Molecular Structure of Glucopyranosylamide Lipid and Nanotube

A series of glucopyranosylamide lipids, N-(X-octadecenoyl)-β-d-glucopyranosylamine [X = 13-cis (1), 11-cis (2), 9-cis (3), 6-cis (4), and 9-cis,12-ci...
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Langmuir 2005, 21, 743-750

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Molecular Structure of Glucopyranosylamide Lipid and Nanotube Morphology Shoko Kamiya,† Hiroyuki Minamikawa,†,‡ Jong Hwa Jung,†,§ Bo Yang,‡ Mitsutoshi Masuda,†,‡ and Toshimi Shimizu*,†,‡ CREST, Japan Science and Technology Corporation (JST), Tsukuba Central 4, 1-1-1 Higashi, Tsukuba, Ibaraki 305-8562, Japan, and Nanoarchitectonics Research Center (NARC), National Institute of Advanced Industrial Science and Technology (AIST), Tsukuba Central 5, 1-1-1 Higashi, Tsukuba, Ibaraki 305-8565, Japan Received September 7, 2004. In Final Form: November 3, 2004 A series of glucopyranosylamide lipids, N-(X-octadecenoyl)-β-D-glucopyranosylamine [X ) 13-cis (1), 11-cis (2), 9-cis (3), 6-cis (4), and 9-cis,12-cis (5)] and their saturated homologue N-octadecanoyl-β-Dglucopyranosylamine (6), which differ in the position of a cis double bond in the C18 hydrocarbon chains, have been synthesized. The effect of the cis double bond position on the chiral self-assembly of each glycolipid has been examined by scanning electron microscopy, transmission electron microscopy, X-ray diffraction, UV, and circular dichroism (CD). The 11-cis derivative 2 was observed to self-assemble in water to form a uniform hollow cylinder structure with about 200-nm outer diameters in >98% yields. The obtained nanotubes from 2 showed the narrowest distribution of outer diameters and also gave a negative CD band around 234-236 nm, showing the largest CD intensity among the glycolipids investigated. Thus, we found that the position of a cis double bond significantly influences the homogeneity of the outer diameters as well as growth behavior of the self-assembled nanotube structures. Chiral molecular packing driven by a possible bending structure of the unsaturated glycolipids is playing a critical role in determining tubular morphology through molecular self-assembly.

Introduction Since the almost concurrent discovery by three research groups,1-3 lipid nanotubes with a hollow cavity submicrometer wide have received much interest because of their structural resemblance to carbon nanotubes in size,4,5 their well-defined hollow cylinders,6-8 and the potential chemical and physical properties.9-11 Once self-assembling can be rationalized, lipid molecules may be created to spontaneously assemble into a desired morphology without need for special instruments and consuming energy. This feature is a fascinating characteristic of molecular selfassembly when we fabricate one-dimensional nanostructures such as tubular structures.12-15 * To whom correspondence should be addressed. Telephone: +8129-861-4544. Fax: +81-29-861-4545. E-mail: tshmz-shimizu@ aist.go.jp. † CREST, JST. ‡ NARC, AIST. § Present address: Nano Material Team, Korea Basic Science Institute (KBSI), 52 Yeoeun-dong, Yusung-gu Daejeon 305-333, Korea. (1) Yager, P.; Schoen, P. E. Mol. Cryst. Liq. Cryst. 1984, 106, 371381. (2) Yamada, K.; Ihara, H.; Ide, T.; Fukumoto, T.; Hirayama, C. Chem. Lett. 1984, 1713-1716. (3) Nakashima, N.; Asakuma, S.; Kunitake, T. J. Am. Chem. Soc. 1985, 107, 509-510. (4) Iijima, S. Nature 1991, 354, 56-58. (5) Singh, A.; Chow, G. M.; Chang, E. L.; Markowitz, M. A. CHEMTECH 1995, 38-43. (6) Schnur, J. M. Science 1993, 262, 1669-1676. (7) Fuhrhop, J. H.; Helfrich, W. Chem. Rev. 1993, 93, 1565-1582. (8) Bommel, K. J. C. v.; Friggeri, A.; Shinkai, S. Angew. Chem., Int. Ed. 2003, 42, 980-999. (9) Evans, E.; Bowman, H.; Leung, A.; Needham, D.; Tirrel, D. Science 1996, 273, 933-935. (10) Karlsson, A.; Karlsson, R.; Karlsson, M.; Cans, a.-S.; Stroemberg, A.; Ryttsen, F.; Orwar, O. Nature 2001, 409, 150-152. (11) Frusawa, H.; Fukagawa, A.; Ikeda, Y.; Araki, J.; Ito, K.; John, G.; Shimizu, T. Angew. Chem., Int. Ed. 2003, 42, 72-74.

Though a limited number of natural and synthetic lipids have been well-documented to self-assemble into tubular structures, there are no definitive guiding principles to regulate nanotube morphology.16-24 Even after lengthy incubation, aqueous dispersions including self-assembled tubular structures contain, in some cases, twisted or helical coils2,25,26 and vesicles,27 which precede nanotube formation as intermediates. In this regard, Kulkarni et al. carefully examined the self-assembly of galactosylceramides into tubular structures depending on the total acyl chain length containing a single cis double bond.28,29 To control outer (12) Fuhrhop, J.-H.; Koening, J. Membranes and Molecular Assemblies: The Synkinetic Approach; The Royal Society of Chemistry: Cambridge, 1994. (13) Feiters, M. C.; Nolte, R. J. M. In Advances in Supramolecular Chemistry; Gokel, G. W., Ed.; JAI Press, Inc.: Stanford, CT, 2000; Vol. 6. (14) Shimizu, T. Macromol. Rapid Commun. 2002, 23, 311-331. (15) Fuhrhop, J.-H.; Wang, T. Chem. Rev. 2004, 104, 2901-2937. (16) Fuhrhop, J. H.; Spiroski, D.; Boettcher, C. J. Am. Chem. Soc. 1993, 115, 1600-1601. (17) Giulieri, F.; Guillod, F.; Greiner, J.; Krafft, M.-P. Chem.sEur. J. 1996, 2, 1335-1339. (18) Shimizu, T.; Kogiso, M.; Masuda, M. Nature 1996, 383, 487488. (19) Jung, J. H.; John, G.; Yoshida, K.; Shimizu, T. J. Am. Chem. Soc. 2002, 124, 10674-10675. (20) John, G.; Masuda, M.; Okada, Y.; Yase, K.; Shimizu, T. Adv. Mater. 2001, 13, 715-718. (21) Shimizu, T.; Masuda, M.; Minamikawa, H. Chem. Rev. 2005, submitted. (22) Georger, J. H.; Singh, A.; Price, R. R.; Schnur, J. M.; Yager, P.; Schoen, P. E. J. Am. Chem. Soc. 1987, 109, 6169-6175. (23) Rudolph, A. S.; Calvert, J. M.; Ayers, M. E.; Schnur, J. M. J. Am. Chem. Soc. 1989, 111, 8516-8517. (24) Schnur, J. M.; Shashidhar, R. Adv. Mater. 1994, 6, 971-974. (25) Jung, J. H.; Kobayashi, H.; Masuda, M.; Shimizu, T.; Shinkai, S. J. Am. Chem. Soc. 2001, 123, 8785-8789. (26) Spector, M. S.; Selinger, J. V.; Singh, A.; Rodriguez, J. M.; Price, R. R.; Schnur, J. M. Langmuir 1998, 14, 3493-3500 and references therein. (27) Spector, M. S.; Singh, A.; Messersmith, P. B.; Schnur, J. M. Nano Lett. 2001, 1, 375-378.

10.1021/la047765v CCC: $30.25 © 2005 American Chemical Society Published on Web 12/24/2004

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Kamiya et al. Chart 1

diameters, several research groups have so far developed new methodologies including metal-ion-assisted selfassembly,30,31 binary mixing of lipids,27,32 and subtle chemical modification of molecular structures.33,34 Although these approaches have produced remarkable changes in the outer diameters, the details about the homogeneity of the gross morphology are unknown. Recently we have reported functionalization of a glucopyranosylamide lipid containing a single cis double bond that self-assembles to form lipid nanotube hollow cylinders.35-37 Here we describe the chiral self-assembly behavior of a series of synthetic glucopyranosylamide lipids into hollow cylinder structures, whose structure strongly depends on the position of a cis double bond in the molecule’s C18 hydrocarbon chains. We were able to optimize the glycolipid structure for uniform nanotube self-assembly in >98% yields by altering the position of the cis double bond. Results and Discussion Phase Transition Behavior of Glycolipids. Previously we found that the presence or absence of a cis double bond in the C15 hydrocarbon tail of cardanyl (glucoside)s significantly affects self-assembled morphologies of highaxial-ratio nanostructures.20,38,39 For example, the incorporation of a cis double bond midway in the alkyl chain caused the formation of a nanotube structure, whereas the saturated homologue gave only helically twisted fibers.40 To follow up this finding, we have examined the effect of the unsaturation degree in the C16 hydrocarbon tails of p-alkylamide glucopyranoside on the self-assembled morphologies. In particular, we have focused on structural optimization of glycolipids by changing the position numbers of a cis double bond.19 Consequently, increasing the position number of a cis double bond from one to three in these derivatives proved to increase the population of self-assembled nanotubes. To get further insight into the correlation between the unsaturation position in the long hydrocarbon chains and the efficient conversion into nanotubular structures, we synthesized glucopyranosylamide lipids 1-6 (Chart 1), which differ in the position of a cis double bond in the C18 hydrocarbon chains. The fluidity of bilayer membranes at a given temperature is an important factor in the transformation from fluid vesicular to solid tubular assemblies.3,41,42 To probe fluidity, we examined the thermodynamic properties (28) Kulkarni, V. S.; Anderson, W. H.; Brown, R. E. Biophys. J. 1995, 69, 1976-1986. (29) Kulkarni, V. S.; Boggs, J. M.; Brown, R. E. Biophys. J 1999, 77, 319-330. (30) Markowitz, M.; Singh, A. Langmuir 1991, 7, 16-18. (31) Markowitz, M. A.; Schnur, J. M.; Singh, A. Chem. Phys. Lipids 1992, 62, 193-204. (32) Singh, A.; Wong, E. M.; Schnur, J. M. Langmuir 2003, 19, 18881898. (33) Thomas, B. N.; Corcoran, R. C.; Cotant, C. L.; Lindemann, C. M.; Kirsch, J. E.; Persichini, P. J. J. Am. Chem. Soc. 1998, 120, 1217812186. (34) Thomas, B. N.; Lindemann, C. M.; Corcoran, R. C.; Cotant, C. L.; Kirshch, J. E.; Persichini, P. J. J. Am. Chem. Soc. 2002, 124, 12271233. (35) Yang, B.; Kamiya, S.; Yui, H.; Masuda, M.; Shimizu, T. Chem. Lett. 2003, 32, 1146-1147. (36) Yang, B.; Kamiya, S.; Yoshida, K.; Shimizu, T. Chem. Commun. 2004, 500-501. (37) Yang, B.; Kamiya, S.; Shimizu, Y.; Koshizaki, N.; Shimizu, T. Chem. Mater. 2004, 16, 2826-2831. (38) John, G.; Minamikawa, H.; Masuda, M.; Shimizu, T. Liq. Cryst. 2003, 30, 747-749. (39) John, G.; Jung, J. H.; Masuda, M.; Shimizu, T. Langmuir 2004, 20, 2060-2065. (40) John, G.; Jung, J. H.; Minamikawa, H.; Yoshida, K.; Shimizu, T. Chem.sEur. J. 2002, 8, 5494-5500.

Table 1. Phase Transition Behavior of the Fully Hydrated and Dried Glycolipids 1-6 in H2Oa 1 2 3 4 5 6

c

in airb c

glycolipid

Tm (°C)

Tm (°C)

Tcld (°C)

ref Tme (°C)

13-cis 11-cis 9-cis 6-cis 9-cis,12-cis sat

88 71 58 f 33 105

159 148 156 133 90 153

189 192 206 201 200 194

27 13 11 29 -5 70

a Determined using DSC. b Determined using an optical microscope equipped with a heating stage. c Melting temperature. d Clearing temperature. e Melting temperature of fatty acids used for the synthesis.43 f Cannot be detected.

of the fully hydrated and dried glycolipids 1-6 by means of differential scanning calorimetry (DSC) and a polarizing optical microscope equipped with a heating stage. Table 1 summarizes the thermodynamic properties obtained for each glycolipid. The melting temperature Tm of each hydrated glycolipid (the gel-to-liquid crystalline phase transition temperature) parallels the melting points of the fatty acids used as starting materials.43 The Tm of the glycolipids 1-6 in water increased in the order 5 < 50 °C < 3 < 2 < 1 < 100 °C < 6. Among the three lipids 1-3, the Tm values give a minimum value for the 9-cis derivative 3. Solubility of each glycolipid into water also strongly depends on both the unsaturation position and numbers of a cis double bond. In particular, the 6-cis 4 and saturated 6 derivatives were found to be the least soluble as a result of their crystalline nature. Above the clearing temperature (Tcl) in air, caramelization took place for all the lipids. It should be noted here that all the glycolipids 1-6 exhibit amphitropic liquid crystallinity at temperatures above Tm, giving a smectic phase based on lamellar organization.38,44,45 Figure 1 shows polarized (41) Nounesis, G.; Ratna, B. R.; Shin, S.; Flugel, R. S.; Sprunt, S. N.; Singh, A.; Rister, J. D.; Shashidhar, R.; Kumar, S. Phys. Rev. Lett. 1996, 76, 3650-3653. (42) Spector, M. S.; Selinger, J. V.; Schnur, J. M. J. Am. Chem. Soc. 1997, 119, 8533-8539. (43) Gunstone, F. D.; Ismail, I. A. Chem. Phys. Lipids 1967, 1, 264269.

Glucopyranosylamide Lipid and Nanotube Morphology

Figure 1. Polarized optical micrographs for the dried and fully hydrated glycolipids 2 showing a typical smectic phase. Each micrograph was observed at (a) 173.3 °C and (b) 88.5 °C.

optical micrographs of the fully hydrated and dried glycolipids 2, giving the oily streak texture typical of the smectic phase. Self-Assembly of Glycolipids. For self-assembly, each glycolipid (1 mg) was dispersed into water (10-30 mL) at temperatures above the corresponding melting point Tm of the hydrated sample. In general, heating the mixture at 95 °C for 30 min was enough to get a homogeneous transparent solution. The obtained aqueous solutions were allowed to cool to an incubation temperature. The incubation temperature and their spent period strongly influence the self-assembled nanotubes.46-48 We compared the distribution of outer diameters for the lipid nanotubes formed from 2, which self-assembled under different incubation conditions, including (a) room temperature for 10 days, (b) 62 °C for 7 days, and (c) 62 °C for 12 days. Figure 2 shows the histograms of the outer diameters for the nanotubes obtained under different conditions. The (44) Jefferey, G. A. Acc. Chem. Res. 1986, 19, 168-173. (45) Wingert, L. M.; Jeffrey, G. A.; Jahangir; Baker, D. C. Liq. Cryst. 1993, 13, 467-470. (46) Chung, D. S.; Benedek, G. B.; Konikoff, F. M.; Donovan, J. M. Proc. Natl. Acad. Sci. U.S.A. 1993, 90, 11341-11345. (47) Thomas, B. N.; Safinya, C. R.; Plano, R. J.; Clark, N. A. Science 1995, 267, 1635-1638. (48) Spector, M. S.; Price, R. R.; Schnur, J. M. Adv. Mater. 1999, 11, 337-340.

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Figure 2. Histograms of the outer diameters for the selfassembled lipid nanotubes from 2, depending on the incubation conditions (a) at room temperature for 10 days, (b) at 62 °C for 7 days, and (c) at 62 °C for 12 days. The average (a.v.) and standard deviation (s.d.) values were evaluated for 250 pieces of nanotubes and are shown in each diagram.

incubation temperature used for conditions b and c is approximately 1 °C lower than the onset temperature associated with the DSC peak for Tm. The histogram pattern for the nanotubes from 2, which were incubated at room temperature [condition a], was found to give the smallest average outer diameter [Dout(a.v.) ) 200 nm] with the smallest standard deviation (s.d. ) 23 nm; Figure 2a). On the contrary, incubation under a higher temperature (62 °C) caused the broadening of the histogram. With an increase in incubation time, lipid nanotubes with somewhat wider outer diameters (>300 nm) were produced (Figure 2b,c). Transmission electron microscopy (TEM) revealed that the self-assembled molecular objects from the lipids 1-3 give nanotube structures as a major product under protocol a (Figure 3a-c). In particular, the lipid 2 proved to selfassemble into nanotube structures exclusively (Figure 3g). TEM images for the nanotubes from 2, as shown in Figure 3b, show no intermediate morphologies such as helical coils. Table 2 summarizes the average outer and inner diameters and membrane wall thicknesses, which were evaluated for 250 pieces of nanotubes by measuring distances on the TEM micrographs. The nanotubes from 2 possess well-defined hollow cylinders with inner diameters of 61 nm in average. Furthermore, scanning electron microscopy (SEM) displayed open-ended nanotube as-

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Figure 3. TEM images for the self-assembled molecular objects from the glycolipids (a) 1 (nanotube), (b) 2 (nanotube), (c) 3 (nanotube), (d) 4 (amorphous aggregate), (e) 4 (fiber), (f) 6 (deformed nanotube), and (g) 2 (magnified image for a single nanotube). Table 2. Self-Assembled Morphologies of 1-6 and the Obtained Size Dimensions for the Self-Assembled Lipid Nanotubes for 1 to 3a inner diamter Din

wall thicknessb

a.v.c

s.d.d

a.v.c

s.d.d

outer diameter Dout self-assembled morphology glycolipid

major

minor

(nm)

(nm)

(nm)

(nm)

a.v.c (nm)

s.d.d (nm)

1

nanotube (∼90%) nanotube (>98%) nanotube (∼90%) amorphous aggregate (∼80%) amorphous aggregate (∼70%) amorphous aggregate (>98%)

amorphous aggregate (∼10%) was not observed amorphous aggregate (∼10%) fiber (∼20%) nanotube (∼30%)

214

55

76

26

60

24

200

23

61

19

54

12

220

71

73

22

63

24

2 3 4 5 6

a Evaluated for 250 pieces of lipid nanotubes. b Thickness was also evaluated by measuring the distances of lipid nanotube walls. Therefore, the subtraction values of Din from Dout are not always consistent with the values twice that of the wall thickness. c Average value. d Standard deviation value.

semblies with very smooth surfaces, indicating no helical markings on the outer surfaces. The absence of helical markings on the nanotube structures is interesting in view of the extensive body of observations relating the chirality of the diacetylenic phospholipid [1,2-bis(tricosa-10,12diynoyl)-sn-glycero-3-phosphocholine] to helical handedness chirality.6,22-24 When the incorporation position of the cis double bond shifts from the C11 to the C13 carbon (closer to the methyl group end) or moved to the C9 carbon (closer to the amide linkage), the size distribution of outer and inner diameters for the obtained nanotubes from 1 or 3 becomes slightly less uniform in comparison to that of 2 (Figure 4a,c). The average outer diameters for the nanotubes 1 and 3 were evaluated to be 214 and 220 nm, respectively. But, the position of a cis double bond had no significant influence on the average outer diameters, whereas it significantly affects the distribution width (Figure 4). Interestingly, the lipid 2 produces the nanotubes with the narrowest distribution among the three lipids 1-3, which makes it the best candidate if uniformity is required. If the monoene

moiety is moved further (closer to the amide linkage), for example, the lipid 4 gave no well-defined tubular structures. Only amorphous aggregates (ca. 80%; Figure 3d) and solid fibrous objects (ca. 20%) are found (Figure 3e). Similarly poor nanotube formation was also observed for the diene 5 and the saturated homologue 6. The glycolipid 5 self-assembled into somewhat deformed nanotube structures in relatively lower yields (∼30%; Figure 3f). No lipid nanotubes were obtained from 6. X-ray Diffraction (XRD) of Lipid Nanotubes. To investigate molecular packing and orientation in the solid nanotubes, we measured powder XRD for freeze-dried lipid nanotubes from 1-3. Figure 5 displays the XRD patterns for the self-assembled nanotubes of 1-3. The XRD for the nanotube 1 contains a set of reflection peaks ascribable to d ) 4.44 (001), 1.12 (004), 0.90 (005), and 0.75 nm (006), supporting the 4.44-nm long-range ordering in the solid bilayer. The higher order of the nanotube formed from 1, as compared to those from 2 or 3, is responsible for this periodic pattern. On the other hand, both the nanotubes of 2 and the nanotubes of 3 gave a single reflection peak

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Figure 4. Comparison of the histograms for the outer diameters of the self-assembled lipid nanotubes from the glycolipids (a) 1, (b) 2, and (c) 3. The histograms were depicted for the 250 pieces of nanotubes.

Figure 6. (a) UV and (b) CD spectra for the molecular assemblies from 1-3 and 5 in water.

Figure 5. Powder XRD patterns in the small-angle region of the freeze-dried lipid nanotubes from (a) 1, (b) 2, and (c) 3.

ascribable to an identical long-range ordering d ) 4.53 nm (001). The analysis of the electron density profile for the hydrated nanotubes of 2 indicated that each bilayer consists of nonpolar (the acyl chain part) and polar parts (interlamellar water + the glucopyranosylamine), 1.6- and 2.9-nm thick, respectively.49 Considering that the extended C18:1 chain length for 2 is 1.8 nm from a space-filling (49) Unpublished results.

molecular model, we can conclude that the chain packs each other in an interdigitated manner. The relatively thicker membrane wall 54-63-nm wide (Table 2) should include 18-21 solid bilayers. Circular Dichroism (CD) of Lipid Nanotubes. Chiral molecular packing within the solid bilayer system is known theoretically50,51 and experimentally3,52,53 to play a crucial role in shaping twisted, coiled, and eventually tubular sheet structures. To confirm the chiral packing order in the self-assemblies formed from 1-3 and 5, we performed CD measurements of the aqueous solutions. Figure 6b displays the CD spectra obtained independent of the placement position or orientation of the cell used. The negative CD band around 234-236 nm and the shoulder around 254-260 nm are characteristic of the CD spectra of the nanotubes from 1-3. UV spectroscopy for the aggregated glycolipids 1-3 in water indicated the existence of UV absorption bands around both 230 and 260 nm (Figure 6a). The aggregated glycolipid 5 gave two (50) Selinger, J. V.; MacKintosh, F. C.; Schnur, J. M. Phys. Rev. E 1996, 53, 3804-3818. (51) Selinger, J. V.; Spector, M. S.; Schnur, J. M. J. Phys. Chem. B 2001, 105, 7157-7169. (52) Schnur, J. M.; Ratna, B. R.; Selinger, J. V.; Singh, A.; Jyothi, G.; Easwaran, K. R. K. Science 1994, 264, 945-947. (53) Spector, M. S.; Easwaran, K. R. K.; Jyothi, G.; Selinger, J. V.; Singh, A.; Schnur, J. M. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 1294312946.

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Figure 7. Differential UV spectra when the volume fraction of methanol gradually increased in the solvent. The obtained spectrum was subtracted from the original UV spectrum in water. The volume fractions of methanol were (a) 29, (b) 38, and (c) 44 vol %.

evident UV absorption bands around both 230 and 280 nm. The 280-nm band is ascribable to the diene moiety in 5. Methanol can dissociate the aggregation state of the glycolipids 1-3. When adding methanol to the aqueous solution of the nanotube 2 from 0 up to 60% fraction volume, while keeping the total lipid concentration constant, we observed a remarkable decrease in the UV absorbance with increasing the concentration of methanol. Figure 7 shows the differential UV spectra caused by the addition of methanol, in which each obtained spectrum was subtracted from that in water. All the differential spectra strongly suggest the presence of UV absorption bands consisting of the 234- and 260-nm bands. This means that the origin of these two bands is closely related to the aggregation of the lipids. UV spectra for the aqueous solution of the aggregated glycolipids 1 or 3 gave a shoulder around 255 nm (Figure 6a). On the other hand, UV spectroscopy for the nonaggregated glycolipids 1-3 in methanol displayed no remarkable absorption bands in the range of 230-270 nm (Figure 8a). To confirm the assignment of the CD band at 234-236 nm, we then measured UV spectra of related compounds, 9-cis-octadecenamide (7) and 9-cis-octadecene-1-ol (8; Chart 1), which append a cis double bond at the C9 position as well as the amide or hydroxyl moiety. The obtained UV spectra of 7 and 8 gave an absorption peak around 234-236 nm (parts b and c of Figure 8, respectively). In particular, the compound 7 carrying an amide linkage displayed a larger absorption coefficient as compared with the compound 8 lacking an amide linkage. These findings suggest that the CD band around 234-236 nm and shoulders at 255 nm are ascribable to strong intermolecular interactions of hybridized amide functionalities with cis double bonds and the intrinsic cis double bond, respectively. The aggregation effect on the UV spectra is more remarkable for the 234-236-nm band as compared to that of the 255nm band. At temperatures above Tm CD activity disappears. For example, at 80 °C, only nanotubes of 1 gave a negative CD band at 240 nm (the Tm of 1 is 88 °C). Figure 9 clearly

Kamiya et al.

Figure 8. UV spectra of (a) 2, (b) 9-cis-octadecenamide 7, and (c) 9-cis-octadecen-1-ol 8 in methanol.

Figure 9. Temperature dependence of the CD spectra for the lipid nanotubes from 2 in water.

displays the dramatic decrease in CD activity of 2 around Tm (71 °C). These results are compatible with the wellknown interpretation that chiral molecular packing in the solid bilayer system is responsible for the nanotube morphologies.27,48,54 If the long hydrocarbon chains of the molecules are in the fluid state, the self-assembled morphologies convert into spherical vesicles showing no CD activity. The CD band intensity at 235 nm increases in the order of 3, 1, and 2 at 25 °C (Figure 6b). Interestingly, this order corresponds well to the tendency in the uniformity of the outer diameters. Chiral packing order in the nanotube of 2 appears to be the highest, resulting in the narrowest distribution. Continuum theory for the lipid nanotube formation predicts that the magnitude of the molecular chirality and the molecular tilt with respect to the bilayer planes could control outer diameters.55-58 (54) Spector, M. S.; Easwaran, K. R. K.; Selinger, J. V.; Singh, A.; Schnur, J. M. Polym. Prepr. (Am. Chem. Soc., Div. Polym. Chem.) 1996, 37, 482-483.

Glucopyranosylamide Lipid and Nanotube Morphology

The 11-cis derivative 2 should be favorable to satisfy these requirements among the six glycolipids synthesized. Conclusion A series of glucopyranosylamide derivatives possessing C18 unsaturated hydrocarbon chains, which differ in the introductory position of a cis double bond, were synthesized as building blocks for nanotube assemblies. We have characterized diameter distributions as a function of the ene position, finding substantial changes in dispersity as the cis double bond moves. Among the six derivatives synthesized, the 11-cis derivative has proven to selfassemble in water to produce uniform nanotube structures exclusively with significantly lower diameter dispersity than those of the other lipids. Thus, we have established limits to which the molecule may be modified (that is, ene position) and still have nanotubes form. CD spectroscopy suggested that the chiral packing order of this molecule in the tubular assembly directs the distribution of outer diameters. Experimental Section General Methods. 1H (400 MHz) and 13C NMR (100 MHz) spectra were recorded with a Varian 400 NMR spectrometer. The melting (Tm) and clearing points (Tcl) for the dried glycolipids were measured on the first heating cycle using a polarized light microscope (Olympus BX50), and optical images were recorded with 3-CCD video camera (Olympus CS520MD). Facile Synthesis of Glycolipids. Without using any protection and deprotection reaction, we synthesized, in only two steps, the glucopyranosylamide lipids 1-6, which differ in the incorporation position of a cis double bond on the C18 hydrocarbon chains. Typically, d-(+)-glucopyranose as a starting material was reacted with ammonium hydrogen carbonate in water at 37 °C for 5 days. The reaction mixture was then desalted to purify glucopyranosylamine. Condensation of the obtained glucopyranosylamine with the corresponding fatty acid possessing different unsaturation positions was carried out with common peptide condensation reagents in dimethyl sulfoxide at room temperature. The glycolipids 1-6 were thus obtained as white solids in 2030% yields after conventional purification. Synthesis of N-(13-cis-Octadecenoyl)-β-D-glucopyranosylamine (1). Solid ammonium hydrogen carbonate (10 g) was added to an aqueous solution of D-(+)-glucose (1.00 g, 5.55 mmol in 25 mL). The mixture was stirred in an open vessel at 37 °C for 5 days. Ammonium hydrogen carbonate (the total amount of 50 g) was added at intervals to keep a portion of solid salt present in the mixture. The reaction mixture was cooled to 5 °C to precipitate ammonium hydrogen carbonate after thin-layer chromatography (TLC; eluent, ethyl acetate/acetic acid/methanol/ water ) 4:3:3:1 by volume) indicated no more conversion (Rf ) 0.40). The supernatant was deionized using a micro-acilyzer G1 (Asahi Chemical Co., Ltd.) equipped with an ion-exchange membrane aciplex cartridge AC-110-10 and then lyophilized. The yield was 0.88 g (88%). To the solution of 13-cis-octadecenoic acid (350 mg, 1.24 mmol) in dimethyl sulfoxide (1 mL) was added a mixture of 1-hydroxybenzotriazol (190 mg, 1.24 mmol) and (benzotriazol-1-yloxy)-tris-(dimethylamino)phosphonium hexafluorophosphate (1.65 g, 3.72 mmol) in dimethyl sulfoxide (0.5 mL), and the reaction mixture was stirred for 10 min. A solution of β-D-glucopyranosylamine in dimethyl sulfoxide (877 mg, 4.9 mmol/1.5 mL) was then added, and the mixture was stirred at 37 °C for 45 h. After TLC (eluent, chloroform/methanol ) 4:1 by volume) indicated no more conversion (Rf ) 0.54), the solution was chromatographed two times on silica gel columns (eluent, dichloromethane/methanol ) 5:1), followed by purification using a TSKgel HW-40S column (eluent, methanol). The yield was 166 mg (30%). (55) Helfrich, W.; Prost, J. Phys. Rev. A 1988, 38, 3065-3068. (56) Ou-Yang, Z.; Liu, J. Phys. Rev. A 1991, 43, 6826-6836. (57) Nelson, P.; Powers, T. Phys. Rev. Lett. 1992, 69, 3409-3412. (58) Selinger, J. V.; Schnur, J. M. Phys. Rev. Lett. 1993, 71, 40914098.

Langmuir, Vol. 21, No. 2, 2005 749 1H NMR (in dimethyl sulfoxide-d , at 25 °C): δ 0.86 (t, CH ); 6 3 1.24 (m, CH2); 1.47 (m, NHCOCH2CH2); 1.98 (m, CHdCHCH2); 2.08 (m, NHCOCH2); 3.03-3.65 (m, H-2, H-3, H-4, H-5, H-6a, H-6b); 4.43-4.95 (m, OH-2, OH-3, OH-4, OH-6); 4.69 (m, J1,2 ) 9.2 Hz, H-1); 5.33 (m, CH2dCH2); 8.24 (d, J1,NH ) 8.8 Hz, NHCO). 13C NMR (in dimethyl sulfoxide-d , at 25 °C): δ 13.9-35.5 6 (hydrocarbon); 61.0 (C-6); 70.1, 72.5, 77.7, and 78.6 (C-2, C-3, C-4, C-5); 79.5 (C-1); 129.7 (CH2dCH2); 171.2 (NHCO). Anal. Calcd for C24H45O6N: C, 64.98; H, 10.22; N, 3.16. Found: C, 63.82; H, 9.97; N, 3.17. FABMS: 444 (M + H)+ (calcd MW ) 443.6). Synthesis of N-(11-cis-Octadecenoyl)-β-D-glucopyranosylamine (2). The reaction was carried out similarly as described above and followed with TLC (eluent, chloroform/methanol ) 4:1 by volume). Rf ) 0.60. 1H NMR (in dimethyl sulfoxide-d , at 25 °C): δ 0.85 (t, CH ); 6 3 1.24 (m, CH2); 1.47 (m, NHCOCH2CH2); 1.98 (m, CHdCHCH2); 2.08 (m, NHCOCH2); 3.00-3.65 (m, H-2, H-3, H-4, H-5, H-6a, H-6b); 4.43-4.94 (m, OH-2, OH-3, OH-4, OH-6); 4.69 (m, J1,2 ) 9.2 Hz, H-1); 5.33 (m, CH2dCH2); 8.22 (d, J1,NH ) 9.2 Hz, NHCO). 13C NMR (in methanol-d , at 25 °C): δ 12.5-35.3 (hydrocarbon); 4 60.8 (C-6); 69.5, 72.0, 77.1, and 77.7 (C-2, C-3, C-4, C-5); 79.1 (C-1); 128.9 (CH2dCH2); 175.5 (NHCO). Anal. Calcd for C24H45O6N: C, 64.98; H, 10.22; N, 3.16. Found: C, 63.68; H, 10.02; N, 3.16. FABMS: 444 (M + H)+ (calcd MW ) 443.6). Synthesis of N-(9-cis-Octadecenoyl)-β-D-glucopyranosylamine (3). β-D-Glucopyranosylamine synthesis was similar to that of 1. Triethylamine (154 mL, 1.10 mmol) was added to a solution of β-D-glucopyranosylamine in methanol (40 mg, 0.22 mmol/10 mL). Oleoyl chloride (514 mL, 1.32 mmol) was then added, and the mixture was stirred at 0 °C for 19 h. After TLC (eluent, chloroform/methanol ) 4:1 by volume) indicated no more conversion (Rf ) 0.51), triethylamine was added for neutralization. The solution was chromatographed on a silica gel column (eluent, chloroform/methanol ) 4:1), followed by purification using a TSKgel HW-40S column (eluent, methanol). The yield was 24 mg (25%). 1H NMR (in dimethyl sulfoxide-d , at 25 °C): δ 0.85 (t, CH ); 6 3 1.24 (m, CH2); 1.47 (m, NHCOCH2CH2); 1.98 (m, CHdCHCH2); 2.07 (m, NHCOCH2); 3.00-3.63 (m, H-2, H-3, H-4, H-5, H-6a, H-6b); 4.45-4.98 (m, OH-2, OH-3, OH-4, OH-6); 4.69 (m, J1,2 ) 9.2 Hz, H-1); 5.32 (m, CH2dCH2); 8.26 (d, J1,NH ) 9.2 Hz, NHCO). Anal. Calcd for C24H45O6N: C, 64.98; H, 10.22; N, 3.16. Found: C, 63.05; H, 9.85; N, 2.93. FABMS: 444 (M + H)+ (calcd MW ) 443.6). Synthesis of N-(6-cis-Octadecenoyl)-β-D-glucopyranosylamine (4). The reaction was carried out similarly and followed with TLC (eluent, chloroform/methanol ) 4:1 by volume). Rf ) 0.57. 1H NMR (in dimethyl sulfoxide-d , at 25 °C): δ 0.85 (t, CH ); 6 3 1.24 (m, CH2); 1.49 (m, NHCOCH2CH2); 1.98 (m, CHdCHCH2); 2.10 (m, NHCOCH2); 3.00-3.64 (m, H-2, H-3, H-4, H-5, H-6a, H-6b); 4.45-4.94 (m, OH-2, OH-3, OH-4, OH-6); 4.69 (m, J1,2 ) 9.2 Hz, H-1); 5.33 (m, CH2dCH2); 8.24 (d, J1,NH ) 9.2 Hz, NHCO). Synthesis of N-(9-cis,12-cis-Octadecenoyl)-β-D-glucopyranosylamine (5). The reaction was carried out similarly and followed with TLC (eluent, chloroform/methanol ) 4:1 by volume). Rf ) 0.48. 1H NMR (in dimethyl sulfoxide-d , at 25 °C): δ 0.86 (t, CH ); 6 3 1.26 (m, CH2); 1.48 (m, NHCOCH2CH2); 2.02 (m, CH2CHd CHCH2CHdCHCH2); 2.08 (m, NHCOCH2); 2.74 (t, CHdCHCH2CHdCH); 3.04-3.65 (m, H-2, H-3, H-4, H-5, H-6a, H-6b); 4.454.95 (m, OH-2, OH-3, OH-4, OH-6); 4.70 (m, J1,2 ) 9.2 Hz, H-1); 5.33 (m, CHdCH); 8.24 (d, J1,NH ) 9.2 Hz, NHCO). Synthesis of N-Octadecanoyl-β-D-glucopyranosylamine (6). The reaction was carried out similarly and followed with TLC (eluent, chloroform/methanol ) 4:1 by volume). Rf ) 0.61. 1H NMR (in dimethyl sulfoxide-d , at 25 °C): δ 0.85 (t, CH ); 6 3 1.23 (m, CH2); 1.47 (m, NHCOCH2CH2); 2.08 (m, NHCOCH2); 3.00-3.63 (m, H-2, H-3, H-4, H-5, H-6a, H-6b); 4.44-4.95 (m, OH-2, OH-3, OH-4, OH-6); 4.69 (m, J1,2 ) 9.2 Hz, H-1); 8.23 (d, J1,NH ) 9.2 Hz, NHCO). DSC Measurement. Glycolipids (0.5-1.6 mg) and distilled water (15-22 mg) were added into an aluminum pan, and the pan was sealed. Heating and cooling scans over the temperature

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range of 15-140 °C were performed on a Seiko DSC 6100 differential scanning calorimeter equipped with a nitrogen gas cooling unit. TEM and SEM Observations. Unstained specimens for TEM were dripped onto a standard TEM grid, and excess water was blotted with filter paper. After drying in air, TEM was done with a Hitachi H-7000 instrument operated at 75 kV. Images were recorded on negative films. For SEM observation, the aqueous dispersions of the nanotubes (0.033-0.1 mg mL-1) were dripped onto an amorphous carbon grid and dried in air overnight. SEM was investigated using a JEOL JSM-5000. XRD Measurement. The XRD of a freeze-dried sample was measured with a Rigaku diffractometer (Type 4037) using graded d-space elliptical side-by-side multilayer optics, monochromated Cu KR radiation (40 kV, 30 mA), and an imaging plate (R-Axis

Kamiya et al. IV). The typical exposure time was 30 min with a 150-mm camera length. CD and UV Measurements. In 2 days after each glycolipid was dispersed in water, the solution containing the self-assembled glycolipids in water was put into a synthetic quartz cell (1 × 1 cm). CD studies were performed on a JASCO J-820 spectropolarimeter operating between 210 and 350 nm. Temperature control was provided by an automated Peltier and circular temperature control system. UV spectra were measured with the same sample cell, using a Hitachi U-3300.

Acknowledgment. B.Y. thanks the JSPS (Japan Society for the Promotion of Science) for financial support. LA047765V