Self-Catalyzed Degradable Cationic Polymer for Release of DNA

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Self-Catalyzed Degradable Cationic Polymer for Release of DNA Nghia P. Truong,† Zhongfan Jia,† Melinda Burgess,‡ Liz Payne,‡ Nigel A. J. McMillan,‡ and Michael J. Monteiro*,† †

Australian Institute for Bioengineering and Nanotechnology and ‡Diamantina Institute for Cancer, Immunology, Metabolic Medicine, The University of Queensland, Brisbane QLD 4072, Australia

bS Supporting Information ABSTRACT: The controlled release of siRNA or DNA complexes from cationic polymers is an important parameter design in polymer-based delivery carriers. In this work, we use the selfcatalyzed degradable poly(2-dimethylaminoethyl acrylate) (PDMAEA) to strongly bind, protect, and then release oligo DNA (a mimic for siRNA) without the need for a cellular or external trigger. This self-catalyzed hydrolysis process of PDMAEA forms poly(acrylic acid) and N,N0 -dimethylamino ethyl ethanol, both of which have little or no toxicity to cells, and offers the advantage of little or no toxicity to off-target cells and tissues. We found that PDMAEA makes an ideal component of a delivery carrier by protecting the oligo DNA for a sufficiently long period of time to transfect most cells (80% transfection after 4 h) and then has the capacity to release the DNA inside the cells after ∼10 h. The PDMAEA formed large nanoparticle complexes with oligo DNA of ∼400 nm that protected the oligo DNA from DNase in serum. The nanoparticle complexes showed no toxicity for all molecular weights at a nitrogen/phosphorus (N/P) ratio of 10. Only the higher molecular weight polymers at very high N/P ratios of 200 showed significant levels of cytotoxicity. These attributes make PDMAEA a promising candidate as a component in the design of a gene delivery carrier without the concern about accumulated toxicity of nanoparticles in the human body after multiadministration, an issue that has become increasingly more important.

’ INTRODUCTION Small interfering RNA (siRNA) holds great potential in the cure of cancers and infectious diseases.1 The siRNA oligonucelotide, a double-stranded RNA consisting of between 19 and 23 nucelotides in length, silences gene expression with high efficiency via the RNA interference mechanism.2 For siRNA-based delivery carriers to reach their full potential, they must be protected from nucleases, have highly efficient cellular uptake, target specific cells and tissues, and have a predetermined and reproducible pharmacokinetics.3 A promising way to achieve successful delivery of siRNA is through the use of either viral or nonviral delivery carriers.4 Viral delivery carriers, although often effective at gene expression knockdown, can induce insertional activation or interruption of genes as well as immunologic responses, making them a safety concern.5 In contrast, nonviral delivery carriers such as lipids, peptides, and polymers are believed to be safer, cheaper, and easier to produce.6 Polycation nanoparticle delivery systems must be designed to package the siRNA into the nanoparticle efficiently and provide serum stability, specific targeting, cell uptake, endolysosomal escape and release of the siRNA within the cell.7 The release of siRNA from the polycation within the cell is a critical step in the silencing process because it allows the siRNA to interact with Dicer and form the RNA-interfering silencing complex (RISC).8 However, the fact that polycations have a high positive charge on the polymer backbone to encapsulate the negatively charged r 2011 American Chemical Society

siRNA makes the release difficult.9 To overcome this difficulty, many polymers have been designed to release via a trigger, including temperature,10 pH,11 redox potential,12 light,13 electric pulse,14 enzymatic degradation,15 and salt levels.16 For example, the incorporation of disulfide linkages in the side chains of different polymers provides, through the intracellular reduction of the disulfide by glutathione and thioredoxin, a trigger release pathway of siRNA into the cytoplasm.17 Some polycations modified with disulfide linkages such as polyethyleneimine (PEI),18 poly(2-dimethylaminoethyl methacrylate),19 and poly(amidoamine)20 have shown high knockdown efficiency but with high toxicity. Shim et al.21 released siRNA using pH (99.5%), and dimethyl sulfoxide (DMSO, >99.9%) were used as received from Sigma-Aldrich. n-Hexane (>98%) and tetrahydrofuran (THF, >99.9%) were used as received from Merck. 2-(Dimethylamino) ethyl acrylate (98%, Sigma-Aldrich) was passed through a column of basic alumina (activity I) to remove the inhibitor. Azobisisobutyronitrile (AIBN) was recrystallized twice from methanol prior to use. Milli-Q Water (18.2 MΩcm1) was generated using a Millipore Milli-Q academic water purification system. 9-27 Oligo DNA was synthesized by Invitrogen (Carlsbad, California), 9-27F+RMW=14998 (23bp),

sense: 50 -GTCAGAAATAGAAACTGGTCATC-30 antisense: 50 -GATGACCAGTTTCTATTTCTGAC30 . Oligofectamine, a standard transfection reagent, was purchased from Invitrogen. All other chemicals and solvents were of at least analytical grade and used as received. Methods. 1H and 13C Nuclear Magnetic Resonance. All nuclear magnetic resonance (NMR) spectra were recorded on a Bruker DRX 500 MHz spectrometer using an external lock (CDCl3 and D2O) and referenced to the residual nondeuterated solvent (CHCl3 and H2O). Size Exclusion Chromatography (SEC). The molecular weight distributions of the polymers was determined using a Waters 2695 separations module, fitted with a Waters 410 refractive index (RI) detector maintained at 35 °C, a Waters 996 photodiode array detector, and two Ultrastyragel linear columns (7.8 300 mm) arranged in series. These columns were maintained at 40 °C for all analyses and are capable of separating polymers in the molecular weight range of 500 to 4 million g/mol with high resolution. All samples were eluted at a flow rate of 1.0 mL/min. Calibration was performed using narrow molecular weight polystyrene (PSTY) standards (PDI e 1.1) ranging from 500 to 2 million g/mol. Data acquisition was performed using Empower software, and molecular weights were calculated relative to polystyrene standards. Dynamic Light Scattering. DLS measurements were performed using a Malvern Zetasizer 3000HS. The sample RI was set at 1.59 for polystyrene. The dispersant viscosity and RI were set to 0.89 and 0.89 Ns/m2, respectively. The number-average particle diameter was measured five times for each sample. Naked oligo DNA (9-27) (2.0 μg) was mixed with polymers at different N/P ratios (10 to 200) in a total volume of 50 μL of water and allowed to complex for 30 min at room temperature. We loaded 40 μL of samples into a microcuvette for measurements. Matrix-Assisted Laser Desorption Ionization Time of Flight (MALDITOF) Mass Spectrometry. Spectra were performed using a Bruker Autoflex III Smartbeam operated in both linear and reflectron mode. Ions were accelerated at a potential of 20 kV with a nitrogen laser emitting at 337 nm.

Synthesis of Poly(2-(dimethylamino)ethyl acrylate) (PDMAEA). All five PDMAEA samples (series A in Table 1 and Scheme 1) were synthesized by reversible additionfragmentation chain transfer (RAFT) polymerization using 2-(butylthiocarbonothioylthio) propanoate (MCEBTTC) as a chain transfer agent.23 PDMAEA with different molecular weights was obtained by changing the ratio of MCEBTTC to the monomer. For a typical polymerization, DMAEA (1 mL, 6.6 103 mol), MCEBTTC (16.6 mg, 6.6 105 mol), and AIBN (1.08 mg, 3541

dx.doi.org/10.1021/bm2007423 |Biomacromolecules 2011, 12, 3540–3548

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Table 1. SEC, 1H NMR, and MALDI-TOF Data for the RAFT Polymerization of DMAEA to Produce Different Molecular Weight (A1A5) at 60 °C in 7 h in Dioxane SECa

H NMRb

1

polymer code [M]:[RAFT]:[I] M (g/mol) PDI M (g/mol) repeating units (n) n n

MALDI-TOF m/z (Δm/z) (amu)c conv. (%)d Mn,theory (g/mol)e

A1

300:10:1

3000

1.26

3300

21

3116.8 (0.1)

77

3555

A2

500:10:1

4000

1.24

4800

32

3546.3 (0.1)

69

5185

A3 A4

700:10:1 900:10:1

5600 6900

1.17 1.20

6100 9700

41 66

6267.9 (0.8) 6696.7 (0.3)

63 59

6558 7845

A5

1000:10:1

8600

1.26

11300

77

8557.8 (0.1)

61

8975

a

SEC data measured in THF solution and using PSTY standards for calibration. b Molecular weight and the number of repeating units determined by 1H NMR (calculated from the integral area of two peaks at 0.9 ppm and 4.1 ppm using the following equation: Mn = (I4.1 3 3/I0.9 3 2) 3 143 + 252). c Molecular weight at the highest visible peak from MALDI-TOF spectra. d Conversion was checked by the gravity method (i.e., by measuring the dry weight of polymer). e Mn theory was calculated from following equation: Mn = conversion 3 ([M]/[RAFT]) 3 143 + 252. 6.6 106 mol) were dissolved in 1 mL of dioxane. The mixture was deoxygenated by purging with Argon for 30 min and then heated to 60 °C for 7 h. The reaction was stopped by cooling to 0 °C and exposure to air. The mixture was precipitated in a large excess of cold n-hexane and then isolated by centrifugation. The operation of redissolving and precipitating was repeated three times in dioxane. The polymer was dried under high vacuum for 48 h at room temperature to give a yellow oily product. The polymers used for this work were named as A1A5, representing an increase in molecular weight. (See Table 1 and Scheme 1.) Quaternization of PDMAEA. All five different molecular weights of PDMAEA (A1A5) were modified by iodomethane to give five quaternized PDMAEA, respectively (denoted as B1B5). For a typical quaternization, PDMAEA (0.286 g, 2.0 mmol of DMAEA repeating units) was dissolved in 2 mL of dioxane. Iodomethane (0.34 g, 2.4 mmol) was added dropwise over 5 min at room temperature. The reaction mixture was diluted and dialyzed using brine over a 48 h period and then Milli-Q water for another 48 h using dialysis tubing with MWCO 3500 Da. After dialysis, the water was removed by freeze-drying to give a yellow solid. Binding Assay. Oligo DNA 9-27 (1.0 μg) was complexed with polymers at different nitrogen-to-phosphorus (N/P) ratios (ranging between 10 and 200) in a total volume of 25 μL of water and allowed to complex without stirring for 30 min at room temperature. Controls included oligo DNA without polymer and oligo DNA complexed with oligofectamine according to the manufacturer’s instruction. The complexed polymer/oligo DNA (25 μL) was mixed with 5 μL of DNA loading dye, loaded into a 2% agarose gel containing TAE buffer, and stained with ethidium bromide (Sigma, Sydney, Australia). The gels were run in 1 3 TAE buffer for ∼10 min at 80 V before being visualized using a Bio-Rad UV transilluminator. Protection Assay. Oligo DNA (2 μg) was complexed with polymers at different N/P ratios (10 to 200 as indicated) in a total volume of 100 μL of water, and the mixtures were allowed to complex for 30 min at room temperature. DNase (1U, NEB, Boston) with 15 μL of 10 DNase buffer and 34.5 μL of H20 were added to each sample without mixing, and the samples were incubated at 37 °C for 10 min. To extract DNA from the polymers, a mixture of phenol/chloroform (200 μL of a 1/1 ratio) was added to each tube, and the samples were vortexed. Tubes were spun at 12 000 rpm for 10 min at 4 °C in a cold room before the aqueous layer was removed. Aqueous layer (5 μL) was mixed with 5 μL of DNA loading dye, samples were loaded into 12% nondenaturing polyarylamide gel containing 1 TBE buffer, and the gel was run in 1 TBE for ∼1 h at 100 V. The gels were stained in ethidium bromide (10 μg/mL) for 10 min and then visualized by using a Bio-Rad UV transilluminator. Transfection Assay. HeLa cells (5 104) were seeded into 24 well plates for 24 h before transfection. FITC-labeled Oligo DNA (0.533 μg,

4 1011 mol, Sigma, Sydney) was complexed with polymers at different N/P ratios (10 to 200, as indicated) in a total volume of 150 μL of water, and the mixtures were allowed to complex without stirring for 30 min at room temperature. OptiMem (Invitrogen, Sydney, Australia) was added to a final volume of 1 mL before polymer/FITC-oligo DNA complexes were added to the cells (250 μL per well). The cells were incubated at 37 °C for 4 h before being washed with 1 mL of sterile PBS, harvested using trypsin-EDTA (Invitrogen, Sydney), collected by centrifugation at 400g for 5 min at 4 °C, resuspended in PBS, and centrifuged again as described above. Supernatant was removed, and cells were resuspended in 1 mL of 2% paraformaldehyde. Uptake was examined on cells by analyzing samples on FACSCanto (BD), and data were analyzed using ACSDiVa software (BD). Cytotoxicity Assay. HeLa cells (1 104) were seeded into 96-well plates for 24 h before transfection. Control wells containing media without cells were used to obtain a value for background luminescence. Oligo DNA (0.533 μg, 4 1011 mol) was mixed with polymers at different N/P ratios (10 to 200) in a total volume of 150 μL of water and allowed to complex for 30 min at room temperature before OptiMem (Invitrogen, Sydney, Australia) was added to a final volume of 1 mL. Oligofectamine (Invitrogen) was used as the positive control. Polymer/ oligo DNA complexes were added to the cells (100 μL per well), and plates were incubated at 37 °C for 4 h before complexes were removed, cells were washed with PBS, and 150 μL complete DMEM (Gibco) was added to each well. Cell Titer Glo (Promega, Sydney, NSW, Australia) cell viability assay was subsequently performed according to the manufacturer’s instructions.

’ RESULTS AND DISCUSSION We used the RAFT polymerization technique to synthesize PDMAEA of five different molecular weights (series A in Table 1).23 The narrow molecular weight distributions (PDI values < 1.26) demonstrated the controlled behavior offered by the RAFT polymerization. The PDMAEA can be readily converted from its partial cationic charge at neutral pH to a permanent cationic charge through a reaction with iodomethane denoted as series B. (See Scheme 1.) Both series were subsequently tested and compared for binding, protection, and transfection of an oligo DNA (to mimic siRNA) to HeLa cells. The cytotoxicity was also tested for the polymer/DNA nanoparticle complexes. Polymer/DNA Binding. The ability of polycations to condense DNA to form nanoparticles plays an important role for successful polymer-based gene delivery. To determine the size of the complexes in water between the oligo DNA and polymers (Series A and B), we used a concentration of 40 μg/mL of oligo 3542


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Table 2. Size of PDMAEA (A1A5)/Oligo DNA and Quaterized PDMAEA (B1B5)/Oligo DNA Complexed in Water at Different N/P Ratios (10, 100, and 200)a hydrodynamic diameter, Dh (nm) (PDI in parentheses) N/P ratio

a

A1

A2

A3

A4 75.24 (0.28)

A5

B1

10

435.20 (0.19) 135.24 (0.13) 295.40 (0.17)

100

457.81 (0.19) 437.25 (0.54) 540.25 (0.44) 430.10 (0.42) 497.04 (0.39) 25.2 (0.45)

200

616.68 (0.28) 440.25 (0.48) 479.17 (0.45) 589.6 (0.43)

B2

B3

B4

B5

94.09 (0.16) 37.09 (0.37) 49.13 (0.27) 25.78 (0.31) 48.64 (0.29) 45.72 (0.30) 648.8 (0.37)

31.09 (0.41) 34.9 (0.46)

28.08 (0.38) 30.31 (0.44)

31.27 (0.13) 29.85 (0.54) 33.82 (0.49) 33.12 (0.32) 46.3 (0.32)

DNA concentration is 40 μg/mL. Dh data is reported as an average of five number percentage measurements.

Figure 1. Agarose gel retardation assays of PDMAEA (A1A5)/Oligo DNA polyplexes (A) after incubation for 30 min and (B) after storage for 1 week and quanternized PDMAEA (B1B5)/Oligo DNA polyplexes (C) after complexation for 30 min and (D) after storage for 1 week; picture (a) N/P ratio 10, (b) N/P ratio 100, and (c) N/P ratio 200.

DNA (9-27), which was far greater than those concentration used in the in vitro experiments. At lower concentrations that would replicate in vitro conditions, the DLS generated poor correlograms leading to highly inaccurate and irreproducible sizes. The sizes of the complexes shown in Table 2 using DLS (see the Supporting Information for particle size distributions based on the distribution function) under the appropriate conditions were all >400 nm for the A series at nitrogen-to-phosphorus (N/P)

ratios of 100 and 200. The greater the N/P ratio, the greater the ratio of polymer to oligo DNA. At an N/P ratio of 10, the sizes were smaller, especially at the higher molecular weights (i.e., A4 and A5). The B series resulted in quite small sizes between 25 to 50 nm, presumably due to the permanent and thus higher cationic charge on the series B compared with series A polymers. These sizes and size distributions, however, could be different at concentrations used in vitro studies. 3543


Biomacromolecules The formation of these nanoparticles does not provide insight into the efficiency of binding between the polymer and oligo DNA. To investigate the binding capacity, we performed agarose gel retardation assays for PDMAEA (A1A5) and quanternized PDMAEA (B1B5) complexed with oligo DNA (9-27) at different N/P ratios. The oligo DNA will not enter the gel while bound in a nanoparticle complex with PDMAEA. The results demonstrate the strong binding with all of the different molecular

Figure 2. Polyacrymide gel retardation assays of polymers/oligo DNA polyplexes at an N/P ratio of 10 incubated with DNase for 10 min, followed by washing DNase out and treatment with a phenol/chloroform mixture: Series A polymer/oligo DNA nanoparticle complexes (A) after complexation for 30 min and (B) after storage for 1 week and series B polymer/oligo DNA nanoparticle complexes (C) after complexation for 30 min and (D) after storage for 1 week.

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weight PDMAEA (i.e., A1A5) and at all N/P ratios tested (10, 100, and 200), as shown by the top band in Figure 1A. The smearing observed in some of the lanes most probably results from incomplete binding of the polymer with the oligo DNA. Even the very low-molecular-weight PDMAEA (A1 and A2) with 20 to 30 monomer repeating units produced stable oligo DNA/ polymer complex nanoparticles. After 1 week of incubation at room temperature, the gel showed the loss of the initial top band and the formation of a new band corresponding to free 9-27 oligo DNA (Figure 1B), indicating the full dissociation (release) of the oligo DNA from the polymer. This release mechanism for the oligo DNA was consistent with the self-catalyzed hydrolysis of PDMAEA in water to form the negatively charged poly(acrylic acid) and N,N0 -dimethylamino ethyl ethanol.23 The quaternized PDMAEA (series B in Scheme 1) binds strongly to the oligo DNA immediately after complexation (Figure 1C) and after 1 week of incubation (Figure 1D). The nonrelease behavior of series B polymers was consistent with its nondegradable properties.23 These results clearly demonstrate the unique ability of PDMAEA (series A) to bind oligo DNA strongly and release it over time through a self-catalyzed hydrolysis of the polymer. The hydrolysis of this polymer was previously found to be independent of pH, salt concentration, or any other conventional trigger stimulus.23 In addition, PDMAEA alone with molecular weights of 5600 and lower gave no cytotoxicty to HeLa cells even at the highest N/P ratio of 200. This previous study also showed that in contrast the permanent cationic polymers (series B) alone showed no hydrolysis and high levels of toxicity, especially at higher molecular weights. Oligo DNA Protection by PDMAEA. The addition of DNase to the nanoparticle mixture assesses the level of protection that the nanoparticles provide to the oligo DNA. DNase will degrade siRNA through the hydrolytic cleavage of phosphodiester linkages in the DNA backbone, resulting in a loss of the oligo DNA gel band, as observed in Figure 2. The protection assay was as follows: the nanoparticle complexes were incubated in the presence of DNase for 10 min at 37 °C; then, the nondegraded oligo DNA was extracted from the polymer in a phenol/chloroform mixture and centrifuged, followed by qualitatively determining the amount of oligo DNA by gel electrophoresis.

Figure 3. Agarose gel retardation assays of Series A polymer/oligo DNA nanoparticle complexes stored for (A) 30 min, (B) 1 day, (C) 2 days, and (D) 3 days at different N/P ratios from 10 to 200. 3544


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Figure 4. Time-dependent particle size of PDMAEA (A5)/FITC-oligo DNA (-9-) and quaternized PDMAEA (B5)/FITC-Oligo DNA (-b(red)) polyplexes at N/P ratio 10. The data are reported as the mean ( standard of deviation of five measurements.

The gels (in Figure 2A) for the nanoparticle complexes (at an N/P ratio of 10) tested immediately after mixing oligo DNA and polymer to form the nanoparticle complex showed excellent protection of the oligo DNA by the polymer for all molecular weights (A1A5). The same result was found at an N/P ratio of 100 and 200 (results not shown). In contrast, naked DNA (9-27+DNase) showed no visible band due to full digestion by the DNase. Storage of the nanoparticle complex for 1 week showed no oligo DNA band on exposure to DNase. This supported the binding studies above that after 1 week the PDMAEA degraded, releasing the oligo DNA into the medium, followed by digestion by the DNase. In contrast, the permanently cationic PDMAEA (B1B5) showed protection of oligo DNA both immediately after the complexation (Figure 2C) and even after 1 week of incubation (Figure 2D). These results highlight the effect of the self-catalyzed degradation of PDMAEA on the disruption of the PDMAEA/ oligo DNA nanoparticles to expose or release the oligo DNA in serum for digestion. Dissociation of Polymer-Oligo DNA Nanoparticle Complexes. For delivery of the siRNA or DNA payload, the nanoparticle complexes have to traffic into the target cells before they degrade and release siRNA or DNA. The data above for our nanoparticle complexes showed release of oligo DNA after 1 week. In the next set of experiments, we analyzed using the gel retardation assay the dissociation of the nanoparticle complexes over 3 days (Figure 3). The nanoparticle complexes consisting of the low-molecular-weight PDMAEA (A1A3) fully dissociated at all N/P ratios within the first 24 h, whereas the higher molecular weight PDMAEA (A4 and A5) showed partial dissociation after 24 h and full dissociation after 2 days. This indicates that for the A4 and A5 polymer complexes there were still some weak interactions between PDMAEA and oligo DNA. We next studied the influence of the oligo DNA release on the size of the nanoparticle complexes using A5 and B5 as the polymers. The DLS data showed that the initial size of the nanoparticle complex using the A5 polymer was close to 100 nm, which remained constant for 10 h (Figure 4). After 10 h, the size decreased dramatically to ∼5 nm, corresponding to the size of the free polymer in solution. However, the size of the nanoparticle

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Figure 5. FITC transfection efficiency of PDMAEA (A5)/Oligo DNA polyplexes at N/P ratio 10 to HeLa cells in pH 7.4 buffer solution with different incubation time; controls used are naked FITC-oligo (untreated) and FITC-Oligo/Oligofectamine complex (Oligofectamine) incubated with cells in 4 h. Concentration of oligo DNA is 40 nM. The data are reported as the mean ( standard of deviation of three replicates.

complex using the B5 polymer remained undissociated at 50 nm, even well past 76 h. These results indicated release of the siRNA from the complex after ∼10 h. This result suggests that for the successful delivery of the siRNA into the cells through complexation with PDMAEA, the complexes should transfect into the cells within the first 10 h before any significant release. The release in 10 h is much faster than the full degradation of all side chains on PDMAEA, suggesting that only a fraction of the side chains need to be converted to the negatively charged poly(acrylic acid) to induce a release. In Vitro Transfection. We next determined whether the PDMAEA nanoparticle complexes could transfect into HeLa cells before they dissociated (i.e., before 10 h). A cell-uptake assay using FITC-labeled oligo DNA (23 bp in length)/ PDMAEA (A5) nanoparticle complexes (N/P = 10) over a 4 h period was analyzed by flow cytometry (Figure 5). The transfection efficiency was close to zero after an incubation time of 1 h. Increasing the incubation time from 2 to 4 h showed a significant increase in transfection from 10 to 90%, respectively, suggesting that at 4 h nearly all cells were transfected with the nanoparticle complexes. Transfection after a 4 h incubation period was further carried out at N/P ratios between 10 to 200 for all series A and B polymer nanoparticle complexes with FITC-oligo DNA (Figure 6). Two sets of experiments were carried out: (1) the polymer and DNA were mixed and stored for 30 min and then incubated with the HeLa cells for 4 h (series A and B in Figure 6A,C, respectively) and (2) the polymer and DNA were mixed and stored for 1 week and then incubated with the HeLa cells for 4 h (series A and B in Figure 6B,D, respectively). Figure 6A shows the very high transfection for A4 and A5 (>85%). Nanoparticle complexes using A1A3 showed transfection between 10 and 30%, suggesting the higher molecular weight PDMAEA (>5600) gave a significant increase in transfection. Increasing the N/P ratio to 200 showed an increase in the transfection for the low-molecularweight polymers (A1A3) and actually a small drop in transfection for A4 and A5 nanoparticle complexes. Incubation of the A series 3545


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Figure 6. FITC transfection assays of polymers/FITC-Oligo DNA polyplexes incubated with HeLa cells at various N/P ratios (10, 100, 200) in buffer solution pH 7.4 for 4 h. PDMAEA (A1A5)/FITC-oligo DNA (A) kept for 30 min after complexation and (B) kept for 1 week after complexation and quanternized PDMAEA (B1B5)/oligo DNA polyplexes mixture (C) kept for 30 min after complexation and (D) kept for 1 week after complexation. Controls used are naked FITC-oligo (untreated) and FITC-oligo/oligofectamine complex (oligofectamine). Concentration of oligo DNA is 40 nM. The data are reported as the mean ( standard of deviation of three replicates.

Figure 7. Cytotoxicity assays of (A) PDMAEA(A1-A5)/oligo DNA and (B) quanternized PDMAEA (B1 to B5)/oligo DNA complexes at various N/P ratios (10, 100, and 200) after 30 min complexation in water and 4 h of transfection to HeLa cells. Controls used are naked oligo DNA (9-27) and oligo DNA/oligofectamine complex (oligofectamine). Concentration of oligo DNA used is 40 nM. The data are reported as the mean ( standard deviation of three replicates.

nanoparticle complexes stored for 1 week, followed by incubating with the cells for 4 h showed little or no transfection (Figure 6B). This demonstrates that the nanoparticle complexes made from PDMAEA transfect readily into the cells before the complexes undergo self-catalyzed dissociation. For the B series, nanoparticle complexes (after mixing for 30 min) and then incubation with the cells for 4 h, as shown in

Figure 6C, gave high transfection even for the low-molecularweight polymers (B1B3). This high transfection may be attributed to the higher loading capacity of quanternized PDMAEA than that of PDMAEA. As expected, we still observed high levels of transfection after the B1B5 nanoparticle complexes had being stirred for 1 week and then incubated with the cells (Figure 6D). 3546


Biomacromolecules Cytotoxicity Assay. Cytotoxicity is also a critical issue for polycations in gene delivery. Therefore, we evaluated toxicity using a Cell Titer Glo, a metabolic assay, that reports cytotoxicity as decreased luminescence for all polyplexes tranfected into HeLa cells. Figure 7A showed the relative cytotoxicity of HeLa cells following the treatment with series A nanoparticle complexes at different N/P ratios. We observed that cytotoxicity increased with increasing N/P ratio and molecular weight. For example, A1A3 nanoparticle complexes showed little or no cytotoxicty, even at the highest N/P ratio of 200 compared with the more toxic A4 and A5 at the same N/P ratio. A similar trend is also apparent from the data presented in Figure 7B for the series B polymers. The high-molecular-weight polymers (B4 and B5) at an N/P ratio of 10 exhibited little toxicity, but at higher N/P ratios the toxicity was high. The results suggest that the polymers (both the PDMAEA and the permanent cationic form of PDMAEA) with molecular weights equal to or below 5600 exhibit little or no toxicity, and such polymer can be used as a component in gene delivery carriers. For molecular weights greater than 5600, one should carefully consider the N/P ratio and maintain this value close to 10.

’ CONCLUSIONS In this work, we show that PDMAEA has the potential to strongly bind, protect, and release via self-catalyzed hydrolysis small oligo DNA sequences into cells. We found that PDMAEA formed large nanoparticle complexes of ∼400 nm with oligo DNA that could protect the DNA from DNase in serum. After 10 h in solution, the PDMAEA nanoparticle complexes fully dissociated into free polymer and oligo DNA, resulting in a significant decrease in the size (to ∼5 nm) and the degradation of oligo DNA by DNase. This dissociation of the complex was catalyzed through a self-hydrolysis process of PDMAEA to form poly(acrylic acid) and N,N0 -dimethylamino ethyl ethanol, both of which have been shown to have little or no toxicity to cells. These PDMAEA nanoparticle complexes rapidly transfected into HeLa cells, with >80% transfection after 4 h. This degree of transfection was dependent on the molecular weight of the polymer; the greater the molecular weight the greater the transfection. The nanoparticle complexes showed no toxicity for all molecular weights at an N/P ratio of 10. Only the higher molecular weight polymers (A4 and A5) at N/P ratios of 200 showed significant levels of cytotoxicity. Our results indicate that PDMAEA has many of the useful attributes of a gene delivery carrier component: it can bind strongly to oligo DNA, it can protect the DNA from DNase, it has high levels of transfection even at a low N/P ratio, and it showed no cytotoxicity at all molecular weights and N/P ratios (even 200) except at the highest molecular weight (8600) and higher N/P ratio of 100 and 200. These attributes make PDMAEA a promising component in a gene delivery carrier without the concern of the accumulation toxicity of nanoparticles in human body after multiadministration. ’ ASSOCIATED CONTENT

bS

Supporting Information. The distributions from DLS data are given. This material is available free of charge via the Internet at http://pubs.acs.org.

’ AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected].

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Self-Catalyzed Degradable Cationic Polymer for Release of DNA

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