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Reversible metamorphosis of hierarchical DNA–inorganic crystals

Nature Nanotechnology volume 20, pages 1813–1821 (2025) Cite this article

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Abstract

Living systems transform their shapes via reversible formation of macromolecular structural complexes, leading to deformations at localized sites. Here we report DNA–inorganic flower-shaped crystals with inscribed deformation modes that enable flowers to shrink and bend reversibly. Template-independent DNA polymerization of pH-responsive and inert blocks tune the hierarchical assembly and spatial localization of DNA within flowers. Experiments and simulations demonstrate that reversible, pH-triggered folding of intraflower DNA strands drives reconfiguration of flowers. By contrast, the subflower localization of these contractile DNA motifs dictates the mode of shape change. As microscale flowers close and open, their nanoscale crystal organization changes reversibly, suggesting that mechanical metamorphosis of flowers is transduced across multiple organizational length scales. The adaptability of flowers to environmental changes activates cascaded biocatalytic reactions and reveals gel-encrypted information. Further variation of the DNA polymer sequence, its subcrystal localization and its reversible folding advances a new class of organic–inorganic shape-shifters.

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Fig. 1: One-pot fabrication of DNA–inorganic crystals via TdT enzymatic DNA polymerization.
The alternative text for this image may have been generated using AI.
Fig. 2: pH-fuelled reversible contraction and swelling of C-DNA flowers.
The alternative text for this image may have been generated using AI.
Fig. 3: Metamorphosis of diblock DNA–inorganic crystals.
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Fig. 4: Programming various actuation mechanisms within modular DNA–inorganic crystals.
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Fig. 5: Transducing shape-shifting modes of DNA–inorganic actuators with coupled biochemical reactions.
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Data availability

All primary data supporting the findings of this study are included in the Article and its Supplementary Information. The simulation code is available via Zenodo at https://doi.org/10.5281/zenodo.16730053 (ref. 48). For additional images or specific details, contact the corresponding author. Source data are provided with this paper.

Code availability

The simulation code is available via Zenodo at https://doi.org/10.5281/zenodo.16730053 (ref. 48).

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Acknowledgements

We are thankful for funding support for this work from the Alfred P. Sloan Foundation, Matter-to-Life Grant G-2021-14197. R.F. acknowledges financial support in the form of a Cottrell Scholar Award (CS-CSA-2023-033), sponsored by Research Corporation for Science Advancement. SEM, TEM, STEM–EDS, powder X-ray diffraction and atomic force microscopy were performed at the Chapel Hill Analytical and Nanofabrication Laboratory (CHANL), a member of the North Carolina Research Triangle Nanotechnology Network (RTNN), which is supported by the National Science Foundation, Grant ECCS-2025064, as part of the National Nanotechnology Coordinated Infrastructure (NNCI). Confocal microscopy and Imaris analysis were performed at the UNC Hooker Imaging Core Facility (HIC), which is supported in part by the National Cancer Institute of the National Institute of Health, Cancer Center Support Grant P30CA016086 to the UNC Lineberger Comprehensive Center. CD was performed at the UNC Macromolecular Interactions Facility (Mac-In-Fac), which is supported by the National Cancer Institute of the National Institute of Health, Grant P30CA016086. We thank J. Hill for the preparation of the graphic illustrations shown in the figures and movies. We thank M. J. Papanikolas for the discussions and technical assistance.

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Authors and Affiliations

  1. Department of Applied Physical Sciences, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA

    Yuan Gao, Wenzheng Shi, Stephen J. Klawa, Margaret L. Daly, Edward T. Samulski, Ehssan Nazockdast & Ronit Freeman

  2. Courant Institute, New York University, New York, NY, USA

    Wenzheng Shi

  3. Department of Chemistry, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA

    Edward T. Samulski

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Contributions

The manuscript was written with contributions and approval from all authors. Y.G. and R.F. conceived the project. R.F. acquired funding and executed project administration. R.F. and E.N. supervised the work. Y.G., W.S., S.J.K., M.L.D., E.T.S., E.N. and R.F. contributed to methodology and visualization of the work. Y.G., W.S., S.J.K., M.L.D., E.N. and R.F. contributed to investigation and writing the original draft. R.F., E.N. and E.T.S. reviewed and edited the final draft.

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Correspondence to Ronit Freeman.

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Nature Nanotechnology thanks Sergii Rudiuk, Ruojie Sha and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.

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Extended data

Extended Data Fig. 1 Structural phase of DNA-inorganic crystals as a function of DNA base, polymerization time, and DNA length.

(a) Representative images of polyN (mixed bases), polyA, polyC, and polyT DNA-inorganic crystals at ranging polymerization times (1, 2, 4, 8, 12, 24 hr) and their respective DNA lengths. [Blue shading indicates flower-shaped crystals shown in main Fig. 1; unshaded area indicates immature crystals]. Scale bars, 2 μm, and identical for images in the same column. Flowers emerge when the DNA strands lengthen to 650 bases, (see magnified SEM images in b, scale bars, 1 μm) and form in a kinetics that follow T>C>A>N, matching the kinetics of enzyme polymerization (shown in c). (c) Agarose gel electrophoresis of polyT, polyC, polyA and polyN. The primers/initiators and monomers ratio is I:M = 1:1000. (d) Control crystal growth experiments show that purley inorganic Co2PPi crystals (I-II) grow after 24 hours upon mixing Na4PPi and CoCl2 (1000 μM), while DNA-templated inorganic crystals emerge at as early as 1 hr with poly T (in a). Mixing pre-polymerized DNA (>650 bases) with cobalt and pyrophosphate (III-IV) accelerates flower formation, but not as much as in situ DNA polymerization and hybrid DNA-inorganic crystallization. Scale bars, 2 μm.

Source data

Extended Data Fig. 2 i-motif formation in C flowers.

Circular dichroism spectra of C flowers (solid lines) compared to T flowers (dashed lines) at (a) varying pH and (b) after 24 hr polymerization. C(8) at pH 7 (light green) shows random coil with a positive band at 275 nm, while C(24) at pH 7 (dark green) shows i-motif signatures of a positive band at 286 nm and negative band at 265 nm signal, suggesting that C flowers show more i-motif folding as the polymerization progresses. However, switching the C(8) from pH 7 to pH 5 shows a transition to i-motif. T(2) to T(24) shows a shift from a positive peak at 273 nm to 278 nm and a negative peak from 248 nm to 252 nm, suggesting a stronger B-form-like character as T polymerization progresses. Moreover, switching T(2) from pH 7 to pH 5 continues to show B-form-like bands. (c) Schematics of a single-stranded polyC with 650 bases at neutral pH with an estimated length of ~420 nm, calculated using 0.63-0.67 nm per base30,31,32,33. At acidic pH, if we estimate that 10 bases of C form an i-motif with stretches of 10 unpaired bases in between, and using ~0.83 nm per width of i-motif and ~0.63-0.67 nm per base in the 10-base ssDNA stretches, the estimated folded i-motif strand for 650 bases is ~240 nm. Insets show schematic representations of fully formed and partially folded i-motif structures and the corresponding chemical structure of the protonated base-pairing of cytosine bases.

Source data

Extended Data Fig. 3 Catalytic activity of flower-confined enzyme cascades.

Enzymatic curves from GOx- and HRP-loaded TC (a) and CT (b) flowers compared to free GOx and HRP in solution. Reactions were run in 1X PBS with 0.5 nM GOX, 0.5 nM HRP, 10 mM glucose, and 2 mM ABTS. DNA flowers at 0.05 nM were used. (c) Enzymatic rates of the catalytic cascade within flowers (and controls). Enzymatic rates were calculated from the first 30 min (green bars and points) and 30-60 min after switching to pH 5 (red bars and points) or keeping the sample at pH 7 (white points) (all ****p < 0.0001). Monitoring the enzymatic efficiency of these constructs at neutral pH (flowers open) showed that the T(GOx)C(HRP) flowers had the highest rate among combinations/cases [free enzymes, free flowers, the mixture of free flowers and enzymes, enzymes encapsulated in T-flowers “T(GOx+HRP)”, enzymes encapsulated in C- flowers “C(GOx+HRP)”, the mixture of T(GOx+HRP) and C(GOx+HRP) flowers, etc]. The on/off switch of the cascade reaction was confirmed by monitoring the efficiency at acidic pH (flower closed). Bars show the mean of n = 4 reactions per sample, and statistics were calculated with an ordinary two-way ANOVA in GraphPad Prism. (d) The fold change in reaction rates during reaction cycles for the catalytic cascade (and the control). To determine the fold change of reaction rate, the rate of each “ON” was compared to the “ON” rate of free enzymes in solution for that cycle (the mixture of free enzymes GOx+HRP). Interestingly, reversing the order of the DNA layers [C(GOx)T(HRP) flowers] did not result in a sizeable improvement when undergoing pH cycles. This highlights the crucial role of bending versus shrinking in facilitating the enzymatic cascade reactions and providing switchable activity. n = 4 separate assays taken from one batch of flowers.

Source data

Supplementary information

Supplementary Information (download PDF )

Supplementary Discussion 1, Supplementary Figs. 1–51, Supplementary Tables 1–7, captions for Supplementary Videos 1–10 and references.

Supplementary Video 1 (download MOV )

pH powered contraction of C-DNA flowers. Time-resolved confocal microscopy of multiple C(8) flowers upon treatment withsequential pH 5 (to close and pH 7 (to open) cycles. Flowers were fabricated with FAM-labeled dCTP (2 mol%) during polyC polymerization (green channel).

Supplementary Video 2 (download MOV )

High-resolution time-lapse of contracting C-DNA flowers. Time-resolved confocal microscopy of 2 representative C(8) flowers upon treatment with sequential pH 5 (to close) and pH 7 (to open) cycles. Timelapse was collected with Airyscan. Flowers were fabricated with FAM-labeled dCTP (2 mol%) during polyC polymerization. Each flower is shown as merged two-channel (green and red) images and combined channel representations in the inferno and inverted grayscale lookup tables from FIJI.

Supplementary Video 3 (download MOV )

High-resolution time-lapse of diblock flowers T(2)C(8) and C(8)T(2). Time-resolved confocal microscopy of T(2)C(8) (left) and C(8)T(2) flowers (right) upon treatment with sequential pH 5 and pH 7 cycles. Timelapses were taken with Airyscan. Flowers were fabricated with FAM-labeled dCTP (2 mol%) during polyC polymerization (green channel) and Cy3-dUTP (2 mol%) during polyT polymerization (red channel) steps. Each flower is shown as merged two-channel (green and red) images and combined channel representations in the inferno and inverted grayscale lookup tables from FIJI.

Supplementary Video 4 (download MOV )

Scheme of C polymerization onto T flowers. Illustration of C polymerization onto pre-formed T flowers and the resulting layering within petals of TC flowers. Schematics were informed by confocal microscopy results.

Supplementary Video 5 (download MOV )

Scheme of T polymerization onto C flowers. Illustration of T polymerization onto pre-formed C flowers and the resulting layering within petals of CT flowers. Schematics were informed by confocal microscopy results.

Supplementary Video 6 (download MOV )

Shape-shifting as a function of di-block DNA polymerization time and layer thickness. Time-resolved confocal microscopy of T(2)C(1) (top left), T(2)C(4) (top right), T(2)C(8) (bottom left) and T(2)C(24) flowers (bottom right) upon treatment with sequential pH 5 and pH 7 cycles. Timelapses were taken with Airyscan. Flowers were fabricated with FAM-labeled dCTP (2 mol%) during polyC polymerization (green channel) and Cy3-dUTP (2 mol%) during polyT polymerization (red channel) steps. Each flower is shown as merged, two-channel (green and red) images and combined channel representations in inverted grayscale.

Supplementary Video 7 (download MOV )

Finite element simulations showing bending mode of (TC) flowers. A 3D model from finite element simulations of TC diblock flowers as linear isotropic elastic materials, using experimental data of flower shape and petal layer thicknesses as inputs. Color scale represents the deflection field, where orange indicates regions with minimal deflection and purple indicates maximum deflection.

Supplementary Video 8 (download MOV )

Finite element simulations showing shrinking mode of (CT) flowers. A 3D model from finite element simulations of CT diblock flowers as linear isotropic elastic materials, using experimental data of flower shape and petal layer thicknesses as inputs. Color scale represents the deflection field, where orange indicates regions with minimal deflection and purple indicates maximum deflection.

Supplementary Video 9 (download MOV )

High-resolution time-lapse of pH-fueled TCT flower. Time-resolved confocal microscopy of a T(2)C(8)T(2) flower upon treatment with sequential pH 5 and pH 7 cycles. Timelapses were taken with Airyscan. Flowers were fabricated with FAM-labeled dCTP (2 mol%) during polyC polymerization (green channel) and Cy3-dUTP (2 mol%) during polyT polymerization (red channel) steps. Each flower is shown as merged two-channel (green and red) images and combined channel representations in the inferno and inverted grayscale lookup tables from FIJI.

Supplementary Video 10 (download MOV )

Reading of encrypted information with DNA flowers ink. Video recording of the gel system with embedded enzyme-encapsulated flowers [T(GOx)C(HRP)] as the environment is adjusted to pH 5 or pH 7 over multiple cycles

Supplementary Gel (download ZIP )

Supplementary gel source data.

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Gao, Y., Shi, W., Klawa, S.J. et al. Reversible metamorphosis of hierarchical DNA–inorganic crystals. Nat. Nanotechnol. 20, 1813–1821 (2025). https://doi.org/10.1038/s41565-025-02026-8

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