Terminating DNA Tile Assembly with Nanostructured Caps - ACS

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Terminating DNA Tile Assembly with Nanostructured Caps Deepak K. Agrawal, Ruoyu Jiang, Seth Reinhart, Abdul Majeed Mohammed, Tyler Dean Jorgenson, and Rebecca Schulman ACS Nano, Just Accepted Manuscript • DOI: 10.1021/acsnano.7b02256 • Publication Date (Web): 13 Sep 2017 Downloaded from http://pubs.acs.org on September 16, 2017

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Terminating DNA Tile Assembly with Nanostructured Caps Deepak K. Agrawal 1,†, Ruoyu Jiang1, Seth Reinhart1, Abdul M. Mohammed1, Tyler D. Jorgenson1†, Rebecca Schulman1,2,* 1

Chemical and Biomolecular Engineering, Johns Hopkins University, Baltimore, MD 21218; 2

Computer Science, Johns Hopkins University, Baltimore, MD 21218

KEYWORDS: DNA nanotubes, DNA origami, self-assembly, nanotubes, nucleation, growth, capping, hierarchical self-assembly

ABSTRACT: Precise control over the nucleation, growth and termination of self-assembly processes are fundamental tools for controlling product yield and assembly dynamics. Mechanisms for altering these processes programmatically could allow the use of simple components to self-assemble complex final products or to design processes allowing for dynamic assembly or reconfiguration. Here we use DNA tile self-assembly to develop general design principles for building complexes that can bind to a growing biomolecular assembly and terminate its growth by systematically characterizing how different DNA origami nanostructures interact with the growing ends of DNA tile nanotubes. We find that nanostructures that present binding interfaces for all of the binding sites on a growing facet can bind selectively to growing ends and stop growth when these interfaces are presented on either a rigid or floppy scaffold. In 1 ACS Paragon Plus Environment

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contrast, nucleation of nanotubes requires the presentation of binding sites in an arrangement that matches the shape of the structure’s facet. As a result, it is possible to build nanostructures that can terminate the growth of existing nanotubes but cannot nucleate a new structure. The resulting design principles for constructing structures that direct nucleation and termination of the growth of one-dimensional nanostructures can also serve as a starting point for programmatically directing 2- and 3-dimensional crystallization processes using nanostructure design.

Molecular self-assembly is an emerging method for inexpensively constructing devices that allows for control over nanoscale and microscale features and massively parallel fabrication. One-dimensional inorganic1, 2 and biomolecular nanostructures3-6 have been assembled that have applications in electronics, optics, chemical sensing and in drug delivery systems,7-10 and selfassembly processes increasingly are being used to produce complex two- and three-dimensional structures.11-16 Methods that use molecules such as DNA that allow for a combinatorial variety of tunable interactions between components have led to self-assembly processes rapidly being developed for structures of greater structural and functional complexity.17-20 While a major focus of self-assembly research has been the development of processes for assembling a specific final product or device,21 biology demonstrates that self-assembly might make it possible to build not only economically fabricated devices with fixed form but also to build devices that can dynamically assemble and reconfigure in response to the state of the current environment. The components of the cytoskeleton, for example, can dynamically assemble into a variety of materials, sensors, propellers, machines and mechanical transducers and then disassemble and reform different structures. This type of organization occurs through continued spatiotemporal control over the nucleation, growth, termination and hierarchical 2 ACS Paragon Plus Environment

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organization of self-assembled filaments. The ability to control the dynamics of structure formation and reorganization could lead to the construction of devices whose structure (and thus function) changes in controlled ways over time and which can adapt to the environment. DNA tile assembly processes, where components are well-defined DNA units such as single strands,22, 23 small nanostructures24-27 or origami structures,28-31 represent a promising route for developing synthetic devices that can dynamically assemble, disassemble and reconfigure. In DNA tile assembly, a set of components that present DNA interfaces, often sticky ends that interact through hybridization, are assembled into lattices, arrays or tubes. The set of tiles and their sticky end sequences can control the structure of what is assembled and the pathway for assembly. Accessory DNA strands or other self-assembled complexes can also programmatically direct nucleation,26, 32, 33 disassembly34 or activate or deactivate tiles,35 and designed DNA reaction cascades or circuits can produce outputs that in turn direct growth or assembly.36 Increasingly precise models of assembly kinetics and structure37 make this control reliable so that such control mechanisms can increasingly be combined to direct more complex assembly and reconfiguration processes.38-40 While control over the nucleation and growth of DNA tile systems26, 32, 33, 38, 39 has been the subject of multiple investigations, relatively little is known about how to terminate growth. Quenching a reaction can stop the growth of all structures,41, 42 but more complex methods are required in order to exert selective control or to terminate the growth of one type of nanostructures while allowing other types of assembly reactions to continue. Here we develop a method to selectively terminate the assembly of growing structures in which a designed complex binds to a facet where growth is occurring, preventing the attachment of further monomers. We use DAE-E DNA tile nanotubes,43 a model system for understanding and controlling the dynamics of DNA tile self-assembly that has been used to study controlled 3 ACS Paragon Plus Environment

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nucleation38,

44

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architecture formation and triggerable growth,36 and driven assembly and

disassembly.45

Figure 1: DNA tiles, nanotubes, seed and caps and schematic of the capping process. (a) DNA tile sequences and structure. Tile sequences here are termed RS. (b) The two tiles in (a) coassemble into nanotubes at room temperature (20 ºC). (c) The RT A seed (top left), a DNA origami structure, serves as a template for nanotube nucleation by emulating the facet of a DAEE tile nanotube at its A interface. The cap structures studied here (top right) attach to sticky ends at the B interface. Gray lines indicate unfolded scaffold regions (not drawn to scale). (d) Nanotubes nucleate at RT A seeds, and grow via tile addition steps at the B interface. Caps can attach to the B interface, stopping further nanotube growth.

We term a complex that binds to the growing end of DNA tile nanotubes and halts nanotube growth a “nanotube cap”. To develop an effective capping structure, we show first how nanostructures that act as templates for nanotube nucleation from specific nanotube facets can 4 ACS Paragon Plus Environment

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also act as caps by binding to that facet on existing nanotubes. While these structures effectively terminate the growth of existing nanotubes, they can also nucleate new nanotubes in their unbound state. To build cap structures which can terminate nanotube growth but not nucleate new nanotubes, we systematically modify the structure of these caps to develop dedicated capping structures that can halt the growth of existing DNA nanotubes by binding to their ends, but do not themselves nucleate new nanotubes. We also show that these caps selectively bind to nanotubes with complementary tile types and that multiple types of caps can selectively act on their respective nanotube tile ends. Together, seeds and caps allow control over both the initiation and termination of nanotube growth. This control could allow precise control over the assembly and disassembly of DNA nanotubes and nanotube architectures. More generally, the systematic design of seeds and caps with DNA origami, whose structure can be precisely controlled, elucidates specific design principles for the design of these structures: nucleation sites might present binding sites for monomers in a well-defined geometry consistent with the structure of a growing homogeneous nucleus, while structures that present binding sites for the growth end on a flexible scaffold can reliably terminate growth but not nucleate new structures. Such principles can guide the construction of complexes that nucleate and terminate growth of a wide variety of nanostructures. The DNA nanotubes we study are assembled from DAE-E tiles that form from 5 component DNA strands (Fig. 1a).43 These DNA tiles assemble into nanotubes through the hybridization of short, single-stranded sticky ends (Fig. 1b). Nanotubes that nucleate homogenously can have a range of circumferences, ranging from 4 to 12 tiles, but nanotubes grown from DNA origami templates have a monodisperse circumference of 6 tiles.38 Growth of nanotubes occurs at free

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nanotube ends; a cap must be able to prevent tile attachment at the multiple potential growth sites at these ends.

RESULTS AND DISCUSSION Templated nucleation of nanotubes from both facets at room temperature. To study nanotube capping, we characterized how caps interact with nanotubes grown from DNA origami seeds. Because these seeds remain attached to nanotubes at one facet throughout the growth process, growth of seeded nanotubes occurs at only one facet. The binding of caps to that facet will therefore stop all growth of nanotubes, making it possible to determine whether caps are functional by determining whether nanotubes stop increasing in length after caps bind to nanotube ends. To template nanotube growth, we use DNA origami seeds that emulate the structure of a nanotube facet.38 A seed is a DNA origami structure consisting of 12 parallel DNA helices connected by crossover points arranged into a cylinder.38 Attached to this structure are adapter strands whose crossover structure matches that of the DNA nanotubes. These adapter strands are designed to minimize structural dissimilarity between the origami and present sticky ends for tile binding (Fig. 1c). It was previously shown that seeds can template nucleation of DNA tile nanotubes at 32 ºC where spontaneous nucleation of nanotubes is very rare.38 At room temperature, however, the tiles used in that study can rapidly nucleate new nanotubes even in the absence of seeds, which limits how control over nucleation can be applied in practice. In particular, it has been necessary to characterize the nucleation and growth of nanotubes in situ at 32 ºC, or to quench the assembly reaction with specifically designed strands before imaging or use under different 6 ACS Paragon Plus Environment

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conditions. Here we modified the tiles and seeds so that the nucleation and growth of nanotubes is controlled by seeds at room temperature so that the self-assembly processes considered here and the characterization of the resulting structures both take place under ambient conditions (Fig. 1a, c Supp. Notes S1 and S3). Because these modified seeds were designed to allow nanotubes to nucleate and grow at room temperature from the A facet of the nanotubes (Fig. 1a), we called them RT (room temperature) A seeds. To test that RT A seeds can direct nanotube nucleation, we annealed and purified RT A seeds labeled with Atto 647N dye and purified them using membrane filters and centrifugation (See Methods, Supp. Note S9). Separately, we annealed aliquots of RS tile strands and after the solutions reached room temperature, added either 2 pM of RT A seeds or an equivalent volume of buffer (for a control) such that the final tile concentration in each mixture was 45 nM (Supp. Note S11). We measured the number of nanotubes that grew and their lengths using two-color fluorescence microscopy images of the reaction mixtures that were pipetted onto a glass surface. Samples containing RT A seeds and tiles had ~6 times more nanotubes after 25 hours than mixtures containing only tiles (Supp. Fig. S10). RT A seeds were visible at most nanotube ends in the seeded sample, consistent with their roles as templates for growth. 81.0 ± 0.4% seeds grew nanotubes. We then modified the seeds to present adapter strands that nucleate nanotubes from the B growth facet rather than the A growth facet to create RT B seeds (Supp. Note S5). Like RT A seeds, RT B seeds increased the number of nanotubes present by a factor of ~6 (Supp. Fig. S11). 77 ± 4% of RT B seeds grew nanotubes. RT A and B seeded nanotubes had similar average lengths. Good control of nanotube nucleation by seeds could also be achieved for range of tile concentrations (Supp. Fig. S12).


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Figure 2: Seeds can cap exposed nanotube ends, halting their growth. Cy3 (green), Atto 647N (red) and Atto 488 (blue) dyes correspond to DNA nanotube tiles, seeds and caps respectively. Scale bars in a-c are 4 µm. Fluorescence micrographs of RS tile nanotubes grown (a) with no seeds (b) with RT A seeds. (c) Nanotubes grown from RT A seeds to which rigid caps were added at 4 hours. (d) The mean lengths of seeded nanotubes stopped increasing after caps were added. (e-f) Distributions of seeded nanotube lengths after different growth times when (e) no caps were added and (f) when caps were added after 4 hours of growth. (g) AFM image of a seeded capped nanotube. Nanotubes open on the surface due to adhesion forces,43 while DNA origami seeds and caps remain intact. (h) Relative numbers of nanotubes with attached seeds and/or caps or neither (unseeded nanotubes) when seeds were added at the beginning of growth and rigid caps were added after 4 hours. Error bars here and elsewhere show 95% confidence intervals and are determined using bootstrapping.

Nanotube seeds can cap growing nanotube ends. Because seeds present sticky ends in a geometry that emulates a complete nanotube facet, we hypothesized that a seed with sticky ends complementary to a nanotube’s growing end could bind to the growing end and prevent further growth. To test this hypothesis, we characterized how RT B seeds would interact with a population of existing nanotube ends grown from RT A seeds.


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We grew nanotubes from 2 pM RT A seeds, and after either 4 or 8 hours, added rigid caps (RT B seeds) or buffer so that the final tile concentration was 45 nM and caps (where added) were present at 10 pM (Supp. Note S12). We measured the density and length of the resulting nanotubes and whether each nanotube had a seed and/or cap using 3-color fluorescence micrographs of reaction aliquots adhered to a microscope slide taken after 4.5, 8.5, 25, 32 and 50 hour incubation times at 20 ºC (Fig. 2b, 2c, Supp. Fig. S13). After 25 hours, at least 85% of seeded nanotubes had caps bound at their growing ends. While seeded nanotubes continued to grow for at least 50 hours in the mixture containing RT A seeds and tiles but no caps, the average length of seeded nanotubes plateaued soon after caps were added in the other mixtures (Fig. 2d, Supp. Figs. S14-S15), suggesting that caps bind to the growing ends of the seeded nanotubes and prevent further nanotube growth. Further, the length distribution of seeded nanotube stops changing significantly after caps were added, consistent with the idea that a cap binds irreversibly to a nanotube end and stops further growth there (Fig. 2e-f, Supp. Fig. S16). At the end of the experiment, the length distribution of seeded nanotubes to which caps were added was significantly more monodisperse than the length distribution of seeded nanotubes to which caps were not added. Finally, atomic force micrographs of nanotubes after seeds and caps were added (See Methods) showed that nanotubes had origami structures with orientations consistent with their binding to the tiles via sticky ends (Fig. 2g). However, while rigid caps effectively stopped nanotube growth, caps that did not bind to nanotube ends often acted as seeds that nucleated the growth of new nanotubes. Because a significant excess of caps was added to the solution to maximize capping yield, after about 25 hours, the majority of the nanotubes in the solution were those that were nucleated by the rigid 9 ACS Paragon Plus Environment

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caps (Fig. 2h, Supp. Fig. S15), suggesting that the use of rigid caps to terminate nanotube growth also produces a large number of undesired side products. At even higher concentrations of rigid caps, there was almost no improvement in the capping yield but the number of nanotubes that have only rigid caps increased significantly (Supp. Fig. S17).

Nanotube seeds missing some or all staple strands function poorly as nanotube nucleation sites. Given the propensity of rigid caps to also nucleate new nanostructures, we asked whether it was possible to assemble a structure that could act as a cap but would otherwise not serve as a nanotube nucleation site. Seeds are designed to emulate the structure of the facet of a growing nanotube by providing a rigid arrangement of sticky ends in the shape of a nanotube. Tiles can attach to this structure by two sticky ends. Many such tile attachments can form a nanotube via additional growth steps. Capping, by contrast, is a reaction between just two species: a nanotube end and a cap. To prevent further nanotube growth, however, the cap must inactivate many sticky ends on the growth end, through hybridization or other means. These observations suggest that a structure that presents many sticky end binding sites arranged in such a way as to make it unfavorable for individual tiles to attach them could act as cap but not as a nanotube nucleation site. We identified two potential modifications to seeds that could produce such a structure. First, a structure that presented sticky ends on a flexible scaffold strand could attach to growing ends by many sticky ends simultaneously, but the attachment of a single tile to the structure only two sticky ends would be unfavorable due to a large loop entropy penalty.46 Second, if only every other adapter strand binding site on the seed were present, no single tile could attach to the


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structure at two adjacent sticky end sites but the set of binding sites on the cap might together be able to bind strongly to an existing nanotube end. We designed a library of modified RT B seeds with these modifications to determine whether any would function as caps that would also not nucleate new nanotubes. The original RT B seed structure and the M13 scaffold strand alone (which should have no nucleation or capping activity) were included as controls (Fig. 3).

Figure 3: The capacity of each of a library of capping structures to nucleate and cap nanotubes. Crossover strand diagrams showing the presence or absence of staples (red strands) and adapters for a library of 8 potential capping structures. Some staples span the top and bottom helices (as shown by curved lines spanning the structure) to form a cylinder. The percentage of each type of cap that nucleated nanotubes after 25 hours when no other seeds were present was measured as a metric for the propensity of structures to act as nanotube nuclei. 2 pM of each structure was used in these experiments. The propensity of each structure to act as a cap was measured using the percentage of nanotubes nucleated by 2 pM RT A seeds that were capped by an excess of caps (10 pM) added after 8 hours. The fraction capped was measured 17 hours after caps were added, after 25 total hours of growth. We tested the propensity of each structure to nucleate nanotubes by using each filter-purified structure (Supp. Notes S9 and S10) as a nanotube seed (Supp. Note S11) and measuring the percentage of each structure that had nucleated nanotubes after 25 hours. 11 ACS Paragon Plus Environment

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Other than the original RT B seed (the Rigid Cap), none of the structures in the library were effective at nucleating nanotubes (Fig. 3). Both increasing the flexibility of structures by removing staples and reducing the number of binding sites for tiles by removing adapter strands sharply decreased nucleation yield. These results suggest that it is important for nucleation templates for DNA nanostructure to both present a large set of monomer binding sites and to present these binding sites on a rigid scaffold that favors the growth of the desired structure. Flexible caps that present tile binding sites on a single-stranded scaffold bind to nanotube ends and stop nanotube growth but cannot nucleate new nanotubes. Our next goal was to determine whether the modified RT B seeds (caps) that could no longer effectively nucleate nanotubes were still able to bind to nanotube ends and prevent further growth. We grew nanotubes from 2 pM of RT A seeds and added each of the structures in the library to one of the nanotube mixtures after 8 hours to final concentration of 10 pM (Supp. Note S12). After 25 hours we measured the percentage of RT A seeded nanotubes with caps attached to their ends. All the cap structures except the structure with neither staples nor adapters were able to bind to and cap the RT A seeded nanotubes with significant yields. All the structures that presented all six pairs of sticky end binding sites (3, 4, 5 and 7) but were missing staples were as effective as the rigid caps at binding to nanotube ends, even when the capping structures contained no staples at all. The structure with no staples, 7, is the scaffold DNA with binding sites presented at sites separated by 192 bp (mostly likely with some folded secondary structure in these separating regions). Cap structures missing half the binding sites (2 and 6) capped nanotubes at significant but lower yields than the cap structures where all the 12 binding sites for sticky ends were present. The reduced number of sticky ends presented by the structure likely reduced the binding energy of the capping structures to nanotube ends, and the reduced binding energy may be the 12 ACS Paragon Plus Environment

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cause of the reduced capping yields. The control cap structure, the M13 scaffold strand alone, which has neither staples nor adapter strands (structure 8 in Fig. 3), capped only a negligible fraction of nanotubes, potentially because of some complementarity between scaffold regions and the sticky ends available on the B interface of the RS tiles (Supp. Note S18). We next measured how often caps missing many staples nucleated new nanotubes when they were added to a solution of nanotubes nucleated by RT A seeds. We found that less than 2% of structures 4, 6 and 7 nucleated nanotubes when they were added after 8 hours of growth at a 5fold excess to RT A seeds (Supp. Fig. S18). Thus, each of these structures can bind to nanotube ends at high yield while being largely unable to nucleate new ones. These results and our studies of nanotube nucleation suggest general design principles for complexes that nucleate or terminate biomolecular assemblies. For a high nucleation yield, a structure should have all the binding sites arranged rigidly such that monomers can bind favorably to the structure while capping or termination requires only that enough binding sites be present to attach to allow the components of the growing facet. The above experiments measured how well different potential capping structures bound to nanotube ends, but did not determine whether this binding stopped nanotube growth. We selected structure 7, which presents 12 sticky ends on an M13 scaffold that is not folded by staples (although secondary structure within the scaffold may affect the structure), to determine how much nanotubes grow after these structures are added and bind to nanotube ends. We termed this structure a “flexible cap” as the structure allows sticky end pairs to access a much broader set of geometrical configurations than the rigid cap does. We grew nanotubes from RT A seeds, then added flexible caps after 4 and 8 hours (Supp. Note S12) and measured how nanotube length changed over time. 13 ACS Paragon Plus Environment

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Figure 4: Flexible caps bind to nanotube ends and stop their growth without nucleating new nanotubes. (a) The mean length of seeded nanotubes did not change significantly after either flexible or rigid caps were added to the reaction solution. (b-c) The relative percentages of nanotubes of the four different types of RS tile nanotubes present after 4.5 and 50 hours of growth when (b) rigid caps and (c) flexible B caps were added after 4 hours of growth. Excess rigid caps nucleate large numbers of new nanotubes, while the same excess of flexible caps did not. (d) The density of different types of nanotubes over time after flexible caps were added after 4 hours. (e) Length distributions of seeded nanotubes and (f) Micrographs of seeded nanotubes to which flexible caps were added after 4 hours. Scale bars are 4 µm.

Flexible caps were as effective as rigid caps at stopping the growth of seeded nanotubes (Fig. 4a, Supp. Figs. S19-S20). And while reaction mixtures to which rigid caps were added contained many nanotubes that were nucleated by excess rigid caps (Fig. 4a-b, Supp. Fig. S21), few new nanotubes formed in reactions where flexible caps were used (Fig. 4c-d, Supp. Figs. S21-S22). Seeded nanotubes in solutions where flexible caps were added were also fairly monodisperse in length (Fig. 4e, Supp. Fig. S23). The flexible cap thus functions reliably as a nanotube cap but does not otherwise observably affect the self-assembly process. While the binding of a flexible 14 ACS Paragon Plus Environment

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cap to a growing nanotube should be essentially irreversible under the conditions studied, the nucleation of nanotubes by flexible caps should be very slow in comparison to RT A or RT B seeds (rigid caps) because of the entropic cost of organizing the flexible cap’s binding sites into a tube-shaped facet (see Supp. Note S20). To understand how to use caps within controlled, dynamic assembly processes, we also measured the rate of nanotube growth and the rate at which flexible caps bind to growing nanotube ends. We attached nanotube seeds to a passivated glass surface using biotinstreptavidin linkers (Supp. Note S13) and used time-lapse fluorescence microscopy to track the state of each of a set of nanotubes throughout a growth or capping process (Supp. Notes S14-15). Under these conditions, slightly different concentrations of tiles were needed to achieve reliable growth and capping, perhaps because of a difference in reagents used to minimize nonspecific adsorption of reaction components and differences that arise because of presence of the dish, rather than Eppendorf tube, surface (Supp. Note S14). At 75 nM of tile monomers, we observed that nanotubes grew at an average rate of 0.165 ± 0.0052 µm/hr (Supp. Figs. S24-S25 and Movie S1), similar to earlier measurements made in other systems. We measured capping by tracking the rate at which caps attached to seeded nanotubes on the surface (Fig. 5, Supp. Figs. S26-S27 and Movie S2) and fitting the fraction of capped nanotubes to find a reaction rate of kj=1.85 × 106 M-1s-1 (Supp. Note S19). This rate is similar to the measured forward rate of tile attachment and of origami hybridization, suggesting that the kinetics of capping are controlled by multivalent hybridization of sticky ends in similar fashion.47 Capping takes a long time to complete because the concentrations of both nanotube ends and caps are both low.


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Figure 5: The fraction of nanotubes that were capped over time characterized using timelapse microscopy following individual seeded nanotubes. Individual A seeded, RS tile nanotubes were attached to a passivated glass dish in a solution of 24 pM caps and 75 nM tiles (Supp. Note S15). The measured fraction of capped nanotubes was used to fit a binding constant between tubes and caps. A 95% confidence interval, determined by bootstrapping, is shown in the shaded light blue region. N=301. Flexible caps selectively bind to and stop the growth only of nanotubes presenting complementary sticky ends. Caps are designed to present nanotube sticky end sites and therefore should bind selectively only to nanotube sticky ends. Sets of caps that each bind only their specific nanotube type could be instrumental in constructing complex networks and structures containing multiple types of nanotube filaments. To ask whether caps selectively bind nanotubes with complementary sticky ends, we designed a new seed (termed C) for tiles with different sequences, termed UV tiles (Supp. Notes S2 and S4).43 Sequence-specific binding of RS and UV tiles ensure that RT A nucleate only RS tile nanotubes and RT C nucleate only UV tile nanotubes.50 We next designed a set of adapters for a flexible cap designed to attach to the growing end of UV tile nanotubes (Supp. Note S7). The flexible caps with these adapters were termed D caps. To test that the binding of B and D caps was selective for nanotube type, we grew RS and UV nanotubes from 2 pM RT A and RT C seeds in four tubes. To the four tubes, we added either no caps, 10 pM B caps, 10 pM D caps or 10 pM of both B and D caps after 8 hours of growth 16 ACS Paragon Plus Environment

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(Supp. Notes S16-S17). We used fluorescence images to determine where binding occurred. UV tiles were labeled so that their fluorescence intensity would be only 25% of the RS tubes, making it possible to distinguish the two types of tubes, and B and D caps were labeled with different combinations of fluorescent dyes so that they could also be distinguished visually. We measured the lengths of the resulting nanotubes and determined whether each nanotube had a seed and/or which cap using 3-color fluorescence micrographs of reaction aliquots adhered to a microscope slide taken after 8.5, 25 and 50 hours of growth (Fig. 6a-d, Supp. Note S28).

Figure 6: Flexible caps bind selectively to nanotubes end with complementary sticky ends. RS tile nanotubes (Cy3 bright green) grew from RT A seeds labeled with Atto 647N (red) and UV tile nanotubes (Cy3 dim green at 25% incorporation) grew from RT C seeds labeled with Atto 647N (red). Flexible cap B was labeled with Atto 488 (blue) and flexible cap D was labeled with 50% Atto 647N and 50% Atto 488 dyes (purple). Example fluorescence micrographs of nanotubes grown in the presence of RT A and C seeds to which (a) buffer was added, (b) flexible B caps and (c) flexible D caps and (d) flexible B and D caps were added at 8 hours. Yellow arrows identify RT C seeded UV nanotubes. (e) RS tile nanotubes stopped growing after B caps are added at 8 hours but UV tile nanotubes did not. (f) UV tile nanotubes stopped growing after D caps were added at 8 hours but RS tile nanotubes did not. (g) Fractions of seeded RS and UV tile nanotubes that were capped by B seeds (AB vs CB) after 25 hours. Caps were added at 8 hours. (h) Fractions of seeded RS and UV tile nanotubes capped by D seeds (AD vs CD) after 25 hours. Caps are added at 8 hours. (i) Fractions of seeded RS and UV tile nanotubes capped by the two types of caps when both B and D caps are added at 8 hours. Scale bars are 4 µm. 17 ACS Paragon Plus Environment

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The average lengths of RT A and C seeded nanotubes plateaued only when their respective flexible caps were present, suggesting that each cap bound only to nanotubes with the complementary growing end to prevent further nanotube growth (Fig. 6e-f, Supp. Fig. S29). When both B and C caps were added, 86 ± 3.6% of RT A seeded nanotubes and 83 ± 3% of RT C seeded nanotubes had B and D caps bound respectively at their growing ends. We observed almost no RT C seeded nanotube with B caps at their ends or RT A seeded nanotube with D caps at their ends (Fig. 6g-i). Thus, the flexible caps selectively terminate the growth of seeded nanotubes. CONCLUSION In this paper we demonstrate how nanostructures can bind to the facets of DNA nanotubes and terminate their growth selectively. Such structures could be used to arrest assembly reactions far from equilibrium such that growth rates and the time over which growth occurs can be independently controlled. The ability to use biomolecules or complexes to controllably stop the growth of filaments is also an essential aspect of many controlled dynamic growth processes in biological systems, such as the assembly of actin-Arp 2/3 networks that can induce motion of cells,48 suggesting that controlling the termination of nanotubes might be an important step toward designing similar synthetic systems, along with nanotube assembly,38 and the formation of interconnects39 and architectures.49, 50 It is straightforward to observe how the structures we have designed could be activated or deactivated by strand displacement reactions and thus controlled by molecular circuits that regulate assembly and reorganization, further expanding the capacity of the resulting dynamic materials. 18 ACS Paragon Plus Environment

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More generally, the systematic approach taken to understanding the capping process in this work elucidates the design principles for assembling structures that terminate the growth of onedimensional nanotubes and other related structures. The use of DNA nanostructures in our investigation made it possible to precisely characterize the relative importance of multivalency and control over the arrangement of binding sites for the functionality of a capping structure. Flexible linkers between binding sites allow a structure to bind to a growing nanotube end but not interact significantly with unassembled monomers enough to grow new structures, whereas structures with enough sticky ends to bind to only some portion of the facet function less effectively, even if the presented sticky ends are arranged to favor binding. These insights may be valuable for designing structures that direct a variety of crystallization processes, including the assembly of DNA-functionalized nanoparticles51,

52

and colloids53 as well as protein

structures,54, 55 carbon nanotubes, metallic and semiconducting one-dimensional nanostructures,56, 57

and for directing biomineralization.58 ----

MATERIALS AND METHODS DNA nanotube, seed and cap components: The sequences of RT tiles, adapter, seeds and caps used in this study are listed in the supporting information. DNA tile, adapter, staple, dye strands, dye attachment strands, biotin attachment and biotin attachment linker strands were synthesized by Integrated DNA Technologies, Inc. M13mp18 was purchased from Bayou Biolabs. Glass bottom dishes (µ-Dish 35mm, high Grid-50 Glass Bottom) were purchased from Ibidi. BiotinPEG-Silane (Biotin-PEG-SIL-3400-500mg) was purchased from Layson Bio, and Neutravidin (31000), Streptavidin (21122) and BSA (Bovine Serum Albumin) were purchased from Thermo Fisher Scientific. Adapter strands with sticky ends and tile strands without fluorescent labels 19 ACS Paragon Plus Environment

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were PAGE purified. Tile and origami strands with fluorescent labels were HPLC purified. All other strands were used directly after desalting. Sequences for seeds and tiles are given in Supp. Notes S1-S4. Sequences for caps are in Supp. Notes S5 and S7 and Fig. S7. RT RS tiles were labeled with Cy3 fluorescent dye to allow fluorescence imaging of nanotubes while RT UV tiles were labeled with 25% Cy3 fluorescent dye, making it possible to distinguish RS tile nanotubes from UV tile nanotubes in the same image (Fig. 6a-d, Supp. Fig. S28). RT A seeds and RT B seeds (or caps) in experiments characterizing their yields when used were labeled with Atto 647N dye and Atto 488 dye respectively (Supp. Note S6). RT C seeds and flexible D caps in experiments characterizing their yields when used were labeled with Atto 647N dye, and a mixture of 50% Atto 647N 50% Atto 488 dyes respectively (Supp. Notes S6,S17). All seed and cap structures used a common set of strands for labeling that attach to unfolded regions of the scaffold strand and present binding sites for a fluorescent strand (Supp. Note S6). All samples were prepared in TAE Mg2+ buffer (40 mM Tris-Acetate, 1 mM EDTA to which 12.5 mM magnesium acetate was added). In experiments with RS tiles performed in Eppendorf tubes, the strands for each tile were present at 45 nM except for the strands presenting sticky ends, which were present at 90 nM to minimize the concentration of malformed tiles (Supp. Note S8). In experiments with RS tiles performed in glass-bottom dishes, the strands for each tile were present at 75 nM except for the strands presenting sticky ends, which were present at 150 nM to minimize the concentration of malformed tiles (Supp. Notes S14-S15). In experiments with UV tiles, the strands for each tile were present at 30 nM except for the strands presenting sticky ends, which were present at 60 nM. Adapter strands presenting sticky end sequences were analogously included at a 2-fold excess over other adapters. Concentrations of tile and adapter strands were


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determined using 260 nm absorbance spectroscopy while the concentrations of staple, attachment and labeling strands concentration were assumed to be those provided by IDT.

Sample preparation: To grow RT A or C seeded nanotubes, 19.7 µL of a tile solution was annealed from 90 ºC to 20 ºC and then 0.3 µL of a solution of purified seeds was added after the solution reached 20 ºC (Supp. Note S11) so that either the RS tile concentration was 45 nM or the UV tile concentration was 30 nM after addition. To grow RT A and C seeded nanotubes in a single pot reaction, 19.7 µL of RS and UV tile solution was annealed from 90 ºC to 20 ºC and then 0.3 µL of a solution of purified seeds was added after the solution reached 20 ºC so that the RS and UV tile concentration were 45 nM and 30 nM respectively after addition (Supp. Note S16). To grow RT A seeded B capped nanotubes, 19.4 µL of a tile solution was annealed from 90 ºC to 20 ºC and then 0.3 µL of a solution of purified seeds was added after the solution reached 20 ºC. To measure the growth rate of seeded nanotubes, purified RT A seeds with linker and attachment strands in 1 × TAE Mg2+ buffer were added to glass bottomed dishes (Supp. Note S14) where the surface prepared as described in Supp. Note S13. This was followed by 3 washes of 1 × TAE Mg2+ buffer to remove the unattached seeds and then 1000 µL of 75 nM of the RS tile mixture were placed into the dish (Supp. Note S14). At each time point (0, 6, 12 and 24 hours) before imaging, the tile mixture in the dish was replaced by 1000 µL of 1 × TAE Mg2+ buffer and once the imaging was over, the same tile mixture was placed back into the dish. Upon each transfer, the dish was sealed with Parafilm to prevent solution evaporation. To measure the capping rate, 19.7 µL of RS tile solution was annealed from 90 ºC to 20 ºC and then 0.3 µL of a solution of purified full RT A seeds with linker and attachment strands was added into the tile mixture so that the final tile and seed concentration will be 75 nM and 2 pM respectively after 21 ACS Paragon Plus Environment

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the addition (Supp. Note S15). Solutions were incubated at 20 ºC in an Eppendorf Mastercycler for 6 hours before transferring into the glass surface made using Supp. Note S13. This was followed by 3 washes of 1 × TAE Mg2+ buffer to remove the unattached seeded nanotubes and then 985 µL of 1 × TAE Mg2+ buffer and 15 µL of the purified flexible B caps (Supp. Notes S9S10) were added so that the cap concentration will be 24 pM after the addition. The dish was then sealed with Parafilm to prevent solution evaporation (Supp. Note S15). Experiments with glass-bottomed dishes where the seed is anchored used the staples for the seed from Mohammed et al. (see Supp. Note S14).38 To grow RT A seeded B capped and RT C seeded D capped nanotubes in a single pot reaction, 19.4 µL of RS and UV tile solution was annealed from 90 ºC to 20 ºC and then 0.3 µL of a solution of purified A and C seeds was added after the solution reached 20 ºC. Flexible B and/or D caps were added at the times described in the text by adding 0.3 µL of a purified cap solution to the resulting mixture (Supp. Note S17) so that the RS and UV tile concentration was 45 nM and 30 nM respectively after addition of seeds and caps. Solutions were incubated at 20 ºC in an Eppendorf Mastercycler. Seed and cap preparation: Seeds and caps were prepared by annealing a solution of 5 nM M13mp18 scaffold, 500 nM of each staple strand, 100 nM adapter strand mix (See Supp. Note 8), 25 nM of each attachment strand and 5000 nM of each labeling strand in TAE Mg2+ buffer and 0.05 mg/ml BSA biotin (A8549, Sigma-Aldrich Co.) to prevent surface absorption in PCR tubes from 90 ºC to 20 ºC using an Eppendorf Mastercycler (Supp. Note S9). After annealing, 50 μL of the seed or cap solution and 350 μL of 1 × TAE Mg2+ buffer was added to a 100 kDA Amicon Ultra-0.5 mL Centrifugal Filter (UFC510096) and centrifuged at 2,000 RCF for 4 min in a fixed angle centrifuge to remove excess staple and adapter strands. The samples were washed three more times by adding 200 μL of 1 × TAE Mg2+ buffer in remaining solution and repeating 22 ACS Paragon Plus Environment

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centrifugation. The remaining solution was recovered by spinning the filter inverted in a fresh tube and the purified mixture was stored at 4 ºC until use. The concentration of seeds or caps in the resulting solution was determined using a calibration method based on fluorescence imaging of different concentrations of structures before and after purification (Supp. Note S10). Fluorescence microscopy: For every time point at which fluorescence microscopy images of nanotubes were analyzed, 2 slides were prepared by adding 6 μL of reaction solution to 2 cover slips and inverting them onto glass slides. We then captured 2-3 images of nanotubes on each slide at randomly chosen locations. Background caused by adsorption of free tiles was reduced by adding 0.3 μL of a solution of 1 μM of 54 bp DNA strand (D01) with a sequence that did not interact with tiles, staples or adapters to each 20 µL sample (Supp. Note S11). This strand acted as a competitor for DNA tile-glass binding. During the experiments performed in glass-bottomed dishes, custom time-lapse image acquisition software was used to collect images at four specific locations, which were selected randomly, over the reaction period. For measurements of nanotube growth rates, 10 200 ms exposure images of both the Cy3 (RS tile nanotubes) and Atto 647N (RT A seed) channels were collected (Supp. Note S14). For the capping rate measurement, once the caps were added into the dish, one image of Atto 647N, three images Atto 488 (flexible B cap) and three images of Cy3 were collected. every ten minutes over 4 hours (Supp. Note S15). Samples were imaged on an inverted microscope (Olympus IX71) using a 60X/1.45 NA oil immersion objective with a cooled CCD camera (iXon3, Andor). Measurement of nanotube length and capping yield: Multicolor images of nanotubes, seeds and caps were prepared by overlaying images of Atto 647, Atto 488 and Cy3 fluorescence that were flattened beforehand using histogram equalization. To measure nanotube lengths, Cy3 channel images were first converted into binary form at a manually determined brightness 23 ACS Paragon Plus Environment

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threshold. The structures were then thinned using the bwmorph Matlab function that reduced the binary objects (nanotubes) into single pixel-thick curves. The length of these curves was then measured using Matlab and converted into microns using the pixel to micron relation (1 pixel = 0.17 μm). The small number of nanotubes that overlapped one another or were at the edges of images were not included in analysis. To measure the lengths of RT A seeded nanotubes that grew in the dishes, nanotube contour length was calculated by assessing 2D projection of DNA nanotube length via ImageJ JFilament 2D (http://athena.physics.lehigh.edu/jfilament/). The measured pixel length was converted into microns using the pixel to micron relation. A nanotube was considered capped when a cap and seed were detected in the same pixel location over multiple consecutive time points. Atomic force microscopy: Seeded, capped nanotubes were prepared from a 70 nM solution of annealed tiles to which 10 pM of seeds at the start of 20ºC incubation and 20 pM of caps were added after 3 hours of growth at 20ºC, and then grown for 4 hours. BSA was not included in the solution to minimize imaging background. 5 μL of reaction solution, then 200 μL of TAE Mg2+ buffer was added onto a freshly cleaved mica surface mounted on a puck with a Teflon sheet. The sample was imaged on a Dimension Icon (Bruker) using Scanasyst mode and Sharp Nitride Lever tip (SNL - 10 C, Bruker) cantilevers. Nanoscope Analysis software was used to flatten the image.

ASSOCIATED CONTENT


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Supporting Information: The supporting information contains sequences for DNA molecules used in our experiments, additional experimental results, and ancillary notes referred to in the paper. This material is available free of charge via the Internet at http://pubs.acs.org AUTHOR INFORMATION Corresponding Author *E-mail: [email protected] Present Addresses † Present Address: Biomedical Engineering Department, Boston University, MA, 02215. † Present Address: Molecular Engineering and Sciences Institute, University of Washington, Seattle, WA 98105. Author Contributions Deepak K. Agrawal, Ruoyu Jiang, Seth Reinhart, Abdul M. Mohammed, Tyler D. Jorgenson and Rebecca Schulman designed the experiments. Deepak K. Agrawal, Ruoyu Jiang and Seth Reinhart conducted the experiments. All the authors discussed the results and Deepak K. Agrawal and Rebecca Schulman wrote the manuscript. Funding Sources This research was supported by DOE grant SC-0010595. ACKNOWLEDGMENT


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56. Stadermann, M.; Sherlock, S. P.; In, J.-B.; Fornasiero, F.; Park, H. G.; Artyukhin, A. B.; Wang, Y.; De Yoreo, J. J.; Grigoropoulos, C. P.; Bakajin, O., Mechanism and Kinetics of Growth Termination in Controlled Chemical Vapor Deposition Growth of Multiwall Carbon Nanotube Arrays. Nano Lett. 2009, 9, 738-744. 57. Huang, Y.; Duan, X.; Wei, Q.; Lieber, C. M., Directed Assembly of One-Dimensional Nanostructures into Functional Networks. Science 2001, 291, 630-633. 58. Weiner, S.; Dove, P. M., An Overview of Biomineralization Processes and the Problem of the Vital Effect. Rev. Mineral. Geochem. 2003, 54, 1-29.

TABLE OF CONTENTS GRAPHIC 33 ACS Paragon Plus Environment

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Terminating DNA Tile Assembly with Nanostructured Caps - ACS

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