Expanding the materials space of DNA via organic-phase ring-opening metathesis polymerization

Summary:

Herein, we develop a facile route to bring DNA to the organic phase, which greatly expands the types of structures accessible using DNA macromonomers. Phosphotriester- and exocyclic amine-protected DNA was synthesized and further modified with a norbornene moiety, which enables homopolymerization via ring-opening metathesis to produce brush-type DNA graft polymers in high yields. Subsequent deprotection cleanly reveals the natural phosphodiester DNA. The method not only achieves high molecular weight DNA graft polymers but when carried out at low monomer:catalyst ratios, yields oligomers that can be further fractionated to molecularly pure, monodisperse entities with one through ten DNA strands per molecule. In addition, we demonstrate substantial simplification in the preparation of traditionally difficult DNA-containing structures, such as DNA/poly(ethylene glycol) diblock graft copolymers and DNA amphiphiles. We envision that the marriage of oligonucleotides with the vast range of organic-phase polymerizations will result in many new classes of materials with yet unknown properties.

eTOC Blurb

Phosphate- and exocyclic amine-protected oligonucleotides are transformed into macromonomers for ring-opening metathesis polymerization in the organic phase. We demonstrate that homo- or copolymerization of the macromonomer followed by an “ultramild” deprotection step cleanly yields four distinct types of DNA-based structures: graft polymers, monodisperse oligomers, diblock graft copolymers, and diblock amphiphiles. These structures represent unprecedented or traditionally difficult DNA- containing polymers.

Graphical Abstract

Oligonucleotides have found extensive applications spanning structural DNA nanotechnology,^1–3 materials assembly,^4,5 DNA-encoded libraries,^6,7 and nucleic acid-based medicine.^8,9 In all of these fields it is often required for the oligonucleotide to be covalently attached to other moieties, be it a small molecule,^10–12 a macromolecule,^13,14 a crosslinked network,^15,16 or a nanoscopic/macroscopic surface.^17–19 These structures play irreplaceable roles in medicine, diagnostics, crystal engineering, drug discovery, among others.^20–24 The success of existing structures underscores the importance of developing new reactions and methods that can covalently arrange nucleic acids into a wide variety of well-defined architectures. Examining the chemical structure of the DNA conjugates reported thus far, two observations can be made, (i) Conjugates with a single oligonucleotide can be synthesized by solid-phase coupling or solution-phase coupling, (ii) Conjugates with multiple oligonucleotides are made by solution-phase coupling almost exclusively.^25,26 These observations reveal an important limitation of the solid-phase methodology: architecturally complex structures involving multiple nucleic acid strands, such as high-density DNA graft polymers and crosslinked networks, are difficult, if not impossible, to achieve on a rigid, two-dimensional surface. Yet, nucleic acid-containing materials with a non-linear, three-dimensional architecture remain at the forefront of exploration in chemistry, materials science, and medicine.^27–29 Solution-phase coupling provides only partial access to these materials. For example, high-density, multivalent DNA constructs are difficult because of the strong electrostatic repulsion of the negatively charged DNA.^30–32 In addition, there is oftentimes a difficulty in finding a common solvent for the DNA and its hydrophobic conjugation partner.^33,34

Among the non-linear architectures, brush-type oligonucleotides have attracted considerable attention due to the increased local nucleic acid density, which has been suggested as a factor that facilitates increased cellular endocytosis and enables carrier-free cellular gene regulation.^35,36 Gianneschi et al. reported the organic-phase polymerization of peptide nucleic acids (PNAs) to prepare brush-type polymers, which takes advantage of the non-charged nature of PNAs and their solubility in dimethyl formamide (DMF).³⁷ Nonetheless, the method cannot be readily extended to natural nucleic acids with phosphodiester backbones. Herrmann et al. adopted cationic surfactants for electrostatic complexation with DNA as a more general means to neutralize DNA negative charge and improve lipophilicity.³⁸ Notwithstanding broader applicability, the reaction yields are moderate, especially for larger oligonucleotides/coupling partners and for polymerization. In addition, the complete removal of surfactants can be challenging. Herein, we report an approach to bypass the common limitations imposed by solubility and nucleobase side reactivities, and expand the toolbox for organic-phase manipulation of oligonucleotides. The method centers around the idea of separating the conventional, single-step reaction to cleave and deprotect the oligonucleotides following solid-phase synthesis into two steps, which allows for the release of nucleobase-protected, charge-masked strands to be used for homogeneous organic-phase reaction. While strategies to protect/deprotect nucleotides have been extensively studied for solid-phase oligonucleotide synthesis and other applications,^39–42 the use of protected oligonucleotides in polymer chemistry remains unexplored.

For a proof-of-concept demonstration, we adopted the ring-opening metathesis polymerization (ROMP) to facilely convert norbornene-modified, protected DNA (protDNA) in high yields into three distinct classes of polymer-DNA nanostructures: brush-type DNA graft polymers, DNA/poly(ethylene glycol) (PEG) diblock graft copolymers, and DNA block copolymer micelles, which represent traditionally challenging DNA nanostructures for aqueous- or solid-phase techniques (Figure 1). The homopolymerization of protDNA not only yields high molecular weight (M_w > 380 kDa) DNA graft polymers but can also be employed to prepare molecularly pure oligomeric DNA with one through ten strands per molecule when a low monomer;catalyst ratio is used. Previously, oligomeric DNA star polymers with four, eight, and twelve arms have been painstakingly synthesized via coupling with a careful selection of multivalent cores, followed by several rounds of HPLC purification.⁴³ These monodisperse entities are scientifically valuable structures for accurately interrogating their interactions with living systems.⁴⁴ In a single reaction, a full series of molecular oligomers (including all odd-numbered ones) up to a decamer is obtained, which allows us to unambiguously verify the DNA density effect on cellular uptake. We envision that the results reported herein are the proverbial tip of an iceberg in DNA-based polymers: the introduction of DNA as a common monomer to the vast knowledge base of polymer chemistry should result in unlimited possibilities.

Figure 1.

Schematic representation of protDNA and its application in constructing DNA nanostructures.

Results and Discussion

Synthesis of the norbornenyl protDNA monomer.

The ester bond is a common base-labile linkage that tethers oligonucleotides to the controlled pore glass (CPG) for solid-phase oligonucleotide synthesis. Hydrolytic cleavage of the ester also results in the deprotection of the oligonucleotide. To release oligonucleotides in their protected form, a disulfide linker was used in place of the ester as it can be cleaved under soft reductive conditions. Tris-(2-carboxyethyl) phosphine (TCEP) quantitatively released the protected oligonucleotides from the CPG under weakly acidic conditions. The exposed sulfhydryl group was then used to couple with iodoacetyl- or maleimide-functionalized norbornene monomers via SN₂ or Michael addition, respectively, to produce the protDNA monomers (Scheme S1). Neutral or slightly basic conditions (e.g. phosphate buffered saline (PBS), pH = 7.4) were required for efficient coupling with the iodoacetyl group, which led to some inadvertent deprotection of the protDNA (mainly the release of the 2-cyanoethyl group) and loss of solubility in organic solvents. As such, the maleimide-sulfhydryl coupling reaction was used in subsequent preparations. To screen for compatibility with the polymerization and deprotection chemistries, a range of phosphoramidites were tested in the synthesis of the protDNA. The “ultramild” phosphoramidites (phenoxyacetyl dA, 4-isopropyl-phenoxyacetyl dG, and acetyl dC) were ultimately selected due to a combination of improved solubility and milder deprotection conditions. One or two 12-methylene (C12) spacers were incorporated at the 3’ to reduce the steric hindrance of the protDNA monomer and facilitate polymerization. Notably, the solubility of protDNA in organic solvents was also improved by the spacer(s). Unless otherwise indicated, the following protDNA macromonomer was used in the polymerization studies: 5’-fluorescein-CTC CAT GGT GCT CAC-(C12)₂-norbornene-3’ (Table S1). Electrospray ionization-mass spectrometry (ESI-MS) confirmed the successful synthesis of the protDNA macromonomer (exact mass: 8252.37 Da, calc. 8260.35 Da; Figure S1).

Homopolymerization of protDNA macromonomers.

Arranging DNA into a dense, highly oriented spherical form (termed spherical nucleic acids, or SNAs) can lead to the emergence of several unusual properties absent from linear or cyclic forms of DNA, such as increased binding affinity to a complementary strand and sharpening of the DNA melting transition (cooperative melting).²⁷ Interestingly, the cellular uptake of SNAs is considerably elevated (by 2–3 orders of magnitude) over free DNA, despite SNAs’ polyanionic nature.²⁷ It is not clear, however, whether there exists a distinct density threshold above which multivalent DNA structures behave like SNAs. The DNA graft polymers can in principle achieve SNA-like properties, and provide definitive answers to the density-uptake relationship question as density is a function of the degree of polymerization (DP), which is tunable. While post-polymerization “grafting-onto” chemistries can be used to access a similar architecture, incomplete grafting due to sterics and the strong charge repulsion ultimately results in limited DNA density.^27,37 A “grafting-through” synthesis using norbornenyl protDNA should produce the highest possible DNA density achievable with graft polymers.

A panel of ROMP conditions were screened for compatibility with protDNA (Table S2). Ultimately, using 3^rd-generation Grubbs’ catalyst to initiate the polymerization in dichloromethane (DCM) at −20 °C, near-quantitative conversion was achieved. Analyzing the raw mixture containing deprotected DNA graft polymers by polyacrylamide gel electrophoresis (PAGE), a collection of individual bands was observed, whichcan be assigned to specific degrees of polymerization (Figure 2a). Of note, while various ultramild deprotection conditions (e.g. K₂CO₃ in methanol and t-butylamine/methanol/water) can be used, some conditions also cleave the graft polymer side chains from the backbone via base-catalyzed retro-Michael reaction (Figure 2b).⁴⁵ Partial cleavage results in lower-density graft polymers. Treatment with methanolic ammonia at 4 °C for 4 h was identified as capable of achieving full deprotection with minimal cleavage, as evidenced by aqueous gel permeation chromatography (GPC) and matrix-assisted laser-desorption ionization-time of flight mass spectrometry (Figures 2b and S2). The GPC peak for the graft polymer is symmetrical in shape (Figure S3), suggesting that chain-terminating events are much slower compared with propagation, and that protDNA iscompatible with the organoruthenium catalyst. Polymerizations with higher monomer:catalyst feed ratios resulted in higher DPs, as detected by GPC and PAGE (Figure 2c–d), with polydispersity indices (PDIs) in therange of 1.37–1.48. Conversion remained ~90% when a 20:1 monomer:catalyst molar ratio was used, but dropped to 68% at 40:1, and 45% at 80:1 (Table S3). The reduced conversion at higher monomer:catalyst ratios limited the number-average DP (DP_n) to ~45, which may be attributed to the steric hindrance of the bulky and stiff protDNA macromonomer, and/or the loss of solubility as the DP increases. The DNA graft polymers were purified by GPC, and their size/morphology were studied by transmission electron microscopy (TEM) and dynamic light scattering (DLS) (Figure 2e and Table S3). TEM images (negatively stained with 2% uranyl acetate) showed spherical or elliptical morphologies approximately 11–17 nm in diameter. DLS showed increasing number-average hydrodynamic diameters (D_h(n) 10–16 nm) with increasing DP_n, and relatively unchanged ζ potential between −25 and −28 mV, corroborating TEM results (Table S3).

Figure 2.

Characterization of DNA graft polymers, (a) PAGE of the raw reaction mixture after ROMP and deprotection (monomer;catalyst molar ratio: 5:1). (b) Deprotection conditions and their effect on the retention of DNA side chains as indicated by aqueous GPC. (c-d) Aqueous GPC chromatograms and PAGE of DNA graft polymers synthesized with increasing monomer initiator feed ratios, (e) Particle morphology observed under TEM. Samples were negatively stained with 2% uranyl acetate. Scale bar: 100 nm.

To further investigate whether the hybridization and dehybridization properties of DNA were influenced by the bottlebrush-like architecture, we adopted a fluorescence quenching assay, where a quencher (dabcyl)-modifιed strand is hybridized with a fluorescein-labeled DNA graft polymer. Hybridization and thermal melting result in decreases and increases of the fluorescence, respectively, and the rate of change reflects their kinetics. The hybridization kinetics of DNA graft polymers was initially as fast as that of the free DNA but slowed slightly as the reaction proceeded, taking longer to reach complete duplex formation. The trend is more obvious for graft polymers with larger DPs (Figure S4). The reduction in hybridization rate towards the end of the reaction may be attributed to the accumulation of negative charges on the nanostructure, which hinders subsequent reaction. On the other hand, thermal stability of DNA duplexes in the graft polymer form increased by ~3 °C compared with the free duplex, and with sharper melting transitions (Figure S5). Similar phenomena have been found with gold-cored SNAs, which can be explained by the elevated local salt concentration associated with densely arranged DNA.^27,46 The increased salt concentration entropically stabilizes duplex DNA by masking their repulsive negative charges, resulting in higher melting temperatures. When a certain duplex within the nanostructure dehybridizes, salt concentration surrounding the duplex is reduced, making neighboring duplexes more prone to dissociate. These interpretations are in principle also applicable to the DNA graft polymers.

Monodisperse DNA oligomers and their cellular uptake.

By carrying out controlled polymerizations at low monomer:catalyst ratios, discreet oligomer libraries can be generated.⁴⁷ The ROMP reaction of protDNA, therefore, can be a powerful method to produce monodisperse multivalent DNA. Octameric and dodecameric DNA have been meticulously synthesized previously by coupling linear DNA strands to a well-defined multivalent core, followed by several rounds of chromatographic purifications to remove partially coupled products.⁴³ The monodisperse structures are poised to enable accurate interrogation of their interactions with living systems. However, due to the need to use a variety of cores to access a full numeric series of oligomers and the difficulty in synthesizing odd-valency cores, there has not been a conclusive study correlating cellular uptake and DNA density.⁴³

In a single ROMP reaction of protDNA (carried out at 2:1 monomer:catalyst molar ratio), oligomeric DNA containing quantized numbers of strands, including all odd-numbered ones, were simultaneously synthesized, and PAGE was used to isolate monodisperse fractions containing unimers through decamers (limited by gel resolution, Figure 3a–b). On the size-density map for SNAs reported thus far (Figure 3c), these molecular, monodisperse entities occupy a DNA density range (0.1–0.9 pmol/cm²) below that of common SNAs, while the polydisperse DNA graft polymers of higher DPs extend into the density range of prototypical SNAs (1.1–2.3 pmol/cm²). Thus, with identical chemical compositions, these well-defined structures allow us to unambiguously evaluate the effect of the DNA surface density on cellular uptake. Human ovarian carcinoma cells (SKOV-3) were treated with varying concentrations (up to 800 nM of DNA) of fluorescein-labeled, monodisperse oligomers as well as polydisperse graft polymers for 4 h, and were studied by confocal microscopy and flow cytometry. Confocal images showed increasing cell-associated fluorescence (mainly from endosomal compartments) with increasing DP (Figure 3d). Flow cytometry showed that the uptake increased linearly with concentration, suggesting that saturation in the uptake is far from being reached (Figures 3e, S6, and Table S4). The relative rate of cell uptake for the multivalent forms of DNA, as determined by the slope of the linear fit, is 90–170× greater than that of free DNA, and increases with increasing DPs. Strikingly, if one plots these slopes as a function of DNA surface density, another linear relationship comes into view (Figure 3f). The relative rate of uptake depends linearly with minimum variation (R²=0.994) on the DNA surface density in the monodisperse oligomer range, and the linear fit predicts the uptake of polydisperse graft polymers reasonably well. These results suggest that there is no threshold density for a multivalent DNA structure to behave like an SNA with regard to cellular uptake; the uptake instead increases cumulatively with increasing DNA surface density, starting as early as a dimer.

Figure 3.

Characterization of monodisperse DNA oligomers and comparison with polydisperse graft polymers in cellular uptake, (a-b) PAGE and aqueous GPC chromatograms of purified DNA oligomers, (c) Size-density map of previously reported SNAs^{30,43,48–51} and DNA graft polymers (with 2× C12 spacers) of varying DP. DNA surface density refers to the density at the outer surface of the particle as opposed to the footprint (see Experimental Procedures). Particle diameter is defined as the diameter of a spherical particle equivalent in volume to that of the graft polymers, which is estimated assuming all linkers and the polymer backbone are in the fully stretched conformation, (d) Laser-scanning confocal microscopy images of SKOV-3 cancer cells treated with different DNA graft polymers at an equal DNA concentration (800 nM; blue channel: DAPI; green channel: DNA). Imaging settings were kept identical for all samples. Scale bar: 20 μm. (e) Mean fluorescence of SKOV-3 cells treated with DNA graft polymers of varying DP as a function of incubation concentration (measured by flow cytometry; total cell counts: 10,000). The fitted slopes reflect the relative rates of cellular uptake, which are summarized in (f) as a function of DNA surface density. Errors (mean ± σ) in DNA surface density for polydisperse DNA graft polymers were calculated based upon aqueous GPC results.⁵²

DNA/PEG diblock graft copolymers.

Covalent attachment of PEG remains a widely used technique to impart better pharmacokinetic properties to many forms of therapeutics. However, a single linear or slightly branched PEG, even with high molecular weight (40–100 kDa), cannot sufficiently shield oligonucleotides from interaction with serum or cell membrane proteins to provide appropriate biopharmaceutical properties for systemic use.⁵³ Recently, we have developed a brush-architectured PEGylated oligonucleotide, termed pacDNA (polymer-assisted compaction of DNA), which consists of oligonucleotides tethered to the backbone of a PEG graft polymer with many (typically >25) shorter (5–10 kDa) PEG side chains.^53,54 The highly branched achitecture provides a “Goldilocks” PEG density: high enough to reduce protein access but not too high to impair DNA hybridization with a complementary sequence.⁵⁵ Such selectivity greatly diminishes unwanted side effects derived from DNA-protein interactions, and enhances pharmacokinetics.^56–59. The current synthesis of the pacDNA involves sequential copolymerization and post-polymerization modifications to obtain a diblock brush polymer, poly(oxanorbornenyl-azide)-b-(polynorbornene-g-PEG), onto which dibenzocyclooctyne (DBCO)-functionalized DNA strands are coupled via copper-free click chemistry.

With the ability to manipulate protDNA in organic solvents, it is possible to prepare pacDNA in one-pot by sequential copolymerization of norbornenyl PEG and protDNA (Figure 4a). Norbornenyl PEG (10 kDa) was polymerized as the first block using 3^rd-generation Grubbs’ catalyst with a 30:1 monomer:catalyst molar ratio at −20 °C in DCM. Upon completion of the PEG block (monitored by GPC), protDNA monomers were introduced to the reaction mixture at a monomer:catalyst molar ratio of 2:1. Three protDNA monomers with zero to two C12 spacers were used to test the effect of linker length on the incorporation yield of protDNA. After deprotection with methanolic ammonia, aqueous GPC showed ~45%, 70%, and 70% incorporation for the protDNAs with zero, one, and two C12 linkers, respectively, indicating that the steric hindrance of protDNA during ROMP can be reduced by lengthening the spacing between norbornene and the protDNA. All three pacDNAs exhibited similar molecular weight (M_n ~250 kDa by aqueous GPC; Figures 4b and S7), number-average hydrodynamic size (−20.0 nm), ζ potential (~−4.0 mV) (Table S5), and morphology (spherical, as observed with TEM; see Figure 4c).

Figure 4.

Synthesis and characterization of pacDNAs. (a) Synthetic scheme for the one-pot pacDNA. (b) PAGE of the pacDNAs with zero, one, and two C12 spacers. Samples form a non-penetrating band due to the large size and near-neutral charge, (c) Typical TEM image of the pacDNA (2× C12 spacer; stained with 2% uranyl acetate), showing a spherical morphology. Scale bar: 100 nm. (d) Enhanced nuclease stability of pacDNAs compared with free DNA. A shorter linker leads to better nuclease resistance.

Hybridization of the one-pot pacDNA with a dabcyl-labeled complementary strand is immediate and rapid, similar to that of free DNA and prototypical pacDNA (Figure S8). To examine the inhibition of protein access, pre-formed free DNA and pacDNA duplexes were treated with DNase I (an endonuclease that non-specifically cleaves dsDNA). Upon degradation, the fluorophore-quencher pair is separated, leading to an increase of fluorescence signal. It was found that all pacDNAs exhibited enhanced enzymatic stability compared to free DNA (2.7−7.8× longer half-life), and the stability increased with decreasing numbers of the C12 linker, which suggests a depth-effect with respect to the graft polymer backbone (Figure 4d).⁶⁰ These results indicate that the one-pot pacDNA possesses the same hallmark features of traditional pacDNAs obtained via multi-step syntheses, and the proDNA approach can greatly shorten the synthetic route.

Amphiphilic DNA block copolymers.

DNA amphiphiles are yet another difficult-to-access class of structures using homogeneous coupling, especially those with a non-polar hydrophobic segment. To demonstrate the feasibility of using protDNA for the synthesis of DNA amphiphiles, two highly hydrophobic monomers (norbornenes coupled to a C₁₂ aliphatic chain, C₁₂NB, or pyrene, pyNB) were used to form the hydrophobic segment via ROMP (Figure 5a). A monomer:catalyst molar ratio of 10:1 was chosen to balance the amphiphilicity of the final copolymer so that micelles can be formed in an aqueous buffer. Once the monomers were consumed (monitored by TLC), a substantially sub-stoichiometric amount of protDNA monomers was added to the mixture. The large excess of the growing polymer chains ensures that amphiphiles containing only one DNA strand are statistically favored. After removal of the catalyst with ethyl vinyl ether (EVE) and deprotection of the protDNA with methanolic ammonia, unreacted polymers were removed by precipitation in water. Micelles suspended in the supernatant were further injected into an aqueous GPC to remove non-micellar residues (Figure S9). TEM showed that the poly(C₁₂NB)-b-DNA and poly(pyNB)-b-DNA formed spherical micelles with dry-state diameters of 18.1 ± 4.6 nm and 15.4 ± 4.3 nm, respectively (Figure 5b), agreeing with DLS hydrodynamic size measurements (D_h(n) 23.1 ± 7.0 nm and 19.5 ± 6.8 nm, respectively) (Table S5). PAGE of the micelles exhibited band smearing, likely due to the dynamic nature of the assemblies (Figure 5c). For poly(pyNB)-b-DNA, the fluorescence emission spectrum of the micelle exhibited characteristic fluorescein (520 nm) and pyrene emissions (380 and 400 nm) (Figure S10). The micellar aggregation numbers were estimated from negatively stained TEM images, assuming the particle contrast is produced only by the hydrophobic core of the micelles. The C12- and pyrene-based micelles consist of −480 and −380 amphiphiles, respectively, which is comparable to other reported DNA micelles.^36,61,62 It is possible that the actual aggregation number is lower than the estimate, due to incomplete removal of excess homopolymers from the reaction mixture. The micellar nanostructures are expected to exhibit SNA-like properties by virtue of their structural similarity. As a demonstration, we tested the nuclease stability of C12- and pyrene-based micelles by treating them with DNase I. The two micelles exhibited 3.9× and 4.5× longer half-lives than that of free DNA (Figure 5d), respectively, which is in line with typical for SNAs studies thus far.^43,63

Figure 5.

Synthesis and characterization of amphiphilic DNA block copolymers, (a) Synthetic scheme of DNA diblock amphiphiles. (b) Spherical micelles assembled from DNA diblock amphiphiles, as observed by TEM (with 2% uranyl acetate staining). Scale bar: 100 nm. (c) PAGE of the DNA micelles dispersed in PBS. The smearing of the bands is likely due to the dynamic nature of the supramolecular assemblies, (d) Increased nuclease resistance of the micelles compared with free duplexes, a hallmark feature of SNAs.

Conclusion

We demonstrate that, by separating the traditionally combined deprotection and cleavage reactions following solid-phase oligonucleotide synthesis into two steps, fully protected, hydrophobic oligonucleotides can be obtained, which enables further manipulation in organic solvents. Using the protected form of DNA, three distinct classes of DNA nanostructures (DNA graft polymers, pacDNAs, and micellar SNAs) can be obtained via ROMP in a one-pot fashion. These structures represent traditionally difficult DNA-based structures due to either architectural complexity or amphiphilicity. By turning DNA into an organics-soluble molecule, this study opens up promising possibilities to greatly expand the material space accessible to DNA. It is also possible to apply the technique to augment existing DNA-based applications, such as DNA-encoded libraries and DNA microarrays.

Experimental Procedures

Synthesis of the protDNA macromonomer.

Solid-phase oligonucleotide synthesis was conducted following standard protocols. After the completion of the synthesis (1 μmol scale), the CPG was air-dried and suspended in a solution containing 800 μL of acetonitrile and 400 μL of aqueous cleavage buffer (25 mg of TCEP•HCl and 15 mg of NaHCO₃ in i960 μL Nanopure™ water). After shaking overnight at room temperature, 400 μL of Nanopure water™ was added to the solution and the CPG solid support was removed by filtration through a 0.45 μm polytetrafluoroethylene (PTFE) filter. The filtrate was then subjected to reverse-phase HPLC to separate thiolated protDNA monomer (C18 column; mobile phases: acetonitrile and pure water; flow rate: 1.0 mL/min; gradient: constant 50/50 vol% of acetonitrile/water from o to 5 min, then 100% acetonitrile for 30 min). The product was lyophilized and redissolved in a solvent consisting of 500 μL of acetonitrile and 500 μL of water. Next, 6 mg of the norbornenyl maleimide was added to the solution, and the mixture was vigorously shaken at 4 °C for 12 h. Last, the mixture was filtered and subjected to reverse-phase HPLC to separate the norbornenyl protDNA monomer from residues using the same gradient. Notably, the product exhibited multiple peaks in HPLC, which may be attributed to the chiral phosphate centers in the DNA backbone. The final solution was lyophilized to yield the protDNA monomer as a white powder.

Estimation of the DNA surface density.

Density is estimated at the outer periphery of the nanoparticle as opposed to the footprint where DNA meets with the polymer backbone. To establish a model for particle size, it is assumed that all chemical bonds in the polymer backbone and linkers are in the fully stretched conformation and occupy the maximum three-dimensional space. With this assumption, the central polymer backbone can be treated as a cylinder with the radius of R + r and a length of (DP_n-1) × L, where R is the length of the DNA, r is the length of the linker, and L is the length of the repeating unit in the backbone. The DNA surface density is therefore a function of DP_n:

$$\text{DNA}\text{surface}\text{density}{\text{(strand/nm}}^{\text{2}}\text{)}=\frac{D{P}_n}{2\pi (R+r)\left(D{P}_n-1\right)L+4\pi {(R+r)}^2}$$

$$1{\text{ strand/nm }}^2=\left[1mol/\left(6.022\times {10}^{23}\right)\right]/{\left({10}^{-7}\text{cm}\right)}^2=166\text{pmol}/{\text{cm}}^2$$

$$\text{DNA}\text{surface}\text{density}\left(\text{pmol}/{\text{cm}}^2\right)=\frac{D{P}_n}{2\pi (R+r)\left(D{P}_n-1\right)L+4\pi {(R+r)}^2}\times 166$$

where r and L are estimated to be 5.79 nm and 0.60 nm, respectively. The DNA has a 15-bp sequence and thus R = 0.34 nm × 15 = 5.1 nm. The calculated DNA surface densities are provided in Figure 3c.

Data availability.

All relevant data are available from the authors.

Highlights.

• Chemically protected DNA was used for polymerization in the organic phase

• Polymerization of DNA macromonomers gave DNA graft polymers in high yields

• Monodisperse DNA oligomers, diblock grafts, and amphiphiles were also demonstrated

• Cell uptake of multivalent DNA was shown to be linearly correlated with DNA density

Bigger Picture.

Short nucleic acid strands are a useful class of materials for a range of applications spanning medical diagnostics/therapeutics, DNA nanotechnology, and DNA-encoded libraries. Incorporation of them as a functional moiety with traditional polymers, however, proved to be difficult due to their insolubility in organic solvents and/or incompatible chemical properties with certain reactions. Overcoming these limitations, as reported herein, opens the door to a diverse range of novel, architecturally complex materials that are otherwise difficult or impossible to obtain. By expanding the materials space accessible to short nucleic acids, their technological potential are expected to be substantially increased.