Modulating the Bond Strength of DNA ... - ACS Publications

Dec 23, 2015 - ABSTRACT: A method is introduced for modulating the bond strength in DNA−programmable nanoparticle (NP) superlattice crystals. This m...
4 downloads 16 Views 4MB Size
Download PDF
Modulating the Bond Strength of DNA− Nanoparticle Superlattices Soyoung E. Seo,†,∥,‡ Mary X. Wang,∥,§,‡ Chad M. Shade,†,∥ Jessica L. Rouge,†,∥ Keith A. Brown,∥ and Chad A. Mirkin*,†,∥,§ †

Department of Chemistry, ∥International Institute for Nanotechnology, §Department of Chemical and Biological Engineering, Northwestern University, Evanston, Illinois 60208, United States S Supporting Information *

ABSTRACT: A method is introduced for modulating the bond strength in DNA−programmable nanoparticle (NP) superlattice crystals. This method utilizes noncovalent interactions between a family of [Ru(dipyrido[2,3-a:3′,2′c]phenazine)(N−N)2]2+-based small molecule intercalators and DNA duplexes to postsynthetically modify DNA−NP superlattices. This dramatically increases the strength of the DNA bonds that hold the nanoparticles together, thereby making the superlattices more resistant to thermal degradation. In this work, we systematically investigate the relationship between the structure of the intercalator and its binding affinity for DNA duplexes and determine how this translates to the increased thermal stability of the intercalated superlattices. We find that intercalator charge and steric profile serve as handles that give us a wide range of tunability and control over DNA−NP bond strength, with the resulting crystal lattices retaining their structure at temperatures more than 50 °C above what nonintercalated structures can withstand. This allows us to subject DNA−NP superlattice crystals to conditions under which they would normally melt, enabling the construction of a core−shell (gold NP-quantum dot NP) superlattice crystal. KEYWORDS: crystallization, DNA intercalator, DNA, nanoparticle, self-assembly with over 24 different crystal symmetries4−8 consisting of various nanoparticle compositions9,10 and interparticle distances.11 In addition, this technique, in certain cases, allows one to control three-dimensional crystal habit (e.g., rhombic dodecahedra single crystals are formed from body centered cubic (bcc) superlattices).12 This method has been used to create a variety of functional materials, including catalysts13 and optical devices.14 While a great deal of work has been done in controlling architecture and building block identity, relatively little work has been conducted on deliberately tuning bond strength. Changing the interparticle bond strength has been explored via

T

he ability to build complex, hierarchical materials with precise control over the identity and placement of each component has long been a goal of materials science.1 Nature provides many elegant examples of self-assembled functional materials with multiple levels of organization. Many synthetic routes for nanoparticle (NP) crystallization have been explored, yet the engineering of versatile building blocks that rationally assemble into complex macroscopic materials remains a challenge. DNA is an attractive candidate for nanoparticle assembly due to its length, sequence, and chemical programmability.2 This allows for the crystallization of DNA−NP conjugates into extended three-dimensional superlattices dictated by wellestablished design rules.3,4 Despite being constructed from semiflexible polymers (i.e., oligonucleotides), these materials have extraordinarily well-defined symmetry. To date, researchers have utilized DNA-mediated NP assembly to make lattices © XXXX American Chemical Society

Received: November 10, 2015 Accepted: December 17, 2015

A

DOI: 10.1021/acsnano.5b07103 ACS Nano XXXX, XXX, XXX−XXX

Article

www.acsnano.org

Article

ACS Nano

Figure 1. RuII complexes with extended ancillary ligands varying in steric profile and charge.

the planar dppz ligand, whereupon π−π stacking interactions between this ligand and the base pairs of the DNA helix enables “molecular light switch” activity.23 Recently, the effect of systematically changing the steric profile of the ancillary ligands and overall charge of this “parent molecule” (Complex 1, Figure 1) on its fluorescence signal in the presence of dsDNA and ssDNA was reported.24 It was hypothesized that these ancillary ligand modifications could be used as a handle to tune DNA−NP superlattice bond strength, reflected in the characteristic melting temperature (Tm) of the material.

changing the identity of the nucleic acid (RNA vs DNA),15 varying counterion concentration, DNA sequence and number of oligonucleotides per particle,16 and introducing ethidium bromide, a well-known DNA intercalator.17 Of these methods, intercalation is the only method that allows bond strength to be altered after superlattice assembly has occurred. This is advantageous since nanoparticle crystallization cannot occur when DNA hybridization strength is above a certain limit.16,18,19 The complex chemical structure of the DNA helix facilitates many types of highly specific molecular interactions. Small molecule DNA intercalators insert between DNA bases and can have an effect on both the thermodynamic and structural properties of the duplex.20,21 The degree to which intercalators affect the DNA duplex is dependent upon their molecular structure, charge, and composition.22 In principle, a family of structurally different intercalator molecules could provide a way to rationally tailor oligonucleotide bond strength in DNA−NP superlattices. Herein, we evaluate the use of a family of structurally different intercalator molecules to noncovalently modulate the bond strength of oligonucleotides in a DNA−NP superlattice. We investigate the effect of interalator charge and steric profile on its binding affinity for double strand deoxyribose nucleic acid (dsDNA) and how this translates to the thermal stability and structure of intercalated bcc DNA−NP superlattices. As a proof of concept, we then selected the most suitable intercalator molecule to facilitate the creation of a core−shell material that has not previously been demonstrated with DNA−NP assembly techniques. This material consists of a DNA−gold NP (AuNP) single crystal core coated via stepwise growth cycles in a shell of DNA−quantum dot NPs (QdNPs). The general architecture of the intercalator molecule is based on work by Barton and co-workers, which showed that [Ru(bpy)2(dppz)]2+ (dppz = dipyrido[3,2-a:2′,3′-c]phenazine, bpy = 2,2′-bipyridine) effectively intercalates into dsDNA via

RESULTS AND DISCUSSION Thermal Stabilization of the DNA−NP Superlattice “Bond”. The potential of using intercalators to thermally stabilize the DNA bonds in a superlattice was studied by performing melt analyses on superlattices incubated with RuII complexes 1−6. Using a family of molecules that possess the same intercalating ligand but different ancillary ligands allowed the relationship between ancillary ligand structure and DNA bond stabilization to be determined. Superlattices were assembled by coating 15 nm diameter AuNPs with a dense monolayer of diisopropylthiol-functionalized oligonucleotides (398 ± 10 strands per particle). These oligonucleotide-coated NPs were then hybridized with 350 equiv of DNA linker strand per particle, resulting in 18 base pair (bp) duplexed regions with unpaired 7 base “sticky ends” at the termini. These particles (25 nM) were then mixed with an equimolar amount of a second species of oligonucleotidecoated AuNP presenting a complementary sticky end sequence. Upon hybridization of the sticky ends of neighboring particles, the conjugates settled out of solution as an amorphous aggregate. To remove excess unhybridized DNA linker strands, the aggregate was washed with 0.5 M NaCl followed by annealing for 30 min at a temperature slightly below Tm, yielding bcc superlattices. These nanoparticle superlattices were then incubated with the racemic RuII intercalators 1−6 (12.1 B

DOI: 10.1021/acsnano.5b07103 ACS Nano XXXX, XXX, XXX−XXX

Article

ACS Nano Scheme 1. RuII Intercalation into DNA−Nanoparticle Superlatticea

a DNA−functionalized gold nanoparticles are assembled with linkers and annealed into a crystalline structure. RuII complex is added and the mixture is stored overnight to maximize intercalation.

Figure 2. DNA−NP superlattice melting temperatures increase after intercalation and correlate to binding affinity of different intercalators. (a) Superlattice samples (50 nM NP total) incubated in 12.1 μM of different RuII complexes exhibit unique shifts in melting transition. (b) Superlattice melting temperatures after intercalation with complexes 1, 3, and 6 as a function of degree of association. (c) The binding affinity of RuII complexes for one dsDNA bp plotted as a function of ΔTm of DNA−programmable superlattices.

μM) in 0.5 M NaCl solution overnight, allowing ample time for intercalation to occur (Scheme 1). The number of intercalators bound per available DNA base pair (degree of association) was determined by quantifying the remaining free RuII complexes in the supernatant of the solution using UV−vis spectroscopy after a brief centrifugation. The difference was assumed to correspond with uptake by the DNA−NP superlattice. The effect of intercalation on the melting behavior of the aggregate was studied by heating the solution at a rate of 0.25 °C/min while monitoring the extinction of the solution at 520 nm, the colorimetric signature associated with the gold nanoparticle plasmon resonance (Figure 2a). The Tm of these superlattices was calculated as the inflection point of the melting transition. Addition of any of the complexes at a constant concentration of 12.1 μM noticeably increased Tm, which can be attributed to intercalation of the dppz moiety into the sticky ends of the DNA, as this is the weak point in the DNA duplex and therefore the lattice. Each of the complexes resulted in a slightly different Tm increase, from 1 °C for complex 6 to 15 °C for complex 1, indicating that intercalator charge and structure significantly influence superlattice thermal stability. This could be due to two factors: either there were a different number of intercalation events occurring at each sticky end or each intercalator complex stabilized dsDNA to a different extent due to its molecular architecture. To determine the contribution of the latter, the Tm values of the DNA−NP superlattice after intercalation with complexes 1, 3, and 6 were compared as a function of degree of association (Figure 2b). This analysis shows that there is a slight contribution from the intrinsic architecture of the molecule on a per intercalator basis,

especially at higher degrees of association. This indicates that the large differences in Tm observed in Figure 2a are due to a difference in the number of molecules intercalating, which correlates to the binding affinity of the complex for DNA. To further explore the relationship between intercalator structure and its binding affinity (K) for free duplex DNA, fluorescence titration was performed to measure K (Figure S1, Table S2). If K values of members of this family of intercalators are compared in a systematic fashion, the relative contributions of the ancillary ligand could be determined. While the binding modes of different intercalating ligands are well characterized in the literature, the effect of extended ancillary ligands on the intercalation behavior of RuII complexes into the DNA duplex has yet to be explored extensively.22,25,26 Our results revealed that the structure and electrostatic charge of the ancillary ligand had a dramatic effect on K (see Supporting Information for detailed analysis). For this family of RuII intercalators, simply changing the number and the nature of the pendant group, which altered the overall charge of the complex, affected K by 2 orders of magnitude at low salt conditions. At higher salt concentrations, electrostatic interactions between the negatively charged DNA backbone and the positively charged complex are screened. A trend of decreasing binding affinity for complexes with more sterically hindered ancillary ligands was observed. The local charge of the pendant group had a more modest effect on binding affinity; complexes with more negatively charged pendant groups exhibited lower binding affinity for DNA. To determine if these trends translated to thermal stabilization of the superlattice, binding affinities were correlated with ΔTm,superlattice upon intercalation in 12.1 μM C

DOI: 10.1021/acsnano.5b07103 ACS Nano XXXX, XXX, XXX−XXX

Article

ACS Nano

Figure 3. Structural effects of intercalation emerge from SAXS and SEM. (a) Scattering patterns of the superlattice in increasing concentrations from 0 to 37.3 μM of complex 1. (b) Isotropic strain resulting from uniform lattice expansion and (c) RMS microstrain induced in superlattice upon intercalation. (d) DNA−NP superlattice single crystals retain structure upon intercalation. Before and after intercalation with complex 1. Scale bars are 2 μm.

nM NP in 40 μM intercalator) were heated at a rate of 1 °C/ min and scans were taken every 2 °C. The characteristic bcc diffraction peaks of the unintercalated control sample disappeared at 46 °C (designated T max ), indicating a dissociation of the nanoparticles due to dehybridization of the DNA bonds. Superlattices that had been incubated with intercalator remained intact at higher temperatures (close to the boiling temperature of water for complex 1). Importantly, the Tmax trends between the complexes mirrored those of Tm from the UV−vis melt data. The absolute differences between Tm and Tmax can be attributed to the different ramp rates used in these two techniques. These results confirm the idea that intercalation strengthens the DNA bonds between nanoparticles against thermal destabilization and that intercalator molecular structure is an important factor. In summary, we have demonstrated that increasing complex binding affinity for dsDNA, which changes depending on the charge and steric profile of the ancillary ligands of the intercalator, translates to increased bond strength. A wide range of thermal stabilities was achieved by intercalation with this family of complexes, suggesting the versatility of using this strategy in postsynthetic modification of DNA−NP superlattices. Structural Effects of Intercalation on the DNA−NP Superlattice. To fully characterize the structural effect of intercalation on both the superlattice bond and the overall

complex solution (Figure 2c). A positive, linear relationship between the binding affinity and ΔTm was observed, revealing that complexes with high binding affinity stabilized superlattices to the greatest extent. This result illustrates that, by modulating the molecular structure of the intercalator, the stability of DNA bonds in superlattices can be tuned in a rational manner. Furthermore, binding affinity of a complex for free dsDNA can serve as an indicator for the degree of bond stabilization. Binding assays confirmed that the intercalators were equilibrated into the DNA−NP superlattice with comparable affinity as for free DNA (Figure S2), providing evidence that the environment of the superlattice does not prevent diffusion of the complexes. As spectroscopic techniques can only provide information about whether particles are bound, small-angle X-ray scattering (SAXS) was used to gain insight into the positioning of nanoparticles within a superlattice after intercalation and during the heating and melting processes. SAXS is a powerful technique analogous to powder diffraction that provides information about the crystal symmetry, interparticle distance (the distance between nearest nanoparticle neighbors), and domain size of a material. By comparing radially averaged onedimensional SAXS data, any structural changes in a lattice can be determined. To probe whether intercalated superlattices retained their structure at higher temperatures, in situ SAXS was used (Figure S4). The annealed and intercalated samples (50 D

DOI: 10.1021/acsnano.5b07103 ACS Nano XXXX, XXX, XXX−XXX

Article

ACS Nano

Scheme 2. Modulation of Superlattice Bond Strength with RuII Intercalation Enables Stepwise Synthesis of Core−Shell Crystalsa

a

DNA−AuNP superlattice single crystals are intercalated with complex 1 overnight before being combined with complementary DNA−QdNPs. Slow cooling allows the DNA−QdNPs to assemble into a shell around core crystals. The presence of the intercalator increases the melting temperature of the AuNP core crystal, allowing it to stay intact during the annealing process that leads to shell formation.

exhibits some defects as the particles move relative to their positioning in a perfect bcc lattice. As additional intercalators are added, a slightly more uniform distribution of particle spacing is evidenced by a slight reduction in the microstrain, a result that we rationalize as arising from the smoothing of the statistical variation of intercalation events as the maximum loading is approached. Synthesis of Core−Shell Superlattice. As a demonstration of the effectiveness of using intercalators to increase the bond strength of DNA−NP superlattices, a core−shell material was synthesized. Previously explored strategies for assembling these materials have been based on methods such as layer-bylayer adsorption using electrostatics.27−29 The use of DNAassembly to create such structures would enable programmable design and synthesis of materials where each component is positioned in a precise manner. In a method analogous to seedmediated growth,30,31 an intercalated superlattice crystal was coated with a shell of another type of DNA−nanoparticle superlattice through subsequent thermal annealing steps (Scheme 2). The presence of the intercalator was crucial for keeping the core superlattice intact during the thermal annealing steps of the secondary growth step. This postsynthetic modification circumvented the need to redesign the DNA strands which could prevent the nanoparticles from reorganizing into their thermodynamic product.16 SEM confirmed that the rhombic dodecahedron crystal habit was preserved after postassembly intercalation (Figure 3d). Rhombic dodecahedron bcc superlattice single crystals were intercalated with complex 1 and then combined with complementary DNA−QdNPs. This mixture was slowly cooled from 45 to 25 °C with additional cycling steps around the Tm of the DNA−QdNPs at a rate of 0.01 °C/min (Scheme 2). The product was embedded in silica and studied using transmission electron microscopy (TEM). Figure 4 shows the resulting core−shell structures, which demonstrate the successful assembly of a superlattice shell of lower electron density (CdSe/ZnS) around a core single crystal of higher electron density (Au). These structures differ strikingly from TEM images of independently crystallized core and shell crystals (Figure S8). SAXS and fast Fourier transforms (FFT) of a TEM image (Figure S9) reveal that the QdNP shell is, in fact, crystalline. When a large excess of DNA−QdNPs was added, single crystals of only DNA−QdNPs formed in addition to the core−shell structures.

material, SAXS and scanning electron microscopy (SEM) were used. First, nanoparticles were functionalized and aggregated as previously mentioned. After aggregation, the amorphous assembly was slow cooled from 55 to 25 °C at a rate of 0.01 °C/min. This yielded micrometer-sized bcc superlattice single crystals. Each of these superlattice samples was titrated with 0.1−40 μM of RuII complex and then stored overnight (ambient temperature, shaking at 750 rpm) before being measured with SAXS. The data (Figure 3a) revealed a gradual shift in the q0 peak (from which interparticle distance is calculated) with increasing quantities of complex 1. Notably, this peak shift occurred without changing the overall bcc character of the structure factor. These changes in interparticle distance (i.e., bond length) can be described in terms of isotropic strain within the superlattice material. Figure 3b shows the calculated isotropic strain (η = ΔL/L0), or change in interparticle distance (ΔL) as a fraction of the original value (L0), after intercalation with complexes 1 and 6. Isotropic strain increased linearly at the same magnitude (2.9 Å/intercalation event) for both complexes before plateauing upon reaching 0.5 degrees of association, or the saturation point of possible intercalation sites. Since complexes 1 and 6 affect superlattice structure in an identical manner, it can be concluded that structural changes in duplexed DNA upon intercalation are solely dependent upon the intercalating dppz ligand and independent of the ancillary ligands. In this particular context, the lengthening bond can be correlated with an increase in bond strength, in contrast with atomic systems, where increased bond strength between any two atoms correlates with a decrease in bond length, e.g., a C−C triple bond is both shorter and stronger than C−C double bond. To probe local defects in the system introduced by intercalation, microstrain in the superlattices was quantified by analyzing peak broadening in the scattering patterns. Deconvoluting the effects of microstrain from peak broadening due to domain size effects can be done by performing a Williamson-Hall peak shape analysis (Figure S6). In the DNA− NP superlattice, microstrain represents the distribution of DNA lengths throughout a crystal. Figure 3c shows the fractional change in root-mean-square (RMS) microstrain as a function of degree of association. It is evident that the initial addition of intercalators increases the microstrain up to a maximum (4.5× the initial microstrain), after which the strain decreases. One possible explanation is that the initial distribution of intercalators (and thus elongation of DNA) throughout the superlattice crystal is not entirely uniform, and the crystal E

DOI: 10.1021/acsnano.5b07103 ACS Nano XXXX, XXX, XXX−XXX

Article

ACS Nano

(DTT, pH 8, Sigma-Aldrich) and then desalted on a NAP5 size exclusion column (GE Life Sciences). The purified thiolated strands were added to 15 nm diameter AuNPs (4 nmol DNA per mL nanoparticles). After 30 min, the solution was brought up to a total concentration of 0.01% sodium dodecyl sulfate. To increase the density of DNA on the nanoparticles without causing them to lose colloidal stability, 5 M sodium chloride (NaCl) was added stepwise over the course of several hours to reach a final concentration of 0.5 M NaCl. Each addition of salt was followed by 10 s of sonication. Following the salting procedure, the solution was placed on a shaker at 140 rpm and 37 °C overnight. The functionalized particles were purified by three rounds of centrifugation (14 000 rpm for 45 min). After each round, the supernatant was removed and the particles resuspended in Milli-Q water. Particle and DNA concentrations were measured on a Cary 5000 UV−vis spectrophotometer (Agilent). Extinction coefficients for the AuNPs and oligonucleotides were obtained from Ted Pella and the IDT Oligo Analyzer, respectively. Particle and DNA concentrations were measured on a Cary 5000 UV− vis spectrophotometer (Agilent). Extinction coefficients for the AuNPs and oligonucleotides were obtained from Ted Pella and the IDT Oligo Analyzer, respectively. Quantum Dot NP Functionalization. CdSe/ZnS QdNPs (Ocean Nanotech) were polymer coated and DNA functionalized as described previously.10 To polymer coat the QdNPs, nanocrystals were dissolved in chloroform (0.1 μM) and heated to 55 °C, after which N3−PMAO in chloroform (0.2 M monomer units) was added. After a gentle shaking period of 2 min, the solution was allowed to cool to room temperature. Solvent was evaporated using a rotary evaporator. Sodium borate buffer (75 mM, pH 9) was added to cover the nanoparticle layer in the flask, and then the mixture was sonicated in ice for 60 min. The solution was then filtered through 0.22 μm cellulose acetate syringe filter. This solution was then concentrated in a 100 kDa cellulose membrane spin filter (Amicon Ultra-15, Millipore) and then ultracentrifuged at 100 000 rcf in a continuous sucrose gradient (density 10% to 60% w/v) for 2 h, or until the nanoparticle band had separated from the floating layer of pure polymer. The layer of nanoparticles was carefully taken out using a syringe and then washed three times through 100 kDa spin filters using deionized water. These coated QdNPs were then functionalized with DBCO-modified DNA strands by combining them at 10-fold excess DNA at 0.5 M NaCl, shaking at room temperature (RT) and 750 rpm. After 5 h, the concentration of salt was increased to 0.5 M NaCl. Superlattice Assembly. DNA−AuNPs were assembled into aggregates by combining 25 nM each of AuNPs functionalized with sequences A and B with 350 equiv per particle of each linker nucleotide in 100 μL of 0.5 M NaCl. After aggregation at room temperature, the amorphous assembly was heated to a temperature slightly below the melting temperature (44 °C for this system) for 30 min. This led to the formation of polycrystalline bcc superlattices. To form single crystal bcc rhombic dodecahedron superlattices, the amorphous aggregates were transferred to PCR tube strips and slowly cooled in a Life Technologies PCR Thermocycler from 55 °C through the melting temperature to 25 °C at a rate of 0.01 °C/min.12 Core−Shell Synthesis. DNA−AuNP superlattice rhombic dodecahedra single crystals (10 nM) were intercalated in 40 μM complex 1 overnight (0.5 M NaCl, 1× PBS). Aggregates of complementary bcc DNA−QdNPs were added (50 and 100 nM total) and subjected to a thermal cycling procedure that slowly decreased from a temperature at least 5 °C above Tm,QdNP, and upon reaching Tm,QdNP, cycled five times to Tm,QdNP − 1 °C before cooling to room temperature. The ramp rate stayed constant at 0.01 °C/min. Fluorescence Binding Assay. Binding isotherms for each complex into free dsDNA were obtained by titrating a fixed quantity of intercalator complex with duplex DNA and measuring the fluorescence response (Figure S1). Duplex DNA was prepared by combining equimolar amounts of 18 bp sequence C with linker C in 5 mM Tris-HCl (pH 7.2) and 50 mM NaCl. This solution was heated to 70 °C for 5 min before allowing it to cool down slowly through the melting temperature, allowing the strands to hybridize. The amount of intercalator complex was fixed at 1 μM and this was titrated with

Figure 4. Core Au−Shell QdNP crystals. TEM images of micron sized core−shell structures synthesized in a stepwise manner. A difference in electron densities between AuNPs (denser) and QdNPs (lighter) results in contrast between the core and shell portions. Scale bars are 1 μm.

CONCLUSION We have demonstrated the effectiveness of using a family of RuII intercalators to increase the DNA bond strength and length of DNA−programmable nanoparticle superlattices. Through rational design of the intercalator ancillary ligands and choice of concentration, the thermal stability of the superlattice was tunable over a wide range. This method was used to increase the strength of the DNA bonds holding DNA programmable materials together, enabling the synthesis of a core−shell superlattice structure. The ability to control the thermal stability of nanoparticle assemblies is important for the creation of tailorable and functional materials. Such tunability in bond strength greatly expands the synthetic capabilities of using DNA-functionalized nanoparticles as building blocks in bottom up assembly of functional materials and provides a foundation for the assembly of increasingly sophisticated structures using DNA-mediated nanoparticle crystallization for plasmonic, photonic, and catalytic applications. METHODS RuII Complex Synthesis. The full synthesis, and chemical and photophysical characterization for the RuII complexes (Figure 1) were performed as described by Shade and Kennedy et al.24 Nanoparticle Functionalization. The oligonucleotides used in this work (Table S1) were synthesized on an MM48 solid-support automated DNA synthesizer (BioAutomation) using standard phosphoramidite chemistry and reagents purchased from Glen Research. All DNA was purified using reverse phase HPLC (Varian ProStar 210) and characterized using matrix-assisted laser desorption ionization time-of-flight mass spectrometry to confirm that the molecular weights matched their theoretical mass. AuNP Functionalization. Gold nanoparticles were DNA functionalized as described previously.3 Briefly, citrate capped AuNPs from Ted Pella were functionalized with a dense nucleotide shell. The 5′dimethoxytrityl-hexyl-mercaptan protecting group of the thiolated DNA strand was treated for 45 min with 100 mM dithiolthrietol F

DOI: 10.1021/acsnano.5b07103 ACS Nano XXXX, XXX, XXX−XXX

Article

ACS Nano dsDNA ranging in concentration from 10−9 to 10−2 M bp (corresponding with 5.6 × 10−11 to 5.6 × 10−4 M oligomer). For the assays performed at 0.5 M NaCl, additional salt was added. Each point in the titration curve was prepared as an individual sample and allowed 30 min to equilibrate before the measurement was taken. Performing a binding assay by titrating dsDNA into a single sample should yield the same results, as no photobleaching was observed for a given sample after running 20 consecutive scans. Fluorescence measurements were performed using a Horiba Jovin-Yvonne Fluorolog fluorometer. The sample temperature was maintained at 25 °C and the Peltier chiller for the detector was set to −20 °C. The sample was excited at the wavelength of 450 nm, and the emission was measured at 633 nm, followed by background subtraction. The observed fluorescence is assumed to be the sum of the weighted contributions of free and bound ligand:

over all types of binding sites on these AuNPs. Thus, we fit the binding isotherm data to the following relationship using nonlinear leastsquares:

B=

The resulting binding affinities of each of these complexes (Table S3) into DNA−programmable superlattices are very similar to their binding affinity into the SNAs. Melting Transition Measurements. AuNPs (1 pmol total) were functionalized and assembled into a bcc superlattice as described above. A defined volume of RuII complex was added to these bcc superlattices in a total volume of 1 mL of 0.5 M NaCl aqueous solution. After the samples were stored overnight at RT shaking at 750 rpm, each solution was transferred to a quartz cuvette containing a stir bar. The melting behavior of the aggregate was studied using UV−vis spectrometer. The extinction of the solution was monitored at 520 and 260 nm while heating the solution from 25 to 85 °C at a rate of 0.25 °C/min. The melting temperature is calculated from the point of inflection on the curve where the extinction at 520 nm increases due to the aggregate dissociating. Small Angle X-ray Scattering. SAXS experiments were performed at sector 5-ID of the DuPont-Northwestern-Dow Collaborative Access Team (DND-CAT) beamline at the Advanced Photon Source (APS) at Argonne National Laboratory. X-rays of wavelength 1.24 Å (10 keV) were used to probe the sample and the scattering angle was calibrated against a silver behenate standard. All SAXS data was collected at RT. To perform a SAXS measurement, roughly 80 μL of sample was loaded into 1.5 mm quartz capillary (Charles Supper Co.) and placed into a sample stage in the X-ray beam path. Two sets of slits were used to define and collimate the Xray beam, and a pinhole was used. The X-ray beam cross section measured 200 μm in diameter and a 0.5 s exposure time was used. A CCD area detector was used and dark current frames were subtracted from all data. An example of these scattering patterns can be seen in Figure S3. To obtain 1D data, two-dimensional SAXS patterns were azimuthally averaged and relative scattering intensity was plotted as a function of scattering vector q, where q is

F = F0(C t − C b) + FbC b where F is the apparent fluorescence at each DNA concentration measured at 633 nm, F0 is the fluorescence of the free ligand only, Fb is the fluorescence of the bound species, Ct is the concentration of total ligand, and Cb is the concentration of bound ligand. The following relationship is used to determine K of an intercalator for one DNA bp, derived from a study of the intercalation of DNA with the small molecule intercalator proflavine:32

F − F0 K[DNA] = Fb − F0 1 + K[DNA] Fluorescence binding isotherms were fit to this equation using nonlinear least-squares, from which the binding affinities were calculated. Absorbance Binding Assay. To ensure that the environment of the DNA−programmable superlattice and its constituent DNA− functionalized nanoparticle did not drastically reduce the affinity of these complexes for dsDNA, we obtained binding isotherms of the complexes into both the superlattice and colloidal DNA−AuNP to compare to that of free DNA (Figure S2). RuII complex was added to solutions containing 0.5 pmol AuNPs total of either superlattices or AuNPs functionalized with sequence B and loaded with linker B in 0.5 M NaCl. The final concentrations of RuII complex in these solutions ranged from 2 to 60 μM. This mixture was stored overnight at RT shaking at 750 rpm. The samples were subsequently vortexed and centrifuged. The DNA−NP superlattice samples were centrifuged for 10 s at 6000 rpm using a microcentrifuge, while the DNA−AuNP samples were centrifuged for 1.5 h at 15 000 rpm. The absorbance of the supernatant of each of these samples was then measured using a Cary 5000 UV−vis-NIR spectrophotometer (Agilent) and the amount of complex free in solution (Cfree) was quantified. This calculation used an extinction coefficient of 12 400 M−1 cm−1 at 445 nm corresponding with the absorbance of the complex’s metal to ligand charge transfer.33 The number of intercalators bound per available DNA base pair (degree of association) was calculated as follows:

Degree of association =

R total = NP ×

Ligand Bound K[L] = Receptor Bound 1 + K[L]

q=

4π sin(θ) λ

where θ is the scattering angle and the λ is the wavelength of X-ray radiation. The structure factor was determined using the relationship:

I(q) =

S(q) F(q)

where I(q) is intensity, S(q) is structure factor, and F(q) is the form factor. The particle form factor is due to individual dispersed particles solution and can be determined from a suspension of free particles. Isotropic Strain upon Intercalation. Nearest-neighbor distance (L, in nm) between the nanoparticles in the lattice is determined from the position of the first order scattering peak, q0 [110] in a SAXS pattern. For bcc symmetry, this equation is used:

C total − Cfree R total

350 oligomers 21.5 bp × NP oligomer

L=

where C refers to the amount of ligand (RuII complex), Rtotal refers to the total number of binding sites, and NP refers to the total moles of nanoparticle in a sample. We make the assumption that, due to the high salt concentration of the solution, all 350 linker oligonucleotides added to the sample hybridize to their complement sequence. Intercalation into dsDNA oriented and packed on a nanoparticle is a very different system than those described in existing models of DNA intercalation.34 Since developing a model to accurately describe such system is beyond the scope of this work, we used the model for a single binding site with a univalent receptor in order to extract an average K that describes the affinity of a complex for one bp averaged

1 6−1/2π 10 q0

Rise per base pair is calculated based on the following equation:

L − rA − rB − 0.75x − 0.8 Rise = 0 bp n where L0 represents the nearest neighbor distance prior to intercalation, rA and rB are the core radii of particles functionalized with sequences A and B, respectively, x is the number of hexaethylene glycol phosphate flexors (sp18) multiplied by a calculated 0.75 nm per sp18 for our system, n is the number of base pairs in between adjacent G

DOI: 10.1021/acsnano.5b07103 ACS Nano XXXX, XXX, XXX−XXX

Article

ACS Nano particles, and 0.8 nm accounts for the combined contribution from the hexylalkyl-thiol moieties on DNA sequences A and B. To determine the effect of intercalation on DNA elongation, RuII complex was added to DNA−programmable single crystal superlattices (0.5 pmol NP total). The concentrations of RuII complex ranged from 0.1 to 40 μM and these samples were stored overnight (RT, 750 rpm). The scattering pattern was taken for each sample and the isotropic strain (ηisotropic) calculated from

ηisotropic =

ization Experimental Center (NUANCE) using a Hitachi SEM SU8030. Images were acquired using an accelerating voltage of 10 kV.

ASSOCIATED CONTENT S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsnano.5b07103. Oligonucleotide sequences, free dsDNA binding isotherms, superlattice and spherical nucleic acid binding isotherms, SAXS patterns and line shape analysis, and Williamson−Hall analysis (PDF)

L − L0 L0

where L0 is as defined above and L is the interparticle distance after the intercalation event. When ηisotropic is plotted against degree of association (Figure 3b), a linear regime is observed from 0 to 0.3 degrees of intercalation. This initial slope (k) represents the steady increase in strain as a function of degree of association. From this, we can calculate the average rise per intercalation event using the following equation:

AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected]. Author Contributions

k × L0 Rise = Intercalation Event n

‡

These authors contributed equally to this work.

Notes

Williamson−Hall Analysis. To perform a Williamson−Hall analysis, S(q) from scattering patterns was fit to Lorentzian profiles (Figure S6). The integral breadth of each peak (β) was obtained and microstrain was calculated using a relationship that combines the Scherrer equation (size broadening) and the Stokes−Wilson equation (strain broadening):

β cos(θ) =

The authors declare no competing financial interest.

ACKNOWLEDGMENTS We thank J. Griffin for helpful discussions and for providing coding expertise for curve fitting analysis. This material is based upon work supported by AFOSR Award FA9550-11-1-0275, and the Centers of Cancer Nanotechnology Excellence (CCNE) initiative of the National Institutes of Health (NIH) under Award U54 CA151880. SAXS experiments were carried out at Sector 5-ID of the DuPont-Northwestern-Dow Collaborative Access Team at the Advanced Photon Source. Use of the Advanced Photon Source at Argonne National Laboratory was supported by the U.S. Department of Energy, Office of Science, Office of Basic Energy Sciences, under Contract No. DE-AC02-06CH11357. This work made use of the EPIC facility (NUANCE Center-Northwestern University), which has received support from the MRSEC program (NSF DMR-1121262) at the Materials Research Center; the International Institute for Nanotechnology (IIN); and the State of Illinois, through the IIN. M. Wang gratefully acknowledges a Graduate Research Fellowship from the National Science Foundation (NSFGRFP) and a Northwestern University Ryan Fellowship. J. Rouge acknowledges a postdoctoral fellowship from the PhRMA foundation. K. Brown gratefully acknowledges support from Northwestern University’s International Institute for Nanotechnology.

kλ + ε sin(θ ) τ

where θ is the scattering angle, β is the integral breadth of the scattering peak, λ is the wavelength of X-ray radiation, τ is the crystal size in the direction perpendicular to the beam, ε is the apparent strain, and k is the shape factor, which is approximated as 0.9 for spherical particles. This can be rewritten as β* =

1 + εq D

where β* = β cos θ/λ and q = 4 sin θ/λ. Plotting β* versus q should result in a straight line. The size of the crystallite (D) can be extracted from the y-intercept and the microstrain (ε) can be extracted from the slope.35−37 Scanning Electron Microscopy. DNA−NP superlattices were transferred to the solid state following the previous literature to allow for scanning electron microscopy (SEM) visualization.38 This method encapsulates solution phase single crystal superlattices in silica. First, the slow-cooled crystals were transferred to a 1.5 mL Eppendorf tube and the volume was brought up to 1 mL with 1× phosphate buffered saline. The quaternary ammonium salt N-trimethoxysilylpropylN,N,N-trimethylammonium chloride (TMSPA, Gelest, Inc.) was added in 1000-fold excess relative to the amount of phosphates in the aggregates (2 μL) to the solution containing the crystals. The tube was placed on a thermomixer (Eppendorf) to shake for 10 min at 750 rpm (RT) to allow the quaternary ammonium to associate with the DNA phosphate backbone. Then, 4 μL of triethoxysilane (TES) was added to the solution to initiate silica growth. The mixture was left shaking at RT and 750 rpm for 4 days. At that point, the crystals had aggregated at the bottom of the tube with a cloudy precipitate of silica formed throughout the solution. This supernatant was carefully removed and discarded. The embedded crystals were then transferred to a new eppendorf tube and dispersed in Milli-Q water. This mixture was purified with three rounds of centrifugation (5 min, 3000 rpm), removal of supernatant, and resuspension in Milli-Q water. Following the final round of centrifugation, the supernatant was removed and the pellet was dried, resuspended, and vortexed in 300 μL of ethanol. This solution (10 μL) was drop-cast onto a silicon wafer (NOVA Electronic Materials) for imaging. SEM images were obtained at the Northwestern University Atomic and Nanoscale Character-

REFERENCES (1) Desiraju, G. R. Supramolecular Synthons in Crystal Engineeringa New Organic Synthesis. Angew. Chem., Int. Ed. Engl. 1995, 34, 2311−2327. (2) Jones, M. R.; Seeman, N. C.; Mirkin, C. A. Programmable Materials and the Nature of the DNA Bond. Science 2015, 347, 204− 208. (3) Macfarlane, R. J.; Jones, M. R.; Senesi, A. J.; Young, K. L.; Lee, B.; Wu, J.; Mirkin, C. A. Establishing the Design Rules for DNA-Mediated Programmable Colloidal Crystallization. Angew. Chem. 2010, 122, 4693−4696. (4) Macfarlane, R. J.; Lee, B.; Jones, M. R.; Harris, N.; Schatz, G. C.; Mirkin, C. A. Nanoparticle Superlattice Engineering with DNA. Science 2011, 334, 204−208. (5) Macfarlane, R. J.; Jones, M. R.; Lee, B.; Auyeung, E.; Mirkin, C. A. Topotactic Interconversion of Nanoparticle Superlattices. Science 2013, 341, 1222−1225. H

DOI: 10.1021/acsnano.5b07103 ACS Nano XXXX, XXX, XXX−XXX

Article

ACS Nano

(26) Erkkila, K. E.; Odom, D. T.; Barton, J. K. Recognition and Reaction of Metallointercalators with DNA. Chem. Rev. 1999, 99, 2777−2796. (27) Liang, Z.; Susha, A.; Caruso, F. Gold Nanoparticle-Based CoreShell and Hollow Spheres and Ordered Assemblies Thereof. Chem. Mater. 2003, 15, 3176−3183. (28) Graf, C.; van Blaaderen, A. Metallodielectric Colloidal CoreShell Particles for Photonic Applications. Langmuir 2002, 18, 524− 534. (29) Dokoutchaev, A.; James, J. T.; Koene, S. C.; Pathak, S.; Prakash, G. S.; Thompson, M. E. Colloidal Metal Deposition onto Functionalized Polystyrene Microspheres. Chem. Mater. 1999, 11, 2389−2399. (30) Langille, M. R.; Zhang, J.; Personick, M. L.; Li, S.; Mirkin, C. A. Stepwise Evolution of Spherical Seeds into 20-Fold Twinned Icosahedra. Science 2012, 337, 954−957. (31) Lofton, C.; Sigmund, W. Mechanisms Controlling Crystal Habits of Gold and Silver Colloids. Adv. Funct. Mater. 2005, 15, 1197− 1208. (32) Schmechel, D.; Crothers, D. Kinetic and Hydrodynamic Studies of the Complex of Proflavine with Poly A·Poly U. Biopolymers 1971, 10, 465−480. (33) Liu, J. G.; Zhang, Q. L.; Shi, X. F.; Ji, L. N. Interaction of [Ru(dmp)2(dppz)]2+ and [Ru(Dmb)2(Dppz)]2+ with DNA: Effects of the Ancillary Ligands on the DNA-Binding Behaviors. Inorg. Chem. 2001, 40, 5045−5050. (34) McGhee, J. D.; von Hippel, P. H. Theoretical Aspects of DNAProtein Interactions: Co-Operative and Non-Co-Operative Binding of Large Ligands to a One-Dimensional Homogeneous Lattice. J. Mol. Biol. 1974, 86, 469−489. (35) Zhao, Y.; Zhang, J. Microstrain and Grain-Size Analysis from Diffraction Peak Width and Graphical Derivation of High-Pressure Thermomechanics. J. Appl. Crystallogr. 2008, 41, 1095−1108. (36) Mittemeijer, E. J.; Welzel, U. The “State of the Art” of the Diffraction Analysis of Crystallite Size and Lattice Strain. Zeitschrift für Kristallographie International Journal for Structural, Physical, and Chemical Aspects of Crystalline Materials 2008, 223, 552−560. (37) Senesi, A. J.; Eichelsdoerfer, D. J.; Brown, K. A.; Lee, B.; Auyeung, E.; Choi, C. H. J.; Macfarlane, R. J.; Young, K. L.; Mirkin, C. A. Oligonucleotide Flexibility Dictates Crystal Quality in DNAProgrammable Nanoparticle Superlattices. Adv. Mater. 2014, 26, 7235−7240. (38) Auyeung, E.; Macfarlane, R. J.; Choi, C. H. J.; Cutler, J. I.; Mirkin, C. A. Transitioning DNA-Engineered Nanoparticle Superlattices from Solution to the Solid State. Adv. Mater. 2012, 24, 5181− 5186.

(6) Jones, M. R.; Macfarlane, R. J.; Lee, B.; Zhang, J.; Young, K. L.; Senesi, A. J.; Mirkin, C. A. DNA-Nanoparticle Superlattices Formed from Anisotropic Building Blocks. Nat. Mater. 2010, 9, 913−917. (7) Auyeung, E.; Cutler, J. I.; Macfarlane, R. J.; Jones, M. R.; Wu, J.; Liu, G.; Zhang, K.; Osberg, K. D.; Mirkin, C. A. Synthetically Programmable Nanoparticle Superlattices Using a Hollow ThreeDimensional Spacer Approach. Nat. Nanotechnol. 2012, 7, 24−28. (8) Senesi, A. J.; Eichelsdoerfer, D. J.; Macfarlane, R. J.; Jones, M. R.; Auyeung, E.; Lee, B.; Mirkin, C. A. Stepwise Evolution of DNAProgrammable Nanoparticle Superlattices. Angew. Chem., Int. Ed. 2013, 52, 6624−6628. (9) Zhang, Y.; Lu, F.; Yager, K. G.; van der Lelie, D.; Gang, O. A. General Strategy for the DNA-Mediated Self-Assembly of Functional Nanoparticles into Heterogeneous Systems. Nat. Nanotechnol. 2013, 8, 865−872. (10) Zhang, C.; Macfarlane, R. J.; Young, K. L.; Choi, C. H. J.; Hao, L.; Auyeung, E.; Liu, G.; Zhou, X.; Mirkin, C. A. A General Approach to DNA-Programmable Atom Equivalents. Nat. Mater. 2013, 12, 741− 746. (11) Park, S. Y.; Lytton-Jean, A. K.; Lee, B.; Weigand, S.; Schatz, G. C.; Mirkin, C. A. DNA-Programmable Nanoparticle Crystallization. Nature 2008, 451, 553−556. (12) Auyeung, E.; Li, T. I.; Senesi, A. J.; Schmucker, A. L.; Pals, B. C.; de La Cruz, M. O.; Mirkin, C. A. DNA-Mediated Nanoparticle Crystallization into Wulff Polyhedra. Nature 2014, 505, 73−77. (13) Auyeung, E.; Morris, W.; Mondloch, J. E.; Hupp, J. T.; Farha, O. K.; Mirkin, C. A. Controlling Structure and Porosity in Catalytic Nanoparticle Superlattices with DNA. J. Am. Chem. Soc. 2015, 137, 1658−1662. (14) Park, D. J.; Zhang, C.; Ku, J. C.; Zhou, Y.; Schatz, G. C.; Mirkin, C. A. Plasmonic Photonic Crystals Realized through DNA-Programmable Assembly. Proc. Natl. Acad. Sci. U. S. A. 2015, 112, 977−981. (15) Barnaby, S. N.; Thaner, R. V.; Ross, M. B.; Brown, K. A.; Schatz, G. C.; Mirkin, C. A. Modular and Chemically Responsive Oligonucleotide “Bonds” in Nanoparticle Superlattices. J. Am. Chem. Soc. 2015, 137, 13566−13571. (16) Macfarlane, R. J.; Thaner, R. V.; Brown, K. A.; Zhang, J.; Lee, B.; Nguyen, S. T.; Mirkin, C. A. Importance of the DNA “Bond” in Programmable Nanoparticle Crystallization. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 14995−15000. (17) Pal, S.; Zhang, Y.; Kumar, S. K.; Gang, O. Dynamic Tuning of DNA-Nanoparticle Superlattices by Molecular Intercalation of Double Helix. J. Am. Chem. Soc. 2015, 137, 4030−4033. (18) Li, T. I.; Sknepnek, R.; Macfarlane, R. J.; Mirkin, C. A.; Olvera de la Cruz, M. Modeling the Crystallization of Spherical Nucleic Acid Nanoparticle Conjugates with Molecular Dynamics Simulations. Nano Lett. 2012, 12, 2509−2514. (19) Dhakal, S.; Kohlstedt, K. L.; Schatz, G. C.; Mirkin, C. A.; Olvera de la Cruz, M. Growth Dynamics for DNA-Guided Nanoparticle Crystallization. ACS Nano 2013, 7, 10948−10959. (20) Zeglis, B. M.; Pierre, V. C.; Barton, J. K. Metallo-Intercalators and Metallo-Insertors. Chem. Commun. 2007, 4565−4579. (21) Song, H.; Kaiser, J. T.; Barton, J. K. Crystal Structure of Δ[Ru(bpy)2dppz]2+ Bound to Mismatched DNA Reveals Side-by-Side Metalloinsertion and Intercalation. Nat. Chem. 2012, 4, 615−620. (22) Pyle, A.; Rehmann, J.; Meshoyrer, R.; Kumar, C.; Turro, N.; Barton, J. K. Mixed-Ligand Complexes of Ruthenium(II): Factors Governing Binding to DNA. J. Am. Chem. Soc. 1989, 111, 3051−3058. (23) Friedman, A. E.; Chambron, J. C.; Sauvage, J. P.; Turro, N. J.; Barton, J. K. A Molecular Light Switch for DNA: Ru(bpy)2(dppz)2+. J. Am. Chem. Soc. 1990, 112, 4960−4962. (24) Shade, C. M.; Kennedy, R. D.; Rouge, J. L.; Rosen, M. S.; Wang, M. X.; Seo, S. E.; Clingerman, D. J.; Mirkin, C. A. Duplex-Selective Ruthenium-Based DNA Intercalators. Chem. - Eur. J. 2015, 21, 10983− 10987. (25) Zhen, Q. X.; Ye, B. H.; Zhang, Q. L.; Liu, J. G.; Li, H.; Ji, L. N.; Wang, L. Synthesis, Characterization and the Effect of Ligand Planarity of [Ru(bpy)2L]2+ on DNA Binding Affinity. J. Inorg. Biochem. 1999, 76, 47−53. I

DOI: 10.1021/acsnano.5b07103 ACS Nano XXXX, XXX, XXX−XXX