sample=50 μm). After the laser ionization, ions were accelerated by a 20 kV electric field into the field-free flight tube and were detected in the positive ion linear detection mode. Spectra were exported as ASCII files and were processed using Origin 8 (OriginLab Corporation).
Preparation of Shrimp Extract.
About 25 g of shrimp was weighed and an equal mass of PBS buffer was added to the sample. The mixture was homogenized in a blender, then centrifuged at 3000 rpm for 1 h, and the supernatant was collected for use.
On-plate digestion and MS analysis of captured CTB. CTB was captured on a PSLB doped with GM1, as described above, and then enzymatically digested by spotting 10 μL of 0.01 μg/μL Trypsin Gold in 25 mM ammonium bicarbonate solution, pH 7.8, over the area of the PSLB that had been incubated with dissolved CTB. The digestion was carried out for 12 hours at 37° C. in a humidified chamber to prevent solvent evaporation after which the wafer was removed from the chamber and allowed to air-dry at room temperature. Different digestion times were tested and 12 hours was found to be optimal to maximize the intensities of the CTB peptide peaks for subsequent MS/MS analysis.
HCCA (20 μg) was dissolved in 250 μL of 50% acetonitrile, 2.5% trifluoroacetic acid and 47.5% nanopure water. One μL of this matrix solution was spotted onto the dried spot where the enzymatic digestion had taken place. One μL of peptide calibration standard II in HCCA solution was also spotted on the wafer. Wafers were mounted onto the MTP MALDI plate and analyzed using the MALDI TOF/TOF mass spectrometer as described above. The digested peptides were ionized, accelerated and detected in the reflectron mode for better resolution at lower mass-to-charge (m/z) ratios. After the full mass spectrum of the digested peptides was obtained, high energy (8 keV) collision induced dissociation was used to fragment the peptides to obtain sequence information (CID-LIFT mode with no added gas; background pressure is sufficient for CID as shown in the literature). Protein Prospector (University of California, San Francisco) was used to determine the theoretical m/z of the peptides generated from CTB digestion based on its amino acid sequence. The experimental peaks were compared to the theoretical peak list to make the assignments. For MS/MS spectral interpretation, b ions and y ions were compared and assigned according to the theoretical m/z values generated by Protein Prospector.
Results and Discussion
MALDI-TOF MS Detection of CTB, LTB and PTB.
Cholera toxin is composed of a dimeric A-subunit (Mr˜27,400) and five identical B-subunits, and the sequence is available. Each B-subunit (Mr˜11,600) contains a binding site for its membrane receptor, GM1. Heat-labile enterotoxin and cholera toxin are very similar with respect to structure, function and immunology; they also share GM1 as the cell surface receptor. Each B-subunit in LTB (Mr˜12,000) is slightly larger than a CTB subunit (and the sequence is given in this reference). Pertussis toxin has an enzymatic component A protomer (S1, Mr˜28,000) noncovalently bound to the B oligomer which is the binding component. Four dissimilar subunits form PTB:S2 (Mr˜21,900), S3 (Mr˜21,900), S4 (Mr˜12,000) and S5 (Mr˜11,750) in a molar ratio of 1:1:2:1, respectively.
To assess the feasibility of using MALDI-TOF MS to detect toxins captured on polymerized PSLBs doped with gangliosides, initial experiments were performed using poly(bis-SorbPC) PSLBs doped with 1 mol % GM1 to capture CTB and LTB under conditions where the toxin concentration (0.24 μM) was sufficient to saturate the GM1 receptors in the PSLB (see Section 9 in SM).
Pertussis toxin B was captured on poly(bis-SorbPC) PSLBs containing 20 mol % GD1a from a 1 μM solution, which was sufficient to saturate the GD1a receptors in the PSLB. The ganglioside mol % was higher than that used for CTB and PTB based on preliminary experiments on poly(bis-SorbPC) PSLBs doped with GD1a at 10-40 mol % (see
The spectra in
Background from Nonspecific Toxin Adsorption and Lipid Membrane Components.
Control experiments were performed to assess the degree of nonspecific adsorption of toxins to PSLBs because in any non-competitive bioaffinity assay, nonspecific adsorption is usually the major source of background. PSLBs composed of only poly(bis-SorbPC) (i.e., lacking gangliosides) were prepared and incubated with 0.24 μM CTB, 0.24 μM LTB, or 1 μM PTB, respectively. Nonspecific adsorption of CTB and PTB was not detectable (see
Another set of control experiments was performed to assess if lipid molecules, either gangliosides or bis-SorbPC monomers and/or oligomers, could be detected. PSLBs composed of 1 mol % GM1 and 20 mol % GD1a in poly(bis-SorbPC) were prepared and analyzed by MALDI-TOF MS, as described above, except that the toxin incubation step was eliminated. HCCA and SA were used as the matrices for low and high m/z ranges, respectively. In
Simultaneous Detection of CTB, LTB and PTB.
A number of label-free assays based on optical and electrochemical transduction principles have been developed for bacterial toxins, most notably for cholera toxin. However, these detection methods lack specificity, i.e., the signals from the analytes and nonspecific binding cannot be distinguished. Additionally, if two bacterial toxins target the same membrane receptor, as in the case of cholera toxin and heat-labile enterotoxin, these label-free methods cannot discern which toxin is present. In contrast, toxins with different molecular weights can be simultaneously captured on an affinity surface and identified in a label-free manner using MALDI mass spectrometry.
To assess the use of a ganglioside-doped lipid bilayer for simultaneous detection of CTB, LTB, and PTB, a poly(bis-SorbPC) PSLB containing 1 mol % GM1 and 20 mol % GD1a was incubated with a solution containing 0.24 μM CTB, 0.24 μM LTB, and 1 μM PTB. The mass spectrum is shown in
The correspondence of the peaks in
Application of the Affinity Capture Platform in Complex Samples.
Typically, biological samples contain analyte(s) in a complex matrix of biomolecules and other components, such as tissue homogenate, fecal matter, blood serum, etc. To assess the utility of the poly(lipid)-based affinity capture platform in complex samples, analysis of CTB in both fetal bovine serum (FBS) and shrimp extract as the sample matrix was performed. CTB was spiked into either 10% (v/v) of FBS or 10% (v/v) of shrimp extract in PBS buffer, at a final CTB concentration of 0.24 μM, and the samples were incubated with poly(bis-SorbPC) PSLBs containing 1 mol % GM1, followed by rinsing, drying, and analysis as described above. Representative mass spectra are shown in
Minimal Detectable Concentration.
The CTB-GM1 pair was used as a model system to estimate the minimal detectable concentration of protein captured on the PSLB-based affinity platform. Due to the semi-quantitative properties of MALDI-TOF MS, an approach based on the frequency with which CTB could be detected was used. After CTB was captured by a poly(bis-SorbPC) PSLB doped with GM1, SA matrix was spotted on different areas on the substrate. Multiple substrates were prepared and up to five matrix spots could be applied on each (the number of samples per substrate varied due to differences in substrate area). Due to the nature of dry-droplet matrix deposition, spatially heterogenous crystal formation occurs across the matrix spot which causes significant variations in signal strength. To overcome this variable, the laser was scanned across entire area of each matrix spot at distance intervals of less than 50 μm (the diameter of the laser spot). Detection of a CTB monomer peak at 11.6 k m/z with a S/N≥3 anywhere on the substrate was recorded as a detectable sample.
The minimal detectable concentration is defined here as the CTB solution concentration that produces a peak with S/N≥3 for 100% of the samples analyzed. In these experiments, the mol % of GM1 was constant (1 mol % in poly(bis-SorbPC)) and the CTB concentration of the solution that was incubated with the PSLB was varied. A large CTB concentration range, from 0.5 nM to 1 μM, was screened, from which the minimal detectable concentration was determined to be in the range of 1-5 nM. A larger number of samples was prepared and analyzed in this concentration range, with each matrix spot counted as one sample. The number of samples that gave a CTB m/z peak with S/N≥3 divided by the total number of samples is reported as a detection frequency in Table 15. At 4 nM CTB, the detection frequency is 100%, so this concentration is a conservative estimate of the minimal detectable CTB concentration. With respect to definitions used in clinical chemistry, 4 nM CTB is the concentration that produces a sensitivity of 100%, and since CTB was never detected in samples lacking the toxin, the specificity is 100%. The estimate of 4 nM is specific to the analysis conditions employed here, for example, the amount of GM1 is finite and the incubation time was selected to achieve an apparent steady state (but not equilibrium which, due to mass transport limitations, would have required much longer times). The minimal detectable concentration of CTB likely could be lowered by increasing the mol % of GM1 and the incubation time. The dissociation constant (KD) for GM1-CTB also plays a role. Apparent KD values reported for CTB-GM1 range from 4.55 μM to 370 nM, a very wide range attributed to significant differences in experimental parameters; thus it is also possible that at 4 nM, the amount of bound CTB is limited by its binding affinity to GM1. Minimal detectable concentrations for cholera toxin using label-free SPR methods are in the nM range, such as the 4 nM reported here.
Table 15 provides estimations of the MALDI-TOF MS minimal detectable concentration for CTB bound to 1 mol % GM1 in poly(bis-SorbPC) PSLBs.a
a This table focuses on a narrow CTB concentration range that was identified from screening samples over a larger concentration range.
b Frequency is reported as the number of samples in which CTB was detected/total samples that were analyzed.
On-Plate Tryptic Digestion of Captured CTB.
Molecular weight information may be adequate to identify multiple analytes with resolvable molecular weights, as demonstrated above. However, when multiple analytes cannot be distinghuished solely based on molecular weight differences, additional steps may be necessary. To further explore the applicability of the PSLB-based affinity capture platform, on-plate tryptic digestion of captured CTB was performed and the fingerprint spectra of CTB peptide fragments were obtained. A solution of 0.24 μM CTB was incubated with a poly(bis-SorbPC) bilayer doped with 1 mol % GM1, rinsed with nanopure water, and dried. The trypsin concentration and on-plate digestion time were varied to obtain maximum amino acid coverage (data not shown). The optimal conditions were found to be 0.01 μg/μL Trypsin Gold and 12 hours, respectively, and MALDI-TOF MS was performed on CTB peptides generated using these conditions.
A typical mass spectrum, in which the CTB peptides produced by digestion are labeled with asterisks, is shown in
Table 16 is a summary of peptide fragments observed in MALDI-TOF mass spectra of a CTB tryptic digest.
In some cases, e.g. when analyzing multiple and/or unknown proteins, the m/z of the constituent peptides may not be sufficient to identify the proteins; in these cases, identification may be possible using MS/MS. Here the amino acid sequences of three peptides produced by on-plate tryptic digestion of captured CTB were obtained using MS/MS. These three peptides were selected for analysis because their relative yields were high in comparison to those of the other nine peptides (which were too low for this analysis). Spectra of sequences with MWs of 1216 Da, 1372 Da and 2042 Da are shown in
Minimal Detectable Surface Coverage of CTB.
Another estimate of the minimal detectable concentration was obtained by maintaining the CTB concentration at a high, constant value (0.24 μM) and varying the mol % of GM1 in the PSLB from 0.005 to 1 mol %. Under these conditions, the concentration of CTB greatly exceeds the GM1 concentration and it is reasonable to assume that one CTB pentamer binds to each GM1; thus this method provides an estimate of the minimal detectable concentration in units of protein surface coverage. PSLBs with a large mol % range of GM1 (0.005-1 mol %) were prepared and screened for CTB detectability. The minimal detectable surface coverage was determined to reside in the range of 0.01-0.05 mol %; thus more samples were prepared in this range. The results are reported in Table 17. At less than 0.05 mol %, the detection frequency was low but reached unity at 0.05 mol % and above. Assuming one bound CTB per GM1 and respective projected molecular areas for GM1 and bis-SorbPC of ˜1 nm2 and ˜0.5 nm2, the surface coverage of CTB bound to 0.05 mol % GM1 in a bis-SorbPC PSLB is 1.7×10−13 mol/cm2. This surface coverage corresponds to ˜3% of a CTB monolayer, assuming that one monolayer is 6.6×10−12 mol/cm2 based on a projected area of ˜25 nm2 per CTB pentamer. Assuming the average matrix spot diameter is ˜2 mm, this surface coverage corresponds to ˜2×10−14 mol of CTB within one matrix spot and ˜5×10−16 mol of CTB within one 50 μm diameter laser spot.
aThis table focuses on a narrow range of GM1 mol % that was identified from a screening samples over a larger range of GM1 mol %.
bFrequency is reported as the number of samples in which CTB was detected/total samples that were analyzed.
Saturable Binding of Toxins to Poly(Bis-SorbPC) PSLBs Doped with Gangliosides.
CTB labeled with Alexa Fluor 488 (Alexa 488-CTB) was purchased from Invitrogen (Eugene, Oreg.). Total internal reflection fluorescence microscopy (TIRFM) was used to measure the binding of Alexa 488-CTB to 1 mol % GM1 in a polymerized bis-SorbPC PSLB. The TIRFM system has been described previously. The 488 nm line of an Ar+ laser (Ion Laser Technology) was coupled into a fused silica slide using a 45° prism (Edmund Optics). Immersion oil (Type FF, Cargille Laboratories, Cedar Grove, N.J.) with a refractive index of 1.4790 was used to couple the prism to the fused silica slide, which formed the lower wall of a liquid flow cell that was mounted on an stage of an inverted microscope (Nikon Diaphot). The PSLB was formed on the upper surface of the slide as described in manuscript. One of the internal reflections at the upper surface of the slide was used to excite fluorescence from Alexa 488-CTB bound to the PSLB. The fluorescence signal was collected using a 4× objective, directed through a 535DF35 (Omega Optical, Brattleboro, Vt.) band pass filter, and detected using CCD (Andor Technology USA, South Windsor, Conn.).
The binding curve in
Conclusions
This work demonstrates that cross-linking lipid polymerization provides the stability necessary for implementation of a receptor-doped PSLB as an affinity capture platform for label-free protein detection using MALDI-TOF MS. Simultaneous capture and detection of CTB, LTB and PTB was performed, showing that differences in ligand molecular weight are sufficient to distinguish among multiple captured proteins. The high resistance of poly(bis-SorbPC) membranes to nonspecific protein adsorption is another feature that makes them useful for analysis of complex biological matrices. In some cases, differences in molecular weights among captured proteins may be inadequate for their identification. As demonstrated here, on-PSLB trypsin digestion can be employed to obtain the molecular weights of peptide fragments using MS/MS, which illustrates the potential use of the PSLB-based platform for proteomic identification of membrane-associated proteins. Finally, it is feasible to incorporate transmembrane proteins, such as bovine rhodopsin and ion channels, into poly(lipid) membranes with retention of activity. This suggests the possibility of using the approaches described herein to capture and identify ligands that bind to transmembrane protein targets.
As used herein, the term “about” refers to plus or minus 10% of the referenced number.
Various modifications of the invention, in addition to those described herein, will be apparent to those skilled in the art from the foregoing description. Such modifications are also intended to fall within the scope of the appended claims. Each reference cited in the present application is incorporated herein by reference in its entirety. Additional advantages and features of the present invention are apparent in U.S. Application No. 62/018,794 and U.S. Application No. 62/018,822, the specifications of which are incorporated herein in their entirety by reference.
Although there has been shown and described the preferred embodiment of the present invention, it will be readily apparent to those skilled in the art that modifications may be made thereto which do not exceed the scope of the appended claims. Therefore, the scope of the invention is only to be limited by the following claims. In some embodiments, the figures presented in this patent application are drawn to scale, including the angles, ratios of dimensions, etc. In some embodiments, the figures are representative only and the claims are not limited by the dimensions of the figures. In some embodiments, descriptions of the inventions described herein using the phrase “comprising” includes embodiments that could be described as “consisting of”, and as such the written description requirement for claiming one or more embodiments of the present invention using the phrase “consisting of” is met.
This application claims priority to U.S. Provisional Patent Application No. 62/018,794 filed Jun. 30, 2014 and U.S. Provisional Patent Application No. 62/018,822 filed Jun. 30, 2014, the specifications of which are incorporated herein in their entirety by reference.
This invention was made with government support under Grant Nos. T32 GM008804, RO1 GM095763 and RO1 EB007047 awarded by NIH and CHE0518702, awarded by NSF. The government has certain rights in the invention.
| Filing Document | Filing Date | Country | Kind |
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| PCT/US2015/038539 | 6/30/2015 | WO | 00 |
| Publishing Document | Publishing Date | Country | Kind |
|---|---|---|---|
| WO2016/004029 | 1/7/2016 | WO | A |
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| Number | Date | Country | |
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| 20170120218 A1 | May 2017 | US |
| Number | Date | Country | |
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| 62018794 | Jun 2014 | US | |
| 62018822 | Jun 2014 | US |
