Peptide Nanotubes

Encyclopedia of Nanoscience and Nanotechnology

www.aspbs.com/enn

Peptide Nanotubes Hiroshi Matsui The City University of New York, New York, USA

CONTENTS 1. Introduction 2. Development of Peptide Nanotubes 3. Characterization of Peptide Nanotubes 4. Peptide Nanotube Applications 5. Summary Glossary References

1. INTRODUCTION Recently, there has been enormous interest in nanomaterials such as nanotubes and nanowires which show superior electronic, magnetic, and mechanical properties that may not be obtained in the bulk states [1–5]. Therefore, it is natural for material scientists to use such excellent nanometer-sized materials as building blocks to build various miniaturized devices. One of the interesting applications is to build these nanotube-based devices in solution [6–11]. If nanotubes are soluble in aqueous solution (i.e., nanotubes are not aggregated in solution), they will be very practical in use for biological and medical applications. The water-soluble nanotubes are also advantageous to organize nanotubes in solution by using self-assembly and supramolecular processes. Among various nanotubes, peptide molecule-assembled nanotubes satisfy those features. Peptide nanotubes are water-soluble and functional groups of peptides such as amide and carboxylic acid can highly enhance their self-assemblies via hydrogen bonding in solution. Peptide nanotubes usually possess high affinity to biological molecules via hydrogen bonding with their amide and carboxylic acid groups and this feature enables peptide nanotubes to decorate with various molecules or to sense biological molecules. This versatile functionalization capability makes peptide nanotubes flexible templates to create various exotic nanotubes. Addition of functionality to peptide nanotubes can also be achieved by incorporating the functional groups into peptide monomers [12] via organic synthesis. Once this modified peptide monomer is assembled ISBN: 1-58883-064-0/$35.00 Copyright © 2004 by American Scientific Publishers All rights of reproduction in any form reserved.

into nanotubes, the functional groups are also incorporated in the nanotubes. However, one needs to be careful because addition of functional groups to peptide monomers may affect their assembled nanotube structures. In other words, the certain additions may totally prevent nanotube formation because molecular self-assembly is very sensitive to the chemical structures of monomers [13]. Therefore, optimization of growth conditions for peptide nanotubes may be necessary when peptide monomers are modified to add new functionalities. Another advantage of peptide nanotubes is that the nanotube structure and size can be controlled by external environments such as surfactant, solvent, temperature, and pH. One of the important factors to determine assembled nanotube structures is the degree of protonation in amide and carboxylic acid groups under these environmental changes. The protonation states of those groups have a significant effect on intermolecular hydrogen bonding and hydrophobicity of peptide monomers, which are driving forces of the peptide nanotube formation. Therefore, the strengths of hydrogen bonding and hydrophobicity of peptide monomers, controlled by the environments, determine the assembled nanotube structure. This feature allows us to synthesize peptide nanotubes in certain dimensions with high reproducibility by controlling experimental conditions. Once peptide monomers are assembled into peptide nanotubes, they are quite stable and can be used as building blocks for device fabrication in relatively harsh environments. Peptide assemblies may seem unstable; however, the peptide monomers are assembled into crystalline nanotubes via three-dimensional hydrogen bonds, which makes peptide nanotubes fairly rigid (See Fig. 4 later in this chapter). For example, the melting point for one of the peptide nanotubes is 235
C which is high enough for various device fabrication conditions [14]. In neutral solution, peptide nanotubes are stable for more than a year, although the solution must be microorganism free because it can consume peptide nanotubes. The third advantage of peptide nanotubes is that the nanotube structure can be well defined in the molecular level [15]. Since the locations and the orientations of specific functional groups of peptide monomers in the peptide nanotubes are understood, modification of peptide Encyclopedia of Nanoscience and Nanotechnology Edited by H. S. Nalwa Volume 8: Pages (445–455)

446 monomers can design chemical properties of inner and outer walls of peptide nanotubes respectively. Of course, peptide monomers have to be functionalized through appropriate chemical structures to be assembled as the nanotube form. For example, cyclic d,l,-peptide nanotubes were designed to have hydrophobic outer walls and hydrophilic inner walls, which were suitable to fabricate artificial ion channels [16]. For some peptide nanotubes, the nanotube structures and the sizes are not consistent with the ones expected from the peptide monomer structures. It seems that the assembled structure of peptide nanotubes should be predictable from the monomer structure since charge distribution and strength of intermolecular hydrogen bonding of peptide monomers at various pHs are well understood. However, the peptide monomer could undergo conformation changes via pH change and the structure of peptide nanotubes is very sensitive to the peptide monomer conformation. As a result, it is not always straightforward to predict the structure and the size of peptide nanotube from the chemical structure of the peptide monomer. It should be noted that the use of peptide nanotubes is not limited to biological applications. Biological recognitions of peptide nanotubes can be used to guide the nanotubes to desired locations in solution for nonbiological devices such as electronics and sensors. In this concept, device designs can be developed by biological interactions between the nanotubes and biological surfaces instead of patterning the designs with photolithography. This method may simplify nanometer-scale device fabrication procedures and reduce fabrication costs because synthesis of most of peptides described here is fairly inexpensive. This chapter consists of three major sections; “Development of Peptide Nanotubes,” “Characterization of Peptide Nanotubes,” and “Peptide Nanotube Applications.” Since this field is extremely young, there are more reports about peptide nanotube synthesis than their applications. However, recently several promising reports about their applications began to appear and they are summarized in Section 4.

Peptide Nanotubes

an important factor to determine the dimension of peptide nanotubes [21]. Figure 1 is an example illustrating how linear peptide monomers self-assemble into nanotubes [22]. The peptide amphiphile monomers in Figure 1a are assembled to orient the hydrophobic alkyl chain to the core and the amide group to the outer surface. While this assembled structure seems to have the same structure as micelles, protonation states of amides still play an important role to organize this structure, which is supported by the observation that the appearance of nanotube depended on pH of solution [23]. Since the nanotube structure from this type of linear peptide monomer is well defined (i.e., tail group inside and polar head group outside), functionalization of peptide nanotubes can be obtained by introducing the functional groups into the peptide monomer via chemical synthesis. If some functional groups are incorporated into the head groups, those groups appear on the outer surfaces of nanotubes, which determines the surface properties of nanotubes. For example, a certain peptide sequence for cell adhesion was incorporated into the head group of peptide to anchor cells onto the peptide nanotubes [22]. Figure 2 shows that nanotube structures are sensitive to chemical structures of peptide

2. DEVELOPMENT OF PEPTIDE NANOTUBES 2.1. From Linear Peptides to Nanotubes Self-assemblies of linear amphiphiles (i.e., molecules with charged head groups and nonpolar tail groups) have been studied to mimic biological membrane structures for decades. Those simple linear amphiphiles were selfassembled by hydrophilic/hydrophobic interactions between the amphiphiles and surfactants. Phase diagrams of those assemblies have been well established as functions of temperature and pressure [17]. Then it was natural to evolve from the simple amphiphile self-assembly to peptidederivatized amphiphile assembly to introduce a new parameter to control self-assemblies, hydrogen bonding [18–20]. Self-assembled peptide amphiphile structures are sensitive to pH that changes the strength of intermolecular hydrogen bonding between amide and carboxylic acid groups. When the peptide nanotube are self-assembled in aqueous solution, the hydrophobicity of peptide monomers also becomes

Figure 1. (A) Chemical structure of the peptide amphiphiles, highlighting five key structural features by 1 through 5. (B) Molecular model of the peptide amphiphiles. C: black, H: white, O: red, N: blue, P: cyan, S: yellow. (C) Schematic showing the self-assembly of the peptide amphiphile molecules into nanotubes. Reprinted with permission from [22], J. D. Hartgerink et al., Science 294, 1684 (2001). © 2001, American Association for the Advancement of Science.

Peptide Nanotubes

Figure 2. TEM images of peptide nanotubes from peptide amphiphiles with (a) 10 carbons in the hydrophobic chain and (b) 16 carbons in the hydrophobic chain. Reprinted with permission from [23], J. D. Hartgerink et al., Proc. Natl. Acad. Sci. USA 99, 5133 (2002). © 2002, National Academy of Science.

monomers. The transmission electron microscope (TEM) image in Figure 2a shows that the peptide nanotubes, formed from peptide amphiphiles with 10 hydrocarbons in the hydrophobic chain, aligned in parallel, while the peptide nanotubes from peptide amphiphiles with 16 hydrocarbons in the hydrophobic chain were grown shorter and aggregated in random orientation as seen in Figure 2b [23]. The comparison of those TEM images indicates that the chemical structure of peptide monomers is very sensitive to the nanotube structure. There is another type of linear peptide monomer with two amide head groups connected by a hydrocarbon tail group (called bolaamphiphile), which showed pH dependence in the assembled structures. This peptide bolaamphiphile monomer assembled into the helical form at pH 8 (Fig. 3a) and the nanotube form at pH 5 (Fig. 3b) [24]. This peptide monomer is assembled into the nanotube and the helical form via three-dimensional hydrogen between amide and carboxylic acid groups (Fig. 4a, c). X-ray diffraction and Raman microscopy indicate that lower pH induces stronger protonation to amide and carboxylic acid groups that results in stronger intermolecular hydrogen bonds [24–26]. This change increases the tilting angle of the peptide monomer alignment and more surface curvature of the

Figure 3. Visible microscopic images of self-assembled structures from the peptide bolaamphiphiles. (a) In a pH = 8 solution, the peptide bolaamphiphile assembled into helical ribbons. The width of the helices is about 2
m and the length varies from 10 to 40
m. The helical structures appeared after one week. (b) In a pH = 5 solution, the peptide bolaamphiphile monomers assembled into nanotube structures. The diameter of the nanotube is 10–1000 nm. The peptide nanotubes appeared after two weeks [24].

447

Figure 4. (a) Chemical structure of peptide bolaamphiphile monomer. (b) Chemical structure of peptide nanotube from the peptide monomer. The nanotube surface has free amide groups (shown by arrows and yellow squares) to immobilize biological molecules and ions to functionalize the peptide nanotubes [28–30, 62].

peptide assembly results in the nanotube formation as if a sheet of paper is rolled up to a cigar [24]. The observation of stripes in zigzags on the nanotubes in Figure 4b supports this nanotube-assembly mechanism. The monomer conformation of the peptide bolaamphiphile also shows a significant effect on the self-assembled form. The helix-to-nanotube transformation was observed in the peptide bolaamphiphile with the seven-hydrocarbon chain (Fig. 4a). However, the same peptide monomer with a six-hydrocarbon chain only forms nanotubes in pH 5 and the helical form did not appear at higher pH in solution. Spectroscopic studies indicate that the peptide nanotubes from the six-hydrocarbon chain peptide monomer are assembled via stronger hydrogen bonding than the peptide monomer with the seven-hydrocarbon chain [27]. It seems that the stronger intermolecular hydrogen bonding between peptide monomers with the sixhydrocarbon chain prevents the structural transformation between the helix and the nanotube. For this type of peptide nanotube, functionalization can be obtained by anchoring molecules onto the nanotube surfaces. While the peptide nanotube is assembled via intermolecular hydrogen bonds between amide groups and carboxylic acid groups, an interesting characteristic of the peptide nanotubes is that their free amide groups can capture and incorporate biological molecules such as DNAs, proteins, and porphyrins via hydrogen bonding (Fig. 4c) [28, 29]. This free amide group also captures metal ions such as Pt, Pd, Cu, Co, and Ni to form square planar complexes and the further electroless plating results in stable metallic coatings [30]. Therefore, instead of chemically modifying peptide monomer structure, this type of peptide nanotube can be used as an efficient template to produce various exotic nanotubes. Another scheme to produce peptide nanotubes is to fold linear peptide monomers to the helical structure (i.e., a shape like a rattled snake). Oligophenylacetylenes were folded into helical forms as shown in Figure 5 [31]. This structural transformation is induced by solvophobical interaction between backbone phenyl rings. The peptide nanotube from the oligophenylacetylenes preserves a hollow coil structure because the turning angle of the helical conformation is wide enough to prevent a close-packed

448

Figure 5. Oligophenylacetylene folding equilibrium between the open state and the helical folded structure [31]. Reprinted with permission from [39], D. T. Bong et al., Angew Chem. Int. Ed. Engl. 40, 988 (2001). © 2001, Wiley–VCH.

core. The folded peptide monomers produce nanotubes of 4 Å in diameter. The ease to add additional derivatives into oligophenylacetylenes is advantageous for this type of folding peptide nanotubes [32]; however, the dimensions of the nanotubes in both diameter and length may be limited by the size of the monomer unless these nanotubes are integrated by noncovalent interactions to form longer tubes [33]. Peptide nanotubes have been produced from various linear peptide monomers [34]. Whereas the peptide nanotube dimension via changes of peptide chain length, functional groups in the chain, and pH of solution has been studied systematically [14, 35, 36], the complexity is that those factors cannot be treated separately. For example, the pH change will alter charge distributions and intermolecular hydrogen bonds between peptide monomers. However, it will also change the peptide monomer conformation via intramolecular charge interactions and/or intramolecular hydrogen bonds [23]. Since self-assembled nanotube dimension is also very sensitive to the peptide monomer conformation, sometimes it is difficult to predict the formation of peptide nanotubes from peptide monomer conformations. However, it is advantageous that the formation of peptide nanotubes via molecular self-assembly is very reproducible and the size of peptide nanotubes is quite monodisperse under the proper nanotube growth conditions. This feature is strength for the use of self-assembly in the nanotube formation [37]. In fact, the sensitivity of the assembled structure changes with experimental conditions may turn to an advantage when the exotic combined structures are aimed to be fabricated by controlling the growth conditions. For example, the monomer in Figure 6, which self-assembled into a microsphere at pH 8 and a nanotube at pH 7, can be formed as the combined structure between the microsphere and the nanotube (i.e., dumbbell) by switching pHs between the spherical growth and the tubular growth conditions [38]. The monomers in Figure 6 were first assembled into microspheres in a pH 8 solution. Once the microspheres were grown in diameter of 1
m, the self-assembly condition was switched to the nanotube formation by tuning pH to 7. After 12 hours, the nanotubes (50 nm in

Peptide Nanotubes

Figure 6. Strategy to produce nanotube-bridge geometry by controlling the molecular self-assembly mechanisms between microspheres and nanotubes. (a) First, the monomers are self-assembled in a pH 8 solution. (b) Then the nanotubes are assembled on the microspheres by changing pH to 7. The nanotubes grow to connect with other microspheres. (c) TEM image of the nanotube bridge between the microspheres [38].

diameter) were observed between the microspheres. Therefore, the adjustment of experimental conditions during self-assembling peptide monomers alters the growth processes in the intermediate time and it will produce exotic nanotube-based structures such as bridges, dumbbells, and multibranches.

2.2. From Cyclic Peptides to Nanotubes Peptide nanotubes can also be self-assembled by stacking cyclic peptide monomers [39–41]. A typical assembling mechanism of a cyclic d,l-peptide monomer is shown in Figure 7 [42]. Those peptide rings were stacked through the backbone–backbone hydrogen bonding between neighboring amide groups. The sequence of octapeptide cyclo[-(l-Glnd-Ala-l-Glu-l-Ala)2 -] was used to prevent subunit association through Columbic repulsion in basic aqueous solution. A cyclic d,l-octapeptide containing bis-aspartic acid was also self-assembled into nanotubes via pH control of solution [43]. One of the simplest cyclic monomers, benzene, can also be stacked to grow as nanotubes if benzene is derivatized with right functional groups at the right positions. When benzene rings were derivatized with secondary

Figure 7. Schematic diagram of peptide nanotube assembly from cyclic d,l--peptides. Reprinted with permission from [39], D. T. Bong et al., Angew Chem. Int. Ed. Engl. 40, 988 (2001). © 2001, Wiley–VCH.

Peptide Nanotubes

amides at the 1,3,5-positions, those monomers stack with amide–amide intermolecular hydrogen bonding and  overlap (Fig. 8) [44]. At higher concentration of the peptide monomer, these stacks self-organized into spherical arrays [45]. This two-dimensional hexagonal array (Fig. 8c) can be directed with external electric fields because the stacked column has a macroscopic dipole moment parallel to the stacking direction. The stacked column can be self-organized into hexagonally arranged fibers or liquid crystalline phases by controlling chirality of the peptide monomer via chemical synthesis. Those types of cyclic peptide nanotubes may have limitations in the nanotube dimensional control over wide ranges. For example, the cyclic d,l-peptide monomer was sequenced between 9 and 13 Å in diameter and nanotubes needed to be bundled as arrays to obtain larger nanotube diameters [39]. To produce a larger diameter of peptide nanotubes from cyclic peptide monomers, the ring diameter of the cyclic peptide monomers needs to be larger. Whereas peptide sequencing may have limitations in the ring size of peptide monomers, combination of peptide molecules into an integrated cyclic structure via hydrogen bonding is possible. These integrated rings can be stacked in the length direction to form nanotubes. For example, a heteroaromatic bicyclic base G ∧ C, possessing the Watson–Crick donor– donor–acceptor of guanine and acceptor–acceptor–donor of cytosine, was self-assembled into nanotubes (Fig. 9a) [46]. This monomer formed a six-membered supermacrocycle held by 18 hydrogen bonds. Then this substantially more hydrophobic supermarcocycle self-assembled as a stack to produce nanotubes. The diameter of the nanotubes is 4 nm and the hydrodynamic radius is between 10 and 100 nm, depending on the monomer concentration. The inner space of the nanotube is determined by the distance separating the hydrogen bonding arrays within G∧C. The peripheral diameter and its chemistry depend on the choice of functional groups conjugated to this motif. Crown ether (Fig. 9a) was conjugated with the motif because of its broad

Figure 8. (a) Chemical structure of the monomer. (b) The intermolecular hydrogen bond conformation between monomers in the stacking direction. (c) The hexagonal arrays. (d) Model of a tetramer showing three helices of hydrogen bonds surrounding the exterior of the column. Reprinted with permission from [45], M. L. Bushey et al., Angew Chem. Int. Ed. Engl. 41, 2828 (2002). © 2002, Wiley–VCH.

449

Figure 9. Hierarchical self-assembly of heteroaromatic bicyclic base G ∧ C into a six-membered supermacrocycle and resulting nanotube. Each crown ether site within the assembled structure provides 328 Å3 of open space for binding of a molecular guest. Reprinted with permission from [49], H. Fenniri et al., J. Am. Chem. Soc. 124, 11064 (2002). © 2002, American Chemical Society.

solvent compatibility, synthetic accessibility, and functionality as versatile receptors [47]. The crown ether plays an important role in controling chiroptical properties by using chiral-promoting molecules, which induce a rapid transition from racemic to chiral nanotubes via electrostatic interactions between the crown ether moiety and the promoter [48]. The promoter molecules could even induce self-assembly of the monomer in the low concentration that the nanotubes were not assembled spontaneously [49]. The degree of chiral induction was sensitive to the chemical structure of the promoter. Similar cyclic peptide nanotube assemblies from noncyclic peptide monomers were reported with series of small dipeptides, l-Leu-l-Leu, l-Leu-l-Phe, l-Phe-l-Leu, and l-Phe-l-Phe [50]. One of characteristic features of the cyclic peptide nanotubes is the precise diameter control of the nanotubes, determined by the chain length, the size of side chains, and the bond angles of the peptide monomer. Proper side chain functionalization also enables those peptide nanotubes to possess distinct surface properties on the inner surfaces and the outer surfaces respectively. As pointed out in Section 1, hydrogen bond-driven self-assemblies are very naive against the environment. However, the peptide nanotubes from cyclic peptide monomers may be less sensitive to the external conditions such as pH and solvent because hydrogen bonds in only one direction need to stack the peptide monomers to produce the nanotubes. Therefore, the optimization of experimental conditions to grow nanotubes from the cyclic peptide monomers may be much more straightforward than the one from linear peptide monomers.

450 The cyclic peptide nanotubes from linear peptide monomers may require more delicate experimental optimization than sequenced cyclic peptide monomers because this system must first form peptide rings from hydrogen-bonding linear peptide monomers and then stack them into tubular structures.

3. CHARACTERIZATION OF PEPTIDE NANOTUBES Since chemical structures of peptide monomers are well defined, it is relatively straightforward to probe peptide nanotube structures if we have additional spectroscopic information. Conventional X-ray diffraction is a powerful spectroscopy to identify the nanotube structures [41, 50]. However, peptide nanotubes are necessary to grow as a perfectly aligned single crystal in the scale of mm3 to resolve fine structures of peptide nanotubes in the X-ray diffraction. Recently, synchrotron X-ray diffraction has been applied to peptide nanotubes. For example, the synchrotron X-ray diffractogram of 1a in Figure 8a resolved a single sharp peak at low angle that indicates their columnar assemblies and the hexagonal lattice structure was indexed by the diffraction peaks up to fifth order (Fig. 10) [44]. The fluidic packing of the side chain was also probed by the diffuse reflection at 4.5 Å [51]. The lateral core-to-core separation was 21 Å, which is consistent with the column separation with extended side chains, 27 Å [44]. Another technique to resolve peptide nanotube structures is vibrational spectroscopy. The most common vibrational spectroscopies to study chemical structures of peptide nanotubes are Fourier transform infrared spectroscopy and Raman spectroscopy [14, 16, 24, 35, 36, 52]. An advantage for these spectroscopies is that no sample preparation is required. Recently, Raman microscopy has been applied to study single carbon nanotubes [53] and peptide nanotubes [24, 54]. For peptide nanotubes, vibrational frequencies of amide groups are very important to monitor because the frequency shifts reflect the strength of hydrogen bonding with the amides. This information helps one to build the structural model: how the hydrogen bonding assembles peptide

Figure 10. Synchrotron X-ray diffraction of the nanotubes from 1a in Figure 8a. Reprinted with permission from [44], M. L. Bushey et al., J. Am. Chem. Soc. 123, 8157 (2001). © 2001, American Chemical Society.

Peptide Nanotubes

monomers into peptide nanotubes. For example, the structure of the peptide nanotube from the monomer (Fig. 4a) was investigated with a Raman microscope. When the structural transformation between the helical form (Fig. 3a) and the nanotube (Fig. 3b) was monitored, the helical phase (Fig. 11a) has a peak for the C O stretch at 1644 cm−1 while the nanotube (Fig. 11b) and the crystalline form (Fig. 11c) have two C O peaks at 1637 and 1660 cm−1 . The C O stretch with the highest frequency is assigned as the free C O of the amide group. The C O frequencies at 1637 and 1644 cm−1 are redshifted from the free C O stretch since these C O stretches bind the amide NH group via intermolecular hydrogen bonds (Fig. 4c). These assignments suggest that the C O group in the nanotube and the crystal phase binds to NH stronger than the C O group in the helical form due to a redshift of the vibrational frequency. This is consistent with the structural transformation model for the peptide bolaamphiphile that shortening amide–amide and acid–acid hydrogen bonds induces further tilt of the peptide monomer arrangement and makes the assembly surface more convex. This greater molecular tilting leads the assembled structure to the nanotubes by scrolling the sheetlike helical form into the tubular form (Fig. 4b). Nuclear magnetic resonance spectroscopy is an alternative way to study hydrogen bonding schemes of peptides in nanotubes [55]. Helicity of the peptide nanotube structure can be analyzed by CD spectroscopy [31, 45, 46]. The monomer concentration dependence of the structural change for the nanotubes stacked from monomer 1b in Figure 8a was probed by CD spectroscopy, as shown in Figure 12 [45]. When the solution of 1b in hexane was diluted with ca. 15% CH2 Cl2 , the helical order of nanotubes was observed

Figure 11. Raman spectra of (a) helical form, (b) nanotube, (c) crystal from the peptide monomer in Figure 4a [24].

451

Peptide Nanotubes

the crystalline minerals were observed on the nanotubes after 20 minutes [22]. The hydroxyapatite crystallographic c axis was aligned with the peptide nanotube long axis, which mimics the natural bone growth with collagen fibrils. The ability of the peptide nanotube to orient crystalline nuclei and subsequent crystal growth in the specific direction can be applied to other mineralized tissue repairs.

4.2. Electronics

Figure 12. CD spectra of the nanotubes from 1b in Figure 8a in hexane. Reprinted with permission from [45], M. L. Bushey et al., Angew Chem. Int. Ed. Engl. 41, 2828 (2002). © 2002, Wiley–VCH.

via an exciton coupling between degenerate chromophores. As the concentration of CH2 Cl2 increased, the helical complex dissociated and the transition dipoles were not observed to couple. The series of CD spectra show that the monomers self-assembles through hydrogen bonds with their side chains close enough to interact with each other only when the solvent is hydrocarbonlike [45].

4. PEPTIDE NANOTUBE APPLICATIONS 4.1. Tissue Engineering In nature, self-assembled fibrils and their biomineralization routinely produce various composite materials. Bone tissue is one of those examples. The collagen fibrils are formed by self-assembled collagen triple helices and the hydroxyapatite crystals grow in these fibrils with the hierarchical organization of their c axes along the long axes of the fibrils [56]. The peptide nanotube was applied as an artificial fibril to serve as crystal nucleation and growth of the inorganic hydroxyapatite crystals [22]. This peptide nanotube was self-assembled from the peptide amphiphile monomer shown in Figure 1a. This synthetic peptide monomer incorporates four important functions. The tail group (region 1 in Fig. 1a) and amide group (region 3 in Figure 1a) stabilize the nanotube structure via hydrophobic interactions and hydrogen bonds. The amide group also plays an important role in promoting the hydroxyapatite crystal growth because their negative charges established the local ion supersaturation [57]. The sulfide group (region 2 in Fig. 1a) provides extra stability to the nanotubes via disulfide bonds with adjacent peptide monomers. The phosphorylated group (region 4 in Fig. 1a) was introduced to nucleate calcium phosphate minerals. As this peptide monomer forms the peptide nanotube, the phosphoserine residue locates at the outer wall of the nanotube. This peptide nanotube with a highly phophorylated nanotube surface resembles the collagen fibrils. To proceed with the biomineralization efficiently, the fibrils should promote the adhesion growth of cells on their surfaces. The peptide sequence Arg–Gly–Asp (RGD), contained in collagen-associated fibronectin, is important for integrin-mediated cell adhesion [22] and this peptide sequence was also incorporated in the peptide monomer shown as region 5 in Figure 1a. When the assembled peptide nanotubes are treated with CaCl2 and Na2 HPO4 solution,

While various nanocomponents have been applied as building blocks to construct nanodevices [9–14], more reproducible methods to assemble them onto precise positions are desirable. The improved fabrication method will be extremely significant to interconnect electronic components with nanotubes. The configurations of nanotubes on those components depend on the type of devices to develop. For example, when two nanowires are crossed perpendicularly at the centers of the nanowires, this configuration acts as an electric switch [58]. More complex nanowire configurations of logic gates such as OR, AND, and NOR have been fabricated [2, 59]. One of the simpler ways to align nanotubes for complex device configurations is to use freestanding nanotubes in solution. This strategy will allow nanotubes to freely move and attach at the target locations in solution via noncovalent interactions, charge interactions, or molecular recognitions. However, it remains highly challenging to fabricate freestanding nanotubes in solution. Biologically compatible peptide nanotubes are soluble in aqueous solution and therefore their aggregations are not a concern. Therefore, the use of peptide nanotubes as building blocks for electronics and sensor devices may be appropriate in the solution-based fabrications. Our strategy has been to use those functionalized peptide nanotubes, which can recognize and selectively bind a well-defined region on patterned substrates, as building blocks to assemble three-dimensional nanoscale architectures at uniquely defined positions [8, 38, 60] and then decorate the nanotubes with various materials such as metals [30, 61], quantum dots [62], and metalloporphyrins [29] for electronics and sensor applications. Complex device configurations may be achieved by using peptide nanotubes incorporating multiple recognition functions such as antibodies and antigens in solution [28]. When a large number of the monodisperse peptide nanotubes are required for practical applications, the peptide nanotubes can be grown in microporous membranes [63]. In this case, the peptide nanotube diameter is controlled by a porous size of polycarbonate membranes. After the nanotubes are grown inside the membrane pores, the membranes are dissolved by CH2 Cl2 to extract the size-controlled nanotubes. When the peptide nanotubes are necessary to have higher mechanical strength for harsher environmental applications, the peptide nanotubes can be bundled via the ligand-ion bridge formations [54]. Patterning of peptide nanotubes in the desired device configurations is achieved before the functionalization of peptide nanotubes. We have demonstrated that the peptide nanotubes can be assembled as an array on Au surfaces via a 4-mercaptobenzoic acid self-assembled monolayer (SAM) [60]. The nanotube array was grown when peptide monomers were assembled to nanotubes with the SAM/Au substrates, as shown in Figure 13a. The SAM, with thiol

452

Figure 13. (a) Visible micrograph of the nanotubes grown on the 4-mercaptobenzoic acid monolayer/Au substrate in a citric acid/sodiumcitrate solution (black arrows shows the positions of nanotubes). (b) Proposed mechanism of the nanotube attachment on a mercaptobenzoic acid SAM/Au substrate [60].

groups at one end and carboxylic acid groups at the other end, serves as a junction between Au and the nanotubes because the thiol groups are attached firmly onto the Au surface while the carboxylic acid groups bind the carboxylic acid groups of the nanotubes via hydrogen bonds (Fig. 13b). To demonstrate the surface-specific immobilization of nanotubes for the nanotube-cross-linking configuration, we connected carboxylic acid groups of SAM/Au surfaces with carboxylic acid groups of the peptide nanotubes via hydrogen bonds (Fig. 14) [60]. All peptide nanotubes were placed on the SAMs parallel to the substrates when we processed the tube immobilization in two steps; first the peptide nanotubes were grown in suspension and then placed in solution containing the 4-mercaptobenzoic acid SAM/Au substrates. The carboxylic acid groups of peptide nanotubes recognized the carboxylic acid groups on the SAM via the

Figure 14. Visible micrograph of the peptide nanotubes immobilized onto two separated 4-mercaptobenzoic acid SAM/Au substrates. Two bright squares of the Au layers (10
m×10
m×0.2
m), patterned on a glass substrate by photolithography, were deposited by the 4-mercaptobenzoic acid SAM. The peptide nanotubes, assembled in suspension, were placed on the carboxylic acid SAM/Au substrate and then complete the immobilization in a pH 6 citric acid/sodium citrate solution [60].

Peptide Nanotubes

acid–acid hydrogen bonds to interconnect two substrates. After the site-specific immobilization of peptide nanotubes on the SAM surfaces, the nanotubes were coated with Ag by an electroless coating process (Fig. 14a). The surface of the nanotube was quite rough due to the nonuniform Ag coating on the nanotube and the concentrations of Ag ions and reducing agents need to be optimized to obtain smooth metallic coatings on the peptide nanotubes. Whereas the peptide nanotubes have been patterned into simple device configurations in solution, those nanotubes must be conductive for use in electronics and sensors. Using the characteristic of the peptide nanotubes that their free amide groups can intercalate metal ions (Fig. 4c) [30], sidewalls of the peptide nanotubes can be coated with metals via electroless plating to form metallic nanowires [24, 30]. The difficulty in organic nanotube metallization is the creation of organic–inorganic junctions on the nanotube surfaces. But the peptide nanotubes can overcome this difficulty through the intercalating of free amide sites. For example, Figure 15 shows an X-ray photoelectron spectroscopy (XPS) microscopic image of the Ni 2p peak on the single peptide nanotube [64]. In Figure 15, brighter colors correspond to higher concentration of Ni. This image indicates that a uniform Ni coating is possible on the peptide nanotubes via a simple electroless coating. Au nanocrystals can also be coated on the peptide nanotube surfaces and they behave as metallic or semiconductor depending on the size and the density of Au nanocrystals [65]. The strategy is to use carboxylic acid-thiol capped Au nanocrystals that can be intercalated by the free amide group on the peptide nanotubes [62]. Figure 16 shows the Au nanocrystal-capped nanotube developed by coating carboxylic acid-thiol capped Au nanocrystals on the peptide nanotube templates. The magnified TEM image in Figure 16c shows multilayer coatings of Au nanocrystals as dots on the peptide nanotubes seen as a dark continuous background. The coated Au nanocrystals have preferred crystal orientations as (111), (200), (220), and (311). Hydrogen bonding between the amide groups of the peptide nanotube and the carboxylic acid groups of the Au nanocrystals was used as a driving force for this fabrication process. While the peptide nanotubes can be conductive by coating metals on the nanotubes, recent computational studies

Figure 15. XPS microscopic image of Ni on the peptide nanotubes via electroless coating. The intensity of the Ni 2p peak on the substrate was mapped [64].

453

Peptide Nanotubes

Figure 16. (a) Scanning electron microscope image of the peptide nanotube coated with the carboxylic acid-terminated thiol-capped Au nanocrystals. (b) TEM image of the peptide nanotube coated with the carboxylic acid-terminated thiol-capped Au nanocrystals in the higher magnification around the surface of nanotube. (c) Electron diffraction pattern of the peptide nanotube coated with the carboxylic acidterminated thiol-capped Au nanocrystals [62].

showed that some of peptide nanotubes may be conductive without any metallic coatings. For example, cyclic d,loctapeptide nanotubes may be conductive due to delocalizations of electrons and holes through the hydrogen bonds between the peptide rings [66]. Because theory suggests that strengths of the intermolecular hydrogen bonds via the nanotube axis affect the electronic band-edge states sensitively [67], proper structures of peptide nanotubes may possess adequate electronic transport properties to build electronics. For example, the highest occupied molecular orbital/lowest unoccupied molecular orbital gap of the cyclic d,l-octapeptide nanotube was computed approximately as 5 eV [68].

4.3. Ion Channels and Biosensors When functionalized peptide nanotubes were incorporated into membranes, Sanchez-Quesada and co-workers demonstrated that this system mimics transmembrane channels [69]. Transmembrane channels involved in electrical signaling in cells display ion selectivity and rectification, gated by their potentials [70]. While monomeric peptide nanotubes incorporated in membranes were used for size selective transmembrane transport of small molecules [52], membranes with heteromeric peptide nanotube channels showed altered conductance and rectification properties. In this demonstration, peptide nanotubes from the cyclic d,l-peptide monomers (Fig. 7) were used as templates to incorporate into membranes. The template nanotubes were capped with two different subunits and these heteromeric peptide nanotubes were assembled in membranes with locating designed functional groups of the nanotubes at the channel ends (Fig. 17). The ion conductivity and conductance of the nanotube channels were modulated via local electrostatic perturbation from the cap subunits bearing ionizable functionalities. When l--phophatidyl-choline (l--lecithin) planar bilayers were applied as membranes and transferred into symmetric KCL solution, the conductance of the peptide nanotubes decreased after capping the nanotubes with a positively charged functional group (cap peptides 2 and

Figure 17. Chemical structures of the cyclic d,l--peptides and schematic representation of the mode of interaction of the tube cap informing heteromeric transmembrane channels. The cyclic peptide monomer self-assembles in lipid membranes to form the core transmembrane channel structure, while the peptides 2–5 serve as the cap subunits creating the heteromeric channel assembles with altered ion conductance properties. Reprinted with permission from [69], J. Sanchez-Quesada et al., J. Am. Chem. Soc. 124, 10004 (2002). © 2002, American Chemical Society.

4 in Fig. 17), while the conductance increased after the nanotubes were capped with a negatively charged functional group (cap peptides 3 and 5 in Fig. 17). If biological recognition functions are incorporated into the capping groups of peptide nanotubes, the membrane– nanotube complex will be applied to biosensors [39]. In this configuration, coupling of target antigen with antibody on the peptide nanotubes modulates the conductance, and the conductivity change will be analyzed to identify the target antigen. Thus, the introduction of the molecular recognition function to the membrane–nanotube complex will dramatically improve the signal-to-noise ratio and sensitivity [71, 72].

4.4. Antibacterial Agents The technique to incorporate peptide nanotubes into membranes as described in the previous section can also be applied to kill bacteria. When cyclic d,l--peptides were functionalized to selectively target bacterial membranes, they were self-assembled in the bacterial membranes and exert antibacterial activity by increasing the membrane permeability [73]. From the library of antibacterial peptides, a series of six- and eight-residue amphiphilic cyclic d,l-peptides were designed to bear at least one basic residue to enhance their target specificity toward negatively charged

454 bacterial membranes [74, 75]. In order to determine the efficacy of the peptide nanotubes with respect to in vivo protection of mice from bacterial infection, groups of mice were infected with lethal doses of methicillin-resistant S. aureus (MRSA) and then each group was treated intraperitoneally with a given bolus dose of the peptide nanotubes at 45–60 min after initial infection [73]. In this study, the amino acid sequences of cyclic peptides to target MRSA were KKKWLWLW, RRKWLWLW, and KKLWLW, where bold letters indicate d-amino-acid residues. After seven days, single appropriate doses of the peptide nanotubes were sufficient to completely protect various groups of mice from MRSA infections. This result will be very useful in developing new therapeutic treatments of existing and emerging infectious diseases by designing peptides rationally via combinatorial peptide libraries.

5. SUMMARY The field of device fabrication based on peptide nanotubes is still very young. At this stage, material development and fundamental mechanistic study of self-assemblies in peptide nanotubes have been very active and there is some progress in organizing peptide nanotubes in device configurations. These two areas are necessary to be coupled to develop real-world devices from peptide nanotubes, which have just started appearing in literature very recently. The biocompatibility of peptide nanotubes makes them valuable to build biotechnology-related devices such as biosensors and biomolecular filters, and their potential to position them into device configurations via biological recognitions may help build improved electronics. Physical properties of peptide nanotubes may not be suitable for certain applications; however, functionalization of peptide nanotubes will give them flexibility in physical properties to overcome it. As explained, functionalization can be achieved via peptide monomer modification or coating on the peptide nanotubes with functional groups.

GLOSSARY Amphiphile A molecule with a functional group as a head and alkyl chain as a tail. Bolaamphiphile A molecule with two functional groups as heads connected by an alkyl chain. Functional group Systematic chemical structures with specific chemical/physical properties. Functionalization Addition of specific chemical/physical properties onto self-assembled structures. Intermolecular hydrogen bond Hydrogen bonding among multiple molecules. Intramolecular hydrogen bond Hydrogen bonding within single molecule. Peptide nanotube A nanometer-sized tubular structure self-assembled from monomers containing amino acids or amino acid-like molecules linked with peptide bonds. Self-assembly Spontaneous organization of molecules into stable, larger-scaled structures.

Peptide Nanotubes

ACKNOWLEDGMENTS I thank my co-workers for the past four years for their hard work in the development of peptide nanotubes and their applications: B. Gologan, S. Pan, G. E. Douberly, S. Jonas, E. Goun, P. Porrata, R. MacCuspie, Dr. Catherine Thaler, and Dr. B. P. Tonner. I also acknowledge the financial support from the National Science Foundation CAREER Award (EIA-0133493), the National Science Foundation NER program (ECS-0103430), the U.S. Department of Energy (DE-FG-02-01ER45935), and Florida Hospital, Gala Endowed Program for Oncologic Research.

REFERENCES 1. Y. Cui, Q. Q. Wei, H. K. Park, and C. M. Lieber, Science 293, 1289 (2001). 2. Y. Huang, X. Duan, Y. Cui, L. J. Lauhon, K. H. Kim, and C. M. Lieber, Science 294, 1313 (2001). 3. M. R. Diehl, S. N. Yaliraki, R. A. Beckman, M. Barahona, and J. R. Heath, Angew. Chem. Int. Ed. Engl. 41, 353 (2001). 4. P. G. Collins, M. S. Arnold, and P. Avouris, Science 292, 706 (2001). 5. A. Bachtold, P. Hadley, T. Nakanishi, and C. Dekker, Science 294, 1317 (2001). 6. E. Braun, Y. Eichen, U. Sivan, and G. Ben-Yoseph, Nature 391, 775 (1998). 7. O. Harnack, W. E. Ford, A. Yasuda, and J. M. Wessels, Nano Lett. 2, 919 (2002). 8. H. Matsui, P. Porrata, and G. E. J. Douberly, Nano Lett. 1, 461 (2001). 9. J. K. N. Mbindyo, B. D. Reiss, E. R. Martin, C. D. Keating, M. J. Natan, and T. E. Mallouk, Adv. Mater. 13, 249 (2001). 10. E. Dujardin and S. Mann, Adv. Mater. 14, 775 (2002). 11. E. Dujardin, L.-B. Hsin, C. R. C. Wang, and S. Mann, Chem. Commun. 1264 (2001). 12. In this chapter, the word “monomer” was used for a peptide molecule from which peptide nanotubes are self-assembled. 13. G. M. Whitesides, E. E. Simanek, J. P. Mathias, C. T. Seto, D. N. Chin, M. Mammen, and D. M. Gordon, Acc. Chem. Res. 28, 37 (1995). 14. M. Kogiso, S. Ohnishi, K. Yase, M. Masuda, and T. Shimizu, Langmuir 14, 4978 (1998). 15. S. Zhang, D. M. Marini, W. Hwang, and S. Santoso, Curr. Opin. Chem. Biol. 6, 865 (2002). 16. J. D. Hartgerink, J. R. Granja, R. A. Milligan, and M. R. Ghadiri, J. Am. Chem. Soc. 118, 43 (1996). 17. J.-H. Fuhrhop and J. Koning, “Membranes and Molecular Assemblies: The Synkinetic Approach.” Royal Society of Chemistry, Cambridge UK, 1994. 18. N. Nakashima, S. Asakuma, and T. Kunitake, J. Am. Chem. Soc. 107, 509 (1985). 19. J.-H. Fuhrhop and W. Helfrich, Chem. Rev. 93, 1565 (1993). 20. T. Shimizu and M. Hato, Biochim. Biophys. Acta 1147, 50 (1993). 21. P. L. Privalov and S. J. Gills, Pure Appl. Chem. 61, 1097 (1989). 22. J. D. Hartgerink, E. Beniash, and S. I. Stupp, Science 294, 1684 (2001). 23. J. D. Hartgerink, E. Beniash, and S. I. Stupp, Proc. Natl. Acad. Sci. USA 99, 5133 (2002). 24. H. Matsui and B. Gologan, J. Phys. Chem. B. 104, 3383 (2000). 25. T. Shimizu, M. Kogiso, and M. Masuda, J. Am. Chem. Soc. 119, 6209 (1997). 26. M. Kogiso, M. Masuda, and T. Shimizu, Supramol. Chem. 9, 183 (1998). 27. B. Gologan and H. Matsui, unpublished results. 28. G. J. Douberly, S. Pan, D. Walters, and H. Matsui, J. Phys. Chem. B. 105, 7612 (2001).

Peptide Nanotubes 29. H. Matsui and R. MacCuspie, Nano Lett. 1, 671 (2001). 30. H. Matsui, S. Pan, B. Gologan, and S. Jonas, J. Phys. Chem. B. 104, 9576 (2000). 31. R. B. Prince, J. S. Barnes, and J. S. Moore, J. Am. Chem. Soc. 122, 2758 (2000). 32. J. C. Nelson, J. K. Young, and J. S. Moore, J. Org. Chem. 61, 8160 (1996). 33. J. S. Moore, Acc. Chem. Res. 30, 402 (1997). 34. S. Vauthey, S. Santoso, H. Gong, N. Watson, and S. Zhang, Proc. Natl. Acad. Sci. USA 99, 5335 (2002). 35. M. Kogiso, Y. Okada, T. Hanada, K. Yase, and T. Shimizu, Biochim. Biophys. Acta 1475, 346 (2000). 36. T. Shimizu, R. Iwaura, M. Masuda, T. Hanada, and K. Yase, J. Am. Chem. Soc. 123, 5947 (2001). 37. G. M. Whitesides, Sci. Am. 273, 146 (1995). 38. H. Matsui and C. Holtman, Nano Lett. 2, 887 (2002). 39. D. T. Bong, T. D. Clark, J. R. Granja, and M. R. Ghadiri, Angew. Chem. Int. Ed. Engl. 40, 988 (2001). 40. D. Seebach, J. L. Matthews, A. Meden, T. Wessels, C. Naerlocher, and L. B. McCusker, Helv. Chim. Acta 80, 173 (1997). 41. D. Ranganathan, C. Lakshmi, and I. L. Karle, J. Am. Chem. Soc. 121, 6103 (1999). 42. M. R. Ghadiri, J. R. Granja, R. A. Milligan, D. E. McRee, and N. Khazanovich, Nature 366, 324 (1993). 43. M. E. Polaskova, N. J. Ede, and J. N. Lambert, Aust. J. Chem. 51, 535 (1998). 44. M. L. Bushey, A. Hwang, P. W. Stephens, and C. Nuckolls, J. Am. Chem. Soc. 123, 8157 (2001). 45. M. L. Bushey, A. Hwang, P. W. Stephens, and C. Nuckolls, Angew. Chem. Int. Ed. Engl. 41, 2828 (2002). 46. H. Fenniri, B.-L. Deng, A. E. Ribbe, K. Hallenga, J. Jacob, and P. Thiyagarajan, Proc. Natl. Acad. Sci. USA 99, 6487 (2002). 47. B. Dietrich, P. Viout, and J.-M. Lehn, “Macrocyclic Chemistry: Aspects of Organic and Inorganic Supramolecular Chemistry.” VCH, Weinheim, 1993. 48. H. Fenniri, P. Mathivanan, K. L. Vidale, D. M. Sherman, K. Hallenga, K. V. Wood, and J. G. Stowell, J. Am. Chem. Soc. 123, 3854 (2001). 49. H. Fenniri, B.-L. Deng, and A. E. Ribbe, J. Am. Chem. Soc. 124, 11064 (2002). 50. C. H. Gorbits, Chem. Eur. J. 7, 5153 (2001). 51. A. M. Levelut, J. Chim. Phys. Phys.-Chim. Biol. 80, 149 (1983). 52. J. Sanchez-Quesada, H. S. Kim, and M. R. Ghadiri, Angew. Chem. Int. Ed. Engl. 40, 2503 (2001).

455 53. X. L. Zhao, Y. Ando, L. C. Qin, H. Kataura, Y. Maniwa, and R. Saito, Appl. Phys. Lett. 81 (2002). 54. H. Matsui and G. E. J. Douberly, Langmuir 17, 7918 (2001). 55. T. D. Clark, K. Kobayashi, and M. R. Ghadiri, Chem. Eur. J. 5, 782 (1999). 56. W. Traub and H. D. Weiner, Proc. Natl. Acad. Sci. USA 86, 9822 (1989). 57. H. D. Weiner and L. Addadi, J. Mater. Chem. 7, 689 (1997). 58. T. Rueckes, K. Kim, E. Joselevich, G. Y. Tseng, C.-L. Cheung, and C. M. Lieber, Science 289, 94 (2000). 59. A. Javey, Q. Wang, A. Ural, Y. M. Li, and H. J. Dai, Nano Lett. 2, 929 (2002). 60. H. Matsui, B. Gologan, S. Pan, and G. J. Douberly, Eur. Phys. J. D. 16, 403 (2001). 61. R. Djalali, Y.-F. Chen, and H. Matsui, J. Am. Chem. Soc. 124, 13660 (2002). 62. H. Matsui, S. Pan, and G. E. J. Douberly, J. Phys. Chem. B. 105, 1683 (2001). 63. P. Porrata, E. Goun, and H. Matsui, Chem. Mater. 14, 4378 (2002). 64. H. Matsui, P. Porrata, E. Goun, M. Kilmov, and B. P. Tonner, unpublished results. 65. J. L. Sample, K. C. Beverly, P. R. Chaudhari, F. Remacle, J. R. Heath, and R. D. Levine, Adv. Mater. 14, 124 (2002). 66. K. Fukusaki, K. Takeda, and K. Shiraishi, J. Phys. Soc. Jpn. 66, 3387 (1997). 67. H. Okamoto, M. Kasahara, K. Takeda, and K. Shiraishi, Pept. Sci. 36, 67 (1999). 68. J. P. Lewis, N. H. Pawley, and O. F. Snakey, J. Phys. Chem. B. 101, 10576 (1997). 69. J. Sanchez-Quesada, M. P. Isler, and M. R. Ghadiri, J. Am. Chem. Soc. 124, 10004 (2002). 70. B. Hille, “Ionic Channels of Excitable Membrane.” Sinauer, Sunderland, MA, 1992. 71. O. Braha, B. Walker, S. Cheley, J. J. Kasianowicz, L. Song, J. E. Gouaux, and H. Bayley, Chem. Biol. 4, 497 (1997). 72. L.-Q. Gu, O. Braha, S. Conlan, S. Cheley, and H. Bayley, Nature 398, 686 (1999). 73. S. Fernandez-Lopez, H. S. Kim, E. C. Choi, M. Delgado, J. R. Granja, A. Khasanov, K. Kraehenbuehl, G. Long, D. A. Weinberger, K. Wilcoxen, and M. R. Ghadiri, Nature, 412, 452 (2001). 74. D. M. Pitterle, J. L. Johnson, and K. V. Rajagopalan, J. Biol. Chem. 268, 13506 (1993). 75. J. E. Walker, M. Saraste, M. J. Runswick, and N. J. Gay, EMBO J. 1, 945 (1982).