Nanostructured Materials Designed for Cell Binding and Transduction

Apr 19, 2001 - Washington University, Department of Chemistry, One Brookings Drive, CB1134, St. Louis, Missouri 63130-4899 ... Ziqing Qian , Agnieszka...
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Communications Nanostructured Materials Designed for Cell Binding and Transduction Jianquan Liu, Qi Zhang, Edward E. Remsen, and Karen L. Wooley* Washington University, Department of Chemistry, One Brookings Drive, CB1134, St. Louis, Missouri 63130-4899 Received January 27, 2001; Revised Manuscript Received March 27, 2001

The surface-functionalization of shell cross-linked (SCK) nanoparticles with the oligomeric peptide sequence YGRKKRRQRRR, the protein transduction domain (PTD) from the human immunodeficiency virus TAT protein, is described, and the cell binding interactions these nanobioconjugates exhibit are demonstrated. A convergent synthetic strategy was employed, whereby the SCK nanoparticles and the PTD were prepared independently and then coupled together during immobilization of the PTD component on a solid support. The SCK nanoparticles were prepared by the micellization of amphiphilic block copolymers of poly(caprolactone-b-acrylic acid), followed by amidation-based cross-linking of the acrylic acid residues located within the micellar corona. The PTD sequence was constructed upon a solid support, from C-terminus to N-terminus, followed by extension with four glycine residues, leaving the amino chain end for subsequent coupling with remaining acrylic acid functionalities present on the surface of the SCK. Finally, cleavage from the solid support was performed, which also facilitated deprotection of the peptide side chain functionalities as well as hydrolysis of the poly(-caprolactone) segments composing the SCK core domain, to yield PTD-derivatized nanocage structures (PTD-nanocage). Covalent labeling of the SCK precursor with fluorescein-5-thiosemicarbazide provided fluorescently tagged PTD-nanocage nanobioconjugates to allow for their detection by fluorescence microscopy. The fluorescent PTD-nanocage bioconjugates were found to interact with CHO cells and HeLa cells, whereas the analogous structure lacking the PTD component did not. CHO cells bound with fluorescent PTD-nanocage bioconjugates were analyzed using flow cytometry and fluorescence activated cell sorting (FACS). Fluorescence confocal microscopy of isolated bioconjugatebound CHO cells indicated that the bioconjugated nanoparticles were primarily located near the cell periphery; however, transduction of the nanoparticle into the cells also occurred. Introduction. As enhanced capabilities are gained for the preparation of well-defined macromolecules, increased attention is being given to ever more complex molecular topologies and morphologies, with guidance often being sought from biological species.1 Given that much of biology is based upon hierarchy, with nanoscale structures playing key roles, synthetic nanostructured materials are being aggressively pursued. The commonality in sizes and general structural features that these materials share with their biological counterparts are then expected to translate into similarities in function. We have focused our efforts toward the preparation of amphiphilic core-shell nanostructures, shell cross-linked nanoparticles (SCKs),1e which are capable of hydrophobic guest sequestration and transport,2 in analogy with lipoproteins. SCKs bearing positively charged surface functional * Corresponding author. Telephone: (314) 935-7136. Fax: (314) 9359844. E-mail: [email protected].

groups have also been shown to package DNA, while maintaining structural integrity,3 in analogy with histone core proteins. These properties suggest that SCKs may be suitable nanoscale containers for the transport and delivery of biologically active agents. However, preliminary studies have shown quite poor gene transfection efficiencies.4 Clearly, in addition to the structure and dimensions of the particular synthetic materials, one must also consider the surface chemistry, as interfacial interactions dominate the fate of materials when placed into any biological environment. The combination of natural and synthetic components into a hybrid entity is the fundamental principle of bioconjugation. Bioconjugation is widely employed in basic and applied research; and is an essential biotechnique utilized by life science companies to produce diagnostic and therapeutic products.5 Polymeric materials functionalized with sugars,6 peptides,7 folate,8 antibodies,9 and other moieties have received considerable interest as a means to generate structures capable of polyvalent, specific binding interactions10

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and the development of intelligent delivery vehicles.11 The ability to readily control particle diameter (10-100 nm) and morphology to establish a polymeric nanocontainer (of similar dimensions as biologically functional transport nanomaterials, e.g., lipoproteins and capsid virus particles) is an important advantage of SCKs relative to linear polymers or dendrimers. Once a particular device reaches the target of interest, the next challenge is to cross the cell membrane. Several peptide sequences, which are biologically active transduction domains, derived from a number of proteins,12 offer the ability to permeate cell membranes. In this report, we investigate the conjugation between our SCK nanoparticles and a biologically active peptide sequence, the protein transduction domain (PTD) derived from HIV. The SCKs are characterized by an intricate amphiphilic, core-shell morphology with overall diameters ranging typically from 10 to 100 nm.13 In this example, the hydrophobic core domain is composed of a hydrolytically degradable material,14 which is extracted to produce ultimately, a nanocage structure15 bearing the PTD sequence. PTD has been used to deliver biologically active proteins of various sizes into mice16 and to transduce 40 nm superparamagnetic (Fe) nanoparticles into mammalian T cells.17 More recently, PTD has been used to translocate paramagnetic MR contrasting agents into HeLa cells.18 Our interests are directed toward the delivery of organic-based synthetic nanomaterials of complex morphologies (to generate various compartments) having hydrogellike surfaces modified with peptide chains to effect cell binding and transduction. Therefore, the construction of a bioconjugate consisting of a SCK nanoparticle bearing a surface-attached protein transduction domain peptide sequence is described, and its binding interactions with and transduction into cells is demonstrated. Experimental Methods. All synthetic reagents were purchased from Aldrich, Sigma, or Fluka unless stated otherwise. Amino acid building blocks for solid-phase peptide synthesis were purchased from NovaBiochem-CalBiochem Corp. Cell culture medium was purchased from Sigma. Chinese hamster ovary (CHO) cells were obtained from Clontech Laboratories, Inc., and HeLa cells were the kind gift of Dr. France Daigle from the laboratory of Professor Roy Curtiss, Department of Biology, Washington University. Ultraviolet spectra were obtained with a Cary 1E UVvis spectrophotometer (Varian). Fluorescence spectra were collected with a FluoroMax Spectrofluorometer (Spex Industries, Inc.). Mass spectra were obtained by Voyager DERP MALDI-TOF mass spectrometer (PE Biosystem). Green fluorescent images were obtained with an Olympus IX70 inverted fluorescent microscope (plain fluorite 10× objective lens; U-MWB cube; excitation filter, BP450-480; dichroic mirror, DM500; and barrier filter, BA515) with a mercury lamp (BH2-RFL-T3) and a built-in 35 mm SLR camera, which was part of a BioScope atomic force microscope (Digital Instruments). Flow cytometry was carried out on FACS Calibur (Benton Dickinson) while fluorescence activated cell sorting (FACS) was accomplished using a FACS Vantage (Benton Dickinson).

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Samples for imaging by atomic force microscopy (AFM) were prepared by placing a 1.0 µL drop of aqueous SCK solution (typical concentration about 0.05 mg/mL) on freshly cleaved mica (Ruby clear mica, New York Mica Co.) and allowing it to dry in air and then further drying the sample in vacuo for 1-2 h. AFM imaging was performed using a BioScope system with a Nanoscope IIIa controller (Digital Instruments) equipped with a J-type vertical engage piezoelectric scanner and operated in tapping mode in air. Tapping mode imaging was performed with the cantilever being oscillated above the surface at a frequency far from its resonance frequency and coming into contact with the surface intermittently, at the peak of each oscillation cycle. The decrease in cantilever oscillation amplitude due to intermittent contacts is used as a control signal for a feedback loop to track the surface topography. Throughout this study, tapping mode imaging was carried out with standard etched silicon probes (l ) 128 µm, spring constant ∼50 N/m, resonance frequency ∼330 kHz) and cantilever oscillation amplitudes of about 10 nm. The typical value of set point (the ratio of damped to free oscillation amplitude) was 0.95. The typical raw signal corresponding to cantilever oscillation was ca. 1.0 V. Typical scan frequencies were between 0.5 and 2.0 Hz, depending on scan fields, which varied from 10 µm × 10 µm to 500 nm × 500 nm. The SCK diameters were taken as the heights of the SCK nanoparticles, which were measured by bearing analysis procedure, using the Nanoscope III software package. Distributions of SCK nanoparticle heights were obtained from individual measurements performed on ∼ 100 particles. The average diameters (Dav) were determined from measurements performed on ∼50 particles without deconvolution of the effects for the silicon tip diameter (ca. 15-30 nm). Values of hydrodynamic diameter distribution were obtained for SCK nanoparticles by dynamic light scattering (DLS) using a Brookhaven Instrument Co. Model 90 Plus dynamic light scattering system. The wavelength of incident light employed was 678 nm with a nominal power of 20 mW. The instrument’s built-in thermoelectric heater was used to equilibrate the temperature of solutions at 20 ( 1 °C. Prior to analysis, solutions were spun at 12 000 rpm for 4 min or filtered through a cotton plug to remove dust particles. Light scattering cells were rinsed with deionized water filtered through a 0.2 µm poly(vinylidene difluoride) membrane filter before being filled with dust-free solutions. Scattered light was collected at a fixed angle of 90°. The digital correlator was operated with 200 channels, a duration time of 3 min, ratio channel spacing, and initial and final delay times of 5 µs and 10 ms, respectively. All measurements were made in triplicate. The calculation of hydrodynamic diameter distribution was provided by built-in particle size distribution analysis software which employed the method of nonnegatively constrained least-squares (NNLS)19 to compute diameter distribution averages. For confocal microscopy,20 preparations were wholemounted on standard slides and imaged with an upright microscope equipped with a Zeiss 63 × 1.4 numerical aperture (NA) planapochromat objective and a Bio-Rad MRC 1024 confocal adapter (Hercules, CA). A krypton-argon

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Figure 1. Solid-phase synthesis strategy involved coupling of the protein transduction domain (PTD) with fluorescein-labeled shell cross-linked (SCK) nanoparticles, followed by cleavage from the support and excavation of the nanoparticle core domain to yield FTSC-labeled PTDnanocage bioconjugates. The SCK derived from PCL105-b-PAA165 was synthesized according to a previous report:14 (1) (a) 1-(3dimethylaminopropyl)-3-ethylcarbodiimide methiodide; (b) 2,2′-(ethylenedioxy)bis(ethylamine). (2) (a) 1-(3-dimethylaminopropyl)-3-ethylcarbodiimide methiodide, H2O, 15 min; (b) fluorescein-5-thiosemicarbazide (FTSC), DMF, overnight. (3) N,N-diisopropylcarbodiimide (DIPCDI), Nhydroxybenzotriazole (HOBt), then resin loaded with side chain-protected and N-terminus free protein transduction domain, overnight. (4) Excess 95% TFA:5% H2O:2.5% TIS (triisopropylsilane), 30 h.

laser was used; epifluorescence filter sets were those designed for Texas Red (SR101 and Cy3) and fluorescein (FITC-V VA and FM1-43). The confocal aperture was set to its smallest (diffraction-limited) diameter; collection amplifier gains and offsets were adjusted for optimum visualization of the fluorescence signal and remained unchanged for each series of experiments unless otherwise stated. A stepper motor attached to the microscope’s focusing knob allowed sequential imaging in different focal planes for threedimensional re-constructions. Usually 15-40 images, at planes separated by 0.25-0.5 µm, were obtained; this increment was approximately half of the instrument’s z-axis resolution and was chosen to provide some signal averaging (smoothing) of depth information via oversampling. Stacks comprising 15-40 one- or two-color images (each 512 × 512 pixels; 8 bits of gray scale per color) were stored on magnetic disk of a computer for subsequent analysis. Synthesis of a Fluorescein-5-thiosemicarbazide (FTSC) Labeled Shell Cross-Linked Nanoparticle (FTSC-Labeled SCK). SCK nanoparticles were prepared from PCL105-bPAA165 as previously described.14 To a solution of the SCKs (0.3 mg/mL) (40 mL, 0.05 mmol of carboxylic acid functional groups), 1-(3-dimethylaminopropyl)-3-ethylcarbodiimide methiodide (18 mg/mL) (0.50 mL, 0.030 mmol) in water was added dropwise. The mixture was allowed to stir for 15 min before FTSC (4.6 mg/mL) (1.0 mL, 0.011 mmol) in DMF was added dropwise. The reaction mixture was allowed to stir overnight and the precipitate that formed was removed by filtration, followed by

purification of the FTSC-labeled SCK by dialysis of the solution against water (5 × 2 L) for 3 d. The solution was then dialyzed against DMF (3 × 1 L) for another 3 d to exchange the aqueous solution to a DMF solution. UV: λmax ) 488 nm (H2O) and 513 nm (DMF). Fluorescence emission: 515 nm (H2O, when excited at 488 nm) and 541 nm (DMF, when excited at 513 nm). The dimensions of the FTSC-labeled SCK nanoparticles were determined with AFM measurement upon a mica substrate to be Hav ) 10 ( 2 nm and Dav ) 55 ( 19 nm. DLS measurement in aqueous solution gave the number distribution percentage (Ni, %) and volume distribution percentage (Vi, %) values for each component in DLS hydrodynamic diameter (Dh) measurement: Dh,i (Ni, %) ) 13 (99+%), Dh,i (Vi, %) ) 13 (68%) and 329 (32%) nm. Synthesis of FTSC-Labeled PTD-Nanocage. GGGGYGRKKRRQRRR16 was synthesized by solid-phase peptide synthesis using standard Fmoc chemistry. MALDI-TOF: [M + 1] ) 1891.6 (calculated [M + 1] ) 1891). The FTSClabeled SCK in DMF was coupled to the peptide sequence, while it remained attached to the solid support beads, using an excess of DIPCDI/HOBt at room temperature overnight. The beads were then filtered and washed. Treatment with 95% TFA:2.5% H2O:2.5% TIS for 30 h cleaved the PTDSCK conjugates from the support, deprotected the peptide, and degraded the SCK core. The resulting FTSC-labeled PTD-nanocage was collected by ether precipitation and purified by dialysis against water for several days. UV

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Figure 2. Atomic force microscope (AFM) images of functionalized shell cross-linked nanoparticles (FTSC-SCK) (a) and FTSC-labeled PTD-nanocage bioconjugates (b). The inset shows the extensive flattening that results for the nanostructures after the removal of the SCK core domain, upon formation of the nanocage material during cleavage of the bioconjugate from the solid support.

(H2O): 440 nm. Fluorescence (H2O): λem ) 513 nm (when excited at 488 nm). Cell Culture, Flow Cytometry, Fluorescence-Activated Cell Sorting (FACS) and Fluorescent Confocal Microscopy: CHO or HeLa cells in a six-well plate were grown in MEM (R-modification) supplemented with G 418 (2 mL) at 37 °C in a 5% CO2 humidified incubator until 50-80% confluence. The medium was removed and replaced with fresh medium (1 mL). FTSC-labeled PTD-nanocage in water (0.3 mg/mL) (1 mL) was added and mixed well with the medium, and the incubation was continued for 30-120 min. The medium was then removed and the cells were washed twice with fresh medium, followed by treatment with trypsin at room temperature for 5-10 min. The cells were spun at 1000 rpm for 10 min. The supernatant was decanted and the cell pellet was resuspended in PBS solution (1 mL) for counting with flow cytometry or sorted by FACS (488 nm). The sorted fluorescent cells (10 µL) were deposited on a standard glass microscope slide, covered with a coverslip, and then subjected to fluorescence confocal microscopy analysis according to the literature procedure.20 Results and Discussion A convergent synthetic approach was employed to construct the PTD-nanocage bioconjugate (Figure 1). SCK

nanoparticles of amphiphilic poly(-caprolactone)-b-poly(acrylic acid-co-acrylamide) (PCL-b-PAA-co-PA) were first prepared by the micellar assembly of poly(-caprolactone)105b-poly(acrylic acid)165 (PCL-b-PAA), followed by covalent cross-linking reactions with 2,2′-(ethylenedioxy)bis(ethylamine) in the presence of 1-(3-dimethylaminopropyl)-3ethylcarbodiimide methiodide.14 Approximately 50% of the acrylic acid groups were consumed in this cross-linking procedure.21 Functionalization of the SCKs with fluorescein5-thiosemicarbazide (FTSC) was accomplished by allowing many of the remaining available -CO2H groups of the SCK to undergo coupling with FTSC, in the presence of 1-(3dimethylaminopropyl)-3-ethylcarbodiimide methiodide. The FTSC derivatization rendered the nanoparticles fluorescent, however, the exact stoichiometry of the FTSC-SCK coupling was not determined. The arginine-rich sequence, YGRKKRRQRRR, of HIV-tat peptide16 was prepared by standard solid phase peptide synthesis (SPPS). The N-terminus was then extended with a four glycine-residue linker to provide a flexible spacer between the PTD and SCK. After the last SPPS coupling, the Fmoc protecting group was removed to free the terminal amine group, which was subsequently coupled with remaining carboxyl groups on the surface of FTSC-labeled SCK. Because both the SCK nanoparticle and the peptide resin are particulate, only interfacial contact between them was expected, allowing for SCK surface attachment and a limited number of couplings per SCK. Moreover, by performing the SCK-to-PTD coupling on the solid support, any nonconjugated nanoparticles were removed by washing of the resin support. After treatment with 95% TFA to afford peptide deprotection and cleavage from the resin, the product was purified by ether precipitation. The resulting yellow precipitate was dialyzed to remove impurities, including unconjugated PTD. Since the deprotection and cleavage of the solid-phase synthesis required a 95%TFA treatment, hydrolysis of the nanoparticle’s PCL core also occurred, to produce a nanometer-scale cagelike structure.14 The AFM images of Figure 2 illustrate the change in the nanoparticle behavior, from core-shell particles having heights and diameters of Hav ) 10 ( 2 nm and Dav ) 55 ( 19 nm, respectively, to a nanocage structure from solutions that spread and collapsed upon adsorption on the mica substrate. The diffuse edges and aggregation events (perhaps caused by electrostatic interactions between the PTD and the nanocage carboxylatecontaining shell) made determination of dimensions inaccurate, however, as can be seen from the inset of Figure 2b, the nanocages give heights less than 5 nm. The recovered FTSC-labeled PTD-nanocage bioconjugate showed the pHdependent spectral characteristics of fluorescein and was characterized by UV-vis spectroscopy, fluorescence spectroscopy and fluorescence microscopy. The PTD-mediated binding of cells by FTSC-labeled PTD-nanocage bioconjugates, following incubation with Chinese hamster ovary (CHO) cells, was evaluated by fluorescence microscopy. Experimentally, 5 × 105 CHO cells were seeded in a six-well plate with MEM (R-modification) supplemented with G 418 and were incubated at 37 °C in a 5% CO2 humidified incubator until 50-80% confluence. The

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Figure 3. Optical microscopy images illustrate the binding of FTSC-labeled PTD-nanocages to CHO cells: (a) image collected under fluorescence using excitation filter BP450-480 for CHO cells following incubation with FTSC-labeled PTD-nanocages and washing; (b) fluorescence image collected for the sample of (a) after treatment with trypsin, removal of cell medium, and washings; (c) fluorescence image of CHO cells incubated with FTSC-labeled PTD-nanocages with medium still present; (d) image showing the sample from part c imaged under blue and visible wavelengths. The lack of interactions between the CHO cells and FTSC-labeled nanocages without the PTD segment is shown in part e for CHO cells imaged under blue and visible wavelengths following incubation with the FTSC-labeled nanocages and washing.

medium was replaced with fresh medium and an equal volume of the aqueous solution of FTSC-labeled PTDnanocage bioconjugate was added and mixed well. The incubation at 37 °C in a 5% CO2 humidified incubator was continued for 30-120 min. The supernatant was then removed and the cells were washed twice with fresh medium to ensure the removal of dead cells and unbound bioconjugates. After less than 60 min of incubation, cells with green fluorescence were observed under the blue light of a fluorescence microscope (Figure 3a). As shown in Figure

3b, trypsin treatment of those plates removed the cells, and along with the cells, the fluorescent PTD-nanocages were also eliminated. Comparison of parts c and d of Figure 3 demonstrate the concentration of the fluorescent PTDnanocages on the cells. These observations indicated the binding of PTD-nanocages to CHO cells. Similar results were observed for the same experiments repeated with HeLa cells. The requirement for the PTD to afford cell binding was established in a control experiment in which the FTSClabeled nanocage, without PTD, was tested under the same

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Figure 4. Fluorescent confocal microscopy images of CHO cells transduced by FTSC-labeled PTD-nanocage bioconjugates. The CHO cells incubated with the bioconjugate for 30-60 min were treated with trypsin and sorted by fluorescence activated cell sorter (FACS). The fluorescent cells were then applied on a microscope slide and imaged under the fluorescence confocal microscope with krypton-argon laser excitation at 488 nm. Twenty-five images, at planes separated by 300 nm, were obtained from top (upper left) to bottom (lower right).

conditions. No evidence of binding between these nanocages and cells was found (see, e.g., Figure 3e). In another set of experiments, the trypsin-treated CHO cells were recovered and analyzed by flow cytometry to quantify the PTD-nanocage-to-cell interaction. Measurements made with CHO cells alone showed no fluorescence above background. After treatment with FTSC-labeled PTDnanocage, the intensity of fluorescence associated with the cells increased more than 100 times relative to background. Up to 14% of the cells were found to fluoresce, as counted by flow cytometry. These data provide further support of PTD-mediated binding of these nanostructures to CHO cells. To locate the nanocages with respect to the cell, the CHO

cells were sorted by fluorescence activated cell sorting (FACS). The fluorescent cells were subsequently analyzed with fluorescence confocal microscopy, which provided optical images of “slices” through a cell. This analysis showed that the majority of FTSC-labeled PTD-nanocages were concentrated along the surface of the cell, but fluorescence was also observed inside the cell (Figure 4), indicating transduction of the nanostructures into cells. Quantitative interpretation of the number of conjugates transduced into the cell based on the relative intensity differences between the surface-bound and internalized fluorophores was not attempted, because of complications with refractive index

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differences, heterogeneity in the labeling, and potential differences in fluorescence quenching effects. Conclusions Polymeric therapeutics and polymeric micelles have shown enhanced permeability and retention effects, characterized by reduced renal excretion, longer blood circulation times, lower toxicities, and higher efficacy.11a,22 As extensions are made from single polymer chains to nanometer scale structures, it is expected that multifaceted scaffolds can be engineered to allow for multiple functions, such as the incorporation of compartments with high capacity guest loading in delivery or sequestration applications, in concert with other regions that are to be presented to intercept and regulate or utilize the biological recognition and signaling machinery. We have demonstrated the conjugation of an important transduction sequence capable of permeating cell membranes to a well-defined nanocage structure. In this scenario, the nanocage is the compartment for delivery or retrieval of bioactives, while the peptide sequence is designed to facilitate cell transduction. The ability to package therapeutic agents, ranging from small molecules to genes, within a nanoscale containment device could potentially provide site- and time-selective drug release in carriers capable of biomimicry and stealth behavior. Further studies are in progress to quantify the thermodynamics and kinetics of PTD-nanocage to cell binding. Much work remains in order to determine the exact fate of the PTD-nanocage structures during in vitro cell culture and, importantly, when subjected to in vivo experiments. Because the mechanism of membrane transduction, although not well understood, appears to occur for a number of protein basic domains without receptor mediation,23 the covalent attachment of receptors to the nanostructures may be required for target-specific delivery. The nanoscale scaffold of the SCKs and nanocages offers a large surface area to accommodate the derivatization with a number of functional groups to produce polyvalent and multivalent interactions. Realization of these materials in biomedical applications will require substantial research efforts at the interface of chemistry, biology, materials science and engineering. Acknowledgment. We thank Professor J. -S. Taylor for his many valuable suggestions. We are also grateful to Dr. F. Daigle and Dr. C. Dozois from Professor R. Curtiss’ laboratory for their generous help with tissue culture experiments; and to Dr. K. B. Thurmond for the use of the dynamic light scattering instrument in his laboratory. The assistance provided by Mr. C. G. Clark, Jr., in the processing of confocal microscopy images and for preparation of 3-D scientific illustrations is appreciated. We also thank Ms. P. Lan, from Professor S. Gilbertson’s laboratory, for assistance with peptide synthesis. The funding support of the National Science Foundation (DMR-9974457) and Monsanto Company is gratefully acknowledged. References and Notes (1) (a) Percec, V.; Holerca, M. N. Biomacromolecules 2000, 1, 6. (b) Piotti, M. E.; Rivera, F., Jr.; Bond, R.; Hawker, C. J.; Fre´chet, J. M. J. J. Am. Chem. Soc. 1999, 121, 9471. (c) Stupp, S. I.; Pralle, M. U.;

(2) (3) (4) (5)

(6)

(7) (8) (9) (10) (11)

(12)

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(14) (15) (16) (17) (18) (19) (20) (21) (22)

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Tew, G. N.; Li, L.; Sayar, M.; Zubarev, E. R. MRS Bull. 2000, 25(4), 42. (d) Cha, J. N.; Shimizu, K.; Zhou, Y.; Christiansen, S. C.; Chmelka, B. F.; Deming, T. J.; Stucky, G. D.; Morse, D. E. Mineralization in Natural and Synthetic Biomaterials. Mater. Res. Soc. Symp. Proc. 2000, 599, 239. (e) Wooley, K. L. J. Polym. Sci., Part A: Polym. Chem. 2000, 38, 1397. Baugher, A. H.; Goetz, J. M.; McDowell, L. M.; Huang, H.; Wooley, K. L.; Schaefer, J. Biophys. J. 1998, 75, 2574. Thurmond, K. B., II; Remsen, E. E.; Kowalewski, T.; Wooley, K. L. Nucleic Acids Res. 1999, 27, 2966. Studelska, D. R.; Thurmond, K. B., II; Liu, J.; Schaefer, J.; Wooley, K. L. Unpublished results. (a) Hermanson, G. T., Ed. Bioconjugate Techniques; Academic Press: San Diego, CA, 1995. (b) Veronese, F. M.; Morpurgo, M. Farmaco 1999, 54, 497. (a) Seymour, L. W.; Duncan, R.; Kopeckova, P.; Kopecek, J. J. Bioact. Compat. Polym. 1987, 2, 97. (b) Kiessling, L. L.; Strong, L. E.; Gestwicki, J. E. Annu. Rep. Med. Chem. 2000, 35, 321. (c) Kiessling, L. L.; Pohl, N. L. Chem. Biol. 1996, 3(2), 71. (d) Reuter, J. D.; Myc, A.; Hayes, M. M.; Gan, Z.; Roy, R.; Qin, D.; Yin, R.; Piehler, L. T.; Esfand, R.; Tomalia, D. A.; Baker, J. R., Jr.; Bioconjugate Chem. 1999, 10, 271. Arap, W.; Pasqualini, R.; Ruoslahti, E. Science 1998, 279, 377. Kono, K.; Liu, M.; Fre´chet, J. M. J. Bioconjugate Chem. 1999, 10, 1115. Seymour, L. W.; Flanagan, P. A.; Al-Shamkhani, A.; Subr, V.; Ulbrich, K.; Cassidy, J.; Duncan, R. Sel.e Cancer Ther. 1991, 7, 59. Mammen, M.; Choi, S.-K.; Whitesides, G. M. Angew. Chem., Int. Ed. 1998, 37, 2754. (a) Langer, R. Nature 1998, 392, 5. (b) Duncan, R. ACS Symp. Ser. 2000, 752, 350. (c) Duncan, R. Pharm. Sci. Technol. Today 1999, 2, 441-449. (d) Kataoka, K. In Controlled Drug DeliVery: Challenges and Strategies; Park, K., Ed.; American Chemical Society: Washington, DC, 1997; p 629. (e) Scholes, P. D.; Coombes, A. G. A.; Davies, M. C.; Illum, L.; Davis, S. S. In Controlled Drug DeliVery: Challenges and Strategies, Park, K., Ed.; American Chemical Society: Washington, DC, 1997; p 629. (f) Ringsdorf, H.; Schmidt, B. Zh. Vses. Khim. O-Va. im. D. I. MendeleeVa 1987, 32, 487-501. Polyakov, V.; Sharma, V.; Dahlheimer, J. L.; Pica, C. M.; Luker, G. D.; Piwnica-Worms, D. Bioconjugate Chem. 2000, 11, 762 and references therein. (a) Thurmond, K. B., II; Kowalewski, T.; Wooley, K. L. J. Am. Chem. Soc. 1997, 119, 6656. (b) Huang, H.; Kowalewski, T.; Remsen, E. E.; Gertzmann, R.; Wooley, K. L. J. Am. Chem. Soc. 1997, 119, 11653. Zhang, Q.; Remsen, E. E.; Wooley, K. L. J. Am. Chem. Soc. 2000, 122, 3642. Huang, H.; Remsen, E. E.; Kowalewski, T.; Wooley, K. L. J. Am. Chem. Soc. 1999, 121, 3805. Schwarze, S. R.; Ho, A.; Vocero-Akbani, A.; Dowdy, S. F. Science 1999, 285, 1569. Lewin, M.; Carlesso, N.; Tung, C.; Tang, X.; Cory, D.; Scadden, D.; Weissleder, R. Nature Biotechnol. 2000, 18, 410. Bhorade, R.; Weissleder, R.; Nakakoshi, T.; Moore, A.; Tung, C.H. Bioconjugate Chem. 2000, 11, 301. Stock, R. S.; Ray, R. J. Polym. Sci., Polym. Phys. 1985, 23, 13931447. Teng, H.; Cole, J. C.; Roberts, R. L.; Wilkinson, R. S. J. Neurosci. 1999, 19, 4855-4866. Huang, H.; Wooley, K. L.; Schaefer, J. Macromolecules 2001, 34, 547. (a) Maeda, H.; Wu, J.; Sawa, T.; Matsumura, Y.; Hori, K. J. Controlled Release 2000, 65, 271. (b) Kataoka, K.; Kwon, G. S.; Yokoyama, M.; Okano, T.; Sakurai, Y. J. Controlled Release 1993, 24, 119. (a) Derossi, D.; Chassaing, G.; Prochiantz, A. Trends Cell Biol. 1998, 8, 84. (b) Wender, P. A.; Mitchell, D. J.; Pattabiraman, K.; Pelkey, E. T.; Steinman, L.; Rothbard, J. B. Proc. Natl. Acad. Sci. U.S.A. 2000, 97, 13003.

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