Designer RNA nanostructures co-transcribed and self-assembled inside human cell nuclei

Domain: chemistry
Published: 2025-12-26
Editorial outcome: minor_revision

Abstract
The use of nucleic acid-based nanostructures as synthetic biological tools to interface with and regulate cell processes remains challenging. A major obstacle lies in nuclear delivery and retention within live eukaryotic cells. Here, we present a platform of single-stranded RNA that can co-transcriptionally fold into defined nanostructures and assemble into rings, ribbons, and nanonet-like architectures. We validate the formation of these structures in vitro using atomic force microscopy. Then, we demonstrate the functional integration of fluorescent aptamers and RNA sensing capability within the single chain by co-folding with these structures. Notably, we show that the RNA nanonets can be co-transcriptionally produced and assembled directly inside the nucleus of live human cells. We use confocal live-cell imaging and transmission electron microscopy to reveal well-defined nanostructure patterns retained in the nucleus. Together, these results establish a genetically encoded, self-assembling RNA nanostructure system with programmable geometry and localization, providing a foundation for the development of RNA-based nanodevices to examine biological properties in live cells and tissues.

Introduction
RNA plays essential roles in genetic regulation, catalysis, and molecular organization1. A critical aspect of RNA function is its subcellular localization, which is tightly regulated in living cells. Most RNA molecules, including messenger RNAs (mRNAs), are transcribed in the nucleus, undergo processing, and are subsequently exported to the cytoplasm. Other types of RNAs, such as long non-coding RNAs (lncRNAs), are often retained in the nucleus, functioning in gene regulatory programs. There are complex mechanisms controlling RNA localization that involve nuclear export signal sequences and specialized RNA structures2,3. Due to its complexity, understanding and potentially controlling the subcellular localization of RNA, particularly synthetic RNA nanostructures that do not occur naturally, has remained a challenge. A promising strategy is co-transcriptional folding, wherein RNA self-folds while being transcribed by RNA polymerase. This approach ensures higher fidelity and yield of desired nanostructures to be produced in large-scale, using cost-effective materials and, more importantly, allows the in vivo assembly of RNA nanostructures by directly interacting with components of living cells.

Previous works in synthetic co-transcriptional RNA design employed multiple strategies, such as antiparallel crossover4–6, branch-kissing loops7, and stem-loops with kissing loops8, to create diverse RNA assemblies. These co-transcriptional folding RNA structures have been expressed and engineered to work in various environments, including E. coli8, synthetic lipid membranes6, and living cells7–12. However, direct visualization of the structure-defined, synthetic RNA nanostructures in living eukaryotic cells, particularly in the nucleus, has not been achieved. The ability to generate and retain synthetic co-transcriptional RNA assemblies inside the nucleus will help us understand the requirements for RNA subcellular localization as well as push forward applications in RNA-based imaging technologies, specific RNA biosensing for disease diagnosis, and new therapeutics based on simultaneous targeting of entire cellular pathways rather than single molecular components.

One strategy that has shown promise in nucleic acid nanotechnology is the use of parallel crossovers, a structural feature that was initially developed in DNA nanotechnology13,14, later adapted to RNA structures15, and recently used to fold single-chain DNA and RNA16–18. This design feature enables the construction of single-stranded motifs without internal kissing loops, offering advantages in both structural stability and modular assembly.

Here, we report a systematic design strategy for the rational creation of a category of co-transcriptionally folded RNA nanostructures that assemble into nanorings, linear arrays, and extended two-dimensional lattices. The design incorporates RNA paranemic cohesion for stable secondary structure formation and terminal kissing loops for higher-order assembly. These synthetic RNA motifs are compact, modular, and able to fold during transcription both in vitro and in live human cells. By integrating light-up RNA aptamers into the designs, we enable fluorescent visualization of these nanostructures in vitro and in living human cells. Furthermore, we demonstrate that RNA arrays can function as sequence-selective sensors by integrating split aptamers to detect RNA targets in vitro. Notably, we demonstrate direct visualization of assembled RNA 2D arrays remaining inside the nuclei of human cells after transient transfection using electron microscopy. This work expands the capabilities of RNA nanotechnology and provides new routes for nuclear-localized RNA materials in biosensing, intracellular imaging, and gene regulation.

Results
Design of co-transcriptional folding RNA nanostructures based on paranemic cohesions
We developed a systematic design method that allowed single-stranded RNA (ssRNA) nanostructures to fold co-transcriptionally by leveraging RNA A-form parallel crossovers and terminal kissing loops (KLs) for higher-order assembly into rings, linear arrays, and lattices. A synthetic structural feature, RNA paranemic cohesion (PC), was employed in the RNA tiles to enable the continuous single-strand routing (Fig. 1A, B and Supplementary Fig. 1). For the higher-order assembly between ssRNA tiles, three types of RNA KLs were employed, including 180° RNA KLs, 120° RNA KLs, and RNA branch kissing loops (BKL), to create a variety of assemblies (Supplementary Figs. 2 and 3). In 180° RNA KLs and 120° RNA KLs interactions, two terminal loops hybridize at angles of 180° and 120°, respectively. The BKL features an additional helix extending from one side of the terminal loop, enabling branching in the assembled structures.

To design a tile, generally, we first determined the overall structure of an RNA tile, including the number and locations of paranemic cohesions. Each RNA PC domain contains top two strands and bottom two strands, forming stable secondary structural units. RNA KLs were incorporated at the terminals not only for the single-stranded routing but also for encoding the matching rules between tiles, enabling programmable formation of lattices and arrays. Figure 1A shows an example of designing a ssRNA tile containing two PC domains and two KL domains. We planned the routing of the single-chain RNA by introducing an additional parallel crossover at the center of the tile to bridge the two helices, allowing us to connect all domains into a single continuous circular strand. Then, we chose the location of the 5’ end to initiate transcription, with the formation of stem loops, by creating a nick position on the circular strand.

After the structural design, the sequence design of a tile was conducted by following several steps. Generally, we first assigned the sequences of certain nucleotides at key locations, such as 5’ end, kissing loops, and PC domains. Then, we will generate sequences domain by domain from 5’ down to the 3’ end (steps 1 to 4 in Fig. 1C) to ensure the formation of the correct secondary structure of each stem loop and to allow these domains to further interact through paranemic cohesion to form the overall shape (detailed sequence assignment protocol in “Materials and Methods”).

We designed five ssRNA tiles with different geometries and interactive motifs by varying tile length, the number and locations of crossovers, and the incorporation of KLs and BKLs, resulting in the formation of nanorings, linear arrays, and 2D lattices (Fig. 2A). In our co-transcriptional protocol, DNA template, T7 polymerase, and NTPs were added to a mica surface and assembled at 37°C. By using this protocol, all ssRNA tiles were capable of assembling co-transcriptionally, generating well-formed structures as shown by atomic force microscopy (AFM) images. Array sizes and structural quality improved when RNA was subjected to post-transcriptional purification and controlled thermal annealing.

For the five designs, AFM revealed distinct assembly behaviors (Fig. 2B and Supplementary Fig. 4). D1 used 120° RNA KL interactions and formed hexagonal rings. D2 and D3 generated zigzag and linear arrays, mediated by 120° and 180° RNA KLs, respectively. D4 and D5, which differ in the number of paranemic cohesion domains, both produced 2D waffle-like lattices, enabled by the orthogonal arrangement of located BKLs and 180° RNA KLs. These results demonstrate the feasibility of using paranemic cohesion and kissing loops to construct structurally diverse artificial ssRNA tiles and assemblies that can fold co-transcriptionally, allowing us to create complex and functional synthetic RNA nanostructures, such as RNA tiles with defined curvature and the integration of functional aptamers.

Curved RNA tiles for nanoring assembly
Curvature is an interesting synthetic structural feature that has been demonstrated in DNA origami19,20, showing precise control over complex nanostructures. We introduced this structural feature into our ssRNA tile system to show the robustness of our design methods. In our ssRNA tiles, each tile body comprises two continuous double-stranded RNA helices, allowing us to introduce curvature by inserting or deleting nucleotides between crossovers. To explore this strategy, we designed and characterized three curved tiles (D6 to D8) with different sizes, bending, and routings (Fig. 2C). D6 and D8 have the same overall length with 11 nt deletion in the bottom helix relative to the top helix, generating asymmetry that introduces curvature. The only difference between them lies in the position of the central crossover. D7 shares the same crossover position as D6 but features longer helical segments between the two PC domains: the upper helix spans 4.5 turns and the lower helix spans 3.5 turns, compared to D6, which has 3.5 turns and 2.5 turns, respectively. AFM images revealed that all three curved tiles successfully assembled into nanorings, confirming that the engineered curvature was sufficient to induce ring formation (Supplementary Figs. 18–23). To compare their assembly behavior quantitatively, we measured the diameters of the resulting nanorings and fitted their distribution curves (Fig. 2D). D8 produced rings with diameters closest to those expected for a pentamer ring, consistent with the angular contribution of the 11 nt deletion. D6 formed rings with smaller diameters, indicating tetrameric or smaller assemblies, potentially due to the specific placement of its crossover. D7 showed a broader size distribution, likely reflecting the increased flexibility of the longer RNA helices in this design.

Although RNA kissing loops have been well-characterized in various biological and synthetic contexts, their behavior within co-transcriptionally folded ssRNA nanostructures, particularly their sensitivity to buffer conditions relative to tile stability and higher-order assemblies, remains less well understood21. Having successfully designed ssRNA tiles with curvature that assemble into nanorings, we leveraged these finite-sized architectures to study the intermolecular interaction of 180o RNA kissing loops under varying ionic conditions. Our parallel crossover-based ssRNA structures are uniquely well-suited for this investigation because the individual tiles contain no internal KLs, and the KLs are used exclusively at the termini for intermolecular assembly. Therefore, any changes in KL interactions affect only the nanoring formation, without compromising the structural integrity of individual ssRNA tiles.

We first test directly annealed D8 samples in a set of TA buffers containing 0, 0.5, 1, 2, 4, or 6 mM Mg2+. The nanoring formation started to be observed when the Mg2+ concentration reached 0.5 mM or higher (Supplementary Figs. 25–27). Notably, D8 monomers folded into the target tile shape with some level of deformations even under 0 mM Mg2+, as confirmed by AFM imaging, indicating that individual ssRNA tiles can fold across a broad range of ionic buffer conditions. This data aligns with the previous report that ssRNA origami based on paranemic cohesion can be stably stored in phosphate-buffered saline (PBS) without the addition of Mg cations for months, remaining intact22. In addition, KL-mediated ring formation required elevated Mg2+ levels, suggesting that KL interactions for higher-order assembly are sensitive to ionic strength. Similar results were also reported in a previous study21, where stem-looped KL tiles were used to study their assembly under 0, 1, and 5 nM of Mg2+, and the results showed that conformation of the higher-order RNA assemblies can be stabilized by the addition of magnesium.

Integration of fluorescence aptamers into RNA assemblies
To introduce the functionality of ssRNA tiles and facilitate their use in vivo, we incorporated fluorescent RNA aptamer sequences into the tile designs. Specifically, we used the Broccoli aptamer and a split Broccoli aptamer, both of which enhance the fluorescence of small-molecule dye DFHBI-1T23,24. To preserve the co-transcriptional folding and structural integrity of the tiles, we tested four designs, D9 to D12, with different aptamer-inserting locations within the RNA tile sequences that were based on the previously validated D3 and D4 tile frameworks (Fig. 3A–C). Each design employed a different strategy for aptamer integration. D9 had an aptamer tagged at the 3’ end of the tile sequence. D10, a larger design, placed the aptamer sequence near the 3’ end but internal to the strand. D11 displayed the aptamer as an extension from a kissing loop, while D12 incorporated the split Broccoli sequence separately at 5’ and 3’ ends. These designs were optimized to maintain robustness during folding while accommodating the aptamer sequences at different locations. All four designs were transcribed and assembled directly without purification and annealing. AFM imaging confirmed the successful formation of the intended assemblies for all four designs (Fig. 3C and Supplementary Fig. 29). D9 and D10 formed 2D lattices, while D11 and D12 assembled into linear arrays. Fluorescent measurements after 1 h of transcription revealed that D9 and D12 produced stronger signals than D10 and D11 (Fig. 3B), indicating possible preferred terminal locations for integrating aptamer sequences.

Scaffolded RNA sensors based on split aptamers
To explore the sensing applications of ssRNA tiles in vitro, we designed D13, a tile incorporating a split Broccoli aptamer and a target-responsive domain for RNA detection. RNA sensing is a useful approach for diseased diagnostics and gene expression studies25,26. We chose a target sequence derived from the KRAS Proto-Oncogene GTPase, a known biomarker for multiple cancer types with a higher impact on pancreatic and colorectal cancers27,28. Inspired by our previous study, that multistranded RNA tiles can incorporate a split Broccoli aptamer and a target-responsive domain for RNA sensing29, we present a ssRNA tile D13 to explore the sensing application in vitro. As shown in Fig. 3B, D13 consists of two RNA helices connected by a single crossover, where one helix features two 180° KLs at both ends and the other helix is capped with two T-junction loops. The target-sensing domain is split across the 5′ and 3′ ends of the tile, flanking the split Broccoli aptamer. Upon hybridization with the target RNA, the complementary sensing regions bring together the split aptamer fragments, reconstituting the aptamer core and activating fluorescence in the presence of DFHBI-1T (Supplementary Fig. 28). AFM images showed the co-transcriptionally assembled D13, with unpaired sensing domains hanging out from the tile body (Supplementary Fig. 40).

To evaluate sensing performance, we measured the dose-response curve of D13 against increasing concentrations of the target RNA (Fig. 3D). A significant increase in fluorescence was observed between 5 and 10 nM target RNA, indicating a low detection threshold. We also tested the dye response curve in the presence of both ssRNA tiles and target RNA, and the RNA target response curve with ssRNA tiles and dye present (Supplementary Fig. 44). Both experiments were performed in a one-pot, co-transcriptional system without any purification or annealing, demonstrating the sensor’s compatibility with rapid, scalable workflows. In addition to sensitivity, we assessed the specificity of D13 by testing various mutated RNA targets, M1 to M7, that were derived from a previous work29. M1 to M4 contained 12 nt, 8 nt, 2 nt, and 3 nt substitutions, respectively, and produced fluorescence levels comparable to the negative control. M5 to M7, with 1 nt, 5 nt, and 2 nt mutations, generated reduced but still detectable signals relative to the fully matched target (Fig. 3E–G and Supplementary Fig. 45). This partial activation likely results from the specific locations or minimal number of mismatches, which may permit partial hybridization to facilitate the aptamer core formation, suggesting that the specificity of our sensor is also influenced by structural factors of the split aptamer in addition to the number of mismatches. Although the current sensing experiments are conducted in a cell-free synthetic system, previous studies of RNA biosensing using split Broccoli30,31 suggest that this sensor design potentially can also be adapted in living cells for visualizing and monitoring RNA assembly.

Folding and assemble RNA structures in human cell nuclei
Building upon the in vitro characterization, we next assessed whether our designs could be applied in biologically relevant settings. We selected D4 and D10, forming large nanonet-like configurations (Figs. 2A D4 and 3B D10) to generate expression plasmids (Supplementary Figs. 72–74) and evaluate their co-transcription and in vivo assembly by transfecting human embryonic kidney 293FT cells (HEK293FT) (Fig. 4A). Their expression was analyzed by quantitative Reverse Transcription-PCR (RT-qPCR) and the nanostructures were visualized by confocal imaging. Prominent expression of both RNA structures was observed 24 h after transfection (Fig. 4B and Supplementary Fig. 46). Since most of the RNA tiles form double-stranded RNA helices upon folding, we used monoclonal J2 antibodies, which detect double-stranded RNA helices larger than 40 bp irrespective of the sequence32. These antibodies are widely used for detecting double-stranded RNA of viruses in infected cells33. To monitor transfection efficiency, we co-transfected HEK293FT cells with the D4 and D10 expression plasmids along with a plasmid expressing mCherry as a transfection marker, followed by immunostaining with J2 monoclonal antibodies. At 24 h post-transfection, we observed 68.50% and 67.09% of cells expressing mCherry showed assembled D4 and D10, respectively (Supplementary Fig. 69). We analyzed the subcellular distribution of these assemblies, classifying them as nucleus only, nucleus and cytosol, or cytosol only. For D4, 48.97% of cells showed nuclear distribution, 47.59% showed both nuclear and cytosolic, and 3.59 % showed cytosolic only localization (Supplementary Fig. 70). For D10, 3.59% showed nucleus only, 88.11% showed nuclear and cytosolic distribution, and 11.15% cytosolic only (Supplementary Fig. 71). Notably, our assemblies localize primarily within the nuclear compartment and we observed distinct dsRNA-positive structures in most cells transfected with either D4 or D10 (Fig. 4C Top left panels, Supplementary Figs. 47 and 48). Both of these nanostructures displayed the expected nanonet-like configuration (Fig. 4C Top right panels, Supplementary Fig. 49 and 50). To further validate the structural assembly, we co-transfected cells with D4, which contains a Broccoli aptamer, and performed live-cell imaging 24 h post-transfection in the presence of DFHBI-1T and Hoechst. These live imaging studies revealed a similar nuclear nanonet-like configuration (Fig. 4C, Bottom panels, Supplementary Figs. 51–54) to the nanostructures observed in immunostained cells using J2 monoclonal antibodies (Fig. 4C, Top panels). To probe for the dynamic properties of the assembled nanostructures, we leveraged the photochemical sensitivity of the DFHBI-1T fluorophore. DFHBI-1T has been shown to undergo rapid isomerization and dissociation from the Broccoli aptamer upon laser illumination34. We therefore performed live-cell photobleaching experiments and observed rapid loss of fluorescent signal within the photobleached region, followed by recovery within 30 seconds (Supplementary Figs. 55 and 56). The observed nanostructures further support the dynamic binding of DFHBI-1T to correctly folded RNA tiles. Collectively, these results validate the co-transcriptional expression and proper self-assembly of our designed RNA tiles within the nucleus of human cells.

Given the resolution limits of confocal microscopy and the nanoscale dimensions of the RNA tile subunits, we sought to further characterize their assembly in HEK293 cells by performing Transmission Electron Microscopy (TEM) on the two selected tiles, D4 and D10 (Fig. 4C, Top panels). TEM analysis revealed large organized synthetic nanostructures with architectural assembly patterns that are consistent with the ones observed by immunofluorescence (Fig. 5A–C and Supplementary Figs. 57–66). We observed distinct waffle-like 2D arrays (Fig. 5B inset 5, Fig. 5C inset 7) that closely resemble those formed by co-transcriptional folding using T7 RNA polymerase visualized by AFM (Figs. 2B D4 and 3B D10). Side-by-side comparisons between AFM and TEM images at the same scales showed that D4 and D10 can efficiently assemble into nanonet-like structures both in vitro and in vivo, inside human cell nuclei. This was further demonstrated by the similar distances measured between the neighboring lines of these arrays in the AFM and TEM images (Supplementary Figs. 67).

Collectively, using both fluorescence cell imaging and TEM, D4 and D10 exhibit a co-transcriptional self-folding into a nanonet-like configuration inside human cell nuclei, with a striking similarity to the in vitro nanostructure demonstrated by AFM. Our TEM images show the integrity of nuclear components in D4-transfected HEK293 cells. For instance, the TEM shows clear nucleolar structures along with well-defined heterochromatin and euchromatin regions (Supplementary Figs. 59–61). Thus, we present a clear demonstration, both in vitro and in vivo, that our designed synthetic RNA nanonet-like structure is co-transcriptionally folded within human cells and is more prominently visible in the nucleus.

Methods
Structural design of RNA tiles
RNA helices were created in Tiamat40 using the CreateStrand function. Helices were rotated and transferred to preferred positions where parallel crossover could be added manually. The number of nucleotides at the paranemic cohesion (PC) should be 8, and at least one turn (11 nt) was used to space between neighboring PCs. Take D3 as an example, we built two paranemic cohesions with an 8 nt length. Between two paranemic cohesions, we made a center parallel crossover to rout this RNA tile into a single strand. The distance between this crossover and the two paranemic cohesions was 11 nt and 14 nt to increase the stability of RNA tiles. The space between PC and kissing loop was chosen at 8 nt to increase the stability of the structure. 180o RNA kissing loops were derived from natural HIV RNA (PDB_ID: 1B8R)41 and 120o RNA kissing loops originated from ColEI plasmid from Escherichia coli (PDB_ID: 2BJ2)42. RNA branch kissing loops originated from this published research7. RNA tiles were drawn in Adobe Illustrator and arrays were built by moving monomers together. Molecular models were rendered by UCSF ChimeraX43 in Fig. 1A.

Sequence assignment of RNA tiles
First, we make the first two nucleotides at 5’ GG to facilitate the RNA transcription. Then we add the sequence to the kissing loops. The total number of G and C bases in every kissing loop is 3. Next, we assign the sequence of PC domains. 4 out of 8 nt at PC are assigned G or C to increase the stability of cohesion. Then we assigned the rest of the RNA tile by the Generate Sequence function of Tiamat with the following settings: Unique Sequence Limit 4; Repetition Limit 4; G repetition Limit 3; G-C Percentage 0.5; Check Sliding Base Yes; Operation Time Out 8. Then G-U wobble pairs were added every 8 to 11 base pairs except for the paranemic cohesion and kissing loop region. After this, we break the tile into domains containing half of a paranemic cohesion and steps in the order of transcription (step 1 in Fig. 1C). NUPACK44 and mFold45 are used to predict the secondary structures of the half paranemic cohesion domain. If the predicted structure differs from the one we designed, we modify the nucleotides near the unexpected secondary structures until NUPACK and mFold output the correct secondary structures. This step ensures that the domain of step 1 of our tile can fold correctly during transcription. Then, we proceed to the step 2 domain (Fig. 1C) and repeat this process until both software outputs the correct folding of our structure at step 4.

Preparation of ssRNA
The subsequent RNA sequences were translated to DNA templates. T7 promoters and PCR primers were added to the 5’ and 3’ (sequence can be found in the Sequence section of this document) and ordered as gBlock DNA from Integrated DNA Technologies (IDT). The gBlock DNA was then diluted to 4 ng/µL in water and amplified by PCR with Taq PCR Kit, from New England Biolabs (NEB). The sample was denatured for 1 min at 95°C, followed by 25 cycles at 95°C for 30 s, 50°C for 1 min, and 70°C for 2 min. The final extension was at 70°C for 5 min. PCR product was subjected to denaturing polyacrylamide gel purification and used for RNA transcription with the HiScribe® T7 High Yield RNA Synthesis Kit (NEB) using its protocol. RNA product was then subjected to another round of denature polyacrylamide gel purification before use.

Co-transcriptional assembly protocol
Co-transcriptional assembly was performed on mica, unless otherwise noted. A freshly peeled mica (Ted Pella) was stuck on a metal disk and was placed in a petri dish with wet filter paper to reduce humidity. This petri dish was kept at 37°C in a PCR chamber with 80 µL of 1× TAE Mg buffer (40 mM Tris base, 20 mM acetic acid, 12.5 mM magnesium acetate, and 1 mM EDTA, with a pH of 8.2) added to the mica. A 20 µL sample was made with 30 ng DNA template, 0.01 µL T7 enzyme, 8 µL NTPs, 1 µL DTT, and 1× TAE Mg buffer instead of T7 buffer. All ingredients are from the T7 kit if otherwise noted. This sample was then transferred to the warm mica and allowed to cool stepwise from 37°C to 20°C over 2 h. D14 and D6 to D8 were co-transcriptionally assembled in solution in a 20 µL sample system with 30 ng DNA template, 1 µL T7 enzyme, 8 µL NTPs, 1 µL DTT, and 1× TAE Mg buffer at 37°C for 2 hrs.

Annealing assembly protocol
Purified RNA was annealed on mica if not specifically noted. For assembly on mica, the mica preparation process was the same as the co-transcriptional protocol. After preparation, the petri dish was placed in a PCR chamber at 70°C with 100 µL of 1× TAE Mg buffer. Prior to assembly, the RNA underwent a quick heating-cooling process (90°C for 3 min, 4°C for 5 min). Then, 500 ng of RNA was diluted in 20 µL of TAE Mg buffer and heated to 70°C before being added to the warm mica. This sample underwent a stepwise cooling ramp from 70°C to 20°C overnight (~12 h). D14 and D6 to D8 were annealed and assembled in solution in a 20 µL sample system with 500 ng of RNA diluted in 20 µL of TAE Mg buffer, and underwent a stepwise cooling ramp from 70°C to 20°C overnight (~12 h).

AFM imaging
AFM imaging was carried out using Dimension FastScan AFM from Bruker in ScanAsyst in Liquid mode and ScanAsyst-Fluid+ tip from Bruker. Sample on mica was washed with 200 µL TAE Mg buffer before imaging and 2 µL 100 mM NiCl2 solution was added to improve contrast. For the sample prepared in solution, 0.5 µL was diluted with TAE Mg buffer to 80 µL and mixed with 2 µL 100 mM NiCl2 solution before adding to the mica. Autocontrolled peak force setting was used for all samples. Ring diameter measurement in Fig. 2D of the manuscript was performed using ImageJ, and the data were processed using the Gaussian fitting function in OriginPro software.

Fluorescence measurements
Sample preparation. Experimentally, D13 was cotranscribed with HiScribe® T7 High Yield RNA Synthesis Kit (NEB) for 1 h. Then, the reaction mixture, without further purification, was directly diluted to 0.5 µM for specificity testing, and the concentration of D13 was calculated based on the OD260 reading from the Nanodrop. Noted that the in vitro transcripts always contain elongated or shortened byproducts, so that the calculated concentration is higher than the concentration of pure D13, and the accurate concentration of D13 was not quantified. M1 to M7 are seven RNA mutants adapted from our multistranded RNA tile study29. Unpurified cotranscriptional products of D13 (0.5 µM) were mixed with each RNA target (0.5 µM, purified) and DFHBI-1T (10 µM), and annealed at 37°C in a thermocycler for 30 min.

Measurement. Fluorescence signals were collected on a Horiba Fluorolog-3 spectrofluorometer with excitation at 470 nm and emission wavelengths at 505 nm. The fluorescence test of the co-transcriptional sample was performed after overnight transcription (~12 h) of the ssRNA tile using the T7 Kit protocol. A 1 µL transcription product was diluted to 50 µL in HEPES buffer containing 100 mM potassium chloride and 1 mM magnesium acetate. 1 µL 0.5 mM (Z)-4-(3,5-difluoro-4-hydroxybenzylidene)-1,2-dimethyl-1H-imidazol-5(4H)-one (DFHBI) was added before testing.

Plasmid construction
Co1-BKL and Co-APT-V2 expression plasmids were generated using standard molecular cloning techniques. Synthetic gBlocks encoding D4 and D10 (IDT) were used as templates for PCR amplification to introduce EcoRI and AgeI restriction sites. The PCR products and the pLKO.1-puro vector (Addgene #8453) was digested with EcoRI-HF and AgeI-HF (New England Biolabs) restriction enzymes and purified using the QIAquick PCR Purification Kit (Qiagen) according to the manufacturer’s instructions.

The purified, digested inserts and vector were ligated using T4 DNA Ligase (NEB) at room temperature for 1 h, followed by heat inactivation at 65°C for 20 min. Four microliters of the ligation reaction were transformed into One Shot Stbl3 chemically competent E. coli (Thermo Fisher Scientific) using heat shock. Colonies were screened by colony PCR using primers flanking the insertion site in the pLKO.1 vector (U6 promoter forward: 5′-CTCGAGCCGCGGCCAAAGT-3′; reverse: 5′-GAGGGCCTATTTCCCATGATTCCTT-3′). Positive clones were expanded overnight in LB medium containing ampicillin, and plasmids were isolated using the CompactPrep Plasmid Kit (Qiagen).

pLKO.1 puro was a gift from Bob Weinberg (Addgene plasmid # 8453; http://n2t.net/addgene:8453; RRID: Addgene_8453)

Cell culture and transfection
HEK293FT cells were obtained from Invitrogen (Cat# R70007) and cultured on Matrigel-coated tissue culture plates in Dulbecco’s Modified Eagle Medium (DMEM; Thermo Fisher Scientific) supplemented with 10% fetal bovine serum (FBS; HyClone), 1% non-essential amino acids (NEAA; Sigma), 1% GlutaMAX (Gibco), 1% sodium pyruvate (Sigma), and 1% penicillin-streptomycin (Gibco). Cells were maintained at 37°C in a humidified incubator with 5% CO₂ and passaged twice per week.

For transfection experiments, cells were trypsinized the day before and plated onto Matrigel-coated coverslips (for immunofluorescence) or chambered dishes (for live imaging) at a density of 2.5×10² to 4.5×10² cells per well. On the day of transfection, 290 ng of expression plasmid and 10 ng of mCherry vector were diluted in Opti-MEM (Gibco) and mixed with polyethyleneimine (PEI) at a final DNA:PEI ratio of 1:4. The mixture was incubated for 20 min at room temperature to allow transfection complexes to form. Cells were washed and fresh antibiotic-free medium was added prior to the addition of transfection complexes. After a 6-hour incubation, the medium was replaced with fresh complete medium. Cells were either fixed or collected for downstream experiments 24 h post-transfection.

Immunofluorescence
Fixed cells were washed three times with PBS and permeabilized with 0.5% Triton X-100 (TX100) in PBS for 10 min at room temperature. To reduce non-specific binding, cells were blocked with a solution containing 2% goat serum, 1% BSA, 0.2% skim milk, 0.1 M glycine, and 0.1% TX100 in PBS for 1 h at room temperature.

Primary antibody incubation was performed overnight at 4°C in a humidified chamber using mouse anti-dsRNA (J2 clone, Nordic-MUbio) diluted 1:250 in 0.1% BSA in PBS. The following day, the cells were washed six times by gently dipping them in a beaker containing 0.1% TX-100 in PBS. Cells were then incubated with Cy2- or Cy5-conjugated secondary antibodies (Jackson ImmunoResearch) for 1 h at room temperature in a dark, humidified chamber.

After secondary incubation, cells were washed again and mounted using Vectashield Vibrance with DAPI (Vector Laboratories). Mounted slides were allowed to cure overnight at room temperature.

Images were acquired using a Zeiss LSM980 confocal microscope, using either a 20× or 63× oil immersion objective. DFHBI-1T was visualized through EGFP filter with shortened detection range (490–552 nm) to reduce mCherry bleed through. Image processing and analysis were performed using ZEN Blue (Zeiss) or ImageJ (NIH) software.

Quantitative PCR
Total RNA was extracted using TriPure Isolation Reagent (Roche) according to the manufacturer’s protocol. RNA concentration was measured using a Quick Drop spectrophotometer (Molecular Devices), and 1 µg of total RNA was used to synthesize cDNA using the LunaScript RT SuperMix Kit (New England Biolabs). cDNA was subsequently diluted 1:10 in TE buffer.

qPCR was performed using Luna Universal qPCR Master Mix (NEB) on a CFX384 Touch Real-Time PCR Detection System (Bio-Rad). Each 10 µL reaction contained 5 µL of Luna Master Mix, 1 µL of diluted cDNA, and 250 nM of each primer.

Gene expression was quantified using the ΔΔCt method, normalized to Gapdh, and reported as fold change relative to the control vector. All reactions were performed in at least three technical replicates, and error bars are presented as SEM.

Transmission electron microscopy
Cultured cells were collected by sedimentation, and the resultant pellets were fixed using 2.5% glutaraldehyde and 2.0% paraformaldehyde in 0.1 M Sodium Cacodylate buffer. After fixation, pellets were post-fixed in 1% osmium tetroxide, and block stained in 1% aqueous uranyl acetate, followed by dehydration and embedding in Embed 812 resin (Electron Microscopy Sciences). Ultrathin sections were cut, collected on copper grids, and post-stained using uranyl acetate followed by Reynolds’ lead citrate solution. Images were acquired using a Thermo Fisher FEI Tecnai 12 BioTwin Transmission Electron Microscope (Thermo Fisher Scientific) equipped with a Gatan OneView CMOS (4K by 4K) camera (Gatan). Image processing and analysis were performed using Gatan Digital Micrograph or ImageJ software.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "Review of \"Designer RNA Nanostructures Co-Transcribed and Self-Assembled inside Human Cell Nuclei\"\nChang et al. present a research study on the design and characterization of RNA nanostructures that can be expressed and\nfolded inside human cells. The study uses the RNA paranemic crossover (PX) motif for cotranscriptional foldning,\ndemonstrates new design concepts such as wireframe PX grids and curvature, and characterize the PX wireframe grids\ninside human cell nuclei. Compared to previous work on expression of RNA nanostructures in bacteria, yeast and human\ncells (Li et al. 2018; Jepsen et al. 2018; Liu et al. 2020; Nguyen et al. 2022; Pothoulakis et al. 2022; Zhang et al. 2024), what\nstands out in the present study is the direct visualization of RNA nanostructures in cells by Transmission Electron\nMicroscopy (TEM) and is therefore a significant contribution to the field. However, there are several issues in the description\nof pervious work, and in the presentation and analysis of experimental results that needs to be addressed before publication\nof the work.\nMajor comments:\n1. The manuscript seems to be overselling that the RNA nanostructures are localized in the nucleus, since this is what is to\nbe expected when using the U6 promoter. It is, however, different from the previous study (Zhang et al. 2024), which was\nalso using the U6 promoter, and found the RNA nanostructures in the cytoplasm. The manuscript would benefit from a better\ndescription of RNA expression strategies with references to previous literature and a discussion of the likely reason for the\ndifferent localizations observed in the current and previous study. Please also provide a plasmid map in the supplementary\ninformation.\n2. The description of the design of the RNA nanostructures is not clear. In the text the design is described as \"carefully\nconducted\" (page 3) and \"carefully designed\" (page 4), but that is not a scientific way to describe the design process. Please\nexplain in general terms in the main text how the RNA nanostructures were designed and reference the Material and\nMethods. The current description of the design process in the Material and Methods is not clear and should contain enough\ndetails so that other researchers can reproduce your design approach. Please explain how the initial sequence was\ngenerated by Tiamat (and remember a reference) and explain the procedure to generate sequences for co-transcriptional\nfolding and the rationale behind what sounds like a manual procedure. As a sidenote, NUPACK and Mfold is not a\n\"simulation\" but a secondary structure prediction using minimum free energy algorithms.\n3. The manuscript is not referencing previous literature sufficiently. Here are some examples: (1) It would be appropriate to\nreference previous studies describing the expression of RNA nanostructures in cells (Li et al. 2018; Jepsen et al. 2018; Liu\net al. 2020; Nguyen et al. 2022; Pothoulakis et al. 2022; Zhang et al. 2024). (2) The following sentence: \"Unlike previously\nreported co-transcriptionally assembled RNA nanostructures, which rely on branch-kissing loops or antiparallel crossover\ninteractions, our design employs parallel crossover interactions, a structural strategy not previously explored in this context.\"\nis not describing that the paranemic crossover has been used before for co-transcriptionally assembled RNA nanostructures\nin the paper by Vallina et al. (Vallina et al. 2023). (3) In relation to the RNA paranemic crossover it would also be appropriate\nto mention the original study of this motif (Afonin et al. 2008). (4) References are missing for previous studies on curvature on\npage 3 and KL sensitivity to buffer conditions on page 4. (5) Concerning the description of the RNA biosensing approach\nusing split Broccoli, previous literature that used similar approaches should be referenced e.g., (Rogers et al. 2015; Alam et\nal. 2017), and the results should be compared.\n4. There is an issue with the Mg-dependence experiments, since TAE buffer is used to make the different dilutions.\nHowever, TAE contains EDTA which chelates Mg2+ and therefore the Mg2+ concentrations are likely misrepresented.\nSince EDTA chelates 2 Mg ions it might be possible to calculate the effective concentration of Mg2+ in your experiments but\nit would be best to repeat experiments without EDTA by using a TA buffer for dilitions. It would be appropriate to reference\nprevious papers that describe the Mg dependence of KLs and compare them to your results. In regard to Mg dependence,\nyou assume that the PX tiles does not require Mg, but it is known that Mg can be important for stabilizing crossovers and for\nalignment of helices. To my eye, the monomer tiles look a bit distorted in the AFM images, which I think should be\naddressed in the discussion of your results.\n5. In the Split-Broccoli sensor experiments you write that \"5 and 10 nM target RNA, indicating a low detection threshold\"\nbut you lack an evaluation of the statistical significance of this detection threshold. It is not clear how many independent\nexperiments were done, and the standard error bars seem very small for independent experiments. The sensor should be\ntested in more detail and the sensitivity reported in relation to target to sensor ratio. In Fig. 3CDE, it is not clear what the\nfluorescence intensity is normalized to and why the fluorescence is not directly reported. In Fig. 3E, the shading is confusing\nin the bar plot and turning the bar plot on its side is not necessary. Please show the M1-M7 mutations directly in the figure,\nwhich will help the reader to understand the effect of the mutations and their closeness to the junction. You write \"M1 to M7\ncontained 12nt, 8nt, 2nt, and 3nt substitutions, respectively, and produced fluorescence levels comparable to the negative\ncontrol.\" This is clearly not true, since M5-M7 shows some fluorescence. Finally, the description of \"moderate single-\nnucleotide specificity\" is unclear. Can you be more quantitative?\n6. The experiments showing expression of RNA nanostructures in cells needs some more details. Why does the mCherry\nexpression look different in the top panels and the bottom panels of Fig. 4C? In the top panels the expression is mainly in\nnucleus, and in the bottom panels the expression is also in the cytoplasm. Is the expression time different between these\nexperiments and could this influence the results?\n7. You write that \"Overlay of the AFM reference regions onto TEM micrographs further confirmed the accurate nuclear\nassembly of D4 in human cells (Figure S67 Supp. Figure 67).\" but you forget to mention that this was also done for D10.\nIn Fig. S67, the drawing on top of the TEM image seems like an overinterpretation, where you draw more than you see.\nIt would be more correct to just draw the lines that you actually see, then measure the distance between the lines, and\ncompare this measurement to the AFM data. If the distance match, you have a good indication that you are looking at the\nsame thing. The connections between lines that you cannot see from the TEM image, you can only see from the AFM. Thus,\nyou can measure these distances in the AFM images and compare them to the theoretical model.\n8. There is an issue with the text about data availability, where you write \"???\" and have not made the Source data files\navailable. The Source data files should be uploaded and made available to the reviewers, so that we can properly evaluate\nthe quality of the data presented.\nMinor comments:\n9. Fig. 1B and Fig. 1C are not comparable and there is confusion about strand direction and colouring scheme. I see from\nFig. S7 that you have the right strand synthesis direction in Fig. 1C. But you need to use the colour gradient consistently.\nE.g. use 5' end as blue and 3' end as red, which corresponds to the standard rainbow colour scheme.\n10. In Fig. 2 and 3 I think it is confusing to have cotranscriptional (CT) AFM images as an inset on top of heat-annealed (HA)\nAFM images, since an inset normally shows a zoom on details of the bigger picture. One possibility is to show them side-by-\nside and then label them clearly in the figure as \"CT\" and \"HA\". Another option is to just show the CT data and leave the HA\ndata in the supplementary information, since CT is the most relevant for the in vivo study.\n11. Fig. 3A is confusing to read: The first part with the test tube in not informative and the branched arrow is misleading.\nWould be better to use the space to show the two different constructs individually: D10 before and after adding the dye and\nD13 before and after adding the target RNA and dye.\n12. It is unclear what is shown in Fig. 3B, since you mention in the text that experiments were done without purification and\nannealing, but annealing data is shown in the figure, so it gives the wrong impression and confuses the reader.\n13. You write \"The unexpected performance of D12 may be attributed to the G-rich sequence at the 5′ end, which could\nenhance transcription efficiency or folding kinetics. Based on structural yield, D10\". The G-rich claim seems like speculation,\nand you will need to be more precise, reference literature, or do experiments to support your claim. Furthermore, I don't know\nwhat you mean by \"structural yield\" and how you evaluate it.\n14. D13 is not really a tile, but just a flexible holiday junction, so why does it fold into neat linear filaments. Please highlight\nthis surprising observation and discuss it.\n15. You write \"the assembled D13, with unpaired sensing domains hanging out from the tile body\" but forget to reference the\nfigure.\n16. There are some issues with language. Please do spelling and grammar check of the document.\n17. There are some issues with \"non-specific/scientific writing\". Please try to be concise.\n18. In Fig 4 it does not make sense to show the grey and green DFHBI icon on the images. That is an overinterpretation. In\nFig. S44 the DFHBI icon should be grey.\n19. Fig. S49-S66: Caption about labelling should either be in each figure caption or be made as a common figure caption for\nall figures.\n20. Please don't show DNA sequences as triplets, which suggests codons.\nReferences:\nAfonin, K. A., D. J. Cieply, and N. B. Leontis. 2008. 'Specific RNA self-assembly with minimal paranemic motifs', J Am Chem\nSoc, 130: 93-102. https://doi.org/10.1021/ja071516m\nAlam, K. K., K. D. Tawiah, M. F. Lichte, D. Porciani, and D. H. Burke. 2017. 'A Fluorescent Split Aptamer for Visualizing\nRNA-RNA Assembly In Vivo', ACS Synth Biol, 6: 1710-21. https://doi.org/10.1021/acssynbio.7b00059\nJepsen, M. D. E., S. M. Sparvath, T. B. Nielsen, A. H. Langvad, G. Grossi, K. V. Gothelf, and E. S. Andersen. 2018.\n'Development of a genetically encodable FRET system using fluorescent RNA aptamers', Nat Commun, 9: 18.\nhttps://doi.org/10.1038/s41467-017-02435-x\nLi, M., M. Zheng, S. Wu, C. Tian, D. Liu, Y. Weizmann, W. Jiang, G. Wang, and C. Mao. 2018. 'In vivo production of RNA\nnanostructures via programmed folding of single-stranded RNAs', Nat Commun, 9: 2196. https://doi.org/10.1038/s41467-\n018-04652-4\nLiu, D., C. W. Geary, G. Chen, Y. Shao, M. Li, C. Mao, E. S. Andersen, J. A. Piccirilli, P. W. K. Rothemund, and Y.\nWeizmann. 2020. 'Branched kissing loops for the construction of diverse RNA homooligomeric nanostructures', Nat Chem,\n12: 249-59. https://doi.org/10.1038/s41557-019-0406-7\nNguyen, M. T. A., G. Pothoulakis, and E. S. Andersen. 2022. 'Synthetic Translational Regulation by Protein-Binding RNA\nOrigami Scaffolds', ACS Synth Biol, 11: 1710-18. https://doi.org/10.1021/acssynbio.1c00608\nPothoulakis, G., M. T. A. Nguyen, and E. S. Andersen. 2022. 'Utilizing RNA origami scaffolds in Saccharomyces cerevisiae\nfor dCas9-mediated transcriptional control', Nucleic Acids Res, 50: 7176-87. https://doi.org/10.1093/nar/gkac470\nRogers, T. A., G. E. Andrews, L. Jaeger, and W. W. Grabow. 2015. 'Fluorescent monitoring of RNA assembly and\nprocessing using the split-spinach aptamer', ACS Synth Biol, 4: 162-6. https://doi.org/10.1021/sb5000725\nVallina, N. S., E. K. S. McRae, C. Geary, and E. S. Andersen. 2023. 'An RNA Paranemic Crossover Triangle as A 3D\nModule for Cotranscriptional Nanoassembly', Small, 19: e2204651. https://doi.org/10.1002/smll.202204651\nZhang, Tianqing, Xinmin Qian, Huangchen Cui, Qingting Li, Xinyu Du, Xiaoyu Fu, Hongzhou Gu, Wenwen Zeng, and Bryan\nWei. 2024. 'Programmable in vitro and in vivo folding of single-stranded nucleic-acid wireframe origami', Chem, 10: 2550-63.\nhttps://doi.org/10.1016/j.chempr.2024.04.023\nBest regards,\nEbbe Sloth Andersen", "reviewer_2": "The authors developed a new RNA nanostructure design strategy in the study. RNA paranemic cohesion were introduced in\nthe folding of individual RNA tiles, and 180° RNA kissing loops, 120° RNA kissing loops, and RNA branch kissing loops\nwere designed for intermolecular RNA tiles interactions. They demonstrated various designs and the AFM images confirmed\nthe desired self-assembly. Then the authors induced broccoli aptamer to their structures in biosensor applications.\nImportantly, these RNA nanostructures could be co-transcriptionally produced and assembled in human cells with a nucleus\nlocalization.\nSpecific points for improvement are listed as below.\n1. In figure 3C, the sample D12 showed the highest fluorescence intensity. When it came to detect RNA and its mutants, why\nD13 instead D12 was used? It would be great if the authors can clarify.\n2. In figures 4 and 5, the authors used D4 and D10 to confirm the localization of co-transcription in the nuclei of cells, and\nexplained the phenomenon by cargo size and structure rigidity for nuclear membrane transport. Other samples of smaller\nsizes, such as D1 and D3, should be applied to prove the hypothesis.", "reviewer_3": "In this study the authors demonstrate a new variation on single-stranded RNA origami tile assembly that form programmable\nRNA nanostructure cotranscriptionally in the nucleus of mammalian cells. Stable expression of RNA origami in human cells\nhas been a goal in this field for years, and this result represents a major advancement as the authors demonstrate both the\nexpression of intricate RNA nanostructures in human cells and also localize the structures to the nucleus. Also, for the first\ntime we are able to see the finer structure of the RNA assemblies in the cell by TEM. The advancement hinges on a new\nRNA architecture is able to form a two-helix tall tile without the internal apical loop-loop interaction that is typically used,\ninstead the authors devise a parallel helix paranemic cohesion (PC) motif to achieve an apparently faster folding structure\n(based on the assumption that loop-loop assembly is slower than paranemic cohesion).\nThey convincingly demonstrate the formation of cotranscriptional RNA nanonets, rings and filaments with extensive atomic\nforce microscopy (AFM) work. The AFM imaging is very high quality, and of sufficient resolution to resolve even minute\ndetails of their structural designs. Numerous different structural variants are constructed and compared, including distorted\ncurved versions of the tiles, providing a broad overview of the assembly space for the new structural tile. The authors further\ninvestigate the effect of integrating light-up aptamers and chemical sensing elements into the designs, and were able to test\nand validate this as well in vivo. Live cell immunofluorescence imaging clearly demonstrates that the RNAs localize to the\nnucleus of human cells, further backed up by a photobleaching experiment on the fluorescent aptamer tags. Lastly, and also\nmost critically for the study, transmission electron microscopy imaging was able to reveal some fine structure details of the\nRNA periodic assemblies in mammalian cell nuclei, demonstrating the formation and retention of programmable RNA nano-\nassemblies in the nucleus. This is a very nice work, and I recommend the paper for publication in Nature Communications\nafter the following revisions/clarifications are made. Here I list my concerns, which I would like to have addressed before\npublication:\n1) I feel that the rationale for RNA structure design could still be better explained. I think you could further explain the logic\nbehind this particular strand routing, how did you come up with it? I spent a long time to try to figure it out, if you untangle the\nPX the secondary structure is a simple X shape. To fold into a tile two 8bp PX form, one on the left and the other on the right\nof the main crossover. The hard part for me to figure out is the proper spacing in bps between the crossovers, with that\nregard I think it would help if Figure 1B could be redrawn in a more accurate way with the proper number of pairs shown. I\nhave attached a very rough illustration in svg format to help show what I mean, but the details could be wrong since I just\ncounted by eye on the 3D model in 1A.\n2) In the Materials and Methods sections, some important details for the design are not fully presented, such as for the\nmolecular modeling and sequence design optimization. I would like to have it written in more detail how the model in Fig 1A\nwas generated, as I think this step being done right is really the key to making successful nanostructure designs.\n3) The presentation of the cotranscriptional folds in the main figures is potentially misleading. Cotranscriptional folds are\nshown as insets, and annealed folds as the larger views in Figure 2B and 3B–this can be misleading as it can make it\nappear that the cotranscriptional assembly products are much larger than they are in fact. I would much prefer that annealed\nand cotranscriptional products be clearly marked and presented in a way that makes them directly comparable at the same\nscale, as is done in Figure S3.\n4) When I look at the SI AFM panels I can see that design D3 assembles dramatically better than the other designs\ncotranscriptionally. Also cofolded D3 look very different from annealed D3, the cofolded ones are thicker and longer.\nTypically the cofolded structures are smaller, but in this case the cofolded structures are larger than the annealed. Of the\ndesigns tested, it looks like D3 assembles into the comparatively largest assemblies cotranscriptionally. It is also notable that\nadding the fluorescent aptamer in D11 and D12 ended up dramatically shortening the assemblies. It would be useful to\nhave some quantification of the length of the fibers in the different experiments.\n5)The PC-tile is closely related to the classic PX tile created by Afonin et al. (JACS 2008), but it differs in an important way\nthat enable the creation of a one-stranded design, so I think you should cite the original work. I think the PC-tile design\npresented here is quite elegant, but I believe it would help to compare its design to other PX assemblies in RNA so it is more\nclear what the architectural development is. In particular, the authors might compare/contrast their design to the larger\nannealed ssOrigami designs of their previous work in Han et al (Science 2017) - is it simply that these tiles are much smaller\nthat allows them to fold cotranscriptionally? –or is there some other feature of the design that is helping to contribute to their\nbetter folding? Also, I'm surprised the authors don't reference their own work on the PX motif, as it's very relevant to the\ncurrent work.\nAnother very recent PX assembly you might not be aware of– a nano-triangle (Sampedro et al. Small 2023), has also been\ncreated using a 6bp rather than 8bp paranemic interaction, and because of the different length of the pseudoknotted\ndomains the helices are not co-planar as they are in your PC-tile design.\n6) Figure 5 vs Figure S67. I feel like the diagramming in S67 is helpful, but the comparison AFM images shown are from\nannealed samples, not cotranscriptional samples. Also after staring at the image in S67 quite a bit, I find I disagree with the\nextent of the arrays drawn. While I admit that interpreting these is a bit of an art form, I think the drawing is really over\noptimistic. A better comparison would be to show the cotranscriptional AFM images here instead of the annealed\nAFM. That being said, I still find the cartoon representations aligned to the EM data to be tremendously helpful compared to what is\nmore conservatively shown in Fig 5. I think finding some middle ground between the two styles might be ultimately the best.\n7) Can you better explain why are the PC-Tile assemblies being localized to the nucleus? If we don't know for sure, at least\nsuggest what experiments would really be needed to reveal/explore this area better. I think the main result of the paper is\nalready solid, that the RNA nanostructures are both well-folded and present in the nucleus. However, the explanation for\nwhy they are retained in the nucleus compared to other similar size RNA assemblies in the literature that end up in\ncytoplasm. Do you think based on this you can now rationally control the localization to either the inside or outside the\nnucleus based on a single design change? Or more specifically, can you devise a control variant of your PC-tile that\nlocalizes to the cytoplasm instead? I think it could greatly improve the impact of the work if you are able to demonstrate what\nis the structural factor that decide if an RNA is retained or exported from the nucleus.\n8) What is the effect of the RNA origami on the health of the transfected human cells? Does the expression of RNA\nnanostructure make the cells sick, grow at a much slower rate, or eventually lyse due to RNA structures? It seems like\nforming giant nanonets in the cell nucleus would be detrimental to the cell in a number of ways. If the cells are not negatively\naffected by the RNA expression this would be worth mentioning. Since the resulting origami-TEM remind me a bit of stress\ngranules, I had wondered if they might produce as a result stress-like effect in the cells.", "reviewer_4": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature\nCommunications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career\nResearchers who co-review manuscripts."}, "author_response": "We thank reviewers for their positive feedback and recognition of the contribution of our work, as\nwell as for the thoughtful and constructive comments that helped us improve the manuscript. We\ntruly appreciate it. We have completed the suggested experiments and revised both the\nmanuscript and supporting information accordingly. The details are summarized below.\nReviewer #1 (Remarks to the Author):\nReview of \"Designer RNA Nanostructures Co-Transcribed and Self-Assembled inside Human Cell\nNuclei\"\nChang et al. present a research study on the design and characterization of RNA nanostructures\nthat can be expressed and folded inside human cells. The study uses the RNA paranemic\ncrossover (PX) motif for cotranscriptional foldning, demonstrates new design concepts such as\nwireframe PX grids and curvature, and characterize the PX wireframe grids inside human cell\nnuclei. Compared to previous work on expression of RNA nanostructures in bacteria, yeast and\nhuman cells (Li et al. 2018; Jepsen et al. 2018; Liu et al. 2020; Nguyen et al. 2022; Pothoulakis\net al. 2022; Zhang et al. 2024), what stands out in the present study is the direct visualization of\nRNA nanostructures in cells by Transmission Electron Microscopy (TEM) and is therefore a\nsignificant contribution to the field. However, there are several issues in the description of pervious\nwork, and in the presentation and analysis of experimental results that needs to be addressed\nbefore publication of the work.\nMajor comments:\n1. The manuscript seems to be overselling that the RNA nanostructures are localized in the\nnucleus, since this is what is to be expected when using the U6 promoter. It is, however, different\nfrom the previous study (Zhang et al. 2024), which was also using the U6 promoter, and found\nthe RNA nanostructures in the cytoplasm. The manuscript would benefit from a better description\nof RNA expression strategies with references to previous literature and a discussion of the likely\nreason for the different localizations observed in the current and previous study. Please also\nprovide a plasmid map in the supplementary figures (Figure S72-S74). The Previous study (Zhang 2024) used the same cell\nline, the same transfection method, and similar cloning strategies, albeit a different vector. We\nconducted a set of experiments (see details below) and concluded that several potential factors\nmay influence the localization of RNA nanostructures. A more comprehensive study should be\nconducted to provide a deeper understanding of this matter. In this work, by following the\nsuggestions from all reviewers, we provide our current understanding based on the experimental\ndata obtained.\nIn our manuscript, we tested Designs 4 and 10 (D4 and D10) for in vivo folding and observed their\nlocalizations. To have a more comprehensive view, we reanalyzed more images of D4 and D10\nand conducted a statistical study to demonstrate the RNA distribution within different cellular\ncompartments of the transfected cells, including cytoplasmic-only, cytoplasmic and nuclear,\nand nuclear-only localizations. The results are included in new supplementary figures (Figure S69-\nS71). The data showed that 68.50% of mCherry-expressing cells formed D4 assemblies, and\n67.09% formed D10 assemblies (Figure S69). Among the mCherry-positive cells, nearly half of\nthe cells showed nuclear-only localization of D4, the other half showed both cytoplasmic and\nnuclear localization, and less than 5% showed cytoplasmic-only (Figure S70). For D10, nearly 90%\nof cells showed both cytoplasmic and nuclear localization (Figure S71). Notably, D4-expressing\ncells exhibiting nuclear and cytosolic localization typically showed large nuclear assembly\naccompanied by small cytosolic aggregates, as seen in representative images. In contrast, D10-\nexpressing cells with nuclear and cytosolic localization displayed three predominant patterns:\na large nuclear assembly with small cytosolic or pronounced assemblies present in both the\nnucleus and cytosol, or a diffused nuclear and cytosolic signal.\nAdditionally, we have recently established a doxycycline-inducible Design 4 expressing cell line.\nIn that cell line, the Design 4 DNA is integrated into the genome of the cells and is only expressed\nupon the addition of doxycycline. Upon doxycycline addition, the expression of Design 4 Structure\nis 20-fold less than transient transfections. Additionally, the generated structures are significantly\nsmaller and detectable in both the nucleus and cytoplasm at 48 hours post-doxycycline addition\n(Data not shown). We have not yet tested whether prolonged stable expression will result in large\nstructures, as shown in this manuscript. But the current data suggest that the predominant nuclear\nlocalization of Design 4 and Design 10 is likely due to the rapid local transcription and assembly\nthat occur under transient transfection conditions.\nWe further examined the previous work (Zhang, 2024) and compared our design with theirs. In\naddition to their smaller-sized structures compared to our design and different design strategy\n(relying on stem-loops and kissing loop interactions), we also identified a possible sequence issue:\nthe lack of a termination sequence directly after their structure design, and the presence of\nmultiple polythymine tracks within their design. We used a well-characterized pLKO1 plasmid and\nincluded a poly T sequence immediately at the end of the ssRNA structure designs. It has been\npreviously shown that RNA polymerase III requires a poly T tract of at least 4 thymine’s to achieve\n75% termination rate of the transcription, 5 thymine’s for a 95% accurate termination, and 6\nthymine’s 99-100% (Gao Z, Herrera-Carrillo E, Berkhout B. Delineation of the Exact Transcription\nTermination Signal for Type 3 Polymerase III. Mol Ther Nucleic Acids. 2018 Mar 2;10:36-44. doi:\n10.1016/j.omtn.2017.11.006. Epub 2017 Nov 21. PMID: 29499947; PMCID: PMC5725217).\nWithout a poly T tract at the end and with a poly T stretch inside the design, the resulting\ntranscripts may vary in length, which would affect the correct folding and assembly rate of the\ntarget structures and, therefore, the localization of the structures.\nTo further investigate the possible structural factors that impact RNA structure localization, we\ncreated a plasmid carrying Design 3, a smaller, linear structure. Noted that both Designs 4 and\n10 are large 2D array assemblies. We confirmed the expression of Design 3 in the cells after 24-\nhour transfection with RT-qPCR. Based on our results, we didn’t directly observe its long linear\nassembly in the cells (see immunofluorescent images in reply to Reviewer 2, question #2).\nUnfortunately, although Design 3 is expressed at similar levels compared to Design 4 (see Figure\nin reply to Riewer 2, question #2 and Figure 4B), it contains a track of 4 and 5 thymines in two\ndifferent locations within the sequence, leading to the possible transcription of three different\ntranscripts. The smaller linear size as well as the non-ideal sequence design in Design 3, would\npotentially interfere with prominent assembly.\nIn summary, all these data and analyses indicate that the factors influencing the localization of de\nnovo designed RNA may be complex and require comprehensive investigation in a separate study.\nSo far, we have identified several possible factors, including structural factors (e.g., size,\ngeometry), sequence design (poly T), and the transfection method (different expression levels\nand transcription rates in transient transfection compared to stable cell lines). It may also involve\nother factors that we have not yet studied. Therefore, in this current work, we focused on the\nparanemic-cohesion-based co-transcriptional RNA folding strategy, and the discussion of\nlocalizations centered on D4 and D10, emphasizing the specific design and experimental\nparameters used to achieve large assemblies in the nucleus. We have also updated the\ndiscussion of the factors that impact the localization of RNA in the revised manuscript.\nFigure S69. Assembly efficiency of D4 and D10 at 24 hours post-transfection. Representative\nfluorescence images and quantification of mCherry-positive cells showing assembled D4 or D10 structures\nat 24 hours post-transfection. Bar graphs indicate that 68.50% of mCherry-expressing cells formed D4\nassemblies and 67.09% formed D10 assemblies. Scale bar: 50 μm.\nFigure S70. Subcellular distribution of D4 assemblies at 24 hours post-transfection. Representative\nimages and quantification showing the localization of assembled D4 structures in mCherry-positive cells.\nCells were categorized based on the subcellular localization of the assemblies as nucleus-only, nucleus\nand cytosol, or cytosol-only. Scale bar: 5 μm.\nFigure S71. Subcellular distribution of D10 assemblies at 24 hours post-transfection. Representative\nimages and quantification showing the localization of assembled D10 structures in mCherry-positive cells.\nCells were classified into nucleus-only, nucleus-and-cytosol, or cytosol-only categories. Scale bar: 5 μm.\n2. The description of the design of the RNA nanostructures is not clear. In the text the design is\ndescribed as “carefully conducted” (page 3) and “carefully designed” (page 4), but that is not a\nscientific way to describe the design process. Please explain in general terms in the main text\nhow the RNA nanostructures were designed and reference the Material and Methods. The current\ndescription of the design process in the Material and Methods is not clear and should contain\nenough details so that other researchers can reproduce your design approach. Please explain\nhow the initial sequence was generated by Tiamat (and remember a reference) and explain the\nprocedure to generate sequences for co-transcriptional folding and the rationale behind what\nsounds like a manual procedure. As a sidenote, NUPACK and Mfold is not a “simulation” but a\nsecondary structure prediction using minimum free energy algorithms.\nResponse: We thank the reviewer’s suggestion. In the revised manuscript, the general\ndescriptions of how we designed our RNA tiles have been added to the Results section of the\nmain text:\n“To design a tile, generally, we first determined the overall structure of an RNA tile, including the\nnumber and locations of paranemic cohesions. Each RNA PC domain contains top two strands and\nbottom two strands, forming stable secondary structural units. RNA KLs were incorporated at the\nterminals not only for the single-stranded routing but also for encoding the matching rules between\ntiles, enabling programmable formation of lattices and arrays. Figure 1A shows an example of\ndesigning a ssRNA tile containing two PC domains and two KL domains. We planned the routing\nof the single-chain RNA by introducing an additional parallel crossover at the center of the tile to\nbridge the two helices, allowing us to connect all domains Into a single continuous circular\nstrand. Then, we chose the location of the 5’ end to initiate transcription, with the formation of stem\nloops, by creating a nick position on the circular strand.\nAfter the structural design, the sequence design of a tile was conducted by following several steps.\nGenerally, we first assigned the sequences of certain nucleotides at key locations, such as 5’ end,\nkissing loops, and PC domains. Then, we will generate sequences domain by domain from 5’ down\nto the 3’ end (steps 1 to 4 in Figure 1C) to ensure the formation of the correct secondary structure\nof each stem loop and to allow these domains to further interact through paranemic cohesion to\nform the overall shape (detailed sequence assignment protocol in “Materials and Methods”).”\nDetailed descriptions of RNA tile design and sequence assignments have been added as\n“Structural design of RNA tiles” and “Sequence assignment of RNA tiles” in the Materials and\nMethods section.\n3. The manuscript is not referencing previous literature sufficiently. Here are some examples: (1)\nIt would be appropriate to reference previous studies describing the expression of RNA\nnanostructures in cells (Li et al. 2018; Jepsen et al. 2018; Liu et al. 2020; Nguyen et al. 2022;\nPothoulakis et al. 2022; Zhang et al. 2024). (2) The following sentence: “Unlike previously reported\nco-transcriptionally assembled RNA nanostructures, which rely on branch-kissing loops or\nantiparallel crossover interactions, our design employs parallel crossover interactions, a structural\nstrategy not previously explored in this context.” Is not describing that the paranemic crossover\nhas been used before for co-transcriptionally assembled RNA nanostructures in the paper by\nVallina et al. (Vallina et al. 2023). (3) In relation to the RNA paranemic crossover it would also be\nappropriate to mention the original study of this motif (Afonin et al. 2008). (4) References are\nmissing for previous studies on curvature on page 3 and KL sensitivity to buffer conditions on\npage 4. (5) Concerning the description of the RNA biosensing approach using split Broccoli,\nprevious literature that used similar approaches should be referenced e.g., (Rogers et al. 2015;\nAlam et al. 2017), and the results should be compared.\nResponse: The corresponding references have been included in the revised manuscript.\nSpecifically,\n(1) The suggested six references were added to the second paragraph of the Introduction section\nas references #7-12 in the main text.\n(2) The suggested paper was cited as reference #18 and the sentence was updated as:\n“Our design employs parallel crossover, a structural feature that was initially developed in DNA\nnanotechnology13, 14, later adapted to RNA structures15, and recently used to fold single-chain DNA\nand RNA16-18.”\n(3) The suggested paper was added in the same sentence above as reference #15.\n(4) The curvature-related references were added as reference #19-20 on page 3, and the papers\nfor KL sensitivity to buffer conditions were cited as reference #21 on page 4.\n(5) The requested references were cited as references #30-31, and we added the following writing\nin the revised manuscript.\n“Although the current sensing experiments are conducted in a cell-free synthetic system, previous\nstudies of RNA biosensing using split Broccoli30, 31 suggest that this sensor design potentially can\nalso be adapted in living cells for visualizing and monitoring RNA assembly.”\n4. There is an issue with the Mg-dependence experiments, since TAE buffer is used to make the\ndifferent dilutions. However, TAE contains EDTA which chelates Mg2+ and therefore the Mg2+\nconcentrations are likely misrepresented. Since EDTA chelates 2 Mg ions it might be possible to\ncalculate the effective concentration of Mg2+ in your experiments but it would be best to repeat\nexperiments without EDTA by using a TA buffer for dilitions. It would be appropriate to reference\nprevious papers that describe the Mg dependence of KLs and compare them to your results. In\nregard to Mg dependence, you assume that the PX tiles does not require Mg, but it is known that\nMg can be important for stabilizing crossovers and for alignment of helices. To my eye, the\nmonomer tiles look a bit distorted in the AFM images, which I think should be addressed in the\ndiscussion of your results.\nResponse: We thank the reviewer for bringing this issue to our attention. A set of new Mg-\ndependence experiments in TA buffer without EDTA was conducted. The results showed that,\nwithout EDTA interruption, nanorings began to form in buffers containing Mg2+ at a concentration\nof 0.5 mM. We added these new data as Figures S25-27 in the revised supporting document and\nupdated the related main text of the manuscript.\nBy comparing the previous papers of studying the Mg dependence of KL (reference #21) and the\nstability of PC-based RNA (reference #22), we added the following paragraph as well as the\ncorresponding references:\n“We first test directly annealed D8 samples in a set of TA buffers containing 0, 0.5, 1, 2, 4, or 6 mM\nMg2+. The nanoring formation started to be observed when the Mg2+ concentration reached 0.5 mM\nor higher (Figure S25-S27). Notably, D8 monomers folded into the target tile shape with some level\nof deformations even under 0 mM Mg2+, as confirmed by AFM imaging, indicating that individual\nssRNA tiles can fold across a broad range of ionic buffer conditions. This data aligns with the\nprevious report that ssRNA origami based on paranemic cohesion can be stably stored in\nphosphate-buffered saline (PBS) without the addition of Mg cations for months, remaining intact22.\nIn addition, KL-mediated ring formation required elevated Mg2+ levels, suggesting that KL\ninteractions for higher-order assembly are sensitive to ionic strength. Similar results were also\nreported in a previous study21, where stem-looped KL tiles were used to study their assembly under\n0, 1, and 5 nM of Mg2+, and the results showed that conformation of the higher-order RNA\nassemblies can be stabilized by the addition of magnesium.”\nFigure S25. AFM images of D8 after purification and annealing in TA buffer containing 0 and 0.5 mM Mg2+.\nWithout EDTA, ring formation is observed in the buffer containing Mg2+ down to 0.5 mM.\nFigure S26. AFM images of D8 after purification and annealing in TA buffer containing 1 and 2 mM Mg2+.\nFigure S27. AFM images of D8 after purification and annealing in TA buffer containing 4 and 6 mM Mg2+.\n5. In the Split-Broccoli sensor experiments you write that “5 and 10 nM target RNA, indicating a\nlow detection threshold” but you lack an evaluation of the statistical significance of this detection\nthreshold. It is not clear how many independent experiments were done, and the standard error\nbars seem very small for independent experiments. The sensor should be tested in more detail\nand the sensitivity reported in relation to target to sensor ratio. In Fig. 3CDE, it is not clear what\nthe fluorescence intensity is normalized to and why the fluorescence is not directly reported. In\nFig. 3E, the shading is confusing in the bar plot and turning the bar plot on its side is not necessary.\nPlease show the M1-M7 mutations directly in the figure, which will help the reader to understand\nthe effect of the mutations and their closeness to the junction. You write “M1 to M7 contained 12nt,\n8nt, 2nt, and 3nt substitutions, respectively, and produced fluorescence levels comparable to the\nnegative control.” This is clearly not true, since M5-M7 shows some fluorescence. Finally, the\ndescription of “moderate single-nucleotide specificity” is unclear. Can you be more quantitative?\nResponse: We thank the reviewer for providing these valuable comments. We have reorganized\nthe discussion of Split-Broccoli sensor experiments and corrected the typos in the original version\nof the manuscript. Below are point-to-point responses to each comment.\nTo evaluate the statistical significance of the detection threshold (5 and 10 nM target RNA), we\nadded the P-value test for them. The results showed that the P < 0.0001 for both of 5 and 10 nM\ntarget RNA compared with the control (0 nM target RNA).\nWe added the corresponding clarification in the legend of Figure 3 that six independent\nexperiments were done for each sample. All the samples contained the same amount of RNA\ntranscripts, which were split from the same reaction mix and split at the same volume. They all\nshared the same tube of DHFBI-1T dye. The normalized fluorescent intensities were summarized\nin the following table.\nTarget RNA concentration / nM Normalized Er±\n0 1 0.00987\n1 1.12919 0.02023\n2.5 1.27044 0.0207\n5 1.73541 0.03056\n10 2.46685 0.0475\n25 5.34942 0.37936\n50 7.8615 0.22388\n100 12.79514 0.65317\n200 19.20402 0.58068\n300 23.46898 0.60779\n400 26.39364 0.76456\n500 29.10901 0.52838\nThe detailed description of how sensors and targets were experimentally tested was added in the\nrevised Materials and Methods. Specifically, two paragraphs were added under the section of\n“Fluorescence measurements”:\n“Sample preparation: Experimentally, D13 was cotranscribed with HiScribe® T7 High Yield RNA\nSynthesis Kit (NEB) for 1h. Then, the reaction mix without further purification was directly diluted\nto 0.5 µM before the specificity testing, and the concentration of D13 was calculated based on the\nOD260 reading of Nanodrop. Note, the in vitro transcripts always contain elongated or shortened\nbyproducts, so that the calculated concentration is higher than the concentration of pure D13, and\nthe accurate concentration of D13 was not quantified.\nM1 to M7 are seven RNA mutants derived from our multistranded RNA tile study8. Unpurified\ncotranscriptional products of D13 (0.5 µM) were mixed with each RNA target (0.5 µM, purified) and\nDHFBI-1T (10 µM), and annealed at 37 oC in a thermocycler for 30 minutes.”\nThe fluorescence intensities in Fig. 3CDE were normalized to the corresponding negative control\n(Sensor D13 with DHFBI-1T dye), as the normalization results directly reflected the increase in\nfluorescence compared to the negative control. The detailed description of the normalized\nfluorescence intensity was provided in the figure legend. The original unnormalized fluorescence\nintensity data were summarized in the source data.\nWe agree that shading in the bar plot is not necessary and have rearranged the color and\norientation of the bar plot in the revised Figure 3E. The M1-M7 mutation information was originally\ndetailed in Figure S45 of the supporting information. Following the reviewer’s suggestion, we also\nadded the sequence information of the mutants to the main figure as Figure 3H.\nThere was a typo for the “M1 to M7 contained 12nt, 8nt, 2nt, and 3nt substitutions, respectively,\nand produced fluorescence levels comparable to the negative control.” It was corrected as “M1\nto M4 contained 12nt, 8nt, 2nt, and 3nt substitutions, respectively, and produced fluorescence\nlevels comparable to the negative control.”\nFinally, we removed the description of “moderate single-nucleotide specificity” since it is confusing.\nIn the revised manuscript, we describe the specificity of our sensor as:\n“This partial activation likely results from the location or minimal number of mismatches, which may\npermit partial hybridization to facilitate the aptamer core formation, suggesting that the specificity\nof our sensor is also influenced by structural factors of the split aptamer in addition to the number\nof mismatches.”\n6. The experiments showing expression of RNA nanostructures in cells needs some more details.\nWhy does the mCherry expression look different in the top panels and the bottom panels of Fig.\n4C? In the top panels the expression is mainly in nucleus, and in the bottom panels the expression\nis also in the cytoplasm. Is the expression time different between these experiments and could\nthis influence the results?\nResponse: In the co-transfection experiments, mCherry is a separate plasmid and was used as\na marker to visualize transfected cells and distinguish them from the non-transfected cells. It was\npreviously shown that co-transfection of two plasmids results in a high percentage of the cells\nreceiving both plasmids, however, in varying stoichiometries. Please refer to the following two\npapers.\n• Georg Hannig & Cordula Jany, 2013, Co-Transfection of Plasmid\nDNA, BioTechniques, 54:1, 47, DOI: 10.2144/000113979,\n• Xie et al. (2013) Co-transfection and tandem transfection of HEK293A cells for overexpression\nand RNAi experiments. Cell Biol Int 35(3):187–192. https://doi.org/10.1042/CBI20100470.\nThe expression of mCherry is irrespective of the expression of the structures; it is solely a mark\nto localize transfected cells.\nOur experiments were conducted 24 hours after transfections. The apparent difference in\nmCherry signal between the top (Fixed cells) and bottom (Live cells) panels is an artifact of fixation.\nFixation of cells with paraformaldehyde results in decreased signal from fluorescent protein.\nPlease see the figure below from Schnell, U., Dijk, F., Sjollema, K. et al. 2012, who show GFP\nprotein localization in live cells (Red arrow) and after permeabilization and fixation with\nparaformaldehyde (Yellow arrow). Although they did not test mCherry, it is expected that it will\nbehave the same way as they are both of similar size and structure\n• https://www.rcsb.org/structure/2H5Q – mCherry crystal structure\n• https://www.rcsb.org/structure/4KW4 – GFP crystal structure)\nPUBLISHED DATA FIGURE REDACTED\n(Schnell, U., Dijk, F., Sjollema, K. et al. Immunolabeling artifacts and the need for live-cell\nimaging. Nat Methods 9, 152–158 (2012). https://doi.org/10.1038/nmeth.1855 ).\n7. You write that “Overlay of the AFM reference regions onto TEM micrographs further confirmed\nthe accurate nuclear assembly of D4 in human cells (Figure S67 Supp. Figure 67).” But you forget\nto mention that this was also done for D10. In Fig. S67, the drawing on top of the TEM image\nseems like an overinterpretation, where you draw more than you see. It would be more correct to\njust draw the lines that you actually see, then measure the distance between the lines, and\ncompare this measurement to the AFM data. If the distance match, you have a good indication\nthat you are looking at the same thing. The connections between lines that you cannot see from\nthe TEM image, you can only see from the AFM. Thus, you can measure these distances in the\nAFM images and compare them to the theoretical model.\nResponse: Thanks for the suggestions. By considering the different opinions from the reviewers,\nwe removed all the artificial drawing of these images and just measure the distance between the\nvisible lines (yellow marks in the figure below). A revised Figure S67 was updated.\nFigure S67. Distance measurement (yellow lines) and comparison between cell EM and RNA AFM\nimages.\n8. There is an issue with the text about data availability, where you write “???” and have not made\nthe Source data files available. The Source data files should be uploaded and made available to\nthe reviewers, so that we can properly evaluate the quality of the data presented.\nResponse: Source data files are provided.\nMinor comments:\n9. Fig. 1B and Fig. 1C are not comparable and there is confusion about strand direction and\ncolouring scheme. I see from Fig. S7 that you have the right strand synthesis direction in Fig. 1C.\nBut you need to use the colour gradient consistently. E.g. use 5’ end as blue and 3’ end as red,\nwhich corresponds to the standard rainbow colour scheme.\nResponse: The color scheme in Figures 1B and 1C has been updated to align with each other,\nwhere the RNA strand starts at the 5’ end in blue color and ends at the 3’ end in red color.\n10. In Fig. 2 and 3 I think it is confusing to have cotranscriptional (CT) AFM images as an inset\non top of heat-annealed (HA) AFM images, since an inset normally shows a zoom on details of\nthe bigger picture. One possibility is to show them side-by-side and then label them clearly in the\nfigure as “CT” and “HA”. Another option is to just show the CT data and leave the HA data in the\nsupplementary information, since CT is the most relevant for the in vivo study.\nResponse: We appreciated the suggestions, and the AFM images of co-transcriptional\nassembled RNA have been used to replace the original heat-annealed images in Figures 2B and\n3C.\n11. Fig. 3A is confusing to read: The first part with the test tube in not informative and the branched\narrow is misleading. Would be better to use the space to show the two different constructs\nindividually: D10 before and after adding the dye and D13 before and after adding the target RNA\nand dye.\nResponse: The schematics in Figure 3A are divided into new Figure 3A and 3D. Figure 3A\nillustrates how the aptamer-carrying RNA designs (e.g., D9-D12) are prepared for fluorescence\nactivation experiments. Figure 3D shows how the sensor design, D13, senses the target KRAS\ngenes. Other details were provided before in the response to question #5.\n12. It is unclear what is shown in Fig. 3B, since you mention in the text that experiments were\ndone without purification and annealing, but annealing data is shown in the figure, so it gives the\nwrong impression and confuses the reader.\nResponse: These misleading AFM images in the original Figure 3B has been updated by using\nthe AFM images of co-transcriptional assembled RNA nanostructures. Please see the revise\nFigure 3.\n13. You write “The unexpected performance of D12 may be attributed to the G-rich sequence at\nthe 5′ end, which could enhance transcription efficiency or folding kinetics. Based on structural\nyield, D10”. The G-rich claim seems like speculation, and you will need to be more precise,\nreference literature, or do experiments to support your claim. Furthermore, I don’t know what you\nmean by “structural yield” and how you evaluate it.\nResponse: Thanks for catching this. We examined our data carefully and found that, in the\nprevious version of Figure 3C, we added the same mass amount of DNA template (10 ng) in each\nsample and normalized the fluorescence to the control sample without DNA template. This is not\ncorrect because we should align these samples to the same molar amount rather than mass due\nto the different molecular weights of the different designs. In the revised manuscript, we calculate\nthe amount of substance (moles) of 10 ng template of each design and normalize the fluorescent\nintensity accordingly. The fluorescent results of D9 to D12 have been updated in Figure 3B. Based\non the new results, D12 has a similar fluorescence as D9. Our previous conclusion of “unexpected\nperformance of D12” is due to this mistake. We really appreciate the reviewer pointing this out.\nAdditionally, the “structural yield” description is not accurate. We removed this sentence. All these\nchanges have been incorporated into the related main text in the revised manuscript.\n“All four designs were transcribed and assembled directly without purification and annealing, and\nAFM imaging confirmed successful formation of the intended assemblies for all four designs (Figure\n3C and Figure S29). D9 and D10 formed 2D lattices, while D11 and D12 assembled into linear\narrays. Fluorescent measurements after 1 hour of transcription revealed that D9 and D12 produced\nstronger signals than D10 and D11 (Figure 3B), indicating possible preferred terminal locations for\nintegrating aptamer sequences.”\n14. D13 is not really a tile, but just a flexible holiday junction, so why does it fold into neat linear\nfilaments. Please highlight this surprising observation and discuss it.\nResponse: D13 is a tile derived from the multi-stranded RNA tile design, TC-RNA-10, in our\npreviously published paper (doi.org/10.1038/s41467-025-58938-5). We modified this design by\nadding KLs to route the whole tile into a single chain and modified the design to fold co-\ntranscriptionally. The structural features, including the holiday junction, remained here. We\nobserved that the tile was able to fold into linear assemblies in both cases. Our hypothesis is that,\nalthough the holiday junction is flexible, two pairs of RNA kissing loops at both ends would lock\nthe tile into the right locations. In addition, the sequence design of the holiday junction will provide\na preferred orientation for it. We also observed that the multi-stranded version of this tile\nassembled better through a surface-assisted assembly strategy, suggesting the substrate may\nalso play a role in platting the structures.\n15. You write “the assembled D13, with unpaired sensing domains hanging out from the tile body”\nbut forget to reference the figure.\nResponse: Figure S40 has been cited in the main text.\n16. There are some issues with language. Please do spelling and grammar check of the document.\nResponse: Thanks for the suggestions. We have checked the revised manuscript for spelling\nand grammar.\n17. There are some issues with “non-specific/scientific writing”. Please try to be concise.\nResponse: We checked our writing by following the suggestion and revised all the suggested\nparts accordingly.\n18. In Fig 4 it does not make sense to show the grey and green DFHBI icon on the images. That\nis an overinterpretation. In Fig. S44 the DFHBI icon should be grey.\nResponse: At the bottom two rows of Figure 4C, we added the DFHBI dye to both control cells\nand the cells transformed with D10. We removed the icon and used DFHBI-1T to indicate the\naddition of this dye in Figure 4C and Figure S44.\nFigure S1. Fluorescence signals of D13 with target RNA when DFHBI-1T is added in real time (left).\nFluorescence signals of D13 with dye when target RNA is added in real time (right). Samples are tested\ndirectly from transcription without purification or annealing.\n19. Fig. S49-S66: Caption about labelling should either be in each figure caption or be made as\na common figure caption for all figures.\nResponse: All the extra paragraphs of Figure S49-S66 have been deleted and have been added\nto the corresponding figure captions.\n20. Please don’t show DNA sequences as triplets, which suggests codons.\nResponse: All the DNA sequences have been changed to a continuous layout in the supporting\ndocument.\nReferences:\nAfonin, K. A., D. J. Cieply, and N. B. Leontis. 2008. ‘Specific RNA self-assembly with minimal\nparanemic motifs’, J Am Chem Soc, 130: 93-102. https://doi.org/10.1021/ja071516m\nAlam, K. K., K. D. Tawiah, M. F. Lichte, D. Porciani, and D. H. Burke. 2017. ‘A Fluorescent Split\nAptamer for Visualizing RNA-RNA Assembly In Vivo’, ACS Synth Biol, 6: 1710-21.\nhttps://doi.org/10.1021/acssynbio.7b00059\nJepsen, M. D. E., S. M. Sparvath, T. B. Nielsen, A. H. Langvad, G. Grossi, K. V. Gothelf, and E.\nS. Andersen. 2018. ‘Development of a genetically encodable FRET system using fluorescent\nRNA aptamers’, Nat Commun, 9: 18. https://doi.org/10.1038/s41467-017-02435-x\nLi, M., M. Zheng, S. Wu, C. Tian, D. Liu, Y. Weizmann, W. Jiang, G. Wang, and C. Mao. 2018.\n‘In vivo production of RNA nanostructures via programmed folding of single-stranded RNAs’,\nNat Commun, 9: 2196. https://doi.org/10.1038/s41467-018-04652-4\nLiu, D., C. W. Geary, G. Chen, Y. Shao, M. Li, C. Mao, E. S. Andersen, J. A. Piccirilli, P. W. K.\nRothemund, and Y. Weizmann. 2020. ‘Branched kissing loops for the construction of diverse\nRNA homooligomeric nanostructures’, Nat Chem, 12: 249-59. https://doi.org/10.1038/s41557-\n019-0406-7\nNguyen, M. T. A., G. Pothoulakis, and E. S. Andersen. 2022. ‘Synthetic Translational Regulation\nby Protein-Binding RNA Origami Scaffolds’, ACS Synth Biol, 11: 1710-18.\nhttps://doi.org/10.1021/acssynbio.1c00608\nPothoulakis, G., M. T. A. Nguyen, and E. S. Andersen. 2022. ‘Utilizing RNA origami scaffolds in\nSaccharomyces cerevisiae for dCas9-mediated transcriptional control’, Nucleic Acids Res, 50:\n7176-87. https://doi.org/10.1093/nar/gkac470\nRogers, T. A., G. E. Andrews, L. Jaeger, and W. W. Grabow. 2015. ‘Fluorescent monitoring of\nRNA assembly and processing using the split-spinach aptamer’, ACS Synth Biol, 4: 162-6.\nhttps://doi.org/10.1021/sb5000725\nVallina, N. S., E. K. S. McRae, C. Geary, and E. S. Andersen. 2023. ‘An RNA Paranemic\nCrossover Triangle as A 3D Module for Cotranscriptional Nanoassembly’, Small, 19: e2204651.\nhttps://doi.org/10.1002/smll.202204651\nZhang, Tianqing, Xinmin Qian, Huangchen Cui, Qingting Li, Xinyu Du, Xiaoyu Fu, Hongzhou\nGu, Wenwen Zeng, and Bryan Wei. 2024. ‘Programmable in vitro and in vivo folding of single-\nstranded nucleic-acid wireframe origami’, Chem, 10: 2550-63.\nhttps://doi.org/10.1016/j.chempr.2024.04.023\nBest regards,\nEbbe Sloth Andersen\nReviewer #2 (Remarks to the Author):\nThe authors developed a new RNA nanostructure design strategy in the study. RNA paranemic\ncohesion were introduced in the folding of individual RNA tiles, and 180° RNA kissing loops,\n120° RNA kissing loops, and RNA branch kissing loops were designed for intermolecular RNA\ntiles interactions. They demonstrated various designs and the AFM images confirmed the\ndesired self-assembly. Then the authors induced broccoli aptamer to their structures in\nbiosensor applications. Importantly, these RNA nanostructures could be co-transcriptionally\nproduced and assembled in human cells with a nucleus localization.\nSpecific points for improvement are listed as below.\n5. In figure 3C, the sample D12 showed the highest fluorescence intensity. When it came to\ndetect RNA and its mutants, why D13 instead D12 was used? It would be great if the\nauthors can clarify.\nResponse: We appreciate the reviewer for pointing out this. We apologize that we didn’t clarify\nthe reason for choosing D13 instead of D12 for RNA sensing application. D13 is a tile derived\nfrom the multi-stranded RNA tile design, TC-RNA-10, in our previously published paper\n(doi.org/10.1038/s41467-025-58938-5). We modified this design by adding KLs to route the whole\ntile into a single chain and modified the design to fold co-transcriptionally. The structural features,\nincluding the holiday junction, remained here. We observed that the tile was able to fold into linear\nassemblies in both cases. We added the following sentence in the revised manuscript:\n“Inspired by our previous study, that multi-stranded RNA tiles can incorporate a split Broccoli\naptamer and a target-responsive domain for RNA sensing29, we present a ssRNA tile D13 to explore\nthe sensing applications in vitro.”\n2. In figures 4 and 5, the authors used D4 and D10 to confirm the localization of co-transcription\nin the nuclei of cells, and explained the phenomenon by cargo size and structure rigidity for\nnuclear membrane transport. Other samples of smaller sizes, such as D1 and D3, should be\napplied to prove the hypothesis.\nResponse: By following the suggestion, we created a plasmid carrying Design 3 to test this\nhypothesis. We confirmed the expression of Design 3 in the cells after 24-hour transfection with\nRT-qPCR, as shown in the graph below.\nHowever, we were unable to confidently detect its assembly in the cells. Please see the\nimmunofluorescent images of dsRNA below. Design 3 should form long linear assemblies, and\nwe could not detect any long linear structures.\nHowever, during this review process, we examined several possible factors that contribute to the\nRNA localization. We realized that although Design 3 is expressed at similar levels as compared\nto Design 4 (Figure 4B), it contains a track of 4 and 5 thymine’s in two different locations within\nthe sequence. It has been previously shown that RNA polymerase III, upon encountering a stretch\nof thymine, terminates the transcription, with 4 thymine’s terminating 75% of the transcripts, 5\nthymine’s 95% and 6 thymine’s 99-100%. This would result in Design 3, transcribed as 3 different\ntranscripts, hence potentially interfering with prominent assembly. Additionally, we have recently\nestablished a doxycycline-inducible Design 4 expressing cell line. In that cell line, the Design 4\nDNA is integrated into the genome of the cells and only expressed upon doxycycline addition.\nUpon doxycycline addition, the expression of Design 4 Structure is 20-fold less than transient\ntransfections. Additionally, the generated structures are much smaller and detectable in the\nnucleus and cytoplasm as well at 48 hours post doxycycline addition (Data not shown). We have\nnot yet tested whether prolonged stable expression will result in large structures, as shown in this\nmanuscript. The predominant nuclear localization of Design 4 and Design 10 is likely due to rapid\nlocal transcription and assembly under transient transfection conditions.\nA more comprehensive discussion was provided above in the reply to Reviewer #1’s question #1.\nWe have adjusted the hypothesis and updated our discussion accordingly in the revised\nmanuscript.\nReviewer #3 (Remarks to the Author):\nIn this study the authors demonstrate a new variation on single-stranded RNA origami tile\nassembly that form programmable RNA nanostructure cotranscriptionally in the nucleus of\nmammalian cells. Stable expression of RNA origami in human cells has been a goal in this field\nfor years, and this result represents a major advancement as the authors demonstrate both the\nexpression of intricate RNA nanostructures in human cells and also localize the structures to the\nnucleus. Also, for the first time we are able to see the finer structure of the RNA assemblies in the\ncell by TEM. The advancement hinges on a new RNA architecture is able to form a two-helix tall\ntile without the internal apical loop-loop interaction that is typically used, instead the authors\ndevise a parallel helix paranemic cohesion (PC) motif to achieve an apparently faster folding\nstructure (based on the assumption that loop-loop assembly is slower than paranemic cohesion).\nThey convincingly demonstrate the formation of cotranscriptional RNA nanonets, rings and\nfilaments with extensive atomic force microscopy (AFM) work. The AFM imaging is very high\nquality, and of sufficient resolution to resolve even minute details of their structural designs.\nNumerous different structural variants are constructed and compared, including distorted\ncurved versions of the tiles, providing a broad overview of the assembly space for the new structural tile.\nThe authors further investigate the effect of integrating light-up aptamers and chemical sensing\nelements into the designs, and were able to test and validate this as well in vivo. Live cell\nimmunofluorescence imaging clearly demonstrates that the RNAs localize to the nucleus of\nhuman cells, further backed up by a photobleaching experiment on the fluorescent aptamer tags.\nLastly, and also most critically for the study, transmission electron microscopy imaging was able\nto reveal some fine structure details of the RNA periodic assemblies in mammalian cell nuclei,\ndemonstrating the formation and retention of programmable RNA nano-assemblies in the nucleus.\nThis is a very nice work, and I recommend the paper for publication in Nature Communications\nafter the following revisions/clarifications are made. Here I list my concerns, which I would like to\nhave addressed before publication:\n5) I feel that the rationale for RNA structure design could still be better explained. I think you\ncould further explain the logic behind this particular strand routing, how did you come up\nwith it? I spent a long time to try to figure it out, if you untangle the PX the secondary\nstructure is a simple X shape. To fold into a tile two 8bp PX form, one on the left and the\nother on the right of the main crossover. The hard part for me to figure out is the proper\nspacing in bps between the crossovers, with that regard I think it would help if Figure 1B\ncould be redrawn in a more accurate way with the proper number of pairs shown. I have\nattached a very rough illustration in svg format to help show what I mean, but the details\ncould be wrong since I just counted by eye on the 3D model in 1A.\nResponse: We thank the reviewer’s constructive suggestions. The schematics in Figure 1B is\nused to show the overall single-strand routing pathway of a single-stranded RNA tile,\ndemonstrating that PX can route such a tile without introducing internal kissing loops. All detailed\nrouting maps with RNA sequences were provided in the supporting figures with details base\ninformation for the PX cohesion, center crossover, and duplexes between crossovers (Figure S5,\nS7, S10, S12, S14, S18, S20, and S22). The tile design that matches the one in Figure 1B is\nlisted in Figure S12.\nFigure S2. Sequence and routing map of D3 tile.\n2) In the Materials and Methods sections, some important details for the design are not fully\npresented, such as for the molecular modeling and sequence design optimization. I would like to\nhave it written in more detail how the model in Fig 1A was generated, as I think this step being\ndone right is really the key to making successful nanostructure designs.\nResponse: New descriptions for the detailed tile design process and sequence assignment\nprotocol were added as “Structural design of RNA tiles” and “Sequence assignment of RNA tiles”\nin the Materials and Methods section.\nIn the revised manuscript, the general descriptions of how we designed our RNA tiles have been\nadded to the Results section of the main text:\n“To design a tile, generally, we first determined the overall structure of an RNA tile, including the\nnumber and locations of parranemic cohesions. Each RNA PC domain contains top two strands\nand bottom two strands, forming stable secondary structural units. RNA KLs were incorporated at\nthe terminals not only for the single-stranded routing but also for encoding the matching rules\nbetween tiles, enabling programmable formation of lattices and arrays. Figure 1A shows an\nexample of designing a ssRNA tile containing two PC domains and two KL domains. We planned\nthe routing of the single-chain RNA by introducing an additional parallel crossover at the center of\nthe tile to bridge the two helices, allowing us to connect all domains Into a single continuous\ncircular strand. Then, we chose the location of the 5’ end to initiate transcription, with the formation of stem\nloops, by creating a nick position on the circular strand.\nAfter the structural design, the sequence design of a tile was conducted by following several steps.\nGenerally, we first assigned the sequences of certain nucleotides at key locations, such as 5’ end,\nkissing loops, and PC domains. Then, we will generate sequences domain by domain from 5’ down\nto the 3’ end (steps 1 to 4 in Figure 1C) to ensure the formation of the correct secondary structure\nof each stem loop and to allow these domains to further interact through paranemic cohesion to\nform the overall shape (detailed sequence assignment protocol in “Materials and Methods”).”\n3) The presentation of the cotranscriptional folds in the main figures is potentially misleading.\nCotranscriptional folds are shown as insets, and annealed folds as the larger views in Figure 2B\nand 3B–this can be misleading as it can make it appear that the cotranscriptional assembly\nproducts are much larger than they are in fact. I would much prefer that annealed and\ncotranscriptional products be clearly marked and presented in a way that makes them directly\ncomparable at the same scale, as is done in Figure S3.\nResponse: We thank the reviewer for pointing this out. By following the suggestion, the AFM\nimages in Figure 2B and Figure 3B have been replaced by the images of the co-transcriptional\nfolded samples (new Figure 2B and new Figure 3C). They were previously included in the Figure\nS4 and Figure S29 of the original submission.\n4) When I look at the SI AFM panels I can see that design D3 assembles dramatically better than\nthe other designs cotranscriptionally. Also cofolded D3 look very different from annealed D3, the\ncofolded ones are thicker and longer. Typically the cofolded structures are smaller, but in this case\nthe cofolded structures are larger than the annealed. Of the designs tested, it looks like D3\nassembles into the comparatively largest assemblies cotranscriptionally. It is also notable that\nadding the fluorescent aptamer in D11 and D12 ended up dramatically shortening the assemblies.\nIt would be useful to have some quantification of the length of the fibers in the different\nexperiments.\nResponse: We believe this difference of AFM images is caused by the AFM sample preparation\nand imaging process. The Co-transcriptional assembly was performed on mica and then followed\nby washing steps for AFM imaging. While the annealed samples were folded in PCR tubes,\ndeposited on mica, and then washing. In addition to the different sample preparation for AFM,\nthere are also some parameters that cannot be fully controlled in this process. For example, we\nscanned multiple areas of one sample, and the collected images can show different “assembly\nyield”. Also, some of the samples are preferred to be attached onto the mica surfaces even by\nusing the same protocol. Therefore, we believe that the “dramatically better” AFM images of D3\nare highly possible just when we obtain the images from a condensed sample area.\nBy following the suggestion of having some ideas of the length of the fibers, we added the\nsupporting figure (Figure S68). We selected a few representative structures and measured their\nlengths. The results showed that D11 may be shorter due to the non-ideal insertion location of the\naptamer sequence.\nFigure S68. Line length measurements of D3, D11 and D12 from AFM images. From left to right are D3,\nD11 and D12. CT represents co-transcriptional folding. PA means purified annealed assembly.\n5)The PC-tile is closely related to the classic PX tile created by Afonin et al. (JACS 2008), but it\ndiffers in an important way that enable the creation of a one-stranded design, so I think you should\ncite the original work. I think the PC-tile design presented here is quite elegant, but I believe it\nwould help to compare its design to other PX assemblies in RNA so it is more clear what the\narchitectural development is. In particular, the authors might compare/contrast their design to the\nlarger annealed ssOrigami designs of their previous work in Han et al (Science 2017) – is it simply\nthat these tiles are much smaller that allows them to fold cotranscriptionally? –or is there some\nother feature of the design that is helping to contribute to their better folding? Also, I’m surprised\nthe authors don’t reference their own work on the PX motif, as it’s very relevant to the current\nwork.\nAnother very recent PX assembly you might not be aware of– a nano-triangle (Sampedro et al.\nSmall 2023), has also been created using a 6bp rather than 8bp paranemic interaction, and\nbecause of the different length of the pseudoknotted domains the helices are not co-planar as\nthey are in your PC-tile design.\nResponse: We appreciate the suggested references and have added the following text in the\nintroduction to including these related works. The JACS 2008 paper was added as reference #15.\nOur previous work (Science 2017) of single-stranded DNA and RNA origami based on PX was\ncited as reference #16. The suggested paper (Small 2023) was cited as reference #1. There is\nalso another related work we published before, which fold PX-based ssDNA and ssRNA into\ntopological structures, was added as reference #17:\n“Our design employs parallel crossover, a structural feature that was initially developed in DNA\nnanotechnology13, 14, later adapted to RNA structures15, and recently used to fold single-chain DNA\nand RNA16-18.”\nParticularly, to compare the current work with our Science 2017 work, the 2017 work formed the\nfoundation for our current work because it solved the single-stranded routing problem by using\nPX. The structures in the 2017 work rely on thermal annealing not co-transcriptional folding, and\ntherefore, the folding pathway design and sequence assignment in this work are significantly\ndifferent from the previous one. In the revised manuscript, the general descriptions of how we\ndesigned our RNA tiles have been added to the Results section of the main text. The detailed\ndescriptions of RNA tile design and sequence assignments have been added as “Structural\ndesign of RNA tiles” and “Sequence assignment of RNA tiles” in the Materials and Methods\nsection. All our structures designed following this process were able to assemble without\nannealing. We never tried if a smaller sized structure in the 2017 work can fold co-transcriptionally,\nwithout using our current folding pathway design and sequence assignment.\n6) Figure 5 vs Figure S67. I feel like the diagramming in S67 is helpful, but the comparison AFM\nimages shown are from annealed samples, not cotranscriptional samples. Also after staring at the\nimage in S67 quite a bit, I find I disagree with the extent of the arrays drawn. While I admit that\ninterpreting these is a bit of an art form, I think the drawing is really over optimistic. A better\ncomparison would be to show the cotranscriptional AFM images here instead of the annealed\nAFM. That being said, I still find the cartoon representations aligned to the EM data to be\ntremendously helpful compared to what is more conservatively shown in Fig 5. I think finding\nsome middle ground between the two styles might be ultimately the best.\nResponse: Thanks for the suggestions. By considering the different opinions from the reviewers,\nwe removed all the artificial drawing of these images and just measure the distance between the\nvisible lines like in Figure 5. A revised Figure S67 was updated.\n7) Can you better explain why are the PC-Tile assemblies being localized to the nucleus? If we\ndon't know for sure, at least suggest what experiments would really be needed to reveal/explore\nthis area better. I think the main result of the paper is already solid, that the RNA nanostructures\nare both well-folded and present in the nucleus. However, the explanation for why they are\nretained in the nucleus compared to other similar size RNA assemblies in the literature that end\nup in cytoplasm. Do you think based on this you can now rationally control the localization to\neither the inside or outside the nucleus based on a single design change? Or more specifically,\ncan you devise a control variant of your PC-tile that localizes to the cytoplasm instead? I think it\ncould greatly improve the impact of the work if you are able to demonstrate what is the structural\nfactor that decide if an RNA is retained or exported from the nucleus.\nResponse: Please refer to the reply to Reviewer #1, Question #1 and Review #2, Question #2.\n8) What is the effect of the RNA origami on the health of the transfected human cells? Does the\nexpression of RNA nanostructure make the cells sick, grow at a much slower rate, or eventually\nlyse due to RNA structures? It seems like forming giant nanonets in the cell nucleus would be\ndetrimental to the cell in a number of ways. If the cells are not negatively affected by the RNA\nexpression this would be worth mentioning. Since the resulting origami-TEM remind me a bit of\nstress granules, I had wondered if they might produce as a result stress-like effect in the cells.\nResponse: We performed a cell viability assay (XTT assay) over the course of 72H after\ntransfections and see no apparent changes in bulk cell viability. XTT is a colorimetric assay that\nmeasures cell viability by assessing the metabolic activity of cells. Viable cells convert the yellow\ntetrazolium salt XTT into a soluble orange formazan dye. The absorbance of formazan is detected\nat 450 nm wavelength. We attribute no differences on bulk population due to nature of transient\ntransfections. Not all cells are transfected, and out of transfected cells 67-68% assemble the RNA\nstructures in varying sizes. (Please see, reviewer #1, question #1). However, we do acknowledge\nand observe that some cells exhibit deformed nuclei and even nuclei rupturing upon too much\naccumulation of the nanostructures, hence we anticipate cells exhibiting large assembly to be\nstressed and apoptotic. In the future, we will construct inducible systems to control the degree of\nassembly.\nReviewer #4 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is\npart of the Nature Communications initiative to facilitate training in peer review and to provide\nappropriate recognition for Early Career Researchers who co-review manuscripts.\nWe sincerely thank all the reviewers for their thoughtful comments and supportive feedback. The\nmanuscript has been significantly improved through the whole revision process, and we truly\nappreciate the reviewers’ efforts in helping us strengthen the quality and clarity of our work. Below\nare our responses to the individual comments:\nREVIEWERS' COMMENTS\nReviewer #1 (Remarks to the Author):\nComments on revision by reviewer 1:\nChang et al. have made additional experiments, revised the manuscript, and answered to all\nreviewer comments to a satisfactory level. I therefore recommend publication and only have one\nadditional comment/suggestion:\nConcerning comment 5, it is still difficult for the reader to understand how the mutations affect the\nstructure. The schematics in Fig. 3E is misleading and not precise. Also, the new Fig. 3H provides\ninformation, but does not help the reader much either to understand how mutations affect the\nstructure. I have made a schematics in an attached pdf file to that shows how one can combine\nshow the information in a simplified but precise manner – by aligning the mutant sequences\nunderneath the structure. So, my suggestion is to combine Fig. 3E and H and to name the\nsequences in the figure. It needs to help the reader understand how the mutations affect the\nstructure and the fluorescence intensity in Fig. 3G.\nResponse: Thank you for this great suggestion and for providing the schematic. We have\nfollowed your suggestion to combine Fig. 3E and 3H into a single schematic. The mutant\nsequences are now aligned underneath the target structure, and their names and mutation sites\nare clearly labeled. This new schematic provides a much clearer visual representation. We\nappreciate your helpful input.\nReviewer #2 (Remarks to the Author):\nThe authors addressed most of the concerns from this reviewer. A publication of this revised\nmanuscript is recommended.\nResponse: Thank you for your constructive suggestions during the review process. We\nappreciate your support and are glad that the revisions addressed your concerns.\nReviewer #3 (Remarks to the Author):\nThe authors have significantly improved their manuscript with their extensive revisions. The text\nreads more clearly, and their elegant design rationale for the primary structural unit is explained\nmuch better. The expanded Materials and Methods, inclusion of plasmid maps, make the\nmethodology more clear and reproducible. All of the issues that were brought up in review were\naddressed.\nI agree the choice to use the helical spacing as a measure in the TEM images is a very good one,\nand it certainly more convincing that the same nanostructures are present in the cell nuclei as the\nin vitro AFM experiments.\nI appreciate the longer explanation on how RNA structure and folding dynamics interplay with\nnuclear trafficking, and I think the explanation provided is a good starting point for further\ninvestigation.\nThe authors demonstrate all claims made in the paper very clearly, and I think the work meets all\nthe requirements for publication. This paper showcases some of the most impressive rational\nRNA design and microscope imaging in our field.\nResponse: We are truly grateful for your thoughtful feedback and generous comments. Your\nrecognition of our structural design, microscopy, and methodological clarity is very encouraging\nto us. Thank you for your support throughout the review process.\nReviewer #4 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is\npart of the Nature Communications initiative to facilitate training in peer review and to provide\nappropriate recognition for Early Career Researchers who co-review manuscripts.\nG--C\nC--G\nU--G\n3’-UAAGCUUUUGUAUUU╯ ╰UUUCUACUCG-5’\n┊┊┊┊┊┊┊┊┊┊┊┊┊┊┊ ┊┊┊┊┊┊┊┊┊┊\nTR 5’-GAGAAAUUCGAAAACAUAAA─GA─AAAGAUGAGCAAAGA-3’\nM7 ....................-U.-.C.............\nM6 .....UAA............-..-........CG.....\nM5 ....................-..-...C...........\nM4 ....................-..-.CUC...........\nM3 ....................-..-.C.C...........\nM2 .........AC.U...AUC.-..-.C.C...........\nM1 .....UAAGC.U.UG.....-..-.....ACUCA....."}]

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