Biomolecular Interactions at Phospholipid-Decorated Surfaces of Liquid Crystals

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Science  19 Dec 2003:
Vol. 302, Issue 5653, pp. 2094-2097
DOI: 10.1126/science.1091749


The spontaneous assembly of phospholipids at planar interfaces between thermotropic liquid crystals and aqueous phases gives rise to patterned orientations of the liquid crystals that reflect the spatial and temporal organization of the phospholipids. Strong and weak specific-binding events involving proteins at these interfaces drive the reorganization of the phospholipids and trigger orientational transitions in the liquid crystals. Because these interfaces are fluid, processes involving the lateral organization of proteins (such as the formation of protein- and phospholipid-rich domains) are also readily imaged by the orientational response of the liquid crystal, as are stereospecific enzymatic events. These results provide principles for label-free monitoring of aqueous streams for molecular and biomolecular species without the need for complex instrumentation.

Some of the most important biomolecular interactions occur at biological membranes, including the binding events that permit entry of protein toxins into cells (1), the binding and enzymatic events that trigger cell signaling pathways (2), the assembly of proteins, lipids, and cholesterol into rafts (3), the crystallization of proteins (4), and the binding events that are the first stage of viral infection (5). Past attempts to provide facile methods of reporting these biomolecular interactions (e.g., for biological sensing) have exploited the self-assembly of the constituents of biological membranes such as phospholipids and proteins at interfaces (111). These systems, however, have been difficult to analyze because they generally require the use of either labeled molecules (e.g., fluorescent labels) or complex instrumentation.

The work reported here was inspired by the observation that most biomolecular interactions at biological membranes are accompanied by a reorganization of the proteins, lipids, and other species that constitute the membranes (111). We report that fluid, phospholipid assemblies formed at interfaces between thermotropic liquid crystals (LCs) and aqueous phases are coupled to the orientations of the thermotropic LCs. The specific binding of proteins to these interfaces and their subsequent formation of organized lateral assemblies, as well as the activities of enzymes, are demonstrated to trigger spatially patterned orientational transitions in the LCs that are readily imaged with polarized light. This coupling permits label-free imaging of a range of dynamic molecular phenomena that occur at these interfaces (12, 13). Because aqueous streams can also flow past the lipid-laden interface of the LC, these principles may also provide the basis of low-cost, passive (zero-power) indicators of the presence of targeted biological species.

Films of nematic LC [4′-pentyl-4-cyanobiphenyl (5CB)] were deposited into the pores (width ∼ 283 μm; depth ∼ 20 μm) of gold grids supported on octadecyltrichlorosilane (OTS)–coated glass substrates (Fig. 1A) (14). The OTS-coated glass caused the LC to assume a perpendicular (homeotropic) orientation at the glass surface. Immediately after immersion under an aqueous solution containing a dispersion of vesicles (hydrodynamic diameter of ∼36 nm) of L-α-dilauroyl phosphatidylcholine (L-DLPC) [melting temperature (Tm) = –1°C], the LC appeared bright when viewed between crossed polars (Fig. 1B). The initial optical appearance of the LC indicates a radial orientation of LC that is parallel (planar) to the LC-aqueous interface and perpendicular to the LC-OTS interface (Fig. 1B and fig. S1). Within 10 min of immersion, black domains were observed to nucleate and grow in the LC (Fig. 1C). These domains correspond to LC that is anchored perpendicular to both the aqueous-LC and LC-OTS interfaces. By using fluorescently labeled lipids, we determined that the patterned orientation of the LC accompanied the assembly of the phospholipid on the interface and the formation of transient phospholipid domains (Fig. 2, A to F). After 2 hours of immersion (15), the optical appearance of the LC was uniformly dark, corresponding to homeotropic anchoring of the LC (16) across the entire aqueous-LC interface (Figs. 1D and 2E). Quantitative fluorescence measurements confirmed the presence of a monolayer of phospholipid at the interface (17). The interface was stable for more than 1 week after the exchange of the solution with a solution that was free of L-DLPC.

Fig. 1.

(A) Schematic illustration of the experimental system. hν, incident light. (B) Optical image and cartoon representation of the anchoring of 5CB and the state of the aqueous-5CB interface immediately after injection of a dispersion of vesicles formed from 0.1 mM l-DLPC in tris-buffered saline (TBS) (aqueous 10 mM Tris, 100 mM NaCl; pH 8.9). The optical image above the cartoon shows the transmission of polarized light (crossed polars) through the 5CB. Scale bar, 300 μm. (C) Optical image and cartoon representation of the anchoring of 5CB after ∼10 to 20 min of contact with the vesicle dispersion of l-DLPC. (D) Optical image and cartoon representation of the anchoring of 5CB after 2 hours of contact with the vesicle dispersion of l-DLPC.

Fig. 2.

(A, C, and E) Optical micrographs (transmission through crossed polars) and (B, D, and F) corresponding fluorescence micrographs of the lipid-laden 5CB-waterinterface afterexposure to aqueous (TBS) dispersions of vesicles of l-DLPC doped with 1 mole percent (mol%) TR-DHPE for[(A) and (B)] 2 min, [(C) and (D)] 30 min, and [(E) and (F)] 120 min. These images were acquired by moving the sample between a polarized light microscope and fluorescence microscope (14). About 1 min elapsed between the acquisition of the corresponding polarized light and fluorescence micrographs. The fluorescence micrograph shown in (D) was digitally enhanced to accentuate the contrast between the lipid-poor (black) and lipid-rich (white) regions at the interface. The actual fluorescence intensities of the lipid-poor and lipid-rich domains were similar to those shown in (B) and (F), respectively. (G) Plot of fluorescence intensity of a lipid layer (l-DLPC doped with 1% TR-DHPE) supported at the aqueous-LC interface immediately after patterned photobleaching (solid line) and 3 hours after patterned photobleaching (dashed line). The fluorescence intensity is plotted as a function of position along the diagonal of the two grid squares shown in the inset (indicated by the arrow). Photobleaching leads to almost complete loss of fluorescence from the lipid in the lower left compartment of the grid (inset, positions 0 to 400 μm in the plot), whereas a gradient in fluorescence intensity was generated in the upper right compartment of the grid (inset, positions 500 to 900 μm in the plot). The plot shows that the gradient in fluorescence intensity in the upper right compartment relaxes over the 3 hours after photobleaching without transport of the fluorescently labeled lipid across the grid separating the two compartments. Scale bars, 150 μm.

Many biologically relevant lipids, such as D-α-dipalmitoyl phosphatidylcholine (D-DPPC), exist in a solid or gel state at room temperature (Tm ∼ 41°C for D-DPPC). Because vesicles of these lipids do not spontaneously form densely packed monolayers at interfaces of LCs, we used mixed micelles of dodecyltrimethylammonium bromide (DTAB) and D-DPPC to deliver the D-DPPC (14, 18). Interestingly, during the transfer process with DTAB and D-DPPC, the optical texture of the 5CB changed continuously from planar to homeotropic (i.e., domains were not observed as they were in Fig. 2C when L-DLPC was used), reflecting different mechanisms of transfer of phospholipids and surfactants to the interface. After the delivery of D-DPPC to the interface, we removed the DTAB from the system by exchanging the mixed micellar solution for a solution that was free of DTAB (14).

The mobility of phospholipids plays a central role in transducing biomolecular interactions at biological membranes because it permits the reorganization of proteins and lipids after binding (e.g., leading to the formation of domains or crystallization) as well as the transport of substrates to enzymes (13, 11, 19). By performing patterned, photobleaching experiments on aqueous-LC interfaces decorated with mixtures of either L-DLPC or D-DPPC and the fluorescently labeled lipid Texas red dihexadecanoyl phosphatidylethanolamine (TR-DHPE), we measured these phospholipids to possess lateral mobilities at the aqueous-LC interface that were similar to lipids in biological membranes (20, 21). The interface was, however, compartmentalized by the gold grid (Fig. 2G). We used phospholipase A2 (PLA2) from Naja mossambica mossambica (22) and neutravidin as model systems to demonstrate that it was possible to exploit the mobility of the phospholipids to couple weak (Kd ∼ 10–4 M, where Kd is the dissociation constant) and strong (Kd ∼ 10–14 M) specific-binding events to the orientation of the LC through protein-induced reorganization of the phospholipids. Calcium-mediated binding of PLA2 to monolayers of D-DPPC (Kd ∼ 10–4 M to 10–3 M) (2325) has been measured to lead to a decrease in the tilt angle of the acyl chains of D-DPPC (19). Whereas the introduction of >100 nM PLA2 (with 5 mM Ca2+) into an aqueous phase contacting a D-DPPC–laden interface of the LC was observed to trigger a change in the optical appearance of the LC from dark to light (Fig. 3, A and B), neither 100 nM PLA2 in the absence of Ca2+ nor Ca2+ in the absence of PLA2 triggered a measurable optical response of the LC. Fluorescence imaging with labeled PLA2 confirmed the association of PLA2 with the D-DPPC–laden aqueous-LC interface as the cause of the orientational transition of the LC from the homeotropic orientation (dark appearance) to a near-planar orientation (light appearance). In contrast, nonspecific adsorption of bovine serum albumin, cytochrome-c, and lysozyme (bulk concentrations 1 to 10 μM) did not perturb the LC under any buffer conditions. In addition, contact of these interfaces with cell culture media (Dulbecco's modified eagle medium) did not perturb the orientation of the LC.

Fig. 3.

Optical images (crossed polars) of 5CB before (A) and after (B) the introduction of 100 nM PLA2 labeled with Alexa Fluor488 (∼1 fluorophore/protein) in the presence of 5 mM Ca2+ into aqueous solutions contacting d-DPPC–laden 5CB interfaces. To prepare the interface, micellar solutions of 3 mM DTAB, 0.1 mM d-DPPC, and 1 μM TR-DHPE in TBS were contacted with 5CB for0.5 hours and then subsequently exchanged with various aqueous buffers that were free of DTAB and DPPC. (C to F) Mixed l-DLPC (98%) and Bi-X-DPPE (2%) monolayers were formed at the aqueous-5CB interface by adsorption from micelles (2.5 mM DTAB, DTAB:lipid ratio of 400:1) for30 min. The micellarsolutions were exchanged forphosphate buffered saline (PBS) with pH 7.4 containing [(C) to (E)] 500 nM fluorescein-labeled NeutrAvidin or (F) 500 nM NeutrAvidin that had been blocked with biotin. After 2 hours, the protein solutions were exchanged for PBS. The optical texture of 5CB was imaged by [(C) and (F)] polarized white light with the sample light without polarizers. The bright domains in (C) correspond to tilted or planar orientations of the LC at the interface between the LC and aqueous phase, and the black regions correspond to a homeotropic orientation. (E) Fluorescein-labeled NeutrAvidin associated with the lipid-laden aqueous-5CB interface imaged by epifluorescence microscopy (14). (F) 500 nM NeutrAvidin blocked with soluble biotin in solution. Scale bars, 150 μm.

We also prepared mixed phospholipid monolayers at the aqueous-LC interface that contained 98% L-DLPC and 2% of N-((6-(biotinoyl)amino)hexanoyl)-1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine (Bi-X-DPPE). These mixed monolayers caused homeotropic orientation of the LC. Contact of these monolayers with aqueous solutions containing 500 nM fluorescently labeled neutravidin [which specifically binds biotin, Kd ∼ 10–14 M (26)] resulted in the formation of dendritic brush textures in the LC that were visible with transmitted white light in the presence (Fig. 3C) and absence (Fig. 3D) of crossed polarizers. Epifluorescence imaging of the interface revealed that the patterned orientation of the LC reported the lateral distribution of proteins at the interface (Fig. 3E). The spatial patterns observed in the LC are similar to the shapes of two-dimensional (2D) protein crystals formed at lipid-laden interfaces (4, 26). The patterned orientational response of the LC was largely eliminated when the neutravidin in solution was blocked with soluble biotin (Fig. 3F).

Whereas PLA2 binds to D-DPPC but does not catalyze the hydrolysis of D-DPPC, PLA2 stereoselectively catalyzes the hydrolysis of the sn-2 ester bond of the glycerol moiety of L-DLPC and L-DPPC to form 1-lysophospholipid and a fatty acid through a Ca2+-regulated pathway (27). We exploited this hydrolysis reaction to demonstrate that it is possible to report the enzymatic activity of PLA2 at surfaces of 5CB decorated with L-phospholipids. We observed concentrations of PLA2 as low as 10 pM in the presence of Ca2+ to trigger patterned orientational transitions in films of LC supporting L-DLPC (Fig. 4, A to C). In the absence of Ca2+, PLA2 did not cause a change in the orientation of the LC. By using fluorescently labeled lipids, we determined that the patterned orientations of the LCs accompanying the enzymatic activity of PLA2 corresponded to regions of the interface that were depleted in the fluorescently labeled lipids (Fig. 4D). The hydrolysis products of L-DLPC are known to desorb from lipid-laden interfaces (28). The time-dependence of both the optical texture of the LC (∼30 to 90 min) and the fluorescence intensity of the lipid layer correlate well with estimates of the reaction time scale for PLA2 and the lipids (29, 30). In a similar experiment, we detected the enzymatic activity of PLA2 dissolved in cell culture media (fig. S2). Whereas the hydrolysis products of L-DLPC desorb from interfaces, the hydrolysis products of L-DPPC do not. In contrast to L-DLPC, we observed that PLA2-catalyzed hydrolysis of L-DPPC led to the formation of small bright domains in the LC (∼10 to 100 μm, Fig. 4E), consistent with past reports of phase separation of lipids and fatty acid products of the hydrolysis of L-DPPC and the accumulation of enzymes at the boundary between the lipid and fatty acid (31). In the absence of calcium, or when D-DPPC was present, these domains did not form (Fig. 4F).

Fig. 4.

(A to C) Optical images (crossed polars) of 5CB following the introduction of 1 nM PLA2 into an aqueous solution of TBS containing 5 mM CaCl2 contacting the 5CB interface laden with l-DLPC and 1 mol% TR-DHPE. Images were obtained at (A) 0 min, (B) 45 min, and (C) 90 min. Scale bar, 150 μm. The black regions correspond to a homeotropic orientation of the LC at the interface between the LC and aqueous phase and the bright domains correspond to tilted or planar orientations. (D) The epifluorescence micrograph of the TR-DHPE in the lipid-laden aqueous-5CB interface corresponding to the optical texture of 5CB shown in (B) (14). (E) Optical image of 5CB after90 min of exposure of a l-DPPC–laden aqueous-5CB (TBS with 5 mM CaCl2) interface to 1 nM PLA2. (F) Optical image of 5CB after 90 min of exposure of its l-DPPC–laden aqueous (TBS with 5 mM ethylenediaminetetraacetic acid) interface to 1 nM PLA2 in the absence of Ca2+.

Because a wide range of membrane events involving proteins, lipids, and other biomolecules lead to dynamic spatial patterns associated with the reorganization of these species, the observed coupling of the interactions of proteins and phospholipids to orientational transitions of films of thermotropic LCs reported here is likely an example of a general mechanism of amplification. This mechanism may offer new approaches for reporting the formation of organized protein and lipid domains, for rapid screening of solution conditions leading to crystallization of proteins at interfaces, or for designing low-cost devices that continuously monitor for the presence of a targeted biological species in an aqueous stream. Because the orientational transitions reported in this paper lead to changes in both optical and dielectric properties of the LCs, these transitions in orientation could be exploited to report targeted analytes in simple (e.g., binary readout) optical or electrical sensors.

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Materials and Methods

Figs. S1 and S2


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