Protection of a Decapeptide from Proteolytic Cleavage by Lipidation

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Langmuir 1999, 15, 5500-5508

Protection of a Decapeptide from Proteolytic Cleavage by Lipidation and Self-Assembly into High-Axial-Ratio Microstructures: A Kinetic and Structural Study Kyujin C. Lee,†,‡ Paul A. Carlson,§ Alex S. Goldstein,† Paul Yager,*,§ and Michael H. Gelb*,† Departments of Chemistry and Biochemistry, University of Washington, Box 351700, Seattle, Washington 98195, Department of Pathobiology, University of Washington, Box 357238, Seattle, Washington 98195, and Department of Bioengineering, University of Washington, Box 352255, Seattle, Washington 98195 Received January 26, 1999. In Final Form: April 30, 1999 We report on the synthesis, kinetics of proteolysis by trypsin, and morphological characterization of a novel lipidated decapeptide that spontaneously self-assembles in aqueous solutions into 0.5 µm diameter hollow tubules and helices that range in length from tens to hundreds of micrometers depending on formation conditions. We also report on an improved method for the tritioacetylation of peptides. Tight molecular packing of the peptide-amphiphile into a crystalline bilayer array forces tight packing between peptide headgroups, which was found to significantly protect the peptides from proteolysis by trypsin. Relief of this steric hindrance between peptide headgroups caused by solubilization of the bilayer into detergent micelles accelerated the rate of trypsin hydrolysis by 32,000-fold. Raman spectroscopy and circular dichroism spectropolarimetry were used to gain molecular-level insight into the difference between hydrolysis rates. Results obtained from these studies suggest that differences in molecular packing and conformation of the peptide headgroups in crystalline tubular and dispersed micellar phases determine the extent of proteolytic protection. Protection from proteolysis is considered a useful feature of lipidated peptide tubules for their potential use as a depot of bioactive peptides and other labile prodrugs at defined biological sites for sustained release.

Introduction Delivery of labile peptide drugs and vaccines to desired biological sites is problematic because of hydrolytic degradation, poor oral absorption, fast clearance, and poor permeation into cells. Significant efforts have been made to protect and deliver peptides to target sites in a controlled manner. Chemical modification of peptides to make them more resistant to degradation such as N-methylation of the peptide bond,1,2 C-terminal esterification,3 the use of infusion pumps for peptide delivery,4 and encapsulation of peptides in various carriers such as liposomes5,6 and bioerodable microspheres7,8 are some recent examples. A potentially useful approach to solve these delivery problems incorporates the chemical derivatization of the peptide with a lipid moiety9-11 to produce a lipidated * To whom correspondence should be addressed. † Departments of Chemistry and Biochemistry. ‡ Department of Pathobiology. § Department of Bioengineering. (1) Simon, R. J.; Kania, R. S.; Zuckerman, R. N.; Huebner, V. D.; Jewell, D. A.; Banville, S.; Ng, S.; Wang, L.; Rosenberg, S.; Marlowe, C. K. Proc. Natl. Acad. Sci. U.S.A. 1992, 89, 9367-9371. (2) Bundgaard, H.; Rasmussen, G. J. Pharm. Res. 1991, 8, 313-322. (3) Mass, J. Peptide prodrugs designed to limit metabolism; Taylor, M. D., Amidon, G. L., Eds.; ACS Professional Reference Book: Washington, DC, 1995; pp 423-448. (4) Brange, J.; Havelund, S. Acta Med. Scand. Suppl. 1993, 671, 135-138. (5) Arya, S. C. Vaccine 1994, 12, 1423-1435. (6) Allen, T. M. Drugs 1997, 54, 8-14. (7) Ogawa, Y.; Yamamoto, M.; Okada, H.; Yashiki, T.; Shimamoto, T. Chem. Pharm. Bull. (Jpn.) 1988, 36, 1095-1103. (8) Couvreur, P.; Vauthier, C. J. Controlled Release 1991, 17, 187. (9) Eichholtz, T.; deBont, D. B. A.; deWidt, J.; Liskamp, R. M. J.; Ploegh, H. L. J. Biol. Chem. 1993, 268, 1982-1986. (10) BenMohamed, L.; Gras-Masse, H.; Tartar, A.; Daubersies, P.; Brahimi, K.; Bossus, M.; Thomas, A.; Druilhe, P. Eur. J. Immunol. 1997, 27, 1242-1253.

prodrug.3 This approach can be very useful to protect the peptide against first-pass metabolism, to facilitate absorption, and to slow clearance of hydrophilic peptides only if the lipidated derivatives can be subsequently converted into the parent peptides through an enzymatic reaction at the site of delivery. Recently, several classes of modified peptide surfactants or lipopeptides that contain two alkyl chains have been found to spontaneously self-assemble into tightly packed crystalline bilayers that form hollow tubules and helical ribbons.12-18 Thesehigh-axial-ratiomicrostructures(HARMs) are of micrometer size, and their potential use for zeroorder sustained drug release has been proposed.17,18 As self-assembling noncovalent aggregates, bilayer microstructures offer a unique model for molecular engineering in which the types of constituent molecules can be varied, chemically modified, and easily assembled. We envisioned the construction of lipopeptides that would self-assemble into tubule HARMs and would function as a general drugdelivery vehicle for peptide therapeutics; lipopeptides tightly packed into HARMs would be protease resistant. Although crystalline packing within a liposomal micro(11) Achiwa, K. Bio. Pharm. Bull. 1996, 19, 1271-1274. (12) Yager, P.; Schoen, P. Mol. Cryst. Liq. Cryst. 1984, 106, 371381. (13) Yamada, K.; Ihara, H.; Ide, T.; Fukumoto, T.; Hirayama, C. Chem. Lett. 1984, 10, 1713-1716. (14) Nakashima, N.; Asakuma, S.; Kim, J. M.; Kunitake, T. Chem. Lett. 1984, 1709-1712. (15) Nakashima, N.; Asakuma, S.; Kunitake, T. J. Am. Chem. Soc. 1985, 107, 510-512. (16) Shimizu, T.; Hato, M. Biochim. Biophys. Acta 1993, 1147, 5058. (17) Carlson, P. A.; Gelb, M. H.; Yager, P. Biophys. J. 1997, 73, 230238. (18) Lee, K. C.; Lukyanov, A. N.; Gelb, M. H.; Yager, P. Biochim. Biophys. Acta 1998, 1371, 168-184.

10.1021/la9900775 CCC: $18.00 © 1999 American Chemical Society Published on Web 06/12/1999

Protection of a Decapeptide

Langmuir, Vol. 15, No. 17, 1999 5501

Figure 1. Chemical structures of the HARM-forming lipidated peptides described in text. Critical aggregation concentrations were determined either by a measure of the activity remaining within the supernatant of centrifuged tritiated HARM suspensions (i.e., 3) or by an estimate based on reported values for analogous lipids having different alkyl chain lengths (i.e., 1 and 2).16,18

structure would also hinder enzymatic hydrolysis, liposomes can have a density of geometric packing defects sufficiently high enough to destroy crystalline periodicity,19 and these defects may provide regions within the microstructure more prone to enzymatic attack.17 Tubules comprised of diacetylenic phospholipids have fewer defects than do multilamellar vesicles (MLVs), and lipid molecules appear to pack more tightly in tubules compared to crystalline MLVs.20 To test this hypothesis, we designed a novel lipopeptide that features a glutamic acid linker modified with bis(tetradecylamide) tails and a decapeptide headgroup containing a trypsin-labile cleavage site. Enhanced protection is considered a useful feature of HARMs for their potential use as a depot of lipidated peptides and other labile drugs at defined biological sites for sustained release. In this article, we report on the synthesis and characterization of this novel lipidated decapeptide that spontaneously self-assembles into hollow tubular and helical ribbon microstructures in aqueous solution. Also reported is an improved method for the tritioacetylation of peptides. Assembly of the lipopeptide into HARMs was found to significantly protect the peptides from hydrolysis by trypsin as compared to the rate of hydrolysis for the same lipopeptide dispersed in detergent micelles. To gain insight into these vastly different rates of hydrolysis, Raman spectroscopy and circular dichroism spectropolarimetry were used to characterize differences in molecular packing and conformation of the peptide headgroups found in crystalline tubular and dispersed-micellar phases.

Materials. DMF (Aldrich, reagent grade) was dried over 4 Å molecular sieves. Unless indicated otherwise, all other chemicals and solvents were obtained from commercial sources (Aldrich, Lancaster, Fischer, Mallinckrodt, J. T. Baker, EM Science), were of reagent grade, and were used without further purification.

Synthesis. Analytical TLC was performed on precoated Silica Gel 60 F-254 plates (EM Science), and visualization was accomplished with UV light, iodine vapor, Cl2/o-tolidine staining, or phosphomolybdic acid stain. Flash chromatography was performed with Silica Gel 60 (230-400 mesh) from EM Science. Electrospray mass spectrometry (ES-MS) was performed using a Kratos HV-3 Profile analytical electrospray magnetic sector mass spectrometer. Preparation of L-Glutamic Acid Bis(tetradecylamide), 1, and Tri-L-proline-L-glutamic Acid Bis(tetradecylamide), 2. The tubuleforming core compounds, E-(NHC14H29)2 (compound 1 in Figure 1) and PPPE-(HNC14H29)2 (compound 2 in Figure 1), were synthesized and characterized according to the methods described in detail earlier.16,18,21 Preparation of Fmoc-GR(Pmc)AGGAAPPP. The peptide-bound resin (520 mg), Fmoc-GR(Pmc)AGGAAPPP-2-chlorotrityl resin (synthesized by SynPep, Inc., Dublin, CA), was dried in vacuo in the presence of P2O5 for 2 h and then treated with 1:1:8 AcOH/ TFE/dichloromethane (16 mL) for 1 h while stirring at room temperature.22 The cleaved peptide was separated from the resin by filtration. The resin was subjected to the same cleavage conditions a second time, and the filtrates were combined. Upon evaporation of the solvent, the residue was taken up in 1:5 MeOH/ CHCl3 (∼5 mL), and this solution was added to ethyl ether (∼100 mL) to precipitate the cleaved peptide. The resulting precipitate was recovered by filtration, washed with ethyl ether, and dried in vacuo for 5 h to give 254 mg of Fmoc-GR(Pmc)AGGAAPPP. TLC (4:1:1 BAW): Rf 0.30. Preparation of Ac-GRAGGAAPPPE-(HNC14H29)2, 3. A solution of Fmoc-GR(Pmc)AGGAAPPP (74.7 µmol, 100 mg) and hydroxybenzotriazole (78.5 µmol, 10.6 mg) in DMF (2 mL) was cooled to 0 °C. 1-(3-(dimethylamino)propyl)-3-ethylcarbodiimide hydrochloride (82.2 µmol, 15.8 mg) was added, and the mixture was stirred for 1 min followed by the addition of 1 (78.5 µmol, 42.2 mg) dissolved in CHCl3 (2.5 mL). The reaction was stirred at 0 °C for 3.5 h and then at room temperature for 1.5 days. Upon evaporation of the solvent, the residue was purified by silica gel flash chromatography with CHCl3/MeOH (9:1) to give FmocGR(Pmc)AGGAAPPPE(NHC14H29)2 in 65% yield. TLC (8.5:1.5 CHCl3/MeOH): Rf 0.36.

(19) Yeagle, P. The structure of biological membranes; CRC Press: Boca Raton, FL, 1992; p 1227. (20) Caffrey, M.; Hogan, J.; Rudolph, A. S. Biochemistry 1991, 30, 2134-2146.

(21) Ihara, H.; Yoshikai, K.; Takafuji, M.; Hirayama, C.; Yamada, K. Kobunshi Ronbunshu 1991, 48, 327-34. (22) Barlos, K.; Chatzi, O.; Gatos, D.; Stavropoulos, G. Int. J. Pept. Protein Res. 1991, 37, 513-520.

Methods

5502 Langmuir, Vol. 15, No. 17, 1999 The N-terminal peptide blocking group was removed by mixing Fmoc-GR(Pmc)AGGAAPPPE(NHC14H29)2 (46.9 µmol, 87.2 mg) with piperidine (15 mL) and stirring the solution at room temperature for 1.5 h under argon. Upon evaporation of the solvent, the residue was purified by Sephadex LH-20 column chromatography with MeOH (100%) to give GR(Pmc)AGGAAPPPE(NHC14H29)2 in 97% yield. TLC (8:2 CHCl3/2 M NH3 in MeOH): Rf 0.23. ES-MS ([M + H]+): 1635.2; calcd, 1634.2. Acetylation of the peptide’s N-terminus was achieved as follows. Briefly, to GR(Pmc)AGGAAPPPE(NHC14H29)2 (17.7 µmol, 29.0 mg) dissolved in 1:1 DMF/CHCl3 (2.5 mL) was added acetic anhydride (19.50 µmol, 1.84 µL) followed by the addition of triethylamine (19.5 µmol, 2.72 µL) and 4-(dimethylamino)pyridine (1.77 µmol, 0.22 mg) dissolved in 1:1 DMF/CHCl3 (400 µL). An additional 0.5 mL of DMF was added to the mixture, and it was stirred overnight at room temperature. H2O (0.5 mL) was added, and the reaction mixture was stirred for 30 min before diluting it with CHCl3 (70 mL). The organic phase was washed with 5% NaHCO3 (2×) and H2O. Upon evaporation of the solvent, the residue was purified by Sephadex LH-20 chromatography to give Ac-GR(Pmc)AGGAAPPPE(NHC14H29)2 in 91% yield. TLC (2:8 MeOH:CHCl3): Rf 0.20. Final deprotection was achieved by treating Ac-GR(Pmc)AGGAAPPPE(NHC14H29)2 with 95:5 TFA/H2O (3 mL). The mixture was stirred at room temperature for 2.5 h. Upon evaporation of the solvent, ethyl ether (5 mL) was added to the product precipitate. The solid was washed with ethyl ether (2×) and reprecipitated from 20% MeOH/H2O (4 mL) with ether. The resulting solid was further purified by HPLC on a Vydac 214TP1010 column with an 80% MeOH/H2O to 100% MeOH gradient containing 0.06% TFA to give a single peak of AcGRAGGAAPPPE(HNC14H29)2 (compound 3 in Figure 1). Yield: 45%. TLC (4:1:1 BAW): Rf 0.27. ES-MS ([M + H]+): 1412.9; calcd, 1412.8. Preparation of [3H]Ac-GRAGGAAPPPE(HNC14H29)2, [3H]-3. Tritioacetylation of the peptide headgroup was achieved using an improved radiolabeling method that avoids using acetic anhydride, which is volatile, difficult to handle safely, and easily hydrolyzed. [3H]Acetic acid, sodium salt (100 mCi, specific activity ) 9.3 Ci/mmol, Amersham), was suspended in ethanol (2 mL) and sonicated in a bath sonicator (Laboratory Supplies Co., Inc, Hicksville, NY) to facilitate dispersion. Half of the tritiated acetic acid solution (∼1 mL) was dried under vacuum in the presence of P2O5 and Dririte followed by further drying from benzene (3 × 0.5 mL) and CHCl3 (1 × 0.5 mL). The dried lipopeptide, GR(Pmc)AGGAAPPPE(NHC14H29)2 (4.8 µmol, 7.9 mg), was dissolved in dry DMF (400 µL) before it was added to the [3H]acetic acid, sodium salt, solution. We added BOP (5.9 µmol, 2.6 mg) dissolved in dry DMF (150 µL) to the reaction mixture, which was rotated under argon at room-temperature overnight. Water (2.1 mL) was added to the reaction mixture, and the precipitate was separated by removing the supernatant. The solid was washed with H2O (2.5 mL), 5% HCl (2.5 mL), H2O, and 5% NaHCO3, by sequential centrifugation and resuspension steps. After a final wash with ethyl ether, a white solid was isolated from the aqueous phase with ∼40% recovery containing ∼5% unacetylated peptide. TLC (1.5:8.5 MeOH/CHCl3): Rf 0.17. To increase the coupling yield, the [3H]acetic acid coupling reaction was repeated with a final 90% recovery and no detectable unacetylated peptide. TLC (25% MeOH in CHCl3): Rf 0.42. The product comigrated with the authentic unlabeled compound. The final deprotection to give [3H]Ac-GRAGGAAPPPE(HNC14H29)2 was achieved as described earlier. The product was recovered by HPLC on a Vydac 214TP1010 column with an 80% MeOH/H2O to 100% MeOH gradient, dissolved in 5 mL DMF/benzene (95:5), and stored at -80 °C. TLC and fluorography revealed a single comigrating spot with authentic unlabeled compound. TLC (6:2:2 BAW): Rf 0.40. Preparation of Ac-GRAGGAAPPP. Ac-GR(Pmc)AGGAAPPP2-chlorotrityl resin (80 mg) was stirred with 1:1:8 AcOH/TFE/ CH2Cl2 (2 mL) at room temperature for 30 min and then filtered. The resin was retreated as before but with 2:2:6 AcOH/TFE/ CH2Cl2 (2 mL), and the filtrates were combined. Upon evaporation of the solvent, H2O was added, and the peptide solution was dried with a speed-vac to obtain the Pmc-protected crude peptide, Ac-GR(Pmc)AGGAAPPP (46.9 mg). TLC (4:2:2 BAW): Rf 0.49.

Lee et al. ES-MS ([M + H]+): 1158.8; calcd, 1157.8. Subsequent Pmc removal from Ac-GR(Pmc)AGGAAPPP (4 mg) was achieved with 0.5 mL TFA/Phenol/H2O/triisopropylsilane (88:5:5:2) by stirring at room temperature for 1.5 h.23 Some 20% aqueous AcOH (1.5 mL) was added, and the aqueous phase was washed with cold ethyl ether (5 × 1 mL). The aqueous layer was lyophilized to give Ac-GRAGGAAPPP. The peptide was purified on a Vydac 218TP1010 HPLC column with 100% H2O (0.07% TFA) to 50% ACN/H2O (0.06% TFA) in 50 min (66% yield). TLC (4:2:2 BAW): Rf 0.13. ES-MS ([M + H]+): 892.98; calcd, 891.98. Preparation of Ac-Gly-Arg. To Ac-Gly in 1.7 mL of THF/DMF (1:1.7) were added BOP (0.5 µmol, 221 mg) and N-methyl morpholine (0.5 µmol, 54.8 µL), and the solution was stirred at room temperature for 10 min. The mixture was then added dropwise to a solution of arginine dissolved in H2O (1 mL) and allowed to stir at room temperature overnight. After removal of the solvent, the residue was taken up in H2O (3 mL) and stored at -20 °C overnight. The solution was filtered, and the H2O layer was evaporated to an oil, which was washed with ethyl ether. The product was further purified on a Vydac 214TP1010 HPLC column with 100% H2O to 35% ACN/H2O gradient containing 0.06% TFA with a single peak eluting at 11.5 min. TLC (6:4:1 CHCl3/MeOH/AcOH): Rf 0.16. ES-MS ([M + H]+): 274.1. Trypsin Hydrolysis Studies. Tubule Preparation for Trypsin Hydrolysis. Prior to tubule formation, [3H]-3 (0.643 mg, specific activity ) 1.86 Ci/mmol) in 95:5 DMF/benzene (283 µL) was dried and then redried from MeOH (400 µL). The residue was then dissolved in MeOH (1 mL), and the solution was added to HEPESbuffered saline (HBS) (2.33 mL) dropwise while vortexing. The suspension was left undisturbed at room temperature for 24 h to anneal the suspension of tubule microstructures. The suspension was centrifuged at 10 000g for 30 min, and the supernatant was removed. The tubules were then washed with MeOH-free HBS (600 µL × 3) by a series of centrifugation and resuspension steps, and on the final exchange of solvent, the tubules were suspended in HBS to the desired lipid concentration. Evaluation of Ac-GRAGGAAPPP and 3 as Substrates for Trypsin. To Ac-GRAGGAAPPP (0.7 mg) dissolved in 400 µL of HBS was added 44 µg of trypsin (Promega, modified, sequencing grade). The solution was mixed and incubated at 37 °C in a water bath for 7 h prior to analysis. Hydrolysis fragments were separated from the reaction mixture with a Vydac 218TP1010 HPLC column using a 100% H2O (0.06% TFA) to 50% ACN/H2O (0.07% TFA) gradient over 50 min. Two fractions eluting at 16.5 and 22.5 min were collected separately and analyzed by ES-MS. The [M + H]+ of fraction 1 was 275.1, which corresponds to the molecular weight of 274.3 calculated for Ac-GR, and the [M + H]+ of fraction 2 was 637.7, which corresponds to the molecular weight of 636.7 calculated for AGGAAPPP. Similarly, 0.55 mM 3 in 250 mM borate buffer (pH 7.4) was reacted with 1.5 µM trypsin at 37 °C for 4 h. Hydrolysis fragments from the reaction mixture were separated on a TLC plate. The presence of TLC hydrolysis product spots confirms that 3 is a substrate of trypsin. TLC (4:1:1 BAW): Rf 0.25 and 0.43. Action of Trypsin Activity on Tubule Suspensions and on Triton X-100 Solubilized [3H]-3 Micelles. Micellar solutions of [3H]-3 were created by solubilizing tubule suspensions in either 2 or 10 mM Triton X-100 containing HBS. After overnight incubation of tubular or micellar [3H]-3 in 80 µL of HBS, 2 µL of trypsin (from porcine pancreas, Promega sequencing-grade, modified) stock solution (either 3.5 or 875 nM depending on the exact experiment) was added, and the reaction mixtures were rocked at room temperature. At each time point throughout hydrolysis, a small aliquot (5 µL) was removed, and any residual trypsin activity was quenched with the addition of TFA to reduce the pH to 2. Unlabeled Ac-GR (10 µg) was added to the reaction aliquots as a carrier, and samples were stored at -20 °C until processed by HPLC. [3H]Ac-GR was separated on a Vydac214TP1010 column, and the collected fractions were submitted to liquid scintillation counting. To assess the loss of activity of trypsin with time, aliquots from the reaction mixtures were assayed periodically using benzoylarginine ethyl ester. Each aliquot was immediately (23) Sole, N. A.; Barany, G. J. Org. Chem. 1992, 57, 5399-5403.

Protection of a Decapeptide assayed for trypsin activity with benzoylarginine ethyl ester (100 µM) by monitoring the absorbance change at 253 nm at 21 ( 2 °C with a circulating water bath. Full trypsin activity was retained over 37 h. Transmission Electron Microscopy. Samples for transmission electron microscopy were prepared by applying 0.5-2 µL of tubule suspensions to Formvar-coated 200 mesh copper grids (Electron Microscopy Sciences, Fort Washington, PA). Lipid tubules within the grid sample were allowed to settle for 30 s before the excess sample buffer was removed with filter paper. The images were obtained using a JEM 1200EXII transmission electron microscope operating at 80 kV (University of Washington, Department of Pathology). Raman Spectroscopy. Raman spectra were collected with a SPEX 500M spectrometer (Edison, NJ) equipped with a SPEX Spectrum-1, 578 × 385 pixel, liquid-N2-cooled CCD detector, a premonochromator, and a holographic notch filter to reject Rayleigh scattering. A Lexel argon-ion laser (Fremont, CA) tuned to 514.5 nm was used for sample excitation. Frequencies were calibrated to the 992 ∆ cm-1 band of benzene. The samples studied, AcGRAGGAAPPP and 1-3, were rehydrated from dry powders with deionized, distilled water. Spectra were collected from concentrated (∼100-300 mg/mL), hydrated lipid tubule suspensions placed within quartz capillary tubes. Sample temperature was controlled by a brass sample block connected to a programmable water bath, and 64 repetitive scans were averaged for each sample. A background spectrum obtained for deionized, distilled water was subtracted from each spectrum prior to analysis. Circular Dichroism. All circular dichroism (CD) spectra were recorded on a Jasco J-720 spectropolarimeter with a thermostated sample cell holder using a 0.5 mm path length quartz cell. For each sample, 16-32 repetitive scans were recorded and averaged with a 1 s response time, a 0.2 nm step size, and a 1.0 nm bandwidth. A sample was prepared of the free peptide AcGRAGGAAPPP in aqueous solution at a concentration of 67.3 µM. The zwitterionic detergent (hexadecyldimethylammonio)propane-1sulfonate (zwittergent 316) was found to have noninterfering optical properties in the UV, and 140 µM lipid-modified peptide 3 was readily solubilized in a 10 mM detergent solution to create a mixed-micellar phase. A background CD spectrum corresponding to the solution in the absence of peptide or lipid-modified peptide was subtracted from each CD spectrum. The measurements were carried out at 25 °C; the CD data are expressed as a mean residue ellipticity. Calorimetry. Differential scanning calorimetry (DSC) studies were performed using a Seiko Instruments, Inc. (Torrance, CA) DSC100 differential scanning calorimeter connected to a Seiko Instruments Inc. (Torrance, CA) SSC-5020 thermal analysis system. Samples were prepared by transferring 70 µL aliquots of the tubule suspensions into silver DSC pans. Each pan was immediately sealed and transferred to the calorimeter, which warmed to 25 °C. A 70 µL aliquot of HBS buffer was used as the reference. Samples were scanned at a rate of 0.5 °C/min for both heating and cooling cycles. The heating transition temperatures were determined from the onset of the transition endotherms.

Results/Discussion Peptide Design and HARM Formation. Our design of a model lipidated prodrug centered around two key criteria. First, the lipid-peptide conjugate should spontaneously self-assemble into tubular microstructures by itself or when codispersed with a known tubule-forming compound. Second, the enzymatic cleavage site should be located at a position far enough away from the lipid anchor that the hydrophobic tails would not interfere with hydrolysis by trypsin to distinguish protection from proteolysis by HARM formation from a simple steric effect of the attached lipid anchoring group. The tubule-forming compounds characterized in this article are shown in Figure 1. We decided to base our design upon the known tubule-forming lipidated tripeptide 2.16,18 As shown by the molecular rendering in Figure 2A, a short tether polypeptide AGGAA was added to the N-terminus of the

Langmuir, Vol. 15, No. 17, 1999 5503

Figure 2. (A) Molecular rendering of Ac-GRAGGAAPPPE(HNC14H29)2 (compound 3) in the binding pocket of porcine pancreatic trypsin (EC 3.4.21.4).50 The arginine of 3 was placed in the specificity pocket of trypsin. The peptide-amphiphile was designed so that the lipid anchoring derivative was well outside of the trypsin binding pocket. (B) Molecular space-filling rendering of a closed-packed monolayer lattice comprised of 3, which is based upon a hexagonal array of fully extended lipidated peptides using the areas per headgroup of related lipid-peptide conjugates determined from Langmuir isotherms.24,25 The access of the trypsin binding sites are sterically blocked.

tubule-forming core so that the hydrophobic dialkylamide tails would extend out from the trypsin binding pocket. Small, nonhydrophobic amino acids were chosen to keep the headgroup area small and to facilitate efficient packing between lipids. The trypsin cleavage product AcGR- was attached to the N-terminus of the polypeptide tether and was radiolabeled for easy quantification of hydrolysis kinetics. Tritioacetylation of the peptide headgroup was achieved using an improved radiolabeling method that employs the sodium salt of [3H]acetic acid instead of using [3H]acetic anhydride, which is volatile, difficult to handle safely, and easily hydrolyzed. To mimic an internal trypsin cleavage site buried within the headgroup, Ac-G was added to cap the N-terminus of arginine within the trypsin cleavage product. The molecular rendering depicted in Figure 2B represents a conceptualized monolayer lattice of closed-packed, fully extended lipidated peptides and is based upon the areas per headgroup of related lipidpeptide conjugates determined from Langmuir isotherms.24,25 Within the bilayer matrix, the enzymatic cleavage sites are clearly blocked by neighboring lipidated peptides. Figure 3 shows a digitized transmission electron micrograph of typical tubule microstructures formed by the precipitation of 3 from an alcoholic solvent (e.g., methanol) upon the addition of water and after several solvent exchange steps, which were required to remove the alcohol. The lipid aggregates in Figure 3 have parallel tracks of bilayer walls running the length of the microstructures, a feature which has become the hallmark signature of hollow tubules in TEM. The tubule diameters are uniform (0.50 ( 0.071 µm), and their lengths range from 4 to 30 µm. Helical ribbon microstructures are found (24) Yu, Y.-C.; Pakalns, T.; Dori, Y.; McCarthy, J. B.; Tirrell, M.; Fields, G. B. Methods Enzymol. 1997, 289, 571-587. (25) Fields, G. B.; Lauer, J. L.; Dori, Y.; Forns, P.; Yu, Y.-C.; Tirrell, M. Biopolymers 1998, 47, 143-151.

5504 Langmuir, Vol. 15, No. 17, 1999

Figure 3. Digitized transmission electron micrographs of AcGRAGGAAPPPE(HNC14H29)2 tubules (100 µM lipid) obtained immediately after the addition of 0.9 µM trypsin. The tubule diameters are uniform (∼0.5 µm), and their lengths range from 4 to 30 µm. The tubule microstructures are virtually unchanged by trypsinolysis (not shown). Scale bar ) 1 µm.

Figure 4. Initial rates of [3H]Ac-GRAGGAAPPPE(HNC14H29)2 (compound 3) trypsinolysis in the presence of low trypsin (3.5 nM) as a function of the concentration (moles of [3H]-3 per total reaction volume) of mixed-micellar [3H]-3 (Triton X-100 concentration was held constant at 2 mM) and tubular [3H]-3.

ensheathing a minority (52 µM (linear response in Figure 4), one would have expected the rate to increase with increasing tubule concentration if enzyme hydrolyzed the tubular lipidated peptide (unless the enzyme binds tubular [3H]-3 much tighter than micellar [3H]-3, which is unlikely). Thus, the invariance of the rate versus tubule concentration (Figure 4) suggests that aggregated lipidated peptide is not hydrolyzed at all by trypsin. If true, the rate of cleavage of substrate presented as tubules should approximately equal the rate of cleavage of detergent-solubilized [3H]-3 at the concentration of lipidated peptide equal to the critical aggregation concentration (CAC) that is in equilibrium with tubular amphiphile. This assumes that the rates of hydrolysis of monodisperse and mixed-micellar [3H]-3 are the same; rates for trypsinolysis of [3H]-3 in the presence of 2 and 10 mM Triton X-100 were identical, showing that the detergent does not inhibit trypsin. The two curves in Figure 4 should intersect at the CAC of [3H]-3. The intersection value of 4 µM is not an unreasonable value for the CAC of [3H]-3 given the CAC values reported for related lipidated peptides (see Figure 1).16 The possibility of the protease interacting with the tubular form of substrate at more disordered, solvent exposed, ends and helical edges was examined by incubating the tubular and mixed micellar forms of [3H]-3 with 250-fold more trypsin than in the above experiments. Figure 5 shows the hydrolysis progress curves measured over 33 h. Since each tubule was made of ∼108 lipidated peptides, ∼106 enzyme molecules per tubule are present. For mixed-micellar [3H]-3, most (∼86%) of the lipidated peptide was hydrolyzed in the first 10 min. With tubular [3H]-3, a small burst of hydrolysis (2% of total [3H]-3), equal to ∼3 µM (again close to the CAC as estimated from the above experiment), was observed in the first 10 min followed by a much slower phase of hydrolysis. Even after 33 h, only 8% of total tubular [3H]-3 was hydrolyzed. The rate of hydrolysis in the slow phase was 0.400 nM‚min-1, which is 32,000-fold slower than the rate of hydrolysis of detergent solubilized [3H]-3 (Figure 5, when one accounts for the different amounts of [3H]-3 used). This enormous protection from proteolysis by self-assembly indicates, at the very least, that the rate of desorption of [3H]-3 from the tubule into the aqueous phase is slow relative to the rate of trypsin-catalyzed depletion of the CAC amount of [3H]-3 in the aqueous phase. The slow phase could

Protection of a Decapeptide

represent hydrolysis only of monodisperse [3H]-3 as it slowly leaves the tubule, slow hydrolysis of tubular [3H]3, or a combination of both. If tubular [3H]-3 is slowly hydrolyzed by trypsin, this could occur in regions of high relative disorder such as tubule ends and helical edges where the cleavage site would be more accessible to trypsin as we observed for hydrolysis of diacetylenic phosphatidylcholine tubules by the enzyme phospholipase A2.17 Both solution-phase and tubular-phase trypsinolysis models are consistent with the observation that the rate of the slow phase increased only 1.5-fold when the concentration of trypsin was increased from 0.42 to 1.7 µM (not shown). For example, if solution-phase 3 is the substrate and if the rate-determining-step is the transfer of 3 from the tubular to the aqueous phase, then an increase in the amount of trypsin would not increase the hydrolysis rate. On the other hand, if lipid within tubules is hydrolyzed, then the “defect sites” on the tubule surface may have become saturated with trypsin and adding more trypsin would not increase the rate. Transmission electron microscope images of tubules of [3H]-3 before and after trypsinolysis (5% of [3H]-3 hydrolyzed) showed no apparent differences in aggregate morphology, as expected (i.e., partially hydrolyzed tubules had similar morphologies to those in Figure 3). Raman Spectroscopic Analysis of Headgroup Packing. The 32,000-fold difference seen in the rates of hydrolysis for the lipidated peptide when dispersed in detergent micelles versus when embedded within a tight, crystalline bilayer matrix could arise, in part, from the relief of steric hindrance to trypsin binding achieved upon bilayer solubilization. Even modest changes in headgroup conformation could affect many membrane properties, such as gross morphology of the aggregate, ligand accessibility to trypsin, mass transport kinetics, and physicochemical integrity of the bilayer. Lattice statistical mechanics models predict that surfaces, including bilayers, should enhance secondary structural formation in attached peptides by minimizing the number of unfolded configurational states,28,29 and experimental data have supported this.24,25 Tight molecular packing of tubuleforming surfactants could constrain the molecular configurations of their polypeptide headgroups and slow hydrolysis. Well-ordered headgroup packing may be a natural consequence of bilayer formation, which is lost upon detergent solubilization, and insights into the configurational states of the peptide headgroup are necessary to fully interpret trypsin hydrolysis data. Raman spectroscopy has been widely used to investigate phospholipid configuration and dynamics in model and biological membranes,30-33 and it has become an equally powerful tool for assessing peptide secondary structure.34-38 (28) Chan, H. S.; Wattenbarger, M. R.; Evans, D. F.; Bloomfield, V. A.; Dill, K. A. J. Chem. Phys. 1991, 94, 8542-8557. (29) Wattenbarger, M. R.; Chan, H. S.; Evans, D. F.; Bloomfield, V. A.; Dill, K. A. J. Chem. Phys. 1990, 93, 8343-8351. (30) Spiker, R. C.; Levin, I. W. Biochim. Biophys. Acta 1976, 455, 560. (31) Gaber, B. P.; Peticolas, W. L. Biochim. Biophys. Acta 1977, 465, 260. (32) Levin, I. W. Vibrational spectroscopy of membrane assemblies; Clark, R. J. H., Hester, R. E., Eds.; John Wiley & Sons: New York, 1984; Vol. 11, pp 1-48. (33) Huang, C.-H.; Lapides, J. R.; Levin, I. W. J. Am. Chem. Soc. 1982, 104, 5926-5930. (34) Spiro, T. G.; Gaber, B. P. Annu. Rev. Biochem. 1977, 46, 553572. (35) Williams, R. W. J. Mol. Biol. 1983, 166, 581-603. (36) Williams, R. W. Methods Enzymol. 1986, 130, 311-331. (37) Williams, R. W.; Chang, A.; Juretic, D.; Loughran, S. Biochim. Biophys. Acta 1987, 916, 200-204. (38) Trabbic, K. A.; Yager, P. Macromolecules 1998, 31, 462-471.

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Figure 6. Raman scattering vibrational spectra obtrained from hydrated capillary samples of the unmodified peptide AcGRAGGAAPPP and the lipidated peptides AcGRAGGAAPPPE-(NC14)2 (compound 3), PPPE-(NC14)2 (compound 2), and E-(NC14)2 (compound 1). The sample temperature was thermostated at 25 °C. (A) The spectral region containing the skeletal C-C stretching modes (i.e., 1050-1150 ∆ cm-1) and the amide I vibrational modes (i.e., 1620-1720 ∆ cm-1). (B) The spectral region containing the C-H stretching vibrational modes.

Figure 6 shows Raman vibrational spectra obtained for the nonmodified peptide AcGRAGGAAPPP and the lipidated peptides 1-3 and at a temperature (∼25 °C) well below their respective lipid chain melting temperatures of 61.0, 51.2, and 39.9 °C, respectively. Consequently, all spectra of lipidated compounds presented in Figure 6 were obtained from suspensions of crystalline tubules. The amide I stretching regions (1620-1720 ∆ cm-1) for all compounds presented in Figure 6A are broad, which is a feature seen in Raman spectra taken of aqueous peptide and protein samples. For the aqueous peptide, the position and shape of the peak at 1660 ∆ cm-1 (amide I band) are consistent with a peptide configuration that is neither fully R-helical nor β-sheet but, rather, more disordered. The amide I stretching modes in spectra obtained for suspensions of lipidated peptides 1-3 are more interesting. Of particular note is the sharp, narrow vibrational band centered at 1660 ∆ cm-1 that is found superimposed upon a more typical broad amide I stretching band and is found only in spectra obtained for lipidated peptides that contain proline (see 2 and 3 in Figure 6A). The sharpness of this spectral feature is unusual compared to spectra of other peptides and proteins, and it implies that a well-ordered molecular headgroup configuration exists within these bilayers. Interestingly, this sharp spectral feature at 1660 ∆ cm-1 is absent in tubule samples composed of the nonproline containing lipid 1, which suggests that its origin may arise from the ordered, crystalline packing of the amide bonds within the peptide headgroup and not from ordering of the amide bonds found within the fatty amide tails.

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The skeletal C-C stretching mode regions from 1050 to 1150 ∆ cm-1 in Figure 6A reflect the intramolecular trans/gauche conformational changes within the bilayer aliphatic chains.31,32,39-42 The three “all-trans” alkyl chain C-C stretching modes are assigned to bands at 1062 ∆ cm-1 (out-of-phase skeletal stretching motions), 1098 ∆ cm-1, and 1128 ∆ cm-1 (in-phase skeletal stretching motions).40,42 These bands are particularly strong features in phospholipid spectra, and they are distinct features in these spectra obtained for lipopeptide suspensions of 1-3 (note their absence in the spectrum obtained for the unmodified peptide in Figure 6A). A feature assigned to gauche conformers along the aliphatic chains appears around 1080 ∆ cm-1,32,40-42 but it is very weak, if not absent entirely, at 25 °C suggesting that all lipid-modified peptide aggregates are crystalline. The free, unmodified decapeptide lacks the distinct skeletal C-C peaks at 1062 and 1128 ∆ cm-1, but a strong C-C stretching peak is visible at 1100 ∆ cm-1. The spectral region from 2700 to 3100 ∆ cm-1 (Figure 6B) is quite complex because it contains several strong, overlapping bands that arise from asymmetric and symmetric C-H stretches.43 Not surprisingly, the ν(CH2) symmetric and ν(CH3) symmetric stretching modes at 2849 and 2880 ∆ cm-1, respectively, are very strong and pronounced in all lipidated compounds due to contributions from the aliphatic hydrocarbon tails. These modes are much weaker, if not absent entirely, in the spectrum obtained for the nonlipidated peptide. The ν(CH2) asymmetric and ν(CH3) asymmetric stretching modes at 2935 and 2985 ∆ cm-1, respectively, are well-defined in all samples. Figure 7 shows the 825-1725 and the 2650-3050 ∆ cm-1 Raman spectral regions obtained for aqueous suspensions of lipid 3 aggregates at various temperatures throughout the main phase transition, which occurs at 39.9 °C (as measured by DSC). The alkyl C-C backbone in 3 and in other lipids has greater mobility and disorder at temperatures above the chain-melting transition (T > 40 °C) when the lipids adopt a melted smecticlike phase having liquidlike in-plane order. The skeletal C-C Raman stretching vibrations reflect this property, as in Figure 7A. As the temperature is increased, the intensity of 1127 ∆ cm-1 band decreases while the intensity of the 1078 ∆ cm-1 band increases. The decrease in the 1127 ∆ cm-1 band is attributed to a decrease in the amount of “alltrans,” fully extended alkyl chains, whereas the 1078 ∆ cm-1 band arises from hydrocarbon chain conformations containing an increased number of gauche rotamers.44 Figure 7B shows temperature-dependent changes in the C-H stretching region for 3. When lipids are in a liquid crystalline state, the symmetrical-Raman stretching vibrations of the amide CH2 groups at about 2849 ∆ cm-1 dominate. However, when lipids are in a crystalline state, with close packing of planar zigzag chains, the intensity of CH3 stretching vibrations at 2880 ∆ cm-1 dominates.44 The spectra in Figure 7B show how the intensity at 2880 ∆ cm-1 is reduced when lateral packing between chains (39) Gaber, B. P.; Yager, P.; Peticolas, W. L. Biophys. J. 1978, 24, 677-688. (40) Gaber, B. P.; Yager, P.; Peticolas, W. L. Biophys. J. 1978, 21, 161-207. (41) Gaber, B. P.; Yager, P.; Peticolas, W. L. In Applications of Raman spectroscopy to biomembrane structure; Theophanides, T. M., Ed.; D. Reidel Publishing Co.: Dordrecht, The Netherlands, 1979. (42) Batenjany, M. M.; Wang, Z.-q.; Huang, C.-h.; Levin, I. W. Biochim. Biophys. Acta 1994, 1192, 205-214. (43) Lawson, E. E.; Anigbogu, A. N. C.; Williams, A. C.; Barry, B. W.; Edwards, H. G. M. Spectrochim. Acta. Part A 1998, 54, 543-558. (44) Tu, A. T. Raman spectroscopy in biology: principles and applications; John Wiley & Sons: New York, 1982.

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Figure 7. Raman scattering vibrational spectra taken for a hydrated sample of AcGRAGGAAP3E-(NC14)2 (3) at several temperatures below and above its main phase transition temperature of 39.9 °C: (A) spectral region containing the skeletal C-C stretching modes (i.e., 1050-1150 ∆ cm-1) and the amide I vibrational modes (1620-1720 ∆ cm-1); (B) spectral region containing the C-H stretching vibrational modes; (C) An expanded view of the amide I vibrational region.

is disrupted, and they reflect a transition of the aggregate from a solid-crystalline to a liquid-crystalline phase. Figure 7C shows the temperature dependence of the amide I stretching region for 3. The molecular origin of the aforementioned sharp band at 1660 ∆ cm-1 with its small shoulder at 1670 ∆ cm-1 is unclear, but both bands decrease in intensity as the hydrocarbon chains melt, which indicates that the peptide headgroups become less ordered and adopt a larger degree of conformations at higher temperatures. At temperatures above the phase transition temperature, the amide I stretching band broadens and becomes more characteristic of the amide I region for water-soluble peptides. Also shown in Figure 7C is the amide I region for 1 that has been normalized to reflect the fewer number of amide bonds. The sharp vibrational band at 1660 ∆ cm-1 found in 3 is noticeably absent in the spectrum of 1. Analogous to what has been demonstrated for the melting of phospholipid acyl chains, the C-H stretching

Protection of a Decapeptide

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Figure 9. Circular dichroism spectra obtained for aqueous, unmodified peptide AcGRAGGAAPPP and lipidated peptide AcGRAGGAAP3E-(NC14)2 codispersed in (hexadecyldimethylammonio)propane-1-sulfonate mixed micelles. The spectra are markedly similar and consistent with CD spectra obtained for other proline-rich peptides.

Figure 8. Temperature dependence of semiqualitative Raman order parameters measured for aqueous tubule suspensions of P3E-(NC16)2 (A, C, E) and AcGRAGGAAP3E-(NC14)2 (B, D, F). The I1078/I1127 parameter (A and B) is sensitive to alkylamide chain trans/gauche rotational isomerization. Both the I2847/I2885 (C and D) parameter and the I2938/I2885 (E and F) parameter are sensitive to interchain order/disorder.

modes of the fatty amide chains are particularly sensitive to molecular reorganization of the bilayer membrane. Several qualitative bilayer order parameters have been constructed from peak-height intensity ratios in phospholipid systems.32,42 In particular, the I2847/I2885 order parameter directly monitors hydrocarbon chain order/ disorder arising from interchain interactions.32,42 Likewise, the I2938/I2885 parameter furnishes an index of the degree of interchain order/disorder, but it is also sensitive to intrachain gauche/trans isomerization effects.32,42 In the alkyl chain skeletal C-C stretching mode region, the appearance of the gauche conformer band at 1078 ∆ cm-1 permits the I1078/I1127 peak-height ratio to be used as a probe for intrachain trans/gauche isomerization.32,42 The temperature dependence of these Raman order parameters are plotted in Figure 8 for aqueous suspensions of 2 and 3 tubules, which have main phase transition temperatures measured by DSC of 51.2 and 39.9 °C, respectively. In all cases, the midpoint of the change in Raman order parameters occurs at temperatures approximately 4-7 °C higher than the phase transition temperature determined calorimetrically. Although melting temperatures have been measured for concentrated samples of hydrated 2 and 3 by DSC, these Raman experiments represent the first direct measurement of the melting temperature in lipid tubules composed of 2 and 3. The disparity between melting temperatures measured by DSC and Raman spectroscopy most likely arises from the effect of pH. For example, compound 2 has a pKa between 6.8 and 7.0 (as determined from titration of 2 with base). At pH 7.4, the Tm for 2 measured by DSC is 51.2 °C. However, if the pH of the suspension is lowered to 6.0, the Tm for 2 increases to 56 °C. While suspensions of HARMs are usually buffered

at pH 7.4, the suspensions used for Raman spectroscopy were left unbuffered, and their pHs may have drifted below pH 7.4. Circular Dichroism Studies. The amide I stretching regions within spectra obtained of lipopeptide 3 at temperatures greater than its melting temperature and within spectra taken of the nonlipidated peptide are similar in shape and peak position, and presumably, the conformations of each compound are equally disordered. Likewise, solubilization of a crystalline HARM bilayer of 3 by detergents should also induce a major reorganization of the lipid headgroup. Unfortunately, satisfactory Raman spectra of detergent-solubilized lipid could not be obtained due to interfering Raman bands of the detergent and a large fluorescence background. As a result, the conformation of a detergent-solubilized lipidated peptide 3 was compared to that of the nonmodified peptide using circular dichroism spectroscopy to assess whether loss of peptide secondary structure (i.e., along with the loss of any steric protection of the trypsin cleavage site) occurs upon solubilization. The CD spectra for AcGRAGGAAPPP and 3 are shown in Figure 9. Qualitatively, both spectra are quite similar; each has a positive band in ellipticity centered at 230 nm (albeit much smaller for the detergent solubilized lipid) and a large negative band at 198 nm. A strong negative band in ellipticity at ∼204 nm and a weaker positive band at ∼229 nm are characteristic spectral features of the poly-Pro-II helical structure;45,46 however, with proline-rich peptides, it is often difficult to distinguish the CD of extended coiled structures such as the poly-Pro-II helix from those that are random coil.47 Nevertheless, these spectral features suggest that the peptide adopts a conformation resembling the poly-Pro-II structure and that the conformation of the detergentsolubilized lipidated peptide is much like the conformation adopted by the nonmodified peptide in solution. We hypothesize that solubilization of the lipidated peptide disorders the headgroup, helps to relieve unfavorable steric interactions between neighboring lipids, increases the extent of molecular motions, and accelerates hydrolysis by trypsin. Closing Remarks The peptide-amphiphile tubule reported here is a model drug delivery system, where the headgroup of the am(45) Pysh, E. S. J. Mol. Biol. 1967, 23, 587-99. (46) Rabanal, F.; Ludevid, M. D.; Pons, M.; Giralt, E. Biopolymers 1993, 33, 1019-1028. (47) Niidome, T.; Mihara, H.; Oka, M.; Hayashi, T.; Saiki, T.; Yoshida, K.; Aoyagi, H. J. Pept. Res. 1998, 51, 337-345.

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phiphile can be any desired peptide or other hydrophilic therapeutic agent. Many other peptides lipidated with glutamic acid dialkylamides form tubules and helical ribbons,13,15,16,18,21 and these tubules may find use as matrices for sustained-release of peptides. While the large dimensions of lipid tubules suggest that they will be of restricted utility in delivery of drugs from within the vasculature, they are ideally suited for use at intramuscular, subcutaneous, intraperitoneal, topical, and intratumor sites in which localization of the drug delivery system is desirable. For some bioactive peptides, lipidation has been shown to enhance permeability through cell membranes thus increasing absorption and bioavailability,48,49 but if cleavage of the peptide from the lipid conjugate is required for pharmacological activity, then such cleavage could occur by enzymes present in extracellular and, more importantly, cytosolic fluids. Lipidated peptides can be taken up by cells after they desorb from HARMs. Peptides could be released from their lipid anchors by intracellular processes such as the action of hydrolytic enzymes in the lysosome on monomeric lipidated peptides; however, peptides in HARMs would remain protected from proteolysis for extended periods. The tubule, then, would act as a depot for continuous release of lipidated drug by dissolution from the bilayer aggregate, and the rate of release could be controlled by varying the length of the lipid hydrocarbon chains and ultimately changing the lipid’s aqueous solubility. The present study establishes the protection of peptides toward proteolysis conferred by their self-assembly into lipid tubules. The concept of using tubule-based systems (48) Tomasi, J., T. B.; Tan, E. M.; Solomon, A.; Prendergast, R. A. J. Exp. Med. 1965, 101. (49) Nardelli, B.; Haser, P. B.; Tam, J. P. Vaccine 1994, 12, 13351339. (50) Huang, Q.; Liu, S.; Tang, Y. J. Mol. Biol. 1993, 229, 1022-36.

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for sustained drug delivery is being further evaluated. For example, lipidated peptide tubules may find utility as a single-shot vaccine delivery system, both by parenteral and oral routes, where protection from premature proteolytic degradation is of great concern. Current work aims at developing this and other self-assembled tubule systems into new drug delivery technologies. How the molecular packing of lipids with larger oligiopeptide headgroups affects tubule formation, the rate of enzymatic hydrolysis, and subsequent drug release by dissolution are research areas currently under investigation. Abbreviations Abbreviations used in this paper are as follows: ACN, acetonitrile; AcOH, acetic acid; BAW, butanol/acetic acid/ water; BOP, ((benzotriazol-1-yl)oxy)tris(dimethylamino)phosphonium hexafluorophosphate; CAC, critical aggregation concentration; CD, circular dichroism; CHCl3, choloroform; DMF, N,N-dimethylformamide; DSC, differential scanning calorimetry; ES-MS, electrospray mass spectrometry; Fmoc, 9-fluorenylmethoxycarbonyl; HBS, 20 mM HEPES, 120 mM NaCl, 1 mM EDTA, 0.2% NaN, pH 7.4; HPLC, high-performance liquid chromatography; MeOH, methanol; Pmc, 2,2,4,6,8-pentamethylchroman6-sulfonyl; TFA, trifluoroacetic acid; TFE: tetraflouroethanol; TLC, thin-layer chromatography. Acknowledgment. The authors thank Anatoly Lukyanov (DSC), Scott Gerber (mass spectrometry), Kim Trabbic Carlson (Raman spectroscopy), and Geeng-Fu Jang for their help and insight. Stewart Hendrickson and Kohei Yokoyama are acknowledged for their support and encouragement. This research was supported by a grant from the Whitaker Foundation to P.Y. and M.H.G. LA9900775