- Email: [email protected]
Transgene regulation system responding to Rho associated coiled-coil kinase (ROCK) activation
GENE DELIVERY
Journal of Controlled Release 155 (2011) 40–46
Contents lists available at ScienceDirect
Journal of Controlled Release j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / j c o n r e l
Transgene regulation system responding to Rho associated coiled-coil kinase (ROCK) activation Akira Tsuchiya a, Jeong-Hun Kang b, Daisuke Asai c, Takeshi Mori a, d, Takuro Niidome a, d, e, Yoshiki Katayama a, d, e,⁎ a
Graduate School of Systems Life Sciences, Kyushu University, 744 Motooka, Nishi-ku, Fukuoka, 819-0395, Japan Department of Biomedical Engineering, National Cerebral and Cardiovascular Center Research Institute, 5-7-1 Fujishiro-dai, Suita, Osaka 565-8565, Japan Department of Microbiology, St. Marianna University School of Medicine, 2-16-1 Sugao, Miyamae-ku, Kawasaki 216-8511, Japan d Department of Applied Chemistry, Faculty of Engineering, Kyushu University, 744 Motooka, Nishi-ku, Fukuoka 819-0395, Japan e Center for Future Chemistry, Kyushu University, 744 Motooka, Nishi-ku, Fukuoka 819-0395, Japan b c
a r t i c l e
i n f o
Article history: Received 12 July 2010 Accepted 1 May 2011 Available online 7 May 2011 Keywords: Gene delivery Rho associated coiled-coil kinase Protein kinase DDS
a b s t r a c t Recently, we have proposed a new system of gene regulation called ‘drug or gene delivery system responding to cellular signals’ (D-RECS). In this system, transgene expression is activated in response to intracellular target protein kinases or proteases for safe, cell-specific gene delivery by using peptide–polymer conjugates. Here we applied this system to an intracellular Rho-associated coiled-coil kinase (ROCK) signal, which is activated abnormally in cardiovascular diseases. A ROCK responsive polymer consisting of neutral polymers in main chain and cationic ROCK substrate peptides in side chains was prepared and could form the complex with plasmid DNA. The complex was transferred into NIH3T3 cells with or without L-α-lysophosphatidic acid (LPA) that increases ROCK activity. At an N/P ratio of 2.0, a significant increase of the gene expression was identified in LPA-treated NIH3T3 cells, but was disappeared in NIH3T3 cells treated with ROCK specific inhibitor, Y-27632. These results suggest that the ROCK responsive polymer can regulate gene expression in response to ROCK activity. © 2011 Elsevier B.V. All rights reserved.
1. Introduction Gene therapy has much potential as a therapeutic approach for many diseases. Cationic polymers or cationic lipids have been studied as gene carriers and a lot of carriers have been developed for in vitro and in vivo use [1–6]. For such treatments to be efficient and safe, these carriers must target therapeutic genes specifically into target disease cells and overcome barriers such as gene degradation by serum components, cellular uptake, endosomal escape, and release from carriers in the cytoplasm or nucleus [7,8]. Stimulus-responsive carriers, that change their properties with decreased pH, increased temperature and/or a changed redox microenvironment, are highly promising strategies
Abbreviations: ζ-potential, zeta-potential; DLS, dynamic light scattering; D-RECS, drug or gene delivery system responding to cellular signals; APS, ammonium persulfate; TEMED, N, N, N′, N′,-tetramethylethylenediamine; LPA, L-α-lysophosphatidic acid; MALDI-TOF-MS, matrix-assisted laser desorption ionization time-offlight mass spectrometry; N/P ratio, amine/phosphate ratio; PDI, polydispersity index; pDNA, plasmid DNA; RFI, relative fluorescence intensity; RLU, relative luminescence units; ROCK, Rho associated coiled-coil kinase; RT-PCR, real-time polymerase chain reaction. ⁎ Corresponding author at: Department of Applied Chemistry, Faculty of Engineering, Kyushu University, 744 Motooka, Nishi-ku, Fukuoka 819-0395, Japan. Tel./fax: + 81 92 802 2850. E-mail address: [email protected] (Y. Katayama). 0168-3659/$ – see front matter © 2011 Elsevier B.V. All rights reserved. doi:10.1016/j.jconrel.2011.05.002
towards developing an efficient gene delivery system [9,10]. In these strategies, therapeutic genes form complexes outside of disease cells that protect them from degradation. For instance, the pH in endosomes is more acidic than that in cytosol. Using a pH-sensitive linkage in the carrier, transgene escape from endosomes into the cytosol is facilitated, enhancing its expression drastically [11,12]. Thermo-sensitive polymers have also been used to exploit hyperthermia to exert spatial and temporal control over drug and gene delivery [13–15]. Similarly, redox microenvironment-responsive carriers are another possibility for the enhancement of gene expression [16,17]. Disulfide linkages in these carriers are cleaved in a reductive environment, triggering the release of genes into the cytosol. Although these systems enhance the disintegration of the carrier– gene complex, such stimuli often have insufficient cell specificity to properly target genes to their required site of action. Carriers may thus need an additional targeting ligand to confer cell specificity. Ideally, both cell specificity and accelerated expression functions should be integrated into a single, stimulus-responsive gene delivery system. We have developed a technology (‘drug or gene delivery system responding to cellular signals’, or D-RECS) that utilizes the abnormal activation of protein kinases as the trigger to release genes from cationic polymeric carriers. Protein kinases have essential roles in communicating signals from the cell surface to the nucleus in cells and thus initiate cellular responses such as cell death, proliferation and
migration. It has been frequently reported that abnormal activation of certain protein kinases is closely related to cancer, inflammation and other diseases. We aimed to exploit such abnormal activation of protein kinases to both ‘signpost’ disease cells and activate transgene expression. For this purpose, we required a polymer grafted with cationic substrate peptides which were phosphorylated by the activated target protein kinase. This cationic polymer should then form a complex with plasmid DNA (pDNA) through electrostatic interactions and efficiently suppress expression of the gene (Fig. 1A). Upon phosphorylation of the substrate peptides by disease-activated protein kinases in cells, pDNA is released from the polymer due to the disintegration of the electrostatic complex and gene expression is activated. In recent studies, we demonstrated transgene regulation in response to protein kinase Cα (PKCα) [18–20], c-AMP dependent protein kinase (PKA) [21] and IκB kinase β (IKKβ) [22] activities. Here, we chose the Rho associated coiled-coil kinase (ROCK) as the target kinase for this system. ROCK is a serine/threonine kinase and plays an important role in various cellular processes such as smooth muscle contraction, actin cytoskeleton organization, cell adhesion and motility, proliferation and cytokinesis [23–25]. These functions are involved in the pathogenesis of cardiovascular diseases [26], Alzheimer's disease [27] and cancer metastasis [28]. Especially, cardiovascular diseases are important diseases but there are little systems to delivery drug or genes to those diseases selectively. Therefore, ROCK is a key target for the therapy of these conditions.
2. Materials and methods 2.1. Synthesis of the substrate peptide In this study, a peptide RAKYKTLRQIR-NH2 was used as ROCK sensitive substrate and RAKYKAKRQIR-NH2 was used as a control peptide. The peptide substrate was originally developed after screening of peptide library (to be submitting). These peptides with a methacryloyl group at the amino-terminus were synthesized according to an Fmoc synthetic strategy. Peptides were cleaved from the resin by treatment with 94% trifluoroacetic acid (TFA) containing 2.5% triisopropylsilane, 2.5% water and 1.0% 1,2-ethanedithiol and purified with reverse-phase high performance liquid chromatography. Mass spectra were obtained by matrix-assisted laser desorption ionization time-of-
41
flight mass spectrometry (MALDI-TOF-MS) to verify the sequences of the synthesized peptides.
2.2. Phosphorylation assay For the substrate peptides, phosphorylation by ROCK (Carna Bioscience Incorporation, Kobe, Japan) was evaluated by means of MALDI-TOF-MS. Phosphorylation reactions were carried out in a total volume of 20 μl containing 20 mM Tris–HCl buffer (pH 7.4), 1 mM MgCl2, 100 μM ATP, 30 μM substrate peptide and 1 μg/ml ROCK2. The assay mixture was incubated at 37 °C for 1 h. The reaction was quenched by addition of 4 μl of 0.1% TFA containing α-cyano-4-hydroxycinnamic acid matrix in 50% water/acetonitrile to 1 μl of the reaction mixture. An aliquot of the mixture (2 μl) was applied to the sample plate, dried to induce crystallization and analyzed by MALDI-TOF-MS.
2.3. Synthesis of the ROCK-responsive polymer The polymer was synthesized in a manner similar to that described previously [18–20] (Fig. 1B). Briefly, a methacryloyl peptide and acrylamide were dissolved in degassed water and allowed to stand at room temperature for 90 min after the addition of ammonium persulfate (APS) and N, N, N′, N′,-tetramethylethylenediamine (TEMED) as the redox initiator couple. The product was purified by dialysis against water for 3 days. The content of the peptides was calculated by elemental analysis and the molecular weight of the polymer was estimated by gel-permeation chromatography using a polyethyleneglycol standard.
2.4. Preparation of pDNA A pDNA containing a firefly luciferase cDNA driven by a CMV promoter was prepared following our previous report [18–20] and was used in all experiments without a cell-free experiment. A pDNA containing a firefly luciferase cDNA driven by a T7 promoter was used in the cell-free expression experiment.
2.5. Dynamic light scattering (DLS) measurement Complexes of the polymer and pDNA at various amine/phosphate (N/P) ratios were prepared by mixing at room temperature for 15 min. The final concentration of pDNA was adjusted to 2.5 μg/ml using 10 mM HEPES buffer (pH 7.4) or PBS (pH 7.4). The diameter and zeta (ζ)-potential of the complexes were determined using a Zetasizer (Malvern Instruments, Worcestershire, UK) with the He/Ne laser at a detection angle of 173° at 25 °C. For the evaluation of complex disintegration during phosphorylation, we prepared the complex by the same method and the final concentration of pDNA was adjusted to 2.5 μg/ml in buffer containing 10 mM HEPES (pH 7.4), 1 mM MgCl2, 200 μM ATP and 2 μg/ml activated or inactivated ROCK2. Inactivation of ROCK2 was performed by heating at 97 °C for 3 min. Dynamic light scattering intensity was then monitored throughout the ROCK reaction.
2.6. Gel electrophoresis retardation assay
Fig. 1. (A) ROCK-responsive gene regulation system. Co-polymers grafted to cationic substrate peptides form complexes with plasmid DNA through electrostatic interactions and suppress gene transcription. Phosphorylation of the grafted substrate peptides in the polymer by activated ROCK in disease cells causes disintegration of the complex and allowing activation of the transgene. (B) Schematic of the chemical reaction whereby substrate peptide is grafted to the polymer.
pDNA (0.3 μg) and various concentrations of polymers were incubated at room temperature for 15 min and then, incubated at 37 °C for 1 h with 20% fetal bovine serum (FBS). One hundred equivalent poly anion (poly (sodium 4-styrenesulfonate)) to pDNA was added to the solutions to exclude pDNA from polymers and the samples were analyzed by electrophoresis using 1% agarose gel at 100 V for 25 min.
GENE DELIVERY
A. Tsuchiya et al. / Journal of Controlled Release 155 (2011) 40–46
GENE DELIVERY
42
A. Tsuchiya et al. / Journal of Controlled Release 155 (2011) 40–46
2.7. Ethidium bromide (EtBr) exclusion assay
2.12. Peptide phosphorylation cellular reporter assay
Complexes between the polymer and pDNA were prepared at various N/P ratios in sterilized water in the presence of EtBr. Solutions were incubated at room temperature for 15 min. The final concentrations of pDNA and EtBr were adjusted to 5 μg/ml and 12.5 μg/ml, respectively, with 10 mM HEPES buffer (pH 7.4). Fluorescence measurements of sample solutions were carried out at 25 °C using the ARVO multilevel counter (Wallac Incorporation, Turku, Finland). Excitation and emission wavelengths were 530 nm and 590 nm, respectively. The relative fluorescence intensity (RFI) was determined using the following equation:
To stimulate ROCK activation, cells were stimulated with 20 μM L-αlysophosphatidic acid (LPA) for 6 h, then scraped and lysed in 100 μl of lysis buffer (20 mM Tris–HCl, pH 7.4, 0.05% Triton-X 100, 2 mM EDTA). After centrifuging the sample at 15,000 g at 4 °C for 10 min, a 10 μl aliquot of the supernatant was used to measure chemiluminescence in a MiniLumat LP9506 (EG & G Berthold, Wildbad, Germany) directly after adding 40 μl of luciferase assay solution. The results are presented as relative luminescence units (RLU)/mg protein.
RFIð%Þ = ðFobs −Fe Þ = ðF0 −Fe Þ × 100
Cells were prepared for transfection as above. Four hours after transfection, the medium was replaced with DMEM and cells were incubated for a further 18 h. The cultured cells were then scraped and lysed in 200 μl of lysis buffer (20 mM Tris–HCl, pH 7.4, 0.05% Triton-X 100, 2 mM EDTA). After centrifuging the sample at 15,000 g at 4 °C for 10 min, pDNA was extracted from cell lysates using DNeasy Blood & Tissue Kits (Qiagen GmbH, Hilden, Germany). Quantitative analysis of pDNA was performed by real time PCR in a protocol similar to our previous report [29]. Sequences of PCR primers were as follows: forward primer, 5′-gcgccttatccggtaactatc-3′; reverse primer, 3′-acttcaccaccggattgatgc-5′. CYBR Green I purchased from Takara (Tokyo, Japan) was used as a fluorescent probe. PCR conditions were: 95 °C for 1 min, followed by 40 cycles at 94 °C for 15 s, 60 °C for 15 s and 72 °C for 30 s.
where Fobs, Fe and F0 are the fluorescence intensities of the polymer– pDNA complexes at each N/P ratio; EtBr without pDNA; and EtBr/pDNA without the polymer, respectively. 2.8. Cell-free gene expression assay Luciferase expression by a cell-free system was carried out using the T7 Quick Coupled Transcription/Translation System (Promega, Madison, WI, USA) as described [18]. Copolymer/pDNA complexes at N/P ratios of 1.0, 2.0 and 4.0 were prepared by mixing both compounds for 15 min. Final volumes were adjusted 10 μl by 10 mM HEPES buffer (pH 7.4). Detection of luciferase expression was according to manufacturer's protocols. Naked-pDNA (0.3 μg) was the controls. To monitor chemiluminescence, 20 μl of reaction solutions was added to the luciferase assay solution (Promega) and luminescence intensity measured by the ARVO multilabeled counter. 2.9. Monitoring complex phosphorylation using a coupled enzyme assay Phosphorylation on the pendant peptide in the polymer–pDNA complex was monitored using a coupled enzyme assay [18]. The complex or the peptide itself was diluted to 100 μl (final concentration of the peptide unit was 30 μM) in 10 mM HEPES (pH 7.4) containing 1 mM phosphoenolpyruvate, 10 U/μl lactate dehydrogenase (LDH), 4 U/μl pyruvate kinase, 200 μM ATP, 1 mM MgCl2, 300 μM nicotinamide adenine dinucleotide (NADH) and 2.0 μg/ml ROCK2. The reaction was carried out at 25 °C. The consumption of NADH was detected by monitoring absorbance at 340 nm with a UV/Vis spectrometer (UV2550; Shimadzu, Tokyo, Japan) equipped with an SPR-8 temperature controller (Shimadzu).
2.13. Real time (RT)-PCR
3. Results and discussion 3.1. Synthesis of the peptide-grafted polymer With five cationic amino acids making it highly suitable for D-RECS, we used the peptide RAKYKTLRQIR-NH2 as the substrate for ROCK. Evaluation using MALDI-TOF-MS confirmed this peptide as an effective ROCK substrate (Fig. 2). The polymer-grafted substrate peptides were then synthesized by radical polymerization using methacryloyl-peptide monomer and acrylamide (Fig. 1B). The molecular weight of the
2.10. Cell culture The NIH3T3 cell line was purchased from Health Science Research Resources Bank (Tokyo, Japan). Cells were cultured in Dulbecco's modified Eagle's medium (DMEM) (Wako, Osaka, Japan) containing 10% FBS, 100 U/ml penicillin, 100 μg/ml streptomycin and 0.25 μg/ml amphotericin-B (all from Invitrogen, New York, USA). Cells were incubated in a humidified atmosphere of 5% CO2 and 95% air at 37 °C. 2.11. Transfection of complexes into cells NIH 3T3 cells were cultured in on 24-well culture plates (20,000 cells/well) at 37 °C for 18 h in 0.5 ml DMEM containing 10% FBS. The medium was changed to serum-free DMEM followed by a further 4 h incubated at 37 °C. The medium was then replaced with 500 μl of Opti-MEM (Invitrogen) containing the complexes (prepared as for DLS assay, above) at N/P ratios of 1.0, 2.0 or 4.0 (pDNA remained constant at 2.0 μg/ml) and cells were incubated at 37 °C for 4 h. The medium was then changed to serum-free DMEM followed by a further 14 h incubation at 37 °C before assaying.
Fig. 2. MALDI-TOF-MS spectra of the substrate peptide before (A) and after (B) phosphorylation by ROCK.
43
resultant polymer was ~130 kDa, with 2.9% methacrylated peptide incorporation. 3.2. Characteristics of the polymer–pDNA complex The diameter and ζ-potential of the complexes are important factors in efficient gene delivery. The suitable diameter of the complex to transfer pDNA into cells via endocytosis is around 100 nm. After mixing the polymer and pDNA for 15 min, diameter and ζ-potential of the complex was measured at N/P ratios of 1.0, 2.0 and 4.0 (Table 1) using DLS. The particle size of this complex decreased as N/P ratios increased from 1.0 to 2.0, becoming constant at around 110 nm in 10 mM HEPES buffer (pH 7.4). This value is smaller than that for our previously reported D-RECS complexes (150–200 nm) [18,20,21]. Particle size at an N/P ratio of 1.0 (229 ± 85.4 nm) was also much smaller than that in ordinary D-RECS complexes, which tend to aggregate into very large (μm range) complexes with pDNA at an N/P ratio of 1.0. These results indicate that the polymer formed tighter complexes with pDNA than do ordinary D-RECS polymers. The diameters of these complexes were within a suitable range to transfer into cells via endocytosis. We then investigated the condensation of pDNA with the cationic polymer using the EtBr exclusion assay. Intercalation of EtBr into the DNA strand increases its fluorescence intensity drastically. Conversely, complexation of the DNA during formation of electrostatic complexes with polycations releases the intercalated EtBr, decreasing its fluorescence intensity. Exploiting this phenomenon, the condensation of DNA has been evaluated in cationic polymer–DNA complexes [30]. EtBr fluorescence intensity in both HEPES buffer (pH 7.4) and PBS (pH 7.4) was measured for each of the complexes formed at different N/P ratios in this study (Table 1). The polymer decreased the EtBr fluorescence intensity by 70% at N/P ratios above 2.0. Furthermore, the values both in HEPES buffer and in PBS were very similar at N/P ratios of above 1.0. This is notable because high salt concentrations such as in PBS or saline tend to cause disintegration of ordinary D-RECS polymer–pDNA complexes. Next, using a cell-free expression system, we evaluated the suppression of gene expression resulting from complexation of the pDNA with the polymer at various N/P ratios. For safe gene therapy, it is critical to minimize gene expression in non-diseased cells to avoid any undesirable side effects. At an N/P ratio of 1.0, the gene expression was suppressed by 40% compared with uncomplexed pDNA (Fig. 3B). However, at N/P ratios of 2.0 and 4.0, gene expression was suppressed by N95% relative to uncomplexed pDNA. Taken together with the results in Table 1 indicating that complex size decreases as N/P ratio increases, this suggests that smaller, tighter complexes produce more effective suppression of gene expression.
Fig. 3. Characterization of the polymer–pDNA complexes. (A) Condensation of pDNA by forming the complex with polymer in the EtBr exclusion assay. Assays were performed in either HEPES buffer or PBS. Data represent RFI of EtBr–DNA interaction at increasing N/P ratios. Decreasing RFI corresponds to elevated DNA condensation as EtBr is excluded from the complex. Data are mean ± SEM for three experiments (B) Luciferase expression from the polymer–pDNA complex at each N/P ratio in a cell-free expression system. Complexes were prepared by mixing pDNA with the polymers. RLUs were calculated by normalizing the luminescence for each complex to that of pDNA alone. Data are mean ± SEM for three experiments.
This decrease in A340 illustrates that grafting the peptide into the polymer chain did not affect its status as a ROCK phosphorylation target. Using DLS, we then evaluated whether the complex disintegrated following phosphorylation by activated ROCK (Fig. 5). At an N/P ratio of 2.0, introducing activated ROCK into the complex solution led to an increase in the particle diameter (from 200 nm to 2000 nm) and a decrease in the count rate (from 6300 kcps to 4100 kcps) within 60 min in phosphorylation reaction solution. In contrast, inactivated ROCK did not alter either the hydrodynamic diameter or the count rate of the complex over the same period. Furthermore, the polydispersity index (PDI) was increased by the addition of activated ROCK (data not shown).
3.3. Phosphorylation and disintegration of the complex In D-RECS, kinase phosphorylation of substrate peptides grafted into the polymer causes complex disintegration, which activates gene expression. To verify that the substrate peptide used here remained a phosphorylation target for ROCK when grafted into the polymer, we used a coupled enzyme assay to assess peptide phosphorylation by ROCK in the polymer–pDNA complex (Fig. 4). Phosphorylation of uncomplexed peptide and polymer–pDNA complex (at N/P ratios of 2.0 and 4.0) was measured, with peptide content in each condition adjusted to 30 μM. Uncomplexed and complexed peptide were phosphorylated comparably, as shown by a similar time course in the decrease in A340. Table 1 Diameter and ζ-potential of polymer/pDNA complexes. N/P ratio
1.0
2.0
4.0
Diameter (nm) ζ-potential (mV)
229.4 ± 85.4 5.9 ± 2.4
110.8 ± 0.6 14.1 ± 2.3
109.4 ± 2.2 14.3 ± 1.0
Fig. 4. Phosphorylation time course of the peptide (black) and the polymer–pDNA complexes at N/P ratios of 2.0 (dark gray) and 4.0 (light gray) monitored by coupled enzyme assay. Consumption of NADH, measured by changes in the absorbance at 340 nm, (A340) was used as a reporter for the phosphorylation status of the polymer.
GENE DELIVERY
A. Tsuchiya et al. / Journal of Controlled Release 155 (2011) 40–46
GENE DELIVERY
44
A. Tsuchiya et al. / Journal of Controlled Release 155 (2011) 40–46
Fig. 5. DLS measurements of the polymer–pDNA complexes in a solution containing either activated or inactivated ROCK. Diameters (A) and count rates (B) of the complex during the phosphorylation by ROCK at an N/P ratio of 2.0 (circles) and 4.0 (squares). Open and closed symbols represent the complex in the presence of activated ROCK and inactivated ROCK, respectively.
Fig. 6. DLS measurements of the polymer–pDNA complexes in PBS. Shown are time courses showing the change in diameter (A) and count rate (B) of the complex at N/P ratios of 1.0 (circles), 2.0 (squares) and 4.0(triangles).
serum. This result suggested that pDNA in serum was little degraded but kept its formation. These results suggest that the complex at an N/P ratio of 2.0 was disintegrated in response to activated ROCK. We showed that complexes formed at an N/P ratio of 4.0 were viable phosphorylation substrates for activated ROCK (Fig. 4). However, at an N/P ratio of 4.0, the diameter and the count rate of the complex stayed constant over 60 min, even in the presence of activated ROCK (Fig. 5). This suggests that at high N/P ratios the complex does not disintegrate following phosphorylation of the substrate peptide. 3.4. Stability of the polymer–pDNA complex To use a polymer–DNA polyplex for gene delivery in vivo, it must remain stable until it reaches its target. If the complex is broken or aggregated in the blood stream or in normal cells, efficient gene delivery cannot be achieved. In this context, the stability of the complex was evaluated using DLS to measure diameter and count rate of complexes in 150 mM saline (Fig. 6). At an N/P ratio of 1.0, the diameter of the complex increased from 250 nm to 700 nm and the count rate decreased from 2000 kcps to 800 kcps. At an N/P ratio of 2.0, the diameter change was negligible in the presence of saline although the count rate increased from 2800 kcps to 4000 kcps. At an N/P ratio of 4.0, both diameter and count rate were unchanged. We conclude that at low N/P ratio the polymer forms an ‘open’ complex with pDNA that can easily disintegrate and aggregate in the presence of saline. However, at N/P ratios above 2.0, the complex was increasingly stable. Next, we evaluated the protection of pDNA from nuclease in serum. Polymer–pDNA complexes were prepared at N/P ratios of 1.0, 2.0 and 4.0 and mixed with 20% serum in 10 mM HEPES buffer. Degradation of pDNA was evaluated by gel electrophoresis assay. In the case of only pDNA, as a control, pDNA was degraded perfectly (Fig. 7). In 20% serum, forming complex with polymers, pDNA was obtained which formation were different from that in the absence of
3.5. In vitro transgene regulation It is known that ROCK is activated in NIH3T3 cells treated with LPA [31] and we also found ROCK-medicated actin stress fiber formation (Fig. S1). We used this system as a model system of ROCK activating cells to investigate whether transgene expression in the polymer–pDNA complex was regulated in response to ROCK activation. A basal level of transgene expression was seen following addition of the complex at all N/P ratios, but decreased as N/P ratio increased (Fig. 8A), reflecting the decreasing propensity of the complex to disintegrate spontaneously at higher N/P ratios. In fact, the highest gene expression was obtained at an N/P ratio of 1.0 and the lowest was obtained at an N/P ratio of 4.0. This propensity was conformed to the result of cell-free expression system. Importantly, at an N/P ratio of 2.0, elevated transgene expression was observed in response to LPA, whereas in the presence of 20 μM ROCK inhibitor, Y-27632, or using control polymers that were exchanged Thr residue to Ala residue, there were no such increase in gene expression
Fig. 7. Protection of pDNA degradation from nuclease in FBS. Complexes were incubated with 20% FBS and then, degradation of pDNA was evaluated by gel electrophoresis shift assay.
45
Furthermore, we investigated whether the absence of any transgene expression in cells exposed to complexes with an N/P ratio of 4.0 was due to impaired uptake of these complexes. Using real-time PCR, we quantified the pDNA transferred into cells at each N/P ratio (1.0, 2.0 and 4.0). Cellular uptake of the complex was highest at an N/P ratio of 1.0 but, although uptake at an N/P of 4.0 was lower, it was comparable to that for N/P ratio 2.0 (Fig. 8B). This suggests that the low level of gene expression in the complex at an N/P ratio of 4.0 is not caused by poor uptake. Taken together, our data support a conclusion that this effect is most likely due to these N/P ratio 4.0 complexes not disintegrating properly. In this scenario, despite being properly endocytosed (Fig. 8B) and phosphorylated (Fig. 4), these complexes are simply too stable to release the transgene from the complex, as suggested by our stability and disintegration data (Figs. 6 and 5, respectively). This hyperstable complex thus cannot express the luciferase gene under either basal or stimulated conditions. On the other hand, complexes at an N/P ratio of 1.0 were not suppressed the gene expression without LPA treating because of that low stability. To get responsibility of the activation of ROCK using ROCK responsive polymer, it is important that we select the optimal N/P ratios. 4. Conclusion
Fig. 8. Transfection of polymer–pDNA complexes into NIH3T3 cells. Complexes were prepared by mixing pDNA with the polymer at N/P ratios of 1.0, 2.0 and 4.0. (A) Luciferase expression of each complex in the absence (black bar) or presence (white bar) of LPA stimulation. As a control, the polymer that Thr residues exchanged to Ala residues was used. Data are presented as the RLU/mg protein, mean ± SEM for six experiments. *P b 0.05 (calculated using t-test). (B) The amounts of pDNA (μg/mg protein) transfected into NIH3T3 cells at each N/P ratio of complex. pDNA was purified using DNeasy blood & tissue kits and quantitative analysis of pDNA was performed by real time PCR. Data represent mean expression level ± SEM for three experiments.
(Fig. 8A). At an N/P ratio of 2.0, when substrate peptides were phosphorylated by ROCK, the complex was disintegrated (Fig. 5). The same phenomenon might be observed in vitro. The complex was phosphorylated in cells which treated with LPA and ROCK was activated, and then the gene expression was activated. Because the gene expression was not activated in the case of treatment with ROCKspecific inhibitor and using control polymers, this activation of the gene expression was depended to ROCK activity and caused by phosphorylation of the complex. The consensus sequence of the ROCK phosphorylation site is considered to be R/KXXS/T or R/KXS/T (R, arginine; K, lysine; X, any amino acid; S, serine; and T, threonine) [25]. These sequences are similar to consensus phosphorylation site motifs for PKC, which were identified to be R/KXXS/T, R/KXS/T, R/KXXS/TXR/K, or R/KXS/TXR/K [32,33]. Thus, we examined whether PKCα-responsive polymer can increase gene expression in the LPA-treated NIH3T3 cells. PKCαresponsive polymer (PPC (S)) and its control polymer (PPC (A)) [18–20] showed no gene expression in the LPA-treated NIH3T3 cells (Fig. S2). Previous studies suggested that PKCα is not involved in LPA-induced actin stress fiber formation [34] and high level of LPA inhibits PKCα activity [35]. These results mean that the gene expression in ROCK substrate-grafted polymer system is solely due to ROCK activity, but not PKCα activity.
We report here a novel gene delivery regulation system that can activate transgene expression in response to ROCK signals. The polymer grafted with ROCK substrate peptides formed a more stable complex with pDNA than ordinary D-RECS systems. This should confer a strong resistance to complex disintegration in serum, blood, or non-target cells, although the responsiveness to ROCK is also lost if the N/P ratio is too high. We have successfully optimized ROCK-responsive transgene activation at an N/P ratio of 2.0. This system may have key applications for therapy and diagnostics in diseases where ROCK activation is observed. Acknowledgments This work was financially supported by a grant-in-aid for Scientific Research from the Ministry of Education, Science, Sports, and Culture in Japan and also supported by A3 Foresight Program in Japan Society for the Promotion of Science. A. T. is grateful to Japan Society for the Promotion of Science (JSPS) for the DC scholarship. Appendix A. Supplementary data Supplementary data to this article can be found online at doi:10.1016/j.jconrel.2011.05.002. References [1] J. Luten, C.F. Nostrum, S.C.D. Smedt, W.E. Hennink, Biodegradable polymers as non-viral carriers for plasmid DNA delivery, J. Control. Release 126 (2008) 97–110. [2] M. Ruponen, S. Arkko, A. Urtti, M. Reinisalo, V.P. Ranta, Intracellular DNA release and elimination correlated poorly with transgene expression after non-viral transfection, J. Control. Release 146 (2009) 226–231. [3] O. Jeon, H.S. Yang, T.J. Lee, B.S. Kim, Heparin-conjugated polyethyleneimine for gene delivery, J. Control. Release 132 (2008) 236–242. [4] W.H. Kong, D.K. Sung, Y.H. Shim, K.H. Bae, P. Dubois, T.G. Park, J.H. Kim, S.W. Seo, Efficient intracellular siRNA delivery strategy through rapid and simple two steps mixing involving noncovalent post-PEGylation, J. Control. Release 138 (2009) 141–147. [5] P.R. Dash, V. Toncheva, E. Schacht, L.W. Seymour, Synthetic polymers for vectorial delivery of DNA: characterization of polymer–DNA complexes by photon correlation spectroscopy and stability to nuclease degradation and disruption by polyanions in vitro, J. Control. Release 48 (1997) 269–276. [6] A.E. Aneed, An overview of current delivery systems in cancer gene therapy, J. Control. Release 94 (2004) 1–14. [7] V. Sokolova, M. Epple, Inorganic nanoparticles as carriers of nucleic acids into cells, Angew. Chem. Int. Ed. 46 (2007) 2–16.
GENE DELIVERY
A. Tsuchiya et al. / Journal of Controlled Release 155 (2011) 40–46
GENE DELIVERY
46
A. Tsuchiya et al. / Journal of Controlled Release 155 (2011) 40–46
[8] I.A. Khalil, K. Kogure, H. Akita, H. Harashima, Uptake pathways and subsequent intracellular trafficking in nonviral gene delivery, Pharmacol. Rev. 58 (2006) 32–45. [9] S. Ganta, H. Devalapally, A. Shahiwala, M. Amiji, A review of stimuli-responsive nanocarriers for drug and gene delivery, J. Control. Release 126 (2008) 187–204. [10] N. Rapoport, Physical stimuli-responsive polymeric micelles for anti-cancer drug delivery, Prog. Polym. Sci. 32 (2007) 962–990. [11] M. Meyer, A. Philipp, R. Oskuee, C. Schmidt, E. Wagner, Breathing life into polycations, functionalization with pH-responsive endosomolytic peptides and polyethylene glycol enables siRNA delivery, J. Am. Chem. Soc. 130 (2008) 3272–3273. [12] Y. Lee, T. Ishii, H. Cabral, H.J. Kim, J.H. Seo, N. Nishiyama, H. Oshima, K. Osada, K. Kataoka, Charge-conversional polyionic complex micelles — efficient nanocarriers for protein delivery into cytoplasm, Angew. Chem. Int. Ed. 48 (2009) 1–5. [13] R. Namgung, S. Nam, S.K. Kim, S. Son, K. Singha, J.S. Kwon, Y. Ahn, M.H. Jeong, I.K. Park, V.K. Garripelli, S. Jo, An acid-labile temperature-responsive sol–gel reversible polymer for enhanced gene delivery to the myocardium and skeletal muscle cells, Biomaterials 30 (2009) 5225–5233. [14] S.H. Lee, S.H. Choi, S.H. Kim, T.G. Park, Thermally sensitive cationic polymer nanocapsules for specific cytosolic delivery and efficient gene silencing of siRNA: swelling induced physical disruption of endosome by cold shock, J. Control. Release 125 (2008) 25–32. [15] J.E. Chung, M. Yokoyama, T. Okano, Inner core segment design for drug delivery control of thermo-responsive polymeric micelles, J. Control. Release 65 (2000) 93–103. [16] M. Breunig, C. Hozsa, U. Lungwitz, K. Watanabe, I. Umeda, H. Kato, A. Goepferich, Mechanistic investigation of poly(ethylene imine)-based siRNA delivery: disulfide bonds boost intracellular release of the cargo, J. Control. Release 130 (2008) 57–63. [17] O. Taratula, O.B. Garbuzenko, P. Kirkpatrick, I. Pandya, R. Savla, V.P. Pozharov, H. He, T. Minko, Surface-engineered targeted PPI dendrimer for efficient intracellular and intratumoral siRNA delivery, J. Control. Release 184 (2009) 284–293. [18] R. Toita, J.H. Kang, J.H. Kim, T. Tomiyama, T. Mori, T. Niidome, B. Jun, Y. Katayama, Protein kinase Cα-specific peptide substrate graft-type copolymer for cancer cellspecific gene regulation systems, J. Control. Release 139 (2009) 133–139. [19] T. Tomiyama, J.H. Kang, R. Toita, T. Niidome, Y. Katayama, Protein kinase Cαresponsive polymeric carrier: its application for gene delivery into human cancers, Cancer Sci. 100 (2009) 1532–1536. [20] J.H. Kang, D. Asai, J.H. Kim, T. Mori, R. Toita, T. Tomiyama, Y. Asami, J. Oishi, Y.T. Sato, T. Niidome, B. Jun, H. Nakashima, Y. Katayama, Design of polymeric carriers for cancer-specific gene targeting: utilization of abnormal protein kinase Cα activation in cancer cells, J. Am. Chem. Soc. 130 (2008) 14906–14907.
[21] J. Oishi, K. Kawamura, J.H. Kang, K. Kodama, T. Sonoda, M. Murata, T. Niidome, Y. Katayama, An intracellular kinase signal-responsive gene carrier for disordered cell-specific gene therapy, J. Control. Release 110 (2006) 431–436. [22] D. Asai, A. Tsuchiya, J.H. Kang, K. Kawamura, J. Oishi, T. Mori, T. Niidome, Y. Shoji, H. Nakashima, Y. Katayama, Inflammatory cell-specific transgene expression system responding to Iκ-B kinase beta activation, J. Gene Med. 11 (2009) 624–632. [23] K. Riento, A.J. Ridley, ROCKS: multifunctional kinases in cell behavior, Mol. Cell. Biol. 4 (2003) 446–456. [24] N.P. Tsai, L.N. Wei, RhoA/ROCK1 signaling regulates stress granule formation and apoptosis, Cell. Signal. 22 (2010) 668–675. [25] J.H. Kang, Y. Jiang, R. Toita, J. Oishi, K. Kawamura, A. Han, T. Mori, T. Niidome, M. Ishida, K. Tatematsu, K. Tanizawa, Y. Katayama, Phosphorylation of Rhoassociated kinase (Rho-kinase/ROCK/ROK) substrates by protein kinases A and C, Biochimie 89 (2007) 39–47. [26] H. Shimokawa, M. Rashid, Development of Rho-kinase inhibitors for cardiovascular medicine, Trends Pharmacol. Sci. 28 (2007) 296–302. [27] A. Salminen, T. Suuronen, K. Kaarniranta, ROCK, PAK, and Toll of synapses in Alzheimer's disease, Biochem. Biophys. Res. Commun. 371 (2008) 587–590. [28] K. Itoh, K. Yoshioka, H. Akedo, M. Uehata, T. Ishizaki, S. Narumiya, An essential part for Rho-associated kinase in the transcellular invasion of tumor cells, Nat. Med. 5 (1999) 221–225. [29] T. Kawano, M. Yamagata, H. Takahashi, Y. Niidome, S. Yamada, Y. Katayama, T. Niidome, Stabilizing of plasmid DNA in vivo by PEG-modified cationic gold nanoparticles and the gene expression assisted with electrical pulses, J. Control. Release 111 (2006) 382–389. [30] K. Miyata, S. Fukushima, N. Nishiyama, Y. Yamasaki, K. Kataoka, PEG-based block catiomers possessing DNA anchoring and endosomal escaping functions to form polyplex micelles with improved stability and high transfection efficacy, J. Control. Release 122 (2007) 252–260. [31] M.P. Iwanicki, T. Vomastek, R.W. Tilghman, K.H. Martin, J. Banerjee, P.B. Wedegaertner, J.T. Parsons, FAK, PDZ-RhoGEF and ROCKII cooperate to regulate adhesion movement and trailing-edge retraction in fibroblasts, J. Cell Sci. 121 (2008) 895–905. [32] R.B. Pearson, B.E. Kemp, Protein kinase phosphorylation site sequences specificity motifs: tabulations, Method Emzymol. 200 (1991) 62–81. [33] H. Hug, T.F. Sarre, Protein kinase C isoenzymes: divergence in signal transduction? Biochem. J. 291 (1993) 329–343. [34] K. Riento, N. Totty, P. Villalonga, R. Garg, R. Guasch, A.J. Ridley, RhoE function is regulated by ROCK I-mediated phosphorylation, EMBO J. 24 (2005) 1170–1180. [35] J.J. Sando, O.I. Chertihin, Activation of protein kinase C by lysophosphatidic acid: dependence on composition of phospholipid vesicles, Biochem. J. 317 (1996) 583–588.
Journal of Controlled Release 155 (2011) 40–46
Contents lists available at ScienceDirect
Journal of Controlled Release j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / j c o n r e l
Transgene regulation system responding to Rho associated coiled-coil kinase (ROCK) activation Akira Tsuchiya a, Jeong-Hun Kang b, Daisuke Asai c, Takeshi Mori a, d, Takuro Niidome a, d, e, Yoshiki Katayama a, d, e,⁎ a
Graduate School of Systems Life Sciences, Kyushu University, 744 Motooka, Nishi-ku, Fukuoka, 819-0395, Japan Department of Biomedical Engineering, National Cerebral and Cardiovascular Center Research Institute, 5-7-1 Fujishiro-dai, Suita, Osaka 565-8565, Japan Department of Microbiology, St. Marianna University School of Medicine, 2-16-1 Sugao, Miyamae-ku, Kawasaki 216-8511, Japan d Department of Applied Chemistry, Faculty of Engineering, Kyushu University, 744 Motooka, Nishi-ku, Fukuoka 819-0395, Japan e Center for Future Chemistry, Kyushu University, 744 Motooka, Nishi-ku, Fukuoka 819-0395, Japan b c
a r t i c l e
i n f o
Article history: Received 12 July 2010 Accepted 1 May 2011 Available online 7 May 2011 Keywords: Gene delivery Rho associated coiled-coil kinase Protein kinase DDS
a b s t r a c t Recently, we have proposed a new system of gene regulation called ‘drug or gene delivery system responding to cellular signals’ (D-RECS). In this system, transgene expression is activated in response to intracellular target protein kinases or proteases for safe, cell-specific gene delivery by using peptide–polymer conjugates. Here we applied this system to an intracellular Rho-associated coiled-coil kinase (ROCK) signal, which is activated abnormally in cardiovascular diseases. A ROCK responsive polymer consisting of neutral polymers in main chain and cationic ROCK substrate peptides in side chains was prepared and could form the complex with plasmid DNA. The complex was transferred into NIH3T3 cells with or without L-α-lysophosphatidic acid (LPA) that increases ROCK activity. At an N/P ratio of 2.0, a significant increase of the gene expression was identified in LPA-treated NIH3T3 cells, but was disappeared in NIH3T3 cells treated with ROCK specific inhibitor, Y-27632. These results suggest that the ROCK responsive polymer can regulate gene expression in response to ROCK activity. © 2011 Elsevier B.V. All rights reserved.
1. Introduction Gene therapy has much potential as a therapeutic approach for many diseases. Cationic polymers or cationic lipids have been studied as gene carriers and a lot of carriers have been developed for in vitro and in vivo use [1–6]. For such treatments to be efficient and safe, these carriers must target therapeutic genes specifically into target disease cells and overcome barriers such as gene degradation by serum components, cellular uptake, endosomal escape, and release from carriers in the cytoplasm or nucleus [7,8]. Stimulus-responsive carriers, that change their properties with decreased pH, increased temperature and/or a changed redox microenvironment, are highly promising strategies
Abbreviations: ζ-potential, zeta-potential; DLS, dynamic light scattering; D-RECS, drug or gene delivery system responding to cellular signals; APS, ammonium persulfate; TEMED, N, N, N′, N′,-tetramethylethylenediamine; LPA, L-α-lysophosphatidic acid; MALDI-TOF-MS, matrix-assisted laser desorption ionization time-offlight mass spectrometry; N/P ratio, amine/phosphate ratio; PDI, polydispersity index; pDNA, plasmid DNA; RFI, relative fluorescence intensity; RLU, relative luminescence units; ROCK, Rho associated coiled-coil kinase; RT-PCR, real-time polymerase chain reaction. ⁎ Corresponding author at: Department of Applied Chemistry, Faculty of Engineering, Kyushu University, 744 Motooka, Nishi-ku, Fukuoka 819-0395, Japan. Tel./fax: + 81 92 802 2850. E-mail address: [email protected] (Y. Katayama). 0168-3659/$ – see front matter © 2011 Elsevier B.V. All rights reserved. doi:10.1016/j.jconrel.2011.05.002
towards developing an efficient gene delivery system [9,10]. In these strategies, therapeutic genes form complexes outside of disease cells that protect them from degradation. For instance, the pH in endosomes is more acidic than that in cytosol. Using a pH-sensitive linkage in the carrier, transgene escape from endosomes into the cytosol is facilitated, enhancing its expression drastically [11,12]. Thermo-sensitive polymers have also been used to exploit hyperthermia to exert spatial and temporal control over drug and gene delivery [13–15]. Similarly, redox microenvironment-responsive carriers are another possibility for the enhancement of gene expression [16,17]. Disulfide linkages in these carriers are cleaved in a reductive environment, triggering the release of genes into the cytosol. Although these systems enhance the disintegration of the carrier– gene complex, such stimuli often have insufficient cell specificity to properly target genes to their required site of action. Carriers may thus need an additional targeting ligand to confer cell specificity. Ideally, both cell specificity and accelerated expression functions should be integrated into a single, stimulus-responsive gene delivery system. We have developed a technology (‘drug or gene delivery system responding to cellular signals’, or D-RECS) that utilizes the abnormal activation of protein kinases as the trigger to release genes from cationic polymeric carriers. Protein kinases have essential roles in communicating signals from the cell surface to the nucleus in cells and thus initiate cellular responses such as cell death, proliferation and
migration. It has been frequently reported that abnormal activation of certain protein kinases is closely related to cancer, inflammation and other diseases. We aimed to exploit such abnormal activation of protein kinases to both ‘signpost’ disease cells and activate transgene expression. For this purpose, we required a polymer grafted with cationic substrate peptides which were phosphorylated by the activated target protein kinase. This cationic polymer should then form a complex with plasmid DNA (pDNA) through electrostatic interactions and efficiently suppress expression of the gene (Fig. 1A). Upon phosphorylation of the substrate peptides by disease-activated protein kinases in cells, pDNA is released from the polymer due to the disintegration of the electrostatic complex and gene expression is activated. In recent studies, we demonstrated transgene regulation in response to protein kinase Cα (PKCα) [18–20], c-AMP dependent protein kinase (PKA) [21] and IκB kinase β (IKKβ) [22] activities. Here, we chose the Rho associated coiled-coil kinase (ROCK) as the target kinase for this system. ROCK is a serine/threonine kinase and plays an important role in various cellular processes such as smooth muscle contraction, actin cytoskeleton organization, cell adhesion and motility, proliferation and cytokinesis [23–25]. These functions are involved in the pathogenesis of cardiovascular diseases [26], Alzheimer's disease [27] and cancer metastasis [28]. Especially, cardiovascular diseases are important diseases but there are little systems to delivery drug or genes to those diseases selectively. Therefore, ROCK is a key target for the therapy of these conditions.
2. Materials and methods 2.1. Synthesis of the substrate peptide In this study, a peptide RAKYKTLRQIR-NH2 was used as ROCK sensitive substrate and RAKYKAKRQIR-NH2 was used as a control peptide. The peptide substrate was originally developed after screening of peptide library (to be submitting). These peptides with a methacryloyl group at the amino-terminus were synthesized according to an Fmoc synthetic strategy. Peptides were cleaved from the resin by treatment with 94% trifluoroacetic acid (TFA) containing 2.5% triisopropylsilane, 2.5% water and 1.0% 1,2-ethanedithiol and purified with reverse-phase high performance liquid chromatography. Mass spectra were obtained by matrix-assisted laser desorption ionization time-of-
41
flight mass spectrometry (MALDI-TOF-MS) to verify the sequences of the synthesized peptides.
2.2. Phosphorylation assay For the substrate peptides, phosphorylation by ROCK (Carna Bioscience Incorporation, Kobe, Japan) was evaluated by means of MALDI-TOF-MS. Phosphorylation reactions were carried out in a total volume of 20 μl containing 20 mM Tris–HCl buffer (pH 7.4), 1 mM MgCl2, 100 μM ATP, 30 μM substrate peptide and 1 μg/ml ROCK2. The assay mixture was incubated at 37 °C for 1 h. The reaction was quenched by addition of 4 μl of 0.1% TFA containing α-cyano-4-hydroxycinnamic acid matrix in 50% water/acetonitrile to 1 μl of the reaction mixture. An aliquot of the mixture (2 μl) was applied to the sample plate, dried to induce crystallization and analyzed by MALDI-TOF-MS.
2.3. Synthesis of the ROCK-responsive polymer The polymer was synthesized in a manner similar to that described previously [18–20] (Fig. 1B). Briefly, a methacryloyl peptide and acrylamide were dissolved in degassed water and allowed to stand at room temperature for 90 min after the addition of ammonium persulfate (APS) and N, N, N′, N′,-tetramethylethylenediamine (TEMED) as the redox initiator couple. The product was purified by dialysis against water for 3 days. The content of the peptides was calculated by elemental analysis and the molecular weight of the polymer was estimated by gel-permeation chromatography using a polyethyleneglycol standard.
2.4. Preparation of pDNA A pDNA containing a firefly luciferase cDNA driven by a CMV promoter was prepared following our previous report [18–20] and was used in all experiments without a cell-free experiment. A pDNA containing a firefly luciferase cDNA driven by a T7 promoter was used in the cell-free expression experiment.
2.5. Dynamic light scattering (DLS) measurement Complexes of the polymer and pDNA at various amine/phosphate (N/P) ratios were prepared by mixing at room temperature for 15 min. The final concentration of pDNA was adjusted to 2.5 μg/ml using 10 mM HEPES buffer (pH 7.4) or PBS (pH 7.4). The diameter and zeta (ζ)-potential of the complexes were determined using a Zetasizer (Malvern Instruments, Worcestershire, UK) with the He/Ne laser at a detection angle of 173° at 25 °C. For the evaluation of complex disintegration during phosphorylation, we prepared the complex by the same method and the final concentration of pDNA was adjusted to 2.5 μg/ml in buffer containing 10 mM HEPES (pH 7.4), 1 mM MgCl2, 200 μM ATP and 2 μg/ml activated or inactivated ROCK2. Inactivation of ROCK2 was performed by heating at 97 °C for 3 min. Dynamic light scattering intensity was then monitored throughout the ROCK reaction.
2.6. Gel electrophoresis retardation assay
Fig. 1. (A) ROCK-responsive gene regulation system. Co-polymers grafted to cationic substrate peptides form complexes with plasmid DNA through electrostatic interactions and suppress gene transcription. Phosphorylation of the grafted substrate peptides in the polymer by activated ROCK in disease cells causes disintegration of the complex and allowing activation of the transgene. (B) Schematic of the chemical reaction whereby substrate peptide is grafted to the polymer.
pDNA (0.3 μg) and various concentrations of polymers were incubated at room temperature for 15 min and then, incubated at 37 °C for 1 h with 20% fetal bovine serum (FBS). One hundred equivalent poly anion (poly (sodium 4-styrenesulfonate)) to pDNA was added to the solutions to exclude pDNA from polymers and the samples were analyzed by electrophoresis using 1% agarose gel at 100 V for 25 min.
GENE DELIVERY
A. Tsuchiya et al. / Journal of Controlled Release 155 (2011) 40–46
GENE DELIVERY
42
A. Tsuchiya et al. / Journal of Controlled Release 155 (2011) 40–46
2.7. Ethidium bromide (EtBr) exclusion assay
2.12. Peptide phosphorylation cellular reporter assay
Complexes between the polymer and pDNA were prepared at various N/P ratios in sterilized water in the presence of EtBr. Solutions were incubated at room temperature for 15 min. The final concentrations of pDNA and EtBr were adjusted to 5 μg/ml and 12.5 μg/ml, respectively, with 10 mM HEPES buffer (pH 7.4). Fluorescence measurements of sample solutions were carried out at 25 °C using the ARVO multilevel counter (Wallac Incorporation, Turku, Finland). Excitation and emission wavelengths were 530 nm and 590 nm, respectively. The relative fluorescence intensity (RFI) was determined using the following equation:
To stimulate ROCK activation, cells were stimulated with 20 μM L-αlysophosphatidic acid (LPA) for 6 h, then scraped and lysed in 100 μl of lysis buffer (20 mM Tris–HCl, pH 7.4, 0.05% Triton-X 100, 2 mM EDTA). After centrifuging the sample at 15,000 g at 4 °C for 10 min, a 10 μl aliquot of the supernatant was used to measure chemiluminescence in a MiniLumat LP9506 (EG & G Berthold, Wildbad, Germany) directly after adding 40 μl of luciferase assay solution. The results are presented as relative luminescence units (RLU)/mg protein.
RFIð%Þ = ðFobs −Fe Þ = ðF0 −Fe Þ × 100
Cells were prepared for transfection as above. Four hours after transfection, the medium was replaced with DMEM and cells were incubated for a further 18 h. The cultured cells were then scraped and lysed in 200 μl of lysis buffer (20 mM Tris–HCl, pH 7.4, 0.05% Triton-X 100, 2 mM EDTA). After centrifuging the sample at 15,000 g at 4 °C for 10 min, pDNA was extracted from cell lysates using DNeasy Blood & Tissue Kits (Qiagen GmbH, Hilden, Germany). Quantitative analysis of pDNA was performed by real time PCR in a protocol similar to our previous report [29]. Sequences of PCR primers were as follows: forward primer, 5′-gcgccttatccggtaactatc-3′; reverse primer, 3′-acttcaccaccggattgatgc-5′. CYBR Green I purchased from Takara (Tokyo, Japan) was used as a fluorescent probe. PCR conditions were: 95 °C for 1 min, followed by 40 cycles at 94 °C for 15 s, 60 °C for 15 s and 72 °C for 30 s.
where Fobs, Fe and F0 are the fluorescence intensities of the polymer– pDNA complexes at each N/P ratio; EtBr without pDNA; and EtBr/pDNA without the polymer, respectively. 2.8. Cell-free gene expression assay Luciferase expression by a cell-free system was carried out using the T7 Quick Coupled Transcription/Translation System (Promega, Madison, WI, USA) as described [18]. Copolymer/pDNA complexes at N/P ratios of 1.0, 2.0 and 4.0 were prepared by mixing both compounds for 15 min. Final volumes were adjusted 10 μl by 10 mM HEPES buffer (pH 7.4). Detection of luciferase expression was according to manufacturer's protocols. Naked-pDNA (0.3 μg) was the controls. To monitor chemiluminescence, 20 μl of reaction solutions was added to the luciferase assay solution (Promega) and luminescence intensity measured by the ARVO multilabeled counter. 2.9. Monitoring complex phosphorylation using a coupled enzyme assay Phosphorylation on the pendant peptide in the polymer–pDNA complex was monitored using a coupled enzyme assay [18]. The complex or the peptide itself was diluted to 100 μl (final concentration of the peptide unit was 30 μM) in 10 mM HEPES (pH 7.4) containing 1 mM phosphoenolpyruvate, 10 U/μl lactate dehydrogenase (LDH), 4 U/μl pyruvate kinase, 200 μM ATP, 1 mM MgCl2, 300 μM nicotinamide adenine dinucleotide (NADH) and 2.0 μg/ml ROCK2. The reaction was carried out at 25 °C. The consumption of NADH was detected by monitoring absorbance at 340 nm with a UV/Vis spectrometer (UV2550; Shimadzu, Tokyo, Japan) equipped with an SPR-8 temperature controller (Shimadzu).
2.13. Real time (RT)-PCR
3. Results and discussion 3.1. Synthesis of the peptide-grafted polymer With five cationic amino acids making it highly suitable for D-RECS, we used the peptide RAKYKTLRQIR-NH2 as the substrate for ROCK. Evaluation using MALDI-TOF-MS confirmed this peptide as an effective ROCK substrate (Fig. 2). The polymer-grafted substrate peptides were then synthesized by radical polymerization using methacryloyl-peptide monomer and acrylamide (Fig. 1B). The molecular weight of the
2.10. Cell culture The NIH3T3 cell line was purchased from Health Science Research Resources Bank (Tokyo, Japan). Cells were cultured in Dulbecco's modified Eagle's medium (DMEM) (Wako, Osaka, Japan) containing 10% FBS, 100 U/ml penicillin, 100 μg/ml streptomycin and 0.25 μg/ml amphotericin-B (all from Invitrogen, New York, USA). Cells were incubated in a humidified atmosphere of 5% CO2 and 95% air at 37 °C. 2.11. Transfection of complexes into cells NIH 3T3 cells were cultured in on 24-well culture plates (20,000 cells/well) at 37 °C for 18 h in 0.5 ml DMEM containing 10% FBS. The medium was changed to serum-free DMEM followed by a further 4 h incubated at 37 °C. The medium was then replaced with 500 μl of Opti-MEM (Invitrogen) containing the complexes (prepared as for DLS assay, above) at N/P ratios of 1.0, 2.0 or 4.0 (pDNA remained constant at 2.0 μg/ml) and cells were incubated at 37 °C for 4 h. The medium was then changed to serum-free DMEM followed by a further 14 h incubation at 37 °C before assaying.
Fig. 2. MALDI-TOF-MS spectra of the substrate peptide before (A) and after (B) phosphorylation by ROCK.
43
resultant polymer was ~130 kDa, with 2.9% methacrylated peptide incorporation. 3.2. Characteristics of the polymer–pDNA complex The diameter and ζ-potential of the complexes are important factors in efficient gene delivery. The suitable diameter of the complex to transfer pDNA into cells via endocytosis is around 100 nm. After mixing the polymer and pDNA for 15 min, diameter and ζ-potential of the complex was measured at N/P ratios of 1.0, 2.0 and 4.0 (Table 1) using DLS. The particle size of this complex decreased as N/P ratios increased from 1.0 to 2.0, becoming constant at around 110 nm in 10 mM HEPES buffer (pH 7.4). This value is smaller than that for our previously reported D-RECS complexes (150–200 nm) [18,20,21]. Particle size at an N/P ratio of 1.0 (229 ± 85.4 nm) was also much smaller than that in ordinary D-RECS complexes, which tend to aggregate into very large (μm range) complexes with pDNA at an N/P ratio of 1.0. These results indicate that the polymer formed tighter complexes with pDNA than do ordinary D-RECS polymers. The diameters of these complexes were within a suitable range to transfer into cells via endocytosis. We then investigated the condensation of pDNA with the cationic polymer using the EtBr exclusion assay. Intercalation of EtBr into the DNA strand increases its fluorescence intensity drastically. Conversely, complexation of the DNA during formation of electrostatic complexes with polycations releases the intercalated EtBr, decreasing its fluorescence intensity. Exploiting this phenomenon, the condensation of DNA has been evaluated in cationic polymer–DNA complexes [30]. EtBr fluorescence intensity in both HEPES buffer (pH 7.4) and PBS (pH 7.4) was measured for each of the complexes formed at different N/P ratios in this study (Table 1). The polymer decreased the EtBr fluorescence intensity by 70% at N/P ratios above 2.0. Furthermore, the values both in HEPES buffer and in PBS were very similar at N/P ratios of above 1.0. This is notable because high salt concentrations such as in PBS or saline tend to cause disintegration of ordinary D-RECS polymer–pDNA complexes. Next, using a cell-free expression system, we evaluated the suppression of gene expression resulting from complexation of the pDNA with the polymer at various N/P ratios. For safe gene therapy, it is critical to minimize gene expression in non-diseased cells to avoid any undesirable side effects. At an N/P ratio of 1.0, the gene expression was suppressed by 40% compared with uncomplexed pDNA (Fig. 3B). However, at N/P ratios of 2.0 and 4.0, gene expression was suppressed by N95% relative to uncomplexed pDNA. Taken together with the results in Table 1 indicating that complex size decreases as N/P ratio increases, this suggests that smaller, tighter complexes produce more effective suppression of gene expression.
Fig. 3. Characterization of the polymer–pDNA complexes. (A) Condensation of pDNA by forming the complex with polymer in the EtBr exclusion assay. Assays were performed in either HEPES buffer or PBS. Data represent RFI of EtBr–DNA interaction at increasing N/P ratios. Decreasing RFI corresponds to elevated DNA condensation as EtBr is excluded from the complex. Data are mean ± SEM for three experiments (B) Luciferase expression from the polymer–pDNA complex at each N/P ratio in a cell-free expression system. Complexes were prepared by mixing pDNA with the polymers. RLUs were calculated by normalizing the luminescence for each complex to that of pDNA alone. Data are mean ± SEM for three experiments.
This decrease in A340 illustrates that grafting the peptide into the polymer chain did not affect its status as a ROCK phosphorylation target. Using DLS, we then evaluated whether the complex disintegrated following phosphorylation by activated ROCK (Fig. 5). At an N/P ratio of 2.0, introducing activated ROCK into the complex solution led to an increase in the particle diameter (from 200 nm to 2000 nm) and a decrease in the count rate (from 6300 kcps to 4100 kcps) within 60 min in phosphorylation reaction solution. In contrast, inactivated ROCK did not alter either the hydrodynamic diameter or the count rate of the complex over the same period. Furthermore, the polydispersity index (PDI) was increased by the addition of activated ROCK (data not shown).
3.3. Phosphorylation and disintegration of the complex In D-RECS, kinase phosphorylation of substrate peptides grafted into the polymer causes complex disintegration, which activates gene expression. To verify that the substrate peptide used here remained a phosphorylation target for ROCK when grafted into the polymer, we used a coupled enzyme assay to assess peptide phosphorylation by ROCK in the polymer–pDNA complex (Fig. 4). Phosphorylation of uncomplexed peptide and polymer–pDNA complex (at N/P ratios of 2.0 and 4.0) was measured, with peptide content in each condition adjusted to 30 μM. Uncomplexed and complexed peptide were phosphorylated comparably, as shown by a similar time course in the decrease in A340. Table 1 Diameter and ζ-potential of polymer/pDNA complexes. N/P ratio
1.0
2.0
4.0
Diameter (nm) ζ-potential (mV)
229.4 ± 85.4 5.9 ± 2.4
110.8 ± 0.6 14.1 ± 2.3
109.4 ± 2.2 14.3 ± 1.0
Fig. 4. Phosphorylation time course of the peptide (black) and the polymer–pDNA complexes at N/P ratios of 2.0 (dark gray) and 4.0 (light gray) monitored by coupled enzyme assay. Consumption of NADH, measured by changes in the absorbance at 340 nm, (A340) was used as a reporter for the phosphorylation status of the polymer.
GENE DELIVERY
A. Tsuchiya et al. / Journal of Controlled Release 155 (2011) 40–46
GENE DELIVERY
44
A. Tsuchiya et al. / Journal of Controlled Release 155 (2011) 40–46
Fig. 5. DLS measurements of the polymer–pDNA complexes in a solution containing either activated or inactivated ROCK. Diameters (A) and count rates (B) of the complex during the phosphorylation by ROCK at an N/P ratio of 2.0 (circles) and 4.0 (squares). Open and closed symbols represent the complex in the presence of activated ROCK and inactivated ROCK, respectively.
Fig. 6. DLS measurements of the polymer–pDNA complexes in PBS. Shown are time courses showing the change in diameter (A) and count rate (B) of the complex at N/P ratios of 1.0 (circles), 2.0 (squares) and 4.0(triangles).
serum. This result suggested that pDNA in serum was little degraded but kept its formation. These results suggest that the complex at an N/P ratio of 2.0 was disintegrated in response to activated ROCK. We showed that complexes formed at an N/P ratio of 4.0 were viable phosphorylation substrates for activated ROCK (Fig. 4). However, at an N/P ratio of 4.0, the diameter and the count rate of the complex stayed constant over 60 min, even in the presence of activated ROCK (Fig. 5). This suggests that at high N/P ratios the complex does not disintegrate following phosphorylation of the substrate peptide. 3.4. Stability of the polymer–pDNA complex To use a polymer–DNA polyplex for gene delivery in vivo, it must remain stable until it reaches its target. If the complex is broken or aggregated in the blood stream or in normal cells, efficient gene delivery cannot be achieved. In this context, the stability of the complex was evaluated using DLS to measure diameter and count rate of complexes in 150 mM saline (Fig. 6). At an N/P ratio of 1.0, the diameter of the complex increased from 250 nm to 700 nm and the count rate decreased from 2000 kcps to 800 kcps. At an N/P ratio of 2.0, the diameter change was negligible in the presence of saline although the count rate increased from 2800 kcps to 4000 kcps. At an N/P ratio of 4.0, both diameter and count rate were unchanged. We conclude that at low N/P ratio the polymer forms an ‘open’ complex with pDNA that can easily disintegrate and aggregate in the presence of saline. However, at N/P ratios above 2.0, the complex was increasingly stable. Next, we evaluated the protection of pDNA from nuclease in serum. Polymer–pDNA complexes were prepared at N/P ratios of 1.0, 2.0 and 4.0 and mixed with 20% serum in 10 mM HEPES buffer. Degradation of pDNA was evaluated by gel electrophoresis assay. In the case of only pDNA, as a control, pDNA was degraded perfectly (Fig. 7). In 20% serum, forming complex with polymers, pDNA was obtained which formation were different from that in the absence of
3.5. In vitro transgene regulation It is known that ROCK is activated in NIH3T3 cells treated with LPA [31] and we also found ROCK-medicated actin stress fiber formation (Fig. S1). We used this system as a model system of ROCK activating cells to investigate whether transgene expression in the polymer–pDNA complex was regulated in response to ROCK activation. A basal level of transgene expression was seen following addition of the complex at all N/P ratios, but decreased as N/P ratio increased (Fig. 8A), reflecting the decreasing propensity of the complex to disintegrate spontaneously at higher N/P ratios. In fact, the highest gene expression was obtained at an N/P ratio of 1.0 and the lowest was obtained at an N/P ratio of 4.0. This propensity was conformed to the result of cell-free expression system. Importantly, at an N/P ratio of 2.0, elevated transgene expression was observed in response to LPA, whereas in the presence of 20 μM ROCK inhibitor, Y-27632, or using control polymers that were exchanged Thr residue to Ala residue, there were no such increase in gene expression
Fig. 7. Protection of pDNA degradation from nuclease in FBS. Complexes were incubated with 20% FBS and then, degradation of pDNA was evaluated by gel electrophoresis shift assay.
45
Furthermore, we investigated whether the absence of any transgene expression in cells exposed to complexes with an N/P ratio of 4.0 was due to impaired uptake of these complexes. Using real-time PCR, we quantified the pDNA transferred into cells at each N/P ratio (1.0, 2.0 and 4.0). Cellular uptake of the complex was highest at an N/P ratio of 1.0 but, although uptake at an N/P of 4.0 was lower, it was comparable to that for N/P ratio 2.0 (Fig. 8B). This suggests that the low level of gene expression in the complex at an N/P ratio of 4.0 is not caused by poor uptake. Taken together, our data support a conclusion that this effect is most likely due to these N/P ratio 4.0 complexes not disintegrating properly. In this scenario, despite being properly endocytosed (Fig. 8B) and phosphorylated (Fig. 4), these complexes are simply too stable to release the transgene from the complex, as suggested by our stability and disintegration data (Figs. 6 and 5, respectively). This hyperstable complex thus cannot express the luciferase gene under either basal or stimulated conditions. On the other hand, complexes at an N/P ratio of 1.0 were not suppressed the gene expression without LPA treating because of that low stability. To get responsibility of the activation of ROCK using ROCK responsive polymer, it is important that we select the optimal N/P ratios. 4. Conclusion
Fig. 8. Transfection of polymer–pDNA complexes into NIH3T3 cells. Complexes were prepared by mixing pDNA with the polymer at N/P ratios of 1.0, 2.0 and 4.0. (A) Luciferase expression of each complex in the absence (black bar) or presence (white bar) of LPA stimulation. As a control, the polymer that Thr residues exchanged to Ala residues was used. Data are presented as the RLU/mg protein, mean ± SEM for six experiments. *P b 0.05 (calculated using t-test). (B) The amounts of pDNA (μg/mg protein) transfected into NIH3T3 cells at each N/P ratio of complex. pDNA was purified using DNeasy blood & tissue kits and quantitative analysis of pDNA was performed by real time PCR. Data represent mean expression level ± SEM for three experiments.
(Fig. 8A). At an N/P ratio of 2.0, when substrate peptides were phosphorylated by ROCK, the complex was disintegrated (Fig. 5). The same phenomenon might be observed in vitro. The complex was phosphorylated in cells which treated with LPA and ROCK was activated, and then the gene expression was activated. Because the gene expression was not activated in the case of treatment with ROCKspecific inhibitor and using control polymers, this activation of the gene expression was depended to ROCK activity and caused by phosphorylation of the complex. The consensus sequence of the ROCK phosphorylation site is considered to be R/KXXS/T or R/KXS/T (R, arginine; K, lysine; X, any amino acid; S, serine; and T, threonine) [25]. These sequences are similar to consensus phosphorylation site motifs for PKC, which were identified to be R/KXXS/T, R/KXS/T, R/KXXS/TXR/K, or R/KXS/TXR/K [32,33]. Thus, we examined whether PKCα-responsive polymer can increase gene expression in the LPA-treated NIH3T3 cells. PKCαresponsive polymer (PPC (S)) and its control polymer (PPC (A)) [18–20] showed no gene expression in the LPA-treated NIH3T3 cells (Fig. S2). Previous studies suggested that PKCα is not involved in LPA-induced actin stress fiber formation [34] and high level of LPA inhibits PKCα activity [35]. These results mean that the gene expression in ROCK substrate-grafted polymer system is solely due to ROCK activity, but not PKCα activity.
We report here a novel gene delivery regulation system that can activate transgene expression in response to ROCK signals. The polymer grafted with ROCK substrate peptides formed a more stable complex with pDNA than ordinary D-RECS systems. This should confer a strong resistance to complex disintegration in serum, blood, or non-target cells, although the responsiveness to ROCK is also lost if the N/P ratio is too high. We have successfully optimized ROCK-responsive transgene activation at an N/P ratio of 2.0. This system may have key applications for therapy and diagnostics in diseases where ROCK activation is observed. Acknowledgments This work was financially supported by a grant-in-aid for Scientific Research from the Ministry of Education, Science, Sports, and Culture in Japan and also supported by A3 Foresight Program in Japan Society for the Promotion of Science. A. T. is grateful to Japan Society for the Promotion of Science (JSPS) for the DC scholarship. Appendix A. Supplementary data Supplementary data to this article can be found online at doi:10.1016/j.jconrel.2011.05.002. References [1] J. Luten, C.F. Nostrum, S.C.D. Smedt, W.E. Hennink, Biodegradable polymers as non-viral carriers for plasmid DNA delivery, J. Control. Release 126 (2008) 97–110. [2] M. Ruponen, S. Arkko, A. Urtti, M. Reinisalo, V.P. Ranta, Intracellular DNA release and elimination correlated poorly with transgene expression after non-viral transfection, J. Control. Release 146 (2009) 226–231. [3] O. Jeon, H.S. Yang, T.J. Lee, B.S. Kim, Heparin-conjugated polyethyleneimine for gene delivery, J. Control. Release 132 (2008) 236–242. [4] W.H. Kong, D.K. Sung, Y.H. Shim, K.H. Bae, P. Dubois, T.G. Park, J.H. Kim, S.W. Seo, Efficient intracellular siRNA delivery strategy through rapid and simple two steps mixing involving noncovalent post-PEGylation, J. Control. Release 138 (2009) 141–147. [5] P.R. Dash, V. Toncheva, E. Schacht, L.W. Seymour, Synthetic polymers for vectorial delivery of DNA: characterization of polymer–DNA complexes by photon correlation spectroscopy and stability to nuclease degradation and disruption by polyanions in vitro, J. Control. Release 48 (1997) 269–276. [6] A.E. Aneed, An overview of current delivery systems in cancer gene therapy, J. Control. Release 94 (2004) 1–14. [7] V. Sokolova, M. Epple, Inorganic nanoparticles as carriers of nucleic acids into cells, Angew. Chem. Int. Ed. 46 (2007) 2–16.
GENE DELIVERY
A. Tsuchiya et al. / Journal of Controlled Release 155 (2011) 40–46
GENE DELIVERY
46
A. Tsuchiya et al. / Journal of Controlled Release 155 (2011) 40–46
[8] I.A. Khalil, K. Kogure, H. Akita, H. Harashima, Uptake pathways and subsequent intracellular trafficking in nonviral gene delivery, Pharmacol. Rev. 58 (2006) 32–45. [9] S. Ganta, H. Devalapally, A. Shahiwala, M. Amiji, A review of stimuli-responsive nanocarriers for drug and gene delivery, J. Control. Release 126 (2008) 187–204. [10] N. Rapoport, Physical stimuli-responsive polymeric micelles for anti-cancer drug delivery, Prog. Polym. Sci. 32 (2007) 962–990. [11] M. Meyer, A. Philipp, R. Oskuee, C. Schmidt, E. Wagner, Breathing life into polycations, functionalization with pH-responsive endosomolytic peptides and polyethylene glycol enables siRNA delivery, J. Am. Chem. Soc. 130 (2008) 3272–3273. [12] Y. Lee, T. Ishii, H. Cabral, H.J. Kim, J.H. Seo, N. Nishiyama, H. Oshima, K. Osada, K. Kataoka, Charge-conversional polyionic complex micelles — efficient nanocarriers for protein delivery into cytoplasm, Angew. Chem. Int. Ed. 48 (2009) 1–5. [13] R. Namgung, S. Nam, S.K. Kim, S. Son, K. Singha, J.S. Kwon, Y. Ahn, M.H. Jeong, I.K. Park, V.K. Garripelli, S. Jo, An acid-labile temperature-responsive sol–gel reversible polymer for enhanced gene delivery to the myocardium and skeletal muscle cells, Biomaterials 30 (2009) 5225–5233. [14] S.H. Lee, S.H. Choi, S.H. Kim, T.G. Park, Thermally sensitive cationic polymer nanocapsules for specific cytosolic delivery and efficient gene silencing of siRNA: swelling induced physical disruption of endosome by cold shock, J. Control. Release 125 (2008) 25–32. [15] J.E. Chung, M. Yokoyama, T. Okano, Inner core segment design for drug delivery control of thermo-responsive polymeric micelles, J. Control. Release 65 (2000) 93–103. [16] M. Breunig, C. Hozsa, U. Lungwitz, K. Watanabe, I. Umeda, H. Kato, A. Goepferich, Mechanistic investigation of poly(ethylene imine)-based siRNA delivery: disulfide bonds boost intracellular release of the cargo, J. Control. Release 130 (2008) 57–63. [17] O. Taratula, O.B. Garbuzenko, P. Kirkpatrick, I. Pandya, R. Savla, V.P. Pozharov, H. He, T. Minko, Surface-engineered targeted PPI dendrimer for efficient intracellular and intratumoral siRNA delivery, J. Control. Release 184 (2009) 284–293. [18] R. Toita, J.H. Kang, J.H. Kim, T. Tomiyama, T. Mori, T. Niidome, B. Jun, Y. Katayama, Protein kinase Cα-specific peptide substrate graft-type copolymer for cancer cellspecific gene regulation systems, J. Control. Release 139 (2009) 133–139. [19] T. Tomiyama, J.H. Kang, R. Toita, T. Niidome, Y. Katayama, Protein kinase Cαresponsive polymeric carrier: its application for gene delivery into human cancers, Cancer Sci. 100 (2009) 1532–1536. [20] J.H. Kang, D. Asai, J.H. Kim, T. Mori, R. Toita, T. Tomiyama, Y. Asami, J. Oishi, Y.T. Sato, T. Niidome, B. Jun, H. Nakashima, Y. Katayama, Design of polymeric carriers for cancer-specific gene targeting: utilization of abnormal protein kinase Cα activation in cancer cells, J. Am. Chem. Soc. 130 (2008) 14906–14907.
[21] J. Oishi, K. Kawamura, J.H. Kang, K. Kodama, T. Sonoda, M. Murata, T. Niidome, Y. Katayama, An intracellular kinase signal-responsive gene carrier for disordered cell-specific gene therapy, J. Control. Release 110 (2006) 431–436. [22] D. Asai, A. Tsuchiya, J.H. Kang, K. Kawamura, J. Oishi, T. Mori, T. Niidome, Y. Shoji, H. Nakashima, Y. Katayama, Inflammatory cell-specific transgene expression system responding to Iκ-B kinase beta activation, J. Gene Med. 11 (2009) 624–632. [23] K. Riento, A.J. Ridley, ROCKS: multifunctional kinases in cell behavior, Mol. Cell. Biol. 4 (2003) 446–456. [24] N.P. Tsai, L.N. Wei, RhoA/ROCK1 signaling regulates stress granule formation and apoptosis, Cell. Signal. 22 (2010) 668–675. [25] J.H. Kang, Y. Jiang, R. Toita, J. Oishi, K. Kawamura, A. Han, T. Mori, T. Niidome, M. Ishida, K. Tatematsu, K. Tanizawa, Y. Katayama, Phosphorylation of Rhoassociated kinase (Rho-kinase/ROCK/ROK) substrates by protein kinases A and C, Biochimie 89 (2007) 39–47. [26] H. Shimokawa, M. Rashid, Development of Rho-kinase inhibitors for cardiovascular medicine, Trends Pharmacol. Sci. 28 (2007) 296–302. [27] A. Salminen, T. Suuronen, K. Kaarniranta, ROCK, PAK, and Toll of synapses in Alzheimer's disease, Biochem. Biophys. Res. Commun. 371 (2008) 587–590. [28] K. Itoh, K. Yoshioka, H. Akedo, M. Uehata, T. Ishizaki, S. Narumiya, An essential part for Rho-associated kinase in the transcellular invasion of tumor cells, Nat. Med. 5 (1999) 221–225. [29] T. Kawano, M. Yamagata, H. Takahashi, Y. Niidome, S. Yamada, Y. Katayama, T. Niidome, Stabilizing of plasmid DNA in vivo by PEG-modified cationic gold nanoparticles and the gene expression assisted with electrical pulses, J. Control. Release 111 (2006) 382–389. [30] K. Miyata, S. Fukushima, N. Nishiyama, Y. Yamasaki, K. Kataoka, PEG-based block catiomers possessing DNA anchoring and endosomal escaping functions to form polyplex micelles with improved stability and high transfection efficacy, J. Control. Release 122 (2007) 252–260. [31] M.P. Iwanicki, T. Vomastek, R.W. Tilghman, K.H. Martin, J. Banerjee, P.B. Wedegaertner, J.T. Parsons, FAK, PDZ-RhoGEF and ROCKII cooperate to regulate adhesion movement and trailing-edge retraction in fibroblasts, J. Cell Sci. 121 (2008) 895–905. [32] R.B. Pearson, B.E. Kemp, Protein kinase phosphorylation site sequences specificity motifs: tabulations, Method Emzymol. 200 (1991) 62–81. [33] H. Hug, T.F. Sarre, Protein kinase C isoenzymes: divergence in signal transduction? Biochem. J. 291 (1993) 329–343. [34] K. Riento, N. Totty, P. Villalonga, R. Garg, R. Guasch, A.J. Ridley, RhoE function is regulated by ROCK I-mediated phosphorylation, EMBO J. 24 (2005) 1170–1180. [35] J.J. Sando, O.I. Chertihin, Activation of protein kinase C by lysophosphatidic acid: dependence on composition of phospholipid vesicles, Biochem. J. 317 (1996) 583–588.