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Angew Chem Int Ed Engl. Author manuscript; available in PMC 2016 June 01. Published in final edited form as: Angew Chem Int Ed Engl. 2016 June 1; 55(23): 6657–6661. doi:10.1002/anie.201601008.
Molecularly Regulated Reversible DNA Polymerization Dr. Niancao Chenb, Mr. Xuechen Shib, and Yong Wanga [Associate Professor] Yong Wang: [email protected] aDepartment
of Biomedical Engineering, Pennsylvania State University, 232 Hallowell Building, University Park, PA 16802
bDepartment
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of Biomedical Engineering, Pennsylvania State University, 202 Hallowell Building, University Park, PA 16802
Abstract Natural polymers are synthesized and decomposed under physiological conditions. However, it is challenging to develop synthetic polymers whose formation and reversibility can be both controlled under physiological conditions. Here we show that both linear and branched DNA polymers can be synthesized via molecular hybridization in aqueous solutions, on the particle surface, and in the extracellular matrix (ECM) without the involvement of any harsh conditions. More importantly, these polymers can be effectively reversed to dissociate under the control of molecular triggers. Since nucleic acids can be conjugated with various molecules or materials, we anticipate that molecularly regulated reversible DNA polymerization holds potential for broad biological and biomedical applications.
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TOC image Regulated reversible DNA polymerization: It is challenging to emulate nature in developing synthetic polymers with dynamic reversibility under physiological conditions. This study demonstrates that linear and branched DNA polymers can be synthesized and reversed under the control of molecular triggers without the involvement of any harsh conditions.
Author Manuscript Correspondence to: Yong Wang, [email protected]. Supporting information for this article is given via a link at the end of the document.
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Keywords DNA; dynamic polymers; hybridization; reversible polymerization; self-assembly
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Synthetic polymers are broadly used in various applications such as medicine, cloth, devices, and tools. However, their synthesis and decomposition usually require harsh conditions (e.g., high temperature or organic solvent). In contrast, natural polymers (e.g., proteins) could be synthesized or decomposed with the aid of enzymes under mild physiological conditions.[1] Thus, great efforts have been made to synthesize dynamic polymers with reversible covalent bonds and non-covalent interactions via diverse mechanisms.[2] For instance, units of 2ureido-4-pyrimidone can form a self-complementary array of four hydrogen bonds for synthesis of unidirectional polymers with the reversibility of formation and decomposition.[3] These polymers not only provide a way of understanding the major processes in nature but also hold potential for broad applications.[4] However, synthetic monomers have been mostly studied in organic solvents and their polymers usually do not have biocompatibility;[5] moreover, these polymers do not exhibit regulatable reversibility under physiological conditions.[6]
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The purpose of this study was to demonstrate the ability to develop dynamic polymers whose synthesis and reversibility can be both controlled at the molecular level under physiological conditions (Figure 1), which has not been reported before. The polymers were synthesized using the principle of hybridization chain reaction;[7] the polymers were reversed using the principle of branch displacement.[8] The synthesis of the linear polymer (LP, Figure 1a, upper panel) involves three molecules including a DNA initiator (DI) and two DNA monomers (DMs). The DI is a linear structure with one functional domain as labeled with i (Figure 1b and Supporting Information, Table S1a). The DMs are hairpin structures. DM1s1 has three domains including i*, j and s1 and DM2 has two domains including i and j*. During the polymerization (Figure S1a), DI opens the hairpin structure of DM1s1 to form an i–i* double helix with j and s1 left as a linear segment. The linear j
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domain further reacts with the Notably, the domain s1 of DM1s1 does not participate in the linear polymerization and is a functional side group of LP. Thus, s1 can hybridize with a molecule carrying a complementary sequence domain s1* (Figures 1b and S1b). The molecular trigger T1 has two functional domains including s1* and j*. With the aid of s1–s1* and j–j* hybridization, T1 hybridizes with the DM1s1 unit of LP and displaces DM2. Resultantly, LP is reversed without the involvement of any non-physiological factors.
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Similar to DM1s1, DM2 can be designed with a functional side group (named as DM2k, Figure 1c and Table S1a) to bear a total of three domains including k, i and j*. Through the polymerization of DM1s1 and DM2k, LP acquires two functional side groups, s1 and k (Figure 1c). Since k can be designed as an initiator for DM3s2 and DM4, these two DMs react with the side group k of LP to form side chains along the backbone of LP (Figures 1c and S2). Resultantly, a branched polymer (BP) is synthesized with LP and two DMs (Figure 1a, lower panel). Because both DM1s1 and DM3s2 have the side groups, i.e., s1 and s2, respectively, the use of two corresponding molecular triggers T1 and T2 can induce the reversible polymerization of BP (Figures 1c and S2).
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The variation of DM structures would lead to the formation of different forms of LPs (Table S1b). Four representative LPs were synthesized (Figure S3). The results showed that DMs formed LPs in the presence of DI. Side groups did not apparently influence the LP formation. The apparent molecular weights of LPs mostly fell in the range between 500 to 3,000 bp. In contrast, DMs did not form LPs in the absence of DI (Figure 2a, lane 4). The result also showed that an inhibiting sequence could effectively terminate LP polymerization (Figure S4), suggesting that LPs were formed through sequential hybridization of DMs (Figure S1a). The increase of the reaction time led to more DM conversion (Figure S5). Specifically, ~60% of DMs were quickly converted into LPs within 0.5–1 h. The polymerization reached plateau at 8 h with a cumulative conversion efficiency of 86%. The successful synthesis of LPs was also confirmed using AFM (Figure 2b). The average contour length of LPs was ~122.8 nm (Figure S6a). The side group s1 had 10 nucleotides (nt). When it was incorporated into the 3′ end of DM1, the formed LP had periodic side functional groups (Figure S1a). T1 had 34 nt with 10 nt complementary to s1 and another 24 nt complementary to the j domain of DM1s1. When T1 and DM1s1 hybridize (Figure S7), the formed double-stranded helix has 34 base pairs (bp). In comparison, DM2 and DM1s1 have a helix of 24 bp. The former is thermodynamically more stable than the latter. Thus, T1 is more competitive than DM2 in hybridization with DM1s1.
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As shown in Figure 2c, the LPs bearing the s1 groups virtually disappeared after reacting with T1 (lane 6). The major reaction products (lane 6) were located in the same position as the hybridization complex of DM1s1, DM2, and T1 (lane 1). By comparison, the LP bands were not affected after mixing with the control triggering sequence T2 (lane 5). When LPs bearing no s1 groups were treated with T1, LPs barely changed (lanes 2 and 3). These results clearly demonstrate that T1 effectively triggered LP depolymerization in a sequence-specific and s1-dependent manner.
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The gel electrophoresis results were confirmed by AFM analysis. The morphologies of LPs with s1 side groups after polymerization and depolymerization are shown in Figure 2b&d and Figure S8a. Quantitatively, the average length of LPs decreased from 122.8 nm to 16.4 nm (Figure S6). The measured length of depolymerization products is consistent with the calculated length (Figure S8b). These data further confirm the success of T1-triggered LP depolymerization.
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We also studied the effects of three different hybridization parameters on depolymerization. The reaction time was varied from 10 min to 8 h (Figure S9). The result showed that 95% LPs disappeared within 10 min, suggesting that depolymerization was fast. The depolymerization efficiency increased with the increasing molar ratio of T1 to DM1s1, exhibiting nearly a linear relationship with the molar ratio (Figure S10). The length of T1 was also varied (Figure S11). Before the hybridization length was reduced to 25 bp, T1 could effectively trigger LP depolymerization. The efficiency gradually decreased with the hybridization length reduced from 25 to 16 bp. This trend corresponds to the ΔG change of the hybridization helixes (Figure S11d).
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BP was synthesized using filtered LP and the mixture of DM3 and DM4. The molar ratio of LP monomers to branch monomers was 1:5. LP had two side groups, s1 and k (Figure S12). The side group k had 29 nt. The 5 nt adjacent to the backbone of LP were designed as a spacer to mitigate potential steric hindrance. The other 24 nt were used as an initiator for reaction with DM3 and DM4. The result showed that k was able to induce polymerization of DM3 and DM4 (Figure S13a) similar to the DI-induced polymerization of DM1 and DM2. We further used k-bearing LP to react with DM3 (or DM3s2) and DM4 for BP synthesis. A clear band shift is shown in the gel images (lane 4 in Figure 3a and Figure S13b). The results showed that a significant amount of BP molecules had apparent molecular weights over 12,000 bp. In contrast, the reaction of LP without k and the two DMs did not lead to a band shift (lane 3 in Figure 3a and Figure S13b). This difference demonstrates that BP could be synthesized through the k-bearing LP and the two DMs. AFM imaging was performed to confirm the gel electrophoresis data (Figures 3b and S14a). Future work can be further performed to quantify the amount of monomer incorporation. DM1s1 and DM3s2 both had a side group (s1 or s2). They were designed to hybridize with their corresponding molecular triggers T1 and T2, respectively. Figure S15 showed that the addition of T1 into LP1 solution led to the disappearance of LP1 band (lane 6 in Figure S15a) and the addition of T2 into LP2 solution led to the disappearance of LP2 band (lane 6 in Figure S15b). These results demonstrate that molecular triggers could reverse LP successfully.
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Figure 3c shows that BP treated with different combinations of T1 and T2 exhibited a remarkably band shift (lanes 2–4) in comparison to the untreated BP (lane 1). For BP treated with only T1, the apparent molecular weights of reversed BP products were shifted into the range of 300 to 1,000 bp (lane 2, population i). It suggests that T1 depolymerized the backbone of BP while not affecting the side chains (Figure S16a). For BP treated with only T2, the apparent molecular weights of reversed BP fell into two ranges. One was from 500 to 3,000 bp (population ii); the other was at ~100 bp (population iii). It suggests that T2
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reversed the side chains of BP while not affecting the backbone (Figure S16b). For BP treated with the mixture of T1 and T2 (lane 4), two bright bands were observed. It suggests that both the backbone and side chains were depolymerized (Figure S2c). By contrast, BP without s1 and s2 did not show any change (lanes 5 and 6). These results were confirmed by AFM imaging (Figures 3d and S17). In addition to demonstrating polymerization and depolymerization of LPs and BPs in the aqueous solution, we further combined these polymeric systems with synthetic microparticle and antibody and validated their working efficiency to demonstrate the potential applications of reversible DNA polymerization.
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Polystyrene microparticles were functionalized with DI via streptavidin-biotin interactions and DM2 was labeled with fluorophore. The flow cytometry analysis showed that the average fluorescence intensity of the microparticles with LP was approximately one order of magnitude higher than that of control with one hybridized DM1–DM2 complex (Figure S18). It demonstrates that LP could be synthesized on the microparticle surface. Similarly, the results showed that BP could be synthesized on the microparticle surface (Figures 4a and S19). Experiments were further carried out to examine the reversibility. Microparticles bearing LP were treated with either T1 or T2 (Figure S20). T1 treatment induced a dramatic decrease of fluorescence intensity on microparticles whereas T2 treatment did not. These data demonstrate that LP was specifically reversed on the microparticle surface. Microparticles bearing BP were also treated with the triggering solution (Figures 4b and S21). Similar to LP, BP was reversed on the microparticle surface.
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We also used the ECM of living cells as a substrate to study LP and BP. An antibody-DI conjugate was prepared via the formation of a sandwich structure of biotinylated antibody, streptavidin and biotinylated DI. The antibody-DI conjugate induced the formation of LP in the ECM (Figure S22), and that LP further induced the formation of BP (Figures 4c and S23). When the ECM was treated by triggering solutions, the fluorescence intensities decreased in both cases of LP (Figure S24) and BP (Figures 4d–f and S25) to virtually the same level as the background. While the intensity of the blue background was slightly decreased during the second time imaging of the ECM after the triggering treatment due to photobleaching, Figure S25 demonstrates that triggering depolymerization rather than photobleaching was the dominant reason for the decrease of green fluorescence intensity. Thus, the data show that LP or BP can be polymerized and depolymerized in the ECM.
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Owing to high fidelity of hybridization and the water-soluble nature,[9] nucleic acids have been extensively explored in constructing complex structures[10] or nanodevices.[11] These systems have been further studied for numerous applications such as drug delivery,[12] molecular detection[13] and bioimaging.[14] However, previous work has not demonstrated the development of DNA polymers whose formation and reversibility can be both regulated under physiological conditions. This work has successfully demonstrated the ability to synthesize and reverse linear and branched DNA polymers in aqueous solutions, on the microparticle surface, and in the ECM, which further advances the development of nucleic acid-based material systems. We realize that increasing temperature[15] or using DNase[16] can lead to depolymerization of the DNA polymers; however, these methods are not
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physiologically biocompatible or do not provide selectivity of reversing a certain DNA polymer if multiple polymers exist. Similar to many other self-assembly systems,[3,17] it is also possible to design short self-complementary DNA oligonucleotides that can be spontaneously self-assembled and concomitantly disassembled. However, the spontaneous self-assembly and concomitant disassembly are not stable or molecularly regulatable.[18] In summary, this work has demonstrated that intermolecular hybrization is an effective mechanism for the synthesis of linear and branched DNA polymers with regulatable reversibility under strict physiological conditions. Since nucleic acids can be conjugated with various molecules or materials, molecularly regulated reversible polymerization and depolymerization are envisioned to hold potential for broad applications.
Methods Author Manuscript
Chemicals and reagents
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Phosphate buffered saline (PBS), Tris-EDTA and agarose were purchased from Fisher Scientific (Suwanee, GA). Oligonucleotides (Supporting Tab. 2) were purchased from Integrated DNA Technologies (Coralville, IA). The streptavidin-coated microparticles (average diameter: 5 μm) was obtained from Spherotech (Lake Forest, IL). The DNA marker, SYBR-Safe, CellTracker™ Blue CMAC dye, bovine serum albumin (BSA), human dermal fibroblasts, neonatal (HDFns) and cell culture reagents were purchased from Invitrogen (Carlsbad, CA). Ultracentrifugal filter (100K cut-off size) and biotinylated anticollagen IgG were purchased from Millipore (Billerica, MA). All other chemicals, such as (3-aminopropyl) triethoxysilane (APTES), were obtained from Sigma-Aldrich (Louis, MO). Mica was purchased from Asheville-Schoonmaker Mica Company (Newport News, VA). Atomic force microscope (AFM) cantilever (SCANASYST-FLUID+) was purchased from Bruker (Camarilla, CA). Reversible DNA polymerization in aqueous solution
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DNA sequences were designed using NUPACK server (www.nupack.org). Their secondary structures were also generated using NUPACK. Before hybridization reactions, DNA macromonomers were heated to 95°C and then cooled to room temperature in PBS for 1 h. During the procedure of LP synthesis, DI was mixed with the two DMs at room temperature. To synthesize BP, LP was filtered twice through an ultracentrifugal filter (100K cut-off size) at 14,000 × g to remove the unreacted DMs. The filtered LP products were incubated with the secondary set of DMs (2 μM each) at room temperature. The reaction time for each polymerization was 12 h unless specified elsewhere. To depolymerize the DNA polymers, molecular triggers were added into the solutions of LP or BP and reacted for predetermined periods of time at room temperature. The molar ratio of triggers to their corresponding monomers was all 2:1 unless specified elsewhere. To prepare hybridized DNA complexes, the mixture of molecular triggers and their corresponding monomers with a molar ratio of 1:1 was heated to 95 °C and then gradually cooled to room temperature. To examine the effect of the terminator on polymerization, DI (initially 0.3 μM), each DM (initially 3.0 μM) and terminator of different concentrations were prepared individually and then subjected to
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the annealing treatment before mixed together. The mixture was examined after 12 h incubation using agarose gel electrophoresis. Gel electrophoresis Agarose gel (1%) was used for gel electrophoresis in Tris-borate-EDTA (TBE) buffer (89 mM boric acid and 2 mM EDTA, pH 8.2) The agarose gel was pre-stained with SYBR-Safe (0.1 μL of stock solution per mL of agarose gel). The gel electrophoresis was run at 100 V for 60 min and the gels were imaged using a CRI Maestro EX System (Woburn, MA) with a blue exciting light (455 nm). The fluorescence of DNA in gels were recorded with a blue filter (Em>488 nm). ΔG calculation
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The ΔG values of hybridized DNA macromonomers were calculated using IDT OligoAnalyzer 3.1 (https://www.idtdna.com/calc/analyzer) under the condition of 0.25 μM of DNA and 140 mM Na+. Atomic force microscopy (AFM)
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Mica was modified to bear positive charges on its surface before sample loading and imaging. Briefly, mica with a dimension of ~1.5 cm in square was cut into an octagon shape and freshly cleaved with a thickness of 0.1 mm using a clean razor blade. The mica was attached with a metal clip on its edge to a glass rod in a desiccator, which was filled with argon gas. Two eppendorf caps were placed on the bottom of the desiccator to hold 90 μL of APTES and 30 μL of N,N-diisopropylethylamine (DIPEA). The mica in the desiccator was reacted with the evaporated chemicals for 2 h. The desiccator was purged with argon for 2 min and the mica was cured in the desiccator before use.
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DNA samples were diluted in TE buffer (20 mM Tris-HCl, 1 mM EDTA and 200 mM NaCl) to a concentration of ~0.3 ng/μL for AFM analysis. The diluted DNA solution of 50 μL was deposited on the modified mica for 2 min to allow the DNA to adsorb on the mica surface. Additional 40 μL TE buffer was added to fill the gap between the mica and AFM cantilever. The silicon tips used for AFM analysis were SCANASYST-FLUID+ (tip radius: ~2 nm; resonance frequency: ~150 kHz; spring constant: ~0.7 N/m; length: 70 μm; width: 10 μm). AFM images were taken under the Dimension Icon AFM (Bruker) in Peak Force Tapping mode in fluid. The scan rate was 1 Hz with 512 pixels per line. The scan size was first set to 2 μm and then 1 μm or 500 nm. The imaging data were flattened using the NanoScope Analysis software (1.4 version). The DNA contour length in the AFM images was calculated using ImageJ. Only molecules that were entirely imaged in the scanned area were measured and the overlapping molecules were excluded41. For each sample, about fifty measurements were recorded on the image. Reversible DNA polymerization on the microparticle surface A typical procedure to study the reversible DNA polymerization on microparticle surface is described as follows. Streptavidin-coated microparticles (0.1 mg) were mixed with biotinylated DI (50 nM) in 20 μL of reaction buffer (PBS, 0.1% v/v Tween 20, 0.02% w/v NaN3) at room temperature for 1 h on a rotator. To synthesize LP on the microparticles, they Angew Chem Int Ed Engl. Author manuscript; available in PMC 2016 June 01.
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were incubated in 20 μL of reaction buffer containing two DMs (0.5 μM) at room temperature overnight on a rotator. To synthesize BP on the microparticles, the secondary set of DMs (1 μM) was reacted with the microparticles with LP for another ~12 h. To depolymerize the DNA polymers on the microparticle surface, molecular triggers were added and the reaction lasted for 1 h. The microparticles were washed via centrifugation (10,000 × g for 8 min, twice) after polymerization and depolymerization before examination. Microparticle suspensions were dropped on a Teflon-coated glass slide and imaged using Maestro system with a blue light (Ex, 455 nm) and a blue filter (Em>488 nm). The microparticles were also examined using guava easyCyte™ flow cytometer (Millipore) with a blue light (Ex, 488 nm) and a green filter (Em, 525±30 nm). The results were post analyzed using guavaSoft 2.6 software and the geometric means were used to represent the mean fluorescence intensity.
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Reversible DNA polymerization in the ECM of living cells
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HDFns were used as the model for the tests of reversible DNA polymerization in the ECM. HDFns were cultured in medium 106 (M106) supplemented with recommended low serum growth supplement, gentamicin (10 μg/mL) and amphotericin B (0.25 μg/mL) at 37°C with 5% CO2. The cells between passage 2~4 were used in all experiments. The cells were seeded into 8-well chamber slide (ibidi) and used at approximately 80% confluence. The cells were treated using the standard immunostaining procedure. Briefly, the cells were counterstained with blue CMAC dye (5 μM) in PBS for 0.5 h and then incubated in M106 for another 0.5 h at 37°C. After washing, the cells were blocked with the binding buffer (M106 with 2% w/v BSA) for 0.5 h at 37°C and treated with biotinylated primary antibody (20 μg/mL in binding buffer) for 1 h at 37°C. After washing with M106, they were sequentially treated with streptavidin (10 μg/mL in binding buffer) for 45 min at 37°C and biotinylated DIL (1 μM in M106) for 45 min at 25°C. Finally they were treated with FAMlabeled DMs (1 μM for linear DNA polymerization and 2.5 μM for branched DNA polymerization) and molecular triggers (2 μM of RL and 5 μM of RB) for 1 h, respectively at 25°C. The cells in the chamber slide were imaged using a laser scanning confocal microscope (Olympus FV1000) equipped with FV10-ASW 3.0 software and 60 × (PlanApo) oil objectives. CMAC were excited with violet laser (405 nm) and FAM was excited with Blue Argon (488 nm). The green fluorescence intensity of each sample (at least 3 images) was quantified using ImageJ software.
Supplementary Material Refer to Web version on PubMed Central for supplementary material.
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Acknowledgments We thank the Huck Institute Microscopy Facilities for technical support and Mrs. Erin Gaddes for editing the manuscript. This work was supported in part by the U.S. NSF INSPIRE program (DMR-1322332), the National Heart, Lung, and Blood Institute of the NIH (R01HL122311) and the Penn State Start-Up Fund.
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References
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1. a) Oshovsky GV, Reinhoudt DN, Verboom W. Angew Chem Int Ed. 2007; 46:2366.b) Lehn JM. Science. 1985; 227:849. [PubMed: 17821215] c) Rowan SJ, Cantrill SJ, Cousins GRL, Sanders JKM, Stoddart JF. Angew Chem, Int Ed Engl. 2002; 41:898. [PubMed: 12491278] d) Jin Y, Yu C, Denman RJ, Zhang W. Chem Soc Rev. 2013; 42:6634. [PubMed: 23749182] e) Imato K, Ohishi T, Nishihara M, Takahara A, Otsuka H. J Am Chem Soc. 2014; 136:11839. [PubMed: 25098856] 2. a) Lehn JM. Prog Polym Sci. 2005; 30:814.b) Mozhdehi D, Ayala S, Cromwell OR, Guan Z. J Am Chem Soc. 2014; 136:16128. [PubMed: 25348857] c) Fan J, Lal Saha M, Song B, Schönherr H, Schmittel M. J Am Chem Soc. 2012; 134:150. [PubMed: 22148543] d) Bapat AP, Roy D, Ray JG, Savin DA, Sumerlin BS. J Am Chem Soc. 2011; 133:19832. [PubMed: 22103352] e) Brunsveld L, Folmer BJB, Meijer EW, Sijbesma RP. Chem Rev. 2001; 101:4071. [PubMed: 11740927] f) Alemdaroglu FE, Safak M, Wang J, Berger R, Herrmann A. Chem Commun. 2007:1358. 3. Sijbesma RP, Beijer FH, Brunsveld L, Folmer BJ, Hirschberg JH, Lange RF, Lowe JK, Meijer EW. Science. 1997; 278:1601. [PubMed: 9374454] 4. a) Ying H, Zhang Y, Cheng J. Nat Commun. 2014; 5:1.b) Wojtecki RJ, Meador MA, Rowan SJ. Nat Mater. 2011; 10:14. [PubMed: 21157495] c) Aida T, Meijer EW, Stupp SI. Science. 2012; 335:813. [PubMed: 22344437] 5. Bischoff F. Clin Chem. 1972; 18:869. [PubMed: 4559470] 6. a) Göpferich A. Biomaterials. 1996; 17:103. [PubMed: 8624387] b) Middleton JC, Tipton AJ. Biomaterials. 2000; 21:2335. [PubMed: 11055281] 7. Dirks RM, Pierce NA. Proc Natl Acad Sci U S A. 2004; 101:15275. [PubMed: 15492210] 8. a) Wetmur JG, Davidson N. J Mol Biol. 1968; 31:349. [PubMed: 5637197] b) Lee CS, Davis RW, Davidson N. J Mol Biol. 1970; 48:1. [PubMed: 4915293] c) Meselson MS, Radding CM. Proc Natl Acad Sci U S A. 1975; 72:358. [PubMed: 1054510] 9. Seeman NC. Nature. 2003; 421:427. [PubMed: 12540916] 10. a) Rothemund PWK. Nature. 2006; 440:297. [PubMed: 16541064] b) Zhang F, Nangreave J, Liu Y, Yan H. J Am Chem Soc. 2014; 136:11198. [PubMed: 25029570] c) Feldkamp U, Niemeyer CM. Angew Chem, Int Ed. 2006; 45:1856.d) Aldaye FA, Palmer AL, Sleiman HF. Science. 2008; 321:1795. [PubMed: 18818351] 11. a) Krishnan Y, Simmel FC. Angew Chem, Int Ed. 2011; 50:3124.b) Seeman NC. Trends Biochem Sci. 2005; 30:119. [PubMed: 15752983] c) Bath J, Turberfield AJ. Nat Nanotechnol. 2007; 2:275. [PubMed: 18654284] d) Surana S, Shenoy AR, Krishnan Y. Nat Nanotechnol. 2015; 10:741. [PubMed: 26329110] e) Chen YJ, Groves B, Muscat RA, Seelig G. Nat Nanotechnol. 2015; 10:748. [PubMed: 26329111] 12. a) Zheng J, Zhu G, Li Y, Li C, You M, Chen T, Song E, Yang R, Tan W. ACS Nano. 2013; 7:6545. [PubMed: 23841478] b) Zhu G, Zheng J, Song E, Donovan M, Zhang K, Liu C, Tan W. Proc Natl Acad Sci U S A. 2013; 110:7998. [PubMed: 23630258] c) Chen N, Huang Y, Wang Y. Biomaterials. 2014; 35:9709. [PubMed: 25176063] 13. a) Choi HMT, Chang JY, Trinh LA, Padilla JE, Fraser SE, Pierce NA. Nat Biotechnol. 2010; 28:1208. [PubMed: 21037591] b) Choi J, Routenberg Love K, Gong Y, Gierahn TM, Love JC. Anal Chem. 2011; 83:6890. [PubMed: 21812465] c) Huang J, Wu Y, Chen Y, Zhu Z, Yang X, Yang CJ, Wang K, Tan W. Angew Chem, Int Ed. 2011; 50:401.d) Niu S, Jiang Y, Zhang S. Chem Commun. 2010; 46:3089.e) Chen Y, Xu J, Su J, Xiang Y, Yuan R, Chai YQ. Anal Chem. 2012; 84:7750. [PubMed: 22924989] f) Bi S, Chen M, Jia X, Dong Y, Wang Z. Angew Chem, Int Ed. 2015; 54:8144. 14. a) Zhu G, Zhang S, Song E, Zheng J, Hu R, Fang X, Tan W. Angew Chem, Int Ed. 2013; 52:5490.b) Chen N, Li S, Battig MR, Wang Y. Small. 2013; 9:3944. [PubMed: 23813897] 15. a) Wu P, Nakano SI, Sugimoto N. Eur J Biochem. 2002; 269:2821. [PubMed: 12071944] b) Dillenback LM, Goodrich GP, Keating CD. Nano Lett. 2006; 6:16. [PubMed: 16402780] c) Van Der Meulen SAJ, Leunissen ME. J Am Chem Soc. 2013; 135:15129. [PubMed: 24040916] d) Song J, Arbona JM, Zhang Z, Liu L, Xie E, Elezgaray J, Aime JP, Gothelf KV, Besenbacher F, Dong M. J Am Chem Soc. 2012; 134:9844. [PubMed: 22646845]
Angew Chem Int Ed Engl. Author manuscript; available in PMC 2016 June 01.
Chen et al.
Page 10
Author Manuscript
16. a) Thomas CAJ. J Am Chem Soc. 1956; 78:1861.b) Addelhady HG, Allen S, Davies MC, Roberts CJ, Tendler SJB, Williams PM. Nucleic Acids Res. 2003; 31:4001. [PubMed: 12853616] c) Bezanilla M, Drake B, Nudler E, Kashlev M, Hansma PK, Hansma HG. Biophys J. 1994; 67:2454. [PubMed: 7696484] d) Hoffecker IT, Takemoto N, Arima Y, Iwata H. Biomaterials. 2015; 53:318. [PubMed: 25890730] 17. a) Cui H, Chen Z, Zhong S, Wooley KL, Pochan DJ. Science. 2007; 317:647. [PubMed: 17673657] b) Kostiainen MA, Kasyutich O, Cornelissen JJLM, Nolte RJM. Nat Chem. 2010; 2:394. [PubMed: 20414241] 18. Kim J, Liu Y, Ahn SJ, Zauscher S, Karty JM, Yamanaka Y, Craig SL. Adv Mater. 2005; 17:1749.
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Schematic illustration of the concept. a) Reversible polymerization of LP (upper panel) and BP (lower panel). P: polymerization; R: regulated depolymerization using the reversing trigger molecules (i.e., T). b) and c) Molecular structures of monomer reactants, LP (b) and BP (c) for reversible polymerization. The labeling letters are described in the text. Branches with the leaf-like structure in (a) were drawn for schematic illustrtion only. In principle, branching can occur from each DM2k unit as shown in (c).
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Examination of LP reversibility. a) and b), Gel electrophoresis image and AFM images showing LP synthesis. All samples in (a) were incubated for the same time before gel electrophoresis. c) and d), Gel electrophoresis image and AFM images showing the reversibility of LP. Lane 1 of c: DM1s1, DM2 and T1 were mixed together and annealed to form the hybridized complex; lane 6 of c: LP with the s1 side group was treated by T1 after the formation of LP. The red boxes in lanes 1 and 6 demonstrate that LP with the s1 side group was reversed by T1. T2 was used herein as the control of T1 to show the sequencespecific reversibility of LP.
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Examination of BP reversibility. a) and b), Gel electrophoresis image and AFM images showing BP synthesis. Lanes 1 and 3 in the gel image (a) show the bands of LP without the k side group before and after the treatment of DM3s2 and DM4, respectively; lanes 2 and 4 show the bands of LP with the k side group before and after the treatment of DM3s2 and DM4, respectively. c) and d), Gel electrophoresis image and AFM images showing the reversibility of BP. Lane 1 in the gel image (c): BP with the s1 and s2 side groups; lanes 2–4: BP with s1 and s2 side groups treated with T1 (lane 2), T2 (lane 3) and T1+T2 (lane 4), respectively; lane 5: control BP without side groups; lane 6: control BP treated with T1+T2. In all cases, triggers were added after the formation of BP.
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Examination of the reversibility on the microparticle surface and in the extracellular matrix. a) Flow cytometry histograms showing syntheses of LP and BP on the microparticle surface. b) Flow cytometry histograms showing the reversibility of BP treated with molecular triggers. T: triggering solution (containing T1 and T2). c) Confocal images showing LP and BP syntheses in the ECM. Green: DNA polymers; blue: CMAC. d) Images of the ECM with BP before and after treatment with the triggering solution. e) Fluorescence profiles of the lines drew in the images shown in d. f) Comparison of fluorescence intensities of the entire confocal images shown in d before and after the triggering treatment (n=3). For both depolymerization on microparticle or cell surface, triggers were added after the formation of polymers on them.
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Angew Chem Int Ed Engl. Author manuscript; available in PMC 2016 June 01. Published in final edited form as: Angew Chem Int Ed Engl. 2016 June 1; 55(23): 6657–6661. doi:10.1002/anie.201601008.
Molecularly Regulated Reversible DNA Polymerization Dr. Niancao Chenb, Mr. Xuechen Shib, and Yong Wanga [Associate Professor] Yong Wang: [email protected] aDepartment
of Biomedical Engineering, Pennsylvania State University, 232 Hallowell Building, University Park, PA 16802
bDepartment
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of Biomedical Engineering, Pennsylvania State University, 202 Hallowell Building, University Park, PA 16802
Abstract Natural polymers are synthesized and decomposed under physiological conditions. However, it is challenging to develop synthetic polymers whose formation and reversibility can be both controlled under physiological conditions. Here we show that both linear and branched DNA polymers can be synthesized via molecular hybridization in aqueous solutions, on the particle surface, and in the extracellular matrix (ECM) without the involvement of any harsh conditions. More importantly, these polymers can be effectively reversed to dissociate under the control of molecular triggers. Since nucleic acids can be conjugated with various molecules or materials, we anticipate that molecularly regulated reversible DNA polymerization holds potential for broad biological and biomedical applications.
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TOC image Regulated reversible DNA polymerization: It is challenging to emulate nature in developing synthetic polymers with dynamic reversibility under physiological conditions. This study demonstrates that linear and branched DNA polymers can be synthesized and reversed under the control of molecular triggers without the involvement of any harsh conditions.
Author Manuscript Correspondence to: Yong Wang, [email protected]. Supporting information for this article is given via a link at the end of the document.
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Keywords DNA; dynamic polymers; hybridization; reversible polymerization; self-assembly
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Synthetic polymers are broadly used in various applications such as medicine, cloth, devices, and tools. However, their synthesis and decomposition usually require harsh conditions (e.g., high temperature or organic solvent). In contrast, natural polymers (e.g., proteins) could be synthesized or decomposed with the aid of enzymes under mild physiological conditions.[1] Thus, great efforts have been made to synthesize dynamic polymers with reversible covalent bonds and non-covalent interactions via diverse mechanisms.[2] For instance, units of 2ureido-4-pyrimidone can form a self-complementary array of four hydrogen bonds for synthesis of unidirectional polymers with the reversibility of formation and decomposition.[3] These polymers not only provide a way of understanding the major processes in nature but also hold potential for broad applications.[4] However, synthetic monomers have been mostly studied in organic solvents and their polymers usually do not have biocompatibility;[5] moreover, these polymers do not exhibit regulatable reversibility under physiological conditions.[6]
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The purpose of this study was to demonstrate the ability to develop dynamic polymers whose synthesis and reversibility can be both controlled at the molecular level under physiological conditions (Figure 1), which has not been reported before. The polymers were synthesized using the principle of hybridization chain reaction;[7] the polymers were reversed using the principle of branch displacement.[8] The synthesis of the linear polymer (LP, Figure 1a, upper panel) involves three molecules including a DNA initiator (DI) and two DNA monomers (DMs). The DI is a linear structure with one functional domain as labeled with i (Figure 1b and Supporting Information, Table S1a). The DMs are hairpin structures. DM1s1 has three domains including i*, j and s1 and DM2 has two domains including i and j*. During the polymerization (Figure S1a), DI opens the hairpin structure of DM1s1 to form an i–i* double helix with j and s1 left as a linear segment. The linear j
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domain further reacts with the Notably, the domain s1 of DM1s1 does not participate in the linear polymerization and is a functional side group of LP. Thus, s1 can hybridize with a molecule carrying a complementary sequence domain s1* (Figures 1b and S1b). The molecular trigger T1 has two functional domains including s1* and j*. With the aid of s1–s1* and j–j* hybridization, T1 hybridizes with the DM1s1 unit of LP and displaces DM2. Resultantly, LP is reversed without the involvement of any non-physiological factors.
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Similar to DM1s1, DM2 can be designed with a functional side group (named as DM2k, Figure 1c and Table S1a) to bear a total of three domains including k, i and j*. Through the polymerization of DM1s1 and DM2k, LP acquires two functional side groups, s1 and k (Figure 1c). Since k can be designed as an initiator for DM3s2 and DM4, these two DMs react with the side group k of LP to form side chains along the backbone of LP (Figures 1c and S2). Resultantly, a branched polymer (BP) is synthesized with LP and two DMs (Figure 1a, lower panel). Because both DM1s1 and DM3s2 have the side groups, i.e., s1 and s2, respectively, the use of two corresponding molecular triggers T1 and T2 can induce the reversible polymerization of BP (Figures 1c and S2).
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The variation of DM structures would lead to the formation of different forms of LPs (Table S1b). Four representative LPs were synthesized (Figure S3). The results showed that DMs formed LPs in the presence of DI. Side groups did not apparently influence the LP formation. The apparent molecular weights of LPs mostly fell in the range between 500 to 3,000 bp. In contrast, DMs did not form LPs in the absence of DI (Figure 2a, lane 4). The result also showed that an inhibiting sequence could effectively terminate LP polymerization (Figure S4), suggesting that LPs were formed through sequential hybridization of DMs (Figure S1a). The increase of the reaction time led to more DM conversion (Figure S5). Specifically, ~60% of DMs were quickly converted into LPs within 0.5–1 h. The polymerization reached plateau at 8 h with a cumulative conversion efficiency of 86%. The successful synthesis of LPs was also confirmed using AFM (Figure 2b). The average contour length of LPs was ~122.8 nm (Figure S6a). The side group s1 had 10 nucleotides (nt). When it was incorporated into the 3′ end of DM1, the formed LP had periodic side functional groups (Figure S1a). T1 had 34 nt with 10 nt complementary to s1 and another 24 nt complementary to the j domain of DM1s1. When T1 and DM1s1 hybridize (Figure S7), the formed double-stranded helix has 34 base pairs (bp). In comparison, DM2 and DM1s1 have a helix of 24 bp. The former is thermodynamically more stable than the latter. Thus, T1 is more competitive than DM2 in hybridization with DM1s1.
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As shown in Figure 2c, the LPs bearing the s1 groups virtually disappeared after reacting with T1 (lane 6). The major reaction products (lane 6) were located in the same position as the hybridization complex of DM1s1, DM2, and T1 (lane 1). By comparison, the LP bands were not affected after mixing with the control triggering sequence T2 (lane 5). When LPs bearing no s1 groups were treated with T1, LPs barely changed (lanes 2 and 3). These results clearly demonstrate that T1 effectively triggered LP depolymerization in a sequence-specific and s1-dependent manner.
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The gel electrophoresis results were confirmed by AFM analysis. The morphologies of LPs with s1 side groups after polymerization and depolymerization are shown in Figure 2b&d and Figure S8a. Quantitatively, the average length of LPs decreased from 122.8 nm to 16.4 nm (Figure S6). The measured length of depolymerization products is consistent with the calculated length (Figure S8b). These data further confirm the success of T1-triggered LP depolymerization.
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We also studied the effects of three different hybridization parameters on depolymerization. The reaction time was varied from 10 min to 8 h (Figure S9). The result showed that 95% LPs disappeared within 10 min, suggesting that depolymerization was fast. The depolymerization efficiency increased with the increasing molar ratio of T1 to DM1s1, exhibiting nearly a linear relationship with the molar ratio (Figure S10). The length of T1 was also varied (Figure S11). Before the hybridization length was reduced to 25 bp, T1 could effectively trigger LP depolymerization. The efficiency gradually decreased with the hybridization length reduced from 25 to 16 bp. This trend corresponds to the ΔG change of the hybridization helixes (Figure S11d).
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BP was synthesized using filtered LP and the mixture of DM3 and DM4. The molar ratio of LP monomers to branch monomers was 1:5. LP had two side groups, s1 and k (Figure S12). The side group k had 29 nt. The 5 nt adjacent to the backbone of LP were designed as a spacer to mitigate potential steric hindrance. The other 24 nt were used as an initiator for reaction with DM3 and DM4. The result showed that k was able to induce polymerization of DM3 and DM4 (Figure S13a) similar to the DI-induced polymerization of DM1 and DM2. We further used k-bearing LP to react with DM3 (or DM3s2) and DM4 for BP synthesis. A clear band shift is shown in the gel images (lane 4 in Figure 3a and Figure S13b). The results showed that a significant amount of BP molecules had apparent molecular weights over 12,000 bp. In contrast, the reaction of LP without k and the two DMs did not lead to a band shift (lane 3 in Figure 3a and Figure S13b). This difference demonstrates that BP could be synthesized through the k-bearing LP and the two DMs. AFM imaging was performed to confirm the gel electrophoresis data (Figures 3b and S14a). Future work can be further performed to quantify the amount of monomer incorporation. DM1s1 and DM3s2 both had a side group (s1 or s2). They were designed to hybridize with their corresponding molecular triggers T1 and T2, respectively. Figure S15 showed that the addition of T1 into LP1 solution led to the disappearance of LP1 band (lane 6 in Figure S15a) and the addition of T2 into LP2 solution led to the disappearance of LP2 band (lane 6 in Figure S15b). These results demonstrate that molecular triggers could reverse LP successfully.
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Figure 3c shows that BP treated with different combinations of T1 and T2 exhibited a remarkably band shift (lanes 2–4) in comparison to the untreated BP (lane 1). For BP treated with only T1, the apparent molecular weights of reversed BP products were shifted into the range of 300 to 1,000 bp (lane 2, population i). It suggests that T1 depolymerized the backbone of BP while not affecting the side chains (Figure S16a). For BP treated with only T2, the apparent molecular weights of reversed BP fell into two ranges. One was from 500 to 3,000 bp (population ii); the other was at ~100 bp (population iii). It suggests that T2
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reversed the side chains of BP while not affecting the backbone (Figure S16b). For BP treated with the mixture of T1 and T2 (lane 4), two bright bands were observed. It suggests that both the backbone and side chains were depolymerized (Figure S2c). By contrast, BP without s1 and s2 did not show any change (lanes 5 and 6). These results were confirmed by AFM imaging (Figures 3d and S17). In addition to demonstrating polymerization and depolymerization of LPs and BPs in the aqueous solution, we further combined these polymeric systems with synthetic microparticle and antibody and validated their working efficiency to demonstrate the potential applications of reversible DNA polymerization.
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Polystyrene microparticles were functionalized with DI via streptavidin-biotin interactions and DM2 was labeled with fluorophore. The flow cytometry analysis showed that the average fluorescence intensity of the microparticles with LP was approximately one order of magnitude higher than that of control with one hybridized DM1–DM2 complex (Figure S18). It demonstrates that LP could be synthesized on the microparticle surface. Similarly, the results showed that BP could be synthesized on the microparticle surface (Figures 4a and S19). Experiments were further carried out to examine the reversibility. Microparticles bearing LP were treated with either T1 or T2 (Figure S20). T1 treatment induced a dramatic decrease of fluorescence intensity on microparticles whereas T2 treatment did not. These data demonstrate that LP was specifically reversed on the microparticle surface. Microparticles bearing BP were also treated with the triggering solution (Figures 4b and S21). Similar to LP, BP was reversed on the microparticle surface.
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We also used the ECM of living cells as a substrate to study LP and BP. An antibody-DI conjugate was prepared via the formation of a sandwich structure of biotinylated antibody, streptavidin and biotinylated DI. The antibody-DI conjugate induced the formation of LP in the ECM (Figure S22), and that LP further induced the formation of BP (Figures 4c and S23). When the ECM was treated by triggering solutions, the fluorescence intensities decreased in both cases of LP (Figure S24) and BP (Figures 4d–f and S25) to virtually the same level as the background. While the intensity of the blue background was slightly decreased during the second time imaging of the ECM after the triggering treatment due to photobleaching, Figure S25 demonstrates that triggering depolymerization rather than photobleaching was the dominant reason for the decrease of green fluorescence intensity. Thus, the data show that LP or BP can be polymerized and depolymerized in the ECM.
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Owing to high fidelity of hybridization and the water-soluble nature,[9] nucleic acids have been extensively explored in constructing complex structures[10] or nanodevices.[11] These systems have been further studied for numerous applications such as drug delivery,[12] molecular detection[13] and bioimaging.[14] However, previous work has not demonstrated the development of DNA polymers whose formation and reversibility can be both regulated under physiological conditions. This work has successfully demonstrated the ability to synthesize and reverse linear and branched DNA polymers in aqueous solutions, on the microparticle surface, and in the ECM, which further advances the development of nucleic acid-based material systems. We realize that increasing temperature[15] or using DNase[16] can lead to depolymerization of the DNA polymers; however, these methods are not
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physiologically biocompatible or do not provide selectivity of reversing a certain DNA polymer if multiple polymers exist. Similar to many other self-assembly systems,[3,17] it is also possible to design short self-complementary DNA oligonucleotides that can be spontaneously self-assembled and concomitantly disassembled. However, the spontaneous self-assembly and concomitant disassembly are not stable or molecularly regulatable.[18] In summary, this work has demonstrated that intermolecular hybrization is an effective mechanism for the synthesis of linear and branched DNA polymers with regulatable reversibility under strict physiological conditions. Since nucleic acids can be conjugated with various molecules or materials, molecularly regulated reversible polymerization and depolymerization are envisioned to hold potential for broad applications.
Methods Author Manuscript
Chemicals and reagents
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Phosphate buffered saline (PBS), Tris-EDTA and agarose were purchased from Fisher Scientific (Suwanee, GA). Oligonucleotides (Supporting Tab. 2) were purchased from Integrated DNA Technologies (Coralville, IA). The streptavidin-coated microparticles (average diameter: 5 μm) was obtained from Spherotech (Lake Forest, IL). The DNA marker, SYBR-Safe, CellTracker™ Blue CMAC dye, bovine serum albumin (BSA), human dermal fibroblasts, neonatal (HDFns) and cell culture reagents were purchased from Invitrogen (Carlsbad, CA). Ultracentrifugal filter (100K cut-off size) and biotinylated anticollagen IgG were purchased from Millipore (Billerica, MA). All other chemicals, such as (3-aminopropyl) triethoxysilane (APTES), were obtained from Sigma-Aldrich (Louis, MO). Mica was purchased from Asheville-Schoonmaker Mica Company (Newport News, VA). Atomic force microscope (AFM) cantilever (SCANASYST-FLUID+) was purchased from Bruker (Camarilla, CA). Reversible DNA polymerization in aqueous solution
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DNA sequences were designed using NUPACK server (www.nupack.org). Their secondary structures were also generated using NUPACK. Before hybridization reactions, DNA macromonomers were heated to 95°C and then cooled to room temperature in PBS for 1 h. During the procedure of LP synthesis, DI was mixed with the two DMs at room temperature. To synthesize BP, LP was filtered twice through an ultracentrifugal filter (100K cut-off size) at 14,000 × g to remove the unreacted DMs. The filtered LP products were incubated with the secondary set of DMs (2 μM each) at room temperature. The reaction time for each polymerization was 12 h unless specified elsewhere. To depolymerize the DNA polymers, molecular triggers were added into the solutions of LP or BP and reacted for predetermined periods of time at room temperature. The molar ratio of triggers to their corresponding monomers was all 2:1 unless specified elsewhere. To prepare hybridized DNA complexes, the mixture of molecular triggers and their corresponding monomers with a molar ratio of 1:1 was heated to 95 °C and then gradually cooled to room temperature. To examine the effect of the terminator on polymerization, DI (initially 0.3 μM), each DM (initially 3.0 μM) and terminator of different concentrations were prepared individually and then subjected to
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the annealing treatment before mixed together. The mixture was examined after 12 h incubation using agarose gel electrophoresis. Gel electrophoresis Agarose gel (1%) was used for gel electrophoresis in Tris-borate-EDTA (TBE) buffer (89 mM boric acid and 2 mM EDTA, pH 8.2) The agarose gel was pre-stained with SYBR-Safe (0.1 μL of stock solution per mL of agarose gel). The gel electrophoresis was run at 100 V for 60 min and the gels were imaged using a CRI Maestro EX System (Woburn, MA) with a blue exciting light (455 nm). The fluorescence of DNA in gels were recorded with a blue filter (Em>488 nm). ΔG calculation
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The ΔG values of hybridized DNA macromonomers were calculated using IDT OligoAnalyzer 3.1 (https://www.idtdna.com/calc/analyzer) under the condition of 0.25 μM of DNA and 140 mM Na+. Atomic force microscopy (AFM)
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Mica was modified to bear positive charges on its surface before sample loading and imaging. Briefly, mica with a dimension of ~1.5 cm in square was cut into an octagon shape and freshly cleaved with a thickness of 0.1 mm using a clean razor blade. The mica was attached with a metal clip on its edge to a glass rod in a desiccator, which was filled with argon gas. Two eppendorf caps were placed on the bottom of the desiccator to hold 90 μL of APTES and 30 μL of N,N-diisopropylethylamine (DIPEA). The mica in the desiccator was reacted with the evaporated chemicals for 2 h. The desiccator was purged with argon for 2 min and the mica was cured in the desiccator before use.
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DNA samples were diluted in TE buffer (20 mM Tris-HCl, 1 mM EDTA and 200 mM NaCl) to a concentration of ~0.3 ng/μL for AFM analysis. The diluted DNA solution of 50 μL was deposited on the modified mica for 2 min to allow the DNA to adsorb on the mica surface. Additional 40 μL TE buffer was added to fill the gap between the mica and AFM cantilever. The silicon tips used for AFM analysis were SCANASYST-FLUID+ (tip radius: ~2 nm; resonance frequency: ~150 kHz; spring constant: ~0.7 N/m; length: 70 μm; width: 10 μm). AFM images were taken under the Dimension Icon AFM (Bruker) in Peak Force Tapping mode in fluid. The scan rate was 1 Hz with 512 pixels per line. The scan size was first set to 2 μm and then 1 μm or 500 nm. The imaging data were flattened using the NanoScope Analysis software (1.4 version). The DNA contour length in the AFM images was calculated using ImageJ. Only molecules that were entirely imaged in the scanned area were measured and the overlapping molecules were excluded41. For each sample, about fifty measurements were recorded on the image. Reversible DNA polymerization on the microparticle surface A typical procedure to study the reversible DNA polymerization on microparticle surface is described as follows. Streptavidin-coated microparticles (0.1 mg) were mixed with biotinylated DI (50 nM) in 20 μL of reaction buffer (PBS, 0.1% v/v Tween 20, 0.02% w/v NaN3) at room temperature for 1 h on a rotator. To synthesize LP on the microparticles, they Angew Chem Int Ed Engl. Author manuscript; available in PMC 2016 June 01.
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were incubated in 20 μL of reaction buffer containing two DMs (0.5 μM) at room temperature overnight on a rotator. To synthesize BP on the microparticles, the secondary set of DMs (1 μM) was reacted with the microparticles with LP for another ~12 h. To depolymerize the DNA polymers on the microparticle surface, molecular triggers were added and the reaction lasted for 1 h. The microparticles were washed via centrifugation (10,000 × g for 8 min, twice) after polymerization and depolymerization before examination. Microparticle suspensions were dropped on a Teflon-coated glass slide and imaged using Maestro system with a blue light (Ex, 455 nm) and a blue filter (Em>488 nm). The microparticles were also examined using guava easyCyte™ flow cytometer (Millipore) with a blue light (Ex, 488 nm) and a green filter (Em, 525±30 nm). The results were post analyzed using guavaSoft 2.6 software and the geometric means were used to represent the mean fluorescence intensity.
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Reversible DNA polymerization in the ECM of living cells
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HDFns were used as the model for the tests of reversible DNA polymerization in the ECM. HDFns were cultured in medium 106 (M106) supplemented with recommended low serum growth supplement, gentamicin (10 μg/mL) and amphotericin B (0.25 μg/mL) at 37°C with 5% CO2. The cells between passage 2~4 were used in all experiments. The cells were seeded into 8-well chamber slide (ibidi) and used at approximately 80% confluence. The cells were treated using the standard immunostaining procedure. Briefly, the cells were counterstained with blue CMAC dye (5 μM) in PBS for 0.5 h and then incubated in M106 for another 0.5 h at 37°C. After washing, the cells were blocked with the binding buffer (M106 with 2% w/v BSA) for 0.5 h at 37°C and treated with biotinylated primary antibody (20 μg/mL in binding buffer) for 1 h at 37°C. After washing with M106, they were sequentially treated with streptavidin (10 μg/mL in binding buffer) for 45 min at 37°C and biotinylated DIL (1 μM in M106) for 45 min at 25°C. Finally they were treated with FAMlabeled DMs (1 μM for linear DNA polymerization and 2.5 μM for branched DNA polymerization) and molecular triggers (2 μM of RL and 5 μM of RB) for 1 h, respectively at 25°C. The cells in the chamber slide were imaged using a laser scanning confocal microscope (Olympus FV1000) equipped with FV10-ASW 3.0 software and 60 × (PlanApo) oil objectives. CMAC were excited with violet laser (405 nm) and FAM was excited with Blue Argon (488 nm). The green fluorescence intensity of each sample (at least 3 images) was quantified using ImageJ software.
Supplementary Material Refer to Web version on PubMed Central for supplementary material.
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Acknowledgments We thank the Huck Institute Microscopy Facilities for technical support and Mrs. Erin Gaddes for editing the manuscript. This work was supported in part by the U.S. NSF INSPIRE program (DMR-1322332), the National Heart, Lung, and Blood Institute of the NIH (R01HL122311) and the Penn State Start-Up Fund.
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References
Author Manuscript Author Manuscript Author Manuscript
1. a) Oshovsky GV, Reinhoudt DN, Verboom W. Angew Chem Int Ed. 2007; 46:2366.b) Lehn JM. Science. 1985; 227:849. [PubMed: 17821215] c) Rowan SJ, Cantrill SJ, Cousins GRL, Sanders JKM, Stoddart JF. Angew Chem, Int Ed Engl. 2002; 41:898. [PubMed: 12491278] d) Jin Y, Yu C, Denman RJ, Zhang W. Chem Soc Rev. 2013; 42:6634. [PubMed: 23749182] e) Imato K, Ohishi T, Nishihara M, Takahara A, Otsuka H. J Am Chem Soc. 2014; 136:11839. [PubMed: 25098856] 2. a) Lehn JM. Prog Polym Sci. 2005; 30:814.b) Mozhdehi D, Ayala S, Cromwell OR, Guan Z. J Am Chem Soc. 2014; 136:16128. [PubMed: 25348857] c) Fan J, Lal Saha M, Song B, Schönherr H, Schmittel M. J Am Chem Soc. 2012; 134:150. [PubMed: 22148543] d) Bapat AP, Roy D, Ray JG, Savin DA, Sumerlin BS. J Am Chem Soc. 2011; 133:19832. [PubMed: 22103352] e) Brunsveld L, Folmer BJB, Meijer EW, Sijbesma RP. Chem Rev. 2001; 101:4071. [PubMed: 11740927] f) Alemdaroglu FE, Safak M, Wang J, Berger R, Herrmann A. Chem Commun. 2007:1358. 3. Sijbesma RP, Beijer FH, Brunsveld L, Folmer BJ, Hirschberg JH, Lange RF, Lowe JK, Meijer EW. Science. 1997; 278:1601. [PubMed: 9374454] 4. a) Ying H, Zhang Y, Cheng J. Nat Commun. 2014; 5:1.b) Wojtecki RJ, Meador MA, Rowan SJ. Nat Mater. 2011; 10:14. [PubMed: 21157495] c) Aida T, Meijer EW, Stupp SI. Science. 2012; 335:813. [PubMed: 22344437] 5. Bischoff F. Clin Chem. 1972; 18:869. [PubMed: 4559470] 6. a) Göpferich A. Biomaterials. 1996; 17:103. [PubMed: 8624387] b) Middleton JC, Tipton AJ. Biomaterials. 2000; 21:2335. [PubMed: 11055281] 7. Dirks RM, Pierce NA. Proc Natl Acad Sci U S A. 2004; 101:15275. [PubMed: 15492210] 8. a) Wetmur JG, Davidson N. J Mol Biol. 1968; 31:349. [PubMed: 5637197] b) Lee CS, Davis RW, Davidson N. J Mol Biol. 1970; 48:1. [PubMed: 4915293] c) Meselson MS, Radding CM. Proc Natl Acad Sci U S A. 1975; 72:358. [PubMed: 1054510] 9. Seeman NC. Nature. 2003; 421:427. [PubMed: 12540916] 10. a) Rothemund PWK. Nature. 2006; 440:297. [PubMed: 16541064] b) Zhang F, Nangreave J, Liu Y, Yan H. J Am Chem Soc. 2014; 136:11198. [PubMed: 25029570] c) Feldkamp U, Niemeyer CM. Angew Chem, Int Ed. 2006; 45:1856.d) Aldaye FA, Palmer AL, Sleiman HF. Science. 2008; 321:1795. [PubMed: 18818351] 11. a) Krishnan Y, Simmel FC. Angew Chem, Int Ed. 2011; 50:3124.b) Seeman NC. Trends Biochem Sci. 2005; 30:119. [PubMed: 15752983] c) Bath J, Turberfield AJ. Nat Nanotechnol. 2007; 2:275. [PubMed: 18654284] d) Surana S, Shenoy AR, Krishnan Y. Nat Nanotechnol. 2015; 10:741. [PubMed: 26329110] e) Chen YJ, Groves B, Muscat RA, Seelig G. Nat Nanotechnol. 2015; 10:748. [PubMed: 26329111] 12. a) Zheng J, Zhu G, Li Y, Li C, You M, Chen T, Song E, Yang R, Tan W. ACS Nano. 2013; 7:6545. [PubMed: 23841478] b) Zhu G, Zheng J, Song E, Donovan M, Zhang K, Liu C, Tan W. Proc Natl Acad Sci U S A. 2013; 110:7998. [PubMed: 23630258] c) Chen N, Huang Y, Wang Y. Biomaterials. 2014; 35:9709. [PubMed: 25176063] 13. a) Choi HMT, Chang JY, Trinh LA, Padilla JE, Fraser SE, Pierce NA. Nat Biotechnol. 2010; 28:1208. [PubMed: 21037591] b) Choi J, Routenberg Love K, Gong Y, Gierahn TM, Love JC. Anal Chem. 2011; 83:6890. [PubMed: 21812465] c) Huang J, Wu Y, Chen Y, Zhu Z, Yang X, Yang CJ, Wang K, Tan W. Angew Chem, Int Ed. 2011; 50:401.d) Niu S, Jiang Y, Zhang S. Chem Commun. 2010; 46:3089.e) Chen Y, Xu J, Su J, Xiang Y, Yuan R, Chai YQ. Anal Chem. 2012; 84:7750. [PubMed: 22924989] f) Bi S, Chen M, Jia X, Dong Y, Wang Z. Angew Chem, Int Ed. 2015; 54:8144. 14. a) Zhu G, Zhang S, Song E, Zheng J, Hu R, Fang X, Tan W. Angew Chem, Int Ed. 2013; 52:5490.b) Chen N, Li S, Battig MR, Wang Y. Small. 2013; 9:3944. [PubMed: 23813897] 15. a) Wu P, Nakano SI, Sugimoto N. Eur J Biochem. 2002; 269:2821. [PubMed: 12071944] b) Dillenback LM, Goodrich GP, Keating CD. Nano Lett. 2006; 6:16. [PubMed: 16402780] c) Van Der Meulen SAJ, Leunissen ME. J Am Chem Soc. 2013; 135:15129. [PubMed: 24040916] d) Song J, Arbona JM, Zhang Z, Liu L, Xie E, Elezgaray J, Aime JP, Gothelf KV, Besenbacher F, Dong M. J Am Chem Soc. 2012; 134:9844. [PubMed: 22646845]
Angew Chem Int Ed Engl. Author manuscript; available in PMC 2016 June 01.
Chen et al.
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16. a) Thomas CAJ. J Am Chem Soc. 1956; 78:1861.b) Addelhady HG, Allen S, Davies MC, Roberts CJ, Tendler SJB, Williams PM. Nucleic Acids Res. 2003; 31:4001. [PubMed: 12853616] c) Bezanilla M, Drake B, Nudler E, Kashlev M, Hansma PK, Hansma HG. Biophys J. 1994; 67:2454. [PubMed: 7696484] d) Hoffecker IT, Takemoto N, Arima Y, Iwata H. Biomaterials. 2015; 53:318. [PubMed: 25890730] 17. a) Cui H, Chen Z, Zhong S, Wooley KL, Pochan DJ. Science. 2007; 317:647. [PubMed: 17673657] b) Kostiainen MA, Kasyutich O, Cornelissen JJLM, Nolte RJM. Nat Chem. 2010; 2:394. [PubMed: 20414241] 18. Kim J, Liu Y, Ahn SJ, Zauscher S, Karty JM, Yamanaka Y, Craig SL. Adv Mater. 2005; 17:1749.
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Schematic illustration of the concept. a) Reversible polymerization of LP (upper panel) and BP (lower panel). P: polymerization; R: regulated depolymerization using the reversing trigger molecules (i.e., T). b) and c) Molecular structures of monomer reactants, LP (b) and BP (c) for reversible polymerization. The labeling letters are described in the text. Branches with the leaf-like structure in (a) were drawn for schematic illustrtion only. In principle, branching can occur from each DM2k unit as shown in (c).
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Examination of LP reversibility. a) and b), Gel electrophoresis image and AFM images showing LP synthesis. All samples in (a) were incubated for the same time before gel electrophoresis. c) and d), Gel electrophoresis image and AFM images showing the reversibility of LP. Lane 1 of c: DM1s1, DM2 and T1 were mixed together and annealed to form the hybridized complex; lane 6 of c: LP with the s1 side group was treated by T1 after the formation of LP. The red boxes in lanes 1 and 6 demonstrate that LP with the s1 side group was reversed by T1. T2 was used herein as the control of T1 to show the sequencespecific reversibility of LP.
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Examination of BP reversibility. a) and b), Gel electrophoresis image and AFM images showing BP synthesis. Lanes 1 and 3 in the gel image (a) show the bands of LP without the k side group before and after the treatment of DM3s2 and DM4, respectively; lanes 2 and 4 show the bands of LP with the k side group before and after the treatment of DM3s2 and DM4, respectively. c) and d), Gel electrophoresis image and AFM images showing the reversibility of BP. Lane 1 in the gel image (c): BP with the s1 and s2 side groups; lanes 2–4: BP with s1 and s2 side groups treated with T1 (lane 2), T2 (lane 3) and T1+T2 (lane 4), respectively; lane 5: control BP without side groups; lane 6: control BP treated with T1+T2. In all cases, triggers were added after the formation of BP.
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Examination of the reversibility on the microparticle surface and in the extracellular matrix. a) Flow cytometry histograms showing syntheses of LP and BP on the microparticle surface. b) Flow cytometry histograms showing the reversibility of BP treated with molecular triggers. T: triggering solution (containing T1 and T2). c) Confocal images showing LP and BP syntheses in the ECM. Green: DNA polymers; blue: CMAC. d) Images of the ECM with BP before and after treatment with the triggering solution. e) Fluorescence profiles of the lines drew in the images shown in d. f) Comparison of fluorescence intensities of the entire confocal images shown in d before and after the triggering treatment (n=3). For both depolymerization on microparticle or cell surface, triggers were added after the formation of polymers on them.
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