Degradation of Poly (glycoamidoamine) DNA Delivery Vehicles

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Biomacromolecules 2010, 11, 316–325

Degradation of Poly(glycoamidoamine) DNA Delivery Vehicles: Polyamide Hydrolysis at Physiological Conditions Promotes DNA Release Yemin Liu and Theresa M. Reineke* Virginia Tech, Department of Chemistry, Macromolecules and Interfaces Institute, Blacksburg, Virginia 24061 Received July 20, 2009; Revised Manuscript Received October 24, 2009

Poly(glycoamidoamine)s (PGAAs) are a group of efficient and degradable gene delivery vehicles that consist of three main functionalities: carbohydrate groups, secondary amines, and amide bonds. Herein, we have created nonhydroxylated models to these structures by polymerizing oxylate, succinate, or adipate groups with pentaethylenehexamine. The resulting polymers (named O4, S4, and A4, respectively) were created to understand how the absence of hydroxyl groups and changes in the amide bond spacing affect polymer degradation, plasmid DNA (pDNA) complexation, toxicity, and transfection efficiency in vitro. An additional model was also created that retains a galactaramide unit, but we have replaced the secondary amines with ethyleneoxide units (GO2) to understand the effects of the amine groups on polymer degradation. We have found that the secondary amines and hydroxyls are necessary to facilitate rapid degradation of these polymers, and analogues lacking hydroxyls or amines did not degrade over the time course of the study. Through electron-withdrawing and hydrogen bonding, the hydroxyls appear to activate the carbonyls of the amide bond to hydrolysis via an inductive electron withdrawing effect. Through titration experiments, PGAA degradation appears not to affect the polymer buffering capacity. Furthermore, we have found that PGAA degradation may enhance gene expression by releasing pDNA from polyplexes (polymer-pDNA complexes) and, thus, exposing it to undergo transcription and translation. The difference in the optimal pH that promotes degradation of the PGAAs and the hydroxylfree analogues may prove to be a useful means to achieve pH-regulated DNA release from polyplexes by specifically modulating the chemical structures.

Introduction Synthetic polymers are a versatile class of nonviral nucleic acid delivery vehicles. The structure and chemical composition of these materials can be easily manipulated via chemical synthesis to understand the biological properties and tune polynucleotide delivery efficiency and polymer toxicity. Indeed, many studies in this field have shown the important relationship between polymer structure, cellular delivery efficacy, and cytotoxicity.1–10 Prior investigations in this field have revealed that the biodegradability of polymeric gene delivery vectors is a key factor that leads to low cytotoxicity11–13 and that degradation of the polymer vectors also enhances DNA release from the polyplexes.1 Biodegradable structural modules, including esters,14–16 disulfides,17,18 ortho esters,19 acetals,20 and hydrazones,21 have been widely used in constructing synthetic gene delivery vectors to promote biocompatibility and nucleic acid release. An important strategy in designing novel biomaterials is the incorporation of natural building blocks into the polymer backbone. Carbohydrates are renewable monomer sources that can readily generate versatile synthetic materials with the advantage of being biocompatible and biodegradable. Toward the development of efficient and benign gene delivery vectors, multiple poly(glycoamidoamine)s (PGAAs, Figure 1) were synthesized by polymerizing oligoethyleneamine monomers with carbohydrate moieties that are linked through an amide bond. These materials have generally revealed high gene delivery efficacy and low cytotoxicity.2,3,22,23 Systematic examination of the structure-biological activity of our various PGAAs * To whom correspondence should be addressed. E-mail: treineke@ vt.edu.

Figure 1. Structures of poly(glycoamidoamine)s D4, G4, M4, and T4, where R is mostly H. Some points of branching likely exist along the backbone.

revealed that the structural features of the comonomers, namely, the carbohydrate type [D-glucaramide (D), meso-galactaramide (G), D-mannaramide (M), or L-tartaramide (T)] in the carbohydrate unit and the number of secondary amines (1-4) in the oligoamine unit, significantly affect the overall biological properties of the final polymers, including the pDNA binding affinity, polyplex stability in serum, nuclease protection, and buffering capacity.2,3,22–26 In all of our studies, we have found that the structures that have four secondary amines in the repeat unit (D4, G4, M4, and T4) yield the highest delivery efficiency. During the course of these previous studies, we noticed that when the polyplexes were formulated immediately after dissolving the PGAAs in water, the transfection efficiency was very high. However, if the polyplexes were formulated 24 h or more after dissolving the polymers in water and keeping the polymer solution at 4 °C, a complete lack of transfection was observed. Although polyamides are typically thought to be very stable from hydrolysis, we hypothesized that the functional

10.1021/bm9008233  2010 American Chemical Society Published on Web 01/08/2010

Poly(glycoamidoamine) DNA Delivery Vehicles

Biomacromolecules, Vol. 11, No. 2, 2010

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Scheme 1. Synthesis of Polymers A4, S4, and O4

groups (hydroxyls and amines) within the PGAA structures could play a synergistic role in contributing to amide bond degradation. Kiely et al.,27,28 Liu and Reineke,2 and others29–31 have shown in previous studies that polyhydroxylated esters are highly reactive toward aminolysis and amine bond formation. Kiely et al. and Ogata et al.32 have shown that this high reactivity is likely due to the electron withdrawing nature of the R-hydroxyl group to the ester carbonyl, which promotes aminolysis. Nonhydroxylated esters such as adipate and succinate do not share this high reactivity with amines. Hydrogen bonding (possibly both intra- and intermolecular) also appears to play a role in facilitating this reactivity. In the current study, we propose that a similar mechanism may contribute to PGAA degradation. To this end, we have investigated the degradability of the PGAAs using a variety of techniques such as gel permeation chromatography, FT-IR, and UV spectroscopy. To help assign the effects of the hydroxyl and secondary amine groups on PGAA degradation, we have synthesized analogous models to the PGAA polymers. The first models lack the hydroxyl units along the polymer backbone by replacing the carbohydrate monomer with either an oxylate, succinate, or adipate unit (O4, S4, and A4). The other analogous model was created to contain ethyleneoxides (GO2) in place of the ethyleneamines. In addition to the physical characterization, we have also studied and compared the potential effects of these new analogues to our original PGAAs for polymer vehicle degradability on plasmid DNA (pDNA) binding, gene delivery efficiency, and cytotoxicity.

Experimental Section I. Preparation of Monomers, Copolymers, and Polyplexes. All reagents, unless specified, were obtained from Acros Chemical Co. (Morris Plains, NJ) and used without further purification. The PGAAs were synthesized according to our previously published methods.2,3 After polymerization of all the polymers, the product was dissolved and dialyzed in ultrapure water using a Spectra Por 500 molecular weight cutoff membrane and lyophilized to dryness with a Flexi-dry MP lyophilizer (Stone Ridge, NY). Polyplexes (polymer-pDNA complexes) were also formed according to these same published methods at various N/P ratios [the ratio of the polymer amine groups (N) to phosphate groups (P)], which is a conventional method to denote the formulation ratio between the polymer and the pDNA in the field of gene delivery. Dialysis membranes were purchased from Spectra Por (Rancho Dominguez, CA). FT-IR spectra of polymers were taken as KBr pellets on a Perkin-Elmer Spectrum One Fourier transform infrared spectrometer. NMR spectra were collected on a Bruker AV400 MHz spectrometer. Compound 1 was synthesized according to our previously reported method, which is detailed in the Supporting Information.33,34 Poly(oxalamidopentaethylenehexamine) (O4). As shown in Scheme 1, a solution of oxalyl chloride (0.10 g, 0.79 mmol/5.00 mL of CH2Cl2)

was added dropwise to a solution of 1 (0.50 g, 0.79 mmol/5.00 mL of CH2Cl2). The mixture was stirred at room temperature for 15 min. After evaporating the CH2Cl2, the reaction mixture was dialyzed against methanol overnight using a Spectra Por 1000 molecular weight cutoff membrane. After evaporating the methanol, the resultant polymer was stirred with 10 mL of 4 M HCl in dioxane for 4 h to remove the ditert-butyl dicarbonate (Boc) groups. After evaporating the dioxane, O4 was precipitated in methanol as a light yellow solid. Yield: 0.13 g, 64%. IR (KBr): 3427 and 3312 cm-1 (N-H stretching), 2946 cm-1 and 2678 cm-1 (C-H stretching), 1668 cm-1 (amide CdO stretching), 1520 cm-1 (amide N-H bending), 1446 cm-1 (CH2 scissoring), 1393 cm-1 and 1374 cm-1 (amide C-N stretching), 1286-1057 cm-1 (amine C-N stretching). 1H NMR (D2O with one drop of DCl): 3.26 (t, 4H), 3.48-3.56 (br, 16H). Poly(succinamidopentaethylenetetramine) (S4). As shown in Scheme 1, a solution of succinyl chloride (0.10 g, 0.64 mmol/2.50 mL of CH2Cl2) was added dropwise to a solution of 1 (0.41 g, 0.64 mmol/ 2.50 mL of CH2Cl2). The mixture was stirred at room temperature for 3 h. After evaporating the CH2Cl2, the residue was dialyzed against methanol overnight using a Spectra Por 1000 molecular weight cutoff membrane. After evaporating the methanol, the resultant polymer was stirred with 8 mL of 4 M HCl in dioxane for 4 h to remove the Boc groups. After evaporating the dioxane, S4 was precipitated in methanol as a white solid. Yield: 0.13 g, 64%. IR (KBr): 3427 cm-1 and 3307 cm-1 (N-H stretching), 2981 cm-1 and 2948 cm-1 (C-H stretching), 2800-2300 cm-1 (broad amine salt N-H stretching), 1648 cm-1 (amide CdO stretching), 1539 cm-1 (amide N-H bending), 1469 cm-1 (CH2 scissoring), 1338 cm-1 (amide C-N stretching), 1254-1059 cm-1 (amine C-N stretching). 1H NMR (D2O with one drop of DCl): δ 2.55 (s, 4H), 3.26 (t, 4H), 3.48-3.56 (br, 16H). Poly(adipamidopentaethylenetetramine) (A4). As shown in Scheme 1, a solution of adipoyl chloride (0.10 g, 0.55 mmol/2.50 mL of CH2Cl2) was added dropwise to a solution of 1 (0.35 g, 0.54 mmol/2.50 mL of CH2Cl2). The mixture was stirred at room temperature for 1 h. After evaporating the CH2Cl2, the residue was dialyzed against methanol overnight using a Spectra Por 1000 molecular weight cutoff membrane. After evaporating the methanol, the resultant polymer was stirred with 6 mL of 4 M HCl in dioxane for 4 h to remove the Boc groups. After evaporating the dioxane, A4 was precipitated in methanol as a white solid. Yield: 0.13 g, 70%. IR (KBr): 3427 cm-1 and 3298 cm-1 (N-H stretching), 2982 cm-1 and 2950 cm-1 (C-H stretching), 2800-2300 cm-1 (broad amine salt N-H stretching), 1659 cm-1 (amide CdO stretching), 1539 cm-1 (amide N-H bending), 1494 cm-1 (CH2 scissoring), 1353 cm-1 (amide C-N stretching), 1251-1060 cm-1 (amine C-N stretching). 1H NMR (D2O with one drop of DCl): δ 1.45 (s, 4H), 2.17 (s, 4H), 3.16 (t, 4H), 3.35-3.50 (br, 16H). Poly(galactaramidotriethylenedioxide) (GO2). As shown in Scheme 2, dimethyl galactarate2,27 (0.20 g, 0.84 mmol) was mixed with 2,2′(ethylenedioxy)bis(ethylamine) (0.124 g, 0.84 mmol/in 4.2 mL of methanol) and stirred at room temperature for 20 h. Yield: 0.10 g, 37%. IR (KBr): 3354 cm-1 (O-H stretching), 2940 cm-1 and 2874 cm-1 (C-H stretching), 1645 cm-1 (amide CdO stretching), 1544 cm-1 (amide N-H bending), 1435 cm-1 (CH2 scissoring), 1353 cm-1 and

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Scheme 2. Synthesis of GO2

1299 cm-1 (CH2 twisting and wagging), 1113 cm-1 (C-O-C asymmetric stretching), 1043 cm-1 (C-O-C symmetric stretching). 1H NMR (D2O): δ 4.31 (s, 2H), 3.91 (s, 2H), 3.56 (br, 8H), 3.36 (br, 4H). II. Polymer Characterization and Degradability Assays. For the polymer degradability study, 5.0 mM of D4, G4, M4, T4, A4, S4, O4, and GO2 (final concentration in terms of polymer repeat unit) were incubated in ultrapure water and in various buffers (PBS, pH 7.4; citric acid-phosphate buffer, pH 5.0) at 37 °C. At the desired time points (0, 12, 24, 48, 72, and 120 h), the incubation mixtures were flash frozen with liquid N2 and stored at -80 °C until further analysis. Gel Permeation Chromatography (GPC). The molecular weight, polydispersity, and the Mark-Houwink-Sakurada (MHS) parameter (R) for all of the samples (for the parent polymer structures and all samples taken during the incubation experiments for D4, G4, M4, T4, A4, S4, O4, and GO2) were measured with a Viscotek GPCmax instrument equipped with a TSK-GEL GMPWXL column (Montgomeryville, PA) coupled to a triple detection system (static light scattering, viscometry, and refractive index). Mobile phase (0.5 M sodium acetate, pH 5.0) was prepared in ultrapure water. Each sample (100 µL) was dissolved in the mobile phase, immediately injected onto the column, and eluted at 0.8 mL/min. Changes in the molecular weights of the polymers after the incubation were measured with a ViscoGel PCMBLMW-3078 column at a flow rate of 0.8 mL/min. The mobile phase was either ultrapure water (adjusted to pH 4.0 with acetic acid) for the water-incubated samples or water/methanol/acetic acid (55/40/5, v/v/ v) solution for the buffer-incubated samples, to separate the salt peak from the polymer peak. Attenuated Total Reflectance (ATR) FT-IR Spectroscopy. Spectra of the samples (both polymer G4 and polyplexes formed with G4 and pDNA) after the water incubation experiment (at 0 and 72 h) were recorded on a Perkin-Elmer Spectrum One instrument (Wellesley, MA) using attenuated total reflectance (ATR) with a 45° ZnSe crystal with an effective path length of 12 µm at room temperature. Spectra were obtained from 900-3000 cm-1 at 4 cm-1 resolution by accumulating 250 interferograms. Subtraction of the solvent was performed using the water combination mode at 2200 cm-1.35 For analysis of polymer degradation, the concentration of G4 (as a function of repeat unit) was kept constant at 5.0 mg/mL. For the experiment observing G4 degradation in polyplex form (at 0 and 72 h), the DNA (from salmon sperm, Sigma, St. Louis, MO) concentration was 3.2 mg/mL, which corresponds to polyplex formation at an N/P ratio of 5. ObserVation of the Primary Amine Groups Via Trinitrobenzenesulfonate (TNBS). 36 Polymer G4 was incubated in water (0, 12, 24, 48, 72, 120 h) in separate vials for each time point. After the incubation time, each vial containing G4 was diluted to 100 µg/mL using 0.1 M sodium bicarbonate water solution. After the addition of TNBS solution (0.01% w/v in 0.1 M sodium bicarbonate), the samples were incubated for 2 h under 37 °C, and this reaction was stopped with 1 M HCl solution. The absorbance of each sample was measured at 335 nm. PicoGreen Assay of Polyplex Stability. Solutions of G4 (5.0 mM in terms of the polymer repeat unit) were incubated in PBS buffer at 0, 12, 24, 72, and 120 h. After the incubation in PBS buffer, polyplexes were then prepared at N/P ) 5 and 30 and allowed to bind with pDNA at room temperature for 30 min. The subsequent “polyplex” solutions were then diluted with water to a concentration of 0.3 µg DNA/100 µL. PicoGreen (Molecular Probes, Eugene, OR) solutions were prepared freshly by 200-fold dilution with 10 mM HEPES (Sigma, St. Louis, MO). Each polyplex solution (100 µL) was added into a Costar black flat-bottom 96-well plate and then 100 µL of the PicoGreen solution was added to each well. The fluorescence of PicoGreen (excitation 485 nm, emission 535 nm) for each sample was measured with a TECAN

US plate reader (Research Triangle Park, NC). Fractional dye exclusion (indicating the ability for G4 to still bind pDNA) was determined by the following relationship: dye exclusion ) 1 - (Fsample - Fblank)/ (FDNAonly - Fblank). Titration.22 Polymer G4 was dissolved in 5 mL of PBS (pH 7.4) at a concentration of 0.068 M in terms of total concentration of amine groups. The polymer solution was separated into two aliquots. One was titrated immediately and the other one was incubated at 37 °C for 24 h before the titration. The titration of each aliquot was performed with an aqueous solution of HCl (0.1 M standard) in thermostat flasks. The temperature was maintained at 25 °C throughout each titration with a Fisher Scientific isotherm circulator Model 3016 (Pittsburgh, PA). The pH values were recorded with an Accumet pH meter model AB15 (Fisher Scientific, Pittsburgh, PA) equipped with an Orion 8103BN electrode (Thermo Electron Corporation). Nonlinear regression of the experimental data was performed with Graphpad Prism 4.0 (San Diego, CA) according to our previously published procedure.22 III. Cell Culture Experiments. Media and supplements were purchased from Gibco BRL (Gaithersburg, MD). Hela cells were purchased from ATCC (Rockville, MD) and cultured according to ATCC specifications in advanced DMEM in 5% CO2 at 37 °C. The media were supplemented with 10% fetal bovine serum, 100 units/mg penicillin, 100 µg/mL streptomycin, and 0.25 µg/mL amphotericin. Transfections were performed in serum-free media (Opti-MEM) for the first four hours. Untransfected cells and naked pDNA were used as negative controls. JetPEI (Avanti Polar Lipids, Alabaster, AL) and lipofectamine 2000 (Invitrogen, Carlsbad, CA) were used as positive controls at the optimized dosages recommended by the manufacturers [JetPEI: N/P ) 5; lipofectamine 2000: lipofectamine 2000 (µL) to DNA (µg) ratio of 2.5]. Flow Cytometry. Hela cells were seeded on six-well plates at 2.0 × 105 cells/well and allowed to incubate in supplemented DMEM at 37 °C and 5% CO2 for 24 h. The polyplexes were prepared by combining 200 µL of Cy5-labeled pDNA (0.02 µg/µL) with 200 µL of D4, G4, M4, T4, A4, S4, or O4 at the N/P of 20 or with 200 µL of JetPEI or lipofectamine 2000. The cells were then transfected with 0.4 mL of the polyplex solutions (4 µg of pDNA/well) in 1.0 mL of serum-free media (Opti-MEM). The cells were incubated with each solution for 4 h to allow endocytosis and internalization of the polyplexes. After transfection, the cells were rinsed with PBS three times. Supplemented DMEM was then added and the cells were incubated for another 30 min to allow further polyplex internalization. A total of 4.5 h after initial transfection, the cells were trypsinized, pelleted, and resuspended in PBS containing 2% FBS for flow cytometry (FACS) analysis. A FACSCanto II (Becton Dickenson, San Jose, CA) equipped with a helium-neon laser to excite Cy5 (633 nm) was used. A total of 10000 events were collected in duplicate for each sample. The positive fluorescence level was established by visual inspection of the histogram of negative control cells such that less than 1% appeared in the positive region. Luciferase Assay. Delivery vectors (D4, G4, M4, T4, A4, S4, and O4) and pDNA (gWiz-Luc, Aldevron, Fargo, ND) solutions were prepared in DNase and RNase free water. The polyplexes were prepared immediately before transfection by combining solutions of each polymer (150 µL) with pDNA (150 µL, 0.02 µg/µL) at N/P values of 10, 20, and 30. As positive controls, polyplexes formed with JetPEI were prepared in water, and pDNA complexes formed with lipofectamine 2000 were prepared in Opti-MEM, both under optimal conditions for transfection recommended by the manufacturer. The mixtures were incubated for 1 h and diluted to 900 µL with reduced serum media

Poly(glycoamidoamine) DNA Delivery Vehicles (pH 7.2). Cells were cultured at the appropriate density in 24-well plates (at 5.0 × 104 cells/well) and incubated for 24 h prior to polyplex exposure. The cells were transfected with 300 µL of polyplex solution or with naked pDNA in media with reduced serum in triplicate. After 4 h, 800 µL of DMEM was added to each well. The media was replaced 24 h after the transfection with 1 mL of DMEM. Cell lysates were analyzed 48 h after the transfection for luciferase activity with Promega’s luciferase assay reagent (Madison, WI) and for cell viability as described below. For each sample, light units were integrated over 10 s in duplicate with a luminometer (GENios Pro, TECAN US, Research Triangle Park, NC), and the average relative light unit (RLU) value was obtained. Cell Viability. Cell viabilities after being treated with polyplexes of D4, G4, M4, T4, A4, S4, and O4 were measured by the amount of protein in the cell lysates obtained 48 h after the transfection as previously described.2,4 Protein was quantified by Bio-Rad’s DC protein assay (Hercules, CA) against a protein standard curve of bovine serum albumin (Sigma, St. Louis, MO) in cell culture lysis buffer.

Results and Discussion Polymer Synthesis and Characterization. PGAAs D4, G4, M4, and T4 (Figure 1) were synthesized by copolymerization of esterified D-glucaric acid (D), dimethyl-meso-galactarate (G), D-mannaro-1,4:6,3-dilactone (M), and dimethyl L-tartarate (T) with pentaethylenehexamine in methanol, as previously described.2,3,26 The higher MHS R values (0.61-0.73) indicate that the predominant products of the polymerization reaction have a lower degree of branching due to the higher reactivity of primary amines in the aminolysis of esters.37 Also, a low degree of branching off the secondary amines with these particular structures have been observed via NMR with these structures containing four ethyleneamines.3 To study the effects of the hydroxyl groups on the degradation and biological properties of the PGAAs, we synthesized three hydroxyl-free analogues, A4, S4, and O4 (Scheme 1). We attempted to use similar conditions to synthesize polymers A4, S4, and O4 via direct polycondensation of dimethyl adipate, dimetyl succinate, or dimethyl oxalate with pentaethylenehexamine. Polymerization of these monomers, however, did not occur even after stirring the reactants for several months due to the low reactivity of these diesters with amines. With these polyamidoamines, structural differences lead to the distinct activities toward polymerization observed between the hydroxylbearing and hydroxyl-free reactants. In the polycondensation of PGAAs,2,3,26 the presence of electron withdrawing hydroxyl groups on the R,R′-carbons of carbohydrate diesters or dilactones activates the carbonyl groups toward nucleophilic attack by primary amines, which makes the polymerization proceed considerably fast at room temperature.2,3,27,32,38,39 The lack of the electron withdrawing groups near the carbonyls of adipate, succinate, or oxalate comonomers prevents polymerization under these mild conditions.40 To facilitate the polycondensation under similar conditions, adipoyl chloride, succinal chloride, and oxalyl chloride (Scheme 1) were polymerized with an oligoethyleneamine monomer containing Boc-protected secondary amines (to avoid creating highly branched structures). The attachment of the chloride increases the electrophilicity of the carbonyl carbon, which easily reacts with a weak nucleophile such as primary or secondary amines in pentaethylenehexamine. As shown in Scheme 1, after the polymerization, the secondary amine groups were deprotected by a solution of HCl in dioxane.41 To study the effects of secondary amines on the degradation of PGAAs, a model structure, poly(galactaramidotriethylenedioxide) (GO2), was able to be synthesized using similar


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Table 1. MHS Parameter (R), Weight Averaged Molecular Weight (Mw), Polydispersity (Mw/Mn), and Degree of Polymerization (n) for the Polymers Used in This Study polymer

R

Mw (kDa)

Mw/Mn

N

D4 G4 M4 T4 A4 S4 O4 GO2

0.61 0.64 0.73 0.70 0.69 0.71 0.72 0.63

4.9 4.6 5.6 4.8 3.8 3.3 4.5 5.3

1.6 1.5 1.2 1.5 1.7 1.2 2.0 1.3

12 11 14 14 11 11 16 17

procedures as the synthesis of D4, G4, M4, and T4 due to the higher reactivity of the R-hydroxy esters to aminolysis. This polymer model replaces the ethyleneamines in the polymer repeat unit with ethyleneoxide functional groups (Scheme 2), allowing us to study the effects of secondary amines on the degradation and nucleic acid binding properties. We examined the degrees of polymerization and viscosity of D4, G4, M4, T4, A4, S4, O4, and GO2 with GPC. The synthetic conditions for all of these structures were optimized to obtain polymers with similar degrees of polymerization. This allowed us to properly compare the chemical, physical, and biological properties between the analogs. As previously mentioned, the similar high R values (all above 0.6) derived from the Mark-Houwink-Sakurada equation and our GPC/ light scattering/viscometry analysis (Table 1) denoted similar structural features (mostly linear, random-coiled structures) regardless of using pentaethylenehexamine or the Boc-protected analog (compound 1) for the polymerization of the polymers. This assured that the differences in the biological parameters or the degradability were not due to the variations in the degree of polymerization or branching. Degradability of PGAAs. We first used G4, as a representative, to investigate the degradability of PGAAs, and attempted to characterize the degradation products. Polymer G4 was incubated in a variety of conditions: ultrapure water, citric acid-phosphate buffer (pH 5.0), and phosphate buffered saline (PBS, pH 7.4) at 37 °C from 0 to 120 h, and its degradation was assayed by GPC/light scattering/viscometry analysis. These different aqueous media conditions were selected so they simulate the conditions that the polymers may encounter during the gene delivery process. For example, the buffer systems with pH values of 5.0 and 7.4 are approximately the pH environments encountered within the endosomes and the cytoplasm of the cell, respectively. As shown in the GPC traces (Figure 2A), the retention volume of the sample increased as a function of the incubation time, indicating the molecular size of G4 decreased as the incubation time proceeded. This result was confirmed by the simultaneous reduction of viscosity (Figure 2B). Similar patterns of changes were also found in pH 5.0 and 7.4 buffers (changes in RI detector responses occurred for some samples incubated in pH 7.4 buffer, and are shown in the Supporting Information, Figure S1). The degradation of G4 has also been observed via 1H NMR when incubated in D2O under 37 °C (Supporting Information, Figure S2). To characterize the nature of the observed degradation, we employed attenuated total reflectance (ATR) FT-IR spectroscopy to monitor the structural changes of polymer G4 (Figure 3A) at two time points. It should be noted that the degradation was observed in ultrapure water to avoid salt precipitation on the ATR window. Before the incubation, the two strong amide absorptions at 1625 and 1543 cm-1 were seen, which correspond to the CdO stretching vibrations (amide I

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Figure 2. G4 degradation in pure water at 37 °C. (A) Overlay views of GPC refractive index (RI) detector response of G4 after incubation for 0, 12, 24, 48, 72, and 120 h. (B) Overlay views of GPC viscometer (IV) detector response of G4 after incubation for 0, 12, 24, 48, 72, and 120 h.

band) and the N-H bending vibrations (amide II band), respectively (Figure 3A). After the incubation at 37 °C, these two characteristic amide signals decreased and were accompanied by the appearance of two absorption bands at 1581 and 1410 cm-1, which are characteristics of the asymmetric and symmetric stretching bands of the carboxylate group. This result suggested that the amide linkers in the polymer structure were hydrolyzed. A similar result was found when we observed the degradation of polymer G4 when it was bound to DNA in polyplex form (Figure 3B). This indicates that polymer degradation could be a method of DNA release during the transfection process. If the amides in the polymer are being degraded via hydrolysis, carboxylates, and primary amines should be generated. Overlaps of the IR signals for the primary amines (asymmetrical and symmetrical bending absorbs around 16001575 and 1550-1504 cm-1) and the amide or the secondary amines in the polymer prevent direct characterization of the newly formed primary amines after degradation by FTIR. For this reason, we utilized trinitrobenzenesulfonate (TNBS) to observe the increase in the newly formed primary amines during the degradation. The reagent, TNBS, reacts with primary amines (but not secondary amines),42 forming a chromophore that absorbs at 335 nm. In this assay, an increase in absorbance indicates the increase in primary amines (and therefore polymer degradation). The overlay of the UV spectra of TNBSderivatized samples (Figure 3C) demonstrated that the amount of primary amines increased as a function of incubation time. Taken together, our FTIR and TNBS assays indicate that the amide bonds were degraded potentially via a hydrolysis mech-

Liu and Reineke

Figure 3. (A) Attenuated total reflectance (ATR) FT-IR spectra overlay of polymer G4 incubated in water (spectra taken at 0 and 72 h of incubation). (B) Spectra overlay of polymer G4 complexed to DNA in a polyplex incubated in water (spectra taken at 0 and 72 h of incubation). G4 polyplexes were formed with DNA in water at N/P ) 5 prior to incubation. (C) Overlay view of the UV absorption spectra of the incubated polymer G4 after addition of trinitrobenzene sulfonate (TNBS). The inserted figure is the change in the absorbance of the solution at 335 nm as a function of polymer incubation time.

anism during the incubation of the polymer in aqueous media. Similar results were obtained when samples of G4 were incubated in the buffered solutions (data not shown). To further model the degradation of the amide bond, we also synthesized a model compound N,N′-di-(N-methyl ethyleneamine)-galactaramide (Supporting Information, Scheme S2), a representative of the repeated structure in polymer G4. 1H NMR studies of this compound incubated in D2O for 5 days supported our hypothesis that the amide bond cleavage occurs via a hydrolysis mechanism. Discrete signals for mucic acid and N-methyl ethynlenediamine were found after this time point (carboxylates and primary amines are observed, Supporting Information Figure S3). Comparing the time courses for the degradation of G4 in pH 5.0 and 7.4 buffers, we noticed that G4 degraded more rapidly in the basic solution than in the acidic solution (Figure 4A). Similar trends were also observed in the degradation of D4, M4, and T4 (Figure 4B-D). Our previous research has established that the number and the stereochemistry of the hydroxyl groups in the carbohydrate unit and the number of secondary amines in the oligoamine unit, significantly affect many properties of the polymers, such as pDNA binding affinity, polyplex stability in serum, nuclease protection, and buffering capacity.2,3,22–26 It is interesting to note that while all four



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Figure 4. Changes in Mw of polymers (reported as the percent of parent polymer Mw) for (A) G4, (B) D4, (C) M4, and (D) T4 incubated at 37 °C in PBS buffer (pH 7.4), and citric acid-phosphate buffer (pH 5.0). The incubation time points are 0, 12, 24, 48, 72, and 120 h.

structures have similar degradation profiles at pH 7.4, the degradation at pH 5 varies (M4 and T4 degrade much slower). At low pH, more of the amines in these structures are protonated (pH 5.0: D4 ) 48% and T4 ) 55.0%),22 however, at higher pH, a lesser fraction of amines are protonated (pH 7.5: D4 ) 19.1% and T4 ) 21.0%). As we have suggested in our previous publication, this could be related to the polymer secondary structure. Both M4 and T4 polymerize in a stereoregular manner (isotactic) and have a stiffer chain structure in solution (as supported by their higher MHS values, Table 1). In contrast, G4 and D4 are atactic and have a more random coil structure in solution (lower MHS values). This is important because these studies point toward the possible involvement of amine nucleophiles in catalyzing the degradation mechanism (vide infra). The stiffer chain structures (M4 and T4) are not as flexible, which could slow down the catalysis rate. To further characterize the contribution of the secondary amines to PGAA degradation, GO2, a model that contains two ethyleneoxide moieties between the amide bonds (in place of the amines) was created. In a similar experiment to the PGAA analysis, this model was incubated in the same buffer systems and the polymer molecular weight versus time was examined. As shown in Figure 5A, minimal degradation was observed both at pH 5.0 and pH 7.4. After 5 days of incubation, about 81% of GO2 remained intact at pH 5.0, and 89% at pH 7.4 buffer. The incubation of GO2 in ultrapure water was virtually identical to the incubation data in the buffers (data not shown). These data support our hypothesis that the secondary amines somehow facilitate PGAA degradation. To elucidate the contribution of the hydroxyls to PGAA degradation, we created hydroxyl-free PGAA analogues (O4, S4, and A4) to observe whether they degraded in the aqueous media. The O4, S4, and A4 polymer structures have a spacer of 0, 2, and 4 methylenes, respectively, between the two neighboring amide bonds (Scheme 2B), which replace the carbohydrate units. As shown in Figure 5, polymers S4 and A4

also remained intact in the pH 5.0 and pH 7.4 buffers. After 5 days of incubation, 84% of S4 remained intact at pH 7.4, and degradation was not observed at pH 5.0 (Figure 5C). For A4, after 5 days of incubation, the polymer structure remained completely intact at both pH 5.0 and pH 7.4 (Figure 5D). However, for polymer O4, degradation was observed, and contrary to the PGAAs, the hydrolysis appeared faster in the acidic buffer and slower in the basic buffer (Figure 5B). Indeed, the kinetics and mechanism of degradation appeared to be different than the PGAAs, and certainly more rapid than S4 and A4. Comparison of the polymer incubation data for O4, S4, and A4 indicated that, within this group of polymers, the spacer between the neighboring amide bonds significantly affects polymer degradation. In the absence of a methylene spacer, the vicinal oxalyl carbonyls in O4 are likely positioned to exhibit an electronic effect on each other that could facilitate an acidcatalyzed hydrolysis mechanism. For example, under acidic conditions, one of the carbonyls can be protonated, thus, creating a “hydroxyl group” in the R position to the amide bond (creating a similar structure to the PGAAs). These results comparing the hydroxylated PGAAs to their nonhydroxylated and ethyleneoxide analogues suggest that both the amines and hydroxyl groups along the backbone of these polyamides play a synergistic role in facilitating amide degradation. While we currently do not understand the full mechanistic pathway, we speculate that the R hydroxyls vicinal to the amide carbonyl group have an electron-withdrawing effect. Keily et al. have also observed such an effect in similar systems.38 This theory that an R-electron withdrawing group promotes a more electrophilic carbonyl prone to attack is also supported by the degradation profile of O4. In addition, possible inter- and intramolecular H-bonding of the amide group with neighboring hydroxyls (possibly the β-OH) and amines can also create a more electrophilic environment on the carbonyl making it prone to hydrolysis. While these theories are difficult to confirm with such flexible polymeric models, these concepts are supported

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Figure 5. The changes in Mw (reported as the percent of parent polymer Mw) for (A) GO2, (B) O4, (C) S4, and (D) A4 incubated at 37 °C in PBS buffer (pH 7.4), and citric acid-phosphate buffer (pH 5.0). Incubation time points are 0, 12, 24, 48, 72, and 120 h.

Figure 6. Polymer degradation effects on pDNA binding, DNase protection, and buffering capacity. (A) PicoGreen exclusion assays of degraded polymer-pDNA complexes at N/P ) 5 or 30 formed with incubated polymer G4. (B) The calculated protonated fraction of amines vs pH for freshly dissolved G4 (not incubated 0 h) and G4 incubated in PBS for 24 h.

by previous literature examining the hydrolysis rates of various amides. For example, the studies of Menger et al.43 and Hansen et al.44 revealed enzyme-like amide hydrolysis rates in various amide derivatives of Kemp’s triacid. In their models, amide hydrolysis is accelerated under mild conditions in the proximity of neighboring carboxylate groups and catalysis of bond cleavage by nucleophiles. In a related study, Bennet et al. showed in a series of formamide derivatives that distortion of the amide bond toward a pyramidal structure and lengthening of the C-N bond also accelerates amide bond cleavage via basecatalyzed hydrolysis.45 Polymer Degradation Affects pDNA Binding, DNase Protection, and Buffering Capacity. After the characterization of PGAA degradation, we started to analyze how PGAA degradation would affect polymer properties that are vital to gene delivery, using G4 as a model compound. A thorough and quantitative examination of the binding mechanisms of the polymers studied herein is provided in the proceeding manuscript.46 First, we studied how polymer degradation would affect the ability of forming polyplexes via the PicoGreen exclusion assay.22 PicoGreen is a dye that intercalates into DNA and

exhibits enhanced fluorescence intensity upon binding. If a species, such as a polycation, binds to DNA and displaces the dye from intercalation, the fluorescence intensity is greatly reduced, thus giving a qualitative assessment of polymer-DNA binding. To qualitatively examine the binding of the PGAAs over time (and the effect of polymer degradation on pDNA binding), G4 was incubated in aqueous solution at 37 °C, and at various time points, the polymer was combined with pDNA to form polyplexes with pDNA at N/P ) 5 or 30. As shown in Figure 6A, it was noticed that, as the polymer incubation time was increased, more PicoGreen intercalated into the pDNA (at both N/P values this is shown as a decrease in PicoGreen dye exclusion as a function of G4 incubation time). These results support our previous data and our hypothesis that polymer hydrolysis decreases the stability of the resulting polyplexes and could promote pDNA release. Next, we explored how hydrolysis would affect the buffering capacity of the polymeric vehicle by titrating samples of G4 before and after incubation at 37 °C for 24 h. The buffering capacity of polymeric nucleic acid delivery vehicles has been hypothesized to be an important parameter in the delivery


efficacy as studies have linked this property to endosomal release and cytosolic entry of nucleic acids via the “proton sponge hypothesis”.22,47 The titration and its nonlinear fitting were performed according to our previously published procedures.22,48 The titration data of the parent G4 polymer structure (prior to incubation/hydrolysis), revealed that 29.1% of the amines were protonated at pH 7.5 and 63.9% of the amines were protonated at pH 4.5 (Figure 6B, solid line), yielding a buffering capacity of 34.8%. When comparing these results to our previously published data (discussed earlier) with D4 and T4,22 this reveals that G4 has a much higher protonated fraction of amines at both pH values and a slightly higher buffering capacity (buffering capacities of D4 ) 29.2% and T4 ) 34.0%). The higher prontonated charge and buffering capacity could be related to the high pDNA binding affinity and delivery efficiency, respectively, that we commonly observe with G4.2,23,25 However, it should be noted here that batch to batch differences in the polymers (that vary in length and degree of branching) can cause differences in the data. After incubating G4 in PBS for 24 h, the protonated fraction of degraded G4 was determined to be 25.6% at pH 7.5 and 65.2% at pH 4.5 (Figure 6B, dotted line). The calculated buffering capacity of the G4 sample after the incubation was about 40%, which is higher than that of the nonincubated counterpart and supports our previous data (Figure 3C) that degradation is occurring via hydrolysis (and some of the parent primary amine monomers could be regained after degradation). These data suggest that, even if degradation of the PGAAs would occur in lysosomes, the buffering capacity would not be reduced and release of polyplexes/pDNA could still be promoted. Gene Delivery and Cell Viability. Comparing the PGAAs (D4, G4, M4, and T4) to their hydroxyl-free analogues (A4, S4, and O4) provides a unique platform to study how structural and chemical properties, such as the degradation profiles, influence cellular uptake, cytotoxicity, and gene delivery efficacy. In this study, JetPEI and lipofectamine 2000, two commercially available gene delivery vectors, as well as pDNA alone, were used as the positive and negative controls, respectively. The cellular uptake of Cy5-labeled pDNA facilitated by these polyamidoamines was investigated using flow cytometry. As shown in Figure 7A, all of our polymers facilitated cellular uptake of pDNA as nearly 100% of cells exposed to the synthetic polymers were positive to Cy5 pDNA. When the number of Cy5 positive cells was compared, all the PGAAs and their analogues performed as efficiently as lipofectamine 2000, and the Cy5-polyplexes revealed higher cellular entry than JetPEI. When the average amount of pDNA taken up by each cell was compared, all the polymers yielded higher efficiency than JetPEI, and G4 was about equivalent to lipofectamine 2000. Comparison among PGAAs (D4, G4, M4, and T4) and their hydroxyl-free analogues (A4, S4, and O4), at the current degrees of polymerization (Table 1), showed that G4 was the most effective synthetic vector in Hela cells, and then vector T4, which supports our previous data.2,23,26 When the results were compared within the hydroxyl-free analogues, polymer A4 revealed the highest and S4 revealed the lowest uptake with Hela cells. Also, when comparing the PGAAs to the hydroxylfree derivatives, there was not a clear trend. It should be mentioned that GO2 was not able to bind pDNA and form polyplexes (likely due to the lack of positive charge in the polymer). Subsequently, this structure was completely inactive to deliver pDNA into cultured cells (data not shown).


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Figure 7. Biological evaluation of polyplexes formed with D4, G4, M4, T4, A4, S4, and O4 in Hela cells. Uncomplexed pDNA was used as a negative control. JetPEI (N/P ) 5) and lipofectamine 2000 (µL to µg of DNA ratio is 2.5) were used as positive controls. The data are reported as a mean ( standard of deviation of two replicates for A and three replicates for B and C. (A) Mean fluorescence intensity (bars) and the percentage of cells positive for Cy5 (line) for Hela cells transfected for polyplexes formed with Cy5-labeled pDNA and D4, G4, M4, T4, A4, S4, and O4 at N/P ) 20. (B) The fraction of cell survival of Hela cells transfected with polyplexes formed with pDNA and D4, G4, M4, T4, A4, S4, and O4 at N/P ) 20. The cell viability values are normalized to untransfected cells (fraction of cell survival ) 1.0). (C) Luciferase gene expression observed with polyplexes formed with pDNA and D4, G4, M4, T4, A4, S4, and O4 at N/P ratios of 10, 20, and 30. The gene expression values are shown as relative light units (RLU) per mg of protein. It should be noted here that GO2 did not transfect cells.

Cell viability profiles were examined after the pDNA delivery with these polyamidoamines by measuring the total protein concentration in the cell lysates and normalizing the data to protein level of the untransfected cells. On average, at an N/P ratio of 20, most of the synthetic structures promoted 80% or greater viability which was much higher than that observed for JetPEI (55%) and lipofectamine 2000 (31%; Figure 7B). When the PGAAs were compared to their nonhydroxylated analogues, we were surprised to find that the cell viability profiles were similar even though A4 and S4 do not degrade. Many literature reports have stressed that degradation of gene delivery vectors results in lower cytotoxicity.11–13 There are two possibilities to reconcile the discrepancy (at the current concentrations of transfection) between our observation and the literature. All our polyamines are comparatively short, on average containing 11-16 repeat units, which translates to an average molecular weight between 4 to 6 kDa (Table 1). With the current structures,

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this could be the dominant factor leading to low cytotoxicity. At these low molecular weights, the advantage of degradability toward reducing cytotoxicity is not observed with our systems at the current concentrations. The synthesis of longer PGAAs is ongoing in our lab; comparing longer analogues may provide us with appropriate means to examine the role of PGAA degradation on cytotoxicity. Another possibility is that polymers A4 and S4 degrade by cellular proteinase due to the presence of an amide linkage (peptide linkage) in the backbone. Enzymatic hydrolysis may contribute to their low cytotoxicity. To complement these cellular studies, we investigated the gene expression efficiency promoted by these analogous vectors using a luciferase reporter gene assay. Generally, for all the examined polymers, the luciferase expression level increased as the N/P value was increased (Figure 7C). At N/P ) 30, all polyamidoamines, with the exception of A4 and S4, led to similar levels of luciferase expression to that observed with JetPEI and lipofectamine 2000. Within the group of hydroxylfree polyamidoamines, it is obvious that O4 rendered higher luciferase expression than A4 and S4 at all N/P values. It is interesting that the nondegradable polymers (A4 and S4) yielded low gene expression. When the comparison was made between PGAAs (D4, G4, M4, and T4) and their hydroxyl-free analogues (A4 and S4), a similar pattern was also observed, indicating vector degradation and thus DNA release is important for enhancing gene expression. These observations support other studies in this field that polymer degradation leads to higher transfection efficiency by promoting pDNA release.14–21 A balanced view, however, should be held in the examination of the biological significance of degradable nucleic acid delivery vectors. In addition to the above-mentioned advantage, degradation of the delivery vectors also exposes the DNA to nucleases, which could be resulting in nucleic acid degradation prior to reaching the intracellular target. Given that successful gene delivery requires trafficking pDNA through multiple cellular compartments (endosome, lysosome, and nucleus) in a wellorchestrated manner, if degradation is to be used as a functional module to control DNA release, more in-depth investigations on the timing and location of delivery vector degradation are necessary for a better control on the gene delivery process. Interestingly, PGAAs (D4, G4, M4, and T4) followed a faster degradation at pH 7.4 (cytosolic pH) than pH 5.0 (lysosomal pH; Figure 4), suggesting that their in vitro degradation could occur in the cytosol or nucleus rather than in the lysosomes. However, O4 is more easily degraded at lysosomal pH than at cytosolic pH (Figure 5B), as this polymer favors an acidcatalyzed degradation mechanism. These preliminary but promising results hint that through simple structural modifications, it may be possible to control the compartment-dependent behavior of gene delivery vectors to promote pDNA release in a controlled fashion.

Conclusion We have investigated the degradation of PGAAs in this study, using GPC, ATR FT-IR, and UV absorbance techniques. The results indicate that the secondary amines and hydroxyls in the polymer backbones are necessary to facilitate rapid degradation. For this reason, the ideal storage condition for these polymers is as a lyophilized powder in desiccated conditions at low temperature. The polymer can be easily reconstituted in water immediately prior to polyplex formation and transfection. Titration experiments show that PGAA degradation does not affect the polymer buffering capacity. However, the PGAA

Liu and Reineke

degradation may enhance gene expression by releasing pDNA from polyplexes (polymer-pDNA complexes), thus exposing it to the intracellular transcriptional machinery. The difference in the optimal pH that promotes degradation of the PGAAs and the hydroxyl-free analogues may prove to be a useful means to achieve pH-regulated DNA release from polyplexes by specifically modulating the chemical structures. Acknowledgment. The funding of this work was provided by the NSF CAREER (CHE-0449774) and the Beckman Young Investigators programs. T.M.R. is a Fellow of the Alfred P. Sloan Foundation. Y.L. sincerely thanks Dr. Yumin Chen, Dr. Josh Bryson, and Antons Sizovs for helpful discussions. Supporting Information Available. Overlay view of the GPC refractive index response of the polymer degradation in buffer and the NMR of the degraded polymer. This material is available free of charge via the Internet at http://pubs.acs.org.

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Degradation of Poly (glycoamidoamine) DNA Delivery Vehicles

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