Lipid Membrane Domains for the Selective Adsorption and Surface

Publication Date (Web): April 1, 2013 ... In addition to randomly formed circular gel phase domains, we also show that predefined features, such as st...
1 downloads 0 Views 5MB Size
Download PDF
Article pubs.acs.org/Langmuir

Lipid Membrane Domains for the Selective Adsorption and Surface Patterning of Conjugated Polyelectrolytes Darryl Y. Sasaki,*,† Nicole Zawada,† Sean F. Gilmore,‡ Prihatha Narasimmaraj,† Mari Angelica A. Sanchez,† Jeanne C. Stachowiak,†,⊥ Carl C. Hayden,† Hsing-Lin Wang,§ Atul N. Parikh,‡ and Andrew P. Shreve∥ †

Sandia National Laboratories, Livermore, California 94550, United States University of California, Davis, Davis, California 95616, United States § Los Alamos National Laboratory, Los Alamos, New Mexico 87544, United States ∥ University of New Mexico, Albuquerque, New Mexico 87131, United States ⊥ University of Texas, Austin, Austin, Texas 78712, United States ‡

S Supporting Information *

ABSTRACT: Conjugated polyelectrolytes (CPEs) are promising materials for generating optoelectronics devices under environmentally friendly processing conditions, but challenges remain to develop methods to define lateral features for improved junction interfaces and direct optoelectronic pathways. We describe here the potential to use a bottom-up approach that employs selfassembly in lipid membranes to form structures to template the selective adsorption of CPEs. Phase separation of gel phase anionic lipids and fluid phase phosphocholine lipids allowed the formation of negatively charged domain assemblies that selectively adsorb a cationic conjugated polyelectrolyte (P2). Spectroscopic studies found the adsorption of P2 to negatively charged membranes resulted in minimal structural change of the solution phase polymer but yielded an enhancement in fluorescence intensity (∼50%) due to loss of quenching pathways. Fluorescence microscopy, dynamic light scattering, and AFM imaging were used to characterize the polymer−membrane interaction and the polymerbound domain structures of the biphasic membranes. In addition to randomly formed circular gel phase domains, we also show that predefined features, such as straight lines, can be directed to form upon etched patterns on the substrate, thus providing potential routes toward the self-organization of optoelectronic architectures.

■

INTRODUCTION

complex soft materials. Studies purportedly aimed at exploring these possibilities, however, are sparse. Creating structure in lipid membranes has been achieved in the past through top-down techniques, such as photolithography, microcontact printing, and electric fields;11,12 however, bottom-up approaches could extend the field by utilizing approaches based on self-assembly to generate defined nano- to microscale architectures. As in biology, directing the formation and organization of lipid domains is key to developing structured and functional materials. Generation of lipid domains in synthetic membrane systems is well understood as a consequence of phase separation of immiscible lipid components.13,14 For example, membranes containing a high phase transition temperature (Tg) (e.g., above room temperature) lipid in a membrane otherwise consisting of a low Tg (e.g., below room temperature) lipid will readily phase separate into gel phase domains in a fluid phase matrix. Similarly, fluid−

Selective affinity of charged macromolecules to lipid membranes is a phenomenon of broad general interest that spans from biology to materials development. In biological systems, Coulombic interactions between the cell membrane and proteins and polypeptides are thought to play important roles in the spatial organization of membrane components and the dynamic formation of lipid microdomains (e.g., lipid rafts). The latter are implicated in a variety of cellular functions, such as protein activation and regulation,1,2 antimicrobial activity,3 and endocytosis.4 For materials, electrostatic coupling of polymers to synthetic lipid membranes has been used to create novel nanocomposites enabling the development of therapeutic delivery vehicles,5−7 tethered membranes on solid substrates,8 chromatography supports,9 and sensor materials.10 These applications highlight the unique interfacial and encapsulating properties of lipid membranes. In this same vein, the ability of lipid membranes to generate defined micro- to nanoscale structures, differentiated in their electrostatic properties, in two and three dimensions should prove useful in designing novel © 2013 American Chemical Society

Received: February 1, 2013 Revised: March 26, 2013 Published: April 1, 2013 5214

dx.doi.org/10.1021/la400454c | Langmuir 2013, 29, 5214−5221

Langmuir

Article

Figure 1. Development of structural order in domain architectures for the adsorption and patterning of conjugated polyelectrolytes.

fluid immiscible phases can also be readily produced using lipid components that organize into discrete fluid phases, i.e., liquid ordered (Lo) and liquid disordered (Ld) phases.15,16 Using such domains in patterned structures to capture materials and direct their spatial organization and lateral aggregation into specific architectures offers a potential bottom-up route toward materials design. We are interested in developing lipid membranes with structured domains for the selective capture and patterning of macromolecules. Particularly interesting targets for surface patterning are conjugated polyelectrolytes (CPEs), which offer a range of optoelectronic properties in a form that allows watersoluble processing. In solution, CPEs are strongly influenced by solvent polarity and ionic strength, which alters conjugation length and aggregational state.17,18 Remarkably efficient exciton migration along the polymer chain leads to dramatic fluorescence responses to quenchers (million-fold decrease) with high sensitivity (subpicomolar range).19,20 As a result of these properties, CPEs have been exploited as “reporter” elements of optical sensors for enzymes,21,22 diseased cells,23 and small molecules.24 CPEs have also been used to fabricate light-emitting diodes25−27 and photovoltaic cells28,29 typically by solution phase deposition onto substrates. Using lipid membranes to build a hierarchy of structure into CPE materials via lateral (in-plane) or vertical (out-of-plane) organization could ultimately lead to increased efficiencies through improved definition of junction interfaces and confinement of electronic and optical propagation pathways. The first step in generating structured architectures of CPEs using lipid bilayers is achieving selective adsorption of the polymers to specific membrane domains. In the current work, we use a cationically charged polyphenylenevinylene (structure shown in Figure 1), denoted as P2,17 as our CPE and demonstrate selective binding of it to lipid domains through Coulombic interactions. Previous work by others has demonstrated that negatively charged lipids, such as phospha-

tidic acid (PA), 30 phosphatidyl glycerol (PG), 31 and phosphatidyl serine (PS),32 can phase separate into domains in phosphocholine (PC) membranes and that their structure and composition are influenced by solution pH. For instance, it has even been shown that the shapes of electrostatically differentiated domains in giant vesicles can be conveniently modulated between circles, stripes, and florets simply by switching solution pH.32 In the following we report on the optical and structural behavior of the membrane−P2 complex and demonstrate the use of negatively charged domain structures as a means to selectively adsorb CPEs into defined geometric patterns. Figure 1 shows a progression of structural order in polymer− membrane assemblies starting from uniform polymer adsorption to dispersed anionic lipids in a fluid phase membrane, then confinement of the polymer into domains randomly formed in the membrane, and ultimately into oriented pathways defined by patterned lipid domains. Spectroscopic (fluorescence, UV− vis), dynamic light scattering (DLS), and atomic force microscopy (AFM) data suggest tight and uniform coverage of P2 onto the negatively charged regions of the membranes. Fluorescence microscopy of giant vesicles and supported lipid bilayers (SLBs) demonstrate the selective affinity for the CPEs to domains enriched in negatively charged, gel phase domains. Formation of domains into defined architectures was achieved through directed assembly of the gel phase domains that followed etched patterns placed on the substrate surface.

■

EXPERIMENTAL METHODS

General. Aqueous solutions were prepared from deionized (DI) water obtained through a Barnstead Type D4700 NANOpure Analytical Deionization System with ORGANICfree cartridge registering ≥18.0 MΩ·cm resistance. Phospholipids [1-palmitoyl-2oleoyl-sn-glycerophosphatidylcholine (POPC), 1,2-dihexadecanoyl-snglycero-3-phosphate (sodium salt) (DPPA), 1,2-dioleoyl-sn-glycero-3phosphate (DOPA), 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC), 1,2-dipalmitoyl-sn-glycero-2-phospho(1′-rac-glycerol) (so5215

dx.doi.org/10.1021/la400454c | Langmuir 2013, 29, 5214−5221

Langmuir

Article

coverslip by vesicle fusion.37 Thermal annealing was achieved by placing the SLB-adhered coverslip on a hot plate set at 45 °C for 2 min. The SLBs were imaged with a 60× oil immersion objective. P2 affinity studies were done by exposing the SLB to concentrations of P2 for 2 min, followed by removing excess liquid from the solution well and rinsing the SLB at least seven times with fresh MOPS buffer solution. Detergent treatment of the SLB−P2 complex was performed by adding 10% Triton X-100 solution (17 μL) to the solution volume (170 μL) in the well. Atomic Force Microscopy. SLBs for AFM were prepared on 20 mm × 20 mm silicon substrates that were previously cleaned using piranha solution using the same procedure as above. Measurements were taken with a Veeco Dimension 3100 (Veeco Metrology, Santa Barbara, CA). A silicon nitride tip, mounted to an in-liquid tip holder, with a spring constant of 0.03 N/m (Veeco Metrology, Santa Barbara, CA) was used for scanning in contact mode. After the tip was mounted, the tip and wet cell were lowered into the MOPS buffer and allowed to sit in solution for 10 min prior to the first scan. For P2treated samples the supported bilayers were exposed to P2 at a concentration of 3 μM, followed by successive washing to remove unbound polymer.

dium salt) (DPPG), 1,2-dioleoyl-sn-glycero-2-phosph(1′-rac-glycerol) (sodium salt) (DOPG)] were purchased from Avanti Polar Lipids (Alabaster, AL). The fluorescent probe β-BODIPY 530/550 C5-HPC was purchased from Invitrogen (Grand Island, NY) and Nile Red from Sigma-Aldrich (St. Louis, MO). Cationic polymer P2 (MW ∼ 10 000) was prepared as described previously.17 All solvents and reagents were purchased from either Fisher Scientific (Pittsburgh, PA) or SigmaAldrich. MOPS buffer (3-(N-morpholino)propanesulfonic acid) (20 mM), NaCl (100 mM)) was adjusted to pH 7.4 using 10% aqueous sodium hydroxide solution. Fluorescence spectra were obtained using a HORIBA Jobin Yvon SPEX Fluoromax 3 (Edison, NJ), UV−vis spectra using a HewlettPackard 8453 (Santa Clara, CA), and dynamic light scattering (DLS) using a Protein Solutions DynaPro LSR (Wyatt Technology, Santa Barbara, CA). Fluorescence imaging of giant vesicles was performed on a Zeiss Axiovert 200 M (Thornwood, NY) inverted microscope, and the imaging of supported lipid bilayers was performed on an Olympus IX71 inverted microscope; both microscopes were equipped with Andor iXon+ CCD cameras (South Windsor, CT). Two sets of filters (Chroma Technology Corp.) were used to image the bilayer and P2 fluorescence: TRITC filter (543(22x)/593(40m), dichroic 562 long pass) for the BODIPY 530/550 HPC and Nile Red, and GFP filter (475(28x)/520(28m), dichroic 495 long pass) for P2 fluorescence. Image processing was done with ImageJ software (National Institutes of Health). As a reference of fluorescence intensity between the GFP and TRITC channels for Figures 5 and 6, the BODIPY 530/550 HPC labeled membrane is ∼50% lower in intensity compared to the intensity in the regions where P2 is bound. Small and Large Unilamellar Vesicles. Small unilamellar vesicles (SUVs) were prepared by probe sonication as described previously33 (40−60 nm diameter) for SLB preparation and large unilamellar vesicles (LUVs), used for spectroscopic studies, were formed by extrusion. Lipid films were prepared from stock solutions of DOPA, DOPG, POPC, and DPPG in chloroform and DPPA in 50% methanol/chloroform. For SUVs used for SLB preparation and for GUVs (see below), 0.03% and 0.1% BODIPY 530/550 HPC was added, respectively, for imaging. For the extruded LUVs, dried lipid films coating the inside of glass conical tubes were hydrated in MOPS buffer and incubated at ∼70 °C for 2 h, followed with gentle vortex stirring, and then the solution was passed 21 times through a stack of two 100 nm pore polycarbonate filters at ∼60 °C. All liposome solutions were used within a week of preparation. All spectroscopic measurements were done with the LUVs. For UV−vis spectroscopy and DLS measurements, P2 concentration was set at 20 μM (per monomer unit) and vesicles were at a nominal concentration of 100 μM total lipid to ensure good signal in the measurements. For the fluorescence studies, P2 concentration was 0.3−3.0 μM with a nominal lipid concentration of 100 μM. Percent error averaged over three trials yielded an SD of ∼10%. Giant Vesicles. Giant unilamellar vesicles (GUVs) were formed via electroformation34,35 in sucrose solution (∼350 mOsm) at elevated temperature (70 °C) to ensure fluidity of all lipid components. Samples of the GUVs were diluted in glucose solution (∼350 mOsm) and placed in fluidic channels for imaging under a wide-field fluorescence microscope with a 100× oil immersion objective. The fluidic channels were assembled using two parallel sets of double-sided tape spaced 2−3 mm apart between a No. 1.5 coverslip (Corning Inc.) and a glass slide. Supported Lipid Bilayers (SLB). SLBs were prepared as described previously36 on glass coverslips and imaged on the Olympus microscope. The coverslips (etched and nonetched) were cleaned in soapy water (Neutrad, Decon Laboratories, Inc.), rinsed with DI water, followed by piranha etch cleaning (25% (30% H2O2)/H2SO4) for 30 min at ∼100 °C (caution: this mixture reacts violently with organic materials and must be handled with extreme care), and a final rinsing with copious amounts of DI water. The cleaned coverslips were then stored in DI water. Etched coverslips were prepared prior to cleaning by using a ruler to scribe a straight line on the coverslip surface with various sharp tools (i.e., microforceps tip, copper wire (18 gauge), 10 μL syringe needle tip (Hamilton Co.)). SLBs were formed on the glass

■

RESULTS We prepared vesicles of POPC membranes incorporating one of the four anionically charged lipids: DPPA, DPPG, DOPA, and DOPG. At neutral pH and room temperature, the low phase transition temperature (Tg) lipids, DOPA (Tg = −8 °C) and DOPG (Tg = −18 °C), should homogeneously disperse in the fluid phase matrix of the POPC (Tg = −2 °C) membrane, while the gel phase lipids DPPA and DPPG, with their higher Tg (67 and 55 °C, respectively), are expected to phase separate due to favorable packing interaction of the lipid tails in spite of the repulsive electrostatic interactions at the headgroup position.30,32 The optical behavior of P2 in the presence of these negatively charged lipid membranes was studied with LUVs in MOPS buffer at pH 7.4. At this pH, P2 absorbs at λabs max 17 = 446 nm and fluoresces at λem max = 525 nm. In the presence of vesicles of electrostatically neutral POPC or phase separated 10−20% DPPC (Tg = 55 °C) in POPC membranes, P2 exhibits no change in absorption or fluorescence properties. With vesicles containing 5−20% mole fraction of anionic lipid (DPPA, DOPA, DPPG, DOPG) there is also no detectable change in UV−vis absorption of P2. This result contrasts with the shift in absorption that occurs for P2 in the presence of the anionic surfactant sodium dodecyl sulfate (SDS).38 P2 fluorescence intensity, on the other hand, increased significantly with a slight bathochromic shift (Figure S1). Figure 2 shows the relationship between mole fraction of the negatively charged lipid in the membrane and fluorescence intensity of P2. A maximum of ∼50% increase is observed at a dopant level of 10%, and no significant increase in fluorescence intensity is observed beyond this concentration. The fluorescence behavior of P2 in the presence of membranes prepared with either gel or fluid phase anionic lipids is similar. The relatively small fluorescence enhancement observed here, compared to other CPE−lipid systems,10,21,39 is believed to be due, in part, to the low molecular weight of P2 (∼10 000) compared to other CPEs, such as MPS−PPV (>100 000).19 Structural changes of the LUVs as a result of P2 binding were evaluated with dynamic light scattering (DLS). It is known that the electrostatic adsorption of polyelectrolytes can induce changes in membrane curvature,40 leading to budding and tubule formation. For the control vesicles of 100% POPC and in vesicles containing the negatively charged lipids DOPA, DOPG, or DPPG at 10 mol %, we find no evidence of change 5216

dx.doi.org/10.1021/la400454c | Langmuir 2013, 29, 5214−5221

Langmuir

Article

Figure 3. Domain ripening of 20% DPPA/POPC SLB (0.03% BODIPY 530/550 HPC) on a glass coverslip by thermal incubation. Fluorescence images show the SLB (A) immediately after formation by vesicle fusion and (B) after incubation at 45 °C for 2 min, followed by cooling to room temperature. (TRITC filter; image size = 135 μm).

Figure 2. Fluorescence emission intensity of P2 (3.0 μM) (λex = 450 nm) measured at wavelength maximum in the presence of POPC vesicles doped with anionic lipids as a function of their mole fraction in the membrane.

As in the GUV samples, the size and frequency of the domains increase with higher mole fraction of the gel phase lipids. The domain structures were further characterized by topographic imaging and fluorescence studies. Figure 4 shows

in the LUV size (Rh = 50 nm) in the presence of P2 at the concentrations examined (20 μM) (Table S1). However, for DPPA containing vesicles, P2 addition leads to the growth of a population of particles with Rh = 77 nm, suggesting a possible change in morphology or dimerization of LUVs. In studies with giant liposomes we can distinguish the formation of micrometer size domains. Electroformed GUVs containing gel phase lipids exhibit darkened domains depleted of BODIPY 530/550 HPC (Figures S2), a membrane dye known to partition away from gel phase domains and into the fluid phase.36 The domain sizes generally correspond to the mole fraction of the gel phase lipids and selective adsorption of P2 onto those domains of DPPA and DPPG in POPC membranes was observed (Figures S2) using fluorescence microscopy. DOPA and DOPG containing vesicles exhibited homogeneous P2 fluorescence with no detectable domain formation. In the domain forming systems, the size of the P2bound domains often appears larger than the darkened domains observed in the absence of P2. This increase in domain size suggests polymer-induced domain formation.40 For the DPPA/ POPC vesicles, a fraction of the population was also found as dimers with P2 concentrated at the interfacial regions (Figure S3). SLBs (doped with 0.03% BODIPY 530/550 HPC) were prepared by vesicle fusion onto glass coverslips and imaged via fluorescence microscopy using the TRITC filter. Bilayers of POPC containing DOPA or DOPG (5−20 mol %) were featureless as characterized by the homogeneous fluorescence observed throughout the sample. Bilayers containing DPPA and DPPG (5−20 mol %), on the other hand, generated phaseseparated domains indicated by the presence of dark patches depleted of membrane dye. Figure 3 shows an SLB of 20% DPPA/POPC after initial formation via vesicle fusion and then following incubation at elevated temperature. Initially, the bilayer contains numerous dark domains ≤1 μm in diameter near the resolution limits of the fluorescence microscope, but after heating the sample for 2 min at 45 °C, the domains grow into larger 4−6 μm structures through an apparent ripening process.41 Under the TRITC filter, the fluorescence intensity of the dark domains is roughly half that emitted by the fluid phase, suggesting that the domains are enriched in the gel phase lipid.

Figure 4. AFM image (top) and height profile (bottom) (line trace shown in topographic image) of 20% DPPA/POPC SLB on a silicon wafer. Height difference between gel phase domains and the surrounding fluid phase membrane is ∼1.2 nm.

an AFM image of a supported membrane of 20% DPPA/POPC on an atomically flat silicon wafer with a domain architecture that suggests the existence of gel phase domains in the fluid phase matrix. The domains are ∼1.2 nm taller than the surrounding membrane, which is consistent with other measurements of gel phase domains in fluid phase membranes (e.g., DPPC/POPC,42 DPPC/DOPC41). The size and shape of 5217

dx.doi.org/10.1021/la400454c | Langmuir 2013, 29, 5214−5221

Langmuir

Article

Figure 5. Fluorescence images show the selective affinity of P2 to gel phase domains of the 20% DPPA/POPC SLB. The SLB was exposed to P2 (3 μM), rinsed, and then imaged under (A) TRITC filter to reveal the dye depleted gel phase domains and (B) GFP filter to observe membrane adsorbed P2. Image C shows the merged images (image size = 135 μm).

Figure 6. Defined architectures of membrane-adsorbed P2 as directed through an etch on the surface created by scribing with a microforceps tip. Fluorescence images of 20% DPPA/POPC SLB on the scribed glass surface following exposure to P2 (3 μM) and imaged under (A) TRITC filter showing the defined gel phase domain and (B) GFP filter showing membrane adsorbed P2. Image C shows the merged images (image size = 135 μm).

the domains were also consistent with those observed by fluorescence microscopy (Figure S4). Upon addition of P2 (3.0 μM), no detectable change in domain height or alterations of domain size or shape was found. Further evidence of the physical state of the domains was obtained by using the preferential partitioning of Nile Red into ordered lipid phases.43 With SLBs of 20% DPPA/POPC doped with 0.1% Nile Red we observed that the domains were brightly fluorescent with the dye relative to the surrounding membrane (Figure S5). The gel phase domains enriched in anionic lipid in the SLBs were found to be highly selective for adsorption of cationic P2. Figure 5 shows fluorescence images of a 20% DPPA/POPC SLB exposed to P2 (3 μM) for 2 min, followed by successive rinsing with MOPS buffer to remove unbound polymer. The dark domains enriched in gel phase DPPA (TRITC filter) were selective in adsorbing P2 (GFP filter) on the membrane surface. Similar domain structures with P2 affinity were also observed with SLBs of DPPG/POPC (data not shown). In control experiments, P2 did not exhibit any affinity to electrostatically neutral SLBs containing POPC alone or phase-separated 20% DPPC/POPC membranes. SLBs of DOPA/POPC, which were void of domains at all mole fractions tested in the absence of P2, did form what appeared to be ∼1 μm size domains at high mole fractions (20%) upon exposure to P2 (Figure S6). Such behavior is consistent with polymer-induced domain formation. P2 bound domains in DPPA/POPC SLBs were also found to be stable to detergent solutions (1% Triton X-100) that readily removed the surrounding lipid membrane leaving only the membrane bound P2 on the substrate (Figure S7). In an effort to provide structural definition to the domain architecture, we took advantage of a serendipitous observation. Typically, the gel phase domains were circular as they grow isotropically in the plane of the bilayer. However, stripelike

domain features were occasionally found in the supported membrane (Figure S8). Just like the circular domains, these stripes also exhibited selective affinity for P2. Since the glass coverslips were thoroughly cleaned of surface-adhered contaminants, it was presumed that these structured domains formed in response to inherent defects in the substrate, such as scratches on the surface. Not knowing what types of defects might induce the domain formation we tested this idea by preparing glass coverslips etched with different scribing materials. Figure 6 shows an image of a stripelike domain that formed over a line etched in glass by the tip of a microforceps, along with the domain’s selective affinity of P2. The continuous domain extends millimeters in length with a width of ∼4 μm. The process seems to be quite general as stripes were formed using the other scribing materials as well (Figure S9). These linear P2-bound structures were similarly resistant to Triton X-100, as described earlier with the circular domains, permitting selective removal of peripheral membrane. These structured linear domains also behaved similarly to the undefined circular domains regarding fluorescence probe partitioning (i.e., BODIPY 530/550 HPC and Nile Red), thus identifying them as lipid domains rather than open voids in the membrane.

■

DISCUSSION The interaction of P2 to negatively charged lipid membranes produced a significant change in the polymer’s optical properties, but with marked differences from that observed previously with negatively charged surfactant SDS.44,18 In the case of SDS, there is considerable shift in the absorption wavelength maximum of P2 and large swings in fluorescence intensity with SDS concentration.38,45 It has been proposed that the binding of negatively charged SDS to cationic P2 reduces the twisting of the polymer’s backbone, thus extending 5218

dx.doi.org/10.1021/la400454c | Langmuir 2013, 29, 5214−5221

Langmuir

Article

chemical characteristics that were consistent with gel phase domains enriched in negative charge. Following vesicle fusion of SUVs to the glass substrate, the domains in the formed SLBs were initially small but could be enlarged through a thermal ripening process. The phase partitioning characteristics of two membrane probes, one that partitions into ordered lipid phases (Nile Red) and the other that partitions into disordered regions of the membrane (BODIPY 530/550 HPC), suggests the domains to be in the gel phase. Topographic images of the biphasic SLB by AFM further supported this conclusion, and selective affinity of the polycationic P2 for those domains, as seen observed in Figure 5 and Figure S4, affirmed their chemistry as regions enriched in anionic lipids. We found that the structure of the membrane domains, and subsequently the surface patterning of P2, could be defined through etches made on the substrate surface. Scratching techniques have been successfully used to define membrane architectures in the past46 but were used as a way to remove material rather than define the spontaneous formation of a domain structure. Since in our experiments on biphasic SLBs on various substrates (e.g., glass, quartz, silicon) many nonrandom domain structures were observed, it became apparent that these structures result from interactions of the membrane with defects in the substrate. We demonstrated that by etching the glass substrate with sharp tools (e.g., microforceps tip, syringe needle tip, 18 gauge wire) we can direct the generation of anionic gel phase domains into scribed features (i.e., lines) and that P2 adsorption faithfully follows the pattern (Figure 6 and Figure S9). The precise interaction between the membrane and the etched surface that drives the domain structure is at present unknown. Efforts to understand the structural and chemical forces on etched substrates that may be playing roles in driving the nucleation and structural confinement of gel phase domains in SLBs are currently underway in our laboratories.

the conjugation length that results in a shift of absorption from 445 nm in the absence of SDS to 482 nm at 100 μM SDS.38 Simultaneously, through the concentration range of 0−100 μM SDS, the polymer’s fluorescence quantum yield drops by more than 80% in value due to aggregation of the surfactantcomplexed polymer chains. At higher concentrations of SDS (above the critical micelle concentration), the absorption shifts back to shorter wavelength while the quantum yield increases by more than an order of magnitude as the micelles decrease polymer aggregation and increase the rigidity of the bound polymer, thereby reducing vibrational relaxation pathways. Compared to studies with SDS, negatively charged liposomes exhibit less influence over the structural properties of P2. Interaction with the negatively charged vesicles, for example, produced essentially no change in the spectroscopic absorption behavior of P2, indicating minimal structural change of the polymer upon adsorption to the membrane. Fluorescence intensity of membrane-bound P2, on the other hand, exhibited an increase of up to ∼50%. This suggests a strong polymer− membrane complex, in a similar vein with SDS micelles. The fluorescence emission shape suggests a single fluorescing species in the presence of liposomes (Figure S1), which differs from the multiple emission peaks with SDS.38 While adsorption to the membrane clearly enhances the fluorescence of the polymer, the observed increase in fluorescence intensity as a function of anionic lipid concentration in the membrane (Figure 2) may be associated with the degree of polymer− membrane coupling. The cationic polymer chains may bind only partially at low densities of negative charge on the membranes but more fully at higher densities, resulting in further losses of quenching pathways. In general, the interaction of P2 on the negatively charged lipid membranes had little effect on membrane morphology or the aggregation of vesicles. A lack of change in vesicle size, as determined by DLS, and minimal change in domain height upon the adsorption of P2, determined by AFM, suggests a tight and uniform coverage of the polymer to the negatively charged membrane surface. The only exception occurred with the DPPA containing membranes, where it was observed that a new population of particles ∼50% larger in size formed upon the presence of P2. Fluorescence images of DPPA/POPC GUVs, which find dimerized vesicles with their interfaces concentrated in adsorbed P2, may provide some insight into the source of this particle formation. Of the four anionic lipids used in this study, there are two important considerations regarding their electrostatic interactions with P2: headgroup charge and charge density in the membrane. While the PA lipids bear two negative charges at the pH of the experiment, the PG lipids carry a single negative charge. The density of charge in the membrane is determined by the nominal mole fraction and dispersity of the anionic lipids. Local charge density, on the other hand, will be higher in the phaseseparated structures compared to the homogeneously dispersed membranes. The combination of the double negative charge and high local concentration of DPPA in lipid domains creates a uniquely high anionic charge density that may facilitate the sandwiching of P2 between opposing vesicles. While the process resulting in the dimerized vesicles is not fully understood, this phenomenon could provide a route toward building out of plane structures via CPE−membrane complexes. In the fluorescence images of the SLBs containing anionic lipids of high Tg we observed domains having physical and

■

CONCLUSION Our results demonstrate that lipid domains in the membrane can be used to create definition of structure to surface adsorbed CPEs. Adsorption of CPEs to homogeneously dispersed negative charges on the lipid membrane provides the first level of confinement from the three-dimensional space of bulk solution to the two-dimensional surface of the membrane. This relatively straightforward complexation has led to new materials advancements in sensing and biomembrane models. Through self-assembly of lipid components in the membrane, we found it possible to further confine and organize adsorbed CPEs into discrete structures for potential future use in optoelectronic materials. Adsorption of the CPE into the confined environment of the membrane domain did not impede enhancement of optical behavior of the membrane-bound polymer, and the complexation resulted in a stable assembly that was resistant to detergent treatment. We are currently investigating the possibilities and limitations of building complex nanoscale patterns through directed assembly of lipid domains and the organization of multiple components into defined architectures using chemical recognition to drive the assembly.

■

ASSOCIATED CONTENT

S Supporting Information *

Images, tables, and graphs of the P2-membrane complexed systems. This material is available free of charge via the Internet at http://pubs.acs.org. 5219

dx.doi.org/10.1021/la400454c | Langmuir 2013, 29, 5214−5221

Langmuir

■

Article

(17) Gao, Y.; Wang, C.-C.; Wang, L.; Wang, H.-L. Conjugated polyelectrolytes with pH-dependent conformations and optical properties. Langmuir 2007, 23, 7760−7767. (18) Kaur, P.; Yue, H.; Wu, M.; Liu, M.; Treece, J.; Waldeck, D. H.; Xue, C.; Liu, H. Solvation and aggregation of polyphenylethynylene based anionic polyelectrolytes in dilute solution. J. Phys. Chem. B 2007, 111, 8589−8596. (19) Chen, L.; McBranch, D. W.; Wang, H.-L.; Helgeson, R.; Wudl, F.; Whitten, D. G. Highly sensitive biological and chemical sensors based on reversible fluorescence quenching in a conjugated polymer. Proc. Natl. Acad. Sci. U. S. A. 1999, 96, 12287−12292. (20) Heeger, P. S.; Heeger, A. J. Making sense of polymer-based biosensors. Proc. Natl. Acad. Sci. U. S. A. 1999, 96, 12219−12221. (21) Liu, Y.; Ogawa, K.; Schanze, K. S. Conjugated polyelectrolyte based real-time fluorescence assay for phospholipase C. Anal. Chem. 2008, 80, 150−158. (22) Chemburu, S.; Ji, E.; Casana, Y.; Wu, Y.; Buranda, T.; Schanze, K. S.; Lopez, G. P.; Whitten, D. G. Conjugated polyelectrolyte supported bead based assays for phospholipase A2 activity. J. Phys. Chem. B 2008, 112, 14492−14499. (23) Bajaj, A.; Miranda, O. R.; Kim, I.-B.; Phillips, R. L.; Jerry, D. J.; Bunz, U. H. F.; Rotello, V. M. Detection and differentiation of normal, cancerous, and metastatic cells using nanoparticle-polymer sensor arrays. Proc. Natl. Acad. Sci. U. S. A. 2009, 106, 10912−10916. (24) Zhao, X.; Liu, Y.; Schanze, K. S. A conjugated polyelectrolytebased fluorescence sensor for pyrophosphate. Chem. Commun. 2001, 2914−2916. (25) Ma, W.; Iyer, P. K.; Gong, X.; Liu, B.; Moses, D.; Bazan, G. C.; Heeger, A. J. Water/methanol-soluble conjugated copolymer as an electron-transport layer in polymer light-emitting diodes. Adv. Mater. 2005, 17, 274−277. (26) Huang, F.; Hou, L.; Wu, H.; Wang, X.; Shen, H.; Cao, W.; Yang, W.; Cao, Y. High-efficiency, environmentally-friendly electroluminescent polymers with stable high work function metal as a cathode: green- and yellow-emitting conjugated polyfluorene polyelectrolytes and their neutral precursors. J. Am. Chem. Soc. 2004, 126, 9845−9853. (27) Baur, J. W.; Kim, S.; Balanda, P. B.; Reynolds, J. R.; Rubner, M. F. Thin-film light-emitting devices based on sequentially adsorbed multilayers of water-soluble poly(p-phenylene)s. Adv. Mater. 1998, 10, 1452−1455. (28) Mwaura, J. K.; Pinto, M. R.; Witker, D.; Ananthakrishnan, N.; Schanze, K. S.; Reynolds, J. R. Photovoltaic cells based on sequentially adsorbed multilayers of conjugated poly(p-phenylene ethynylene)s and a water-soluble fullerene derivative. Langmuir 2005, 21, 10119− 10126. (29) Mwaura, J. K.; Zhao, X.; Jiang, H.; Schanze, K. S.; Reynolds, J. R. Spectral broadening in nanocrystalline TiO2 solar cells based on poly(p-phenylene ethynylene) and polythiophene sensitizers. Chem. Mater. 2006, 18, 6109−6111. (30) Garidel, P.; Johann, C.; Blume, A. Non-ideal mixing and fluidfluid immiscibility in phosphatidic acid-phosphatidylethanolamine mixed bilayers. Eur. Biophys. J. 2011, 40, 891−905. (31) Garidel, P.; Johann, C.; Mennicke, L.; Blume, A. The mixing behavior of pseudobinary phosphatidylcholine-phosphatidylglycerol mixtures as a function of pH and chain length. Eur. Biophys. J. 1997, 26, 447−459. (32) Bandekar, A.; Sofou, S. Floret-shaped solid domains on giant fluid lipid vesicles induced by pH. Langmuir 2012, 28, 4113−4122. (33) Last, J. A.; Waggoner, T. A.; Sasaki, D. Y. Lipid membrane reorganization induced by chemical recognition. Biophy. J. 2001, 81, 2737−2742. (34) Angelova, M. I.; Dimitrov, D. S. Liposome electroformation. Faraday Discuss. Chem. Soc. 1986, 81, 303−311. (35) Stachowiak, J. C.; Hayden, C. C.; Sasaki, D. Y. Steric confinement of proteins on lipid membranes can drive curvature and tubulation. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 7781−7786. (36) Hayden, C. C.; Hwang, J. S.; Abate, E. A.; Kent, M. S.; Sasaki, D. Y. Directed formation of lipid membrane microdomains as high

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS The authors thank Dr. Jennifer Martinez for her insightful comments in the manuscript preparation and Dr. Julie Last for her guidance on the nanoscale imaging of the SLBs. This work was supported by the US Department of Energy, Office of Basic Energy Sciences, Division of Materials Science and Engineering. Sandia National Laboratories is a multiprogram laboratory managed and operated by Sandia Corporation, a wholly owned subsidiary of Lockheed Martin Corporation, for the U.S. Department of Energy’s National Nuclear Security Administration under Contract DE-AC04-94AL85000.

■

REFERENCES

(1) Lemmon, M. A. Membrane recognition by phospholipid-binding domains. Nat. Rev. 2008, 9, 99−111. (2) McLaughlin, S.; Murray, D. Plasma membrane phosphoinositide organization by protein electrostatics. Nature 2005, 438, 605−611. (3) Epand, R. M.; Epand, R. F. Lipid domains in bacterial membranes and the action of antimicrobial agents. Biochim. Biophys. Acta 2009, 1788, 289−294. (4) Blood, P. D.; Voth, G. A. Direct observation of Bin/ amphiphysin/Rvs (BAR) domain-induced membrane curvature by means of molecular dynamics simulations. Proc. Natl. Acad. Sci. U. S. A. 2006, 103, 15068−15072. (5) Tong, A. W.; Jay, C. M.; Senzer, N.; Maples, P. B.; Nemunaitis, J. Systematic therapeutic gene delivery for cancer: crafting Paris’ arrow. Curr. Gene Therapy 2009, 9, 45−60. (6) Ewert, K. K.; Ahmad, A.; Bouxsein, N. F.; Evans, H. M.; Safinya, C. R. Non-viral gene delivery with cationic liposome-DNA complexes. Methods Mol. Biol. 2008, 433, 159−175. (7) Yessine, M.-A.; Leroux, J.-C. Membrane-destabilizing polyanions: interaction with lipid bilayers and endosomal escape of biomacromolecules. Adv. Drug Delivery Rev. 2004, 56, 999−1021. (8) Knoll, W.; Bender, K.; Förch, R.; Frank, C.; Götz, H.; Heibel, C.; Jenkins, T.; Jonas, U.; Kibrom, A.; Kügler, R.; Naumann, C.; Naumann, R.; Reisinger, A.; Rühe, J.; Schiller, S.; Sinner, E.-K. Polymer-tethered biomolecular lipid membranes. Adv. Polym. Sci. 2010, 224, 87−111. (9) Loidl-Stahlhofen, A.; Kaufmann, S.; Braunschweig, T.; Bayerl, T. M. The thermodynamic control of protein binding to lipid bilayers for protein chromatography. Nat. Biotechnol. 1996, 14, 999−1002. (10) Ngo, A. T.; Karam, P.; Cosa, G. Conjugated polyelectrolyte-lipid interactions: Opportunities in biosensing. Pure Appl. Chem. 2011, 83, 43−55. (11) Oliver, A. E.; Parikh, A. N. Templating membrane assembly, structure, and dynamics using engineered interfaces. Biochim. Biophys. Acta 2010, 1798, 839−850. (12) Tanaka, M.; Sackmann, E. Polymer-supported membranes as models of the cell surface. Nature 2005, 437, 656−663. (13) Marsh, D. Thermodynamics of phospholipid self-assembly. Biophys. J. 2012, 102, 1079−1087. (14) Almeida, P. F. F. Thermodynamics of lipid interactions in complex bilayers. Biochim. Biophys. Acta 2009, 1788, 72−85. (15) Honerkamp-Smith, A. R.; Veatch, S. L.; Keller, S. L. An introduction to critical points for biophysicists; observations of compositional heterogeneity in lipid membranes. Biochim. Biophys. Acta 2009, 1788, 53−63. (16) van Meer, G.; Voelker, D. R.; Feigenson, G. W. Membrane lipids: where they are and how they behave. Nat. Rev. 2008, 9, 112− 124. 5220

dx.doi.org/10.1021/la400454c | Langmuir 2013, 29, 5214−5221

Langmuir

Article

affinity sites for His-tagged proteins. J. Am. Chem. Soc. 2009, 131, 8728−8729. (37) Anderson, T. H.; Min, Y.; Weirich, K. L.; Zeng, H.; Fygenson, D.; Israelachvili, J. N. Formation of supported bilayers on silica substrates. Langmuir 2009, 12, 6997−7005. (38) Treger, J. S.; Ma, V. Y.; Gao, Y.; Wang, C.-C.; Wang, H.-L.; Johal, M. S. Tuning the optical properties of a water-soluble cationic poly(p-phenylenevinylene):surfactant complexation with a conjugated polyelectrolyte. J. Phys. Chem. B 2008, 112, 760−763. (39) Ngo, A. T.; Cosa, G. Assembly of zwitterionic phospholipid/ conjugated polyelectrolyte complexes: Structure and photophysical properties. Langmuir 2010, 26, 6746−6754. (40) Tribet, C.; Vial, F. Flexible macromolecules attached to lipid bilayers: impact on fluidity, curvature, permeability and stability of the membranes. Soft Matter 2008, 4, 68−81. (41) Giocondi, M.-C.; Vié, V.; Lesniewska, E.; Milhiet, P.-E.; ZinkeAllmang, M.; Le Grimellec, C. Phase topology and growth of single domains in lipid bilayers. Langmuir 2001, 17, 1653−1659. (42) Rinia, H. A.; de Kruijff, B. Imaging domains in model membranes with atomic force microscopy. FEBS Lett. 2001, 504, 194−199. (43) Gao, F.; Mei, E.; Lim, M.; Hochstrasser, R. M. Probing lipid vesicles by bimolecular association and dissociation trajectories of single molecules. J. Am. Chem. Soc. 2006, 128, 4814−4822. (44) Chen, L.; Xu, S.; McBranch, D.; Whitten, D. Tuning the properties of conjugated polyelectrolytes through surfactant complexation. J. Am. Chem. Soc. 2000, 122, 9302−9303. (45) Treger, J. S.; Ma, V. Y.; Gao, Y.; Wang, C.-C.; Jeon, S.; Robinson, J. M.; Wang, H.-L.; Johal, M. S. Controlling layer thickness and photostability of water-soluble cationic poly(p-phenylenevinylene) in multilayer thin films by surfactant complexation. Langmuir 2008, 24, 13127−13131. (46) Cremer, P. S.; Groves, J. T.; Kung, L. A.; Boxer, S. G. Writing and erasing barriers to lateral mobility into fluid phospholipid bilayers. Langmuir 1999, 15, 3893−3896.

5221

dx.doi.org/10.1021/la400454c | Langmuir 2013, 29, 5214−5221