Self-Assembly of Bioconjugated Amphiphilic Mesogens Having

Jan 26, 2016 - Thermotropic liquid crystal films for biosensors and beyond. Piotr Popov , Elizabeth K. Mann , Antal Jákli. Journal of Materials Chemis...
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Self-Assembly of Bioconjugated Amphiphilic Mesogens Having Specific Binding Moieties at Aqueous−Liquid Crystal Interfaces Hiroki Eimura,† Daniel S. Miller,‡ Xiaoguang Wang,‡ Nicholas L. Abbott,*,‡ and Takashi Kato*,† †

Department of Chemistry and Biotechnology, School of Engineering, The University of Tokyo, Hongo, Bunkyo-ku, Tokyo 113-8656, Japan ‡ Department of Chemical and Biological Engineering, University of WisconsinMadison, 1415 Engineering Drive, Madison, Wisconsin 53706, United States S Supporting Information *

ABSTRACT: New bioconjugated amphiphilic mesogens with recognition groups have been synthesized, and their selfassembly behavior has been characterized at aqueous−liquid crystal (LC) interfaces. Specifically, the rod-shaped 2,3difluoro-4′-(4-trans-pentylcyclohexyl)biphenyl-based mesogen was conjugated with either biotin or the arginine−glycine− aspartic acid (RGD) peptide sequence through a tetraethylene glycol chain. Langmuir film measurements revealed that the two biorecognition moieties lead to very different surface pressure−area isotherms, indicating that biotin and RGD have distinct effects on self-assembled monolayers formed by these bioconjugated mesogens at aqueous surfaces. Measurements of the surface-induced orientations of LCs exhibited by biotin-conjugated mesogens mixed with the room-temperature nematic LC 4cyano-4′-pentylbiphenyl (5CB) revealed formation of cholesteric phases (consistent with the chiral nature of biotin) and evidence of the presence of the conjugated mesogen at aqueous interfaces. Preliminary measurements based on fluorescence measurements using Texas Red-labeled streptavidin confirmed that the biotin mesogen is located at the interface and capable of specific recognition of streptavidin. Overall, these results demonstrate that bioconjugated mesogens provide the basis of a general and facile method for the introduction of biological recognition functionality at aqueous−LC interface. These LC interfaces have mobility, elastic properties and responsiveness that are distinct from past studies of biorecognition groups presented at the interface of a solid or isotropic liquid, and the results thus provide a new approach for the introduction of biorecognition groups into this important and promising class of interface for biological and analytical applications.

1. INTRODUCTION Liquid crystals (LCs) have been extensively studied as functional soft materials due to their fluidity and anisotropy.1−15 These properties enable the design of LC-containing materials that respond to a wide range of stimuli, including electric fields,16,17 light,18−22 ions,9,23 bases,24 and acids.25,26 In addition, it is well-known that the ordering6,7,27−37 and phase38,39 of LCs are strongly influenced by the chemical functionality and topography of the surfaces confining them. This surface sensitivity provides the basis for one approach to the design of LC sensors.7,40,41 In particular, aqueous−LC interfaces6,7,42−53 have attracted much attention as promising responsive interfaces at which a wide range of biological phenomena can be reported by ordering the transitions in the LC. For example, aqueous−LC interfaces have been used to couple specific binding events between biomolecules,42−45 enzymatic reactions,42,46−48 and the interactions of living cells with surfaces49 with the ordering transition in LCs, which can be readily transduced using optical methods. We emphasize that a key motivation for the use of LCs in this context is that LC-based sensing does not require the use of fluorescent or radioactive labels. © 2016 American Chemical Society

While the above-described studies serve to illustrate the promise of LC−aqueous interfaces for reporting biological phenomena at interfaces, there still exists a need for the development of general and facile approaches that permit the presentation of biological recognition groups (that mediate interactions with biological species, including proteins; Figure 1) from LC interfaces. In particular, the biological recognition group must be introduced into the interface in a manner that permits its interaction with the targeted biological species. Furthermore, it is necessary to understand how the presence of the biological recognition group influences the ordering of the LC. In many cases, due to the distinct optical appearance of homeotropically aligned LCs (they appear dark between crossed polarizers), it is advantageous to have the initial states of the LC interface exhibit a homeotropic orientation. A previous study42 has focused on the use of biological lipids and proteins adsorbed onto LC interfaces as a means of introducing biological recognition groups. Our intention is to explore an Received: December 8, 2015 Revised: January 26, 2016 Published: January 26, 2016 1170

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this article (Figure 2). We characterized the influence of the introduction of the binding groups (biotin and RGD) on the self-assembly of these mesogens at interfaces and explored the influence of these self-assembly processes on the ordering of the nematic phases of 4-cyano-4′-pentylbiphenyl (5CB) containing mesogens 1 and 2. Specifically, the self-assembly behavior of 1 and 2 at air−aqueous, air−LC, and aqueous−LC interfaces was examined by Langmuir monolayer experiments and polarized optical microscopic observations. Specific and nonspecific interactions of streptavidin to aqueous−LC interfaces containing 1 and 3 were studied by epifluorescence microscopic measurements.

Figure 1. Schematic illustration of our approach to the design of aqueous-liquid crystal (LC) interfaces prepared using tailored mesogens having biorecognition epitopes for targeted proteins. For this purpose, the interface needs to present the specific binding moieties to the aqueous phase and induce homeotropic anchoring in the absence of targeted proteins.

2. RESULTS AND DISCUSSION 2.1. Molecular Design and Synthesis. Compounds 1 and 2 were designed from 3. Compound 3 is an LC molecule having a hydrophobic mesogenic core and a hydrophilic tetra(ethylene oxide) chain.59 The amphiphilic properties of this molecule lead to spontaneous partitioning of 3 from the bulk of an LC phase onto the aqueous−LC interface.51 As noted above, an understanding of the interfacial self-assembly behavior of these molecules is needed to rationally prepare aqueous−LC interfaces that present specific binding moieties. In particular, by coupling the recognition group to a mesogen, we anticipated that the presentation of the recognition group at the LC interface would influence the surface anchoring of the LC.51 Thus, amphiphilic mesogen 3 was used to design bioconjugated amphiphilic mesogens 1 and 2. Compounds 1 and 2 have a biotin moiety and RGD sequence as specific binding moieties, respectively. Past studies have established that biotin forms strong noncovalent interactions with streptavidin.54 The binding constant between biotin and streptavidin is 1015 M−1. Due to this high affinity, the biotin−streptavidin interaction is a widely used model interaction in studies of specific binding events in biological systems. The RGD peptide sequence is found in extracellular matrices, and it is known to

approach based on the introduction of a functional mesogen into a LC material and its subsequent self-assembly at the LC interface. The approach builds on our previous study in which we demonstrated that nonspecific binding and alignment of LCs at aqueous−LC interfaces can be tuned by using a mesogen conjugated to a tetraethylene oxide moiety.51 However, the design of bioconjugated mesogens and their self-assembly at aqueous−LC interfaces has not been studied. Herein, we report the design and synthesis of two bioconjugated amphiphilic mesogens, 1 and 2, and characterize their self-assembly behavior at aqueous−LC interfaces. The focus of this article is not directed to the characterization of biological phenomena at these interfaces but rather to the development of an understanding of how these new bioconjugated mesogens assemble at LC−aqueous interfaces and how they impact the ordering of the LC. This knowledge will enable future studies of biological phenomena at LC− aqueous interfaces.42 Mesogens 1 and 2, which include a biotin54 moiety and an arginine−glycine−aspartic acid (RGD) motif,55−58 respectively, are the focus of the study reported in

Figure 2. Chemical structures of compounds 1−3 and 5CB. 1171

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Chemistry of Materials bind to cell surface proteins called integrins.55 The interaction of the RGD sequence with integrins has been widely studied in the context of cell biology and regenerative medicine because RGD-mediated binding underlies the attachment of cells to surfaces and in turn influences a wide range of cell behaviors.56−58 Our focus on RGD-conjugated mesogens is motivated by the potential future use of these LC systems for reporting biochemical and physical interactions of cells with surfaces without the need for fluorescent or isotopic labels. It is expected that cell adhesion between RGD motifs and integrins can be coupled with an LC ordering transition induced by the binding event, which is followed by a change in optical appearance. Compound 1 was synthesized by 1-ethyl-3-(3dimethylaminopropyl)carbodiimide hydrochloride (EDC) coupling of 3 and biotin (Scheme S1). Compound 2 was synthesized by coupling protected RGD motifs60 with carboxylic derivatives of 3, followed by deprotection using trifluoroacetic acid (Scheme S2). 2.2. Langmuir Film of Amphiphilic Mesogens at Air− Aqueous Interfaces. We first report measurements of monolayer properties of 1−3 at air−aqueous interfaces because the monolayer behavior of these molecules is used when interpreting the effects of the amphiphilic mesogens on anchoring at aqueous−LC interfaces in subsequent sections of this article.61,62 Specifically, the surface pressure (π)−area isotherms of monolayers of 1−3 (Figure 3 and Table 1) were

the interface, inspection of Figure 3 reveals that the surface isotherm of 1 is similar to that of compound 3, whereas the surface isotherm of 1 is identical to that of 2 in the expanded state. We note that past studies of compound 3 have been interpreted to indicate that compound 3 assumes an orientation that is close to perpendicular to the aqueous interface. This result leads us to conclude that compound 1 likely exists in two distinct configurational states, depending on whether the monolayer is in the expanded or condensed phase. This result is significant because the configurational state of the mesogen at the interface will influence the presentation of the biotin group. Specifically, when the monolayer is in the condensed state, the terminal biotin moieties of compound 1 will be presented to the aqueous phase (Figure 4), as was observed previously for

Figure 4. Schematic illustration of self-assembled structures of 1 consistent with Langmuir film measurements in the condensed state.

the hydroxyl groups of 3. These results suggest that the aqueous−LC interfaces prepared using compound 1 will likely present compound 1 in a manner that permits biorecognition to be performed via the biotin group. On the other hand, compound 2 does not show evidence of a phase transition in isotherm measurements, which suggests that the RGD group may sustain the monolayer in the expanded state and prevent the formation of a condensed phase. This may be because the molecular shape of 2 is bulkier than those of 1 and 3 due to the side chain in the RGD motif, leading to a different conformation at high area density. A difference in molecular shape is also confirmed by the larger molecular area of the π− area isotherm for 2 at collapse compared to that of 1 and 3. Accordingly, these results suggest that the RGD moieties of 2 may not be fully accessible from the aqueous phase. We emphasize that the monolayers formed by compounds 1−3 are stable below the above-mentioned collapse pressures. Specifically, we measured the time-dependent changes in surface pressure for monolayers of 1−3 at a high density of molecules. As shown in Figure 5, the surface pressures generated by monolayers of 1 and 3 are maintained for 10 min at area per molecule values of 40 Å2. Similar results were obtained with monolayers of 2 (see Figure S1). Overall, these results indicate that the monolayers of 1−3 are stable at the air−water interface over a range of surface densities.

Figure 3. Surface pressure (π)−area density isotherms of monolayers of compounds 1 (solid blue line), 2 (solid red line), and 3 (solid black line). Aqueous subphase is phosphate buffered saline (PBS) solution.

Table 1. Comparison of Three Key Parameters Obtained from the Langmuir Isotherms for Monolayers Formed by Compounds 1−3

1 2 3

surface pressure at collapse (mN m−1)

molecular area at collapse (Å2)

onset of surface pressure (Å2)

46.0 ± 1.3 35.9 ± 1.6 46.5 ± 0.4

29.0 ± 0.4 44.7 ± 1.1 28.6 ± 2.1

150.9 ± 7.6 196.8 ± 4.8 66.6 ± 5.4

measured to provide insight into the orientation of these molecules at the interface and thus the likely presentation of the recognition groups. We first note that monolayers of 1 and 3 (46.0 ± 1.3 and 46.5 ± 0.4 mN m−1, respectively) collapse at surface pressures that are higher than those of 2 (35.9 ± 1.6 mN m−1). This first difference hints that compounds 1 and 3 form more closely packed and better organized assemblies at the interface than compound 2. Significantly, for the isotherm of compound 1, a phase transition is observed at molecular areas of approximately 80 Å2. Within the condensed state on 1172

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polarized optical micrographs of 5CB or 5CB/1 mixtures exposed to air, thus documenting the effects of 1 on the bulk LC phase and alignment at the air−LC interface. These measurements were performed by filling the pores of 20 μm thick gold electron microscopy grids, which were supported on octadecyltrichlorosilane (OTS)-treated glass, with the LC (Figure 6a).51 This optical cell enables LC alignment at LC interfaces to be characterized easily because sample thickness (20 μm) and LC alignment are defined. We observed that 5CB and 5CB containing 0.01 and 0.1 wt % of 1 generated a dark optical appearance under crossed polarizers except for the region contacting the gold grid (Figures 6b and S2a,b). In addition to the dark appearance, the cross-shaped patterns obtained when using conoscopy (Figures 6c and S2c,d) confirm that the LC is within the wells of the grid and that the LC at the air−LC interface assumes homeotropic (perpendicular) anchoring. However, 5CB with 1 wt % of 1 exhibited a fingerprint texture under crossed polarizers (Figure 6d,e), indicating the formation of a cholesteric phase. This is because 1 (the biotin group of 1) acts as a chiral dopant and induces formation of a helical LC phase. Fingerprint textures have been widely observed when cholesteric LCs are confined by two surfaces that cause homeotropic anchoring.63 Thus, the observation suggests that the cholesteric phase formed by 5CB with 1 also assumes a homeotropic orientation at the air− LC interface. Compared with the solubility of 1, that of 2 in 5CB is low. When using polarized light microscopy, we observed precipitates in 5CB containing 0.5 wt % of 2 (Figure S3). Furthermore, mixtures of 5CB/2 do not exhibit cholesteric phases due to the low miscibility of 2 with 5CB. The formation of hydrogen bonding or ionic interactions between RGD motifs of 2 likely underlies the low solubility and aggregation of 2 in 5CB. Due to the low solubility of 2 in 5CB, we subsequently focus on characterization of the surface properties of 5CB/1 rather than 5CB/2. 2.4. Effects of Amphiphilic Mesogens on LC Ordering at Aqueous Interfaces. Next, we examined the influence of 1 on the orientation ordering of mixtures of 5CB and 1 at aqueous interfaces. Figure 7 shows polarized optical micrographs of 5CB and 5CB/1 in contact with phosphate buffered saline (PBS) solution. When 5CB hosted in a TEM grid (see above) is immersed in PBS solution, the LC exhibits green and pink colors under crossed polarizers and white light illumination (Figure 7a). For mixtures of 5CB/1, we observe that the interference colors of the LC change with the increasing fraction of 1 (Figure 7b,c). These color changes are caused by changes in the optical retardance of the samples, suggesting that addition of 1 to 5CB induces a change in alignment of the LC at the aqueous−LC interface. We quantified the optical retardance of each LC film with a Berek compensator. The tilt angle of the LC relative to the aqueous−LC interfaces can be determined from the optical retardance, using previously detailed methods.51 As shown in Figure 8, the tilt angle at the aqueous interface changes continuously from 90° to approximately 10° as the fraction of 1 in the LC mixtures increases from 0 to 0.5 wt %. Here, we note that this simple analysis neglects the possible effects of twisting of the cholesteric phase (containing low concentrations of 1) on the optical retardance; thus, the angles in Figure 8 should be viewed as approximate. When combined with results obtained from Langmuir film measurements, our observation that 0.5 wt % of 1 in 5CB causes near-homeotropic anchoring at aqueous

Figure 5. Stability of Langmuir films for 1 and 3. Changes in the area density (dashed line) and surface pressure (solid line) for monolayers formed by compounds 1 (a) and 3 (b) over time. Aqueous subphase is phosphate buffered saline (PBS) solution.

2.3. Bulk Amphiphilic properties of bioconjugated

Properties of Mixtures of 5CB and Mesogens. Next, we investigated the bulk the nematic phases of 5CB into which the mesogens were dissolved. Figure 6 shows

Figure 6. Schematic illustration of an optical LC cell and orientation of LCs inside the cell exposed to air (a). In the optical LC cell, LCs are hosted within a 20 μm thick gold electron microscopy (EM) grid and octadecyltrichlorosilane (OTS)-treated glass. Optical micrographs (crossed polarizers) and conoscopic images of an optical LC cell for 5CB in contact with air (b, c). Optical micrograph (crossed polarizers) of an optical LC cell for 5CB with 1 wt % of 1 in contact with air (d, e). 1173

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Figure 9. Optical micrograph (crossed polarizers) of 5CB containing 1 wt % of 1 between rubbed polyimide-coated glass and air.

6e). To obtain 5CB/1 under hybrid anchoring conditions, rubbed polyimide-coated glass was used instead of OTS-treated glass and the period length of 5CB/1 in optical cells was measured, exposed to air. For homeotropic−homeotropic anchoring conditions, the periodic length of 5CB/1 between OTS-coated glass and air was observed. The periodic length of 5CB/1 (1 wt %) immersed in aqueous solution is 17.5 ± 3.5 μm (Figures 7d and S4). The value was similar to that of 5CB/ 1 under homeotropic−homeotropic anchoring rather than that of hybrid anchoring. From the comparison of the period lengths, therefore, we conclude that mixtures of 5CB and 1 (1 wt %) assume a homeotropic orientation at aqueous interfaces. We emphasize here that the conclusion that compound 1 causes homeotropic orientation of the LC is consistent with the interpretation of our Langmuir film studies in which compound 1 was found to adopt a conformation in which the hydrophobic tail was oriented close to the surface normal. 2.5. Binding Properties of Aqueous−LC Interfaces to Proteins in Aqueous Solution. To characterize the biorecognition properties of aqueous−LC interfaces prepared using 1, we performed fluorescence microscopic measurements of aqueous−LC interfaces for 5CB alone and 5CB containing 2 wt % of 1. Fluorescence images were captured after incubation of 1.0 μM Texas Red-conjugated streptavidin (TR−streptavidin) against the LC interface and subsequent rinsing of the interface by PBS. For 5CB alone, fluorescence intensity was detected because 5CB adsorbs streptavidin via nonspecific interactions (including, likely, hydrophobic interactions), as shown in a previous study.44 However, when 5CB contains 2 wt % of 1, the LC interface exhibits fluorescence intensity that is 1.5 times higher than that of 5CB alone (Figure 10). Two interpretations are possible for the increase of fluorescence intensity. First, there is an increase in the amount of streptavidin adsorbed at the aqueous−LC interface due to binding to 1. The other is that there is a change in the state of the bound streptavidin that leads to a change in the fluorescence yield from the adsorbed molecules. Specifically, past studies have reported that binding of biotin to fluorescent streptavidin can lead to a change in the photo quantum yield.66−68 However, when fluorescent streptavidin blocked with biotin was used in our experiments instead of nonblocked streptavidin as a control experiment, the difference in fluorescence intensity between 5CB and 5CB with 2 wt % of 1 disappeared. These results thus support the interpretation that compound 1 promotes binding of streptavidin to the LC interface through specific binding of the streptavidin and biotin moiety of 1, although the fluorescent intensity of blocked TR− streptavidin was lower than that of unblocked TR−streptavidin. To explain the decrease, we consider two possible reasons. The

Figure 7. Change of LC alignment with increasing fraction of 1. Optical micrographs (crossed polarizers) of an optical LC cell for 5CB (a) and 5CB/1 containing 0.01 wt % (b), 0.1 wt % (c), and 1 wt % (d) of 1 in contact with PBS solution.

Figure 8. Effects of the fraction of 1 on anchoring at the aqueous−LC interface. Tilt angle of LC at aqueous interface as a function of the fraction of 1 (black circle) added to 5CB. The tilt angle is defined in the inset and calculated from the retardance of each optical micrograph (crossed polarizers).

interfaces suggests that the biotin groups of 1 are likely presented toward the aqueous phase at the LC−aqueous interface and thus available for specific binding by proteins added to the aqueous solution. At higher concentrations of 1 (where the fingerprint pattern is seen), the relatively short pitch of the cholesteric phases prevents use of the methods employed above to determine the surface orientation of the LC. Alternatively, for those samples showing a fingerprint pattern, the period length is strongly affected by the anchoring conditions at interfaces.63−65 Specifically, the period length seen in a cholesteric phase with homeotropic−planar (hybrid) anchoring is approximately 3 times as long as that under homeotropic−homeotropic anchoring.64 We confirmed that cholesteric 5CB/1 (1 wt %) also showed a similar effect of anchoring on the period length. For 5CB/1, the period length under hybrid anchoring is 39.7 ± 1.5 μm (Figure 9), whereas the period length under homeotropic−homeotropic anchoring is 12.3 ± 1.0 μm (Figure 1174

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that streptavidin can bind specifically to 1 at the LC interface is consistent with the conformation of 1 (biotin extended toward the aqueous phase) interpreted from our Langmuir film studies. Finally, we made some exploratory observations to determine if the protein binding events described above resulted in changes in the ordering of the LC at the aqueous−5CB/1 interface. In these measurements, we observed LC samples containing 0.01 wt % 1 to exhibit a change in retardance upon incubation in PBS solution containing 1.0 μM streptavidin for 90 min (Figure 12). At other concentrations of 1 in 5CB,

Figure 10. Fluorescence intensities of Texas Red-labeled streptavidin unblocked (gray) and blocked (white) with biotin adsorbed at aqueous−LC interfaces of 5CB and 5CB containing 2 wt % of 1.

first possible reason is that binding of biotin to streptavidin decreases the nonspecific adsorption of streptavidin on the aqueous−LC interface. The other is that Texas Red in blocked TR−streptavidin was more highly affected by photoquenching derived from the interaction of the fluorophores with 5CB than that of unblocked TR−streptavidin.69 A previous study has shown that addition of 3 into 5CB is an effective strategy for minimizing nonspecific binding of bovine serum albumin (BSA) at aqueous−LC interfaces.51 We also confirmed that this approach was applicable to nonspecific binding of streptavidin. Lower fluorescent intensity at aqueous−LC interfaces was obtained when using 5CB with 5 wt % of 3 as compared to that of 5CB alone (Figure 11).

Figure 12. Optical micrographs (crossed polarizers) of an optical cell of 5CB containing 0.01 wt % of 1 before (a) and after (b) a 90 min incubation of 1.0 μM streptavidin in PBS.

however, we did not observe a significant change of optical appearance or period length after incubation of PBS solution containing streptavidin. For the 5CB sample containing 0.01 wt % of 1, the change in retardance corresponds to only about a 7° tilt angle change. Here, we emphasize that these measurements were exploratory in nature and that further optimization of the composition of the LC is needed to couple the LC ordering transitions to the protein binding event. The key result reported above with compound 1 is that LC interfaces containing compound 1 do present biotin in a way that is accessible to proteins and that compound 1 does induce the LC to adopt a well-defined orientation at aqueous interfaces (Figure 11).

3. CONCLUSIONS We have designed and synthesized two mesogens, 1 and 2, having a biotin and RGD motif, respectively. We characterized the self-assembly of these mesogens at the surface of water and also at aqueous−LC interfaces to provide fundamental insight into the manner by which the introduction of biological recognition groups changes the properties of mesogens at LC interfaces. Our results, when combined, indicate that compound 1 can be used to create aqueous interfaces of LCs that show specific binding properties for streptavidin. The specific binding behavior of the LC interface was observed by simple mixing of 5CB and 1 and the spontaneous adsorption and self-assembly of 1 at the aqueous interface of the LC. Overall, the results described in this article suggest the basis of a new approach to the realization of functional aqueous−LC interfaces through synthesis of tailored amphiphilic mesogens for a specific biological interaction. Further studies, which will involve optimized molecules and LC compositions, are required to fully control the adsorption of biomolecules at aqueous−LC interfaces and to generate ordering transitions in the LCs triggered by protein binding events. However, this study provides essential foundational knowledge for the design rule of

Figure 11. Fluorescence intensities of Texas Red-labeled streptavidin adsorbed at aqueous−LC interfaces of 5CB, 5CB with 2 wt % of 1, 5CB with 5 wt % of 3, and 5CB containing 2 wt % of 1 and 5 wt % of 3.

Furthermore, the introduction of 1 into 5CB containing 5 wt % of 3 and subsequent incubation of the interface again with labeled streptavidin result in a fluorescence intensity that was comparable to the intensity of 5CB containing 1 without 3. From these fluorescence measurements, we concluded that 1 and 3, when combined as additives to 5CB, enable control of both specific and nonspecific binding of streptavidin to aqueous−LC interfaces. We also note that the conclusion 1175

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4.7. Epifluorescence Microscopic Observation of Fluorescently Labeled Proteins at the Aqueous−LC Interfaces. TR− streptavidin was adsorbed to an aqueous−LC interface by a 90 min incubation of LC mixtures in 400 μL of PBS solution containing 1.0 μM TR−streptavidin. Prior to observation, the incubated PBS solution was diluted with 2 mL of PBS solution in order to remove the fluorescently labeled streptavidin in the solution. An Olympus IX 81 or IX71 was used to capture images of aqueous−LC interfaces adsorbing TR−streptavidin. Fluorescence intensity of each aqueous−LC interface was quantified using ImageJ (U.S. National Institutes of Health). Fluorescence intensity of an aqueous−5CB interface without TR− streptavidin was used as background fluorescence intensity. The fluorescence intensity for each LC interface in this article is the average of four different random locations away from the edges of the grids in each of two independent grids.

biorecognition LC materials for aqueous−LC interfaces having responsive properties to specific biological events.

4. EXPERIMENTAL SECTION 4.1. General. A JEOL JNM-ECX400 spectrometer was used for NMR measurements. Elemental analyses were performed on a PerkinElmer CHNS/O 2400 apparatus. MALDI TOF mass spectra were recorded on a Bruker Autoflex Speed TOF/TOF. Fourier transform infrared (FT-IR) spectra were carried out on a Jasco FT/IR6100 spectrometer on KBr plates. Olympus BX51 and BX60 polarizing optical microscopes equipped with a Mettler FP82HT hot stage were used for optical microscope observation. 4.2. Materials. All reagents for compounds synthesis were purchased from Aldrich, Kanto, Tokyo Kasei, or Wako. N,NDimethylformamide (DMF) and dichloromethane were purified using a Nikko Hansen ultimate solvent system before using them as reaction solvents. The other reagents were used without further purification. Unless otherwise mentioned, all reactions were performed under an argon atmosphere in dry solvents. 4-Cyano-4′-pentylbiphenyl (5CB) was purchased from EM Sciences or Tokyo Kasei. 5CB obtained from Tokyo Kasei was used after distillation with a SHIBATA GTO-1000 glass tube oven. Glass microscope slides were Fisher’s finest premium grade. Gold specimen grids were obtained from Electron Microscopy Sciences or Nissin EM. Texas Red (TR)− streptavidin was purchased from Invitrogen. Streptavidin was purchased from Aldrich. 4.3. Isotherm Measurements of Langmuir Films. A Nima 602A film balance (Coventry, England) equipped with a filter paper Wilhelmy plate was used for Langmuir film experiments. The temperature of the aqueous subphase was maintained at 25.0 °C by circulating water at a constant temperature. Each Langmuir film was prepared by spreading a chloroform solution and 20 min evaporation of chloroform at 25.0 °C. Isotherm measurements were performed by compression of the Langmuir films at a constant rate of 20 cm2 min−1. 4.4. Preparation of Mixtures for 5CB and Amphiphilic Mesogens. LC mixtures were prepared by vacuum evaporation of a chloroform solution containing 5CB and amphiphilic mesogens at room temperature. To remove chloroform, all samples were put under vacuum conditions for longer than 4 h. 4.5. Preparation of Optical Cells Composed of LCs. The optical cells filled with LCs were prepared by following the procedures described in one of our past reports.51 Briefly, OTS-treated glass was prepared by cleaning and coating with octadecyltrichlorosilane (OTS) as described in a published study. The LCs were hosted in the pores (283 μm × 283 μm) of a 20 μm thick gold specimen grid placed on the OTS-treated glass. To obtain a thin LC film with a constant film thickness, approximately 1 μL of the LC was put on the grid supported on the OTS-treated glass and excess LC was removed with a syringe. The LC film on the OTS-treated glass was immersed into 400 μL of PBS solution, forming a stable aqueous−LC interface. 4.6. Quantification of the Tilt Angle for the LC Molecules at the Aqueous−LC Interface. In order to quantitatively estimate the alignment of LC molecules at the aqueous−LC interface, the optical retardance (Δr) of the LC films in aqueous solution was measured using an Olympus U-CTB compensator. As described in our report,51 the tilt angle (θ) of the director at an aqueous−LC interface can be calculated from a following equation

⎛ d⎜ Δr ≈ ⎜ 0 ⎜ 2 2 ⎝ no sin

∫

none

( dz θ) + ne2 cos2( dz θ)

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.chemmater.5b04736. Synthesis and characterization of amphiphiles 1 and 2, area per molecule and surface pressure in a Langmuir film experiment of 2 as a function of time, and polarized optical micrographs of 5CB/1 and 5CB/2 in contact with air and aqueous solution (PDF)

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AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected] (N.L.A.). *E-mail: [email protected] (T.K.). Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was partially supported by a Grant-in-Aid for Scientific Research (No. 22107003) in the Innovative Area of “Fusion Materials” (Area No. 2206) from the Ministry of Education, Culture, Sports, Science and Technology (MEXT). Additional support was received from the National Science Foundation (under awards DMR-1121288 (MRSEC) and CBET-1263970) and the Army Research Office (W911-NF11-1-0251 and W911-NF-14-1-0140). We would like to thank Dr. Tatsuya Nishimura for help with the synthesis of compounds and Prof. Teruyuki Nagamune and Dr. Kosuke Minamihata for epifluorescence microscopic measurements at The University of Tokyo. H.E. is grateful for financial support from a Japan Society for the Promotion of Science (JSPS) Research Fellowship for Young Scientists and JSPS Program for Leading Graduate Schools (MERIT).

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⎞ ⎟ − no⎟dz ⎟ ⎠

REFERENCES

(1) Goodby, J. W.; Collings, P. J.; Kato, T.; Tschierske, C.; Gleeson, H.; Raynes, P. Handbook of Liquid Crystals, 2nd ed.; Wiley-VCH: Weinheim, Germany, 2014. (2) Saez, I. M.; Goodby, J. W. Supermolecular Liquid Crystals. Struct. Bonding (Berlin) 2008, 128, 1−62. (3) Tschierske, C. Development of Structural Complexity by Liquid Crystal Self-Assembly. Angew. Chem., Int. Ed. 2013, 52, 8828−8878. (4) Kato, T. Self-Assembly of Phase-Segregated Liquid Crystal Structures. Science 2002, 295, 2414−2418. (5) Broer, D. J.; Bastiaansen, C. M. W.; Debije, M. G.; Schenning, A. P. H. J. Functional Organic Materials Based on Polymerized Liquid-

where ne and no are the indices of refraction parallel and perpendicular to the optical axis of the LCs, respectively, an d is film thickness, which is 20 μm in this article. The reported values for indices of refraction of 5CB, ne = 1.711 and no = 1.5296 (λ = 632 nm at 25 °C),70 were used as constant values. The tilt angles in this article are the average of four different random locations from each of two independent grids. 1176

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Chemistry of Materials Crystal Monomers: Supramolecular Hydrogen-Bonded Systems. Angew. Chem., Int. Ed. 2012, 51, 7102−7109. (6) Lowe, A. M.; Abbott, N. L. Liquid Crystalline Materials for Biological Applications. Chem. Mater. 2012, 24, 746−758. (7) Carlton, R. J.; Hunter, J. T.; Miller, D. S.; Abbasi, R.; Mushenheim, P. C.; Tan, L. N.; Abbott, N. L. Chemical and Biological Sensing Using Liquid Crystals. Liq. Cryst. Rev. 2013, 1, 29−51. (8) Lester, C. L.; Smith, S. M.; Colson, C. D.; Guymon, C. A. Physical Properties of Hydrogels Synthesized from Lyotropic Liquid Crystalline Templates. Chem. Mater. 2003, 15, 3376−3384. (9) Binnemans, K. Ionic Liquid Crystals. Chem. Rev. 2005, 105, 4148−4204. (10) Gin, D. L.; Lu, X.; Nemade, P. R.; Pecinovsky, C. S.; Xu, Y.; Zhou, M. Recent Advances in the Design of Polymerizable Lyotropic Liquid-Crystal Assemblies for Heterogeneous Catalysis and Selective Separations. Adv. Funct. Mater. 2006, 16, 865−878. (11) Kato, T.; Mizoshita, N.; Kishimoto, K. Functional LiquidCrystalline Assemblies: Self-Organized Soft Materials. Angew. Chem., Int. Ed. 2006, 45, 38−68. (12) Hegmann, T.; Qi, H.; Marx, V. M. Nanoparticles in Liquid Crystals: Synthesis, Self-Assembly, Defect Formation and Potential Applications. J. Inorg. Organomet. Polym. Mater. 2007, 17, 483−508. (13) Woltman, S. J.; Jay, G. D.; Crawford, G. P. Liquid-Crystal Materials Find a New Order in Biomedical Applications. Nat. Mater. 2007, 6, 929−938. (14) Rowan, S. J.; Mather, P. T. Supramolecular Interactions in the Formation of Thermotropic Liquid Crystalline Polymers. Struct. Bonding (Berlin) 2008, 128, 119−149. (15) Kato, T. From Nanostructured Liquid Crystals to PolymerBased Electrolytes. Angew. Chem., Int. Ed. 2010, 49, 7847−7848. (16) Bremer, M.; Kirsch, P.; Klasen-Memmer, M.; Tarumi, K. The TV in Your Pocket: Development of Liquid-Crystal Materials for the New Millennium. Angew. Chem., Int. Ed. 2013, 52, 8880−8896. (17) Shimura, H.; Yoshio, M.; Hamasaki, A.; Mukai, T.; Ohno, H.; Kato, T. Electric-Field-Responsive Lithium-Ion Conductors of Propylenecarbonate-Based Columnar Liquid Crystals. Adv. Mater. 2009, 21, 1591−1594. (18) Ikeda, T. Photomodulation of Liquid Crystal Orientations for Photonic Applications. J. Mater. Chem. 2003, 13, 2037−2057. (19) Kawatsuki, N. Photoalignment and Photoinduced Molecular Reorientation of Photosensitive Materials. Chem. Lett. 2011, 40, 548− 554. (20) Ichimura, K.; Suzuki, Y.; Seki, T.; Hosoki, A.; Aoki, K. Reversible Change in Alignment Mode of Nematic Liquid Crystals Regulated Photochemically by Command Surfaces Modified with an Azobenzene Monolayer. Langmuir 1988, 4, 1214−1216. (21) Kawamoto, M.; Aoki, T.; Shiga, N.; Wada, T. Thermo and Photoresponsive Behavior of Liquid-Crystalline Helical Structures with the Aid of Dual Molecular Motions. Chem. Mater. 2009, 21, 564−572. (22) Soberats, B.; Uchida, E.; Yoshio, M.; Kagimoto, J.; Ohno, H.; Kato, T. Macroscopic Photocontrol of Ion-Transporting Pathways of a Nanostructured Imidazolium-Based Photoresponsive Liquid Crystal. J. Am. Chem. Soc. 2014, 136, 9552−9555. (23) Kanie, K.; Nishii, M.; Yasuda, T.; Taki, T.; Ujiie, S.; Kato, T. Self-Assembly of Thermotropic Liquid-Crystalline Folic Acid Derivatives: Hydrogen-Bonded Complexes Forming Layers and Columns. J. Mater. Chem. 2001, 11, 2875−2886. (24) Gonzalez, C. L.; Bastiaansen, C. W. M.; Lub, J.; Loos, J.; Lu, K.; Wondergem, H. J.; Broer, D. J. Nanoporous Membranes of HydrogenBridged Smectic Networks with Nanometer Transverse Pore Dimensions. Adv. Mater. 2008, 20, 1246−1252. (25) Tan, B.-H.; Yoshio, M.; Kato, T. Induction of Columnar and Smectic Phases for Spiropyran Derivatives: Effects of Acidichromism and Photochromism. Chem. - Asian J. 2008, 3, 534−541. (26) Su, X.; Voskian, S.; Hughes, R. P.; Aprahamian, I. Manipulating Liquid-Crystal Properties Using a pH Activated Hydrazone Switch. Angew. Chem., Int. Ed. 2013, 52, 10734−10739. (27) Jérôme, B. Surface Effects and Anchoring in Liquid Crystals. Rep. Prog. Phys. 1991, 54, 391−451.

(28) Patel, J. S.; Yokoyama, H. Continuous Anchoring Transition in Liquid Crystals. Nature 1993, 362, 525−527. (29) Ichimura, K. Photoalignment of Liquid-Crystal Systems. Chem. Rev. 2000, 100, 1847−1874. (30) Seki, T. New Strategies and Implications for the Photoalignment of Liquid Crystalline Polymers. Polym. J. 2014, 46, 751−768. (31) Kawata, K. Orientation Control and Fixation of Discotic Liquid Crystal. Chem. Rec. 2002, 2, 59−80. (32) Luk, Y.-Y.; Abbott, N. L. Surface-Driven Switching of Liquid Crystals Using Redox-Active Groups on Electrodes. Science 2003, 301, 623−626. (33) Price, A. D.; Schwartz, D. K. Anchoring of a Nematic Liquid Crystal on a Wettability Gradient. Langmuir 2006, 22, 9753−9759. (34) De Cupere, V.; Tant, J.; Viville, P.; Lazzaroni, R.; Osikowicz, W.; Salaneck, W. R.; Geerts, Y. H. Effect of Interfaces on the Alignment of a Discotic Liquid-Crystalline Phthalocyanine. Langmuir 2006, 22, 7798−7806. (35) Iinuma, Y.; Kishimoto, K.; Sagara, Y.; Yoshio, M.; Mukai, T.; Kobayashi, I.; Ohno, H.; Kato, T. Uniaxially Parallel Alignment of a Smectic a Liquid-Crystalline Rod-Coil Molecule and Its Lithium Salt Complexes Using Rubbed Polyimides. Macromolecules 2007, 40, 4874−4878. (36) Honglawan, A.; Beller, D. A.; Cavallaro, M.; Kamien, R. D.; Stebe, K. J.; Yang, S. Pillar-Assisted Epitaxial Assembly of Toric Focal Conic Domains of Smectic-A Liquid Crystals. Adv. Mater. 2011, 23, 5519−5523. (37) Zhang, R.; Zeng, X.; Prehm, M.; Liu, F.; Grimm, S.; Geuss, M.; Steinhart, M.; Tschierske, C.; Ungar, G. Honeycombs in Honeycombs: Complex Liquid Crystal Alumina Composite Mesostructures. ACS Nano 2014, 8, 4500−4509. (38) Boamfa, M. I.; Kim, M. W.; Maan, J. C.; Rasing, T. Observation of Surface and Bulk Phase Transitions in Nematic Liquid Crystals. Nature 2003, 421, 149−152. (39) Gbabode, G.; Dumont, N.; Quist, F.; Schweicher, G.; Moser, A.; Viville, P.; Lazzaroni, R.; Geerts, Y. H. Substrate-Induced Crystal Plastic Phase of a Discotic Liquid Crystal. Adv. Mater. 2012, 24, 658− 662. (40) Herzer, N.; Guneysu, H.; Davies, D. J.; Yildirim, D.; Vaccaro, A. R.; Broer, D. J.; Bastiaansen, C. W. M.; Schenning, A. P. H. J. Printable Optical Sensors Based on H-Bonded Supramolecular Cholesteric Liquid Crystal Networks. J. Am. Chem. Soc. 2012, 134, 7608−7611. (41) Jin, O.; Fu, D.; Ge, Y.; Wei, J.; Guo, J. Hydrogen-Bonded Chiral Molecular Switches: Photo-and Thermally-Reversible Switchable Full Range Color in the Self-Organized Helical Superstructure. New J. Chem. 2015, 39, 254−261. (42) Brake, J. M.; Daschner, M. K.; Luk, Y.-Y.; Abbott, N. L. Biomolecular Interactions at Phospholipid-Decorated Surfaces of Liquid Crystals. Science 2003, 302, 2094−2097. (43) Price, A. D.; Schwartz, D. K. DNA Hybridization-Induced Reorientation of Liquid Crystal Anchoring at the Nematic Liquid Crystal/Aqueous Interface. J. Am. Chem. Soc. 2008, 130, 8188−8194. (44) Tan, L. N.; Orler, V. J.; Abbott, N. L. Ordering Transitions Triggered by Specific Binding of Vesicles to Protein-Decorated Interfaces of Thermotropic Liquid Crystals. Langmuir 2012, 28, 6364−6376. (45) Noonan, P. S.; Roberts, R. H.; Schwartz, D. K. Liquid Crystal Reorientation Induced by Aptamer Conformational Changes. J. Am. Chem. Soc. 2013, 135, 5183−5189. (46) Park, J.-S.; Abbott, N. L. Ordering Transitions in Thermotropic Liquid Crystals Induced by the Interfacial Assembly and Enzymatic Processing of Oligopeptide Amphiphiles. Adv. Mater. 2008, 20, 1185− 1190. (47) Hartono, D.; Bi, X.; Yang, K.-L.; Yung, L.-Y. L. An AirSupported Liquid Crystal System for Real-Time and Label-Free Characterization of Phospholipases and Their Inhibitors. Adv. Funct. Mater. 2008, 18, 2938−2945. (48) Hartono, D.; Xue, C.-Y.; Yang, K.-L.; Yung, L.-Y. L. Decorating Liquid Crystal Surfaces with Proteins for Real-Time Detection of 1177

DOI: 10.1021/acs.chemmater.5b04736 Chem. Mater. 2016, 28, 1170−1178

Article

Chemistry of Materials Specific Protein−Protein Binding. Adv. Funct. Mater. 2009, 19, 3574− 3579. (49) Lockwood, N. A.; Mohr, J. C.; Ji, L.; Murphy, C. J.; Palecek, S. P.; de Pablo, J. J.; Abbott, N. L. Thermotropic Liquid Crystals as Substrates for Imaging the Reorganization of Matrigel by Human Embryonic Stem Cells. Adv. Funct. Mater. 2006, 16, 618−624. (50) Brake, J. M.; Mezera, A. D.; Abbott, N. L. Effect of Surfactant Structure on the Orientation of Liquid Crystals at Aqueous-Liquid Crystal Interfaces. Langmuir 2003, 19, 6436−6442. (51) Yang, Z.; Gupta, J. K.; Kishimoto, K.; Shoji, Y.; Kato, T.; Abbott, N. L. Design of Biomolecular Interfaces Using Liquid Crystals Containing Oligomeric Ethylene Glycol. Adv. Funct. Mater. 2010, 20, 2098−2106. (52) Khan, W.; Park, S.-Y. Liquid Crystal-Based Proton Sensitive Glucose Biosensor. Anal. Chem. 2014, 86, 1493−1501. (53) Zuo, F.; Liao, Z.; Zhao, C.; Qin, Z.; Li, X.; Zhang, C.; Liu, D. An Air-Supported Liquid Crystal System for Real-Time Reporting of Host−Guest Inclusion Events. Chem. Commun. 2014, 50, 1857−1860. (54) Blankenburg, R.; Meller, P.; Ringsdorf, H.; Salesse, C. Interaction Between Biotin Lipids and Streptavidin in Monolayers: Formation of Oriented Two-Dimensional Protein Domains Induced by Surface Recognition. Biochemistry 1989, 28, 8214−8221. (55) Pierschbacher, M. D.; Ruoslahti, E. Cell Attachment Activity of Fibronectin can be Duplicated by Small Synthetic Fragments of the Molecule. Nature 1984, 309, 30−33. (56) Luo, B.-H.; Carman, C. V.; Springer, T. A. Structural Basis of Integrin Regulation and Signaling. Annu. Rev. Immunol. 2007, 25, 619− 647. (57) Perlin, L.; MacNeil, S.; Rimmer, S. Production and Performance of Biomaterials Containing RGD Peptides. Soft Matter 2008, 4, 2331− 2349. (58) Huang, J.; Ding, J. Nanostructured Interfaces with RGD Arrays to Control Cell−Matrix Interaction. Soft Matter 2010, 6, 3395−3401. (59) Kishimoto, K.; Suzawa, T.; Yokota, T.; Mukai, T.; Ohno, H.; Kato, T. Nano-Segregated Polymeric Film Exhibiting High Ionic Conductivities. J. Am. Chem. Soc. 2005, 127, 15618−15623. (60) Welsh, D. J.; Smith, D. K. Comparing Dendritic and SelfAssembly Strategies to MultivalencyRGD Peptide−Integrin Interactions. Org. Biomol. Chem. 2011, 9, 4795−4801. (61) de Mul, M. N. G.; Mann, J. A. Multilayer Formation in Thin Films of Thermotropic Liquid Crystals at the Air-Water Interface. Langmuir 1994, 10, 2311−2316. (62) Carlton, R. J.; Ma, C. D.; Gupta, J. K.; Abbott, N. L. Influence of Specific Anions on the Orientational Ordering of Thermotropic Liquid Crystals at Aqueous Interfaces. Langmuir 2012, 28, 12796−12805. (63) Dierking, I. Textures of Liquid Crystals; Wiley-VCH: Weinheim, Germany, 2003. (64) Baudry, J.; Brazovskaia, M.; Lejcek, L.; Oswald, P.; Pirkl, S. Arch-Texture in Cholesteric Liquid Crystals. Liq. Cryst. 1996, 21, 893−901. (65) Liu, D.; Broer, D. J. Self-Assembled Dynamic 3D Fingerprints in Liquid-Crystal Coatings Towards Controllable Friction and Adhesion. Angew. Chem., Int. Ed. 2014, 53, 4542−4546. (66) Emans, N.; Biwersi, J.; Verkman, A. S. Imaging of Endosome Fusion in BHK Fibroblasts Based on a Novel Fluorimetric AvidinBiotin Binding Assay. Biophys. J. 1995, 69, 716−728. (67) Nicol, F.; Nir, S.; Szoka, F. C. Orientation of the Pore-Forming Peptide GALA in POPC Vesicles Determined by a BODIPY-Avidin/ Biotin Binding Assay. Biophys. J. 1999, 76, 2121−2141. (68) Guler, M. O.; Soukasene, S.; Hulvat, J. F.; Stupp, S. I. Presentation and Recognition of Biotin on Nanofibers Formed by Branched Peptide Amphiphiles. Nano Lett. 2005, 5, 249−252. (69) Meli, M.-V.; Lin, I.-H.; Abbott, N. L. Preparation of Microscopic and Planar Oil-Water Interfaces That Are Decorated with Prescribed Densities of Insoluble Amphiphiles. J. Am. Chem. Soc. 2008, 130, 4326−4333. (70) Blinov, L. M.; Chigrinov, V. G. Electrooptic Effects in Liquid Crystal Materials; Springer: New York, 1993.

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