Contribution of Temperature to Deformation of Adsorbed Vesicles

Dec 22, 2014 - With increasing temperature, biological macromolecules and nanometer-sized aggregates typically undergo complex and poorly understood r...
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Contribution of Temperature to Deformation of Adsorbed Vesicles Studied by Nanoplasmonic Biosensing Eunkyul Oh,+,†,‡,⊥ Joshua A. Jackman,+,†,‡ Saziye Yorulmaz,†,‡ Vladimir P. Zhdanov,†,‡,∥ Haiwon Lee,⊥,# and Nam-Joon Cho*,†,‡,§ †

School of Materials Science and Engineering, Nanyang Technological University, 50 Nanyang Avenue 639798, Singapore Centre for Biomimetic Sensor Science, Nanyang Technological University, 50 Nanyang Drive 637553, Singapore § School of Chemical and Biomedical Engineering, Nanyang Technological University, 62 Nanyang Drive 637459, Singapore ∥ Boreskov Institute of Catalysis, Russian Academy of Sciences, Novosibirsk 630090, Russia ⊥ Department of Chemistry, College of Natural Science, Hanyang University, Seoul 133791, Korea # Department of Convergence Nanoscience, College of Natural Science, Hanyang University, Seoul 133070, Korea ‡

ABSTRACT: With increasing temperature, biological macromolecules and nanometer-sized aggregates typically undergo complex and poorly understood reconfigurations, especially in the adsorbed state. Herein, we demonstrate the strong potential of using localized surface plasmon resonance (LSPR) sensors to address challenging questions related to this topic. By employing an LSPR-based gold nanodisk array platform, we have studied the adsorption of sub-100-nm diameter 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) and 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) lipid vesicles on titanium oxide at two temperatures, 23 and 50 °C. Inside this temperature range, DPPC lipid vesicles undergo the gel-to-fluid phase transition accompanied by membrane area expansion, while DOPC lipid vesicles remain in the fluid-phase state. To interpret the corresponding measurement results, we have derived general equations describing the effect of deformation of adsorbed vesicles on the LSPR signal. At the two temperatures, the shape of adsorbed DPPC lipid vesicles on titanium oxide remains nearly equivalent, while DOPC lipid vesicles become less deformed at higher temperature. Adsorption and rupture of DPPC lipid vesicles on silicon oxide were also studied for comparison. In contrast to the results obtained on titanium oxide, adsorbed vesicles on silicon oxide become more deformed at higher temperature. Collectively, the findings demonstrate that increasing temperature may ultimately promote, hinder, or have negligible effect on the deformation of adsorbed vesicles. The physics behind these observations is discussed, and helps to clarify the interplay of various, often hidden, factors involved in adsorption of biological macromolecules at interfaces.

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inside nanowells7,8 as well as on the surface of nanorod,9 nanocube,10,11 nanodisk,12,13 and nanomehir14 structures. One of the advantages of the LSPR technique is that it has a relatively short penetration depth13,15 (about 10−20 nm compared to, e.g., 100−600 nm and around 250 nm for conventional SPR16 and quartz crystal microbalance17 (QCM), respectively). The LSPR penetration depth is comparable to the length scale of a single lipid bilayer (∼5 nm) and shorter than the length scale of lipid vesicles (∼50−70 nm diameter). This high surface-sensitivity has proven useful for tracking the kinetics of lipid vesicle adsorption and rupture on different substrates.17,19,22−24 On silicon oxide-coated nanostructures,12,13,18,19 acceleration in the time-resolved peak shift is observed when adsorbed vesicles fuse and rupture to form a

INTRODUCTION Nanoplasmonic biosensing platforms have emerged as a powerful tool to investigate phospholipid membranes on solid supports, taking advantage of optical phenomena such as extraordinary optical transmission1,2 and localized surface plasmon resonance (LSPR)3,4 for high-sensitivity measurements. The basic principle of LSPR measurements is that metallic nanostructures5,6 have a characteristic maximumextinction wavelength, λmax, which depends on nanoparticle properties, array configuration, and local dielectric environment. In biosensing measurements, the binding of organic aggregates (e.g., lipid bilayer patches or vesicles) or biomacromolecules (e.g., proteins) typically elicits a redshift in the maximum-extinction wavelength due to changes in the local refractive index (within the nanoparticle’s evanescent field) and the corresponding shift, Δλmax, is tracked. This technique is naturally compatible with various fabrication strategies. In particular, lipid membranes have been formed © 2014 American Chemical Society

Received: October 29, 2014 Revised: December 21, 2014 Published: December 22, 2014 771

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thermodynamically favorable for fluid-phase lipid vesicles to undergo shape deformation as compared to gel-phase lipid vesicles. The particular Tm value of a phospholipid varies depending on factors that influence membrane packing such as properties of the lipid headgroup and hydrocarbon chains (e.g., chain length and number of unsaturated bonds). 1,2-Dioleoyl-snglycero-3-phosphocholine (DOPC, Tm of −17 °C) and related low-Tm lipids are commonly used for fabricating phospholipid membranes on solid supports because typical unilamellar vesicle preparation methods (e.g., extrusion) require fluidphase membrane compositions 36 and vesicles of such compositions are able to rupture on certain substrates (e.g., silicon oxide37 and mica22). In other cases, higher-Tm lipids such as 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC, Tm of 23 °C) and 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC, Tm of 41 °C) are used in lipid membrane studies, especially to investigate the behavior of membranes in different phase states (e.g., in supported lipid bilayers38 and adsorbed vesicles25,26,39). Unilamellar vesicle preparation for higher-Tm lipid compositions is more difficult (albeit still feasible) because extrusion must occur above the highest Tm of lipids in the membrane composition.36 In addition to the effects of temperature on lipid membrane properties, it also strongly influences the diffusion rate of lipid vesicles in solution and the transformation of adsorbed vesicles into a supported lipid bilayer, because the latter process is thermally activated.40−42 As such, the role of temperature in the kinetics of vesicle adsorption and rupture is complex and we primarily focus our attention here on vesicle adsorption and shape deformation. To facilitate such studies, higher-Tm lipid vesicles are typically used because control over the membrane phase state at experimentally convenient temperatures allows one to track vesicle adsorption in different phase states. To the best of our knowledge, nearly all vesicle adsorption studies involving higher-Tm lipid vesicles have employed the QCM technique for tracking the adsorption kinetics. In particular, measurements by Reviakine et al.26 first indicated that at room temperature (∼23 °C) an adlayer of gel-phase DPPC lipid vesicles induced a significantly larger measurement response (frequency and dissipation shifts proportional to adsorbed mass37) than a fluid-phase DOPC lipid vesicle adlayer, and it was inferred that the equivalent thickness of the former is greater. Both vesicle populations had similar size in solution, leading to the conclusion that adsorbed DPPC lipid vesicles undergo less shape deformation than adsorbed DOPC lipid vesicles. Motivated by this finding, gel-phase DPPC lipid vesicles have been explored as a model system for colloidal particle adsorption with QCM model-independent analysis39 and there has been increasing attention to investigate temperature-dependent effects for high-Tm lipid compositions. In another study, Reviakine et al.25 investigated the adsorption of extruded DMPC lipid vesicles onto titanium oxide as a function of temperature, and they reported that gel-phase lipid vesicle adlayers have greater equivalent thickness than fluidphase lipid vesicle adlayers. Jing et al.43 demonstrated that extruded fluid-phase DPPC lipid vesicles (below a certain size, ∼90 nm diameter) rupture to form a complete supported lipid bilayer on silicon oxide. Gel-phase DPPC lipid vesicles may also be deposited on silicon oxide at room temperature, followed by a temperature ramp that induces vesicle rupture and bilayer formation starting at Tm. Likewise, tip-sonicated, fluid-phase DPPC lipid vesicles form supported lipid bilayers on silicon

supported lipid bilayer. This acceleration has been attributed to the fact that lipids in a supported lipid bilayer are, on average, nearer to the substrate than lipids in an adsorbed vesicle. Recently, the LSPR technique has also provided detailed information about vesicle adsorption on titanium oxide-coated nanostructures.15 In the latter case, vesicles adsorb but do not rupture, instead forming an adsorbed vesicle layer. Scaling laws were developed for the dependence of Δλmax on lipid concentration or vesicle size in the case of adsorption of nondeformed vesicles, and deviations from the expected scaling laws were attributed to shape deformation of adsorbed vesicles. The extent of vesicle shape deformation in different coverage regimes was qualitatively interpreted from the LSPR measurements, and the findings provide motivation for continued LSPR-based studies on lipid vesicles as a model for soft matter adsorption at the liquid−solid interface. In this regard, there are many experimental parameters which influence the behavior of adsorbed vesicles, including vesicle characteristics, solution conditions, and environmental factors (see refs 20 and 21 and references therein). Understanding how different parameters contribute to the deformation of adsorbed vesicles is still incomplete. In general, adsorbed vesicles undergo shape deformation,22−26 which is governed by the vesicle−substrate interaction and vesicle bending energy. The relative contribution of each factor is characterized by a dimensionless parameter ϱ ≡ Wr2/κ, where W is the contact energy per unit area, κ is the membrane bending rigidity, and r is the vesicle radius in solution in the nondeformed state.27,28 If ϱ is relatively large, then vesicles deform appreciably, which can lead to vesicle destabilization, rupture, and bilayer formation. On the other hand, if ϱ is relatively small, then vesicle deformation will be minimal and adsorbed vesicles are stable. Despite the apparent simplicity of this criterion, its use is often far from straightforward, because the extent of the influence of governing parameters (e.g., temperature, pH, or ionic strength) on κ and/or W is hidden. The role of temperature in the shape deformation of adsorbed vesicles deserves special attention because it influences the phase state of lipid membranes in general and κ and W in particular. With increasing temperature, a lipid membrane passes through the first-order gel-to-fluid phase transition temperature (Tm); that is, the phospholipids in the membrane change from a rigid, ordered state to a fluid, disordered state.29,30 Some of the earliest studies (see, e.g., ref 31) on this phase transition were focused on bacterial membranes of complex lipid composition, and the corresponding phase transition temperature range was reported to be rather broad because each type of phospholipid has a particular Tm value. Subsequent characterization of the phase transition for single types of synthetic phospholipid proved advantageous in order to understand the thermodynamics of this process, including a much sharper temperature-dependent transition for pure, uniform phospholipids.29,32 Importantly, the phase transition of a lipid membrane to the fluid state is reported to lead to an appreciable decrease in the bending modulus,33 typically from ∼10‑18 J (gel-phase at T < Tm) to ∼10‑19 J (fluidphase at T > Tm); note that the exact values obtained for the bending modulus depend on the measurement technique34 and membrane curvature.35 For highly curved membranes (e.g., vesicles below 40 nm diameter), the Tm value may be slightly lower.35 The contact energy is also expected to depend on temperature although to a much lesser extent. Hence, one could expect that, in the adsorbed state, it would be more 772

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to saturation and accordingly can be identified with the diffusion flux, J, toward the substrate surface. For a channel with a rectangular cross-section, this transient regime is described as15,48

oxide while gel-phase variants may also rupture to some extent on silicon oxide, yielding a combination of supported lipid bilayer patches and intact vesicles (as confirmed by atomic force microscopy).44−46 Taken together, these QCM-based studies describe the behavior of adsorbed high-Tm lipid vesicles on solid supports and the corresponding adsorption kinetics remain to be scrutinized in detail. Indeed, while the QCM technique has many advantages, it is also known that its measurement responses are not linearly proportional to the surface coverage of adsorbed vesicles.26 One of the problems complicating the interpretation of the data in such cases is that the QCM frequency shift depends on the amount of trapped water, which can hardly be accurately taken into account. Comparing experimental data obtained at different temperatures is especially difficult, because the viscosity of water strongly depends on temperature and accordingly the amount of trapped water changes appreciably with increasing temperature. Herein, we investigated the adsorption kinetics of DPPC lipid vesicles on titanium oxide and silicon oxide with temperature-controlled LSPR extinction maximum wavelength shift measurements. By performing the experiments below and above the corresponding Tm, gel- and fluid-phase DPPC lipid vesicles of otherwise equivalent properties were studied, allowing us to determine the relative contribution of membrane phase, temperature, and shape deformation to the adsorption process. In some cases, DOPC lipid vesicles were also tested under identical temperature conditions for comparison. Furthermore, the extent of shape deformation of adsorbed vesicles on titanium oxide and silicon oxide is discussed in the context of vesicle adsorption and rupture. To facilitate the interpretation of our experimental results, we first extend our previous theoretical analysis15 of the effect of adsorbed vesicles on the LSPR signal with emphasis on the role of vesicle deformation. Then, the experimental results are presented.

⎛ v D2 ⎞1/3 C = Jt ≅ ⎜ 0 ⎟ nt ⎝ hx ⎠

where D and n are the vesicle diffusion coefficient and number concentration in solution, respectively, v0 is the average flow velocity, h is the channel height, and x (0 ≤ x ≤ L) is the coordinate along the channel. According to hydrodynamics, we have

D=

kBT 6πηr

(3)

where η is the solution viscosity. In conventional SPR spectroscopy, interpretation of the results is based on the fact that the decay of the evanescent field in solution is exponential.16,47 In the LSPR case, the decay is, however, of the power-law type, and in the case of spherically shaped vesicles, as explained in detail elsewhere,15 eq 1 can be specified as 2r

dz (R + z)6 (4) * where l is the thickness of the lipid bilayer, z is the coordinate perpendicular to the substrate surface (z = 0 corresponds to the vesicle−substrate contact), and R* is the length scale characterizing the distance between the center of a gold nanoparticle and adjacent vesicles contributing to Δλmax. By definition, R* includes the dielectric layer covering the nanoparticles, and taking into account that the substrate layer covering the gold nanoparticles is relatively thin, R* is expected to be comparable to the average length scale of a gold nanoparticle. To show the effect of deformation of adsorbed vesicles on the LSPR signal, we represent a vesicle as a truncated sphere with a circular vesicle−substrate contact area of radius a (Figure 1A). During deformation, the total vesicle area is conserved. For this reason, its radius, r*, is slightly larger than that in solution. In this simple model, the vesicle shape characteristics Δλmax ∝ Clr

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THEORETICAL ASPECTS In the experimental part of our work, we used the LSPR wavelength shift, Δλmax, to track the kinetics of vesicle adsorption onto the substrate surface and their state in the adsorbed overlayer at different temperatures. Here, we show theoretically how the deformation of vesicles can influence Δλmax. In the context of our work, we focus on two types of deformation. The first one is expansion of vesicle surface area which may be significant with increasing temperature (e.g., in the DPPC lipid case). This expansion occurs irrespective of whether a vesicle is in the bulk solution or in the adsorbed state. The second one, related to adsorption, is shape deformation itself. Below, we (i) specify the general relation between Δλmax and adsorption kinetics [eqs 1−3], (ii) analyze the effect of vesicle shape deformation on Δλmax [eqs 5−13], and (iii) scrutinize the role of expansion of vesicle surface area [eqs 14−18]. In analogy with conventional SPR spectroscopy,16,47 the wavelength shift measured in LSPR during vesicle adsorption can be represented as Δλmax ∝ CF(r )

(2)

∫0

(1)

where C is the surface concentration of vesicles, F(r) is the function describing the contribution of a single vesicle to the signal, and r is the vesicle radius in solution in the nondeformed state. The specifics of vesicle adsorption is that the corresponding rate is usually limited by diffusion almost up

Figure 1. Illustration of the influence of deformation of adsorbed vesicles on the LSPR Signal. (A) A deformed vesicle is represented as a truncated sphere. (B) Measures of the effect, P1 [eq 10] and P2 [eq 13], as a function of a/r for r = 36 nm and R* = 38 nm. 773

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corresponds to appreciably deformed vesicles). With increasing a/r, P1 and P2 rapidly increase. The difference between P1 and P2 remains, however, minor. In the complementary part of our analysis, we treat the effect of vesicle surface area expansion on Δλmax under the condition that the shape of vesicles remains the same. Let us associate the initial and expanded states with subscripts 1 and 2, respectively. According to eq 9, the corresponding LSPR peak shifts are represented as

(5)

while the vesicle height is expressed via r* and a as h = r + (r 2 − a 2)1/2 (6) * * With this specification, the contribution of deformed vesicles to Δλmax can be calculated in analogy with how it was earlier done for spherically shaped vesicles.15 In particular, eq 4 can be generalized as ⎛ a2 Δλmax ∝ Cl⎜ 6 + r * ⎝ R*

Δλmax ∝

Δλmax ∝

Clr R5 *

(14)

⎞ Cl 2 ⎛ 5a 22 ⎜ ⎟ + r 2 * R 5 ⎝ R* ⎠ (15) * Taking into account that during surface area expansion the vesicle mass does not change and assuming that the change of the lipid density in the bilayer is negligible, we have r2 = αr1 and l2 = l1/α2, where α > 1 is the linear expansion coefficient. Substituting these relations into eq 15 yields Δλmax ∝

⎞ dz ⎟ 6 (R + z ) ⎠ (7) * where the first and second terms on the right-hand side correspond to the contact and out of contact areas, respectively. As shown earlier,15 the scale of the interval making the main contribution to the integrals in eqs 4 and 7 is Δz ≅ R*/5. This interval is usually appreciably smaller than the vesicle size. For this reason, the integrations in eqs 4 and 7 can be extended up to infinity, and one can rewrite eqs 4 and 7, respectively, as

∫h

⎞ Cl1 ⎛ 5a12 ⎜ + r1 *⎟ 5 R ⎝ R* ⎠ *

0

Cl1 ⎛ 5a12 r ⎞ ⎜ + 1* ⎟ 5 α⎠ R ⎝ R* (16) * Concerning eqs 14 and 16, we conclude that in the case of single vesicles the area expansion does not change the contribution of the vesicle−substrate contact area to Δλmax, while the contribution of the remaining part of a vesicle is reduced by a factor of 1/α. At saturation, we respectively have C ∝ 1/r21* and C ∝ 1/r22* = 1/r21*α2. Substituting these expressions into eqs 14 and 16, we obtain Δλmax ∝

(8)

⎞ Cl ⎛ 5a 2 +r⎟ 5⎜ *⎠ R ⎝ R* (9) * Let us now consider the situation when the thickness of the lipid bilayer does not change. In this case, the ratio of eqs 9 and 8, defined as Δλmax ∝

Δλmax ∝

2

r 5a + * rR r (10) * is a measure of the effect of deformation of single vesicles on the LSPR signal. By definition, the effect is calculated with respect to nondeformed vesicles. Due to deformation, vesicles become closer to the gold nanoparticles, and their contributions to Δλmax increase, and accordingly P1 ≥ 1. At saturation of nondeformed and deformed vesicles, we have C ∝ 1/r2 and C ∝ 1/r2*, respectively. Substituting these expressions into eqs 8 and 9 yields, respectively, P1 ≡

⎞ l1 ⎛ 5a12 ⎜ ⎟ + r 1 * r12*R 5 ⎝ R * ⎠ *

(17)

⎞ l ⎛ 5a 2 +r⎟ 2 5⎜ * r R ⎝ R* ⎠ (12) * * Assuming again that l does not change, we can represent the ratio of eqs 12 and 11 as

⎛ 5a 2 r1 * ⎞ 1 ⎟ ⎜ + α⎠ α 2r12*R 5 ⎝ R * (18) * Due to the presence of α in the denominator of eq 18, the area expansion is seen to reduce Δλmax. The equations derived above can help to interpret the influence of T on Δλmax during the transient adsorption phase and at saturation. In the former case, the change of Δλ with increasing T depends on D and on vesicle deformation. In the latter case, this change depends exclusively on vesicle deformation. Concerning the dependence of D on temperature, we may notice that according to eq 3 D is proportional to T. The role of this factor is, however, minor. A more important factor is that η depends on T in the inverse Arrhenius fashion. Although the corresponding activation energy is relatively small (about 4.8 kcal/mol for water42), it results in a significant increase of D with increasing T.

5a 2r r + r r 2R (13) * * * This is a measure of the effect of deformation of vesicles on the LSPR signal in the case of saturation. In analogy with P1, we have P2 ≥ 1. The ratios P1 [eq 10] and P2 [eq 13] are shown in Figure 1B as a function of a/r changing in the range from 0 to 1 (the latter

Vesicle Preparation. Small unilamellar vesicles composed of DPPC or DOPC lipid (Avanti Polar Lipids, Alabaster, AL) were prepared by the extrusion method. As-supplied lipids in chloroform were dried with nitrogen air in order to form a lipid film. After drying in a vacuum desiccator for a minimum of 12 h, the lipid film was hydrated in 10 mM Tris buffer [pH 7.5] with 150 mM NaCl to a nominal lipid concentration of 4 mg/mL. For DOPC lipid samples,

Δλmax ∝

l rR 5 *

Δλmax ∝

(11)

Δλmax ∝

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P2 ≡

774

l1

MATERIALS AND METHODS

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Figure 2. Schematic of temperature-controlled nanoplasmonic biosensing. (A) Illustration of gel- and fluid-phase lipid vesicle adsorption onto titanium oxide-coated gold nanodisks. Top: Adsorption of gel-phase lipid vesicles with weak shape deformation. Bottom: Adsorption of fluid-phase lipid vesicles at higher temperature with more appreciable shape deformation. Additional deformation, accompanying the phase transition, and a greater shift in the LSPR signal per vesicle are expected to be related to decrease of the lipid bilayer bending modulus. In reality, the situation may be different (see Conclusion). (B) Representative extinction spectrum of titanium oxide-coated gold nanodisks. LSPR measurements were recorded in aqueous buffer solution at 23 °C (black curve), and then a heating ramp was applied to increase the temperature to 50 °C (red curve). A blueshift of ∼0.8 nm is observed upon heating. by high-order polynomial fitting, and the centroid position was calculated from the fit.50 Experiments at room temperature (23 °C) were conducted under ambient conditions. For high-temperature experiments, a thermocouple wire in the measurement chamber provided a feedback loop to keep the temperature constant at 50 ± 0.1 °C. In this case, samples were incubated in a 50 °C water bath (at least 15 min equilibration) before addition to the measurement chamber. All baseline LSPR signals were recorded in aqueous buffer solution at the desired temperature. During experiment, liquid sample was continually introduced by peristaltic pump at a constant flow rate of 10 μL/min, and the average flow velocity was 5 mm/min. Note that the flow rate is ten-fold less than in our recent LSPR report15 in order to ensure proper heating of the liquid in the measurement chamber. The channel cross-section and length is 1 × 2 and 5 mm, respectively.

extrusion was then performed at room temperature with 17 cycles through 50 nm track-etched polycarbonate membranes, followed by 17 cycles through 30 nm membranes. For DPPC lipid samples, the same extrusion procedure was followed except that it was conducted at a higher temperature above the Tm of 41 °C. Before extrusion, hydrated DPPC lipid samples were heated to 60 °C using a water bath and the extruder block was heated to 80 °C on a hot plate. After extrusion, vesicle solutions were stored at 4 °C until experiment and used within 48 h. Dilution to the experimental lipid concentration occurred immediately before experiment. All aqueous solutions were prepared using high-resistivity (>18.2 MΩ·cm) deionized water that had been treated with the Milli-Q filtration system (Millipore, Billerica, MA). Dynamic Light Scattering. The technique was employed in order to calculate the vesicle size distribution based on the diffusion properties of extruded lipid vesicles in solution. Specifically, the diffusion coefficient of vesicles was obtained and is inversely proportional to the hydrodynamic radius of vesicles according to eq 3. To perform the measurements, a 90Plus particle size analyzer (Brookhaven Instruments, Holtsville, NY) with a 658.0 nm monochromatic laser was employed. All measurements were taken at the scattering angle of 90° and the intensity autocorrelation function was measured, which can be fit directly in order to yield the intensityweighted size distribution of vesicles in solution. The autocorrelation function was deconvoluted by using the Method of Cumulants in order to calculate the intensity-weighted Gaussian profile of the vesicle size distribution (including average effective diameter and polydispersity). Measurements were performed at 23 or 50 °C, and the solution viscosity and refractive index were automatically adjusted in the BIC Particle Sizing software package (Brookhaven Instruments). The temperature in the measurement chamber is maintained via an active feedback loop, although the probe is not directly embedded in the liquid solution. Hence, for measurements recorded at 50 °C, optimal results were obtained by preheating the liquid sample in a 50 °C water bath (at least 15 min equilibration) prior to experiment. LSPR Measurements. Ensemble-averaged LSPR measurements on nanodisk arrays were performed in optical transmission mode by using an Insplorion XNano instrument (Insplorion AB, Göteborg, Sweden), as previously described.15 Titanium oxide- and silicon oxidecoated gold nanodisk arrays on glass substrates were purchased from Insplorion AB, and used as-supplied following water and ethanol rinses and oxygen plasma treatment for 1 min at maximum radiofrequency power (Harrick Plasma, Ithaca, NY, USA). During experiment, a white light beam entered the measurement chamber, passed through the sensor chip (∼4 mm2 circular spot), and exited through a quartz glass window. The transmitted light was collected by a spectrophotometer, and data analysis was performed with the Insplorer software package (Insplorion AB). The frequency resolution of the LSPR signal was 1 Hz. The spectral resolution of the plasmon resonance was determined

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RESULTS AND DISCUSSION Temperature-Controlled Nanoplasmonic Biosensing. Previous studies on temperature-controlled nanoplasmonic sensing have typically focused on measurements in gas/air environments. There have been a few biosensing reports51,52 describing the effects of temperature on measurements in liquid environment mainly relating to issues such as detection sensitivity, but there have been no such investigations related to the effect of temperature on the kinetics of biomacromolecular interactions. Here, we explored lipid vesicle adsorption onto a titanium oxide-coated substrate consisting of deposited gold nanodisks on a glass substrate (Figure 2A). By controlling the temperature, we could adjust the membrane phase state of lipid vesicles and monitor the corresponding adsorption kinetics. In our system, the deposited gold nanodisks (100 nm diameter and 20 nm height) had an ∼8% surface coverage with random arrangement and were fabricated by hole-mask colloidal lithography.53 After fabrication of the gold nanodisk array on glass, the array was sputter-coated with a 10 nm thick coating of a dielectric material, either titanium oxide or silicon oxide, across the entire substrate. After fabrication was completed, each nanodisk had characteristic dimensions of around 22 nm height and 150 nm diameter inclusive of the 10nm-thick sputtered titanium oxide or silicon oxide coat (see ref 15 for corresponding atomic force microscopy images). At 23 °C (room temperature), a representative plasmon peak recorded in aqueous buffer solution for a titanium oxide-coated gold nanodisk array was around 758 nm and the ensembleaveraged LSPR signal was stable for biosensing measurements (drift was less than 0.1 nm in 15 min) (Figure 2B). 775

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Figure 3. Influence of temperature on lipid vesicle adsorption onto titanium oxide. (A) Time-resolved shift in maximum-extinction wavelength for 0.1 mg/mL extruded DOPC lipid vesicles at two different temperatures. From t = 10 min, vesicles were added under continuous flow. (B) Derivative of the time-resolved shift in maximum-extinction wavelength (nm/min) presented for the data from Panel (A). Panels (C) and (D) depict the same series of measurements recorded for 0.1 mg/mL DPPC lipid vesicles. The black and red lines correspond to measurements recorded at 23 and 50 °C, respectively.

Experiments were also performed at higher temperature (50 °C). In this case, the baseline measurement was again recorded in aqueous buffer solution at room temperature, followed by a temperature ramp (2 °C/min) to reach the desired temperature. There was a consistent blueshift of ∼0.8 nm due to heating (this shift is related to the dependence of the bulk refractive index of water on temperature54), and the LSPR signal was stable for biosensing measurements thereafter. Adsorption on Titanium Oxide. We first measured the adsorption kinetics of 0.1 mg/mL DOPC lipid vesicles onto titanium oxide (Figure 3A). As described in the Introduction, the Tm of DOPC lipid is −17 °C so DOPC lipid vesicles exhibit fluid-phase behavior across the experimental range of temperatures. The average diameter of extruded DOPC lipid vesicles, determined at room temperature, was 86.2 nm. After establishing a measurement baseline in aqueous buffer solution at the desired temperature, 0.1 mg/mL DOPC lipid vesicles were added at t = 10 min. A redshift was observed and the corresponding peak shift was recorded as a function of time. At 23 °C, monotonic adsorption of fluid-phase DOPC lipid vesicles was observed. The final peak shift was 2.4 nm, and the time scale was 20 min. At 50 °C, there was similar adsorption behavior with a time scale of around 25 min, but the final peak shift was only 1.7 nm at saturation. That is, the peak shift decreased by about 1.4-fold at the higher temperature. To scrutinize the kinetics of vesicle adsorption, we also show the time derivative of the peak shift (Figure 3B) which more explicitly characterizes the rate of this process. We next measured the kinetics of DPPC lipid vesicle adsorption onto titanium oxide (Figure 3C,D). The average diameter of extruded DPPC lipid vesicles used in the LSPR measurements was 71.5 nm, as determined by dynamic light scattering measurements at room temperature. At 23 °C, monotonic adsorption of DPPC lipid vesicles occurred until reaching saturation. The final peak shift was 4.1 nm, and the

time scale of the whole process was 22 min. Similar adsorption behavior was also observed for DPPC lipid vesicles at 50 °C. In the latter case, the final peak shift at saturation was 2.4 nm and the time scale was 15 min, reflecting a 1.7-fold decrease in the final peak shift at the higher temperature. Comparing Panels A and C of Figure 3, the first impression might be that the temperature dependences of the LSPRtracked adsorption kinetics of DOPC and DPPC lipid vesicles are qualitatively similar and that the extent of vesicle deformation might be similar as well. Although the former is apparently the case, the latter can be debated because in order to draw the right conclusions from Figure 3A,B and 3C,D, we should take into account that temperature influences the diffusion rate, size, and lipid bilayer thickness of DOPC and DPPC lipid vesicles in different ways, and these factors can in turn influence interpretation of the observed adsorption kinetics. To interpret the kinetics, we should first choose the key features which can be quantified. From this perspective, we note that three stages of vesicle adsorption (Figure 3) are captured in the LSPR signal: (i) initial rate of the peak shift when vesicles are first added to the measurement chamber; (ii) the constant rate of the peak shift at low vesicle surface coverage; and (iii) the gradually declining rate of the peak shift at moderate and high surface coverages. Here, we focus on analyzing mainly the rate of the LSPR signal shift during stage (ii) and the final shift at stage (iii) and also partly on the duration of the whole transient period. Stage (ii) is especially convenient for the analysis because, during this stage, adsorbed vesicles are expected to occupy energetically preferable areas and there are no vesicle−vesicle lateral interactions. To quantify this stage, we take into account that according to eqs 2 and 3 the dependence of the vesicle adsorption rate, v, on temperature is determined by temper776

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Langmuir ature itself and also by the temperature dependence of η and r. In particular, we have ⎛ T ⎞2/3 v∝⎜ ⎟ ⎝ ηr ⎠

appreciably deformed. The former value is slightly larger than 0.74, supporting that the vesicles at low coverage are slightly less deformed at the higher temperature. In the end of stage (iii), on the other hand, the ratio of the LSPR signals is 2.5 nm/ 4.1 nm = 0.61 (see Figure 3C). According to eq 18, the area expansion-related decrease is either ≅ 1/α3 = 1/1.243 = 0.52 or ≅ 1/α2 = 1/1.222 = 0.65 depending on whether vesicles are nearly spherical or appreciably deformed. Both of these values are close to 0.61 and accordingly indicative that temperatureinduced shape deformation is nearly negligible. Taken together, the data in this case support the conclusion that adsorbed DPPC lipid vesicles are nearly spherical at room temperature and undergo modest shape deformation at higher temperature. Effect of DPPC Lipid Concentration on Vesicle Adsorption. In order to characterize DPPC lipid vesicle adsorption on titanium oxide in more detail, the corresponding kinetics were next investigated as a function of lipid concentration between 0.025 and 0.1 mg/mL. The final peak shifts at each respective lipid concentration and temperature are reported in Table 1. At 23 °C, the final peak shift at low lipid

(19)

The interpretation of the adsorption kinetics is simpler in the case of DOPC lipid vesicles, because with increasing temperature from 23 °C up to 50 °C the change of the size and bilayer thickness of vesicles is nearly negligible (the corresponding membrane surface area expansion55 is only ∼5%). This means that in this case the increase of the adsorption rate with increasing temperature is determined by the ratio (T/η)2/3. As already mentioned in the Theoretical Aspects section, η depends on T in the inverse Arrhenius fashion. At 23 °C, η is around 930 μPa/s and decreases to approximately 550 μPa/s at 50 °C (see, e.g., ref 56 for standard reference values). With these values, eq 19 predicts that the rate of adsorption increases by a factor of 1.5. During stage (ii), the rate of the change of the LSPR peak shift was ∼0.18 and ∼0.24 nm/min at 23 and 50 °C, respectively (Figure 3B); that is, the LSPR-tracked rate increased by a factor of 0.24/0.18 = 1.3. This increase is smaller than that predicted by eq 19 by a factor of 1.3/1.5 = 0.87. This difference is attributed to the fact that the DPPC vesicles are slightly less deformed at the higher temperature. With this deformation, the change of the vesicle radius (r*) in the adsorbed state is nearly negligible (see Theoretical Aspects section). This means that the duration of the whole adsorption process should be inversely proportional to (T/η)2/3; that is, it should be longer by a factor of 1.5 at the lower temperature. This prediction is in line with the experiment (see Figure 3B). The difference 1.3/1.5 = 0.87 can also be identified with the expected decrease of the asymptotic LSPR signal at the higher temperature. The corresponding experimental ratio at the end of stage (iii) is 1.7 nm/2.4 nm = 0.71. The fact that the latter value is slightly smaller than 0.87 is not surprising, because these ratios correspond to different coverages. Thus, interpretation of various data for DOPC vesicle adsorption appears to be self-consistent, and the key conclusion in this case is that adsorbed DOPC lipid vesicles are less deformed at the higher temperature. In particular, characteristic features in the related LSPR signal decrease by a factor of 0.7−0.9. The corresponding extent of deformation can be inferred from Figure 1B. In the case of DPPC lipid vesicles, the complicating factor is that due to the gel-to-fluid phase transition the vesicle radius is larger at the higher temperature. For spherical vesicles in solution, the diameter was reported to increase by a factor of 1.22 (page 280 in ref 57). We performed DLS measurements on DPPC lipid vesicles at 23 and 50 °C and observed a similar 1.24-fold increase in vesicle diameter from 65.5 to 81.0 nm. In combination with the ratio (T/η)2/3 discussed above, this means that according to eq 19 the vesicle adsorption rate is higher by a factor of 1.3. In contrast, the experiment showed that during stage (ii) the rate of the change of the LSPR peak shift was ∼0.25 and ∼0.24 nm/min at 23 and 50 °C, respectively (Figure 3D); that is, the LSPR-tracked rate decreased by a factor of 0.24/0.25 = 0.96. This is smaller than that predicted by eq 19 by a factor of 0.96/1.3 = 0.74. This difference arises from the vesicle area expansion and deformation. According to eq 16, the area expansion-related decrease of the LSPR signal is either ≅ 1/α = 1/1.24 = 0.81 or close to 1 depending on whether vesicles are nearly spherical or

Table 1. Experimental Summary for DPPC Lipid Vesicle Adsorption onto Titanium Oxidea Temp

Lipid Composition

0.1 mg/mL

0.05 mg/mL

0.025 mg/mL

23 °C 50 °C

DPPC DPPC

4.1 2.4

3.5 2.7

3.5 2.2

a

Final peak shift values from maximum-extinction wavelength measurements are reported from single measurements at each lipid concentration.

concentrations (0.025 and 0.05 mg/mL) was 3.5 nm, which is smaller than the final peak shift of 4.1 nm at 0.1 mg/mL lipid concentration (Figure 4A). The results are consistent with our previous findings on the role of lipid concentration and support that the diffusion flux influences the packing of vesicles in the adlayer.15 At low and moderate coverages, the kinetic curves constructed as a function of ct for the measurement results obtained at 23 °C are similar and nearly linear, both of which are consistent with diffusion-limited adsorption (Figure 4B). On the contrary, there was no clear trend in the effect of lipid concentration on DPPC lipid vesicle adsorption at 50 °C, although the final peak shifts were consistently around 2.4 nm (Figure 4C). In this case, the kinetic curves constructed as a function of ct did not coincide for all lipid concentrations (Figure 4D). Comparatively, the final peak shifts were always at least 3.5 nm for DPPC lipid vesicle adsorption at 23 °C and no greater than 2.7 nm for DPPC lipid vesicle adsorption at 50 °C. This finding is important because it supports that the effects of temperature on the kinetics of vesicle adsorption are consistent across a range of lipid concentrations. Adsorption on Silicon Oxide. While fluid-phase DPPC lipid vesicles below a certain size (i.e., less than 90 nm diameter) rupture on silicon oxide, gel-phase DPPC lipid vesicles typically adsorb and remain intact on silicon oxide.43 These findings motivated us to also investigate the kinetics of DPPC lipid vesicle adsorption on silicon oxide (Figure 5). After establishing a measurement baseline, 0.1 mg/mL DPPC lipid vesicles were added at t = 10 min, as presented in Figure 5A. At 23 °C, the lipid vesicles adsorbed onto silicon oxide until reaching saturation and yielded a final peak shift of 3.7 nm. The adsorption kinetics showed monotonic behavior without acceleration over a time scale of greater than 40 min. At 50 777

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Figure 4. Effect of DPPC lipid concentration on vesicle adsorption onto titanium oxide. Time-resolved shift in maximum-extinction wavelength for three lipid concentrations in solution and the same shift scaled according to ct, presented in Panels (A) and (B), respectively, for LSPR measurements recorded at 23 °C. Panels (C) and (D) represent the same series of measurements performed at 50 °C.

expected, the time scale of bilayer formation was appreciably quicker than that leading to an adsorbed vesicle layer, and occurred over 10 min. The corresponding time derivatives of the peak shifts are also presented (Figure 5B). For vesicle adsorption without rupture, the main feature to quantify was again the constant rate associated with stage (ii) as described above for the titanium oxide case. At 23 °C, the diffusion-limited rate of the peak shift increase was 0.12 nm/min. For vesicle adsorption with rupture (i.e., bilayer formation), the main feature was the constant rate observed in the LSPR-tracked signal before rate acceleration due to vesicle rupture. The rate at this stage was 0.18 nm/min and is related to diffusion-limited vesicle adsorption12 and corresponding shape deformation. While the rate of diffusion of vesicles in solution increases by a factor of ∼1.3 at the higher of these two temperatures, the LSPR-tracked signal increased by a factor of 1.5, indicating that adsorbed vesicles (before rupture) are more deformed at higher temperature, which is in contrast to the titanium oxide case. Furthermore, the LSPR measurements conducted on silicon oxide at the two temperatures also show that the final peak shift associated with an adsorbed layer of nondeformed vesicles is greater than the peak shift of a supported lipid bilayer. One could expect the opposite because the jamming limit (maximum packing fraction) of adsorbed vesicles in a twodimensional array is ∼0.77,15 indicating that there is not full coverage of lipids attached to the surface in this case, as compared to full coverage of the substrate by a supported lipid bilayer. Hence, while the LSPR measurement is highly surfacesensitive, the penetration depth is probably at least on the order of 10−20 nm (in line with our previous theoretical estimate15) in order to explain how an adsorbed vesicle layer induces a larger peak shift than a supported lipid bilayer.

Figure 5. Influence of temperature on lipid vesicle adsorption onto silicon oxide. (A) Time-resolved shift in maximum-extinction wavelength for 0.1 mg/mL extruded DPPC lipid vesicles at two temperatures. (B) Derivative of the time-resolved shifts shown in Panel (A). The black and red lines correspond to measurements performed at 23 and 50 °C, respectively.

°C, lipid vesicle adsorption was nearly constant until reaching a peak shift of 1.5 nm, followed by acceleration that yielded a final peak shift of 2.8 nm. The rate acceleration along with the magnitude of the final peak shift is consistent with the formation of a supported lipid bilayer on silicon oxide.12,13,18 As 778

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CONCLUSION In this work, we have investigated the effect of temperature on the deformation of adsorbed vesicles on two solid supports, namely titanium oxide and silicon oxide. A phenomenological model describing the LSPR-related physics of vesicle adsorption has been extended to account for shape deformation and membrane area size expansion. By interpreting the experimentally observed adsorption kinetics in accordance with this model, we observed on titanium oxide that with increasing temperature the shape deformation of adsorbed DOPC vesicles at low coverage decreases but not dramatically while the shape deformation of adsorbed DPPC vesicles remains nearly the same. The former vesicles were in the fluidstate across the range of experimental temperatures, while the latter ones were in the gel-state at low temperature and the fluid-state at high temperature. Concerning these findings, we may notice that due to the presence of the plasmonic gold nanodisks the surface of the support used in our work to study vesicle adsorption is slightly heterogeneous (topographically flat substrates can be fabricated as described in ref 58). Although the role of this factor is not fully negligible (as discussed in our previous study15), it does not seem to influence the interpretation of our present results because the effects reported here are more pronounced. To rationalize these observations, we again (cf. the introduction) refer to the relevant dimensionless parameter, ϱ ≡ Wr2/κ, and note that it was derived by describing vesicles phenomenologically at the level of the membrane bending energy and contact vesicle−substrate interaction.27,28 This model does not explicitly contain temperature, and one could consider that it corresponds to the low-temperature limit. The model can, however, predict general trends if one takes into account the dependence of W, κ, and r on temperature, provided that this dependence is appreciable. One of the modifications of this model consists in a more explicit description of the vesicle−substrate interaction by including temperature.59 With this modification, the model predicts that a vesicle becomes less deformed with increasing temperature even if the model ingredients (e.g., the expression for the vesicle−substrate interaction) are considered to be independent of temperature. The effect predicted is modest, and consistent with what we observe in the case of adsorption of DOPC lipid vesicle on titanium oxide. In general, the situation seems, however, to be more complex especially for DPPC lipid vesicles, because, as already noticed, all three parameters, W, κ, and r, may or do depend on temperature. For example, the simplest phenomenological DLVO-type model of the vesicle−substrate interaction includes the van der Waals, double-layer electrostatic, and hydration forces (see, e.g., ref 60 and references therein), and each of these forces depends on temperature at least to some extent. According to this model, the repulsive part of the potential is, e.g., often controlled by the hydration forces,61 which strongly depend on the short-range order in the location of water molecules and accordingly on temperature. In the case of DPPC lipid vesicles, the bending constant is also often expected to be dependent on temperature. From these perspectives (see Introduction), our observation that the shape of DPPC lipid vesicles is nearly independent of temperature might not initially appear to be what one might expect. In principle, this observation can, however, be rationalized in at least two ways. First, different temperature-dependent factors may

compensate one another. Our second remark is that the lipid bilayer bending constants are usually measured for essentially planar membranes with very small curvatures. In contrast, the vesicles used in our experiments are small and have relatively large curvature. The bending constant may be much higher in this case, and its dependence on temperature may be weaker than that expected for DPPC lipid vesicles. On silicon oxide, the shape deformation of adsorbed DPPC lipid vesicles increased with increasing temperature. This effect is in line with what one would expect and can be explained by the temperature dependence of one of the factors discussed above. In combination, the results obtained for silicon oxide and titanium oxide are consistent because fluid-phase lipid vesicles generally rupture on silicon oxide, but remain intact on titanium oxide. Furthermore, the results indicate that additional factors (e.g., vesicle−substrate contact energy) may strongly influence the manner in which temperature affects the deformation of adsorbed vesicles. To the best of our knowledge, our study is the first report explicitly illustrating this complexity in detail. In summary, we conclude that the effect of temperature on the deformation of adsorbed vesicles on solid supports depends on an interplay of various, often hidden factors, justifying the need for experimental investigations of this kind. Indeed, the role of temperature is multifaceted and the collective sum of its contributions may promote, hinder, or have negligible effect on the deformation of adsorbed vesicles. Taken together, our findings underscore the high surface-sensitivity of the LSPR technique to scrutinize detailed aspects of vesicle adsorption and demonstrate its potential for clarifying related aspects of soft matter adsorption in general.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Author Contributions +

E.O. and J.A.J. contributed equally to this work.

Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors wish to acknowledge support from the National Research Foundation (NRF -NRFF2011-01), the National Medical Research Council (NMRC/CBRG/0005/2012), and Nanyang Technological University to N.J.C. J.A.J. is a recipient of the Nanyang President’s Graduate Scholarship. V.P.Zh. is a recipient of the Tan Chin Tuan Exchange Fellowship at Nanyang Technological University.

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