Kinetics of Strand Displacement and Hybridization on Wireframe DNA

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Kinetics of Strand Displacement and Hybridization on Wireframe DNA Nanostructures: Dissecting the Roles of Size, Morphology, and Rigidity Casey M. Platnich, Amani A. Hariri, Janane F. Rahbani, Jesse B. Gordon, Hanadi F. Sleiman, and Gonzalo Cosa ACS Nano, Just Accepted Manuscript • DOI: 10.1021/acsnano.8b08016 • Publication Date (Web): 28 Nov 2018 Downloaded from http://pubs.acs.org on December 1, 2018

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Kinetics of Strand Displacement and Hybridization on Wireframe DNA Nanostructures: Dissecting the Roles of Size, Morphology, and Rigidity

Casey M. Platnich,1 Amani A. Hariri,1 Janane F. Rahbani, Jesse B. Gordon, Hanadi F. Sleiman*, Gonzalo Cosa* Department of Chemistry, McGill University, 801 Sherbrooke Street West, Montreal, Quebec H3A 0B8, Canada

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These authors contributed equally to this work

KEYWORDS single molecule fluorescence, dynamic DNA nanotechnology, structural cooperativity, size-dependent kinetics, stimuli-responsive nanomaterials

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ABSTRACT Dynamic wireframe DNA structures have gained significant attention in recent years, with research aimed towards using these architectures for sensing and encapsulation applications. For these assemblies to reach their full potential, however, knowledge on the rates of strand displacement and hybridization on these constructs is required. Herein, we report the use of single molecule fluorescence methodologies to observe the reversible switching between double- and single-stranded forms of triangular wireframe DNA nanotubes. Specifically, by using fluorescently labeled DNA strands, we were able to monitor changes in intensity over time as we introduced different sequences. This allowed us to extract detailed kinetic information on the strand displacement and hybridization processes. Due to the polymeric NT structure, the ability to individually address each of the three sides, and the inherent polydispersity of our samples as a result of the step polymerization by which they are formed, a library of compounds could be studied independently yet simultaneously. Kinetic models relying on simple exponential decays, multi-exponential decays or sigmoidal behavior were adjusted to the different constructs to retrieve erasing and refilling kinetics. Correlations were made between the kinetic behavior observed, the site accessibility, the nanotube length, and the structural robustness of wireframe DNA nanostructures, including fully single-stranded analogs. Overall, our results reveal how the length, morphology, and rigidity of the DNA framework modulate the kinetics of strand displacement and hybridization, as well as the overall addressability and structural stability of the structures under study.


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Three-dimensional wireframe DNA nanostructures are minimalistic, DNA-economical constructs that provide architecturally simple yet robust scaffolds attractive for many applications.1 Due to their relatively low cost of production and simplified assembly,2 in combination with the inherent addressability and biocompatibility of DNA, these cage-like structures have been used as programmable host materials, where encapsulation of many guests, including metal nanoparticles,3,4 siRNA,5,6 quantum dots,7 proteins,8,9 and anti-cancer drugs10 has been achieved. Importantly, these structures are stable in serum11 and have been delivered to cells,12,13 supporting their potential as drug delivery vehicles. One strategy that has been employed to release cargo from both wireframe DNA nanostructures3, 5, 14 and DNA origami structures15,16 is strand displacement, wherein a constituent DNA strand bearing a single-stranded overhanging portion (called the toehold or overhang) is removed from the structure upon hybridization to a fully complementary DNA stand.17 As a result of this stimulus, wireframe structures become partially single-stranded, gaining flexibility (Figure 1A. Note that the bending shown in the figures is only intended to aid the reader in visualizing the possible collapsed conformations of the nanotubes, while in reality, there are likely multiple conformations available to these structures), which has been used to selectively release cargo from DNA nanotubes,3, 18 as well as to modulate DNA nanotube bending.19


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Figure 1. Erasing and refilling of ATTO647N-labelled NTs. (A) Schematic representation (not to scale) of the erasing and refilling process of DNA NTs immobilized on pegylated glass coverslip via biotin-streptavidin interactions. (B) Removal of the labeled linking strand leaves the NT partly single-stranded as evidenced by the loss of ATTO647N signal, while the green fluorescence from Cy3 on the NT rung remains constant throughout, indicating structural stability and confirming that the labeled strands are binding specifically to the NTs. (C) Corresponding single molecule


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intensity-time trajectories for the erasing of the labeled linking strand. (D) Re-introduction of the labeled strand results in refilling of the single-stranded region and the recovery of the red fluorescent signal. (E) Corresponding single molecule intensity-time trajectories for the refilling of the linking strand.

Key to unraveling the full potential of DNA wireframe structures is knowledge on the rates associated with DNA strand mediated stimuli. While stand displacement kinetics are well documented for simple duplexes (relying on variables such as the strand length and sequence)20-24 the interplay between individual kinetic events in a multivalent DNA nanostructure remains, to the best of our knowledge, unexplored. Given its intrinsic complexity, it is plausible that the structure itself may play a major role in the rates of reaction, where steric crowding and tension forces are involved. Equally critical to exploiting DNA wireframe structures to their fullest, we posit, is achieving reversible structures, where strand displacement and hybridization can be used to enact structural and morphological changes with spatial and temporal control. Knowledge on the kinetics associated with these processes and their impact on the structural features of the construct are thus required to gaining greater control of dynamic DNA wireframe structures. Here we report single molecule fluorescence imaging studies on the kinetics of strand displacement and re-attachment of DNA strands along single wireframe DNA nanotubes (NTs). We chose as a structure of interest NTs with a triangular cross section consisting of repeat units (rungs) linked using three double-stranded struts (each consisting of a “bearing” and “non-bearing” strand that may be independently addressed (Figure 1A)).19 Our experiments involved introducing fluorescently-tagged overhangs to the non-bearing strand in the struts, yielding “erasable” strands along the NT that could be independently addressed and readily monitored via single molecule


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total internal reflection fluorescence microscopy (TIRFM). By delivering invading strands concurrently or sequentially onto surface immobilized NTs, we were able to visualize, in real time, one, two, or three sides of the NT undergoing deletion, observed as a loss of intensity. Partially or fully single-stranded struts were thus obtained that could then be refilled in a subsequent step with labeled strands re-introduced into the flow-cell, resulting in an intensity enhancement. From the resulting intensity-time trajectories, the kinetics of strand re-hybridization were determined. Overall, we found that the resulting rates were dependent on several factors, including NT length, the degree of double- or single-stranded character, and the local microenvironment of the overhangs/hybridization sites in question. Given the elongated structure of the NT, the ability to selectively modulate each of the three sides, and the inherent polydispersity of the samples (due to the step polymerization by which they are formed), a complex library of compounds could be studied individually yet simultaneously. By monitoring the intensity of each particle in a field of view as a function of time, erasing and refilling kinetics were retrieved. Kinetic models relying on simple exponential decays, multiexponential decays or sigmoidal behavior were adjusted to the different constructs. Correlations were made between the kinetic behavior observed (i.e. the presence or absence binding cooperativity or anticooperativity at erasing and refilling stages), the values of the rates, nanotube length, and the structural robustness of wireframe DNA nanostructures, including fully single-stranded analogs. Our findings show that while small NTs exhibit similar kinetic behavior to simple duplexes, large NTs are highly complex, with the steric hindrance and the flexibility or rigidity of the system altering the strand displacement/hybridization kinetics. Indeed, we found that while erasing follows a stochastic behavior for rigid structures, the erasing of more flexible constructs


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lead to the evolution of slower kinetics over time, reflecting anticooperativity for large NTs. Our data also illustrated that the surface may provide an additional steric constraint toward addressing strands buried under the structure. Interestingly, we found that varying the number of strands being hybridized simultaneously to the wireframe structure changes the kinetic behaviour significantly, with the refilling of one strand being a cooperative process while the refilling behavior for two strands shows no interdependence. Beyond structural cooperativity and size-dependent kinetics, our work further addressed the robustness of the DNA assemblies using single molecule two-color co-localization analysis. Our study revealed that the DNA NTs are generally durable and are capable of undergoing erasing and refilling cycles with minimal degradation. A small number (< 5%) of presumably malformed NTs is nonetheless observed and were destroyed or dislodged from the surface during flow. Overall, this work demonstrates how the size, morphology, and rigidity of the DNA framework may play a concerted role in steering the kinetics of strand displacement and the overall addressability of the structures under study. Properly exploited, one may conceive ways to harness this regio-specific chemistry in order to better control NT actuation. The work also highlights how single molecule fluorescence methodologies are uniquely poised to understand the complex kinetics and the structural robustness of DNA nanostructures, providing essential information towards the rational design and fabrication of DNA-based devices with improved performance.

RESULTS AND DISCUSSION Nanotube design and synthesis


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Working with a minimal set of only 12 DNA sequences, robust wireframe DNA NTs with controlled geometry were built, as previously reported by us (Figure 2A-C).19,25 These nanostructures require significantly fewer sequences to produce when compared to other DNA nanotubes that have been formed using single-stranded DNA tiles26 or DNA origami sheets rolled into tubes,27, 28 for example. The NT architecture was chosen as these structures are of interest for many applications due to their high aspect ratio, encapsulating potential, rigidity, and site-specific addressability.29 This last quality allows for reversible switching of the nanotube from fully double-stranded (ds) to partially single-stranded (ss) forms using toehold-mediated strand displacement strategies. These nanotube structures were thoroughly characterized in our previous work using gel electrophoresis and AFM.19


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Figure 2. DNA nanotube construction. (A) DNA rung, made up of six strands, and its dimensions (B) Linking strands (C) Fully formed nanotube with three rungs (grey-scale) and linking strands. Overhangs of 10 bases can be added to the linking strands to allow for strand displacement to occur at one, two, or three sides of the NT. The possible locations for the overhangs labeled with ATTO647N are indicated. In this diagram, R2 is modified with a 3’ Cy3 moiety for imaging, while R1 can be extended with a 10-base overhang to allow for the attachment of the complementary biotinylated strand. (D) For dimers, only one rung contains a biotin moiety. (E) For the 5% biotin NTs, attachment can occur at any point along the structure. Shown are possible attachment points in the middle of the structure, resulting in NTs laying along the surface, or at either end, resulting in NTs standing upright on the surface. Data for 100% biotin NTs is included in the Supporting Information.

Briefly, the NTs used here were composed of triangular “rung” units (Figure 2A) held together by three struts or “bearing strands” (Figure 2B), which were then made double-stranded, forming a fully double stranded triangular prism-shaped structure (Figure 2C). The doublestranded struts are referred to from here on as linking strands, or LS (Figure 2B), with the nonstructural, “erasable” strand indicated with an asterisk. In order to study the effects of nanotube length, we generated two types of structures: dimers (with only two rungs) and multiunit NTs (Figure 2D and E) that exhibited a range of sizes as a result of the step polymerization by which they are prepared.

Single molecule fluorescence erasing/refilling studies


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To serve as an indicator of structural stability we modified the structural core of the NT by labeling the rung with the green-absorbing fluorophore Cy3 (Figure 2C). To monitor the kinetics of strand displacement and hybridization, 10-base overhangs featuring an ATTO647N fluorophore were added to the 5’ end of any or all of the LS* as specified (Figure 2C). All overhangs consisted of an identical sequence (10T) to remove any artifacts in the kinetic data arising from intrinsic differences in the toehold length or sequence.21, 30, 31 Additionally, the use of poly-T overhangs minimized additional, unwanted hybridization interactions during NT formation.32 Of note, negligible FRET between Cy3 and ATTO647N was expected to occur, due to the large separation between them as dictated by the dimensions of the NT design (Figure S3).33 Only structures with both Cy3 and ATTO647N co-localizing were considered for analysis (Figure S4). NT were immobilized onto the coverslip surface via biotin streptavidin interactions:34 to this end rungs were prepared with strands bearing a biotin moiety. To understand how the attachment of the NT to the surface affected its mobility and kinetic properties, we made NTs with either 5% or 100% of the rungs being modified with a biotin. No major differences were found between these two species overall, and we focus the discussion exclusively on the 5% biotin samples (data for 100% biotin NTs is included in the Supporting Information). To immobilize the dimers, only the bottom rung was biotinylated (Figure 2D). Erasing and refilling solutions were prepared by diluting the DNA strand of interest in an oxygen scavenging cocktail. These solutions were then added to the coverslip surface using a flowcell35 and a syringe pump while monitoring the change in the ATTO647N signal using an EMCCD camera (see Supporting Information for details on the imaging hardware and oxygen scavenger solutions).


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Dimer erasing kinetics We initially tackled the kinetics of the simplest possible structure, a dimer (Figure 3A-C). Removal of one (or two simultaneous) fluorophore-labeled strand/s in a dimer yielded single molecule intensity time trajectories with one (or two) step decays in the intensity profile occurring randomly over time (Figures 3D-F). To determine the rate of erasing for the dimer species, the individual intensity-time trajectories for each molecule within a field of view (~150 molecules) were next combined to generate the corresponding ensemble decay trajectory (Figure 3G-I). The ensemble decay trajectories for one or two simultaneous strand removals followed a single exponential function, consistent with a stochastic process wherein each dimer acts as an independent unit. (Figure 3G-I). Of note, experimentally a “pre-pulse” period was recorded in the ensemble trajectory, prior to the erasing strand arrival. This pre-pulse period served to monitor the dye photobleaching for each experiment, to ensure it did not affect the erasing kinetic measurements (Figure 3G-I, greyed out region). Separate control experiments were also conducted to measure the photobleaching lifetime, on the order of 500 s and over an order of magnitude slower than the erasing lifetimes (see below, and Figure S5).


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Figure 3. Erasing of ATTO647N-labelled dimers. Scheme depicting general erasing process for biotin-immobilized DNA dimers with LS1* overhang (A), LS2* overhang (B), or LS2*/3* overhang (C). (D-F) Examples of SM intensity-time trajectories for the three samples listed previously. (G-I) Ensemble intensity-time plots constructed from combining over 100 single molecule trajectories. Greyed out region corresponds to the imaging period prior to the arrival of the erasing strand.


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Similar rates of strand displacement were recorded whether removing only LS2 or LS2 and LS3 simultaneously (Figure 3, H-I), with lifetime values of 11.30 ± 0.05 s and 10.69 ± 0.07 s, respectively (Table S2). This finding suggests there was no cross-talk between these laterally arranged sites on the dimer, where both were equally available to their incoming erasing strands. The removal of LS1, along the bottom, was found to be ~ 2-fold faster than the erasing of the strands along the top (Figures 3H and 3G, respectively, see also Table S2). While LS1* LS2* and LS3* have the same overhang (10T bases), LS1* has fewer base pairs to its bearing strand in the NT scaffold than LS2*/3* (36 versus 42 base pairs, respectively), resulting in its faster displacement in the dimer.23 The results on kinetics of erasing illustrate that for the dimer, all three sides of the structure are equally available for strand displacement. Notably, the dimer remained stable throughout, as evidenced by analysis of the Cy3 signal intensity before and after erasing for these samples (Figure S8).

Nanotube erasing kinetics Erasing of small NTs consisting of less than 10 repeat units resulted in a series of discrete steps in the single molecule intensity-time trajectories (Figure 4D-F), with each step corresponding to the removal of a single strand along the NT. The size of the NT involved in terms of number of repeat units may thus be inferred from the number of steps and the stoichiometry of the reaction (one vs two strands per repeat unit).29 Due to an insufficient number of small NTs within the samples, these traces were normalized and combined with large NTs to generate ensemble intensity-time trajectories for each experiment. Large NTs consisting of over 10 repeat units provided a unique perspective as each structure behaved as an ensemble in its own, displaying a continuous, rather than stepwise decrease


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in intensity (Figure 4G-I). By monitoring the intensity decay over time, the erasing lifetime for each single structure could thus be obtained and in turn correlated with its size (Figure 4). In turn, an average decay trajectory for large nanotubes and its associated decay function could be obtained from combining the decay trajectory of all nanotubes upon normalization. The normalization ensures that large and small NTs contribute with the same weight to the average, regardless of their number of repeat units (Figure 4J-L).


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Figure 4. Erasing of ATTO647N-labelled NTs. Scheme depicitng general erasing process for biotin-immobilized DNA NTs with LS1 overhang (A), LS2 overhang (B), or LS2/3 overhang (C). Sample SM intensity-time trajectory for small (D-F) and large NTs (G-I), for each sample type. (J-L) Ensemble intensity-time plots for each sample type, constructed by accumulating previously normalized single molecule trajectories. Greyed out region corresponds to the imaging period prior to the arrival of the erasing strand.

The ensemble average lifetimes of erasing the top strands (LS2* or LS2*/3*) we retrieved were equal whether one or two strands were being removed simultaneously (Figure 4K and L), indicating that this factor did not significantly impact the kinetics of erasing of the NT. Importantly, large NTs consistently exhibited a longer ensemble average lifetime (approximately double) as compared to either their corresponding dimers for the erasing of LS2 *and LS2*/3*. We hypothesize that the transition from fully double-stranded to partially single-stranded NTs induces bending within the NT, allowing it to adopt a collapsed structure on the surface, rendering a subset of the linking strands inaccessible to the incoming eraser strand.19 The shape of the single molecule intensity-time trajectories supports this theory for large NTs, where a small residual intensity subsists after erasing (Figure 4 H and I), indicating that some of the labeled strands are not erased. The residual intensity is also apparent though not nearly as prominent within the ensemble average decay. The largest nanotubes represent a small fraction of the total population (Figure 5), and have a minor contribution in the ensemble average decay constructed from normalized trajectories. When erasing of LS1* was executed, the steric hindrance of a subset of strands, partly associated to the surface close proximity and the bending structure, reflected prominently in the


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SM trajectories and in the ensemble trace ((Figures 4G and 4J, respectively) in the form of a residual intensity corresponding to a non-erasable fraction. Additionally, the ensemble average lifetime we recorded for erasing of LS1* on the NTs was ~ 8-fold larger than that of its corresponding dimer counterpart (39.2 ± 0.2 s versus 4.81 ± 0.04 s, respectively). Furthermore, removing LS1* on the large NTs was also approximately 2-fold slower than removing a strand (LS2*) from the top face (Figure 4B and C). Both of these facts indicate that this site is hindered by the coverslip surface and the bending of the NT. In summary, while erasing one or two sides from the top of the NT does not depend on NT size, the erasing of LS1* is more complex, with the bending of the NT onto the coverslip surface radically changing the accessibility of the overhang. By extracting the lifetime of erasing for fifty randomly selected individual NTs from each of the NT samples and correlating them with their size, we found a small dispersion of values around the ensemble average lifetime for erasing LS2* and LS2*/3*. Remarkably, for LS1* erasing, a significantly larger dispersion around the mean was observed, see Figure 5A. We posit that the larger dispersion of erasing lifetime values is due to differences in overhang orientation (and therefore accessibility) relative to the surface. This finding is important, as it suggests that the kinetics of strands displacement on wireframe DNA nanostructures can be manipulated by modulating the steric environment of the overhang.


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Figure 5. Correlation of the erasing lifetimes retrieved for single NT with their initial intensity. (A) LS1, (B) LS2, (C) LS2/3. The red dashed line denotes the average ensemble lifetime calculated from the normalized intensities for each sample. Also shown as single data points are the lifetimes for the corresponding dimer (orange).

Fully single-stranded NTs In order to better understand the steric environment of LS1 and the conformational changes of the NT, we sought to generate a fully single-stranded analog of the NT (Figure 6A). This was achieved by first erasing LS2* and LS3* to obtain partially single-stranded NTs, and then flowing the erasing strand for LS1*, resulting in fully single-stranded NTs. The ensemble average intensity-time trajectory for the erasing of LS1* from partially single-stranded NTs was adjusted with a bi-exponential decay function (Figure 6B). Two lifetimes were extracted from this fitting: a very fast initial lifetime of erasing for LS1* (13.0 ± 0.1 s) and a second, much slower decay with


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a lifetime of 395.1 ± 0.8 s, similar to the lifetimes we observed for photobleaching. The corresponding SM trajectories show that the larger NTs in a given field of view were not fully erased (Figure 6E). We hypothesize that, upon removal of the third set of linking strands, the larger NTs become sufficiently flexible to completely fold in on themselves, preventing invading strands from hybridizing. The slow decay is therefore dominated by the photobleaching of these inaccessible overhangs (Figure 6A). As shown in Figure 6C, the Cy3 intensity, a marker of structural resilience, for small NTs dropped slightly after erasing the third side, while the intensity remained constant for the majority of large NTs. This is most likely because the large structures never reached a plausibly more fragile fully single-stranded analog due to collapse.


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Figure 6. Erasing LS1 on partially single-stranded NTs. (A) Scheme illustrating erasing on small and large NTs. (B) Ensemble intensity-time trajectory from normalized SM traces, showing residual intensity decay following rapid initial erasing. (C) The slope of Cy3 intensity recorded before and after erasing for each fully single-stranded NT is smaller than that recorded for partially single-stranded NTs (Figure S8 for comparison), illustrating that the former are more fragile than the latter and may shed a fraction of their rungs (see inset) (D) Small NTs show single molecule intensity-time trajectories that rapidly fall to zero upon erasing while (E) large NTs show sustained residual intensity after initial fast erasing. Greyed out region corresponds to the imaging period prior to the arrival of the erasing strand.

As a general conclusion to the erasing experiments, our results show that the rates are determined by the availability of the overhangs, which is dictated by steric effects. For this reason, the side of the NT facing the surface exhibited slower erasing kinetics compared to its counterparts directed away from the surface, despite the fact that fewer base pairing interactions have to be disrupted in order to remove LS1*. In contrast, removal of LS1* occurs at a faster rate than LS2*/3* in the case of the dimer, where all three sides are equally accessible and steric hindrance does not play a considerable role. When looking in detail at the rates, NT may be seen as a collection of “n” units undergoing reaction, units that may or may not be equivalent. Accordingly, the observed erasing kinetics may show a similar stochastic behavior and comparable rates to those recorded for a dimer (all NT units are equivalent). Alternatively, as was the case for large NT, anti-cooperative phenomena may arise as erasing takes place, implying that the units along the NT are non-equivalent.


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Altogether, these results demonstrate how the size, morphology, and rigidity of the DNA framework may play a concerted role in steering the kinetics of strand displacement and the overall addressability of the structures under study. These properties offer opportunities, in addition to varying the sequence and length of the DNA duplex in question, to better control NT actuation.

Dimer refilling kinetics The same structures that were initially addressed in the erasing studies were next also studied in refilling experiments. For dimers, the refilling intensity-time trajectories appeared as a single step (Figure 7A) or two steps (Figure 7B), depending on the number of sites being refilled. These trajectories were next combined to generate the corresponding ensemble trajectory which was adjusted with a single-exponential growth function. The ensemble lifetime for refilling two strands was double (73.4 ± 0.5 s) that of refilling a single strand on the dimer (40.4 ± 0.6 s), see also Figures 7A and 7B. We hypothesize that this may be due to a more collapsed configuration with increased single-stranded content in the structure impeding the arrival of the refilling strand. Intriguingly, LS1 refilling for the dimer spanned over a period of hours rather than seconds, meaning that this site is not available for refilling, likely due to collapse of the single-stranded LS1 onto the coverslip surface (see Figure S13).


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Figure 7. Refilling of dimers. (A) with LS2 single-stranded or (B) with LS2 and LS3 singlestranded. Single molecule intensity-time trajectories are shown on the left, while the corresponding ensemble trajectories are shown on the right. Note that the background increase due to flow of labelled strands into the imaging chamber was subtracted from the single molecule trajectories. Greyed out region corresponds to the imaging period where the erasing strand is flowed in and initates binding.

NT refilling kinetics As for the erasing experiments, small NTs (ca. < 10 subunits) showed stepwise refilling intensity-time trajectories (Figure 8A and E), while large NTs showed a continuous increase in


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intensity each behaving as an ensemble in their own, see Figures 8B and 8F. Intriguingly, large NTs displayed two different trajectory shapes depending on the number of strands being refilled simultaneously: refilling one side produced a curve that followed a sigmoidal shape (Figure 8B), whereas refilling two sides generated a curve following an exponential growth shape (Figure 8F). We found that these observations were consistent for all large (>10 rungs) NTs within these sample sets. The sigmoidal shape of the SM traces for refilling one side indicates cooperativity in the refilling process: We posit that because the available binding sites are all linearly arranged, each site that is refilled and rigidified in turn helps to pre-organize adjacent sites for binding. In comparison, when both LS2 and LS3 are single-stranded, the entire structure becomes more flexible and filling one binding site has less of a pre-organizing effect on its surrounding sites, resulting in a stochastic refilling.


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Figure 8. Refilling partially single-stranded NTs. For each sample, a single molecule trajectory is shown for small NTs (A and E) and large NTs (B and F). Ensembles were constructed upon combining the normalized intensity time trajectories for 50 individual NTs are given in (C) and (G). The distribution of half-lives/lifetimes as a function of the intensity (size) is also shown (D and H). The red dashed line represents the ensemble half-life/lifetime for that sample. Also shown as single data points are the lifetimes for the corresponding dimer (orange). Greyed out region corresponds to the imaging period where the erasing strand is flowed in and initates binding.


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Refilling of the NT was substantially slower than that of their dimer counterparts in all cases, indicating that steric hindrance may slow down the rate of this reaction, see Figures 8C and 8F. In general, however, the lifetime/half-life data does not show a strong correlation with size, see Figures 8D and 8H. It is plausible that the kinetics of refilling are more dependent on the local microenvironment of each docking site and cannot be generalized based on NT size. The refilling of LS1, along the bottom of the NTs, occurred very slowly compared to those for LS2 and LS2/3: Only by incubating the NTs overnight with the LS1*-ATTO647N refilling solution was the fluorescence fully restored, see Figure 9. This result suggests that while the 10base overhangs along the bottom are accessible for LS1* strand displacement, the binding site left behind along the bottom of the NT is sterically unavailable for the incoming strand. Importantly, this reveals a clear difference between the erasing and refilling processes in NTs. While erasing kinetics only require accessibility of overhangs, which does not change dramatically with rigidity or proximity to the surface, refilling requires the availability of single-stranded sites within the nanotube, which is strongly dependent on steric access. Thus, we conclude that hybridization kinetics are strongly impacted by and can be modified upon manipulating the steric environment of nanostructures relative to a surface.


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Figure 9. Slow refilling of LS1 on nanotubes. (A) Schematic showing the refilling of LS1. (B-E) Red and green intensity correlations for each NT in one field of view. Fields of view were changed between each time point to reduce the impact of photobleaching in the analysis.

In summary, our results demonstrate that steric effects due to surface immobilization can be used to modulate the kinetics of strand displacement and hybridization on DNA nanostructures. By changing the orientation of the overhangs/docking sites relative to the surface, the rates can be altered, thus adding new elements to the toolbox of available strategies for controlling the kinetics of strand displacement and DNA hybridization.

CONCLUSION We have reported herein SMF methodologies for the study of the kinetics of opening and closing of dynamic wireframe DNA nanostructures, based on strand displacement and subsequent


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re-hybridization of the displaced species. Our results demonstrate the usefulness of SMF techniques to study systems with a high degree of polydispersity. In particular, these methods allowed us to correlate NT size with the kinetics of erasing and refilling strands from the NT. The strategies employed herein also allowed us to probe the various sides of the NT independently from one another, providing insight into the different local environments created by the interactions of these nanostructures with the modified coverslip surface to which they are attached. Our findings also suggest that by anchoring a DNA nanostructure to a surface, one can substantially change the kinetics properties of the various sides in a site-specific manner. Even though chemically equivalent sites would theoretically exhibit similar rates in solution, by placing the structure on a surface, the symmetry can be broken. Just as an understanding of the interaction energies and structural properties of DNA duplexes enabled their use in a variety of applications, supramolecular DNA structures will only reach their full potential when we understand - and therefore can exploit - their structural intricacies. Globally, this work reveals how the size, morphology, and rigidity of a DNA nanostructure influence the kinetics of strand displacement and re-hybridization. This knowledge will be especially important, we posit, to those studying nanostructures using methods based on DNA-PAINT, wherein the accessibility of these different structural entities may impact the experimental outcomes. Furthermore, this information will allow for greater addressability of the structures under study, which may in turn allow for enhanced control over NT actuation for applications in sensing and drug delivery.

MATERIALS AND METHODS


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Materials.

Acetic

acid,

boric

acid,

EDTA,

urea,

magnesium

chloride,

GelRed,

tris(hydroxymethyl)aminomethane (Tris), D(+) glucose, β-mercaptoethanol, and streptavidin were purchased from Aldrich. Nucleoside (1000 Å)-derivatized LCAA CPG solid supports with loading densities of 25-40 μmol/g, Sephadex G-25 (super fine DNA grade), and reagents for automated DNA synthesis were used as purchased from BioAutomation. Acrylamide (40%)/bis-acrylamide 19:1 solution was purchased from BioShop. For TIRFM surface sample preparation, 1% v/v Vectabond/acetone was purchased from Vector Laboratories, while poly-(ethylene glycol) succinimidyl valerate MW 5000 (mPEG-SVA) and biotin-PEG-SVA were purchased from Laysan Bio, Inc. Imaging chamber components were custom-made, purchased from Grace Bio-Lab. TBE buffer is composed of 90 mM Tris and boric acid and 1.1 mM EDTA, with a pH of ∼8.3. TAMg buffer is composed of 45 mM Tris and 12.5 mM MgCl2 with a pH of ~7.8 adjusted by glacial acetic acid. DNA nanotube/dimer design and assembly. The nanotube design and synthesis are based on the work of the Sleiman group described in detail in previously published protocols.17 Details on the synthesis and purification of oligonucleotides can be found in the Supporting Information. Single molecule surface preparation. Glass coverslips were cleaned and passivated using previously reported methods.29,34,35 These techniques are detailed in the Supporting Information. Single molecule imaging. Single molecule fluorescence imaging was carried out on a Nikon Eclipse Ti inverted microscope. Details on our imaging configurations and the procedures we used are listed in the Supporting Information.

SUPPORTING INFORMATION


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The Supporting Information is available free of charge on the ACS Publications website. The Supporting Information includes DNA sequences, detailed descriptions of nanotube design and synthesis, single molecule surface preparation and imaging information, co-localization images, photobleaching experiments, intensity histograms of all samples, Cy3 intensity analysis for NT robustness for all samples, all data pertaining to 100% biotin samples, erasing data for fully single stranded dimers, LS1 refilling data for dimers, details on how data analysis and curve fitting was carried out, and a table of all rate constants.

AUTHOR INFORMATION Corresponding Author Correspondence to Hanadi F. Sleiman ([email protected]) or Gonzalo Cosa ([email protected])

Present Addresses Amani A. Hariri: Department of Radiology, Stanford School of Medicine, 350 Serra Mall, Stanford, CA 94305-5105 Janane F. Rahbani: Rosalind and Morris Goodman Cancer Research Centre, Department of Biochemistry, McGill University, 3655 Promenade Sir William Osler, Montreal, QC, H3G1Y6 Jesse B. Gordon: Department of Chemistry, Massachusetts Institute of Technology, 77 Massachusetts Avenue, 18-393, Cambridge, MA 02139

Author Contributions


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C.M.P and A.A.H. carried out the single molecule imaging and all associated data analysis with assistance from J.B.G. C.M.P., A.A.H. and J.F.R. contributed DNA sequences and prepared samples. C.M.P., A.A.H., H.F.S., and G.C. designed the experiments and wrote the manuscript. H.F.S and G.C. coordinated and oversaw the study. All authors have given approval to the final version of the manuscript.

ACKNOWLEDGMENT G.C. and H.F.S. are thankful to the National Science and Engineering Research Council of Canada (NSERC), the Canada Foundation for Innovation (CFI), the Fonds de Recherche Nature et Technologies (FRQNT) and the Canada Institute for Health Research (CIHR). H.F.S. is also thankful to the Canada Research Chairs program and is a Cottrell Scholar of the Research Corporation. We are thankful to the McGill CIHR Drug Development Training Program (C.M.P. and A.A.H.), GRASP (A.A.H.), FQRNT, (A.A.H.) and NSERC (C.M.P) for postgraduate scholarships and to NSERC (J.B.G) for an undergraduate scholarship. We are also thankful to Y. Gidi for insightful discussions.


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For table of contents only. 82x44mm (300 x 300 DPI)


Kinetics of Strand Displacement and Hybridization on Wireframe DNA

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