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Polymerized Vesicles Containing Molecular Recognition Sites Markus Biesalski,*,†,‡ Raymond Tu,‡ and Matthew V. Tirrell‡ Institute for Microsystem Technology (IMTEK), Department of Chemistry and Physics of Interfaces, University of Freiburg, 79085 Freiburg, Germany, and Departments of Chemical Engineering and Materials, University of California Santa Barbara, Santa Barbara, California 93106 Received February 21, 2005. In Final Form: May 3, 2005 Vesicles are prepared from diacetylenic peptide amphiphiles that expose a molecular recognition site at the surface. The amphiphiles can be polymerized using UV light, and the resulting polymeric vesicles exhibit interesting chromatic responses that can be used for label-free detection of the interaction with a distinct protein in solution.
An ongoing challenge within the field of bioengineered materials is the development of novel “synthetic” materials that interact with biological materials in a desired specific fashion.1 Areas of applications for such materials progress from templates for engineering tissues 2 and targeted drug delivery vehicles3 to model systems for the investigation of complex biological phenomena4 and novel biosensor devices.5 With respect to the latter, researchers have started to simplify biological recognition phenomena to the interplay of small biological modules.6 An example for such an interplay of modules is the interaction of small peptide ligands found in extracellular matrix proteins possessing a distinct biospecific activity. In particular, engineering nature’s lock-and-key mechanism into synthetic materials may perhaps be the critical parameter to construct novel bioactive materials and devices. Amphiphilic molecules such as lipids or simple fatty acids, modified with a bioactive peptide sequence, so-called peptide amphiphiles,7,8 have been shown to be a versatile tool to construct systems that spontaneously self-assemble into organized two- (i.e., monolayers, bilayers, etc.)9-11 as well as three-dimensional structures (i.e., vesicles and rodlike micelles)12 capable of mimicking biological processes. The construction of biomimetic materials via a * To whom correspondence should be addressed. E-mail:
[email protected]. † University of Freiburg. ‡ University of California Santa Barbara. (1) Tirrell, M.; Kokkoli, E.; Biesalski, M. Surf. Sci. 2002, 500 (1-3), 61. (2) Drury, J. L.; Mooney, D. J. Biomaterials 2003, 24 (24), 4337. (3) Langer, R. AIChE J. 2000, 46, 1286. (4) Mammen, M.; Choi, S. K.; Whitesides, G. M. Angew. Chem., Int. Ed. 1998, 37, 2754. (5) Zieziulewicz, T. J.; Unfricht, D. W.; Hadjout, N.; Lynes, M. A.; Lawrence, D. A. Toxicol. Sci. 2003, 74 (2), 235. (6) Hartwell, L. H.; Hopfield, J. J.; Leibler, S.; Murray, A. W. Nature 1999, 402, C47. (7) Berndt, P.; Fields, G. B.; Tirrell, M. J. Am. Chem. Soc. 1995, 117, 9515. (8) Yu, Y. C.; Berndt, P.; Tirrell, M.; Fields, G. B. J. Am. Chem. Soc. 1996, 118, 12515. (9) Yu, Y. C.; Pakalns, T.; Dori, Y.; McCarthy, J. B.; Tirrell, M.; Fields, G. B. Methods Enzymol. 1997, 289, 571. (10) Fields, G. B.; Lauer, J. L.; Dori, Y.; Forns, P.; Yu, Y. C.; Tirrell, M. Biopolymers 1998, 47, 143. (11) Dori, Y.; Bianco-Peled, H.; Satija, S. K.; Fields, G. B.; McCarthy, J. B.; Tirrell, M. J. Biomed. Mater. Res. 2000, 50 (1), 75. (12) (a) Hartgerink, J. D.; Beniash, E.; Stupp, S. I. Science 2001, 294 (5547), 1684. (b) Tu, R.; Biesalski, M.; Tirrell, M. Biophys. J. 2002, 82 (1), 791.
self-assembly process of pre-synthesized molecular building blocks allows for a precise control of parameters such as peptide ligand surface concentration and presentation, that is, distance of the bioactive block from the surface as well as peptide conformation.7,11 Despite the versatility of bioactive bilayer films formed from peptide amphiphiles, for the use in any prospective biomedical or biosensing application one must consider the stability of these architectures. For example, it is wellknown that “supported” bilayer films once exposed to air can easily “roll-up” into nondefined aggregates.13 In addition, vesicles consisting of peptide amphiphilic molecules undergo thermodynamically driven aggregation, which ultimately leads to phase separation. A possible strategy to overcome this problem is to stabilize the assemblies by chemically cross-linking the amphiphilic molecules. Among others, an interesting class of polymerizable amphiphiles are diacetylenic fatty acids.14,15 Diacetylenic fatty acids can be polymerized under the influence of UV light, where the polymerization reaction is a 1-4 addition polymerization. It is further known that this reaction is topochemically controlled; that is, polymerization only occurs in the ordered (crystalline) state of the fatty acid. Once the fatty acid is polymerized in various assemblies (e.g., vesicles, monolayers, or bilayers), the polymer exhibits a strong absorption of visible light due to the electron-rich backbone that is formed. Moreover, the polymerized diacetylenic assemblies are known to show strong colorimetric responses upon changes in the environment, such as the temperature, pH, or addition of low molecular weight salts (for details see ref 14 and references therein). This chromism is attributed to a change of the backbone conformation of the polymer upon changes in the interaction of the respective headgroups or upon mechanical stress.14,15 Recently, it has been shown how one can make use of the chromism of polydiacetylenic assemblies to construct novel biosensing devices.16-20 Charych and co-workers have modified diacetylenic amphiphiles with specific (sugar) ligands that are known to interact with various (13) Dori, Y. Ph.D. Thesis, University of Minnesota, Minneapolis, MN, 1998. (14) Okada, S.; Peng, S.; Spevak, W.; Charych, D. Acc. Chem. Res. 1998, 31, 229. (15) Charych, D.; Nagy, J. O.; Spevak, W.; Bednarski, M. D. Science 1993, 261, 585.
10.1021/la0504558 CCC: $30.25 © 2005 American Chemical Society Published on Web 05/25/2005
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Figure 1. Schematic description of the solid-phase organic synthesis of the peptide amphiphile TDA-GRGDSP.
pathogens such as influenza viruses or bacterial toxins (a class of proteins).16-19 Both polymerized planar monolayers as well as polymerized vesicles have been used as novel bioanalytic devices to target biological materials. In addition, Stevens and Cheng have used polydiacetylenic assemblies to construct a novel “label-free” glucose sensor.20 In all cases the polydiacetylenic assemblies exhibit a “biochromic” effect upon binding of an analyte to the surface-bound ligand. However, in many specific interactions of proteins with their respective ligands, the latter consist of small bioactive peptide modules. So far it has not been shown that peptides can be immobilized on such polymeric assemblies in the same manner, with the goal to target specific protein-protein interactions, by the interplay of the attached peptide with their biological counterparts. In this letter we transfer the concept described by Charych and co-workers onto peptide-modified diacetylenic fatty acids that eventually can be used to mimic and sense protein-protein interactions. In particular, we report on the synthesis of a polymerizable peptide amphiphile that consists of a bioresponsive peptide headgroup, which is attached to a diacetylenic fatty acid. This polymerizable peptide amphiphile is further assembled into vesicular aggregates and stabilized via light-induced polymerization. Finally, in the first studies we test the bioactivity of the prepared peptide decorated polymerized vesicles, by using the vesicles as a sensor for a peptide receptor interaction. The peptide chosen for the preparation of the polymerizable bioactive amphiphile is the well-known tri-peptide arginine-glycine-aspartate (RGD), flanked by a glycine residue on one side and a serine-proline di-peptide on the other. Of course, we are aware of the fact that RGD peptides do not bind only one but rather a variety of (integrin) receptors found on the surfaces of different cells. However, as a proof-of-concept model the use of RGD peptides in this approach offers distinct advantages such as a large amount of references and literature-known responses to peptide-modified interfaces.1 The peptide amphiphile is prepared by Fmoc solid-phase organic synthesis as summarized in Figure 1. The synthesis of “non-polymerizable” peptide amphiphiles following this strategy has been described in detail elsewhere.7,8 Here we used a similar strategy for the (16) Reichert, A.; Nagy, J. O.; Spevak, W.; Charych D. J. Am. Chem. Soc. 1995, 117, 829. (17) Charych, D.; Cheng, Q.; Reichert, A.; Kuziemko, G.; Stroh, M.; Nagy, J. O.; Spevak, W.; Stevens, R. C. Chem. Biol. 1996, 3, 113. (18) Pan, J.; Charych, D. Langmuir 1997, 13, 1365. (19) Spevak, W.; Nagy, J. O.; Charych, D. H. Adv. Mater. 1995, 7, 85. (20) Cheng, Q.; Stevens, R. C. Adv. Mater. 1997, 9, 481.
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construction of the diacetylenic peptide amphiphile. Briefly, tricosadiynoic acid (TDA) is linked to the deprotected amine group of the first glycine residue. Subsequently, the whole peptide amphiphile as well as the sidechain protecting groups are cleaved of the solid phase, and the amphiphile is purified on a C4-RP-HPLC. After purification, the peptide amphiphile TDA-GRGDSP is characterized with Fourier transform infrared spectroscopy, 13C/1H NMR, and electrospray ionization quadrupole time-of-flight mass spectrometry, respectively (for details on the synthesis and characterization see Supporting Information). The diacetylenic peptide amphiphile is assembled into mixed vesicles (i.e., diacetylenic peptide amphiphile + nonmodified diacetylenic amphiphile) of defined composition and subsequently polymerized to yield stable bioactive colloids. As the background amphiphile for this purpose, the nonmodified TDA fatty acid has been used. The preparation of polymerized diacetylenic vesicles has been described extensively in the literature. Briefly, the monomeric amphiphiles are hydrated above Tm (Tm ≈ 56 °C), and the vesicular suspension is formed by disruption of the hydrated amphiphile films with ultrasound. After cooling below Tm, the vesicles are polymerized with UV light (t ) 2 min). The appearance of a blue color after the polymerization suggests the formation of the polymerized assemblies. Vesicles containing 100% peptide amphiphile also appear to be blue-colored after polymerization, indicating that the modification of the TDA headgroup with the peptide does not hinder the organization into the polymerizable assembly (see Supporting Information for details). The polymerized vesicles are further characterized with dynamic light scattering (DLS) and transmission electron microscopy (TEM). DLS results typically yield vesicle diameters of about 70-150 nm. Complementary studies with TEM reveal that the vesicles are more or less spherical in shape with vesicle diameters of about 100150 nm. The polymerized vesicles appear to be stable for weeks (no aggregation within the vesicular solution detectable, as evidenced from visible inspection and light scattering measurements). We have chosen to explore the polymerized vesicle’s ability to specifically recognize a (peptide) ligand-receptor interaction. In particular, here we have studied the interaction of the RGD ligand located on the surface of the vesicles with integrin receptors dissolved in phosphatebuffered solution (PBS, pH ) 7.4). A schematic description of this recognition event is shown in Figure 2. The receptor that has been used to this purpose is the R5β1 integrin, solubilized with small amounts of the surfactant TritonX100 in PBS. Note, to study the interaction of the receptor with its ligand, the receptor needs to be activated prior to the experiment. In nature calcium ions activate the receptor.21 However, in cases where no calcium ions are present, or where such bivalent ions are unfavorable to use, such as is the case with polydiacetylenic assemblies, the receptor can also be activated by using a specific antibody called TS2/16.22 Polymerized vesicles are prepared that contain 10% peptide amphiphile, and the vesicles are incubated for 30 min at 37 °C with the antibody-activated integrins at two different concentrations. As reference tests, we additionally incubated polymerized vesicles with just the antibody, (21) Lauffenburger, D. A., Lindermann, J. J., Eds. Receptors - Models for binding trafficking and signaling; Oxford Press: New York, 1993. (22) Davison, E.; Diaz, R. M.; Hart, I. R.; Santis, G.; Marshall, J. F. J. Virol. 1997, 71 (8), 6204.
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Figure 4. Visible-absorption spectra of polymerized vesicles containing 10 mol % peptide ligand on the surface. Polymerized vesicles (A); after incubation with free peptide GRGDSP (B); after incubation with “blocked” R5β1 integrins (C), after incubation with the activated integrins (D).
Figure 2. Schematic description of the molecular recognition of the peptide ligands located on the surface of the polymerized vesicles by the R5β1 integrin in solution (figure not drawn to scale).
Figure 3. Visible-absorption spectra of polymerized vesicles containing 10 mol % peptide ligand on the surface. Polymerized vesicles (A); after incubation with the antibody TS2/16 (B); after incubation with nonactivated R5β1 integrins (C); after incubation with activated R5β1 integrins (D, c ) 0.6 µmol/L; and E, c ) 1 µmol/L).
or nonactivated receptors, to test for any nonspecific interactions that might also induce some colorimetric responses of the vesicles. Figure 3 shows visible spectra taken 30 min after incubation. The pure polymerized vesicles (“raw TDA-peptide vesicle”, denoted as “A” in the figure) exhibit one large absorption maximum at 640 nm as well as a smaller maximum at 540 nm. Because of the strong absorbance at 640 nm the vesicular solution appeared blue-to-purple. The visible spectrum does not change significantly, if either the nonactivated integrins (“B”) or the antibody TS2/16 (“C”) alone is added to the polymerized vesicles. However, if the integrin receptor is activated prior to the incubation, the visible spectrum changes significantly. If 0.6 µmol/L of the activated R5β1 integrin is added (“D”), the absorbance at 640 nm decreases, while the absorbance at 540 nm increases. If the concentration of the receptor is increased to 1.0 µmol/ L, the change in the absorbency, that is, the colorimetric response, is even more significant. The change of the bluepurple solution to a red solution is even visible by the naked eye. Note, even at the highest integrin concentration chosen (1.0µmol/L) in the experiment, we stay below the approximate concentration of the peptide ligand, which was in this case about 12 µmol/L.
To test whether this colorimetric response observed with the activated integrins is due to a specific molecular recognition, we have incubated the active integrin receptors with free GRGDSP peptide in solution prior to the addition to the polymerized vesicles to “block” any specific interactions. Figure 4 shows the visible spectrum of the polymerized vesicles after incubation with “blocked” integrins (“C”) as well as the spectrum of the vesicles that are incubated with only free peptide (“B”). For comparison the spectrum of the pure TDA-peptide vesicles (“A”) as well as the spectrum of the vesicles incubated with the active integrins (“D”), both taken from Figure 3, are shown. It can be seen that the colorimetric response of the vesicles upon binding of the integrins to the peptide is significantly reduced, when the integrins have been blocked by the free peptide prior to the incubation. Hence, the latter result gives evidence that the colorimetric response of the peptide-modified polymerized vesicles is due to the molecular recognition of the receptor by the ligand. Charych and co-workers define the relative change of the blue phase (i.e., a stronger absorbance at 640 nm) to the red phase (stronger absorbance at 540 nm) as “colorimetric response” (CR):
CR ) (PB0 - PBf)/PB0 × 100%
(1)
Here, PB0 ) Ablue/(Ablue + Ared) × 100% is the initial percentage of the blue-form vesicles, with A being the integral absorption of either the blue (“640 nm”) or the red (“540 nm”) band. PBf is the final percentage of the blue form. Note, here the “blue” and the “red” vesicles are attributed to the appearance of the colored solutions rather than to the relative spectra measured. Figure 5 shows the colorimetric response of the polymerized vesicles as calculated from the visible spectra shown in Figures 3 and 4, using eq 1.23 It can be seen that the vesicles hardly show any response upon the addition of nonactivated integrins or upon the addition of the activator (TS2/16) alone. If activated integrins are considered, the vesicles show a significant colorimetric response up to about 35% of the initial “blue” phase. Finally, when the bioanalyte is blocked by the free peptide, prior to the sensing event, or when the free peptide is considered alone, the colorimetric response decreases significantly. (23) The error bars are estimated errors from the measurements (concentration of added analyte and concentrations of polymerized vesicles, as well as initial concentrations of peptide amphiphiles, respectively), as well as the analysis of the spectra (absorbance measurements).
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Upon incubation with activated integrins, the polymerized vesicles exhibit a colorimetric response that was even visible by the naked eye. Moreover, reference experiments show that this response is due to a specific molecular recognition event between the peptide ligand and the receptor. Further studies will be directed to show that this system is a promising candidate for the construction of (“label-free”) bioanalytic devices that might be useful to target specific protein interactions, in particular, if the ligands used consist of various small peptide motifs. Finally, in a separate communication we will report on the use of this polymerizable peptide amphiphile for the construction of stable planar surface coatings that can be used to manipulate cellular adhesion and spreading.24 Figure 5. Colorimetric response (“CR”) of polymerized TDApeptide vesicles upon incubation with nonactivated R5β1 integrin receptor (A); antibody TS2/16 (B), activated integrins (C, c ) 0.6 µmol/L; and D, c ) 1.0 µmol/L); free peptide GRGDSP (E); and “blocked” integrins (F).
In conclusion, we have synthesized a polymerizable peptide amphiphile that consists of a diacetylenic fatty acid attached to a peptide headgroup containing the peptide sequence RGD that is known to specifically interact with integrin receptors. The peptide amphiphile can be mixed with a polymerizable background amphiphile, and stable vesicles can be prepared by lightinduced polymerization of the assemblies in solution. In these initial studies on the bioactivity of the prepared vesicles, we have focused on the interaction of the peptide ligand located on the surface of the polymerized vesicles with antibody-activated R5β1 integrin receptors in solution.
Acknowledgment. M.B. would like to acknowledge financial support from the Deutsche Forschungsgemeinschaft (DFG) within the “Emmy-Noether-Program” (Award Nos. BI738/1-1, BI738/1-2) and the Fonds der chemischen Industrie. At UCSB, this work was supported by the NIRT and MRSEC Program of the National Science Foundation under Award Nos. CTS-0103516 and DMR-0080034. Supporting Information Available: Details of the synthesis and characterization of the polymerizable peptide amphiphile as well as the preparation, polymerization, and characterization of the vesicles can be found. This material is available free of charge via the Internet at http://pubs.acs.org. LA0504558 (24) Biesalski, M. A.; Knaebel, A.; Tu, R.; Tirrell, M. V. Submitted for publication.
