Supported Lipid Bilayer Templated J-Aggregate Growth: Role of

Jul 31, 2009 - New type of protein interaction observed. Biochemists have long thought that biomolecules must have or develop well-defined structures ...
1 downloads 11 Views 9MB Size
Download PDF
pubs.acs.org/Langmuir © 2009 American Chemical Society

Supported Lipid Bilayer Templated J-Aggregate Growth: Role of Stabilizing Cation-π Interactions and Headgroup Packing Gary C. H. Mo‡ and Christopher M. Yip*,†,‡,§ †

Department of Biochemistry and ‡Department of Chemical Engineering and Applied Chemistry and §Institute of Biomaterials and Biomedical Engineering, Terrence Donnelly Centre for Cellular and Biomolecular Research, University of Toronto, 160 College St, Toronto, Canada M5S 3E1 Received April 7, 2009. Revised Manuscript Received July 16, 2009

Controlling the self-assembly of molecules into specific structural motifs has important implications for the design of materials with specific optical properties. We report here the results of a correlated confocal fluorescence-atomic force microscopy (AFM) study of pseudoisocyanine iodide (PIC) self-assembly on supported lipid bilayers. Through judicious selection of bilayer headgroup packing and chemistry, two types of PIC J-aggregates, distinguishable by their absorbance spectra, and both exhibiting strong resonant fluorescence and bathochromic shifts in absorbance relative to the monomer, were isolated. Remarkably, selective templating can be achieved using different zwitterionic headgroups, producing J-aggregates that display a larger bathochromic shift than their solution counterparts. Our correlated confocal-AFM studies coupled with FT-IR spectroscopy suggested that zwitterionic phospholipids mediate J-aggregate formation through specific cation-π interactions between PIC and the lipid headgroups with the PIC molecules oriented largely perpendicular to the bilayer normal. The existence of the two isoforms further suggests that bilayer headgroup packing plays a key role in controlling interchromophore organization and subsequent aggregate nucleation and growth.

Introduction Molecular self-assembly describes the process by which molecules associate to form coherent, supramolecular architectures through noncovalent interactions. In many cases, the resulting optical and electronic properties of these structures are derived from the characteristics of the constituent molecules and their specific arrangement in the solid state. In order to design materials with specific optical, electronic, or magnetic properties, the mechanisms that govern the self-assembly process and the roles that chemical structure, conformation, and orientation of the constituent molecules play in defining these mechanisms need to be clearly identified. It is particularly critical that the structural characteristics of the first stable nuclei be identified since it necessarily serves as the template for subsequent growth. Since surfaces and interfaces are critical to the initial nucleation stage, exercising control over surface chemistry or structure provides a means of favoring the formation of specific or previously unknown structures. A well-known characteristic of cyanine dyes is their ability to self-assemble into so-called Jelley (J-) or Scheibe aggregates. Alternatively, dyes also assemble into H-aggregates (Figure 1). These new packing motifs create new electronic contacts between the constituent molecules and thus alternative pathways for exciton relaxation. In J-aggregates, the chromophores are arranged in a staircase motif as a series of ordered, onedimensional arrays. In the two-dimensional analogues, the one-dimensional arrays are packed in a herringbone or brickstone motif.1 Ordered cyanine J-aggregates are characterized by a sharp, bathochromic shift in absorption and an accompanying resonant fluorescence relative to the monomer, an effect thought to arise due to excitonic delocalization across the *Corresponding author: Tel 416-978-7853; Fax 416-978-4317; e-mail [email protected]. (1) Birkan, B.; Gulen, D.; Ozcelik, S. J. Phys. Chem. B 2006, 110, 10805.

Langmuir 2009, 25(18), 10719–10729

cyanine molecules in the aggregate.2 The self-assembly of cyanine dyes into the J-aggregate state has been reported to occur in solution,3,4 as a consequence of laser-induced aggregation,5 on polyelectrolyte multilayers,6,7 Langmuir-Blodgett films,8,9 nucleic acid,10 DNA helices,11 and mica or glass substrates.12-14 J-aggregates have been formed as LangmuirSchaefer films,15 in micellar dispersions,16,17 as crystals,18 on silicate cerasomes,19 functionalized quantum dots and nanotubes,20,21 and between amphiphiles.22,23 The formation of (2) Fidder, H.; Terpstra, J.; Wiersma, D. A. J. Chem. Phys. 1991, 94, 6895–6907. (3) Jelley, E. E. Nature 1936, 138, 1009. (4) Scheibe, G. Angew. Chem. 1936, 49, 563. (5) Tanaka, Y.; Yoshikawa, H.; Masuhara, H. J. Phys. Chem. C 2007, 111, 18457–18460. (6) Peyratout, C.; Donath, E.; Daehne, L. Photochem. Photobiol. Sci. 2002, 1, 87. (7) Liu, M.; Kira, A. Thin Solid Films 2000, 359, 104. (8) Czikkely, V.; Forsterling, H. D.; Kuhn, H. Chem. Phys. Lett. 1970, 6, 11. (9) Hada, H.; Hanawa, R.; Haraguchi, A.; Yonezawa, Y. J. Phys. Chem. 1985, 89, 560. (10) Achyuthan, K. E.; McClain, J. L.; Zhou, Z. J.; Whitten, D. G.; Branch, D. W. Anal. Sci. 2009, 25, 469–474. (11) Wang, M.; Silva, G. L.; Armitage, B. A. J. Am. Chem. Soc. 2000, 122, 9977. (12) Ono, S. S.; Yao, H.; Matsuoka, O.; Kawabata, R.; Kitamura, N.; Yamamoto, S. J. Phys. Chem. B 1999, 103, 6909. (13) Yao, H.; Sugiyama, S.; Kawabata, R.; Ikeda, H.; Matsuoka, O.; Yamamoto, S.; Kitamura, N. J. Phys. Chem. B 1999, 103, 4452. (14) Yao, H.; Ikeda, H.; Kitamura, N. J. Phys. Chem. B 1998, 102, 7691. (15) Tian, C. H.; Zoriniants, G.; Gronheid, R.; Van Der Auweraer, M.; De Schryver, F. C. Langmuir 2003, 19, 9831. (16) Kato, N.; Prime, J.; Katagiri, K.; Caruso, F. Langmuir 2004, 20, 5718. (17) Tatikolov, A. S.; Costa, S. M. B. Chem. Phys. Lett. 2001, 346, 233. (18) Gao, Y. N.; Zhang, X. M.; Ma, C. Q.; Li, X. Y.; Jiang, J. Z. J. Am. Chem. Soc. 2008, 130, 17044–17052. (19) Dai, Z. F.; Tian, W. J.; Yue, X. L.; Zheng, Z. Z.; Qi, J. J.; Tamai, N.; Kikuchi, J. Chem. Commun. 2009, 2032–2034. (20) Halpert, J. E.; Tischler, J. R.; Nair, G.; Walker, B. J.; Liu, W. H.; Bulovic, V.; Bawendi, M. G. J. Phys. Chem. C 2009, 113, 9986–9992. (21) Magadur, G.; Lauret, J. S.; Alain-Rizzo, V.; Voisin, C.; Roussignol, P.; Deleporte, E.; Delaire, J. A. ChemPhysChem 2008, 9, 1250–1253. (22) Li, X. W.; Zheng, Z. L.; Han, M. Y.; Chen, Z. P.; Zou, G. L. J. Phys. Chem. B 2007, 111, 4342–4348. (23) Seki, T.; Ichimura, K.; Ando, E. Langmuir 1988, 4, 1068.

Published on Web 07/31/2009

DOI: 10.1021/la901227h

10719

Article

Figure 1. Schematic representation of excitonic delocalization leading to the creation of new energy levels associated with J- or H-aggregate formation.

Mo and Yip

diffraction-limited, and while they may be able to confirm that a diffraction-limited region of interest has a specific optical characteristic, they are unable to confirm the size or number of the aggregate(s) responsible for that signal. We have previously exploited these attributes in correlated confocal or total internal reflection fluorescence-AFM imaging of protein assembly and dynamics in supported lipid bilayers.31-35 Using both coupled confocal-AFM microscopy and FT-IR spectroscopy, we obtained direct evidence of the role of bilayer phase and local chemistry on the stability and growth of oriented PIC-derived J-aggregates. Our results suggest that judicious selection of bilayer phase and chemistry can be used to nucleate the growth of specific J-aggregate types. These insights are critical for the design of substrates to facilitate J-aggregate formation as well as the de novo design of chromophores with specific optical and electronic properties.

Methods and Materials

J-aggregates is the principle behind membrane potential-sensitive dyes used to image excitable cells.24,25 Traditionally, J-aggregates have been identified by their characteristic fluorescence and absorbance properties; however, little work has been performed to correlate these characteristics with specific structural or morphological details. For example, in situ atomic force microscopy (AFM) and ex situ spectroscopy have been used to examine the orientation and structure of pseudoisocyanine iodide (PIC) aggregates grown from aqueous solution and deposited onto mica.13 Key questions, however, regarding J-aggregate formation from the solvated chromophore remain unanswered, including the critical nuclei size associated with the onset of J-aggregate behavior and how molecular packing within the nuclei influences exciton propagation. For our study, substrates with controlled chemistries and structures are necessary since we are particularly interested in the role of interfacial chemistry and structure on J-aggregate nucleation/growth and how such interfaces could be manipulated to guide the formation of aggregates with controlled optical properties. This prompted us to draw upon the success of others in using supported planar lipid bilayers (SPB) as templates for protein crystal growth26 and protein adsorption27,28 for our study of the initial stages of J-aggregate formation. In order to address the key questions of how local structure determines the characteristics of the J-aggregates, and the role of the substrate in facilitating J-aggregate formation, we applied a coupled imaging strategy that integrates confocal microscopy with in situ atomic force microscopy (AFM). The coupling of these two approaches provides a unique perspective since it affords direct mapping of topographical features by AFM with the spectral imaging capabilities of confocal fluorescence microscopy. This approach also addresses key limitations of the individual techniques. Conventionally, while in situ AFM has been shown to be a powerful tool for characterizing the selfassembly of molecular constituents into two- and three-dimensional structures,29,30 it only provides topographical information and is unable to obtain compositional or spectral information. Conventional confocal and related fluorescence techniques are

Pseudoisocyanine iodide (PIC; 1,10 -diethyl-2,20 -cyanine iodide; IUPAC: 1-ethyl-2-[(1-ethyl-2(1H)-quinolinylidene)methyl]quinolinium iodide) was purchased from Sigma-Aldrich (Oakville, Ontario) and used without further purification. Aqueous 364 μM PIC stock solutions were prepared, the concentration of which was determined by absorbance spectroscopy using a measured PIC extinction coefficient at 523 nm of 2.52  104 M-1 cm-1. Stock solutions were stored in the dark at room temperature and used within 4 days of preparation. All lipids were purchased from Avanti Polar Lipids (Alabaster, AL) either in chloroform solution or in powder form: 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC), 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1,2-distearoylsn-glycero-3-phosphate (DSPA), 1,2-dioleoyl-sn-glycero-3-phosphate (DOPA), 1,2-distearoyl-sn-glycero-3-[phospho-rac-(1glycerol)] (DSPG), 1,2-dioleoyl-sn-glycero-3-[phospho-rac-(1glycerol)] (DOPG), 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE), 1,2-distearoyl-sn-glycero-3-phosphoethanolamine (DSPE), 1,2-dimyristoyl-3-trimethylammonium propane (DMTAP), and 1,2-dioleoyl-3-trimethylammonium propane (DOTAP). All lipids were stored in chloroform at a stock concentration of 10 mg/mL. Lissamine Rhodamine B headgrouplabeled dihexadecylphosphatidylethanolamine (Rho-DHPE) was purchased from Molecular Probes (Invitrogen, Oakville, Ontario) and stored in chloroform at a stock concentration of 1-10 mg/mL. All aqueous solutions were prepared using 18 MΩ 3 cm water, doubly purified by an Elix 3/Milli-Q Pro system (MilliPore, Billerica, MA). Supported Lipid Bilayer Preparation. Supported planar bilayers were formed by in situ fusion of small unilamellar vesicles (SUV) in 10 mM HEPES, 150 mM NaCl, pH 7.4 buffer (HEPES buffer) directly onto freshly cleaved muscovite mica. To prepare the SUV suspension, 1 μmol of the desired lipid mixture was dissolved in chloroform and placed into a clean glass test tube. The chloroform was then removed by rotary evaporation under vacuum. The dry lipid film was rehydrated in 1 mL of HEPES buffer, and the resulting liposome suspension was subjected to ultrasonication in a water bath at a temperature above the phase transition temperature of the lipid to create small unilamellar vesicles ∼80 nm in diameter, as determined by dynamic light scattering. To prepare the bilayers, 100 μL of the liposome solution and 400 μL HEPES buffer were deposited onto freshly

(24) Sims, P. J.; Waggoner, A. S.; Wang, C. H.; Hoffman, J. F. Biochemistry 1974, 13, 3315. (25) Reers, M. Biochemistry 1991, 30, 4480. (26) Muller, D. J.; Janovjak, H.; Lehto, T.; Kuerschner, L.; Anderson, K. Prog. Biophys. Mol. Biol. 2002, 79, 1. (27) Mueller, H. J. Phys. Chem. B 2000, 104, 4552–4559. (28) Ross, E. E.; Spratt, T.; Liu, S. C.; Rozanski, L. J.; O’Brien, D. F.; Saavedra, S. S. Langmuir 2003, 19, 1766–1774. (29) Hillier, A. C.; Ward, M. D. Science 1994, 263, 1261–1264. (30) Last, J. A.; Hillier, A. C.; Hooks, D. E.; Maxson, J. B.; Ward, M. D. Chem. Mater. 1998, 10, 422–437.

(31) Shaw, J. E.; Alattia, J. R.; Verity, J. E.; Prive, G. G.; Yip, C. M. J. Struct. Biol. 2006, 154, 42–58. (32) Slade, A. L.; Schoeniger, J. S.; Sasaki, D. Y.; Yip, C. M. Biophys. J. 2006, 91, 4565–74. (33) Shaw, J. E.; Epand, R. F.; Epand, R. M.; Li, Z. G.; Bittman, R.; Yip, C. M. Biophys. J. 2006, 90, 2170–2178. (34) Alattia, J. R.; Shaw, J. E.; Yip, C. M.; Prive, G. G. J. Mol. Biol. 2006, 362, 943–53. (35) Shaw, J. E.; Epand, R. F.; Sinnathamby, K.; Li, Z. G.; Bittman, R.; Epand, R. M.; Yip, C. M. J. Struct. Biol. 2006, 155, 458–469.

10720 DOI: 10.1021/la901227h

Langmuir 2009, 25(18), 10719–10729

Mo and Yip

Article

cleaved muscovite mica, heated to 65 °C, slowly cooled, and flushed with 1 mL of a 4 mM CaCl2 HEPES buffer solution to remove excess liposomes and promote fusion. Excess calcium was removed via chelation with 1 mL of a 4 mM EDTA HEPES buffer solution. To minimize optical and chromatic artifacts, the mica substrates were ∼20-50 μm thick. All the bilayer experiments were conducted at room temperature using a custom-designed fluid cell holder. J-Aggregate Preparation. Aqueous PIC solution was added to the prepared bilayer sample as 100 μL aliquots of a 364 μM PIC stock solution to a final PIC concentration of 72.8 μM. To determine PIC mobility on the lipid bilayer, fluorescence recovery after photobleaching (FRAP) experiments were performed by confocal microscopy. To verify the stability of templated J-aggregates, 100 μL aliquots of a 5 M NaCl stock solution were added to the aggregates to a final concentration of 4 M. The resulting samples were examined by confocal fluorescence microscopy and visible light spectroscopy.

Visible Light Absorption and Fluorescence Spectroscopy. For solution calibration, a 1 mL aliquot of the solution was placed in QS-grade 10 mm path length quartz cuvettes (Hellma GmbH, Mullheim, Germany). All other samples were placed in a custom holder. Spectra were collected using an Ocean Optics (Dunedin, Fl) S2000 spectrometer equipped with either a tungsten-halogen or mercury-arc source. Final spectra were averaged without binning over at least 32 samples using the OOIBase32 software (Version 2.0.6.3, Ocean Optics). FT-IR Spectroscopy. Transmission IR spectra were obtained on a pressed KBr sample using a Nicolet Continuum IR microscope (Nicolet/Thermo Scientific, Madison WI) interfaced with a Nexus 670 FT-IR spectrometer equipped with a mid-IR source, KBr beamsplitter, and MCT-A liquid nitrogen cooled detector. All bilayer ATR-FT-IR measurements were performed using a rectangular Ge IRE element fitted to a homemade fluid cell mounted in the sample compartment of a Magna 750 (Nicolet/ Thermo Scientific) spectrometer equipped with a mid-IR source, KBr beamsplitter, and DTGS detector. All spectra were collected as 512 scans at a resolution of 1.0 cm-1 using the OMNIC version 5.2a software. For polarized data, a wire-grid polarizer (KRS-5, Harrick Scientific Products, Pleasantville, NY) was fitted to the entrance port of the spectrometer. Confocal Microscopy. Confocal fluorescence images were acquired using either an Olympus FluoView 300 or FluoView 500 (Olympus Canada, Markham, ON) scanning laser confocal microscope equipped with a Plan-APO 60, 1.40 NA, oil immersion objective as 12-bit, 1024  1024 pixel data sets using the Fluoview version 4.3 software at a typical line scan rate of ∼105 Hz with a 150 μm confocal pinhole setting. All images are shown without further manipulation, with the exception of cropping/scaling against the AFM images during correlation. A blue (effective power 640 μW, 488 nm, argon ion, Melles Griot) laser was used as the excitation source. The emitted light was passed through a 570 nm dichroic mirror into two photomultiplier channels set at isovoltage with unit gains (channel 1: 505-575 nm; channel 2: 575-635 nm; Figure 1 of the Supporting Information). The images are presented as superimposed intensity maps from the two channels. The merged images are pseudo-colored to indicate relative intensities and provide qualitative information on the emission spectrum. J-aggregates with a resonant emission at 574 nm are shown in cyan while those that emit at 580 nm appear red. Pseudo-colors were applied postacquisition with the FV10ASW Viewer software (Version 01.03.02.09, Olympus Canada). Relative yield values were obtained as ratio between the excitation and emission intensities and were compared to the corresponding values obtained for J-aggregates on DSPC. Spectral imaging was performed using the wavelength λ-scan mode of an Olympus FV1000 confocal microscope. Atomic Force Microscopy Imaging and Analysis. Contactand tapping-mode AFM imaging in fluid was performed with Langmuir 2009, 25(18), 10719–10729

Figure 2. Normalized spectra absorbance (solid line): J-aggregates templated by DSPC, DSPE bilayers. NaCl-complexed PIC in solution. Monomeric PIC in solution. Fluorescence (dashed line): J-aggregates templated by DSPC, DSPE bilayers, NaClcomplexed PIC in solution; and monomeric PIC in solution. The spectra are normalized to their respective maxima. The monomer (PIC) fluorescence spectrum is shifted vertically for clarity. Inset: structure of PIC. DNP-S C tips (manufacturer’s specification: length = 100 μm, k = 0.32 N/m; Veeco Instruments, Woodbury, NY) using a Digital Instruments Bioscope scanning probe microscope equipped with a Nanoscope IIIa controller and a dual-range J-scanner with a maximum possible scan size of 90 μm  90 μm (Veeco Instruments, Santa Barbara, CA). This system is mounted on an Olympus FV500 confocal microscope, making it possible to directly correlate the confocal and AFM images.36 All AFM images shown were acquired as 16-bit, 512  512 pixel images in tapping mode under amplitude feedback or in contact mode under deflection feedback, at a typical line scan rate of 1.2 Hz. All AFM images were analyzed with Digital Instruments Nanoscope software (version 5.30r3). Height images were subjected to zeroorder flatten and second-order X-axis plane-fit filters with scanline removal as necessary. Section analysis was performed using the Nanoscope software function. Manual correlation of the AFM and confocal images was performed using Adobe Photoshop (Version CS).

Results Spectroscopic Characteristics of Solution-Formed J-Aggregates. In solution, visible light spectroscopy revealed the expected PIC monomer absorption bands at 523 and 489 nm, corresponding to the well-known room temperature 0 r 0 and 1 r 0 transitions, respectively (Figure 2).37 In the presence of 4 M (36) Shaw, J. E.; Slade, A.; Yip, C. M. J. Am. Chem. Soc. 2003, 125, 11838–9. (37) Cooper, W. Chem. Phys. Lett. 1970, 7, 73.

DOI: 10.1021/la901227h

10721

Article

Mo and Yip

Figure 3. Confocal microscopy images of PIC on various zwitterionic lipid bilayers (DSPC, DPPC,DOPC, DSPE, POPE, DOPE). On DSPC, DSPE, and POPE, the strong sharp localized fluorescence is consistent with J-aggregates. The diffuse fluorescence seen on DPPC, DOPC, and DOPE is consistent with nonspecifically adsorbed PIC. Scale bar: 50 μm.

NaCl, the 0 r 0 transition was shifted to 530 nm, with the concomitant appearance of a new, sharp absorbance band at 574 nm (fwhm: 4.5 nm), a strong resonant fluorescence (fwhm: 7 nm), as has been previously reported.38 These spectroscopic characteristics define what we will call type I aggregates. It is worth noting that the J-aggregation reported by others on mica does not occur in the presence of HEPES buffer.13 Spectroscopic Characterization of Bilayer-Formed J-Aggregates. Confocal fluorescence microscopy revealed that, while J-aggregates readily formed on gel-phase zwitterionic DSPC, DSPE, and POPE bilayers, this did not occur on gel-phase charged bilayers (DSPG, DSPA, DMTAP) (Figures 3 and 4). Visible light spectroscopy revealed that aggregates formed on DSPC exhibited a strong absorbance at 580 nm, which was not observed in solution-grown J-aggregates. We describe these as type II aggregates. Interestingly, while on DSPC, only type II aggregates were seen, there was spectroscopic evidence of both type I and II aggregates on DSPE. We did not observe a hypsochromic shift in absorbance, which would be indicative (38) Lebedenko, A. N.; Guralchuk, G. Y.; Sorokin, A. V.; Yeflmova, S. L.; Malyukin, Y. V. J. Phys. Chem. B 2006, 110, 17772.

10722 DOI: 10.1021/la901227h

of H-aggregate formation and herringbone arrangements of J-aggregates, in any of these cases. FRAP experiments revealed that the gel-phase DSPC- and DSPE-grown J-aggregates were largely immobile in contrast to the freely diffusing aggregates observed on fluid-phase bilayers (data not shown). Among similar lipid headgroups, the effect of bilayer fluidity and order on aggregation was qualitatively assessed by their relative fluorescent yields. The stability of the templated aggregates was tested by the addition of 4 M NaCl, which is known to induce type I J-aggregate formation. While the addition of NaCl did not affect the morphology or the fluorescent characteristics of existing aggregates on the DSPC and DSPE bilayers (Figure 5), it did result in the deposition of solution-grown type I J-aggregates onto the DSPG, DSPA, and DMTAP bilayers. Substrate Preference for PIC J-Aggregation. To examine the effect of bilayer phase on J-aggregation, single-component DSPE and mixed gel-fluid phase DSPC and DOPC bilayers were used. While fusion of the DSPE SUV suspension onto mica resulted in homogeneous, molecularly smooth bilayers, the DSPC:DOPC bilayers formed by SUV fusion were characterized by the presence of topographically distinct domains. Correlated confocal-AFM imaging revealed that the Rho-DHPE labeled Langmuir 2009, 25(18), 10719–10729

Mo and Yip

Article

Figure 4. Confocal microscopy images showing the presence or lack of fluorescent J-aggregates on various single-component charged bilayers (DSPG, DSPA, DMTAP, DOPG, DOPA, and DOTAP). Scale bar: 50 μm.

fluid-phase DOPC domains were ∼5 nm tall, relative to the mica support, with the gel-phase DSPC domains extending ∼6.5 nm above the mica surface. Introduction of the PIC solution led to the formation of ∼1.5 nm thick type II J-aggregates on the gel-state DSPC domains (Figure 6a) whereas on the single-component DSPE bilayers, both type II and ∼2.5 nm thick type I J-aggregates formed (Figure 7). On DSPC, nonfluorescent PIC aggregates were seen as irregularly shaped ∼20-100 nm tall features on the supporting bilayer. These nonfluorescent aggregates were not colocalized with any fluorescence feature. On the DSPC bilayers, AFM imaging also revealed small ∼50 nm diameter, subdiffraction limit aggregates that we propose are type II PIC J-aggregates. It is important to note that these assignments were based on the AFM images, as these aggregates were not resolved by correlated confocal microscopy or spectroscopy. Since AFM imaging of several regions revealed that the surface coverage of these individual aggregates was on the order of 4% (by area), we cannot distinguish whether the absence of an optical signal was due to too few optically active aggregates within a diffractionlimited region or whether the individual aggregates themselves were too small to individually yield an excitonic signal. Polarized ATR FT-IR Spectroscopy of Bilayer-Associated PIC. To determine the approximate orientation of the Langmuir 2009, 25(18), 10719–10729

bilayer-associated PIC molecules, polarized ATR FT-IR spectroscopy was performed on samples prepared on a Ge ATR IRE element (Figure 8). The main spectral regions of interest are 2800-3000 cm-1 (lipid: -CH2-), 1730-1740 cm-1 (lipid CdO), 1400-1600 cm-1 (monomeric PIC aromatic), and 1160 cm-1 (monomeric PIC aliphatic C-N), with absorbances at 1610, 1558, 1510, and 1450 cm-1 corresponding to the ring-breathing modes of the central quinoline rings.39-41 Reference transmission IR spectra of PIC obtained in the absence of lipid were consistent with monomeric PIC. While ATR-IR spectra of DSPC, DSPE, DMTAP, and DOTAP bilayers after PIC exposure were largely inconclusive, spectra of samples prepared on DOPA, DOPG, DOPC, and DOPE bilayers supported the presence of monomeric PIC. It is important to note that we were unable to conclusively distinguish J-aggregates from other possible aggregate states from the IR spectra alone. Close inspection and comparison of the p- and s-polarized DOPG, DOPA, and DOPE ATR-IR spectra did reveal features consistent with a PIC orientation that is largely (39) Ilharco, L. M.; de Barros, R. B. Langmuir 2000, 16, 9331–9337. (40) Guo, C.; Aydin, M.; Zhu, H. R.; Akins, D. L. J. Phys. Chem. B 2002, 106, 5447–5454. (41) Bellamy, L. J. The Infrared Spectra of Complex Molecules, 3rd ed.; Chapman and Hall: London, 1975.

DOI: 10.1021/la901227h

10723

Article

Mo and Yip

Figure 5. Confocal microscopy images of fluorescent J-aggregates on gel-phase bilayers (DSPG, DSPA, DMTAP, DSPE, and DSPG) after addition of 4 M NaCl. Scale bar: 50 μm.

upright with the molecule’s central axis (along the aromatic core) perpendicular to the bilayer normal and its short alkyl tails oriented largely parallel with the bilayer normal and likely inserted into the bilayer (Figure 9). We would note that DFT models of the ground-state configuration of PIC describe an approximate C2 symmetry with the quinoline planes oriented at ∼46° relative to each other.40

Discussion It is clear that in order to favor J-aggregate formation the substrate must both stabilize the associating monomers and allow them to self-assemble into the appropriate motifs. This suggests a balance between monomer-substrate and monomer-monomer interactions. Bilayer-mediated PIC aggregation resulted in the formation of two distinct structural motifs. Spectroscopically, the type I J-aggregates resemble the solution-grown J-aggregates while the type II J-aggregates exhibit a larger bathochromic shift in absorbance relative to the monomer. Both types form preferentially on gel-phase domains. Fluorescent type II aggregates grown on the gel-phase domains of a mixed bilayer had a serrated appearance while those that formed on pure singlecomponent gel-state lipids tended to be small ∼100 nm ellipsoids. 10724 DOI: 10.1021/la901227h

Morphologically, these 2D structures are quite distinct from the structures previously reported for J-aggregates formed on mica. The formation and coalescence of 2D ellipsoidal islands into serrated grains suggests that they grew via a cooperative sequential adsorption mechanism.42 Our data support a model wherein growth of the optically active J-aggregates occurs through PIC monomer stacking at specific nucleation sites. Aggregate growth did not occur at the boundary between the gel- and fluid-phase domains but appeared to start at the center of (42) Evans, J. W. Rev. Mod. Phys. 1993, 65, 1281–1329. (43) Lis, L. J.; McAlister, M.; Fuller, N.; Rand, R. P.; Parsegian, V. A. Biophys. J. 1982, 37, 657. (44) Lindblom, G.; Rilfors, L.; Hauksson, J. B.; Brentel, I.; Sjolund, M.; Bergenstahl, B. Biochemistry 1991, 30, 10938–48. (45) Zuidam, N. J.; Barenholz, Y. Biochim. Biophys. Acta 1997, 1329, 211–22. (46) Gruner, S. M.; Tate, M. W.; Kirk, G. L.; So, P. T.; Turner, D. C.; Keane, D. T.; Tilcock, C. P.; Cullis, P. R. Biochemistry 1988, 27, 2853–66. (47) Rand, R. P.; Fuller, N.; Parsegian, V. A.; Rau, D. C. Biochemistry 1988, 27, 7711–22. (48) Chapman, D.; Williams, R. M.; Ladbrooke, B. D. Chem. Phys. Lipids 1967, 1, 445–475. (49) Harlos, K. Biochim. Biophys. Acta 1978, 511, 348–55. (50) Watts, A.; Harlos, K.; Marsh, D. Biochim. Biophys. Acta 1981, 645, 91–6. (51) Jahnig, F.; Harlos, K.; Vogel, H.; Eibl, H. Biochemistry 1979, 18, 1459–68. (52) Lewis, R. N.; Tristram-Nagle, S.; Nagle, J. F.; McElhaney, R. N. Biochim. Biophys. Acta 2001, 1510, 70–82.

Langmuir 2009, 25(18), 10719–10729

Mo and Yip

Article

Figure 6. (a) In situ tapping mode AFM image showing preferential growth of PIC aggregates on DSPC gel-phase domains of a DSPCDOPC-RhoDHPE mixed bilayer. The height profile is intentionally overscaled to emphasize the difference between J-aggregates and nonfluorescent aggregates (saturated peak). The growth of both (b) type II (height 1.5 nm) and (c) type I aggregates (height 2.5 nm) on the same DSPE bilayer can clearly be seen.

the gel-phase domains. This suggests that lateral stabilization by the substrate is critical for stable nuclei formation. Visible light spectroscopy and fluorescence microscopy of gel-state bilayers confirmed that the J-aggregates originated from surface-mediated templating rather than adsorption of solution-nucleated aggregates. The presence of templated type I J-aggregates, commonly associated with NaCl-induced aggregation in solution, further supports the role of the substrate in selective templating. Specifically, while POPE bilayers yielded exclusively type I J-aggregates, the DSPC bilayers afforded only type II J-aggregates. Such specificity suggests that templating on these substrates was significantly influenced by the lateral structure of the bilayers. To examine the role of lipid order and packing on aggregation, we then considered the phase behavior and area per headgroup (Ah) of our substrate lipids. While ionic strength has been reported to alter the phase behavior of certain charged lipids,53 this effect is relatively slight, requiring roughly 3 orders of magnitude change in ionic strength to effect a 5 °C change in transition temperature.54 Remarkably, gel-phase DSPC bilayers (Ah = 54.7 A˚2) facilitated type II aggregates,55,43 while gel-phase DPPC bilayers (Ah = 48.0 A˚2)48 could only support weakly

fluorescent non-J-type aggregates, which were usually found on fluid-phase bilayers such as DOPC (Ah = 82.0 A˚2). This sensitivity to headgroup spacing suggests that J-aggregate growth on bilayers is a consequence of specific, steric interactions between the PIC monomers and the lipid headgroups. Polarized ATR FT-IR spectroscopy revealed that monomeric PIC was associated with the lipid headgroups of the anionic (PA and PG) and zwitterionic (PC and PE) bilayers. The absence of spectral features that could be uniquely assigned to J-aggregates on the gel-phase bilayers suggests that these vibrational modes were either too weak or infrared-inactive due to the symmetry of PIC and/or the aggregate. In the case of PIC, it has been shown in a non-J-aggregate form that stabilization in the solid state occurs though π-stacking (Cambridge Structural Database code: DAQYEZ) and that in the J-aggregate motif the quinoline rings of adjacent PIC molecules interact through π-bonds.56 It is also known that stacked aromatic rings can have inactive ringbreathing modes.57 To characterize the orientation of the bilayer-associated PIC molecules, we examined the aliphatic C-N stretching mode at ∼1160 cm-1. In the case of PIC, this particular mode is oriented

(53) Lamy-Freund, M. T.; Riske, K. A. Chem. Phys. Lipids 2003, 122, 19. (54) Copeland, B. R.; Andersen, H. C. Biochemistry 1982, 21, 2811–2820. (55) Harvey, R. D.; Heenan, R. K.; Barlow, D. J.; Lawrence, M. J. Langmuir 2004, 20, 9282.

(56) Akins, D. L.; Macklin, J. W.; Zhu, H. R. J. Phys. Chem. 1991, 95, 793–798. (57) Cai, W. B.; Wan, L. J.; Noda, H.; Hibino, Y.; Ataka, K.; Osawa, M. Langmuir 1998, 14, 6992–6998.

Langmuir 2009, 25(18), 10719–10729

DOI: 10.1021/la901227h

10725

Article

Mo and Yip

Figure 7. (top) Fluorescence, (middle) correlated, and (bottom) AFM images showing preferential growth of PIC J-aggregates on (a-c) DSPC gel domains of a DSPC-DOPC-RhoDHPE bilayer. Scale bar: 5 μm. (d) DSPE bilayer; scale bar: 2 μm.

Figure 8. Polarized FT-IR difference spectra of different monocomponent bilayers. Signature spectra for monomeric PIC comprise four ring-breathing bands from 1400 to 1600 cm-1. Polarized spectra are vertically shifted for clarity. Bulk PIC powder transmission spectra are shown for comparison.

perpendicular to the molecule’s long axis and is not associated with the central quinoline rings. On both DOPC and DOPE 10726 DOI: 10.1021/la901227h

bilayers, this band was only observed in the p-polarized spectra, whereas on DOPG and DOPA, this mode was present in both Langmuir 2009, 25(18), 10719–10729

Mo and Yip

Article

Figure 9. Proposed cation-π stabilization scheme and molecular orientation based on polarized FT-IR data. (a) Interaction of the choline headgroup (blue) with quinoline rings of PIC. (b) Direction of aliphatic C-N bond (red) stretching with respect to the PIC molecule. Table 1. Summary of Lipid Bilayer Phase, Measured Area per Lipid in Excess Hydration, Phase Behavior at Room Temperature Prior to the Interaction with PIC, Charge at pH 7.4 (from pKai of Each Lipid at the Experimental Ionic Strength), and Corresponding J-Aggregate Types lipid

area/lipid (A˚2)

phase

charge at pH 7.4

aggregate type

43

LR neutral DOPC 82.0 DOPG NA LR LR DOPA 69.044 LR þ DOTAP 65.045 46 LR neutral DOPE 65.0 Lβ neutral type I POPE 56.647 Lβ’ neutral type II DSPC 54.743 Lβ’ neutral DPPC 48.048 49 Lβ neutral type I and II DSPE 39.0 Lβ DSPG 48.050 39.251 Lβ DSPAa Lβ þ DMTAP 40.052 a The DSPA headgroup area was estimated using data available for DPPA.

s- and p-polarized spectra. These data suggest that on bilayers that support J-aggregate formation the PIC molecules are oriented such that their hydrocarbon tails are aligned largely parallel to the bilayer normal, while on the nonaggregate forming bilayers the PIC molecules are largely disordered (Figure 9). We would note that the PIC quinoline rings are twisted relative to each other, which would complicate assigning an orientation to the PIC monomer based solely on polarized IR spectra of the representative ring modes.40 Given this orientation, the observed type I and type II aggregate thicknesses of 2.5 and 1.5 nm would correspond to ∼4 and 2 molecular layers, respectively. Attempts to resolve in-plane molecular-scale periodicities on these aggregate structures by high-resolution in situ AFM were unsuccessful. To assess the mechanism associated with PIC-bilayer binding, we examined the PIC-lipid headgroup interactions. Studies of lipid headgroup orientation by 2H NMR have shown that, in the presence of cations, the PC headgroup is oriented at ∼40° (phosphate-amine, P-N vector measured from the bilayer normal), relative to the bilayer plane.58,59 The zwitterionic (58) Seelig, J.; Macdonald, P. M.; Scherer, P. G. Biochemistry 1987, 26, 7535– 7541. (59) Bechinger, B.; Seelig, J. Chem. Phys. Lipids 1991, 58, 1–5.

Langmuir 2009, 25(18), 10719–10729

bilayers would therefore expose the choline or ethanolamine headgroup. At the experimental pH, the PA/PG/TAP headgroups present phosphate anions/hydroxyl groups/cations at the bilayer surface, respectively. On zwitterionic bilayers, the ionic distribution is unaffected by surface charge. These considerations suggest that PIC association with zwitterionic PE and PC bilayers is likely facilitated by cation-π interactions. Studies of PIC and its aggregates using density functional theory have reported that the net positive charge on PIC is delocalized about the central conjugated carbon chain, resulting in alternating arrangement of slightly positive and negative carbon atoms.40 This effect coupled with delocalization of the chromophore’s HOMO π-orbitals, as has been reported in a related study of indocyanine dyes,60 suggests that PIC could participate in cation-π interactions. We and others have described the role of cation-π interactions in driving the association of the indole sidegroup of tryptophan-rich peptides with model membranes both computationally61,62 and experimentally.31,63 Our data would suggest that PIC follows a similar initial adsorption and binding pathway. In support of our model of cation-π interactions, recent experimental and computational modeling studies that examined the interaction between tryptophan and phosphocholine were considered.64,65 In this work, H-bonding, carbonyl-cation, and cation-π interactions were all found to influence tryptophan-phosphocholine association; however, it was also argued that conformational considerations are in place with respect to the relative magnitude of the individual contributions. In the context of the present work, the close structural similarity between PIC and the tryptophan moiety modeled in this work provides compelling support for our suggestion of strong cation-π interactions, including preferred cation-π interactions with the 6-membered aromatic core.66 Furthermore, it has been reported that the free energy associated with tryptophan-phosphocholine interactions can be significant, ranging from ∼1.7 to 5 kcal/mol.64 This suggests that there is sufficient interaction between the adsorbing PIC and the lipid to induce local rearrangement of the lipid molecules themselves. Indeed, subtransitions within gel lipid phases have relatively low enthalpic requirements (∼0.17 to 0.82 kcal/mol)67 while the stacking of quinolinium carbocations is energetically favorable (3.6 to 11.8 kcal/mol).68 Electrostatic interactions alone would favor PIC binding to the negatively charged PA and PG bilayers, compared to positively charged TAP bilayers, which is consistent with our observations. These data suggest the following model for J-aggregate formation on supported bilayers (Figures 10 and 11). Initial absorption of PIC to the bilayer surface is driven through electrostatic and cation-π stabilization with the lipid headgroups. Inter-PIC interactions then stabilize the formation of a type I or type II aggregate, depending on the local bilayer headgroup packing and phase. Local reorganization of the lipids resulting in a transition to a subgel or LC phase could then occur due to the low energetic requirements of these transitions and the high binding energy between the lipid headgroups and the PIC monomers. We noted (60) Fu, Y. L.; Huang, W.; Li, C. L.; Wang, L. Y.; Wei, Y. S.; Huang, Y.; Zhang, X. H.; Wen, Z. Y.; Zhang, Z. X. Dyes Pigm. 2009, 82, 409–415. (61) Khandelia, H.; Kaznessis, Y. N. J. Phys. Chem. B 2007, 111, 242–50. (62) Hsu, J. C.; Yip, C. M. Biophys. J. 2007, 92, L100–2. (63) Shaw, J. E.; Epand, R. F.; Hsu, J. C.; Mo, G. C.; Epand, R. M.; Yip, C. M. J. Struct. Biol. 2008, 162, 121–38. (64) Sanderson, J. M. Org. Biomol. Chem. 2007, 5, 3276–86. (65) Sanderson, J. M.; Whelan, E. J. Phys. Chem. Chem. Phys. 2004, 6, 1012– 1017. (66) Ma, J. C.; Dougherty, D. A. Chem. Rev. 1997, 97, 1303–1324. (67) Tenchov, B.; Koynova, R.; Rapp, G. Biophys. J. 2001, 80, 1873–1890. (68) Buisine, E.; de Villiers, K.; Egan, T. J.; Biot, C. J. Am. Chem. Soc. 2006, 128, 12122–12128.

DOI: 10.1021/la901227h

10727

Article

Figure 10. Schematics of the proposed J-aggregate templating mechanism. The mechanism differs for each phase due to headgroup and tail order. In the case of the Lc phase, the nontilted lipid tails result in two possible packing motifs. Open circles: lipid tails; green rectangle: headgroup; orange oval: first layer PIC molecule; filled circles: packing of lipid tails.

Mo and Yip

PIC monomers to the bilayer. On the basis of Lβ phase headgroup area, which is similar to that in Lc phase, the optimal headgroup lateral spacing is 54.7 A˚2 for tilted bilayers and 39 A˚2 for nontilted bilayers. If we consider the tilt of the lipid headgroups, the area perpendicular to the lipid tail is 54.7 cos(40°) = 42 A˚2, corresponding to an intrachromophore headgroup-headgroup distance of 6.24-6.47 A˚, in good agreement with the distance between the two quinoline rings of PIC (Figure 9). These data would suggest that cyanine dyes with longer conjugated chains separating the quinoline rings would not fulfill these steric requirements. This was confirmed when we found no evidence of fluorescent J-aggregates while examining the self-assembly of the cationic carbocyanine dye, pinacyanol chloride (1,10 -diethyl2,20 -carbocyanine chloride; IUPAC:1-ethyl-2-[3-(1-ethyl-2(1H)quinolinylidene)propenyl]quinolinium chloride), which is known to exhibit bathochromic shifts of the monomer band when associated with micelles,69 and gel-phase bilayers (data not shown). We also noted that heating the bilayers resulted in a loss of fluorescence and disintegration of the J-aggregates, with the onset of disruption occurring at or slightly below the lipid pretransition temperatures. These data support our model that stabilization of the J-aggregates requires a structural match with the substrate and that the inter-PIC interactions are relatively weak. The interchromophore distance is also affected by lipid chain tilt. In the Lc phase, the chain tilt increases the separation between headgroups such that the arrangements of dye molecules in several directions are degenerate. A lack of degeneracy in possible dye packing motifs on nontilted bilayers results in two distinct dye packing motifs. Indeed, both type I and type II J-aggregates formed on the nontilted DSPE bilayers. To our knowledge, this is the first time that these distinct J-aggregates were imaged on the same substrate. A herringbone arrangement of the dye was unlikely since the characteristic hypsochromic shift was not observed in these aggregates nor did the J-peak undergo Davydov splitting.70

Conclusion

Figure 11. Schematic of proposed J-aggregation mechanism with respect to bilayer phase and headgroup chemistry. (top) Templated growth of J-aggregates occurs through a combination of cation-π interactions and specific lipid headgroup packing and phase motifs. PIC does not adsorb on cationic lipids due to electrostatic repulsion and cannot accumulate at the bilayer interface. (middle) Zwitterionic PC and PE bilayers satisfy all requirements and yield templated J-aggregates. (bottom) Anionic lipids such as PG and PA lack specific binding interaction and cannot support epitaxial packing.

that PIC binding to the bilayers resulted in a slight decrease in the monomer:dimer solution absorbance spectra ratio (data not shown). This effect is consistent with preferential binding of 10728 DOI: 10.1021/la901227h

In situ correlated confocal fluorescence-AFM imaging has revealed that the nucleation and growth of PIC-derived J-aggregates on supported lipid bilayers is a consequence of both structural and electrostatic interactions. While cation-π interactions favor stabilization with the lipid headgroups, lateral headgroup order and packing geometry are responsible for the organization of the dye aggregates. Moreover, we found that the form of the J-aggregates was dependent on the bilayer phase and lipid tail tilt, suggesting that judicious selection of bilayer chemistry and local structure may provide a route for the isolation of previously unknown aggregate isoforms. Similar approaches have been used by others to examine the ability of substrates, including single crystals, self-assembled monolayers with tailored chemistries, and confined structures to nucleate polymorphs of various small molecule dye and pharmaceutical compounds.71-74 These studies illustrate the need to create an environment that favors formation of a critically sized and/or oriented nuclei that then facilitates stable polymorph growth. (69) Sabate, R.; Estelrich, J. J. Phys. Chem. B 2003, 107, 4137–4142. (70) Saito, K.; Ikegami, K.; Kuroda, S.-i.; Tabe, Y.; Sugi, M. Jpn. J. Appl. Phys., Part 1 1991, 30, 1836. (71) Soegiarto, A. C.; Ward, M. D. Cryst. Growth Des. 2009, Article ASAP. (72) Singh, A.; Lee, I. S.; Myerson, A. S. Cryst. Growth Des. 2009, 9, 1182–1185. (73) Mitchell, C. A.; Yu, L.; Ward, M. D. J. Am. Chem. Soc. 2001, 123, 10830–9. (74) Plaut, D. J.; Martin, S. M.; Kjaer, K.; Weygand, M. J.; Lahav, M.; Leiserowitz, L.; Weissbuch, I.; Ward, M. D. J. Am. Chem. Soc. 2003, 125, 15922–34.

Langmuir 2009, 25(18), 10719–10729

Mo and Yip

In the present work, we have shown how a subtle interplay exists in which the formation of that molecular nuclei can be driven by both an initial substrate-facilitated stabilization and that the subsequent intermolecular interactions within the growing aggregate then facilitate a local reordering within the substrate itself. An analogous energetic model had been previously extended by Last et al., in which the epitaxial formation of molecular overlayers was facilitated by a balance between the intermolecular interactions within the layer(s) themselves and the stabilizing influence of the crystalline substrate.30 Our studies also suggest that the formation of specific J-aggregate states could be used to report on the local order and/or structure within the supporting substrate. Our present work illustrates the need to carefully consider not only intermolecular interactions associated with dye aggregation, as is often studied in solution, but also, in the creation of solid-state structures, the role that the nucleating substrate (or environment) may have in directing the formation, alignment, and state of the resulting architecture. This latter issue is particularly relevant for the synthesis of lowdimensional J-aggregate structures and understanding the mechanism associated with exciton propagation in optoelectronic applications.75,76 Controlling or understanding the self-assembly of these dye aggregates has importance in the cellular context. For example, J-aggregate-forming dyes, such as JC-1, are often used to track membrane depolarization in mitochondria and report on the early stages of apoptosis.77,78 However, these dye aggregates can play multiple roles. Indeed, J- and related dye aggregates have (75) Chan, J. M.; Tischler, J. R.; Kooi, S. E.; Bulovic, V.; Swager, T. M. J. Am. Chem. Soc. 2009. (76) Lagoudakis, P. G.; de Souza, M. M.; Schindler, F.; Lupton, J. M.; Feldmann, J.; Wenus, J.; Lidzey, D. G. Phys. Rev. Lett. 2004, 93, 257401. (77) Smiley, S. T.; Reers, M.; Mottola-Hartshorn, C.; Lin, M.; Chen, A.; Smith, T. W.; Steele, G. D. Jr.; Chen, L. B. Proc. Natl. Acad. Sci. U.S.A. 1991, 88, 3671–5. (78) Reers, M.; Smiley, S. T.; Mottola-Hartshorn, C.; Chen, A.; Lin, M.; Chen, L. B. Methods Enzymol. 1995, 260, 406–17.

Langmuir 2009, 25(18), 10719–10729

Article

been shown to exert a protective role in cells by acting as singlet oxygen quenching agents.79-81 Perhaps more intriguingly lies the potential that the aggregated state may reflect the transition between a compound that can act as either a photoprotective or a phototoxic agent, as has been explored recently in studies of cyanine dyes for photodynamic therapeutic agents for the treatment of cancer.82-85 This suggests that judicious design of the dye molecule with due consideration for the environment that it will encounter and the role that that environment may play in the molecule’s self-association characteristics may provide opportunities for tunable or switchable therapeutic agents. Acknowledgment. G.C.H.M. thanks NSERC for scholarship support. C.M.Y. acknowledges support from the Canada Research Chairs program. The equipment used in this study were provided by a grant from the Canada Foundation for Innovation (Grant # 3131) and supported by a Research Resource grant from CIHR (Grant # PRG-80174). The authors thank Chantal Blanchard and George Sakellaropoulos of Olympus Canada, and Andrew Elia, Princess Margaret Hospital, University Health Network, Toronto, for access to their Olympus FV1000 confocal microscope. Supporting Information Available: Normalized fluorescence spectra and corresponding confocal images of the J-aggregates (Figure 1). This material is available free of charge via the Internet at http://pubs.acs.org. (79) Kanofsky, J. R.; Sima, P. D. Photochem. Photobiol. 2000, 71, 361–8. (80) Kanofsky, J. R.; Sima, P. D. Photochem. Photobiol. 2009, 85, 391–9. (81) Sima, P. D.; Kanofsky, J. R. Photochem. Photobiol. 2000, 71, 413–21. (82) Chen, Y.; Gryshuk, A.; Achilefu, S.; Ohulchansky, T.; Potter, W.; Zhong, T.; Morgan, J.; Chance, B.; Prasad, P. N.; Henderson, B. W.; Oseroff, A.; Pandey, R. K. Bioconjugate Chem. 2005, 16, 1264–74. (83) Kassab, K. J. Photochem. Photobiol. B 2002, 68, 15–22. (84) Ara, G.; Aprille, J. R.; Malis, C. D.; Kane, S. B.; Cincotta, L.; Foley, J.; Bonventre, J. V.; Oseroff, A. R. Cancer Res. 1987, 47, 6580–5. (85) Oseroff, A. R.; Ohuoha, D.; Ara, G.; McAuliffe, D.; Foley, J.; Cincotta, L. Proc. Natl. Acad. Sci. U.S.A. 1986, 83, 9729–33.

DOI: 10.1021/la901227h

10729