Supported Lipid Bilayers at Skeletonized Surfaces for the Study of

Jan 9, 2012 - Department of Chemistry, University of Florida, Gainesville, Florida 32611-7200, United States. Langmuir , 2012, 28 (5), pp 2835–2841...
2 downloads 0 Views 3MB Size
Download PDF
Article pubs.acs.org/Langmuir

Supported Lipid Bilayers at Skeletonized Surfaces for the Study of Transmembrane Proteins Roxane M. Fabre, George O. Okeyo, and Daniel R. Talham* Department of Chemistry, University of Florida, Gainesville, Florida 32611-7200, United States S Supporting Information *

ABSTRACT: Skeletonized zirconium phosphonate surfaces are used to support planar lipid bilayers and are shown to be viable substrates for studying transmembrane proteins. The skeletonized surfaces provide space between the bilayer and the solid support to enable protein insertion and avoid denaturation. The skeletonized zirconium octadecylphosphonate surfaces were prepared using Langmuir−Blodgett techniques by mixing octadecanol with octadecylphosphonic acid. After zirconation of the transferred monolayer, rinsing the coating with organic solvent removes the octadecanol, leaving holes in the film ranging from ∼50 to ∼500 nm in diameter, depending on the octadecanol content. Upon subsequent deposition of a lipid bilayer, either by vesicle fusion or by Langmuir−Blodgett/Langmuir−Schaefer techniques, the lipid assemblies span the holes providing reservoirs beneath the bilayer. The viability of the supported bilayers as model membranes for transmembrane proteins was demonstrated by examining two approaches for incorporating the proteins. The BK channel protein inserts directly into a preformed bilayer on the skeletonized surface, in contrast to a bilayer on a nonskeletonized film, for which the protein associates only weakly. As a second approach, the integrin α5β1 was reconstituted in lipid vesicles, and its inclusion in supported bilayers on the skeletonized surface was achieved by vesicle fusion. The integrin retains its ability to recognize the extracellular matrix protein fibronectin when supported on the skeletonized film, again in contrast to the response if the bilayer is supported on a nonskeletonized film.

■

INTRODUCTION Approximately 20−30% of the genome of organisms encodes for membrane-bound proteins,1,2 and the importance of membrane proteins is reflected in the fact that they currently account for two-thirds of all protein drug targets. 3−5 Unfortunately, the high complexity of cells makes it difficult to study membrane proteins in their natural environment, so viable model systems are critical. Supported lipid bilayers have proven to be useful for studies of many aspects of cell membrane dynamics such as lipid and protein diffusion, domain formation,6 and ion transport trough membranes,7,8 as well as membrane surface processes such as cell adhesion,9 ligand−receptor interactions, and signal transduction. Tamm and McConnell developed the process of generating supported lipid bilayers on substrates by vesicle fusion,10 a process that is now widely used to form bilayer models with ever-increasing complexity.11 Many surfaces have been used to support lipid bilayers, such as glass, mica, silicon, metal films, and polymers.8,12−14 It is well understood that interactions between the bilayers and the support can perturb membrane components and membrane dynamics. Transmembrane proteins, in particular, with complex structures that can include large cytoplasmic and extracellular domains that protrude from the membrane can be adversely influenced by the close proximity of the substrate, © 2012 American Chemical Society

hindering their incorporation, decreasing mobility due to frictional coupling with the support, or even leading to denaturation. Methods have been developed to increase the spacing between the membrane and the support to reduce the bilayer−substrate interactions. Strategies include forming bilayers on cellulose films,12 polymer cushions,15−19 and nanoporous microbeads,20 or by using tethered molecules that help suspend the bilayer above the surface.15,21−28 These strategies are highly effective, but at the same time are not all generally applicable or easy to use, sometimes requiring specialized molecules, or, in many cases, limited to a specific type of solid support. Therefore, significant motivation remains to develop new strategies for preparing supported bilayers that address the issues of membrane mobility and stability, while avoiding adverse effects of the substrate. The strategy described in the present study approaches the problem from the point of view of the solid support, pursuing a surface or coating that will support bilayers while addressing the problems outlined above without the need to modify the membrane or utilize specialized molecules. Ideally the same surface treatment or coating is transferable to varied supports, Received: November 14, 2011 Revised: January 5, 2012 Published: January 9, 2012 2835

dx.doi.org/10.1021/la204485n | Langmuir 2012, 28, 2835−2841

Langmuir

Article

(KSV Instruments, Stratford, CT). A hydrophobic slide was dipped down through a Langmuir monolayer of either octadecylphosphonic acid, to make a nonskeletonized surface, or a mixture of octadecanol and octadecylphosphonic acid to form skeletonized films. For transfer, monolayers were held at a constant pressure of 20 mN min−1, and slides were dipped into a vial beneath the subphase. Different mixtures were studied, from 0 to 50% in volume of octadecanol. The vial containing the slide was removed from the trough, and zirconyl chloride was added to bring the Zr4+-ion concentration to 3 mM. After 4 days of incubation, the slide was rinsed with water and an additional ethanol rinse was performed on the mixed monolayers to remove the free alcohol molecules. Characterization of the skeletonized films was performed by atomic force microscopy. Supported Lipid Bilayers. All experiments were performed at room temperature. Unilamellar vesicles were formed following the same procedure as described previously.29 Briefly, the chloroform lipid mixture was dried via nitrogen, and the film was hydrated in tris buffer composed of 10 mM trizma hydrochloride and 100 mM sodium chloride at pH 7.4 to obtain a lipid concentration of 0.5 mg mL−1. After five freeze−thaw cycles, the lipid suspension was extruded 11 times trough polycarbonate membranes with a pore diameter of 100 nm. Supported lipid bilayers were formed by the adsorption and fusion of the vesicles at 0.5 mg mL−1 concentration on the hydrophilic modified surface. After formation of the bilayers visualized by the increase and stabilization of the signal, the surface was washed with buffer for 30 min. The BK channel protein was incorporated by flowing a solution of the protein, 662 μg/mL at 100 μL/min, over the supported bilayer and then rinsing with buffer after the SPREE signal stabilized. BK Ion Channel Expression. Plasmid DNA encoding a histidinetagged BK channel constructed with a C-terminal deletion from amino acid 347 was transcribed in vitro by the mMessage mMachine T7 ultra kit (Ambion, Austin, Texas). RNA was precipitated with LiCl, washed and centrifuged in ethanol (70%), dissolved in RNase-free water, and injected (46 nL per oocyte, ∼50 ng of RNA) into defolliculated stage V or VI oocytes maintained at 19 °C in ND 96 oocyte culture medium (96 mM NaCl, 2 mM KCl, 1 mM MgCl2, 10 mM HEPES, 1.8 mM CaCl2, pH 7.4) enriched with sodium pyruvate (2.5%), penicillin/ streptomycin (1%), and horse serum (5%). Functional channel expression was verified by two-electrode voltage clamping (TEVC). Oocytes expressing BK channel constructs were rinsed in high K buffer (400 mM KCl, 5 mM PIPES, pH 6.8) supplemented with 100 μM phenylmethylsulfonylfluoride, 1 μM pepstatin, 1 μg mL−1 aprotinin, 1 μg mL−1 leupeptine, 1 μM p-aminobenzamidine and transferred to a 1 mL ground glass tissue grinder (Kontes Duall). Ground oocytes were solubilized using a buffer solution at a final concentration of 10 mM βoctyl glucoside containing 20 mM Tris buffer, 500 mM KCl, and 20 mM imidazole, pH 7.5 with 5 μL/mL mammalian protease inhibitor cocktail (Sigma-Aldrich, St. Louis, MO) added. The suspension was then agitated gently for 1 h at 200 rpm on a platform shaker followed by centrifugation at 14 000 rpm at 4 °C to separate the solubilized mixed micelles from cellular debris. Two milliliters of the supernatant solutions was loaded onto 1 mL Histrap FF columns (GE Healthcare Life Sciences, PA) for purification by immobilized metal ion affinity chromatography. Columns were equilibrated with binding buffer (10 mM β-octyl glucoside, 20 mM Tris buffer, 500 mM KCl, and 20 mM imidazole, adjusted to pH 7.5) at a flow rate of 1 mL min−1 and washed with the same buffer. Unbound material was washed off using 5 column volumes of binding buffer, and then fractions of 1.5 mL eluted in a stepwise manner with an elution buffer containing 500 mM imidazole and all other components of the binding buffer at pH 7.4 and a flow rate of 1 mL min−1. Immunoblotting was used to determine the identity of purified protein samples. Reconstitution of Integrin. Integrin-containing proteoliposomes were fused directly onto the zirconated surfaces to form bilayers. The proteoliposome preparation is based on a procedure described by Sinner et al.23 A solution of 1 mg mL−1 POPC in chloroform was injected into a vial and dried via a nitrogen stream to a uniform dry lipid film. A volume of 20 μL at 100 μg mL−1 integrin α5β1 was then added with 3 mL of tris-Mg buffer (10 mM trizma hydrochloride, 100

enabling use of different surface-limited analytical techniques without changing the coating chemistry. Recent papers introduced the idea of using zirconium phosphonate or zirconium phosphate modified surfaces to support lipid bilayers.29,30 The zirconated surfaces are known to have a strong affinity for divalent phosphate, to the exclusion of common monovalent groups, and we demonstrated that by incorporating a low percentage of phosphatidic acid lipids into the bilayer, the strong binding of these lipids provides anchoring points to stabilize the bilayers even when removed from water. The viability of bilayers on the zirconium phosphonate surface as membrane models was demonstrated by quantifying binding of the membrane protein, mellitin.29 In the present work, we address the issue of providing space for transmembrane species and separating the bilayer from the surface. In contrast to other approaches that elevate supported lipid bilayers from the support, the strategy involves the creation of reservoirs beneath the membrane to incorporate transmembrane proteins. To achieve the reservoirs, skeletonized zirconium phosphonate films are prepared, building on concepts from the early days of Langmuir−Blodgett films. The skeletonized films yield holes in the zirconium phosphonate surface that the membrane can span, allowing transmembrane proteins to be incorporated. To demonstrate the viability of the approach, the membrane binding behavior is evaluated for two different proteins that play key roles in biological processes. The first, BK ion channel, is expressed in a wide variety of fundamental cells including nerve cells and is involved in controlling firing patterns in neurons, muscle cells, and endocrine cells.31 The BK channel is shown to insert directly into bilayers supported on the skeletonized surfaces. The second protein, integrin α5β1, is part of a family of cell adhesion molecules that interact with the extracellular matrix (ECM). Integrin-mediated adhesion leads to intracellular signaling processes that regulate cell survival, proliferation, and migration.32 Some integrins can recognize a single ECM ligand, while others bind several different ECM proteins. The integrin α5β1 binds to several ligands including fibronectin,32 which is used here in a ligand binding assay to assess the transmembrane function. The interaction mechanisms of these two proteins with the supported lipid bilayers on the skeletonized surface were analyzed using surface plasmon resonance enhanced ellipsometry (SPREE). Behavior is compared to bilayers on the nonskeletonized surface and results from other commonly used supported lipid bilayer strategies.

■

EXPERIMENTAL SECTION

Materials. The lipids 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) and 1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-Lserine (POPS) were purchased from Avanti Polar Lipids (Alabaster, AL). Detergent-solubilized human integrin α5β1 and bovine fibronectin were obtained from Chemicon (Temecula, CA). The refractive index matching fluid diodomethane and zirconyl chloride (98%) were purchased from Sigma-Aldrich (St. Louis, MO). Slides used for SPREE experiments were SF10 glass (Schott Glass) with 28.5 nm of gold evaporated onto a 2 nm chromium adhesion layer. Glass microscope slides for the AFM experiments were from Gold Seal (Portsmouth, NH). Zirconium Phosphonate-Modified Surfaces. 33 Gold slides were cleaned and then rendered hydrophobic by immersion in a 1 mM octadecylmercaptan solution in ethanol for 16 h. Glass slides were rendered hydrophobic with an octadecyltrichlorosilane layer. Langmuir−Blodgett film depositions were performed using a KSV 3000 Teflon-coated LB trough with hydrophobic barriers 2836

dx.doi.org/10.1021/la204485n | Langmuir 2012, 28, 2835−2841

Langmuir

Article

Figure 1. Skeletonized zirconium phosphonate surfaces for supporting lipid bilayers. (a) A mixed monolayer is deposited by the LB technique on a hydrophobic surface. (b) After incubation with zirconium ions, the surface is washed with ethanol to remove the noncovalently bound molecules. (c) Lipid bilayers can be formed on the skeletonized surfaces by vesicle fusion or by Langmuir−Blodgett/Langmuir−Schaefer techniques. mM sodium chloride, 1 mM magnesium chloride, 0.2 mM calcium chloride, pH 7.4) to give a protein to lipid ratio of 1:5000, and the mixture was vortexed for a few hours. The proteoliposome solution was then extruded 11 times through polycarbonate membranes with a pore diameter of 100 nm. A volume of 2 mL of the proteoliposome suspension was used directly for vesicle fusion procedures performed in the SPREE flow cell using a flow rate of 150 μL min−1. Integrin binding assays23 were conducted by adding fibronectin at a concentration of 15 μg mL−1 to the SPREE cell. After incubation for 30 min, the surface was rinsed with buffer until the signal stabilized. Instrumentation. Surface plasmon resonance enhanced ellipsometry (SPREE) measurements were performed on a commercial EP3SW imaging system (Nanofilm Surface Analysis, Germany). The ellipsometer employed a frequency-doubled Nd:YAG laser (adjustable power up to 20 mW) at 532 nm. Linearly p-polarized light was directed through a 60° equilateral SF10 prism coupled to a gold-coated SF10 slide via diiodomethane index matching oil in the Kretschmann configuration. The angle of incidence was kept at 64° for all experiments because this condition provided the highest sensitivity. For all experiments, a flow cell with a 100 μL sample volume was used. Curve fitting of the experimental data used the analysis programs AnalysR (Nanofilm) and BIAevaluation (Biacore) for the two-state model. Atomic force microscopy imaging was performed in tapping mode in air using a Multimode AFM with a Nanoscope IIIa controller (Digital Instruments, Santa Barbara, CA). Standard silicon tips were used, with a nominal force constant of 42 N m−1 and a nominal resonance frequency of 330 kHz (Nanosensors, Switzerland).

Figure 2. AFM images of skeletonized surfaces on glass slides. The supports were rendered hydrophobic by an OTS layer, and the skeletonized film was formed on the surface with different 1octadecanol concentrations of (a,a′) 20% and (b,b′) 10%.

■

RESULTS Characterization of the Skeletonized Zirconium Phosphonate Surfaces. The Langmuir−Blodgett (LB) technique was used to form the skeletonized zirconium octadecylphosphonate modified surfaces (Figure 1). A mixed monolayer of ODPA and octadecanol was formed on the LB trough and deposited as a monolayer onto a hydrophobic support. Exposing the surface to Zr4+ binds the metal ions to the surface, cross-linking the phosphonate groups, leading to a stable monolayer.34 However, the alcohol molecules do not bind Zr4+ ions so that an ethanol rinse removes the free octadecanol leading to the skeletonized film. The skeletonized surfaces were then characterized by atomic force microscopy. Two different surfaces are shown in Figure 2, based on using

10% and 20% octadecanol, which produced holes of approximately 100 and 500 nm diameter, respectively. Most holes are 2−3 nm deep, consistent with the absence of the zirconated octadecanol/ODPA layer. A few of the larger holes experience deeper pits when coincident with defects in the priming octadecylmercaptan or OTS layer. The holes in the surface can be spanned by lipid layers, as demonstrated upon deposition of an LB monolayer of POPC. Figure 3 shows AFM images of the lipid layer on the two surfaces derived from 10% and 20% octadecanol. The lipid layer spans the holes, creating homogeneous surfaces. If the holes are too large, such as those formed from a 50% octadecanol mixture, the lipid monolayer 2837

dx.doi.org/10.1021/la204485n | Langmuir 2012, 28, 2835−2841

Langmuir

Article

Figure 3. AFM images of a POPC lipid monolayer on skeletonized surfaces on glass. The lipid monolayer was deposited by the LB technique on the skeletonized surfaces derived from (a) 20% and (b) 10% 1-octadecanol. The holes from the skeletonized zirconium phosphonate surfaces (Figure 2) can no longer be seen, indicating that the lipid layer spans the holes.

conforms to the holes, so further studies were limited to mixtures of 10% and 20% octadecanol. Supported Lipid Bilayers and Incorporating the BK Channel. To test the viability of the skeletonized support, mixed POPC/POPS (80/20) lipid bilayers were formed by vesicle fusion and then exposed to the BK ion channel protein. Both the vesicle fusion steps and the protein insertion steps were monitored using SPREE. Figure 4 compares the processes on three surfaces, the nonskeletonized film and skeletonized surfaces derived from 10% and 20% octadecanol. For each support, vesicle fusion can be observed as an increase in ψ corresponding to an increase in layer thickness. After the systems stabilize, the increase corresponds to a thickness of near 5 nm, which is consistent with the formation of lipid bilayers, using a value of 1.45 for the refractive index of the lipid membrane. There is a slightly longer time constant for vesicle fusion on the skeletonized films, but the equilibrated bilayer thickness is the same on each surface. After rinsing away the free vesicles with buffer, the BK channel was introduced above the supported lipid bilayer. An increase in ψ is again observed, confirming the adsorption of the ion channel at the lipid bilayers. After the signal stabilized, desorption of nonincorporated proteins was performed by buffer rinsing. Whereas the protein is completely removed from the bilayers on the nonskeletonized films, the protein is retained on the skeletonized surfaces. After rinsing, 35% of the originally adsorbed protein is retained on the skeletonized surface derived from 10% octadecanol, and 62% of the ion channel remains on the skeletonized surface derived from 20% octadecanol. Full desorption of the transmembrane protein from the nonskeletonized surface suggests that the protein could not incorporate into the membrane that is in close contact with the inorganic support. On the other hand, protein insertion into the membrane is apparent on the skeletonized surfaces. The time constant for the protein adsorption step is longer on the skeletonized films, further suggesting a different mechanism of protein−membrane interaction on these surfaces. To better understand the differences, the interaction kinetics measured with SPREE were analyzed by curve fitting using numerical integration analysis. A simple one-step adsorption model did not provide a good description of the data. On the other hand, as a transmembrane protein, the insertion of BK channel into the lipid membrane can be represented as a two-step interaction.35 The first step describes the adsorption of the

Figure 4. SPREE analysis of BK ion channel incorporation into lipid membranes supported on zirconium phosphonate modified surfaces prepared using (a) 20%, (b) 10%, and (c) 0% octadecanol. The kinetics study shows the formation of supported lipid bilayers followed by the insertion of the transmembrane protein. After formation of the POPC/POPS lipid bilayers, free vesicles were then removed with buffer rinsing (*). The membrane was then incubated with the ion channel and then again rinsed with buffer. The buffer used was trizma hydrochloride and sodium chloride at pH 7.4.

protein to the membrane surface, and the second is the incorporation of the BK channel into the membrane. The twostep reaction model can be represented as: ka1

ka2

kd1

kd2

P + L XooY PL XoooY PL* (1)

#tab;where in the first step, the protein, P, associates with the lipid, L, to give PL, which is then changed to PL* in the second step. The species PL* cannot dissociate directly to P and L. The association and dissociation rates for the first and second steps are ka1, kd1, ka2, and kd2. The affinity constants for the first and second steps are K1 and K2, and equal ka/kd for the respective steps. The total affinity constant KA (M−1) is the product K1K2. Kinetic parameters derived using this two-step interaction model are presented in Table 1. The kinetic constants indicate that the second step is far faster for the membrane supported on the skeletonized surfaces. In addition, the affinity constant of the protein is much higher on the skeletonized films. Fusion of Protein-Loaded Vesicles and the Binding of Fibronectin to Integrin α5β1. As a further test of the utility of the skeletonized surfaces for studying transmembrane proteins, 2838

dx.doi.org/10.1021/la204485n | Langmuir 2012, 28, 2835−2841

Langmuir

Article

Table 1. Equilibrium and Kinetic Parameters for the Adsorption of BK Ion Channel to POPC/POPS Lipid Membranes Using the Two-Step Binding Modela 20% skel. 10% skel. control a

ka1 (×103 M−1 s−1)

kd1 (×10−1 s−1)

K1 (×104 M−1)

ka2 (×10−2 s−1)

kd2 (×10−5 s−1)

K2

KA (×106 M−1)

5.25 ± 0.17 7.66 ± 0.22 30.3 ± 0.65

1.31 ± 0.12 2.69 ± 0.09 1.08 ± 0.11

4.01 ± 0.19 2.85 ± 0.08 28.1 ± 0.34

1.82 ± 0.06 1.04. ± 0.11 ∼0

0.912 ± 0.012 5.48 ± 0.19 1.10 ± 0.07

2000 ± 75 190 ± 15 1.75 ± 0.08

80.0 ± 0.23 5.40 ± 0.21 0.49 ± 0.01

Data were averaged over three experiments.

supported bilayers were formed by fusion of protein-loaded vesicles onto the zirconated surface. In this experiment, the integrin α5β1 was incorporated into POPC vesicles, and following bilayer vesicle fusion to form the supported bilayers, the binding capacity of the integrin receptor to the ligand fibronectin was studied by SPREE (Figure 5). In the first step,

Γ = 3d

(n2 − nb2) (n2 + 2)(r(nb2 + 2) − υ(nb2 − 1))

(2)

where Γ is the surface concentration of the protein, d is the layer thickness, nb and n are the refractive indexes of the buffer layer and the adsorbed layer, respectively, r is the specific refractivity of fibronectin (0.243 mL g−1), and υ is the partial specific volume of fibronectin (0.729 mL g−1).37,38 After free proteoliposomes are rinsed away, addition of fibronectin results in a binding signal of 0.69°, corresponding to 51 ng cm−2 (Figure 5). This surface concentration represents fibronectin partial coverage of approximately 19% of a complete monolayer, which reasonably corresponds to the integrin composition. Uptake of fibronectin indicates that transmembrane protein receptors are functional. For comparison, an insignificant binding signal is observed when the integrinfunctionalized lipid bilayer was formed on the nonskeletonized surface (Figure 5). Also, as a further control, a POPC lipid bilayer without the integrin on the skeletonized surface was incubated with fibronectin. As expected, the ligand does not bind the membrane surface without the integrin receptor (Supporting Information).

■

DISCUSSION Several strategies have been developed to provide space between the support and the membrane to address the problem of incorporating transmembrane proteins. A popular approach is to use polymer cushions to separate the bilayer from the support. Polyethyleneglycol (PEG) polymer is commonly used as it prevents nonspecific adsorption of proteins to supports. For example, transmembrane proteins such as annexin V and Cytochrome b5 were shown to conserve their lateral fluidity in lipid bilayers supported on PEG-polymer cushions.15,17 Tethered lipid bilayers make up another common design. Lipid membranes are attached to the support via different tethering molecules such as 2,3-di-O-phytanyl-snglycerol-1-tetraethylene glycol-D,L-α-lipoic acid ester (DPTL) or by lipopolymers. As an example, the integrin αIIbβ3 has been studied in a membrane supported via the lipopolymer tethered molecule, synthesized from 2-methyl-2oxalines with a silane group at one end and a lipid anchor on the other end.16 A peptide-tethered artificial lipid membrane system was also created to study the incorporation of integrins αvβ3 and α1β1.23 Yet a different approach uses nanoporous silica microspheres as a support for lipid bilayers, and membrane proteins were shown to be functional.20 In the work described here, the lipid membrane is separated from the support by skeletonized surfaces that provide hollow spaces for the incorporation of transmembrane proteins. Skeletonized LB films were first reported by Blodgett,39,40 taking advantage of the fact that fatty acids are much more soluble in organic solvents than are their corresponding fatty acid salts. By varying the pH and metal ion content of the

Figure 5. Binding of fibronectin to integrin α5β1-functionalized membranes supported on (a) skeletonized and (b) nonskeletonized surfaces. The SPREE plot shows the formation of the functionalized membrane followed by incubation of fibronectin with the surface. The free proteoliposomes and unbound fibronectin were removed by buffer rinsing (*). When the membrane is supported on the skeletonized surface, the signal increases upon binding of fibronectin to the integrin α5β1. The buffer was of trizma hydrochloride, sodium chloride, magnesium chloride, and calcium chloride at pH 7.4.

the skeletonized surface was exposed to a solution of integrin α5β1-POPC vesicles to form the protein-containing bilayer, and the experimental response was fit with AnalysR using a sevenlayer model based on the Fresnel equations. The optical parameters for the fit are included in Table S1 (Supporting Information). The optical thickness of the integrin-functionalized membrane over multiple experiments is 10.1 nm, which corresponds well to related measurements.23 Upon exposure to fibronectin, the surface concentration of the bound ligand is calculated according to:36 2839

dx.doi.org/10.1021/la204485n | Langmuir 2012, 28, 2835−2841

Langmuir

Article

Another difference between the skeletonized and nonskeletonized supports is that the association kinetics of the BK channel appear to be slower at the skeletonized surfaces, which is likely a consequence of the more complicated insertion process on the skeletonized film. Using a simple two-step model (eq 1), the association rate for the first step, ka1, decreases slightly at the skeletonized film, but the association rate for the second step, ka2, significantly increases. Intuitively, the first step, governed by electrostatic interactions with the bilayer interface, should be similar at the two supports. The second step, insertion into the membrane, is only significant on the skeletonized support. However, significant restructuring of the protein is expected upon insertion, including reorganization of the protein solvent sphere. The simple kinetic analysis represented by eq 1 assumes no mass change associated with the insertion step, so to the extent that SPREE is sensitive to these solvolysis changes, the insertion step may appear to lower the overall association rate. The binding kinetics are quite similar to those observed for the same proteins on the DPTL supported lipid bilayer system.49 Also, similar responses have been reported for other systems characterized by protein insertion into a bilayer. For example, insertion of the instrinsic membrane protein CyaA into polymer tethered membranes showed slower apparent kinetics than when associated with a non tethered control.51 The fusion of vesicles that have been reconstituted with a transmembrane protein represents an alternative route to protein-containing supported lipid bilayers. This approach was tested on the skeletonized supports with the integrin α5β1, using the protein’s ability to recognize fibronectin as an indicator of the viability of the supported lipid bilayer. The SPREE responses in Figure 5 are consistent with vesicle fusion on both the skeletonized and the nonskeletonized supports. However, upon exposing the supported bilayers to fibronectin, ligand binding is only observed at the bilayer on the skeletonized film. The result indicates that the integrin inserts into the bilayer and retains its binding function when supported on the skeletonized surfaces. Ligand binding assays have been used to study the functionality of other integrins. Sinner et al. confirmed the functionality of the transmembrane receptors by the binding of vitronectin and collagen type IV to the integrins αvβ3 and α1β1, respectively, by surface plasmon spectroscopy.23 Binding of the ligand vitronectin to integrin immobilized onto gold via a calix[4]crown-ether monolayer called Prolinker was also studied by SPR.36 In this case, the integrin was immobilized as a single molecular monolayer, and binding to vitronectin was observed. The behavior is quite similar to what we observe here for fibronectin binding to integrin in bilayers supported on the skeletonized surface.

subphase, metal carboxylate LB films can be deposited with different fractions of free acid. Upon rinsing the films with organic solvents, Blodgett reported skeletonized films, with tunable refractive index.39 More recently, the skeletonization process has been used for other interesting applications such as vapor sorption at LB multilayers41 and to pattern surfaces with contrasting surface properties.42 The addition of zirconium ions to a phosphonate monolayer results in cross-linking of the phosphonate molecules to form an extended zirconium phosphonate network at the surface.43 Zirconation renders the monolayer extremely stable and is not removed by either organic or aqueous washes and is stable to drying.34,44 Furthermore, the surface readily chemically binds divalent phosphate or phosphonate, while other common ligands, such as carboxylate, phosphate diester, or amines, only adsorb weakly to the surface, unable to displace the oxide and hydroxide groups that terminate the zirconium layer. The selective binding of terminal phosphate has been shown to be useful in applications that include binding of oligonucleotides for DNA arrays, and for stabilizing lipid bilayers that contain phosphatidic acid groups.29,45,46 When preparing the zirconium phosphonate monolayers, metal ion binding occurs in a separate step after monolayer transfer. Therefore, to achieve skeletonization, a mixed monolayer with octadecanol was used. After the zirconation step, the octadecanol can be washed away, leaving voids in the film. AFM analysis shows the creation of holes with diameters ranging from 80 to 700 nm across with depths of 2−3 nm. Two compositions using 10% and 20% octadecanol were selected for further study, as both proved to be viable supports for homogeneous lipid layers. Generally, transmembrane proteins can be incorporated in the bilayer by either inserting directly into the preformed lipid bilayers47 or by reconstitution into the lipid vesicles that are then fused on the substrates.48 Both mechanisms were investigated to demonstrate that the model bilayers can support membrane-bound proteins. To observe insertion into the bilayer, the BK channel protein was chosen. BK ion channels are calcium-activated potassium channels responsible for translocation of potassium across membranes.31 Structurally, BK channels comprise a pore-forming tetrameric α domain made of seven putative transmembrane segments and a β domain comprising two α-helical transmembrane domains connected by a large glycosylated extracellular loop, with intracellular amino and carboxy termini.31 In the present study, a modified BK channel was used with a C-terminal deletion at amino acid 347. An earlier study showed that the modified protein inserts into a mixture of 1,2-diphyanoyl-sn-glycero-3phosphocholine (DPhPC) and 1,2-diphytanoyl-sn-glycero-3phosphoethanolamine (DPhPE) bilayers supported on the well-established self-assembled monolayer DPTL.49,50 The affinity of the BK channel is stronger when lipid bilayers are supported on a skeletonized surface, Table 1. At the nonskeletonized support, the protein interacts with the POPC/ POPS bilayer, but it is quickly removed upon buffer rinsing, Figure 4. On the other hand, with the skeletonized supports, a significant amount of the original mass increase is retained upon rinsing. To determine if the protein is fully functional will require further physiological studies, but the result indicates that the presence of reservoirs between the support and the membrane allows the insertion of the transmembrane protein into the bilayer.

■

CONCLUSIONS Skeletonized zirconium phosphonate modified surfaces can be used to successfully support planar lipid bilayers and have been shown to be suitable for studies of transmembrane proteins. The holes in the skeletonized film provide reservoirs beneath the bilayer providing opportunity for proteins to span the bilayer. Among the potential advantages of this strategy for preparing supported lipid bilayers is that the coating can be deposited onto a wide range of surfaces, so that once a protocol for preparing a model membrane is developed, it need not be changed to accommodate the different surfaces used for different bioanalytical techniques. For example, the same procedures for preparing a lipid bilayer structure can be 2840

dx.doi.org/10.1021/la204485n | Langmuir 2012, 28, 2835−2841

Langmuir

Article

(16) Purrucker, O.; Fortig, A.; Jordan, R.; Tanaka, M. ChemPhysChem 2004, 5, 327. (17) Diaz, A. J.; Albertorio, F.; Daniel, S.; Cremer, P. S. Langmuir 2008, 24, 6820. (18) Kunding, A.; Stamou, D. J. Am. Chem. Soc. 2006, 128, 11328. (19) Shen, W. W.; Boxer, S. G.; Knoll, W.; Frank, C. W. Biomacromolecules 2001, 2, 70. (20) Davis, R. W.; Flores, A.; Barrick, T. A.; Cox, J. M.; Brozik, S. M.; Lopez, G. P.; Brozik, J. A. Langmuir 2007, 23, 3864. (21) Naumann, C. A.; Frank, C. W.; Prucker, O.; Lehmann, T.; Ruehe, J.; Knoll, W. Biophys. J. 2000, 78, 1616. (22) Schiller, S. M.; Naumann, R.; Lovejoy, K.; Kunz, H.; Knoll, W. Angew. Chem., Int. Ed. 2003, 42, 208. (23) Sinner, E. K.; Reuning, U.; Kok, F. N.; Sacca, B.; Moroder, L.; Knoll, W.; Oesterhelt, D. Anal. Biochem. 2004, 333, 216. (24) He, L. H.; Robertson, J. W. F.; Li, J.; Karcher, I.; Schiller, S. M.; Knoll, W.; Naumann, R. Langmuir 2005, 21, 11666. (25) Bunjes, N.; Schmidt, E. K.; Jonczyk, A.; Rippmann, F.; Beyer, D.; Ringsdorf, H.; Graber, P.; Knoll, W.; Naumann, R. Langmuir 1997, 13, 6188. (26) Sinner, E. K.; Knoll, W. Curr. Opin. Chem. Biol. 2001, 5, 705. (27) Terrettaz, S.; Mayer, M.; Vogel, H. Langmuir 2003, 19, 5567. (28) Sackmann, E.; Tanaka, M. Trends Biotechnol. 2000, 18, 58. (29) Oberts, B. P.; Blanchard, G. J. Langmuir 2009, 25, 2962. (30) Fabre, R. M.; Talham, D. R. Langmuir 2009, 25, 12644. (31) Ghatta, S.; Nimmagadda, D.; Xu, X. P.; O’Rourke, S. T. Pharmacol. Ther. 2006, 110, 103. (32) Zimmermann, D.; Guthohrlein, E. W.; Malesevic, M.; Sewald, K.; Wobbe, L.; Heggemann, C.; Sewald, N. ChemBioChem 2005, 6, 272. (33) Byrd, H.; Pike, J. K.; Showalter, M. L.; Whipps, S.; Talham, D. R. Interfacial Des. Chem. Sensing 1994, 561, 49. (34) Byrd, H.; Pike, J. K.; Talham, D. R. Chem. Mater. 1993, 5, 709. (35) Ma, C.; Srinivasan, M. P.; Waring, A. J.; Lehrer, R. I.; Longo, M. L.; Stroeve, P. Colloids Surf., B 2003, 28, 319. (36) Chen, H. X.; Lee, M. S.; Lee, J. B.; An, W. G.; Choi, H. J.; Kim, S. H.; Koh, K. Talanta 2008, 75, 99. (37) Corsel, J. W.; Willems, G. M.; Kop, J. M. M.; Cuypers, P. A.; Hermens, W. T. J. Colloid Interface Sci. 1986, 111, 544. (38) Jung, L. S.; Campbell, C. T.; Chinowsky, T. M.; Mar, M. N.; Yee, S. S. Langmuir 1998, 14, 5636. (39) Blodgett, K. B. J. Phys. Chem. 1937, 41, 975. (40) Blodgett, K. B.; Langmuir, I. Phys. Rev. 1937, 51, 0964. (41) Girard, K. P.; Hanley, C. M.; Vanderlick, T. K.; Quinn, J. A. Ind. Eng. Chem. Res. 2001, 40, 4283. (42) Duschl, C.; Liley, M.; Corradin, G.; Vogel, H. Biophys. J. 1994, 67, 1229. (43) Byrd, H.; Whipps, S.; Pike, J. K.; Ma, J. F.; Nagler, S. E.; Talham, D. R. J. Am. Chem. Soc. 1994, 116, 295. (44) Petruska, M. A.; Talham, D. R. Langmuir 2000, 16, 5123. (45) Monot, J.; Petit, M.; Lane, S. M.; Guisle, I.; Leger, J.; Tellier, C.; Talham, D. R.; Bujoli, B. J. Am. Chem. Soc. 2008, 130, 6243. (46) Nonglaton, G.; Benitez, I. O.; Guisle, I.; Pipelier, M.; Leger, J.; Dubreuil, D.; Tellier, C.; Talham, D. R.; Bujoli, B. J. Am. Chem. Soc. 2004, 126, 1497. (47) Salamon, Z.; Wang, Y.; Soulages, J. L.; Brown, M. F.; Tollin, G. Biophys. J. 1996, 71, 283. (48) Jeuken, L. J. C.; Connell, S. D.; Henderson, P. J. F.; Gennis, R. B.; Evans, S. D.; Bushby, R. J. J. Am. Chem. Soc. 2006, 128, 1711. (49) Okeyo, G. O. University of Florida, 2009. (50) Keizer, H. M.; Dorvel, B. R.; Andersson, M.; Fine, D.; Price, R. B.; Long, J. R.; Dodabalapur, A.; Koper, I.; Knoll, W.; Anderson, P. A. V.; Duran, R. S. ChemBioChem 2007, 8, 1246. (51) Rossi, C.; Homand, J.; Bauche, C.; Hamdi, H.; Ladant, D.; Chopineau, J. Biochemistry 2003, 42, 15273.

applied to glass for optical studies and gold for SPR or electrochemical studies. Also, the skeletonized surfaces avoid the need for specialized molecules or polymers. Furthermore, once the zirconium phosphonate surface is prepared, experience shows that they are stable for months or more, so modified slides can be kept for long periods of time and remain ready for use. In the present system, the reservoirs are 2−3 nm deep, but the fabrication protocol could be changed to adjust the depth if needed to incorporate proteins with larger extramembrane domains. Finally, although the example illustrated here uses a skeletonized Langmuir−Blodgett layer to generate the reservoirs, the concept should be general, for example extendable to other methods of preparing heterogeneous zirconium phosphonate or zirconium phosphate coatings.

■

ASSOCIATED CONTENT

S Supporting Information *

A table listing the optical parameters for the integrinfunctionalized membrane on the multilayered support, including refractive indexes, extinction coefficients, and thicknesses, and the SPREE analysis of the control experiment of fibronectin interacting with the supported lipid bilayers in the absence of integrin. This material is available free of charge via the Internet at http://pubs.acs.org.

■

AUTHOR INFORMATION

Corresponding Author

*Tel.: (352) 392-9016. E-mail: [email protected].

■

ACKNOWLEDGMENTS This material is based upon work supported by the National Science Foundation Division of Chemistry under Grant No. 0957155 (D.R.T.), cofunded by the MPS/CHE and the Office of International Science and Engineering. We thank Professor Peter A. V. Anderson of the Whitney Laboratory for Marine Bioscience helpful comments and for assistance with isolating the BK channel protein, and Professor Gail E. Fanucci for her insights and encouragement.

■

REFERENCES

(1) Gerstein, M.; Hegyi, H. FEMS Microbiol. Rev. 1998, 22, 277. (2) Muller, D. J. Biophys. J. 2006, 91, 3133. (3) Scheffer, G. L.; Maliepaard, M.; Pijnenborg, A.; van Gastelen, M. A.; de Jong, M. C.; Schroeijers, A. B.; van der Kolk, D. M.; Allen, J. D.; Ross, D. D.; van der Valk, P.; Dalton, W. S.; Schellens, J. H. M.; Scheper, R. J. Cancer Res. 2000, 60, 2589. (4) Bramhill, D. Annu. Rev. Cell Dev. Biol. 1997, 13, 395. (5) Hopkins, A. L.; Groom, C. R. Nat. Rev. Drug Discovery 2002, 1, 727. (6) Shaw, J. E.; Slade, A.; Yip, C. M. J. Am. Chem. Soc. 2003, 125, 11838. (7) Ottovaleitmannova, A.; Tien, H. T. Prog. Surf. Sci. 1992, 41, 337. (8) Sackmann, E. Science 1996, 271, 43. (9) Kam, L.; Boxer, S. G. J. Biomed. Mater. Res. 2001, 55, 487. (10) Tamm, L. K.; McConnell, H. M. Biophys. J. 1985, 47, 105. (11) Wong, J. Y.; Majewski, J.; Seitz, M.; Park, C. K.; Israelachvili, J. N.; Smith, G. S. Biophys. J. 1999, 77, 1445. (12) Goennenwein, S.; Tanaka, M.; Hu, B.; Moroder, L.; Sackmann, E. Biophys. J. 2003, 85, 646. (13) Srinivasan, M. P.; Ratto, T. V.; Stroeve, P.; Longo, M. L. Langmuir 2001, 17, 7951. (14) Keller, C. A.; Kasemo, B. Biophys. J. 1998, 75, 1397. (15) Wagner, M. L.; Tamm, L. K. Biophys. J. 2000, 79, 1400. 2841

dx.doi.org/10.1021/la204485n | Langmuir 2012, 28, 2835−2841