Aptamer-Functionalized In Situ Injectable Hydrogel for Controlled

Sep 1, 2010 - B.S. is in part supported by the Royal Thai Government Scholarship. This work is supported by the NSF Grant DMR-0955358 to Y.W...
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Aptamer-Functionalized In Situ Injectable Hydrogel for Controlled Protein Release Boonchoy Soontornworajit, Jing Zhou, Zhaoyang Zhang, and Yong Wang* Department of Chemical, Materials, and Biomolecular Engineering, University of Connecticut, Storrs, Connecticut 06269-3222 Received July 10, 2010; Revised Manuscript Received August 8, 2010

Various in situ injectable hydrogels have been developed for protein delivery in treating human diseases. However, most hydrogels are highly permeable, which can lead to the rapid release of loaded proteins. The purpose of this study is to apply nucleic acid aptamers to functionalize an in situ injectable hydrogel model to control the release of proteins. The aptamers were studied using secondary structural predictions and binding analyses. The results showed that the structural predictions were different from the experimental measurements in numerous cases. The affinity of the aptamer was significantly affected by the mutations of the essential nucleotides, whereas it was not significantly affected by the variations of the nonessential nucleotides. The mutated aptamers were then used to functionalize the injectable hydrogel model. The results showed that the aptamer-functionalized hydrogel could prolong protein release. Moreover, the release rates could be controlled by adjusting the affinity of the aptamer.

Introduction Tremendous efforts have been made in developing proteins as therapeutic reagents for the treatment of human diseases because proteins are essential biomolecules of organisms and are involved in virtually every process in a cell.1 Despite the great potential of protein drugs in disease treatments, protein delivery remains a long-standing challenge. The development of innovative polymeric systems has received significant attention in the field of protein delivery and has been proposed as a promising strategy for controlling the release of protein drugs at a target site with a desired rate.2 One of the most appealing polymeric systems is hydrogels because hydrogels have a number of advantages (e.g., structural similarity to human tissues) for in vivo applications over other polymeric systems.2-7 Hydrogels are made of hydrophilic polymers3 that can be either natural biomolecules8,9 or synthetic materials.10-15 The hydrogel networks are usually formed by the cross-linking of polymer chains via covalent bonds, hydrogen bonds, or ionic interactions.3 Hydrogels can be synthesized outside of the body for in vivo implantation. Alternatively, a polymer solution can be rationally designed and directly injected into the desired site where the solution is transformed into a hydrogel (i.e., in situ gelation).16 In comparison to hydrogel implants, the in situ gelation of polymer solutions has special merits for in vivo applications. An injectable hydrogel can be delivered in vivo with minimal surgery and, in principle, can fill cavities with any geometry.16 Thus, the delivery of in situ injectable hydrogel formulations will not only improve patient compliance and quality of life, but also avoid the need for fabricating patient-specific hydrogel implants. The in situ gelation can be simply achieved through diverse mechanisms, such as temperature-induced phase transition, UV-mediated polymerization, polyelectrolyte complexation, solvent exchange, and self-assembly.16-18 Because of these merits, the in situ injectable hydrogels have been widely studied for protein * To whom correspondence should be addressed. Tel.: (860) 486-4072. Fax: (860) 486-2959. E-mail: [email protected].

delivery.16 However, most hydrogels are highly permeable, which can lead to the rapid release of loaded proteins.7 Thus, there is a clear need to develop novel methods for improving the properties of injectable hydrogels to achieve desired protein release kinetics. Here we demonstrate a novel strategy for synthesizing in situ injectable hydrogels to control the release of proteins by using nucleic acid aptamers as affinity sites for the proteins in the hydrogel. Aptamers are single-stranded DNA or RNA oligonucleotides that are discovered from nucleic acid libraries with the technology named systematic evolution of ligands by exponential enrichment (SELEX).19,20 In principle, a specific aptamer can be selected to recognize any protein drug with high affinity.21 Aptamers can also be chemically modified to be tolerant of harsh chemical and biological conditions.21 Moreover, aptamers are small and can be synthesized with a standard chemical route.21 Because of these merits, aptamers have been studied for various biological and biomedical applications.22-38 For instance, we have demonstrated that aptamers can be chemically incorporated into a polyacrylamide hydrogel to slow the protein release.39 The model aptamer used in this study was previously selected against platelet-derived growth factor B (PDGF-B) by Green et al. from a DNA library by using a gel electrophoresis-based SELEX approach.40 To understand the effect of sequence modifications on the binding functionality of this model aptamer, we generated a series of anti-PDGF aptamers either by randomizing the nonessential nucleotide tail or by mutating the essential nucleotides. The functionality of these aptamers was studied by the examination of their secondary structures and dissociation constants. Based on these studies, several aptamer sequences were selected to investigate the protein release from an aptamer-functionalized in situ injectable poloxamer hydrogel. Poloxamer is a block copolymer that has been proposed for a variety of pharmaceutical applications,18 such as the delivery of growth factors41 and viruses.42 The results demonstrated that the aptamer-functionalized poloxamer hydrogels could slowly release PDGF-BB with tunable kinetics.

10.1021/bm100774t  2010 American Chemical Society Published on Web 09/01/2010

Injectable Hydrogel for Controlled Protein Release Table 1. List of DNA Sequencesa sequence ID S1 S2 S3 S4 S5 S6 S7 S8 S9 S10 M1 M2 M3 S-S1 FAM-CO

nucleotide sequence (5′f3′) GCGATACTCC ACAGGCTACGGCACGTAGAGCATCACCATGATCCTG CAATTCCGCG ACAGGCTACGGCACGTAGAGCATCACCATGATCCTG CCACGGTCTA ACAGGCTACGGCACGTAGAGCATCACCATGATCCTG CGCCATTCAG ACAGGCTACGGCACGTAGAGCATCACCATGATCCTG CGCATGCTCA ACAGGCTACGGCACGTAGAGCATCACCATGATCCTG TCGCACATGC ACAGGCTACGGCACGTAGAGCATCACCATGATCCTG GCCGTTCCAA ACAGGCTACGGCACGTAGAGCATCACCATGATCCTG TGCCATGCCA ACAGGCTACGGCACGTAGAGCATCACCATGATCCTG GCAACTGCTC ACAGGCTACGGCACGTAGAGCATCACCATGATCCTG CATGAGCCCT ACAGGCTACGGCACGTAGAGCATCACCATGATCCTG GCGATACTCC ACAGGCTACGGCACGTAGAGCATCACCATGATCCTA GCGATACTCC ACAGGCTACGGCACGTAGAGCATCACCATGATCCCA GCGATACTCC ACAGGCTACGGCACGTAGAGCATCACCATGATCTCA GCGATACTCC ATCAATGGACCGCGCACTCGCCAGTGCTAATGGCAA FAM-CAGGATCATGGTGATGCT CTACGTGCCGTA

a The nonessential nucleotide tail is underlined. The mutated nucleotides are italic and bold. CO: complementary oligonucleotide.

Experimental Section Materials. All the DNA molecules were purchased from Integrated DNA Technologies (Coralville, IA) and listed in Table 1. Recombinant human PDGF-BB and bovine serum albumin (BSA) were purchased from R & D Systems (Minneapolis, MN) and Invitrogen (Carlsbad, CA), respectively. Poloxamer 407 (Pluronic F-127), biotinyl-N-hydroxysuccinimide (NHS-biotin), and 2-(N-morpholino) ethanesulfonic acid (MES) were purchased from Sigma-Aldrich (St. Louis, MO). The 5K membrane filter unit was purchased from Millipore (Billerica, MA). Streptavidin-coated microparticles (1.3 µm) were purchased from Spherotech (Lake Forest, IL). N-Ethyl-N-(3-diethylaminopropyl) carbodiimide (EDC), N-hydroxysuccinimide (NHS), phosphate buffered saline (PBS), disodium hydrogen phosphate (Na2HPO4), Tween 20, and sodium azide (NaN3) were purchased from Fisher Scientific (Suwanee, GA). Modification of Aptamer Sequence. The sequence of the aptamer was modified through either tail variation or stem mutation. For the tail variation, the tail was generated by sequence randomization with A:C:G:T ratio maintained the same. For the stem mutation, the first three nucleotides G, T, and C at the 3′-end of the aptamer S1 were replaced with A, C, and T, respectively. The aptamers with one, two, and three mutation sites were denoted as M1, M2, and M3, respectively. The sequences of modified aptamers are shown in Table 1. Structure Prediction. The secondary structures of these aptamer sequences were predicted by using the program RNAstructure, version 4.6 (http://rna.urmc.rochester.edu/rnastructure.html). This program can be used to generated the secondary structures of both RNA and DNA oligonucleotides. The secondary structures with the lowest free energies were used for presentation and analysis. Measurement of Binding Affinity. The affinities of the aptamers were measured with surface plasmon resonance (SPR) spectrometry (SR7000DC, Reichert Analytical Instrument, Depew, NY). Carboxyl group-functionalized sensor chips were purchased from the Reichert Analytical Instrument. The chips were initially activated with 0.2 M EDC/0.1 M NHS for 10 min. After the activation step, PDGF-BB (20 µg/mL in 25 mM Na2HPO4 at pH 8.5), was flowed over the activated sensor chip for protein immobilization. Before the test, the system was equilibrated with the running buffer for 30 min. The running buffer was made of PBS buffer (pH 7.4) containing 0.05% Tween 20. During

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the test, the aptamer solution was flowed over the sensor chip at a flow rate of 30 µL/min for 5 min. Subsequently, the flowing solution was switched to the running buffer for molecular dissociation. After each test, the sensor chip was regenerated by flowing 1 M NaCl in the channel for 2 min and then washing with the running buffer. To determine the dissociation constants, a series of aptamer solutions were prepared, with the concentration ranging from 3.13 to 200 nM. The data were processed with the Scrubber 2.0 software (BioLogic Software, Australia). The responses from the reference channel were subtracted from the responses from the analyte channel before data analysis to minimize the effects of reflective index changes, nonspecific binding, and instrument drift. The dissociation constant (KD) was determined by fitting the data to a simple biomolecular “1:1 binding” model. Preparation of Aptamer-Functionalized Poloxamer Hydrogel. The aptamers with a primary amine group at the 5′-end were reacted with NHS-biotin at pH 7.0 overnight. The free biotin was removed from the reaction mixture by filtration through a 5K membrane filter unit. The aptamer solution containing a total of 2.5 nmol of aptamers was mixed with 1 mg of streptavidin-coated polystyrene particles in 100 µL of PBS. After a 30 min incubation, the functionalized particles were washed with PBS four times. To prepare the affinity poloxamer hydrogel, 80 µg of aptamer-functionalized particles were first incubated with 4 ng of PDGF-BB in 20 µL PBS for 30 min at room temperature. The suspension was then mixed with 1000 µL of 20% w/w poloxamer solution at 4 °C. Finally, 250 µL of suspension was transferred into a 2 mL tube and allowed to form the hydrogel at 37 °C for 30 min before the hydrogel characterization and the release studies. Flow Cytometry. Flow cytometry experiments were performed to examine the functionalization of the particles with the aptamers. Approximately 2 × 107 particles were incubated in 100 nM of complementary oligonucleotide labeled with 6-carboxy-fluorescein (denoted as FAM-CO) for 1 h. The particles were then washed twice with 500 µL of PBS before subjected to flow cytometry analysis with a BD FACSCalibur flow cytometer (San Jose, CA). A total of 10000 events were collected for data analysis. Microscopic Examination. The particle suspensions were spread on the surface of glass slides, covered with a coverslip, and examined by using a Leica SP2 spectral confocal microscope with a 100× objective. The images were processed using the Leica Confocal software provided by the manufacturer. Rheology Characterization. The storage (G′) and loss (G′′) moduli of the hydrogels were measured with an AR-G2 rheometer (TA Instruments, New Castle, DE). Approximately 200 µL of cold particle-hydrogel suspension was loaded into the chamber. The experiments were performed with an oscillation mode. To ensure the validity of the data, a linear viscoelastic regime was first determined by performing a stress-sweep experiment at both 4 and 37 °C. The oscillation stress was varied from 0.01 to 1000 Pa at a fixed frequency of 0.1 Hz. The temperature-dependent moduli were measured from 4 to 45 °C with a fixed oscillation stress (6 Pa) and a constant heating rate (2 °C/min). The gelation point was defined as the crossover point of G′ and G′′. In addition, the time-sweep modulus of the poloxamer solution with or without particles was measured at 37 °C for 1 h. The oscillation stress was fixed at 6 Pa during the measurement. Measurement of PDGF-BB Release. The release medium (i.e., PBS containing 0.1% BSA, 0.05% Tween-20, and 0.05% NaN3) was first prewarmed to 37 °C. Next, 500 µL of release medium was carefully added into the tube containing 250 µL of poloxamer hydrogel. The tubes were incubated at 37 °C with a shaking rate of 60 rpm. At predetermined time points, 500 µL of release medium was carefully collected from the solution and replenished with 500 µL of fresh release medium. PDGF-BB in the release medium was quantified with a human PDGF-BB ELISA kit (PeproTech, Rocky Hill, NJ). The experiments were performed in triplicate. The total amount of PDGF-BB in the native hydrogel was analyzed at the end of the release experiment to calculate the cumulative release of PDGF-BB.

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Figure 1. Secondary structures of anti-PDGF-BB aptamers with different tail compositions. The 10-nt tail is marked in blue.

Results and Discussion The model aptamer can bind PDGF-BB with high affinity and specificity. Its original sequence contains 86 nucleotides.40 In general, approximately 10-15 nucleotides of an aptamer form a functional structure to interact with the target.43 These nucleotides exhibit the structures such as hairpin loops, quartet loops, bulges, or pseudoknots.43 However, these nucleotides require presentation in the context of the “parent” aptamer to achieve a sufficient binding capability. Thus, the essential nucleotides of an aptamer include not only the nucleotides forming a functional structure, but also those providing the right context to facilitate the formation of the functional structure. An aptamer usually contains 25-40 essential nucleotides.21 The other nucleotides are nonessential because they do not bind the target or facilitate the binding of the aptamer. Our previous study has shown that the anti-PDGF-BB aptamer with 36 essential nucleotides can bind PDGF-BB with high affinity.39 One of the main purposes of this study is to understand the effect of sequence modifications on the binding functionality of the aptamer. Thus, we used two methods to tune the binding functionality of the anti-PDGF-BB aptamer: changing the context of the essential nucleotides by varying the composition of a nonessential nucleotide tail (Figures 1 and 2) and mutating the stem of the aptamer (Figures 3 and 4). Effects of Tail Composition on Secondary Structure and Binding Functionality. Because the binding capability of a nucleic acid aptamer is dependent on its functional structure,21 we first varied the sequence and structure of the 36-nt aptamer by attaching a nonessential nucleotide tail to its 5′-end. The tail contained 10 nucleotides. Our hypothesis was that the nonessential nucleotide tail could form intramolecular base pairs with the essential nucleotides. As a result, the variation of the tail would change the context of the essential nucleotides and the binding affinity of the aptamer. The hypothesis was first tested by analyzing the secondary structures predicted with the program RNAstructure. As shown in Figure 1, the 36 essential nucleotides adopt a secondary structure that has three stem regions radiating from a common junction. The 36-nt aptamer and the 36 essential nucleotides in the aptamer S1 (i.e., the sequence truncated from the full-length

aptamer without randomization; Figure 1) virtually exhibit the same structure. The 5′- and 3′-ends form a four-base-pair stem. Totally, we studied 10 aptamers with differential tail compositions and one scrambled aptamer. The representative structures of the aptamers are shown in Figure 1. The aptamer S1 is composed of 36 essential nucleotides and 10 nonessential nucleotides of the “parent” aptamer. The other nine aptamers are composed of 36 essential nucleotides and a randomized sequence of the 10-nt tail. The scrambled aptamer S-S1 was generated by randomizing the 36 essential nucleotides of aptamer S1. Two aptamers, S2 and S3, exhibit the same threestem structure as the 36-nt aptamer. Others exhibit a completely different structure. Because the structure of an aptamer plays a

Figure 2. Effect of tail variation on the aptamer-protein interaction. (A) SPR sensorgrams. (B) Normalized equilibrium responses. The normalization was performed by dividing the equilibrium response of each aptamer by that of S1.

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Figure 3. Secondary structures of anti-PDGF-BB aptamers with different mutations. The mutated nucleotides are marked in green. The 10-nt tail is marked in blue.

Figure 4. Effect of stem mutation on the aptamer-protein interaction. (A) SPR sensorgrams. (B) Normalized binding responses.

critical role in determining its binding capability,21 the structural prediction indicates that aptamer S2 and aptamer S3 can bind PDGF-BB, whereas the others may not. However, as shown in Figure 2, the results from SPR analysis are not in full agreement with the predictions of the secondary structures. Six aptamers (S2, S3, S4, S6, S9, and S10) virtually exhibited the same binding capability as the aptamer S1. The other three aptamers (S5, S7, and S8) exhibited weaker binding capability. The difference between the structural predictions and the experimental measurements may result from different working conditions. Current algorithms (e.g., the RNAstructure program) were developed to predict the secondary structures of aptamers in a clean system. A clean system accounts for only the aptamers in a buffer solution. However, the SPR analysis is used to analyze the interactions of a molecular pair in a buffer solution. Thus, in addition to the aptamers, the system also has proteins. The proteins will compete with the nucleotide tail in binding to the essential nucleotides. If the intermolecular interaction between the proteins and the essential nucleotides is much stronger than the intramolecular base paring between the nucleotide tail and the essential nucleotides, the tail may not interfere with the structural formation of the essential nucleotides in the presence of the target protein. In fact, as shown in the predicted structures (Figure 1), the intramolecular interactions did not result in a large number of Watson-Crick base pairs in some aptamers (e.g., S9), which indicates that the nucleotide

tail interacts weakly with the essential nucleotides. Previous studies have shown that aptamers can undergo conformational changes upon binding to a target.44,45 In addition, the essential nucleotides of aptamers originally hybridized with shorter complementary oligonucleotides can change their structures in the presence of target molecules and still bind their target.37,46 Thus, both the previous studies and this study indicate that the essential nucleotides may have the capability of forming the functional structure in the presence of their target molecules and that the variation of a nonessential nucleotide tail, at least in the current system, may not significantly change the affinity of the aptamer. Effects of Stem Mutation on Secondary Structure and Binding Functionality. Because the randomization of a nonessential nucleotide tail did not result in significant interference with the aptamer-protein interactions, we further mutated the essential nucleotides in the four-base-pair stem formed at the 5′- and 3′-ends. The difference between these two methods is that the former one did not change the sequence of the essential nucleotides, whereas the later one directly changed it. Three aptamer mutants were generated: M1, M2, and M3. Their sequences are shown in Table 1. These mutants have one, two, or three mutated nucleotides at the 3′-end, respectively. These three aptamer mutants exhibit the same structural format as predicted by the program RNAstructure (Figure 3). However, these secondary structures do not resemble that of aptamer S1 (Figure 1). Aptamer S1 has a four-base-pair stem at the 5′- and 3′-ends, whereas none of the mutants have a base pair at the 5′- and 3′-ends according to the structural prediction. Even aptamer M1, with only one mutated nucleotide, does not form a stem at its 5′- and 3′-ends (Figure 3). Interestingly, though the predicted stem-loop structures of the three mutants are the same, the SPR analysis showed that each exhibited a different capability of binding PDGF-BB (Figure 4). The maximal SPR response decreased with an increasing number of mutations, indicating that the binding capability decreased. As discussed earlier, some of the essential nucleotides play the role of forming the functional structure for binding the target, while the others provide a context to stabilize the structure. It is likely that the stem at the 5′- and 3′-ends of the sequence of the essential nucleotides plays a role in stabilizing the functional structure. When this region has mutations, the stem stability is decreased. Though the presence of the target protein might aid in facilitating the formation of the functional structure, it was not enough to overcome the instability resulting from the mutations in the stem region. As a result, the overall structural stability is decreased. The degree of decreased stability is dependent on the number of mutations. The interaction of a molecular pair is determined by not only molecular association but also by molecular dissociation. The dissociation rate constant divided by the association rate constant

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Figure 5. Determination of dissociation constants (KD). (A) Concentration-dependent binding sensorgrams. (B) Equilibrium binding plot. (C) Table for dissociation constants.

is defined as the equilibrium dissociation constant (KD). Thus, we further prepared a series of aptamer solutions and characterized the aptamer-protein interactions by calculating the dissociation constants (Figure 5). The concentration-dependent responses were processed to calculate KD using an equilibrium analysis.47,48 This analysis approach minimizes the inaccuracy due to the limitation of mass transport on the biochip surface.49 The KD values of M1, M2, and M3 were 27.6, 109, and 354 nM, respectively. These data clearly show that the binding affinity of the aptamer decreases with the increasing number of mutations. Preparation and Characterization of Aptamer-Functionalized Poloxamer Hydrogel. The affinity hydrogel was synthesized by mixing the aptamer-functionalized particles with a poloxamer solution (20% w/w). First, the properties of the particles were characterized by microscopy and flow cytometry (Figure 6). The microscopic observation indicated that the overall particle morphology did not change after the functionalization with the aptamers or after the adsorption of PDGF-BB and that there was no significant particle cross-linking or aggregation. To confirm that the anti-PDGF-BB aptamers were tethered to the particles, FAM-labeled complementary oligonucleotides were used to treat the particles. The flow cytometry data showed the presence of the complementary oligonucleotides on the particle surface, indicating that the aptamers were tethered to the particles. We then characterized the properties of the affinity hydrogel with microscopic observation and rheology analysis. The microscopy images showed that the particles were well distributed in the hydrogel (Figure 7A). The rheology data demonstrated that the mechanical properties were not changed after the incorporation of the particles into the poloxamer hydrogel (Figure 7B and 7C). Affinity hydrogels are usually synthesized by the chemical conjugation of the ligands to the backbone of polymers.39,50,51 However, previous studies showed that a significant amount of affinity ligands could not be incorporated into the hydrogel network during the formation of hydrogels.39,52 This would negatively affect the loading efficiency of the proteins into the system. In addition, the free affinity ligands would diffuse out

Figure 6. Particle characterization. (A) Microscopic observation of the particles in PBS: A1, streptavidin-coated polystyrene particles; A2, aptamer-coated particles; and A3, aptamer-coated particles treated with PDGF-BB. Scale bar: 10 µm. (B) Flow cytometry histograms.

of the hydrogel, and could bind and inactivate protein drugs during the protein release. This problem can be significant if a higher concentration of affinity ligands is required. One may propose to wash free ligands out of the hydrogel. However, it is not possible to prewash hydrogels for in situ applications. This study shows a different approach for developing affinity hydrogels because the preparation of the hydrogels does not need any chemical conjugation between the aptamers and the hydrogel. This system only requires the physical mixing of the affinity particles, the protein drugs, and the polymer solution. Free aptamers can be removed before the particles are mixed with protein drugs and polymer solution. Thus, it is possible that any type of injectable hydrogel can be functionalized with nucleic acid aptamers by using this method for preparing an in situ injectable affinity hydrogel for controlled protein release. Protein Release from Aptamer-Functionalized Poloxamer Hydrogel. A variety of in situ injectable hydrogels have been studied for the delivery of protein drugs.16,17 One of them is

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Figure 7. Hydrogel characterization. (A) Examination of particle distribution in the poloxamer hydrogel: A1, with particles; A2, without particles. Scale bar: 10 µm. (B) Characterization of storage (G′) and loss (G′′) modulus vs temperature. (C) Characterization of storage (G′) modulus at 37 °C.

thermosensitive hydrogels.17 Because their solutions can be easily transformed into a gel state at body temperature, they have been widely used for protein delivery in the field of regenerative medicine and tissue engineering.17,18 In this study, poloxamer 407 was used as the model to investigate the capability of aptamers in controlling the protein release. Poloxamer 407 was a suitable model because it has been wellstudied for drug delivery18 and its solution can be transformed into a gel state by increasing the temperature from a low degree to a high degree (e.g., body temperature). For instance, the 20% w/w solution formed a hydrogel with the temperature increased to approximately 20 °C as determined by the crossover point of G′ and G′′ (Figure 7B). Drug release from a native hydrogel is governed by both drug diffusion and poloxamer dissolution.18 The poloxamer hydrogel was directly incubated in the release medium with no membrane to separate the hydrogel from the release medium. Thus, water uptake, poloxamer dissolution, and protein release could happen simultaneously. When the poloxamer hydrogel starts to dissolve in the release medium, its hydrogel concentration decreases.18 Because the formation of a poloxamer hydrogel is not only a function of temperature, but of concentration as well, the decrease of the concentration of the hydrogel will in turn accelerate its dissolution.18 The drug release rate will increase

Figure 8. (A) PDGF-BB release from aptamer-functionalized poloxamer hydrogels. (B) The first-day release rate as a function of KD value. (C) The cumulative release for 14 days as a function of KD value.

in parallel. Previous studies have shown that the drug release rate and poloxamer dissolution follow zero-order kinetics.53 Our data also showed that PDGF-BB release from a native poloxamer hydrogel (i.e., poloxamer hydrogel without aptamer-coated particles) was fast and exhibited apparent zero-order kinetics during the first day (Figure 8). More than 80% of the loaded PDGF-BB was released during the first day. The fast release will not only raise the cost of treatments, but will also lead to a wide distribution of protein drugs in nontarget tissues and cause in vivo side effects.1 This is a particularly important issue in the delivery of a protein drug (e.g., interleukin 2) with a narrow therapeutic index.54 In contrast, the PDGF-BB release from the aptamer-functionalized poloxamer hydrogels (i.e., poloxamer hydrogels with aptamer-coated particles) was significantly decreased (Figure 8). For instance, less than 10% of the loaded PDGF-BB was released from the S1-functionalized hydrogel during the first day. A total of 14.5% was released within the first two weeks. It was not surprising to observe the slow release of proteins

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from the aptamer-functionalized poloxamer hydrogels because the aptamers functioned as the affinity sites of the proteins in the hydrogels. The strong molecular interaction between proteins and aptamers creates a significant barrier for protein diffusion and therefore retards the release of proteins from the hydrogels. The results also showed that the release rates of PDGF-BB could be controlled by varying the affinity of the anti-PDGF-BB aptamer (Figure 8). The first-day release rate and the cumulative release at the end of the experiment decreased with the decrease of the KD value (Figure 8B,C). The higher affinity indicates the stronger molecular interaction. The stronger molecular interaction will retard the diffusion of proteins from the particle surface and the hydrogel more significantly. Thus, the release was slower when poloxamer hydrogels were functionalized with higher affinity aptamers.

Conclusion Structural predictions and binding analysis were used to study a number of anti-PDGF-BB aptamers that were generated by the mutation of the essential nucleotides and the variation of the nonessential nucleotides. The results showed that the mutation of the essential nucleotides in the stem region significantly altered the binding affinity of the aptamer, depending on the number of mutations. In contrast, the sequence modifications of the nonessential nucleotide tail did not significantly alter the affinity of the aptamer. The difference between the structural predictions and the experimental measurements indicated that the aptamers could undergo structural changes in the presence of their target molecules. The aptamers were further used to functionalize the poloxamer hydrogel. The functionalization did not need any specific chemical modification of the hydrogel. The release tests demonstrated that PDGF-BB was rapidly released from the native poloxamer hydrogel. In contrast, its release from the aptamer-functionalized hydrogels was significantly prolonged. The release rate could be controlled by adjusting the affinity of the aptamer. Therefore, the results demonstrate that nucleic acid aptamers, in principle, can be applied to functionalize any in situ injectable hydrogel for controlled protein release. Acknowledgment. The authors greatly acknowledge Dr. Carol Norris for technical support in using the flow cytometer and the confocal microscope. The authors also thank Mr. Mark Battig for reading the manuscript. B.S. is in part supported by the Royal Thai Government Scholarship. This work is supported by the NSF Grant DMR-0955358 to Y.W.

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