Non-Cross-Linking Aggregation of DNA-Carrying Polymer Micelles

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Interface Components: Nanoparticles, Colloids, Emulsions, Surfactants, Proteins, Polymers

Non-Cross-Linking Aggregation of DNA-Carrying Polymer Micelles Triggered by Duplex Formation Zhonglan Tang, Tohru Takarada, and Mizuo Maeda Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.8b01840 • Publication Date (Web): 07 Aug 2018 Downloaded from http://pubs.acs.org on August 9, 2018

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Non-Cross-Linking Aggregation of DNA-Carrying Polymer Micelles Triggered by Duplex Formation Zhonglan Tang,† Tohru Takarada,*,‡ and Mizuo Maeda*,‡

†

National Engineering Research Center for Biomaterials, Sichuan University, 29 Wangjiang Road, Chengdu 610064, China

‡

Bioengineering Laboratory, RIKEN Cluster for Pioneering Research, 2-1 Hirosawa, Wako, Saitama 351-0198, Japan

ABSTRACT

Colloidal behaviors of particles functionalized with biomolecules are generally complicated. This study describes that colloidal behaviors of double-stranded (ds) DNA-carrying polymer micelles are well controlled by altering the molar ratio of single-stranded (ss) DNA moiety in the dsDNA shell. ssDNA-carrying micelles composed of a poly(N-isopropylacrylamide) (PNIPAAm) core surrounded by a dense shell of ssDNAs were prepared through self-assembly of PNIPAAm grafted with ssDNA by incubating its solution above the lower critical solution temperature. Spontaneous, non-cross-linking aggregation of the micelles was triggered by DNA

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duplex formation on the surface. Comparison of the critical coagulation concentration of NaCl among a series of the DNA-carrying micelles revealed the relationship between the helical structure of the surface-bound DNA and the colloidal stability of the micelles. The electrophoretic mobility analysis of the micelles indicated that the duplex formation reduced the structural flexibility of the surface-bound DNA, thereby decreasing the interparticle entropic repulsion. It is also suggested that the augmented rigidity of the surface-bound DNA increases the number of terminal base pairs facing the solvent, which could lead to multiple blunt-end stacking interaction among the micelles. Therefore, small DNA molecules could be considered unique surface-modifiers capable of controlling interactions between the surfaces of materials.

INTRODUCTION Addition of salts to an aqueous dispersion of charged hydrophobic colloids induces their spontaneous aggregation, which can be quantitatively described by the DLVO theory.1 When the surfaces of colloids are modified with biomolecules such as proteins, their colloidal behaviors are highly complicated, mainly due to the fact that intrinsic properties of the surface-bound molecules greatly affect the colloidal stability.2–5 For example, DNA-functionalized particles, including microbeads,6–11 gold nanoparticles,12–16 silver nanoparticles,17–20 and quantum dots,21–24 are readily aggregated by using DNA duplex formation in a cross-linking manner. Both turbidity changes and solution color changes induced by colloidal aggregation allow the naked-eye detection of DNA duplex formation.25–33 Therefore, such particles can be used as a powerful tool in medical diagnosis. Moreover, careful design of base sequence of surface-grafted DNA can


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afford colloidal crystals34–37 and nanoparticle superlattices,38–43 both of which are potentially applicable to various devices. We previously demonstrated that the colloidal stability of DNA-carrying polymer micelles was highly dependent on the DNA structure (Figure 1a).44 When the surface-grafted DNA was single-stranded (ss), the polymer micelles remained stably dispersed even in an aqueous medium of high ionic strength, due to interparticle electrostatic and entropic repulsion between the surface-grafted ssDNAs. However, when complementary ssDNA was added to form a duplex, the resulting double-stranded (ds) DNA-carrying micelles were spontaneously aggregated in a non-cross-linking manner, thereby causing a sharp increase in the turbidity of the dispersion. This colloidal behavior was used to develop a naked-eye method for detecting single-nucleotide polymorphism45 and ATP.46 The DNA-carrying polymer micelles were prepared from thermoresponsive copolymers consisting of poly(N-isopropylacrylamide) (PNIPAAm) and ssDNA.47 The micelles were formed from graft copolymers,44–47 block copolymers,48–51 and starshaped copolymers51 simply by raising the solution temperature above the lower critical solution temperature of the PNIPAAm segment (ca. 32°C). The as-prepared micelles were composed of a hydrophobic PNIPAAm core surrounded by a dense shell of hydrophilic ssDNAs.52 The noncross-linking aggregation of dsDNA-modified particles has been widely observed with various hydrophobic cores, including polystyrene beads,53 gold nanospheres,54 gold nanorods,55 and gold nanotriangles.55 Therefore, the colloidal stability should be determined based on the properties of the DNA shell, because the non-cross-linking aggregation occurs irrespective of the composition, size, and shape of the hydrophobic core. The present study focused on the relationship between the helical structure of the surfacebound DNA and the colloidal stability of the micelles. We used ssDNA-carrying polymer


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micelles that were hybridized with longer complementary ssDNAs to afford singly or doubly nicked dsDNA. This procedure allowed us to readily prepare various DNA-carrying polymer micelles with strictly identical hydrophobic cores and strictly identical DNA grafting densities, which differed only in the helical structure of the surface-grafted DNA. Our conclusions could contribute not only to elucidation of the mechanism for the non-cross-linking aggregation, but also to the rational design of functional materials, including colloidal probes for bioassay and building blocks of self-assembled nanodevices.

EXPERIMENTAL SECTION Materials. All chemical reagents were purchased from Wako Pure Chemical Industries unless otherwise noted. N-isopropylacrylamide (NIPAAm) was obtained from Tokyo Chemical Industry

and

recrystallized

from

a

mixture

of

benzene

and

hexane.

N-

(menthacryloyloxy)succinimide was purchased from Kokusan Chemical Works. Ammonium persulfate (APS) and N,N,N′,N′-tetramethylethylenediamine (TEMED) were obtained from Kanto Chemical and Nacalai Tesque, respectively. All chemically synthesized DNAs were purchased from Espec Oligo Service. Synthesis of Copolymers. The DNA macromonomer was synthesized by the coupling reaction between N-(menthacryloyloxy)succinimide and ssDNA modified with an amino group at the 5´-terminus by following the reported procedure.44 The PNIPAAm grafted with DNA (PNIPAAm-g-DNA) was prepared by radical polymerization between NIPAAm and the DNA macromonomer as reported previously.46 The composition of the copolymers was determined by


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gravimetry and based on the UV absorption at 260 nm of the DNA macromonomer units using the molar extinction coefficient, which was calculated from the nearest-neighbor model. Cloud-Point Measurements. The transmittance curve of an aqueous solution of the copolymer (0.1 mg/mL) in 10 mM Tris–HCl buffer (pH 7.4) containing 500 mM NaCl was monitored at 500 nm at a heating rate of 0.5°C/min using a UV-vis spectrophotometer (Shimadzu UV-2500). The cloud point of the copolymer was determined as the temperature at which the transmittance at 500 nm started to decrease from 100%. Preparation of Micelles. The copolymer (0.1 mg/mL) was dissolved in 10 mM Tris–HCl buffer (pH 7.4) containing NaCl (300–700 mM). The copolymer solution was incubated at 40°C for 30 min, resulting in the spontaneous formation of micelles through the phase transition of the PNIPAAm-g-DNA. Light Scattering Measurements. Both static light scattering (SLS) and dynamic light scattering (DLS) measurements were conducted with DLS-7000 instrument (Otsuka Electronics). The light source was an Ar ion laser (488 nm, 75mW). The dn/dC of copolymer was calculated from the additive equation, dn/dC = wPNIPAAm(dn/dC)PNIPAAm + wDNA(dn/dC)DNA, where w represents the weight fraction.44 The dn/dC of DNA macromonomer, (dn/dC)DNA, was measured by DEM-1021 (Otsuka Electronics) at 488 nm and determined to be 0.247 mL/g at 25°C and 0.226 mL/g at 40°C. The literature values were used for the dn/dC of PNIPAAm, (dn/dC)PNIPAAm. To determine the weight-average molecular weight (Mw) value of the copolymer, the SLS measurements were carried out with an angle ranging from 40° to 150° and a concentration ranging from 0.5 mg/mL to 2.3 mg/mL at 25°C in 10 mM Tris–HCl buffer (pH 7.4). For analysis of the polymer micelles, the SLS measurements were performed at a fixed


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copolymer concentration (0.1 mg/mL) at 40°C in 10 mM Tris–HCl buffer (pH 7.4) containing 300 mM NaCl. To determine the average hydrodynamic diameter of the micelle, the DLS measurements were conducted at 90°. Melting Curve Measurements. The melting curve of free dsDNA in solution was obtained by measuring the change of absorbance at 260 nm with the UV-vis spectrophotometer. The temperature ramp was 0.5°C/min. The melting temperature (Tm) of the duplex was determined from the maximum in the first derivative of the melting curve. The degree of hypochromic effect was estimated by the difference of absorbance (∆Abs) between two values, whose corresponding temperatures gave maximal values in the second derivative of the melting curve. Transmittance Measurements. The complementary ssDNA was added to the dispersion of the ssDNA-carrying micelles. After incubation at 40°C for 30 min, an aqueous solution of salt was further added to the dispersion. After subsequent incubation at 40°C for 30 min, the transmittance at 500 nm was measured with the UV-vis spectrophotometer. For the polymer micelles having singly or doubly nicked DNA duplexes, the 24-nucleotide (nt)-long or 39-ntlong ssDNA was hybridized with one or two short complementary ssDNA fragment(s) and then added into the micellar dispersion at 40°C. The transmittance change of the dispersion was monitored by the same method. Electrophoretic Mobility Measurements. An equimolar mixture of the PNIPAAm grafted with 15-nt-long ssDNA and a PNIPAAm homopolymer with the Mw of 9.5 × 104 was incubated in 10 mM Tris–HCl buffer (pH 7.4) containing 10 mM NaCl at 45°C for 30 min to prepare the polymer micelles. The concentration of surface-grafted ssDNA was 1.3 µM. A 15-nt-long complementary ssDNA (1.3 µM) was added to the micellar dispersion. After incubation at 45°C


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for 30 min, an NaCl solution was further added to the dispersion. After subsequent incubation at 45°C for 30 min, the electrophoretic mobility of the as-prepared polymer micelles was measured with Zetasizer 3000 HS (Malvern). The measurement was repeated at least 25 times to ascertain variability.

RESULTS Preparation of DNA-Carrying Micelles. PNIPAAm grafted with 9-nt-long ssDNA (PNIPAAm-g-DNA) was synthesized by radical copolymerization according to the reported method (Figure 1b).44 The Mw value was determined to be 4.0 × 105 by the SLS measurements. The fraction of DNA macromonomer units in the copolymer was determined to be 0.36 mol% by UV-vis absorption spectroscopy. Based on these data, the average DNA graft number of the copolymer was calculated to be 11. The DNA concentration in an aqueous solution of the copolymer (0.1 mg/mL) was 3.0 µM. The cloud-point temperature of PNIPAAm-g-DNA in 10 mM Tris–HCl buffer (pH 7.4) containing 300 mM NaCl was determined to be 30°C, which was higher than that of PNIPAAm (26°C), due to the greater hydrophilicity. When the same buffer solution of the copolymer was incubated at 40°C, the copolymers self-assembled to form a polymer micelle (PM1) that was composed of a PNIPAAm core surrounded by a dense shell of the 9-nt-long ssDNAs (Figure 1c).52 The hydrodynamic diameter and the molecular weight of PM1 were determined to be 36 nm and 6.6 × 106 from the DLS and SLS measurements, respectively. Thus, the average number of copolymers involved in a single PM1 was calculated to be 17, which corresponded to an average of 187 ssDNAs per particle in a single PM1 (one DNA strand/22 nm2).


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Aggregation of Micelles Triggered by Duplex Formation. Our previous study demonstrated that formation of the fully matched DNA duplex on the surface of the micelles caused colloidal destabilization (Figure 1a).44 Interestingly, the melting curve of the duplex between the surfacegrafted ssDNA and its complementary ssDNA was undistinguishable from the melting curve of the duplex between the unmodified ssDNA with the identical base sequence and its complementary ssDNA; namely, the same Tm values and a similar degree of hypochromic effect (∆Abs) were obtained.46 The results indicated that the surface-grafted ssDNA had the same ability as that of free ssDNA in solution to engage in the hybridization event. In the present study, we investigated in further detail the relationship between the colloidal stability of the DNA-carrying polymer micelles and the helical structure of the surface-bound DNA. Initially, all surface-grafted ssDNAs of PM1 were hybridized to the complementary ssDNAs to afford PM2 (Figure 2a), which was constructed by adding an equimolar complementary ssDNA to the surface-grafted ssDNA on PM1. The colloidal stability of PM1 and PM2 was evaluated by the critical coagulation concentration (CCC) of the supporting electrolyte (NaCl). The CCC was determined by measuring the transmittance at 500 nm of the micellar dispersion at 40°C. The measurement temperature was selected to be above the cloud-point temperature of PNIPAAm-gDNA (30°C) but below the Tm value of the surface-bound dsDNA (57°C). As shown in Figure 2b, the PM1 micelles remained dispersed even at the highest NaCl concentration tested; the CCC value was above 1300 mM NaCl. In sharp contrast, the CCC value of PM2 was determined to be 370 mM NaCl. Moreover, when the amount of complementary ssDNA added to the PM1 dispersion was increased from 0.2 equivalents (eq) to 0.8 eq, compared to the surface-grafted ssDNA number, the resultant micelles were gradually destabilized. Figure 2c shows the relationship between the amount of the complementary ssDNA and the CCC. The CCC value


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was gradually decreased from 900 mM to 370 mM NaCl, when the amount of the complementary ssDNA was increased from 0.2 eq to 1.0 eq. Noticeably, no change of CCC was observed when the DNA ratio was further increased from 1.0 eq to 2.0 eq. These results imply that almost all of the ssDNA segments in the copolymer existed on the surface of the hydrophobic core of PM1, consistent with a previous study in which the same Tm and ∆Abs values were observed between the surface-bound dsDNA and the free dsDNA in solution. In addition, the non-cross-linking aggregation of the micelles took place immediately after the complementary ssDNA was added to the dispersion (Figure 2d). When the amount of the complementary ssDNA was more than 0.5 eq in the presence of 500 mM NaCl, the transmittance at 500 nm rapidly decreased below 80% in 3 min, which was clearly recognized as turbidity by the naked eye. Next, we compared the colloidal stability of PM1 and PM2 by using various supporting electrolytes. After both micelles were prepared in 10 mM Tris–HCl buffer (pH 7.4) containing 300 mM NaCl, various salts were added to the micellar dispersion to induce the non-crosslinking aggregation. Initially, we focused on the effect of the cation. The colloidal stability of PM1 and PM2 in the presence of NaCl (Figure 3a) was almost identical to the stability of PM1 and PM2 with KCl (Figure 3b), respectively. Similarly, the stability profiles of PM1 and PM2 in the presence of MgCl2 (Figure 3c) were the almost same as those with CaCl2 (Figure 3d). We observed a remarkable difference of CCC between the monovalent and divalent metal ions. As for PM 2, the CCC value of the monovalent cation was approximately 20 times as high as that of the divalent cation. The divalent cations were permitted to trigger the non-cross-linking aggregation of the PM1 micelles within the concentration range tested. The CCC was highly dependent on an ionic valence number of metal, but almost independent of the type of cation


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with the same valence number. We then examined the effect of the counter anion. As shown in Figure 3e, the CCC of NaCl and NaNO3 for PM2 were almost identical, suggesting that the CCC was also independent of the type of anion with the same valence number. We also tested the effect of single-base mismatch in the surface-bound dsDNA on the colloidal stability. By adding the single-base-substituted complementary ssDNA to the PM1 dispersion, we prepared three kinds of micelle: PM2a, PM2c, and PM2t (Figure 4a). Both the PM2a and PM 2c micelles were stably dispersed even at high ionic strength (Figure 4b), because the singlebase-mismatched DNA duplex was not allowed to form at 40°C; the Tm values for the surfacebound dsDNA of PM2a and PM2c were much lower than 40°C (Figure 4a). This means that the actual state of the PM2a and PM2c surfaces was identical to that of PM1. Unexpectedly, we observed that the PM2t micelles were spontaneously aggregated at high NaCl concentration above 1000 mM. One possible explanation is that surface-bound dsDNAs containing two bulge moieties were partially formed at elevated NaCl concentrations, thereby inducing the non-crosslinking aggregation. PNIPAAm grafted with 15-nt-long ssDNA was also synthesized using the same method.44 The Mw, the fraction of the DNA macromonomer, and the average DNA graft number were 1.1 × 105, 0.30 mol%, and 4, respectively. The DNA concentration of the copolymer (0.1 mg/mL) was 2.5 µM. The cloud-point temperature in 10 mM Tris–HCl buffer (pH 7.4) containing 200 mM NaCl was determined to be 31°C. When the same buffer solution of the copolymer was incubated at 40°C, the copolymers self-assembled to form PM1´ with a hydrodynamic diameter of 41 nm. The addition of the complementary ssDNA and its single-base-substituted one to the PM1´dispersion produced PM2´ and PM2´t, respectively (Figure 5a). In sharp contrast with PM2 and PM2t (Figure 4b), both PM2´ and PM2´t underwent the non-cross-linking aggregation at a


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low NaCl concentration; the CCC value of PM2´t was almost the same as that of PM2´ (Figure 5b). This is because the Tm value for the surface-grafted dsDNA containing a single G-T mismatch was sufficiently higher than 40°C. This result strongly suggested that a single-base mismatch involved in the middle of surface-bound long dsDNA had a negligibly small effect on the micellar stability. The duplex formation on the micellar surface can trigger the colloidal destabilization of micelles. Colloidal Destabilization by Augmented Rigidity of DNA. To gain insight into changes of the colloidal property induced by duplex formation on the micellar surface, we evaluated the electrophoretic mobility of the DNA-carrying polymer micelles. We made an equimolar mixture of the PNIPAAm grafted with 15-nt-long ssDNA and a PNIPAAm homopolymer with the Mw of 9.5 × 104. We prepared PM1´´ by incubating an aqueous solution of the polymers in 10 mM Tris–HCl buffer (pH 7.4) containing 10 mM NaCl at 45°C (Figure 6a). The hydrodynamic diameter, the Mw value, and the DNA grafting number per particle were 174 nm, 2.5 × 108, and 3000, respectively. Addition of the complementary ssDNA at 1.0 eq to the PM1´´ dispersion afforded PM2´´. The stabilities of PM1´´and PM2´´ are shown in Figure 6b. The PM1´´ micelles remained stably dispersed at the highest NaCl concentrations tested, whereas the PM2´´ micelles were spontaneously aggregated with the CCC value of 200 mM NaCl. We then measured the electrophoretic mobility of PM1´´and PM2´´ under the conditions by which both micelles kept dispersing (Figure 6c). When the ionic strength was increased, the mobility of both micelles approached zero, mainly due to neutralization of the negative surface charge by Na+. The absolute value for the mobility of PM2´´ was smaller than that of PM1´´ at high ionic strength (>0.05 M). To estimate the difference in surface properties between PM1´´ and PM2´´, we analyzed the mobility data with the soft particle theory developed by


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Ohshima.56,57 The mobility of particles whose surface was covered by an ion-penetrable layer was expressed as the following equation:

µ=

εrε 0 Ψ 0 κm + ΨDON λ zeN + 2 η 1 κm + 1 λ ηλ ,

(1)

with

ΨDON

1/ 2 2   kT  zN  zN  ln = +   + 1  ve  2vn  2vn     ,

1/ 2 1/ 2 2    2vn   zN  2    kT   zN  zN  1 −  ln Ψ0 = +   + 1  +  + 1   ve   2vn  2vn    zN   2vn       ,  

  zN  2  κ m = κ 1 +      2vn  

 2ne 2 v 2   κ =  kT ε ε  r 0 

(2)

(3)

1/ 4

,

(4)

1/ 2

.

(5)

Here, µ is the electrophoretic mobility of the polymer micelle covered with a DNA shell, in which ionized groups of valence z are uniformly distributed at a number density of N (m–3). n (m–3) is the bulk concentration of Na+ with valence v (= 1) in the disperse medium. η is the viscosity, εris the relative permittivity of the solution, ε0 is the permittivity of a vacuum, ΨDON


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is the Donnan potential of the micellar surface layer, Ψ0 is the potential at the boundary between the micellar surface layer and the surrounding solution, κ is the Debye-Hückel parameter of the surrounding solution, k is the Boltzmann constant, and T is the measurement temperature. We call Ψ0 the surface potential of the DNA-carrying micelle. κm can be interpreted as the DebyeHückel parameter of the DNA layer. The reciprocal of λ (1/λ) has the dimension of length (nm) and can be considered to be the softness parameter. Equation 1 directly relates the measured values of electrophoretic mobility to the charge density (zN) and the softness (1/λ), both of which were determined by a curve-fitting procedure. The results are summarized in Figure 6a. The duplex formation of the surface-grafted DNA caused an increase of the absolute value of zN and a decrease of 1/λ. These changes of the charge density and the softness related to the surface property were in line with the physicochemical property changes at the molecular level; namely, the duplex formation of free DNA in solution generally increases the charge density and decreases the structural flexibility of the DNA strand. In addition, the absolute value of zN for PM2´´ was larger than that for PM1´´, whereas the absolute value of the mobility for PM2´´ was lower than that for PM1´´ at high ionic strength. This result suggests that a decrease in mobility of the polymer micelle arose mainly from a decrease in softness of the DNA shell. Therefore, we hypothesized that the lower colloidal stability of PM2´´ compared to the stability of PM1´´ was due to augmented rigidity of the surface-bound DNA strands caused by the duplex formation.

Aggregation of Micelles with Singly Nicked Duplexes. To examine the working hypothesis described above, we further investigated the DNA-structure dependency of the micellar stability. Specifically, a singly nicked 24-base-pair (bp)-long dsDNA was constructed on the micellar


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surface (Figure 7a). The stability data of each micelle are shown in Figure 7b. The CCC value for PM3 obtained by hybridizing the 24-nt-long complementary ssDNA to the surface-grafted ssDNA of PM1 was determined to be >1300 mM NaCl; the micelle had a protruding long ssDNA moiety at the solution-facing terminus, which caused the micelle to behave like a ssDNA-carrying micelle (PM1 or PM1´). By sharp contrast, when the 15-nt-long ssDNA complementary to the protruding ssDNA moiety was further added to form a singly nicked DNA duplex on the surface, the resultant micelle (PM4) was significantly destabilized to exhibit a CCC value of 400 mM NaCl. Next, we introduced a single-base mismatch into the 15-bp-long dsDNA region highlighted by a rectangular frame in Figure 7a. The CCC value for PM5, which was constructed by adding a terminal-base-substituted 15-nt-long ssDNA to the PM3 dispersion, was almost identical to the CCC value for PM4. This is because the thermodynamic stability of the dsDNA that had a terminal T-G mismatch was almost equal to the stability of the fully matched counterpart of PM4 at 40°C (Figure 7a–7c). However, the CCC value for PM6 having a single-base C-A mismatch at the third position from the nick site was increased to >1300 mM NaCl. The melting curve for the free dsDNA whose sequence was identical to that of PM6 was shifted towards a lower temperature (Tm = 54°C) and exhibited half the hypochromic effect (∆Abs = 0.08) of the fully matched counterpart (Figure 7a and 7c). This result suggests that the surface-bound dsDNA of PM6 was partially dehybridized around the mismatch site. Similar results were obtained for PM7, which had a C-A mismatch at the sixth position from the nick site. In contrast, the CCC value for PM8 having a C-A mismatch at the eighth position from the nick site was 400 mM NaCl, which was identical to that for PM4 having the fully matched dsDNA. The formation of the DNA duplex involving the C-A mismatch was thermodynamically allowed at 40°C, which


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was supported by the fact that the degree of the hypochromic effect observed in the melting curve for the dsDNA with the single-base mismatch (∆Abs = 0.15) was almost the same as that for the fully matched counterpart (∆Abs = 0.16). These results are consistent with the fact that PM2´t exhibited colloidal stability as low as that of PM2´ (Figure 5b). Overall, the data shown in Figure 7 strongly suggest that the colloidal stability of dsDNA-carrying polymer micelles is increased when the short ssDNA region is introduced into the middle of the surface-grafted dsDNA.

Aggregation of Micelles with Doubly Nicked Duplexes. The validity of the insight described above was directly assessed by using the micelles having a surface-bound dsDNA with two nick sites (Figure 8). For this purpose, we prepared DNA-carrying polymer micelles composed of a larger hydrophobic PNIPAAm core surrounded by a longer DNA duplex. Specifically, polymer micelles with a hydrodynamic diameter of 80 nm were prepared by incubating an aqueous solution of PNIPAAm-g-DNA involving 0.28 mol% DNA macromonomer in 10 mM Tris–HCl buffer in the presence of 700 mM NaCl at 40°C. First, we hybridized a 39-nt-long ssDNA to the as-prepared larger micelle to make a 30-ntlong ssDNA moiety protruding from the surface. As expected, the micelles were dispersed at high ionic strength to exhibit a large CCC value (>1300 mM NaCl), because the long ssDNA moiety endowed the micelle with high stability. By contrast, an addition of a 30-nt-long ssDNA complementary to the protruding ssDNA moiety into the micellar dispersion produced the less stable PM9 micelle, which had a singly nicked 39-bp-long dsDNA (Figure 8a); the CCC value was 700 mM NaCl (Figure 8b). Next, we trimmed the long complementary ssDNA from 30 nt to 24 nt to obtain the micelles PM10–13, which contained an internal ssDNA region ranging from 1


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nt to 6 nt (Figure 8a). Concurrently, the micelles were gradually stabilized to show the CCC value from 700 mM to 950 mM NaCl (Figure 8b). Further shortening of the complementary ssDNA from 22 nt to 18 nt produced highly stable micelles PM14–16, all of which exhibited the high CCC value (>1300 mM NaCl). These results indicated that the colloidal stabilization of the dsDNA-carrying polymer micelles was caused by introducing the ssDNA moiety to the middle region of the surface-grafted dsDNA. To confirm this hypothesis, the internal ssDNA region within the surface-grafted dsDNA of PM14–16 was hybridized to a short ssDNA fragment (8, 10, and 12 nt) to construct a doubly nicked surface-grafted dsDNA of PM17–19 (Figure 9a). Interestingly, all the CCC values for PM17–19 were decreased from >1300 mM to 700–750 mM NaCl (Figure 9b). Our hypothesis was further supported by the following experimental results. We introduced a single-base substitution (T to A and G to A) to the middle of the 39-nt-long ssDNA on the surface of PM17 and PM18 to construct PM20 and PM21, respectively (Figure 10a). As shown in Figure 10b and 10c, the PM20 and PM21 micelles were stably dispersed because the 8-nt-long and 10-nt-long ssDNA fragments were not permitted to bind to the 39-nt-long ssDNA on the surface at 40°C owing to the mismatch. The Tm values for the corresponding dsDNA regions were much lower than 40°C (Figure 10a). On the other hand, the similar introduction of the single-base mismatch to PM19 produced PM22 (Figure 10a), which was moderately stabilized to exhibit a CCC value of 800 mM NaCl (Figure 10d). This is because the hybridization between the 12-nt-long ssDNA and the single-base-substituted 39-nt-long ssDNA (Tm = 46°C) was partially permitted to proceed at 40°C. This result is also in line with the colloidal destabilization of PM2´t (Figure 5b).


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Since the 39-nt-long ssDNA would be used as an analyte, the current colloidal system would be used to discriminate a single-base difference in genetic samples with the naked eye; namely, the polymer micelles carrying the doubly nicked dsDNA could be applied to develop instrumentfree gene diagnostics.45 The base sequence of the short ssDNA fragment, which is essentially important to detect single-base substitution, should be rationally designed such that the Tm value for the fully matched duplex between the short ssDNA fragment and the wild-type ssDNA analyte is higher than the experimental temperature (40°C), while the Tm value for the singlebase-mismatched duplex between the short ssDNA fragment and the point-mutated ssDNA analyte is lower than 40°C. In the current system, the set of PM18 and PM21 satisfied these requirements.

DISCUSSION The CCC values of the dsDNA-carrying polymer micelles were independent of the type of cation and anion (Figure 3). Provided that the duplex formation of the surface-bound DNA was thermodynamically forbidden due to a single-base mismatch, the non-cross-linking aggregation was severely inhibited (Figure 4). When the surface-bound dsDNA containing the single-base mismatch was stably formed, the non-cross-linking aggregation took place (Figure 5b, 7b, and 10d). These data indicate that duplex formation triggers spontaneous aggregation of DNAcarrying micelles. The analysis of the electrophoretic mobility of the micelles revealed that the duplex formation of the surface-grafted ssDNA reduced the softness of the DNA shell surrounding the PNIPAAm core, thereby causing the electrophoretic mobility to approach zero (Figure 6c). This means that stiffness of the surface-bound DNA at the molecular level accounts


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for the surface property of the polymer micelle at the mesoscopic level; namely, the augmented rigidity of the surface-bound DNA reduces the interparticle entropic repulsion, which confers thermodynamic advantages on the aggregating process. Our recent study suggested that multiple blunt-end stacking interaction could be responsible for the non-cross-linking aggregation of dsDNA-carrying nanoparticles. The force–distance curve analyses between dsDNA layers using a colloid probe-AFM technique revealed that the outermost fully matched dsDNA layers attract each other at a high NaCl concentration.58 In the current study, the polymer micelles having a smaller number of dsDNAs on the surface were more stabilized (Figure 2b), probably due to a reduction of the blunt-end stacking interaction along with augmentation of the interparticle entropic repulsion (Figure 11a). A smaller number of terminal base pairs facing the solution imposes substantial entropic penalties to multiple bluntend stacking interaction. Figure 7b (PM 6 and 7) and Figure 8 (PM10–16) demonstrate that a longer ssDNA moiety in the middle of surface-bound dsDNA provides higher colloidal stability of the polymer micelle (Figure 11b). For these micelles, the total number of terminal base pairs potentially engaging in the blunt-end stacking interaction is unchanged by altering the number and length of the ssDNA moiety. However, the greater flexibility of the surface-bound dsDNA achieved by inserting the ssDNA moiety could lead to a decrease in the number of the terminal base pairs that preferably face the solution, thereby generating less blunt-end stacking interaction. In conclusion, the colloidal stability of dsDNA-carrying micelles was determined by the molar ratio of the ssDNA moiety within the dsDNA shell surrounding the hydrophobic core. The hypothesis that a greater number of terminal base pairs facing the solution results in more efficient non-cross-linking aggregation of the micelles can explain all experimental results obtain


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in the current study. The colloidal behaviors of particles functionalized with various biomolecules are generally complicated; however, the behaviors of the dsDNA-carrying particles are relatively simple and thus predictable by using the molar ratio of the ssDNA moiety in the dsDNA shell under given conditions. In other words, small DNA molecules could be considered unique surface-modifiers capable of controlling interactions emerging between the surfaces of materials. Applications of the DNA interfaces to bioassays, medical diagnosis, and nanomaterials assembly are highly expected.

AUTHOR INFORMATION Corresponding Authors *E-mail: [email protected] (T.T.). *E-mail: [email protected] (M.M.). ORCID Tohru Takarada: 0000-0001-6906-5812 Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT We are grateful to Mr. Yoshikuni Sato (Kyushu University) and Ms. Emi Imaizumi (Saitama Institute of Technology) for their technical support. This study was financially supported in part


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by JSPS KAKENHI Grant Number JP25220204. T.T. and M.M. are grateful for a grant for “Fundamental principles underlying the hierarchy of matter: A comprehensive experimental study” provided by RIKEN.

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Figure 1. (a) Schematic representation of colloidal behaviors of DNA-carrying polymer micelles in a medium of high ionic strength. (b) Chemical structure of a graft copolymer made of poly(Nisopropylacrylamide) and single-stranded DNA (PNIPAAm-g-DNA) used in this study. (c) Schematic representation of a DNA-carrying polymer micelle formed via self-assembly of PNIPAAm-g-DNA.


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Figure 2. (a) Schematic representation of the polymer micelle carrying 9-nt-long ssDNA or 9bp-long dsDNA. (b) Transmittance at 500 nm of the dispersion of PM1 with various amounts of the complementary ssDNA as a function of the NaCl concentration at 40ºC. The concentration of complementary ssDNA was described as a molar ratio to the surface-grafted ssDNA of PM1 (3.0 µM). Lines are drawn as a guide. The same applies hereafter unless otherwise noted. (c) Critical coagulation concentrations of NaCl for PM1 as a function of the molar ratio of the complementary ssDNA. (d) Rapid aggregation of PM1 with various amounts of the complementary ssDNA in 10 mM Tris–HCl buffer (pH 7.4) containing 500 mM NaCl at 40ºC.


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Figure 3. Transmittance at 500 nm of the dispersion of PM1 and PM2 as a function of the concentration of (a) NaCl, (b) KCl, (c) MgCl2, (d) CaCl2, and (e) NaNO3 in 10 mM Tris–HCl buffer (pH 7.4) at 40ºC. Prior to adding each salt, all colloidal systems contained 300 mM NaCl so that PM1 was formed from the graft copolymers.


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Figure 4. (a) Schematic representation of the polymer micelle carrying 9-bp-long dsDNA with or without a single-base mismatch. The Tm values for the free dsDNA were measured in 10 mM Tris–HCl buffer (pH 7.4) containing 1 M NaCl. The concentration of each DNA was 3.0 µM. (b) Transmittance at 500 nm of the dispersion of PM2, PM2a, PM2c, and PM2t as a function of the NaCl concentration at 40ºC.


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Figure 5. (a) Schematic representation of the polymer micelles carrying 15-nt-long ssDNA and 15-bp-long dsDNA with or without a single-base mismatch. The Tm values for the free dsDNA were measured in 10 mM Tris–HCl buffer (pH 7.4) containing 500 mM NaCl. The concentration of each DNA was 2.4 µM. (b) Transmittance at 500 nm of the dispersion of PM1´, PM2´, and PM2´t as a function of the NaCl concentration at 40°C.


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Figure 6. (a) Schematic representation of the polymer micelles used for the electrophoretic mobility analysis. Parameters determined by the best fitting of eq 1 are also shown. (b) Transmittance at 500 nm of the dispersion of PM1´´ and PM2´´ as a function of the NaCl concentration at 45ºC. The NaCl concentration range used for the electrophoresis is highlighted with a gray background. (c) Ionic-strength dependence of the electrophoretic mobility of the micelles at 45ºC. Lines represent the best fitting of eq 1 using the parameters shown in (a).


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Figure 7. (a) Schematic representation of the polymer micelles carrying singly nicked 24-bplong dsDNA with or without single-base mismatch. Measured Tm values and absorbance changes (∆Abs) induced by the hyperchromic effect are given for the 15-bp-long dsDNA moiety highlighted by a rectangular frame for PM4. These values for the corresponding single-basemismatched dsDNA moieties are also provided for PM5−PM8. (b) Transmittance at 500 nm of the dispersion of PM3–PM8 as a function of the NaCl concentration at 40°C. (c) Melting curves at 260 nm for the 15-bp-long dsDNA moiety highlighted by a rectangular frame for PM4 and the corresponding single-base-mismatched dsDNA moieties for PM5–PM8 in 10 mM Tris–HCl


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buffer (pH 7.4) containing 1 M NaCl. The free dsDNA version was used for the measurement; the concentration of each DNA was 3.0 µM.


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Figure 8. (a) Schematic representation of the polymer micelles carrying 39-bp-long dsDNA with or without an ssDNA moiety. (b) Transmittance at 500 nm of the dispersion of PM9–PM16 as a function of the NaCl concentration at 40°C.


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Figure 9. (a) Schematic representation of the polymer micelles carrying 39-bp-long dsDNA with or without an ssDNA moiety. PM17, PM18, and PM19 were prepared from PM14, PM15, and PM16, respectively, by using the short complementary ssDNA. (b) Transmittance at 500 nm of the dispersion of PM17, PM18, and PM19 as a function of the NaCl concentration at 40°C.


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Figure 10. (a) Schematic representation of the polymer micelles carrying doubly nicked 39-bplong dsDNA with or without a single-base mismatch. Calculated Tm values for the dsDNA


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moiety highlighted by a rectangular frame are given under the same conditions as for the colloidal experiments, except for [NaCl] = 1200 mM. (b–d) Transmittance at 500 nm of the dispersion of PM17 and PM20 (b), PM 18 and PM 21 (c), or PM19 and PM22 (d) as a function of the NaCl concentration at 40°C.


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Figure 11. Schematic representation of the surface of DNA-carrying polymer micelles at high ionic strength. (a) Strong attraction between the short-dsDNA-carrying polymer micelles for non-cross-linking aggregation (upper) and weak attraction between the short-dsDNA-carrying polymer micelles having unhybridized surface-grafted ssDNA (lower). (b) Strong attraction between the long-dsDNA-carrying polymer micelles for non-crosslinking aggregation (upper) and weak attraction between the long-dsDNA-carrying polymer micelles containing an ssDNA region in the middle of the surface-bound DNA (lower).


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Non-Cross-Linking Aggregation of DNA-Carrying Polymer Micelles

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