www.sciencemag.org/content/351/6275/841/suppl/DC1

Supplementary Material for

DNA-controlled dynamic colloidal nanoparticle systems for mediating cellular interaction

Seiichi Ohta, Dylan Glancy, Warren C. W. Chan*

*Corresponding author. E-mail: warren.chan@utoronto.ca

Published 12 February 2016, Science 351, 841 (2016)

DOI: 10.1126/science.aad8142

This PDF file includes:

Materials and Methods Figs. S1 to S16 Full Reference List

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Materials and Methods Materials

Gold (III) chloride (HAuCl4), sodium citrate tribasic, tannic acid, potassium carbonate, Bis(p-sulfonatophenyl)phenylphosphine dihydrate dipotassium salt (BSPP), magnesium chloride (MgCl2), and fetal bovine serum (FBS) was purchased from Sigma-Aldrich (St. Louis, MO, USA). Tween 20, dithiothreitol (DTT)E and phosphate buffered saline (PBS) were purchased from BioShop Canada Inc. (Burlington, ON, Canada). Dulbecco’s modified Eagle’s medium (DMEM), SYBR® gold nucleic acid stain and Trypsin-EDTA (0.25 %) were purchased from Life Technologies (Carlsbad, CA, USA). Thiolated methoxypolyethylene glycol (PEG, MW = 5 kDa) was purchased from Laysan BIO (Arab, AL, USA). Folic acid (FA)-functionalized PEG (MW = 5 kDa) terminated with thiol was purchased from Nanocs Inc. (New York, NY, USA). Hydrochloric acid (HCl) and nitric acid were purchased from Caledon Laboratories (Georgetown, ON, Canada). All DNA sequences used in this study were synthesized, labeled, and purified by Bio Basic Inc. (Markham, ON, Canada). Gold nanoparticle synthesis and characterization

Gold nanoparticles were synthesized according to previous works (28). In brief, 13-nm gold nanoparticles were synthesized by first adding 1.0 ml (1.0%w/v) of HAuCl4 to NanopureTM water (98 ml) in a 250 ml Erlenmeyer flask. The solution was brought to a boil on a stir plate set to 350 ˚C. Then, 1.0 ml of sodium citrate tribasic (1.0 ml, 3.0 %w/v) was injected swiftly into the boiling solution under vigorous stirring. The reaction was allowed to proceed for 10 minutes, followed by cooling on ice. Obtained nanoparticles were stored at 4 ˚C prior to use.

To synthesize 3- and 6-nm nanoparticles, 1.0 ml of HAuCl4 (1.0 %w/v) was added to 80ml of NanopureTM water in a 250 ml Erlenmeyer flaks and brought to 60 ˚C on a stir plate. A reducing solution containing sodium citrate tribasic, tannic acid, and potassium carbonate was prepared according to the following table and warmed to 60 ˚C in a water bath for 50 minutes:

Nanoparticle size (nm) 3-nm 6-nm 1.0 %w/v Sodium citrate tribasic (ml) 6.0 6.0 1.0 %w/v Tannic acid (ml) 7.5 0.38 3.46 mg/ml Potassium carbonate (ml) 7.5 --- NanopureTM water (ml) 9.0 24

Under vigorous stirring, 20 ml of the reducing solution was swiftly injected into the Erlenmeyer flask and the reaction was kept at 60 ˚C for 30 minutes and then at 90 ˚C for 10 minutes. After cooling on ice, 1.0 ml of 80 mg/ml BSPP was added to the nanoparticle solutions and stirred overnight to improve particle stability. The obtained nanoparticles were washed three times by 30 minutes centrifugation using 0.01% sodium citrate aqueous solution containing 0.01% Tween 20. The centrifugation speed was 240,000 g for 3-nm and 120,000 g for 6-nm particles. The purified particles were stored at 4 ˚C prior to use.

Synthesized nanoparticles were observed by transmission electron microscopy (TEM). 2.0 μl of nanoparticle solution (ca. 1-10 nM) was dropped on TEM grids, left for 10 minutes,

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and then blotted. The prepared samples were observed by TEM at 200 kV (Techni 20, FEI, Hillsboro, OR, USA). Diameter of nanoparticles was analyzed from the obtained TEM images by using ImageJ software. Nanoparticles were also characterized by dynamic light scattering (DLS) (Zetasizer Nano-ZS; Malvern Instruments Ltd, Worcestershire, UK), and ultraviolet-visible (UV-vis) spectroscopy (UV-1601PC; Shimadzu, Kyoto, Japan). Nanoparticle concentration was determined by UV-vis absorbance via the Beer-Lambert law, assuming that the extinction coefficient ε for gold nanoparticles can be described as a function of particle diameter D as follows (29):

+

⋅

=0935.4

23ln0643.1 3

10Dπ

ε (S1) List of DNA sequences used in this study

DNA sequences used in this study are as follows: DNA strands conjugated to gold nanoparticles Name Sequence NP1 5’- HS−AAAAAAAAAACCTATCGACCATGCT - 3’ NP2 5’- HS−AAAAAAAAAAATGGCCGATGTATGT - 3’ NP3 5’- HS−AAAAAAAAAACGAGTGAGTGCGACG - 3’ NP4 5’- AGAGGAAAGTAGGCTAAAAAAAAAA−SH - 3’ NP5 5’- TAACAACGATCCCTCAAAAAAAAAA−SH - 3’ NP6 5’- GGTGGCTTACAGTCAAAAAAAAAAA−SH - 3’

Linker, attaching, and detaching DNA strands Name Sequence L1 5’- GAGGGATCGTTGTTATACAGTTCAGGCAGTGTAGCATGGTCGATAGG- 3’ L2 5’- AGCCTACTTTCCTCTACATACATCGGCCAT - 3’ A1 5’- TGACTGTAAGCCACCCATTGAACTCGGTGAGTCGTCGCACTCACTCG - 3’ L1comp 5’- CCTATCGACCATGCTACACTGCCTGAACTGTATAACAACGATCCCTC - 3’ A1comp 5’- CGAGTGAGTGCGACGACTCACCGAGTTCAATGGGTGGCTTACAGTCA - 3’

Fluorophore-labeled DNA strands

Name Sequence NP1-FAM 5’- HS−AAAAAAAAAACCTATCGACCATGCT−FAM - 3’ NP3-Cy5 5’- HS−AAAAAAAAAACGAGTGAGTGCGACG−Cy5 - 3’ NP5-Cy3 5’- Cy3−TAACAACGATCCCTCAAAAAAAAAA−SH - 3’ NP6-Cy3 5’- Cy3− GGTGGCTTACAGTCAAAAAAAAAAA−SH - 3’

Cy5-labelled linker DNA strands

Name Sequence NP1comp -Cy5

5’-Cy5 ACAAACAAACAAACAAACAAACAAACAAACAAACAAACAAACAAA GCATGGTCGATAGG - 3’

NP2comp -Cy5

5’-Cy5-CAAACAAAAAAACAAACAAACAAACAAACAAACAAACAAACAAAA CATACATCGGCCAT - 3’

NP3comp -Cy5

5’-Cy5-AAACAAACAAACAAACAAACAAACAAACAAACAAACAAACAAAAC GTCGCACTCACTCG - 3’

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NP4comp -Cy5

5’-AGCCTACTTTCCTCTCCCTCCCTCCCTCCCTCCCTCCCTCCCCTCCCTCCCC TCCCTCC-Cy5 - 3’

NP5comp -Cy5

5’-GAGGGATCGTTGTTATTTGTGGGTGGGTGGGTGGGTGGGTGGGTGGGTGG GTGGGTGGG-Cy5 - 3’

NP6comp -Cy5

5’-GACTGTAAGCCACCAAAGAAAAGAAAAGAAAAGAAAAGAAAAGAAAAG AAAGAAAGAAA-Cy5 - 3’

DNA strand that has nonsense sequence

NS 5’-CACCTGAATGTCAGTAAACAATGCCAGAGTGGAAGGCGAAGTCAGTA- 3’

We chose above DNA sequences in a random manner. We tested all sequences using IDT's OligoAnalyzer service (http://sg.idtdna.com/calc/analyzer) and used only sequences where no secondary structures formed with delta G < -8 kcal/mole. DNA functionalization of gold nanoparticles

Gold nanoparticles were functionalized with single stranded, thiolated single-stranded DNA using a previously published method (15). Briefly, gold nanoparticles were mixed with thiolated DNA strands in 750 μl of 0.01% Tween 20 aqueous solution according to the following table:

Nanoparticle size 13-nm 6-nm 3-nm Particle conc. (nM) 8.0 100 500

DNA sequence NP1 NP2 NP3 NP4 NP5 NP6 DNA conc. (nM) 900 300 1200 400 2000 2000

Nanoparticle : DNA 1:112.5 1:37.5 1:12 1:4 1:4 1:4 Thiolated PEG (MW = 5 kDa) was also added to the mixture (400 nM for 13- and 6-nm, and 1.0 μM for 3-nm particles) to improve particle stability. After 5 minutes incubation, 250 μl of sodium citrate buffer (100 mM of sodium citrate tribasic, pH of which was adjusted to 3.0 by HCl) containing 0.01% Tween 20 was further added to enhance the DNA loading. The mixture was incubated for 30 minutes at room temperature and subsequently purified by centrifugation for 30 minutes at 4˚C. The centrifugation speed was 15,000 g for 13-nm, 120,000 g for 6-nm, and 240,000 g for 3-nm particles (we use these speeds in all of the following centrifugations). After removing the supernatant, DNA-functionalized nanoparticles were further purified by centrifugation using PBS containing Tween 20 (0.01 %w/v) as a solvent (we describe this washing buffer as PBST hereafter).

Hydrodynamic size and UV-vis absorbance of the DNA-functionalized nanoparticles was measured by DLS and UV-vis spectroscopy, respectively. The amount of loaded DNA strands was measured as described in the following section. Quantification of DNA loading on gold nanoparticles

The absolute number of DNA strands conjugated to particles was measured using SYBR® gold nucleic acid stain. Nanoparticles were incubated with one sequence of oligonucleotides at a concentration equal to that of the combined concentrations of the two DNA sequences used for each nanoparticle size in the manner described above. After

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conjugation and washing, thiolated groups were stripped using 10% 1 M DTT at 60 ˚C for one hour. A standard curve was prepared with the same sequences in identical solvent conditions to that of the sample. A working solution of nucleic acid stain was prepared by making a 5000-fold dilution of the concentrated stock in PBS. 50 µL of nucleic acid stain was mixed with 50 μL of sample and incubated for 5 minutes in the dark. Fluorescence of samples and standards was measured by a fluorescence microplate reader (PHERAstar FS; BMG Labtech GmbH, Ortenberg, Germany) using excitation 495 nm and 537 nm emission. The average number of oligonucleotides per nanoparticle of a particular size was assumed to be the absolute number. In order to determine the proportion of each oligonucleotide sequence making up the absolute number, oligonucleotides were conjugated to the nanoparticles using the exact procedure described previously. Linker strands labelled with Cy5 by the supplier (Bio Basic, Inc.) were added to different microtubes containing DNA-conjugated particles such that one sequence in each microtube was hybridized with fluorescent linkers. Fluorescence (excitation 605 nm, emission 670 nm) was measured sequentially for samples of the same nanoparticle size by a spectrofluorometer (FluoroMax-3; Horiba, Kyoto, Japan) and the ratio of fluorescence intensities between the two sequences of each nanoparticle size was used to determine the number of each oligonucleotide on the nanoparticle surface. Assembly of “core-satellite” structure

DNA-functionalized 13-nm particles were first hybridized with linker DNA strands. 6.0 nM of the functionalized 13-nm particle solution was prepared in 1.0 ml of a hybridization buffer consisting of 3X PBS, 5 mM MgCl2, and 0.01% Tween20. L1 and L2 strands were added to the solution with various stoichiometry so that the final concentration of total DNA was 128 nM, leading to the stoichiometries of 64 DNAs per nanoparticle. 32 nM of L2, equal to 4 L2 strands per particle, was used for Figs. 1, 2a, 3, and 4, while variety of stoichiometries (from 4 to 40 L2 strands per particle) was used for Fig. 2a. The mixture was warmed to 60 ˚C for 10 minutes in a water bath and then incubated at 37 ˚C for 2 hours. Linker-functionalized nanoparticles were washed twice by centrifugation at 4 ˚C and 15,000 g for 30 minutes using 1X PBST. After that, 4.0 nM of purified linker-functionalized nanoparticles were mixed with DNA-functionalized satellite nanoparticles at 100:1 molar excess relative to the 13-nm core nanoparticles in 1.0 ml of the hybridization buffer. The ratio of 3-nm satellites to 6-nm satellites was the same as that of L1 to L2 strands used in the previous step. The mixture was incubated at 37 ˚C for 2 hours and then washed 4 times by centrifugation at 4 ˚C and 8,500 g for 30 minutes using 1X PBST. After the last wash, the obtained nanoassemblies (assembly morphology 1) were stored at 4 ˚C prior to use. Shape shift of assembled nanostructures mediated by DNA

The shape of nanoassemblies was changed by the sequential addition of attaching (A1) and detaching (L1comp) DNA strands. After the nanostructure assembly, we cannot apply eq. (S1) to calculate the concentration because of the complex geometry of the nanoassemblies. Instead, UV-vis absorbance around 525 nm was used as a semi-quantitative indicator for the concentration of the assemblies. First, the nanoassemblies whose 525 nm absorbance was 0.12, which corresponds to 0.5 nM of 13-nm particles, were mixed with 12 nM A1 strands in 8 mL of the hybridization buffer. In this step, relatively

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low nanoparticle concentration, ca. 10 times lower than other step, was used in order to prevent possible inter-nanoassembly crosslinking. The mixture was incubated at 37 ˚C for 2 hours and then washed 3 times by centrifugation at 4 ˚C and 8,500 g for 30 minutes using 1X PBST. The obtained nanoassemblies of intermediate shape were further mixed with L1comp strands in 1.0 ml of the hybridization buffer. The 525 nm absorbance of the nanoassembly was 0.70, which corresponds to 3 nM of 13-nm particles, while concentration of L1comp was 300 nM in the reaction solution. The mixture was incubated at 37 ˚C for 2 hours, followed by 3 times washing by centrifugation at 4 ˚C and 8,500 g for 30 minutes using 1X PBST. To reverse the shape change, the obtained nanoassembly of morphology 2 was further mixed with extra attaching and detaching strands, L1 and A1comp, in the manner described above. The obtained nanoassemblies were stored at 4 ˚C prior to use.

To examine specificity of the shape change, a DNA strand that has a nonsense sequence (NS) was also used instead of A1 or L1comp strand. All of the procedure was the same as described above except for the use of NS strand.

Hydrodynamic size and UV-vis absorbance of the nanoassemblies before and after the shape change was measured by DLS and UV-vis spectroscopy, respectively. TEM analysis of the shape-shifting nanoassemblies was carried out as described in the following section. TEM analysis of the shape change of the nanoassemblies

Morphology of the nanoassemblies at each shape-shifting step was observed by TEM. Samples were prepared in the same way as described above. The efficiency of the shape change was evaluated from the obtained images. We defined assembly morphology 1 and 2 by particle distance as illustrated in Fig. S6. First, for each nanoassembly in the TEM images, distance from the center of each 3-nm particle to that of 13-nm particle (r13) and to that of 6-nm (r6) was measured by using ImageJ software. Then, if more 3-nm particles were closer to 13-nm particle in a nanoassembly, the structure was categorized as morphology 1. On the other hand, if more 3-nm particles were closer to 6-nm particles, it was categorized as morphology 2. Nanoassemblies which did not meet both of the above criteria were categorized as “others”, which was mostly composed of primary 13-nm particles generated during the shape-shifting procedures. Primary 3- and 6-nm particles were hardly observed in TEM images because they can easily be removed by centrifugation. > 100 assemblies were analyzed to evaluate the shape change efficiency. Number of 3- and 6-nm satellites on a nanoassembly was also counted from the TEM images and plotted as histograms.

Radial distribution function analysis from TEM images

Radial distribution function (RDF) analysis was performed to quantitatively evaluate the shape change. RDF of satellites from a core particle was calculated from the TEM images. In the case of two dimensions, RDF g(r) can be defined as (21):

( ) ( )ρπ ⋅⋅

=drrrNrg

2 (S2)

where N(r) is the mean number of satellite particles which can be found in a ring of width dr at distance r from one core particle, and ρ is the mean density of nanoassemblies. For

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each core particle, N(r) was analyzed by using ImageJ software with dr = 2 nm. Obtained N(r) was then averaged to obtain g(r). ρ = 6.2×10-5 nm-2 was used.

Fluorophore labeling of the shape-shifting nanoassemblies

To introduce fluorophores to the nanoassemblies, we used fluorophore-labeled DNA strands. NP1, NP3, NP5, and NP6 strands were end-labeled with FAM, Cy5, Cy3, and Cy3 respectively by the supplier (Bio Basic Inc.). These fluorophore-labeled DNA strands and non-labeled NP2 and NP4 strands were conjugated to nanoparticle building blocks in the manner described above to obtain FAM-labeled 13-nm, Cy5-labeled 6-nm, and Cy3-labeled 3-nm particles. The conjugation was confirmed by photoluminescent spectra of DNA-functionalized nanoparticles obtained by a spectrofluorometer (FluoroMax-3; Horiba, Kyoto, Japan). 13-, 6-, and 3-nm particles fluoresced when excited at 460, 605, and 515 nm, respectively.

FRET measurement

By using the fluorophore-labeled nanoparticle building blocks, assembly and shape change of the nanoassemblies were conducted in the same way as described above. At each shape-shifting step (before the washing procedure), FRET signals were measured by the spectrofluorometer. 480 and 520 nm was used for the excitation of FAM and Cy3, respectively. The obtained spectra were normalized to the peak from donors (FAM or Cy3) and then compared.

For the real time FRET measurement, the nanoassembly of morphology 1 was dispersed in 198 μl of the hybridization buffer in a quartz cuvette, the 525 nm absorbance of which was 0.24 (equal to 1.0 nM of 13-nm particles). The cuvette was set on the spectrofluorometer and then the real time monitoring was started. The fluorescent signal at 561nm by 480 nm excitation was tracked as the signal from Cy3 (FRET 1), whereas 665 nm by the excitation at 520 nm was tracked as the signal from Cy5 (FRET 2). After 5 minutes, 2 μl of A1 strands (2.4 μM) were added to the cuvette, followed by immediate mixing. After 60 minutes, 2 μl of L1comp strands (10 μM) was further added to the cuvette with immediate mixing. The monitoring was carried out for 2 hours at room temperature. The obtained signals I were normalized according to the following equation and then plotted against time:

minmax

min

IIIII norm −

−= (S3)

where Imax and Imin represent the maximum and minimum value of I obtained during the real time monitoring.

Assuming first order reaction for each shape-shifting step, time change of I norm for FRET 1 and FRET 2, I1, norm and I2, norm, can be described as follows:

( )11,1 exp0.1 tkI norm −−= (S4)

( )22,2 exp tkI norm −= (S5)

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where k1 and k2 represent the rate constant for the first and the second step of the shape change. t1 and t2 represent the time after the addition of A1 and L1comp strand, respectively. By fitting these equations to the experimental results via the least-squares method, we obtained the values of k1 and k2. FA functionalization of the shape-shifting nanoassemblies

The FA functionalization was achieved by concurrently conjugating FA-functionalized PEG (MW = 5 kDa) terminated with thiol and DNA strands to 13-nm particles. All of the conjugation procedure, including DNA stoichiometry and the ratio of PEG, was the same as described before except for the use of the FA-functionalized PEG instead of non-functionalized PEG. The amount of loaded FA-functionalized PEG was determined by a depletion assay using optical absorbance of FA (30). After nanoparticle incubation, particles were pelleted by centrifugation and the supernatant containing excess FA-functionalized PEG was collected and its absorbance at 363 nm (the extinction coefficient = 6197 M−1cm−1) was measured by UV-Vis and compared to a standard curve. The absorbance of the initial solution was also measured and the calculated difference in concentrations was used to determine the number of molecules per nanoparticle.

Cell culture

U87-MG human glioblastoma-astrocytoma cells were grown in DMEM supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin. Cells were maintained in a humidified incubator at 37 °C with 5% CO2.

Cellular uptake and transmission electron microscopy

U87-MG cells grown to 80-90% confluency in a T75 cell culture flask (NEST Biotechnology Co., Wuxi, China) were harvested using 0.25 % trypsin-EDTA and then seeded on 6-well plates at a density of 2 × 105 cells/well. Cells were incubated overnight at 37°C and 5% CO2 atmosphere. On the day of the experiment, nanoparticles (either the nanoassemblies or their building blocks) were first equilibrated in serum-free media at 37 ˚C for 1 hour. For FA competition assay, free FA (1.0 or 2.0 mM) was also added to the serum-free media. In parallel, cells were washed with PBS, and then incubated in serum-free media for 30 minutes at 37 ˚C for serum starvation. For FA competition assay, cells were incubated with the FA-containing media for 2 hours at 37 ˚C for pretreatment. After that, media was decanted and cells were incubated in 1.0 ml of 6.0 μg/ml nanoparticle-containing medium per well. Cells were exposed to nanoparticle-containing media for 4 hours, followed by 3 times washing by PBS without calcium/magnesium. For each condition, we used at least three independent replicates. Upon the first washing, the nanoparticle-containing medium was collected to observe the nanoassemblies after the incubation. For TEM observation, cells were harvested using 0.25 % trypsin-EDTA, transferred into a 1.5 ml micro-centrifuge tube, and pelleted by centrifugation at 180 g for 3 minutes. After removing the supernatant, cells were fixed in an electron microscopy fixative for 1 hour at room temperature and sliced onto a TEM grid for visualization. The collected nanoparticle-containing medium was also concentrated and dropped on a TEM grid as described above. For ICP-MS analysis, washed cells were stored at -20 ˚C prior to ICP-MS analysis.

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ICP-MS The cellular uptake amount of the nanoassemblies and their building blocks was

quantified by ICP-MS (NexION; Perkin Elmer, Waltham, MA, USA). 6-well plates containing frozen cell samples were first thawed at room temperature. 100 μl of nitric acid was added to each well to collect cell samples. After 30 minutes reaction at room temperature, the cell-suspended nitric acid was transferred into 1.5 ml micro-centrifuge tubes, followed by 1 hour incubation at 70 ˚C to digest the cells. Samples were cooled to room temperature and then transferred into 4.875 ml of water and 25 μl of HCl in 15 ml conical bottom tubes for ICP-MS measurements. A double standard containing gold and magnesium was prepared to measure the concentration of both elements within the cell samples. Magnesium concentration was used to estimate total cell content according to previous reports (28, 31). The t-test was used for the statistical analysis using Kareida Graph software.

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Fig. S1 TEM image and size distribution of post-synthesized building block gold nanoparticles. (A) 13-nm (B) 6-nm and (C) 3-nm. Scale bar is 20 nm. Diameter of >100 nanoparticles was analyzed from TEM images and plotted as histograms. Average diameters ± standard deviations are also shown.

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Fig. S2 Quantification of DNA loading on gold nanoparticles. (A) DNA grafting density of each nanoparticle type. The grafting density was almost the same regardless of nanoparticle size. (B) Number of DNA grafted to each nanoparticle type. The number of DNA increased with nanoparticles size because of larger surface area. (C) Breakdown of DNA sequences grafted for each nanoparticle type. Data are averages ± standard deviations (n = 4).

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Fig. S3 Low magnification TEM images of shape-shifting nanoassemblies. Shape of multiple nanoassemblies was successfully changed. The images also suggest that these nanoassemblies are monodispersed. Scale bar is 100 nm.

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Fig. S4 UV-vis absorbance spectra of nanoparticle building blocks and shape-shifting nanoassemblies. No significant difference was observed after either DNA functionalization or shape-shifting from assembly morphology 1 to 2. These spectra show no signs of aggregation (no shift with the spectra and no peaks in the red), suggesting high colloidal stability of the building blocks and the nanoassemblies.

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Fig. S5 Hydrodynamic sizes of nanoparticle building blocks and shape-changing nanoassemblies measured by DLS. The measurement was conducted in water for post-synthesized nanoparticles, while PBS was used for other nanoparticle types. Hydrodynamic size of building blocks increased after the DNA functionalization due to relatively large size of DNA. Hydrodynamic size of the nanoassemblies was consistent with the TEM observation, suggesting their high colloidal stability in PBS.

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Fig. S6 Definition of nanoassembly shape by nanoparticle distance. Distance from the center of each 3-nm particle to that of the 13-nm particle (r13) and to that of the 6-nm particle (r6) was analyzed from TEM images. If more 3-nm particles are closer to the 13-nm particle in a nanoassembly, the structure is categorized as assembly morphology 1. On the other hand, if more 3-nm particles are closer to 6-nm particles, it is categorized as assembly morphology 2.

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Fig. S7. The population of 3- and 6-nm particles on a nanoassembly of the dominant shape at each shape-shifting step. These populations were analyzed from TEM images.

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Fig. S8 Effect of nonsense DNA strand (NS) on the shape change of nanoassemblies. NS strand was added to nanoassemblies instead of attaching (A1) or detaching (L1comp) strand, and its effect was evaluated by TEM observation (top right of each panel) and RDF analysis (bottom of each panel). Expected morphology of nanoassembly is also schematically shown in top left of each panel.

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Fig. S9 Controlling the number of 3- and 6-nm satellites in a nanoassembly (morphology 1) by the stoichiometry of linker strands L1 and L2. The stoichiometry of L1 and L2 strands were varied while keeping the total DNA stoichiometry to core as 64. The number of 6-nm particles increased with increasing L2 stoichiometry, which accompanied the decrease in the number of 3-nm particles. Data are averages ± standard deviations (n > 100).

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Fig. S10 Photoluminescent spectra of fluorophore-modified nanoparticle building blocks. (A) 13-nm particles were functionalized with FAM-labeled NP1 and non-labeled NP2 strands. (B) 6-nm particles were functionalized with Cy5-labeled NP3 and non-labeled NP4 strands. (C) 3-nm particles were functionalized with NP5 and NP6 strands both of which were labeled with Cy3. 13-, 6-, and 3-nm particles were excited at 460, 605, and 515 nm, respectively. These photoluminescent spectra confirm the successful conjugation of the fluorophore-labeled DNA strands.

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Fig. S11 Time change of FRET signals during the shape change predicted by fist order reaction kinetics. Plots and solid lines represent the results of experiment and kinetic model calculation, respectively. Equations S4 and 5 were fitted to experimental results in Fig. 3D to estimate the rate constants k1 and k2.

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Fig. S12 Extra TEM images of the FA-functionalized nanoassemblies incubated with U87-MG cells. (A) and (B) represent the result of the nanoassembly internalization for morphologies 1 and 2, respectively. The same cell as used in Fig. 4 was observed.

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Fig. S13 TEM images of the non-FA-functionalized nanoassemblies incubated with U87-MG cells. (A) and (B) represent the result of the internalization of nanoassemblies in morphology 1 and morphology 2, respectively. In each panel, the left image shows overall picture of a cell that internalized the nanoassemblies. The right images (i)-(iii) show the enlarged images of their corresponding part in the overall picture. The nanoassemblies found in the culture media after the incubation are also shown. The scale bar is 2.0 μm in the low magnification images and 100 nm in enlarged images.

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Fig. S14 Effect of DNA sequence on the cellular uptake of DNA-functionalized nanoparticles. 13-nm AuNPs were functionalized with each DNA sequence (NP1-NP6) at the stoichiometry of 1:150 and their cellular uptake by U87-MG cells was quantified by ICP-MS. The results were normalized to the cellular uptake amount of NP1-functionalized particles for comparison. Data are averages ± standard deviations (n >3 experimental replicates). Cellular uptake amount of nanoparticles differs according to the surface DNA sequences, suggesting that changes in the surface DNA presentation induced by the shape change could affect the cellular uptake of the nanoassemblies.

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Fig. S15 Cellular uptake of building block nanoparticles relative to their assembled structures. (A) and (B) shows the result of control (without FA functionalization) and FA-functionalized nanoassemblies, respectively. Cellular uptake of the building block particles were quantified by ICP-MS and then normalized to their respective assemblies with morphology 1 for comparison. Data are averages ± standard deviations (n >3 experimental replicates). Comparing (A) with (B), relative cellular uptake of 13-nm building blocks increased by the FA functionalization, while that of 6- and 3-nm particles was almost the same.

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Fig. S16 Effect of free FA on the amount of cellular uptake of the FA-functionalized nanoassemblies between assembly morphology 1 and 2. Nanoassemblies without FA functionalization were also used as control. Data are averages ± standard deviations (n > 3 experimental replicates). Statistical comparisons are shown as follows: **p < 0.01, *p < 0.05, N.S: not significant. These data were used to calculate the ON/OFF ratio in Fig. 4E.

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