Langmuir 2008, 24, 6881-6888
6881
Plasmid DNA Complexation with Phosphorylcholine Diblock Copolymers and Its Effect on Cell Transfection XiuBo Zhao, ZhuoQi Zhang, Fang Pan, Thomas A. Waigh, and Jian R. Lu* Biological Physics Group, School of Physics and Astronomy, The UniVersity of Manchester, Schuster Building, Manchester M13 9PL, United Kingdom ReceiVed February 25, 2008. ReVised Manuscript ReceiVed April 4, 2008 We examined a series of novel cationic MPC-based (2-methacryloyloxyethyl phosphorylcholine) copolymers as vectors for gene delivery, with emphasis on the assessment of the effects of the charge ratio (administered via pH variation) on the complex (polyplex) formation and the subsequent transfection efficiency. A combination of electrophoresis, dynamic light scattering, and small angle neutron scattering was used to characterize the structure and charge distribution of the polyplexes formed between the copolymer and the luciferase plasmid DNA. Polymers with larger hydrophobic side chains had lower pKa values and tended to aggregate more strongly. For a given copolymer, electrostatic interaction was the main driving force for the formation of the nanopolyplexes. When the cationic copolymers were in excess, the majority of the polyplexes formed was neutral, and only a small faction of them carried net positive charges. Polyplexes formed under excess copolymer protected the DNA from restriction enzyme digestion. As the copolymers were weak polyelectrolytes, the pH had a distinct effect on the structure and charge distribution of the polyplexes formed. Below the pKa, the copolymers were found to bind with the plasmid DNA in the form of unimers, while above the pKa, the copolymers self-aggregated and complexed with DNA in the form of micelles. It was subsequently found that unimer/DNA polyplexes were far more effective in the transfection of HEK293 cells than micellar DNA polyplexes. The results thus revealed that different hydrophobicities of the side chains in the copolymer series led to different nanostructuring and charge characteristics, which had a consequential effect on the transfection efficiency. This study provided useful insight into the molecular processes underlying polyplex formation and demonstrated a strong link between structural and physical properties of polyplexes and cell transfection efficiency.
1. Introduction DNA delivery has stimulated a great deal of interest over the past decade because it holds great promise for curing many acquired and inherited diseases. Encapsulation of DNA using protein shells from naturally occurring viruses appeared to be an ideal candidate for the transportation of target genes into the cell nucleus and other subcellular compartments. However, viral gene delivery still suffers from major setbacks (e.g., unacceptable immune responses) in spite of some tremendous advances that have been achieved.1–5 The drawbacks associated with viral DNA delivery have stimulated studies of nonviral gene delivery. In nonviral gene delivery, physical methods such as microinjection, gene gun, electroporation, laser irradiation, and magnetofection were developed to deliver genes into cells.4,6,7 The more commonly adopted approaches to aid the delivery processes use chemicals such as cationic lipids (surfactants) and natural and synthetic polymers. Lipids and polymers form complexes with DNA (termed lipoplexes and polyplexes, respectively) that improve the internalization of DNA molecules into cells. To reach cellular nuclei, the external genes have to overcome a series of intracellular barriers including escape from * Corresponding author. Tel.: manchester.ac.uk.
44-161-3063926; e-mail: J.Lu@
(1) Pack, D. W.; Hoffman, A. S.; Pun, S.; Stayton, P. S. Nat. ReV. Drug DiscoVery 2005, 4, 581–593. (2) Segura, T.; Shea, L. D. Ann. ReV. Mater. Res. 2001, 31, 25–46. (3) El-Aneed, A. J. Controlled Release 2004, 94, 1–14. (4) Mehier-Humbert, S.; Guy, R. H. AdV. Drug DeliVery ReV. 2005, 57, 733– 753. (5) Goverdhana, S.; Puntel, M.; Xiong, W.; Zirger, J. M.; Barcia, C.; Curtin, J. F.; Soffer, E. B.; Mondkar, S.; King, G. D.; Hu, J.; Sciascia, S. A.; Candolfi, M.; Greengold, D. S.; Lowenstein, P. R.; Castro, M. G. Mol. Ther. 2005, 12, 189–211. (6) Tan, P. H.; Chan, C. L. H.; Chan, C.; George, A. J. T. Br. J. Surgery 2005, 92, 1466–1480. (7) Niidome, T.; Huang, L. Gene Ther. 2002, 9, 1647–1652.
enzymatic disintegration.8–10 Since the introduction of lipofection in 1987, various cationic lipid formulations have been developed and tested as vectors for gene delivery.11 However, because most of them are cell specific, organic solvent-based, and have a varying degree of cytotoxicity, they are unsuitable for further development toward in vivo applications. The development of water-based formulations from cationic polymers and polypeptides has been extensively pursued and may offer promising alternatives.12–14 Poly(L-lysine) (PLL), poly(ethylenimine) (PEI), and related compounds (such as poly(ethylene glycol)-grafted-poly(L-lysine) and poly(ethylene glycol)-grafted-poly(ethylenimine)) have been most extensively studied as nonviral gene delivery systems.3,15–17 However, PLL/DNA complexes are rapidly bound to plasma proteins and have a short circulation half-life. The transfection efficiency is often low, and chloroquine needs to be added to improve its behavior. The proton sponge effect of PEI could, in principle, efficiently help PEI/DNA complexes to escape from endosome degradation and result in a relatively high transfection efficiency. However, most PEIs used so far have poorly defined (8) Smedt, S. C. D.; Remaut, K.; Lucas, B.; Braeckmans, K.; Sanders, N. N.; Demeester, J. AdV. Drug DeliVery ReV. 2005, 57, 191–210. (9) Zhang, S.; Xu, Y.; Wang, B.; Qiao, W.; Liu, D.; Li, Z. J. Controlled Release 2004, 100, 165–180. (10) Lechardeur, D.; Verkman, A. S.; Lukacs, G. L. AdV. Drug DeliVery ReV. 2005, 57, 755–767. (11) Felgner, P. L.; Gadek, T. R.; Holm, M.; Roman, R.; Chan, H. W.; Wenz, M.; Northrop, J. P.; Ringold, G. M.; Danielsen, M. Proc. Natl. Acad. Sci. U.S.A. 1987, 84, 7413–7417. (12) Mahato, R. I. AdV. Drug DeliVery ReV. 2005, 57, 699–712. (13) Smedt, S. C. D.; Demeester, J.; Hennink, W. E. Pharm. Res. 2000, 17, 113–126. (14) Smith, L. C.; Duguid, J.; Wadhwa, M. S.; Logan, M. J.; Tung, C. H.; Edwards, V.; Sparrow, J. T. AdV. Drug DeliVery ReV. 1998, 30, 115–131. (15) Nah, J. W.; Yu, L.; Han, S. O.; Ahn, C. H.; Kim, S. W. J. Controlled Release 2002, 78, 273–284. (16) Kichler, A. J. Gene Med. 2004, 6, 3-10. (17) Lee, M.; Kim, S. W. Pharm. Res. 2005, 22, 1–10.
10.1021/la800593q CCC: $40.75 2008 American Chemical Society Published on Web 05/24/2008
6882 Langmuir, Vol. 24, No. 13, 2008
molecular structures (e.g., in a wide range of molecular weights and linear and branched topologies).16,18,19 While factors such as molecular weight, polyplex particle size, degree of polymerization, charge ratio, and protocol by which the complexes are formed all may influence cell transfection efficiency and cytotoxicity, there is a lack of fundamental studies to look into the mechanistic processes underlying complex formation and to link molecular information with cell transfection and expression.3,9,20 For example, although it is widely known that large complexes enter cells by receptor- and clathrin-independent endocytosis and smaller ones through a nonspecific, clathrindependent process, there is a lack of detailed quantitative studies to examine as to how the structure of the complexes affects the cell transfection efficiency.10 A number of studies have reported interesting properties of interfacial adsorption and solution aggregation from diblock copolymers incorporating the MPC block (where MPC stands for 2-methacryloyloxyethyl phosphorylcholine (PC)).21–25 PC containing polymers have a high biocompatibility,26–32 and some studies recently reported attractive benefits of PC containing polymers in gene delivery.33,34 MPC-based diblock copolymers were used in this work as model gene delivery vectors due to their better-defined structures. The MPC block is zwitterionic and hydrophilic, whereas the second tertiary amine block is weakly cationic and is comprised of either 2-(diethylamino)ethyl methacrylate (DEA) or 2-(diisopropylamino)ethyl methacrylate (DPA). Both dynamic light scattering (DLS) and small angle neutron scattering (SANS) revealed strong pH-dependent selfaggregation of the copolymers. Different copolymer aggregation states subsequently implicated the structures of the complexes formed. Our results have further demonstrated the strong effects of size and charge characteristics of the polyplexes on the transfection efficiency of HEK293 cells, thereby providing a useful illustration of the link between physiochemical properties of the polyplexes and cell gene transfection into cells.
2. Experimental Procedures 2.1. Materials. The synthesis of the diblock copolymers has been described previously.35 Atom transfer radical polymerization (ATRP) was used to synthesize MPC blocks. DEA (or DPA) was subsequently added onto the MPC block at high conversion in methanol, resulting in further chain growth and the formation of MPC-DEA (or MPCDPA) diblock copolymers. A stock PC copolymer solution was (18) Densmore, C. L.; Orson, F. M.; Xu, B.; Kinsey, B. M.; Waldrep, J. C.; Hua, P.; Bhogal, B.; Knight, V. Mol. Ther. 2000, 1, 180–188. (19) Godbey, W. T.; Wu, K. K.; Mikos, A. G. J. Controlled Release 1999, 60, 149–160. (20) Haider, M.; Megeed, Z.; Ghandehari, H. J. Controlled Release 2004, 95, 1–26. (21) Zhao, X.; Zhang, Z.; Pan, F.; Ma, Y.; Armes, S. P.; Lewis, A. L.; Lu, J. R. Langmuir 2005, 21, 9597–9603. (22) Zhao, X.; Pan, F.; Lu, J. R. Annu. Rep. Prog. Chem., Sect. C: Phys. Chem. 2007, 103, 261–286. (23) Zhao, X.; Pan, F.; Lu, J. R. Langmuir, submitted. (24) Zhao, X.; Zhang, Z.; Pan, F.; Ma, Y.; Armes, S. P.; Lewis, A. L.; Lu, J. R. Surf. Interface Anal. 2006, 38, 548–551. (25) Mu, Q. S.; Zhao, X.; Lu, J. R.; Armes, S. P.; Lewis, A. L.; Thomas, R. K. J. Phys. Chem. B, in press. (26) Tang, Y.; Su, T. J.; Armstrong, J.; Lu, J. R.; Lewis, A. L.; Vick, T. A.; Stratford, P. W.; Heenan, R. K.; Penford, J. Macromolecules 2003, 36, 8440– 8448. (27) Tang, Y.; Lu, J. R.; Lewis, A. L.; Vick, T. A.; Stratford, P. W. Macromolecules 2002, 35, 3955–3964. (28) Zhang, Z.; Cao, X.; Zhao, X.; Withers, S. B.; Holt, C. M.; Lewis, A. L.; Lu, J. R. Biomacromolecules 2006, 7, 784–791. (29) Lewis, A. L. Colloids Surf., B 2000, 18, 261–275. (30) Lewis, A. L.; Freeman, R. N. T.; Redman, R. P.; Tolhurst, L. A.; Kirkwood, L. C.; Grey, D. M.; Vick, T. A. J. Mater. Sci: Mater. Med. 2003, 39–45. (31) Lewis, A. L.; Hughes, P. D.; Kirkwood, L. C.; Leppard, S. W.; Redman, R. P.; Tolhurst, L. A.; Stratford, P. W. Biomaterials 2000, 21, 1847–1859. (32) Lewis, A. L.; Tolhurst, L. A.; Stratford, P. W. Biomaterials 2002, 23, 1697–1706.
Zhao et al. typically made by dissolving 0.1 g of polymer sample in 10 mL of ultrahigh quality (UHQ) water (Purelab UHQ, Vivendi Water Systems Ltd.) or D2O (99+% D, Sigma) for SANS and was then stirred overnight, filtered with 0.2 µm syringe filters, and subsequently diluted to the desired concentrations. The solution pH was adjusted by a minimal amount of HCl or NaOH solution (in UHQ or D2O) to provide the desired pH. Luciferase (Luc) plasmid DNA (pCMV-Luc, 5.1 kb, Clontech) was extracted from Escherichia coli using a Qiagen Plasmid DNA Purification Mega Kit following the manufacturer’s protocol.37,36 Restriction enzyme digestion and electrophoresis were carried out to check the extracted plasmid DNA. The plasmid concentration was determined using a Genesys 6 spectrophotometer (Thermo Spectronic), assuming that 50 µg/mL corresponded to one absorbance unit at 260 nm. 2.2. Fabrication of Polymer/DNA Complexes. Copolymer/DNA complexes were prepared by adding the DNA solution to the polymer solutions. Stock solutions were prepared at appropriate concentrations such that the final solutions were diluted to a constant DNA concentration of 0.25 mg/mL after mixing with polymer. Copolymer solutions were diluted to the desired concentrations to achieve different charge ratios (positive/negative) between copolymer and DNA (from 20:1 to 1:10). These polyplex solutions were mostly obtained by mixing 0.25 mL of DNA solution with 0.25 mL of polymer solution. All the solutions were gently shaken and were left for 30 min at room temperature for equilibration before use. 2.3. Agarose Gel Electrophoresis. A 4 µL solution from each complex sample (1 µg of DNA equivalence) was mixed with 1 µL of DNA loading buffer. The new samples were briefly mixed by pipetting and were pulse spun in a benchtop microcentrifuge. These samples were then loaded into 1% agarose gel containing 0.5 µg/mL ethidium bromide (EtBr, Sigma). Electrophoresis was carried out at 100 V in 1X TBE buffer (Sigma) for 30 min. The DNA band was visualized under UV transillumination (Fisher). Copolymers were stained by immersing the gel in a Coomassie blue staining solution (10:45:45 glacial acetic acid/methanol/doubly distilled water (ddH2O) and 0.1% w/v Coomassie bright blue G250 (Fisher) for 30 min in a shaker followed by washing with destaining solution (10:10:80 glacial acetic acid/methanol/ddH2O) overnight. All photos were taken using a Sony DSC T1 digital camera. 2.4. Restriction Enzyme Digestion. Luc plasmid DNA solution and copolymer/DNA complexes (0.5 µg of DNA equivalence) were mixed with 1 µL of 10X buffer D, 0.5 µL of BSA, 0.5 µL of Not I, QG (Promega), and ddH2O to give a total volume of 10 µL each. They were then incubated in a water bath at 37 °C for 1.5 h. Agarose gel electrophoresis was subsequently performed on these samples following the same procedures as described previously. 2.5. DLS. DLS was used to determine the average size of polymer aggregates or complexes under different solution conditions. In DLS experiments, particles or aggregates in solution are illuminated with light of a given wavelength, and the intensity fluctuations from the scattered light are measured over a time course between 1 ns and 1 ms. Intensity fluctuations occur due to the random diffusion of particles through the solvent (Brownian motion). Changes in signal intensity over the time course arising from these fluctuations are described through an autocorrelation function that is related to intraand interparticle scattering factors.38,39 If the particles studied are small as compared to the wavelength used, the translational diffusion (33) Chim, Y. T. A.; Lam, J. K. W.; Ma, Y.; Armes, S. P.; Lewis, A. L.; Roberts, C. J.; Stolnik, S.; Tendler, S. J. B.; Davies, M. C. Langmuir 2005, 21, 3591–3598. (34) Zhao, X.; Pan, F.; Zhang, Z.; Grant, C.; Ma, Y.; Armes, S. P.; Tang, Y.; Lewis, A. L.; Waigh, T.; Lu, J. R. Biomacromolecules 2007, 8, 3493–3502. (35) Lobb, E. J.; Ma, I.; Billingham, N. C.; Armes, S. P.; Lewis, A. L. J. Am. Chem. Soc. 2001, 123, 7913–7914. (36) Ma, Y.; Tang, Y.; Billingham, N. C.; Armes, S. P.; Lewis, A. L.; Lloyd, A. W.; Salvage, J. P. Macromolecules 2003, 36, 3475–3484. (37) QIAGEN Plasmid Purification Handbook, 2nd ed.; QIAGEN: Hilden, Germany, 2003. (38) Berne, B. J.; Percora, R. Dynamic Light Scattering; Wiley: New York, 1976. (39) Vertesse, B. G.; Magazu`, S.; Mangione, A.; Migliardo, F.; Brandt, A. Macromol. Biosci. 2003, 3, 477–481.
DNA Complexation with Phosphorylcholine Copolymers
Langmuir, Vol. 24, No. 13, 2008 6883
coefficient, D0, can be determined through Laplace inversion of the autocorrelation function. From the diffusion coefficient, the hydrodynamic radius of the scattering particles can be calculated from the Stokes-Einstein equation
Rh ) kBT/6πηD0 where kB denotes the Boltzmann constant, T is the absolute temperature, and η is the viscosity of the solvent at the same temperature. Rh is an experimental measure of the radius of a hydrated object. In the case of a nonspherical object, it approximates to the largest rotational radius. DLS experiments were carried out using the Zetasizer Nano-S Instrument from Malvern Instruments Ltd. (model no. ZEN1600). This instrument can measure particles ranging from 0.6 nm to 6 µm and in a wide range of concentrations (0.1 mg/mL to 40 % w/v) at a temperature range of 2-90 °C. In this work, copolymer solutions (0.1 wt %) at different pH values were measured using a quartz cell at 25 °C. 2.6. SANS. SANS measures the differential scattering crosssection as a function of wave vector (Q) and is more sensitive to the size and shape of the nanosized particles in the sample than the DLS technique used.40 As in the case of light scattering, the scattering intensity also is related to intra- and interparticle scattering form factors, but when the concentration is sufficiently low, the interparticle form factor approximates to 1. Analytical expressions can be written for particles of different shapes, with and without size distributions. Information about the structure of scattering objects usually is obtained by comparing a scattering profile calculated from a presumed geometrical shape with the measured data, and the process is iterated until an acceptable fit is produced. The data were fitted using the FISH2 program supplied by Dr. Richard Heenan at RAL. Copolymer and copolymer/DNA complex samples for SANS were prepared in D2O as mentioned previously. The experiments were carried out at the ISIS Neutron Facility, Rutherford Appleton Laboratory, Oxford, U.K. 2.7. Cell Culture. HEK293 cells (a product of ATCC and a gift from Dr. John Garland, Manchester Medical School, The University of Manchester, U.K.) were cultured at 37 °C, 5% CO2 in Dulbecco’s Modified Eagle’s Medium (DMEM, Sigma) supplemented with 10% heat-inactivated fetal bovine serum (FBS, Sigma), 100 U/mL penicillin, and 50 µg/mL streptomycin (Sigma, UK). 2.8. Cell Transfection Assay. Cells were seeded in 24-well cell culture plates at a density of 4 × 104/well and grown overnight at 37 °C, 5% CO2. Copolymer/DNA complexes (0.5 µg of DNA equivalence) at different N/P ratios were loaded, and each plate carried positive controls using TransIT-LT1 transfection reagent (Cambridge Bioscience). Loaded plates were incubated for 24 and 48 h, respectively, with and without 10% FBS. After these treatments, the cells were washed with PBS (Sigma), lysed using 50 µL of buffer (Triton X-100 0.1-1%, 50 µL of protease inhibitor cocktail (Sigma), and 10 mL of PBS). A total of 20 µL of lysate of each sample was mixed with 50 µL of Luc assay substrate (Promega) and then measured under a luminometer (Lumat LB 9507).
Figure 1. DLS measurements of hydrodynamic diameters as a function of pH from MPC30-DEA60 (9) and MPC30-DPA60 (2), all at 0.1 wt % and 25 °C. Lines through the points are to help guide the eye.
3.1. Response of Self-Aggregation of Copolymers to Solution pH. DLS was first used to determine the hydrodynamic size (diameter) of the scattering objects from copolymer solutions. Figure 1 shows the pH-dependent variation of the hydrodynamic size from the two copolymers: MPC30-DEA60 and MPC30DPA60. Below pH 6, the size measured was ca. 10 nm for both copolymers and was equivalent to the hydrodynamic diameter of a polymeric molecule. Above pH 6, the apparent hydrodynamic size increased, indicating the formation of polymeric micelles. The middle points shown in Figure 1 correspond to pH values
of 6.3 and 7.6 for MPC30-DPA60 and MPC30-DEA60 diblock copolymers, respectively, consistent with results obtained from pyrene emission spectra by Ma et al.36 The pKa values for DPA and DEA homopolymers reported by Bu¨tu¨n et al.41 are 6.0 and 7.3, respectively, and are close to the middle points of aggregation for the copolymers. As the pKa values are likely to be similar for the copolymers, the results show that copolymer micellization starts immediately above their pKa values, indicating the strong effect of deprotonation of the amine groups. The low pKa and its early micellization for MPC30-DPA60 are consistent with its greater side chain hydrophobicity. This observation is also consistent with the much greater size of the aggregates formed. The micellization process thus is driven by a combined action of pH related deprotonation and the hydrophobicity of the amine block. It is, however, useful to explore as to whether the pHdependent self-aggregation from the two copolymers affects their complexation with DNA and the subsequent gene transfection. 3.2. Agarose Gel Electrophoresis of Copolymer/DNA Complexes. We demonstrated elsewhere34 that MPC30-DEA70 formed interesting nanopolyplexes with a small antisense oligonucleotide (ODN, single strand with 15 bases) and that the size and shape of the polyplexes varied with the charge ratio. These polyplexes produced a range of transfection efficiencies for HeLa cells, showing a combined effect of charge and size. As in the case of the ODN study, we again focused on examining the role of charge interaction under the same experimental conditions. Figure 2 shows the agarose gel electrophoresis using MPC30-DEA70/DNA complexes produced at different cationic/ anionic (N/P) ratios. The gel was stained by ethidium bromide to view DNA and Coomassie bright blue dye to view the copolymer, respectively. As the amine groups in the copolymer are cationically charged at pH 7, pure copolymers migrate to the cathode (lane 1 in Figure 2A,B), while the negatively charged plasmid DNA migrates to the opposite direction (lane 8 in Figure 2A and lane 12 in Figure 2B). However, the MPC30-DEA70/ Luc plasmid DNA complexes formed at different N/P ratios migrate in both directions, indicating different net charges on these complexes. Specifically, the complexes displayed in lanes 2-4 in Figure 2A and lanes 2-6 in Figure 2B where the copolymer was in excess are overall positively charged. The brightness of the sample wells indicates the presence of the complexes, but because they are overall neutral, they remain in the wells. It can also be seen from Figure 2 that instead of a distinct DNA band of migration, a continuous stripe of the complexes is observed, indicating that there is a distribution of different net charges on the complexes. At the N/P ratio of unity
(40) King, S. M. Small Angle Neutron Scattering. In Modern Techniques for Polymer Characterization; Pethrick, R. A., Dawkins, J. V., Eds.; Wiley: New York, 1999; Ch. 7, pp 171-231.
(41) Bu¨tu¨n, V.; Armes, S. P.; Billingham, N. C. Polymer 2001, 42, 5993– 6008. (42) Lynn, D. M.; Langer, R. J. Am. Chem. Soc. 2000, 122, 10761–10768.
3. Results and Discussion
6884 Langmuir, Vol. 24, No. 13, 2008
Zhao et al.
Figure 2. Agarose gel electrophoresis of copolymer (MPC30-DEA70)/DNA (Luc plasmid) complexes. (A) Lanes 1 and 8 are copolymer and DNA controls. Lanes 2-7 correspond to 20:1, 10:1, 5:1, 1:1, 0.2:1, and 0.1:1 (cationic/anionic charge ratios (N/P) between copolymer and DNA), respectively. (B) Lanes 1 and 12 are copolymer and DNA controls. Lanes 2-11 correspond to 5:1, 4:1, 3:1, 2:1, 1.5:1, 1:1, 0.5:1, 0.33:1, 0.25:1, and 0.2:1, respectively.
(lane 5 in Figure 2A and lane 7 in Figure 2B), most complexes prefer to stay in the well due to the dominant charge neutrality, but a smaller fraction still moves out, again indicating the polydispersity of the charge distribution. The situation, however, becomes different when DNA is in excess. While some of the complexes still remain in the wells, quite a large fraction of them migrates to the anode and is widely distributed, just as in the case of copolymer excess, but in the opposite direction. Distinct DNA bands can be seen with the same migration distance as the DNA control. These observations again show a wide distribution of net charges at a given charge ratio, but when DNA is in excess, the net charges of the complexes become predominantly negative with some DNA remaining pure and uncomplexed. Because the DNA concentration was fixed at 1 µg/well, the brighter DNA band in lane 7 than in lane 6 in Figure 2A indicates less complex formation due to the smaller amount of copolymer present in sample 7. The electrophoresis results thus provide an overall view of the net charge distributions in the complexes with varying N/P ratios. These observations are broadly similar to those obtained from the complexes formed between MPC30-DEA70 and small ODN, showing that the larger DNA size has not affected the broad charge distribution of the charge driven complexation.34 For comparison, the same electrophoresis measurement was performed for complexes formed between Luc DNA and MPC30DPA60 at pH 5 and 7, and the results are shown in Figure 3. The results at pH 5 are quite consistent with those taken between Luc DNA and MPC30-DEA70, indicating the formation of copolymer/ DNA complexes under this condition. However, it can be seen from Figure 3 that the fraction of positively charged complexes formed at high N/P ratios (e.g., see lane 2 in Figure 3) is low and that the complexes are almost entirely neutral. Thus, it seems that even at high N/P ratios, this copolymer did not complex strongly with Luc DNA and that the number of net positively charged polyplexes is small. However, the features observed at pH 7 for this copolymer/ DNA system are very different, indicating a different pattern of interaction. Unlike the interaction at pH 5, there is only a small amount of neutral complexes left inside the wells. Instead, the copolymer/DNA polyplexes formed at high N/P ratios migrate in the same direction as the DNA, indicating the formation of almost all net negative complexes with the DNA. In light of the
Figure 3. Agarose gel electrophoresis from different copolymer/DNA complexes formed between MPC30-DPA60 and Luc plasmid DNA at pH 5 (lanes 1-6) and pH 7 (lanes 7-12) in 10 mM PBS. At pH 5: Lanes 1 and 6 are copolymer and DNA controls. Lanes 2-5 correspond to copolymer/DNA complexes with charge ratios (N/P) of 5:1, 2:1, 1:1, and 0.5:1, respectively. At pH 7: Lanes 7 and 12 are copolymer and DNA controls. Lanes 8-11 correspond to copolymer/DNA complexes with N/P ratios of 5:1, 2:1, 1:1, and 0.5:1, respectively.
results shown in Figure 1 from DLS, MPC30-DPA60 already formed micelles under pH 7 before being mixed with DNA. The complexation can only occur between copolymer micelles and DNA molecules. As pH 7 is above the pKa value for this copolymer, the net positive charges each micelle carries are low. Furthermore, upon the formation of micelles, the tertiary amine groups are encapsulated inside the hydrophobic cores, with hydrophilic PC groups forming the corona outside,21,36 making it difficult to complex with DNA. Some of the residual cationic charges inside the copolymer micelles are sterically inaccessible. These factors may work together and cause incomplete neutralization. Several DNA molecules may bind to one copolymer micelle, and the net charges per polyplex become negative, consistent with what was observed.
DNA Complexation with Phosphorylcholine Copolymers
Langmuir, Vol. 24, No. 13, 2008 6885
Figure 4. SANS scattering intensity (I) plotted against wave vector (K) from 2.5 mg/mL pure copolymer (MPC30-DEA70) solution (+), 0.25 mg/mL pure luciferase plasmid DNA solution (*), and complex (MPC30DEA70/Luc plasmid DNA) solutions at ratios (cationic/anionic) of 20:1 (0), 10:1 ()), 5:1 (∆), and 1:1 (O). DNA concentrations were fixed at 0.25 mg/mL for each sample, all at pH 7.
3.3. SANS Study of Copolymer/DNA Complexes. SANS experiments were performed to gain more direct structural information on the complexation between MPC30-DEA70 and plasmid DNA. In our previous work,34 SANS was successfully used to reveal the size and shape of the copolymer/oligonucleotide complexes formed under different solution conditions. The work undertaken in this study allows us to examine as to how different sizes of DNA affect complexation. The fitted profiles under different charge ratios are shown as continuous lines in Figure 4, and the parameters obtained are listed in Table 1. Pure copolymer and pure plasmid DNA were used as controls. All the measured data were modeled with a solid cylindrical morphology. Pure copolymers were fitted as cylinders with a radius of 15 ( 2 Å and length of 200 ( 50 Å. The radius and length for pure DNA were 165 ( 10 and 508 ( 50 Å, respectively. For complexes at N/P ratios above 1:1, two kinds of cylinders were detected by SANS. The radius and length for small cylinders are the same as the pure copolymers, indicating that excess copolymers are free and that they do not associate with the complexes. The radius and length for the large cylinders are slightly smaller than the pure plasmid DNA. This may be caused by charge condensation during complexation. As the charge ratio approaches 1:1, only one type of cylinder was detected, indicating complete complexation between copolymer and plasmid DNA. It is interesting to note that the fitted scale factors were in proportion to the N/P ratio. At a N/P ratio of 20:1, the ratio of the scale factors between excess copolymer and copolymer/DNA complex was 19:1. As N/P ratios dropped to 10:1 and 5:1, the corresponding ratios between excess copolymer and complex became 9:1 and 4:1. It is thus clear that no matter as to how much copolymers were in excess, only the portion equal to the amount of DNA participated in complexation. While the stochastic complexation was consistent with the charge driven interaction, the results from electrophoresis as shown in Figure 2 indicate
Figure 5. (a) SANS scattering intensity (I) plotted against wave vector (Q) from 3 mg/mL pure copolymer (MPC30-DPA60) solution (+), 0.25 mg/mL pure luciferase plasmid DNA solution (*), and complex (MPC30DPA60/Luc plasmid DNA) solutions at (cationic/anionic) ratios of 5:1 ()), 2:1 (0), 1:1 (O), and 0.5:1 (∆). DNA concentrations were fixed at 0.25 mg/mL for each sample, all at pH 5. (b) SANS scattering intensity (I) plotted against wave vector (K) from 3 mg/mL pure copolymer (MPC30-DPA60) solution (+) and complex (MPC30-DPA60/Luc plasmid DNA) solutions at (cationic/anionic) ratios of 5:1 ()), 2:1 (0), 1:1 (O), and 0.5:1 (∆). DNA concentrations were fixed at 0.25 mg/mL for each sample, all at pH 7.
that at each N/P ratio, the net charges on some of the complexes are skewed toward positive distributions. This is, however, expected from statistical analysis with a small kinetic barrier to the equilibration of the complexes’ charges. The SANS data together with the electrophoresis data indicate that the formation of complexes bearing net positive charges does not cause any measurable structural changes from those carrying net neutral charges. It is interesting to note that this binding model is very different from that observed between copolymers and ODN, where ODN molecules were much smaller than copolymers and bind around the copolymers.34 This means that the relative size difference between copolymer and DNA could result in different mechanistic processes of binding and different polyplex morphologies. Many authors studied changes in the size and zetapotential with N/P ratios using DLS, but because the polymers were poorly defined, it was often difficult to ascertain the exact
Table 1. Best-Fit SANS Parameters from Scattering Profiles Shown in Figure 4a charge ratio
polymer -1
scale factor 1 (cm ) radius 1 (Å) length 1 (Å) scale factor 2 (cm-1) radius 2 (Å) length 2 (Å) a
2.51 × 10 15 ( 2 200 ( 50
-6
20:1
10:1
5:1
1:1
DNA
1.93 × 15 ( 2 200 ( 50 1.02 × 10-7 155 ( 10 490 ( 50
9.32 × 15 ( 2 200 ( 50 1.07 × 10-7 155 ( 10 490 ( 50
4.42 × 15 ( 2 200 ( 50 1.09 × 10-7 155 ( 10 490 ( 50
2.71 × 10-8 165 ( 10 500 ( 50
6.15 × 10-9 165 ( 10 500 ( 50
10-6
10-6
10-6
Scale factor provides a useful indication of the number concentration of particles in the aqueous dispersion.
6886 Langmuir, Vol. 24, No. 13, 2008
Zhao et al.
Table 2. Best-Fit SANS Parameters from Scattering Profiles Shown in Figure 5a,b Figure 5a ratio
polymer
5:1
2:1
1:1
0.5:1
DNA
scale factor 1 (cm-1) radius 1 (Å) length 1 (Å) scale factor 2 (cm-1) radius 2 (Å) length 2 (Å)
1.76 × 10-6 17 ( 3 255 ( 30
6.84 × 10-7 17 ( 3 255 ( 30 1.78 × 10-7 165 ( 10 500 ( 50
1.88 × 10-7 17 ( 3 255 ( 30 1.5 × 10-7 165 ( 10 500 ( 50
5.36 × 10-8 165 ( 10 500 ( 50
5.8 × 10-8 165 ( 10 500 ( 50
1.17 × 10-8 165 ( 10 500 ( 50
Figure 5b ratio
polymer
5:1
2:1
1:1
0.5:1
DNA
scale factor 1 (cm-1) radius 1 (Å) length 1 (Å) scale factor 2 (cm-1) radius 2 (Å) length 2 (Å)
5.22 × 10-6 220 ( 30 145 ( 5
1.08 × 10-6 250 ( 30 180 ( 10 6.38 × 10-7 160 ( 10 500 ( 50
1.04 × 10-7 250 ( 30 180 ( 10 2.31 × 10-7 160 ( 10 500 ( 50
2.05 × 10-8 250 ( 30 180 ( 10 4.6 × 10-8 160 ( 10 500 ( 50
2.2 × 10-8 250 ( 30 180 ( 10 2.41 × 10-8 160 ( 10 500 ( 50
6.20 × 10-9 165 ( 10 500 ( 50
mode of complexation.42 Furthermore, because DLS detects the overall hydrodynamic size, which is insensitive to shape and is dominated by the large end of the complexes, the true structural information might be different. Chim et al.33 used AFM to examine complexes formed between MPC30-DMAx (where x is the mean degree of polymerization of the DMA block) and gWiz Luc plasmid DNA and observed a range of structural features depending on the copolymer structure, N/P ratio, etc. They observed rod and rectangular block-like structures for x ) 60, and at low N/P ratios, the sizes are consistent with our results from SANS studies. SANS experiments also were carried out for complexes formed between MPC30-DPA60 copolymer and Luc plasmid DNA at both pH 5 and pH 7. The scattering profiles together with the best fits are shown in Figure 5a,b, and the parameters obtained are shown in Table 2. The results at pH 5 are quite similar to the complexation observed between MPC30-DEA70 and plasmid DNA. The free excess copolymer appears to be longer, but the diameter is similar to that obtained from MPC30-DEA70, showing that the increased chain size and hydrophobicity may force the chain to extend further in aqueous solution. Upon complexation with DNA, the size and shape of the complexes and the N/Pdependent stochastic behavior are almost exactly the same as observed for MPC30-DEA70. However, when the pH is increased to 7, disk-like polymer micelles were detected at all N/P ratios (see Table 2b). These
polymeric micelles were fitted with a radius of 220 ( 30 Å and height of 145 ( 5 Å. Once mixed with the plasmid DNA, both disk-like copolymer micelles and cylinder-like plasmid DNA were detected. But, in the presence of DNA, the copolymer micelles become slightly larger than those formed from the pure copolymer, indicative of the occurrence of binding and complexation. Although most of the copolymers at pH 7 form disklike micelles, there are still some free polymer molecules left in solution. These free molecules bind with DNA in preference to the binding between micelles and DNA. But, as already stated, free polymer binding to DNA does not appear to alter the size of DNA significantly due to the size difference. Any binding or association occurring between polymeric micelles and DNA could lead to the formation of larger aggregates, but this was not detected by SANS. Figure 6 shows the schematic representation of complexes formed between MPC30-DPA60 and plasmid DNA at pH 5 and 7. In contrast to ODN, plasmid DNA (around 5.1 kb) is much larger than the copolymer, while ODN (15 bases) is significantly smaller. In terms of charge neutralization, a number of copolymers (more than 170) are required to bind to the plasmid DNA. However, at pH 7, the copolymers self-aggregate to form micelles first. They may then interact with the plasmid DNA. As each plasmid DNA is now roughly the same size as a copolymer micelle, it is difficult to neutralize the DNA due to the steric effect and the significantly reduced charge density. Any weak
Figure 6. Schematic diagram of complexes formed by MPC30-DPA60 and plasmid DNA at pH 5 and 7.
DNA Complexation with Phosphorylcholine Copolymers
Figure 7. Agarose gel electrophoresis of restriction enzyme digested DNA/copolymer polyplexes. Complexation was carried out between MPC30-DEA70 and Luc plasmid at different ratios. Lanes 2-5 correspond to 10:1, 5:1, 1:1, and 0.5:1 (cationic/anionic charge ratios), respectively. Lane 6 is DNA digested by the enzyme. Lane 1 represents complexes formed between copolymer and digested DNA (from the sample of lane 6) at a ratio of 10:1. Lane 7 is the Luc plasmid DNA control.
association between polymeric micelles and DNA would result in incomplete neutralization and net negative charges on the polyplexes formed. 3.4. Restriction Enzyme Digestions of Copolymer/DNA Polyplexes. We further investigated the capability of MPC30DEA70 to protect plasmid DNA from restriction enzyme digestion after complex formation. This is one of the basic requirements for a desired gene transfer vector. Figure 7 compares the different effects of restriction enzyme digestion using an enzyme called Not I, QG. The digestion was carried out at different N/P ratios for 1.5 h at 37 °C. Lanes 2-5 in Figure 7 correspond to N/P ratios of 10:1, 5:1, 1:1, and 0.5:1, respectively. Lane 6 in Figure 7 is the DNA digested by the enzyme. Lane 1 in Figure 7 represents the complexes formed after DNA has been digested (from the sample of lane 6) and the N/P ratio was fixed at 10:1. Lane 7 in Figure 7 is the Luc plasmid DNA control only. Complexes formed above a N/P ratio of 1:1 (lanes 1-3 in Figure 7) migrate in the direction opposite to the DNA, indicating the formation of complexes carrying a net positive charge. It is unclear at this point as to whether DNA has been digested or not. However, it can be observed in Figure 7 that at N/P ratios of unity or lower, the DNA samples have been digested because of the presence of two distinct bands on the gel plate. The continued retardation of the DNA in lanes 4 and 5 in Figure 7 indicates the polydispersity of the net charge distribution corresponding to different DNA fragments. 3.5. Cell Transfection by Copolymer/Plasmid DNA Complexes. To test any consequential effect of exposure of the copolymer/DNA complexes to the restriction enzyme, transfection experiments using representative enzyme treated samples were carried out using HEK293 cells. Figure 8 shows the relative transfection rates from these samples. As lane 1 in Figure 7 represents digested DNA before complexation, 10:1 (*) in Figure 8 works as a negative control or background. The transfection from samples from lane 2 (10:1 in Figure 8) and lane 3 (5:1 in Figure 8) shows little difference with and without the restriction enzyme digestion of the complexes. This observation confirms that the copolymer can protect DNA from the enzyme digestion when N/P ratios are above unity. However, at or below the unity charge ratio, gel electrophoresis showed strong digestion (Figure 7). This is entirely consistent with significantly reduced Luc expressions when compared to the control experiments without treatment of the restriction enzyme digestion.
Langmuir, Vol. 24, No. 13, 2008 6887
Figure 8. Transfection efficiencies of copolymer/DNA (MPC30-DEA70/ Luc plasmid DNA) at different charge ratios. Blue columns represent transfection by polymer/DNA complexes, and purple columns represent transfection by polymer/DNA complexes that have been treated with the Not I, QG restriction enzyme for 1.5 h at 37 °C.
Figure 9. Cell (HEK293) transfection by copolymer/Luc plasmid DNA complexes at N/P ratios of 0.5:1, 1:1, 2:1, 5:1, and 10:1 at pH 7 and TransIT-LT1/Luc plasmid DNA complexes as positive control and plasmid only as negative control. (A) With 10% FBS and (B) without FBS. (Blue columns for 24 h and purple for 48 h.)
To better examine the effect of N/P ratios on cell transfection for these copolymer/plasmid DNA polyplexes, a more systematic study was carried out using the same cell line, and the results are shown in Figure 9. To obtain a useful perspective of the effectiveness of our own model copolymer vector, a comparison also was made with the popular commercial lipid transfection vector LT1. It can be seen from Figure 9 that the transfection rate goes down with decreasing N/P ratios due to the reduced population of positively charged complexes formed. The trans-
6888 Langmuir, Vol. 24, No. 13, 2008
fection results without FBS look better than those with 10% FBS, indicating that the presence of serum in the process of transfection has some negative effect. This may be due to the binding of complexes to some of the serum components. Furthermore, the transfection rates at 48 h look better than those at 24 h, with and without FBS. Overall, the transfection rates, even at the high N/P ratios, are relatively low when compared to the positive LT1 control, although this model copolymer works far more effectively than most other model polymeric vectors reported. One of the technical issues is how to increase the population of cationically charged complexes when using these copolymers. As described previously, even when the copolymers are in large excess, there is still a high proportion of net neutral polyplexes formed, as is evident from gel electrophoresis data. Our transfection results suggest that neutral polyplexes are far less effective. Although an increase in the copolymer excess can increase the portion of net positively charged complexes formed, this approach often leads to an increased cytotoxicity. The same issue also exists with LT1, but it was found in this work that LT1 often produces a transfection efficiency 10-50 times greater than our model copolymers under similar levels of cytotoxicity. Future work will need to address the structural design of the copolymer to promote its binding with DNA, taking into account findings from this study. The effect of solution pH on transfection efficiency was studied using MPC30-DPA60. We first used complexes formed between copolymer and DNA at pH 7. We note from the previous description that at pH 7, the copolymers self-aggregate to form micelles, and thus, complexation occurs between copolymer micelles and plasmid DNA. Because of weak cationic charges on each micelle and steric constraints, complexation between copolymer micelles and DNA molecules is weak and inefficient. Gel electrophoresis already indicated the net negative charge distribution on the complexes formed. Transfection from copolymer/DNA complexes formed at pH 7 resulted in a low transfection efficiency, consistent with results obtained from electrophoresis and SANS. However, when using complexes formed at pH 5, the expression of Luc was detected. At this pH, complexation did occur in the form of molecular interaction through the binding of a number of copolymers to a single DNA, and thus, more effective neutralization was achieved. The subsequent transfection study did result in better efficiency than what was observed using polyplexes formed at pH 7. However, transfection was still very low when compared to that obtained using MPC30-DEA70. This is because of a lack of polyplexes carrying net positive charges. Although some of the polyplexes formed at pH 5 carried net positive charges, they became net neutral or negative when the pH was brought back to 7 for transfection, making the transfection efficiency extremely low.
Zhao et al.
4. Conclusion In this study, we examined the structure of complexes formed between MPC-based diblock copolymers and Luc plasmid DNA by a combined approach of electrophoresis, DLS, and SANS. The results show that the complexation processes and the resulting size and shape of the polyplexes are affected by the hydrophobicity of the weak cationic blocks and their pH-dependent ionization. MPC30-DEA70 formed polyplexes with Luc plasmid DNA via electrostatic interactions and could protect the DNA from restriction enzyme digestion at N/P ratios above unity. Transfection results indicated that the transfection efficiency increased with N/P ratios. On the basis of these observations, we speculate that the transfection efficiency arose mainly from positively charged polyplexes. However, the fraction of positively charged polyplexes was quite low as compared to neutral polyplexes even when the copolymer was in high excess. The effect of solution pH on complexation and the subsequent transfection was well-demonstrated with MPC30-DPA60. This copolymer could form the same polyplexes with plasmid DNA as MPC30-DEA70 at pH 5. However, at pH 7, MPC30-DPA60 self-aggregated to form micelles because the pKa was 6.3. While a small amount of free copolymer molecules could complex with DNA as usual, the complexation between polymeric micelles and DNA at pH 7 was expected to be difficult. As the polyplexes formed were poorly neutralized, they contained net negative charges, leading to a very low transfection efficiency. It also was found that although complexation at pH 5 occurred between MPC30-DPA60 via molecular interactions and better neutralization occurred, the complexes became either neutral or negatively charged when the pH was shifted to 7 for transfection, making the transfection efficiency low again. As this part of the work demonstrated that the formation of polymeric micelles reduces complexation and transfection, it is essential to select polymeric vectors with pKa values close or above physiological pH. For a given copolymer vector, the charge ratio between transfecting vector and plasmid DNA is important in regulating transfection efficiency. Its alteration can lead to optimization between production of positively charged complexes and reduction of cytotoxicity, while ensuring efficient transfection. Acknowledgment. We thank Prof. Steve P. Armes and Dr. Andrew L. Lewis for provision of cationic copolymer samples and EPSRC and Impact Faraday Partnership for funding this work. We also thank the ISIS neutron facility for beam-time and Dr. R. K. Heenan for helpful advice. X.Z. thanks Biocompatibles UK Ltd. for a studentship. LA800593Q
