Results

1. AFM: Verification Of The DNA Origami Structure

In our design, the successful assembly of DNA origami was the prerequisite for the function of the DNA origami-based gene editing system. Therefore, verifying the correct assembly of DNA origami was an indispensable foundation for subsequent experiments.

The shapes of DNA origami nanostructures are typically analysed using single-molecule microscopy techniques, the most important being atomic force microscopy (AFM) and transmission electron microscopy (TEM). TEM can yield high-resolution images of single nanostructures but requires sophisticated sample preparation and image analysis. AFM, as a powerful technique to visualise nano- to micrometre-scale structures with subnanometer resolution, has the advantage that it can be operated under atmospheric conditions and even in liquids, and the requirements for sample preparation are minimal (1,2). Hence, AFM imaging was selected to visualise the structural information of the basic plane of DNA origami (DO_PAM).

The DO_PAM nanostructures were assembled in TAE/Mg²⁺ buffer using M13 scaffold, helper strands (S-x, S-PAM-cap-x), and functional strand (PAM-rich) through PCR programs. After purification by filtering three times, the DO_PAM solution was added to the mica sheet to prepare the dry AFM sample. AFM imaging was performed via a tapping mode by using an Asylum Research Cypher ES environmental AFM (Oxford Instruments) equipped with AC-240TSA-R3 tips (f = 70 kH, k = 2 N/m, no tip coating, Oxford Instrument). All images were analysed using the Gwyddion 2.69 and Origin 2025b software. Detailed method and operation procedures can be found in the following protocol (4 Verification of the DNA origami structure by AFM). (AFM handbook)

Nano-sized aggregates with side lengths around 100 nm and heights under 4 nm were considered as the desired DNA nanotiles, as the design specifications for the rectangular DNA origami were 100 nm × 80 nm × 2 nm. In AFM imaging, the DO_PAM maintained the rectangular shape (Figure 1A). Height analysis of a selected single rectangular DO_PAM showed that the length of DO_PAM was around 100nm, the width was around 80nm (Figure 1B), keeping in line with our expectations. These structural values indicated the successful assembly of DO_PAM.

Figure 1. AFM image and Height analysis of a single rectangular DO_PAM. A. A schematic diagram of the designed DO_PAM (100nm × 80nm × 2nm) and an AFM image showing a selected assembled rectangular nanostructure. B. Height analysis for the length (upper) and width (below) of DO_PAM by AFM. The left pictures showed the drawn lines, and the right pictures showed the height variation curve along the line, which changed with the distance. Scale bar: 100nm.

With a larger scanning area and a relatively smaller magnification factor, monodisperse and uniform DNA origami were observed by AFM (Figure 2). In Figure 2, four representative structures were marked. Structure 1 was the correct structure we were expecting, which was shown and analysed in Figure 1. The only less satisfactory aspect was the presence of a small gap in the upper right corner of the rectangle, suggesting that the assembly may not have been fully completed. Structure 2 was a very typical, incomplete assembled rectangle, with a size approximately half that of the normal one. Structure 3 was presumed to be a DNA origami that had completed the loading of all the links but failed to complete the final contraction and shaping steps. Structure 4 was a large cluster, which was hypothesised to be a polymer formed by several DNA nanotiles. Quite noisy background signals were displayed in Figure 2, which were considered to be unfiltered staples. However, due to the relatively high filtration loss rate and the required yield, our team maintained three-time filtration and did not conduct more rounds of filtration or any further purification steps.

Figure 2. AFM images with a larger scanning area. In the figure, four representative structures were circled and numbered in red. Structure 1 was a rectangular shape that exhibited the correct size and a relatively complete structure. Structures 2 and 3 both had the correct height, but they did not present a rectangular shape. Structure 2 was slightly smaller in size, while structure 3 was slightly larger. The height and size of Structure 4 were both relatively large, and under the microscope, it appeared as a large aggregate. Numerous small molecules of a certain height (yellow dots) were widely distributed in the background. Scale bar: 500nm.

2. Loading sgRNA_L/Cas9 Complex On DNA Origami

Fluorescent Modification of sgRNA_L

In the above experiment, we successfully assembled our DNA origami. The next step was to load the sgRNA_L/Cas9 complex onto the basic plane of the DNA origami. To verify the loading of the sgRNA_L/Cas9 complex, we employed a strategy of fluorescently labeling the sgRNA_L/Cas9 complex to assess its successful incorporation onto the DNA origami.

Fluorescein isothiocyanate (FITC, green fluorescence, excitation wavelength = 495nm, maximum emission wavelength = 525nm) is a common fluorescent-modified molecule (comparatively, the reason why we don't directly use EGFP is that it is too large and does not meet the size requirements of our DNA origami structure). There are two ways to modify sgRNA_L. One approach is to modify the amino groups attached to the tail end of the sgRNA_L. The other method is to directly modify the amino groups on the A and C bases (3). The latter method has a lower labeling efficiency but more potential labeling sites, which were selected for use. Meanwhile, to enhance the modification efficiency, we replaced FITC with FITC-NHS ester.

According to Mirus's nucleic acid labelling protocol and previous research (3), the labeling and purification were conducted. The pre-synthesized sgRNA_L and other labeling reagents were mixed and incubated at 37°C for an hour, and the labeled nucleic acid was subsequently purified by the ethanol precipitation method. During the first purification step, the unreacted fluorescein will remain in the supernatant, while the fluorescein-labeled precipitate will appear dark brown. The presence of a white precipitate indicates that the reaction was unsuccessful. This could act as secondary evidence to examine if the labeling is successful. Similarly, the absorbance values at 260 nm and 495 nm were measured using a nanodrop for the product. Details of the procedure can be seen in the protocol (5 Fluorescent modification of sgRNA_L).

In our experiment, during the purification and separation process, we observed a clear separation of solid and liquid phases, as well as a dark brown precipitate (Figure 3). The purified product was also observed to have two distinct absorption peaks at 260nm and 495nm wavelength, respectively corresponding to sgRNA_L and FITC (Figure 4). These results confirmed that FITC successfully labeled sgRNA_L. The molarity of 11.43 μM and labeling efficiency of 6.7 were achieved, reaching the ideal values that satisfy the requirements of DNA origami assembly (15 nM) or fluorescence imaging (Table 1).

Figure 3. Fluorescence during the ethanol purification process. The three pictures from left to right correspond respectively to situations after the first centrifugation, after the supernatant has been removed, and after the second washing solution has been added. The presence of dark brown sediment can be clearly seen in all pictures.

Figure 4. UV absorption spectrum of one example FITC-sgRNA_L product. There were two distinct absorption peaks at 260nm and 495nm, respectively, corresponding to sgRNA_L and FITC.

	Molarity	Volume	Labeling Efficiency	Recovery rate
FITC-sgRNAL	11.43 µM	5 µl	6.70	66.70%

Table 1. Profiles of one example set of FITC-labeled sgRNA_L product. The molar concentration (absorbance/ extinction coefficient) was measured to be 11.43 μM, with a purification yield of 5 μL volume. The labeling efficiency of FITC (molarity of FITC/ molarity of sgRNA_L) was calculated to be about 6.70. The recovery rate of sgRNA_L (mass after purification/ mass before purification) was about 66.70%. Details of the calculation can be seen in the protocol.

Loading of FITC-sgRNA_L/Cas9 Complex

In the preceding experiment, we have prepared the FITC-sgRNA_L/Cas9 complex. The next step is to use this FITC-sgRNA_L/Cas9 complex to verify its successful loading onto the DNA origami.

To confirm the loading of FITC-sgRNA_L/Cas9 complex, we first labeled DNA origami with F-H conjugated with Cy5. We then loaded FITC-sgRNA_L/Cas9 complex onto the Cy5-labeled DNA origami as the experimental group (Cy5-labeled DO_PAMR_FITCC). Two additional control groups were established: one containing only Cy5-labeled DNA origami (Cy5-labeled DO_PAM) and the other containing only FITC-sgRNA_L/Cas9 complex (R_FITCC). As FITC and Cy5 exhibit absorbance peaks at 495nm and 650nm, respectively, we used Thermo Fisher Scientific NanoDrop One 2.12.0 and Python 3.13 to visualize the absorbance spectrum of each group. We then exported the absorbance data for these three groups and plotted the absorption spectrum for visualization (Figure 5). Details of the procedure can be seen in the protocol (6 Loading of FITC-sgRNA_L_Cas9 complex).

Cy5-labeled DO_PAM exhibited two absorption peaks: one at 260 nm, corresponding to DNA, and another at 650 nm, corresponding to Cy5 (Figure 5), indicating the successful Cy5-labeling of the DNA origami. In the group of Cy5-labeled DO_PAMR_FITCC, three distinct absorption peaks were observed: 260 nm for DNA/RNA, 495 nm for FITC, and 650 nm for Cy5 (Figure 5). These results confirmed successful loading of the FITC-sgRNA_L/Cas9 complex onto the Cy5-labeled DNA origami.

Figure 5. The UV absorption spectra of Cy5-labeled DO_PAMR_FITCC, Cy5-labeled DO_PAM, and R_FITCC. Cy5-labeled DO_PAM (red) exhibited characteristic peaks at 260 nm (DNA) and 650 nm (Cy5), while R_FITCC (green) showed a peak at 495 nm (FITC). The combined Cy5-labeled DO_PAMR_FITCC (orange) displayed all three peaks (260, 495, and 650 nm).

3. Loading G-Quadruplex/Hemin On DNA Origami

Dimerization

Up to this point, we have successfully loaded the sgRNA_L/Cas9 complex onto the basic plane of our DNA origami. The subsequent objective was to endow the DNA origami with a membrane-permeabilizing capability. To achieve this, we incorporated G-quadruplexes (G4s) onto the DNA origami through slow annealing. However, G4s have a known tendency to dimerize through stacking interactions (4). Considering spatial limitations and the increased local concentration of G4s on the DNA origami, it was necessary to investigate whether these conditions would promote significant dimerization.

To load G4s onto the DNA origami, we subjected the G4-DNA origami complex (DO_PAMG) to slow annealing, cooling from 95 °C to 20 °C. This served as the experimental group. Two additional control groups were established: the M13 scaffold solution and the DNA origami (DO_PAM) solution. Agarose gel electrophoresis was then employed to assess the extent of dimerization by evaluating the size of the resulting bands and assessing the extent of dimerization. Details are provided in the protocol (7 Dimerization).

Agarose gel electrophoresis analysis revealed the bands of DO_PAMG and DO_PAM with approximate sizes double that of M13, confirming the successful assembly of DNA origami (Figure 6). Notably, no significant difference was observed between the DO_PAM and DO_PAMG groups. Both groups displayed a single distinct band, suggesting that the attachment of G4s did not induce detectable dimerization (Figure 6).

Figure 6. 1% agarose gel electrophoresis analysis of M13, DO_PAM, and DO_PAMG. A. M13 scaffold solution. B. DO_PAM solution. C. DO_PAMG solution.

Loading of Hemin

So far, G4s have been successfully incorporated onto the DNA origami. However, only G4s did not exhibit peroxidase-mimicking activity. Instead, this catalytic activity arose from the formation of G4/hemin DNAzymes. Hence, the aim of the following experiment was to load hemin onto DNA origami and verify its proper incorporation.

To confirm the loading of hemin, we first incubated hemin with the G4-loaded DNA origami (DO_PAMGH) at room temperature, serving as the experimental group. A control group was established, with G4-loaded DNA origami alone (DO_PAMG) serving as a negative control. As hemin exhibits an absorbance peak around 400nm in the presence of DO_PAMG (5), we used Thermo Fisher Scientific NanoDrop One 2.12.0 and Python 3.13 to visualize the absorbance spectrum of hemin-loaded DNA origami. We then exported the absorbance data for these three groups and plotted the absorption spectrum for visualization (Figure 7). Details are provided in the protocol (8 Loading of hemin).

DO_PAMG exhibited a single absorption peak at 260 nm, corresponding to DNA (Figure 7). In contrast, the DO_PAMGH exhibited two distinct absorption peaks: one at 260 nm, corresponding to DNA, and the other around 400nm, attributed to hemin (Figure 7). These results confirmed successful loading of the hemin onto the DO_PAMG.

Figure 7. The UV absorption spectra of DO_PAMG and DO_PAMGH. DO_PAMG (blue) exhibited an absorption peak at 260nm (DNA), while DO_PAMGH (orange) exhibited two absorption peaks at 260nm and 400nm, corresponding to DNA and hemin, respectively.

ABTS Assay: Detecting Catalytic Activity of G4/hemin

With G4/hemin DNAzymes successfully incorporated, the DNA origami was expected to display peroxidase-mimicking activity, which was essential for membrane permeabilization. Accordingly, the core objective of the next experiment was to examine the peroxidase-mimicking activity of G4/hemin DNAzymes on DNA origami.

To evaluate the peroxidase-mimicking activities of G4/hemin DNAzymes, we used 2,2′-azinobis (3-ethylbenzthiazolin-6-sulfonic acid) (ABTS) assay. In this assay, ABTS is oxidised by hydrogen peroxide in the presence of peroxidase, forming ABTS cation-radical (ABTS^•+), a blue-green chromophore with a maximal absorption at 415nm (6) (Figure 8). We designed four groups as follows: (i) buffer, ABTS, and H₂O₂. (ii) hemin, ABTS, and H₂O₂. (iii) free G4/hemin, ABTS, and H₂O₂. (iv) DO_PAMGH, ABTS, and H₂O₂. By measuring the absorbance of the reaction systems at 415nm, we can indirectly evaluate the peroxidase-mimicking activities of G4/hemin. Details of the ABTS assay are provided in the protocol (9 ABTS assay of G4_hemin DNAzymes).

Figure 8. ABTS reaction system. ABTS is oxidised by hydrogen peroxide in the presence of G4/Hemin DNAzymes, forming ABTS cation-radical (ABTS^•+).

The absorbance of the reaction system was monitored at 415 nm, corresponding to the maximum absorption wavelength of ABTS^•+. Group (iv) treated with DO_PAMGH exhibited significantly higher absorbance values compared with those treated with buffer, hemin alone, or free G4/hemin, indicating the enhanced peroxidase-mimicking activity of G4/hemin DNAzymes when assembled on DNA origami (Figure 9).

Figure 9. The absorbance of each group at 415nm. The bar chart showed the average absorbance values obtained from the ABTS assay for various treatments, including buffer, hemin, free G4/hemin, and DO_PAMGH. The DO_PAMGH complex exhibited a significantly higher absorbance compared to all other groups (buffer, hemin, free G4/hemin). Data represented the mean ± s.d. from three independent replicates. Statistical significance was determined using a one-way ANOVA and post-hoc analysis, with **** representing p < 0.0001.

NPN Uptake Assay: Membrane Permeabilization

Having confirmed the peroxidase-mimicking activity of G4/hemin DNAzymes on DNA origami, we sought to investigate the effect of this catalytic activity on the membrane permeability of bacteria.

The hydrophobic fluorescent probe N-Phenyl-1-naphthylamine (NPN), which fluoresces weakly in aqueous environments but exhibits strong fluorescence in phospholipid conditions, has been widely used to evaluate the membrane integrity of bacteria (7). Due to the presence of lipopolysaccharide in the outer leaflet of the outer membrane, the lipophilic fluorescent probe cannot thoroughly permeate the intact bacteria (Figure 10). When the outer membrane is disrupted by permeabilizers, NPN can access the phospholipids and exhibits surges in fluorescent intensity upon excitation at 350nm wavelength (7) (Figure 10).

Figure 10. Schematic representation of NPN fluorescence in bacterial membranes. In intact bacteria, NPN exhibits weak fluorescence due to the lipopolysaccharide layer on the outer membrane, preventing the probe from permeating. However, when the outer membrane is disrupted by permeabilizers, NPN gains access to the phospholipids, resulting in a significant increase in fluorescence upon excitation at 350nm wavelength.

Therefore, to evaluate the efficacy of G4/hemin DNAzymes, we used the NPN fluorescent probe to reflect the permeability of the bacterial outer membrane and to assess the effects of various membrane permeabilizers. We designed four groups as follows: (i) buffer and NPN, and (ii) buffer, bacterial suspension, and NPN, both serving as control groups. (iii) Buffer, bacterial suspension, EDTA Na₂, and NPN were used as the positive control group. (iv) Buffer, bacterial suspension, DO_PAMGH, and NPN were designated as the experimental group. After incubation, we measured the relative fluorescent units under a microplate reader and quantified the membrane permeabilization of each group via fluorescence values after background subtraction and NPN uptake factors. The fluorescence value after background subtraction is defined as the fluorescence value of each group minus the corresponding value of the same group without NPN (Table 2). The NPN uptake factor indicates the ratio of background-subtracted fluorescence values to that of the buffer value (Table 2). Details of the NPN uptake assay are provided in the protocol (10 NPN uptake assay).

In the experimental group (iv) treated with DO_PAMGH, the fluorescence value after background subtraction and the NPN uptake factor were significantly higher than those of the control groups (i, ii) treated with buffer, bacterial suspension alone (Figures 11 and 12). This confirmed the membrane-permeabilizing activity of DO_PAMGH. At the same time, we found that compared to the positive control group treated with EDTA Na₂, the experimental group treated with DO_PAMGH yielded a significant increase in fluorescence value and NPN uptake factor (Figures 11 and 12). This demonstrated the remarkable effect of DO_PAMGH on membrane permeabilization.

Figure 11. Bar graph showing the background-subtracted fluorescence values of four groups. The bar graph showed the average background-subtracted fluorescence values of four groups, including buffer, buffer and cell, EDTA, and DO_PAMGH. The background-subtracted fluorescence values were expressed as relative fluorescent units (RFU). The group treated by DO_PAMGH exhibited a significantly higher RFU compared to all other groups. Data represent the mean ± s.d. from three independent replicates. Statistical significance was determined using a one-way ANOVA and post-hoc analysis, with **** representing p < 0.0001.

Figure 12. Bar graph showing the NPN uptake factors of four groups. The NPN uptake factor indicated the ratio of background-subtracted fluorescence values to that of the buffer value. The NPN uptake factor correlated with the permeability of the bacterial outer membrane in the presence of different treatments, with the experimental group (DO_PAMGH) showing the highest NPN uptake factor compared to the control and positive control groups.

Sample	NPN	Fluorescence value (mean±SD)	Fluorescence value after background subtraction	NPN uptake factor
Buffer	-	191 ± 2.6
Buffer	+	1635 ± 3.3	1444	1
+ Cells	-	836 ± 6.1
+ Cells	+	5564 ± 89.4	4728	3.3
+ Cells + EDTA (1mmol l-1)	-	861 ± 3.3
+ Cells + EDTA (1mmol l-1)	+	6677 ± 18.5	5816	4.0
+ Cells + DOPAMGH (5nmol l-1)	-	884 ± 1.9
+ Cells + DOPAMGH (5nmol l-1)	+	8880 ± 73.3	7996	5.5

Table 2. Fluorescence values and NPN uptake factors obtained in the NPN uptake assay for membrane permeabilization using SpectraMax® iD5 Microplate Reader with E. coli ATCC 25922 as the test organism.

4. Loading Aptamer On DNA Origami

NPN Uptake Assay: Targeting Validation

In addition to the membrane-permeabilizing activity mediated by G4/hemin DNAzymes, we endowed our DNA origami with recognition capability by incorporating DNA aptamers. DNA aptamers are oligonucleotides that bind to a specific target molecule (8). In our project, by adding aptamers to both sides of the DNA origami, its targeting ability can be equipped, allowing the drug to accumulate around the target and minimizing the damage to other cells, such as human cells. The aptamer specifically targeting MRSA was selected and optimized by our team (see Design: Aptamer Screening and Optimization). However, experiments regarding MRSA are not allowed to be carried out in the wet lab. So, the classic and secure E. coli ATCC25922 and a known aptamer obtained from previous research were alternatively used (9).

Theoretically, DNA origami loaded with aptamers (DO^A) would accumulate around E. coli, enhancing the effect of membrane permeabilization. Therefore, we continued to use the lipophilic fluorescent probe NPN to reflect the permeability of the bacterial outer membrane, indirectly determining the targeting ability of DO^A. We established two groups: one treated with G4/hemin DNAzymes on DNA origami (DO_PAMGH) as the control group, and the other treated with G4/hemin DNAzymes on aptamer-loaded DNA origami (DO^A_PAMGH) as the experimental group. Fluorescence intensity was measured using a microplate reader, and membrane permeabilization was quantified by background-subtracted fluorescence values and NPN uptake factors (Table 3). Details of the NPN uptake assay are provided in the protocol (10 NPN uptake assay).

The background-subtracted fluorescence and NPN uptake factor in the DO^A_PAMGH-treated group were significantly higher than those in the DO_PAMGH control group (Figures 13 and 14). This result confirmed the enhanced targeting ability of DO^A_PAMGH, demonstrating the effectiveness of DNA aptamers in facilitating targeted delivery and membrane permeabilization.

Figure 13. Bar graph showing the background-subtracted fluorescence values of two groups. The bar chart shows the average background-subtracted fluorescence values of two groups, including DO_PAMGH and DO^A_PAMGH. The background-subtracted fluorescence values were expressed as relative fluorescent units (RFU). The group treated by DO^A_PAMGH exhibited a significantly higher RFU compared to the DO_PAMGH. Data represent the mean ± s.d. from three independent replicates. Statistical significance was determined using a one-way ANOVA and post-hoc analysis, with **** representing p < 0.0001.

Figure 14. Bar graph showing the NPN uptake factors of two groups. The NPN uptake factor indicated the ratio of background-subtracted fluorescence values to that of the buffer value. The NPN uptake factor correlated with the permeability of the bacterial outer membrane in the presence of different treatments, with the experimental group (DO^A_PAMGH) showing the highest NPN uptake factor compared to the control group (DO_PAMGH).

Sample	NPN	Fluorescence value (mean±SD)	Fluorescence value after background subtraction	NPN uptake factor
+ Cells + DOPAMGH (5nmol l-1)	-	884 ± 1.9
+ Cells + DOPAMGH (5nmol l-1)	+	8880 ± 73.3	7996	5.5
+ Cells + DOAPAMGH (5nmol l-1)	-	885 ± 2.2
+ Cells + DOAPAMGH (5nmol l-1)	+	11791 ± 97.6	10906	7.6

Table 3. Fluorescence values and NPN uptake factors obtained in the NPN uptake assay for targeting using SpectraMax® iD5 Microplate Reader with E. coli ATCC 25922 as the test organism.

LSM: Targeting Validation

To more intuitively visualize the targeting ability of aptamer-loaded DNA origami, we employed confocal laser scanning microscopy (LSM) for validation.

In this experiment, we focused on directly visualizing the targeting of aptamer-loaded DNA origami, without assessing the permeability of the bacterial outer membrane. Therefore, G4s were not loaded onto the DNA origami in this experiment. To visualize the DNA origami, Cy5 was used as a fluorescent label, which could be detected under a fluorescence microscope. Two groups were designed: Cy5-labeled DNA origami (Cy5-labeled DO_PAM) as the control group and Cy5-labeled aptamer-loaded DNA origami (Cy5-labeled DO^A_PAM) as the experimental group. After assembly and incubation with E. coli for 15min at 37°C, the mixture was washed with PBS once and observed under LSM 880 (Zeiss, Germany). The signals without co-localization with the bacteria were regarded as uncleaned Cy5 or Cy5-labeled DNA origami, which were subtracted as background. The signals observed on or around the bacteria were considered to represent the DNA origami that had successfully bound. These areas were selected for mean fluorescence intensity (MFI) quantitative analysis. By comparing and analyzing the fluorescence aggregation around the bacteria under the microscope, we could qualitatively and quantitatively assess the targeting effect of DO^A_PAM. Detailed information can be found in the following protocol (11 Targeting validation by LSM) and handbook (Zeiss Confocal Microscope - LSM880 Handbook).

Under the LSM microscope, the Cy5-labeled DO^A_PAM was obviously observed to be more likely to bind to the surface of the bacteria (Figure 15). MFI analysis further revealed that the MFI of the Cy5-labeled DO^A_PAM group was significantly higher than that of the Cy5-labeled DO_PAM group (Figure 16). These results provided relatively strong evidence for the targeted binding ability of DO^A_PAM, highlighting the effectiveness of the aptamer (targeting component).

Figure 15. Representative confocal microscopy images of E. coli cells after the incubation of Cy5-labeled DO_PAM with or without the modification of the aptamer. Scale bar: 20µm x 20µm.

Figure 16. Quantification of the mean fluorescence intensity (MFI) of E. coli cells after the incubation of Cy5-labeled DO_PAM with or without the modification of the aptamer. - Aptamer: Cy5-labeled DO_PAM and + Aptamer: Cy5-labeled DO^A_PAM. Data represented the mean ± s.d.. Statistical significance was calculated by the unpaired two-tailed Student's t-test, with *** representing p < 0.001.

Protocol

3. Preparation (click me to open the protocol!)

4. Verification of the DNA origami structure by AFM (click me to open the protocol!)

4.1 Handbook: AFM cypher ES (click me to open the handbook!)

5. Fluorescent modification of sgRNA_L (click me to open the protocol!)

6. Loading of FITC-sgRNA_L/Cas9 complex (click me to open the protocol!)

7. Dimerization (click me to open the protocol!)

8. Loading of hemin (click me to open the protocol!)

9. ABTS assay of G4/hemin DNAzymes (click me to open the protocol!)

10. NPN uptake assay (click me to open the protocol!)

11. Targeting validation by LSM (click me to open the protocol!)

11.2 Handbook: Zeiss LSM 880 (click me to open the handbook!)

Notebook

Notebook for Assembly (click me to open the Notebook!)

Reference

1. Kolbeck PJ, Dass M, Martynenko IV, van Dijk-Moes RJA, Brouwer KJH, van Blaaderen A, et al. DNA Origami Fiducial for Accurate 3D Atomic Force Microscopy Imaging. Nano Lett. 2023 Feb 6;23(4):1236–43.
2. Bald I, Keller A. Molecular Processes Studied at a Single-Molecule Level Using DNA Origami Nanostructures and Atomic Force Microscopy. Molecules. 2014 Sep 3;19(9):13803–23.
3. Nasri M, Mir P, Dannenmann B, Amend D, Skroblyn T, Xu Y, et al. Fluorescent labeling of CRISPR/Cas9 RNP for gene knockout in HSPCs and iPSCs reveals an essential role for GADD45b in stress response. Blood Adv. 2019 Jan 8;3(1):63–71.
4. Kogut M, Kleist C, Czub J. Why do G-quadruplexes dimerize through the 5'-ends? Driving forces for G4 DNA dimerization examined in atomic detail. PLoS Comput Biol. 2019 Sep 20;15(9):e1007383.
5. Fabrication of Defined Polydopamine Nanostructures by DNA Origami‐Templated Polymerization - Tokura - 2018 - Angewandte Chemie International Edition - Wiley Online Library [Internet]. [cited 2025 Sep 29]. Available from: https://onlinelibrary.wiley.com/doi/10.1002/anie.201711560
6. Antioxidant activity applying an improved ABTS radical cation decolorization assay. Free Radical Biology and Medicine. 1999 May 1;26(9–10):1231–7.
7. Helander IM, Mattila‐Sandholm T. Fluorometric assessment of Gram‐negative bacterial permeabilization. J Appl Microbiol. 2000 Feb 1;88(2):213–9.
8. Hanžek A. Development of the aptamers for detection of ovarian cancer biomarkers in urine. 2023 [cited 2025 Sep 29]; Available from: https://rgdoi.net/10.13140/RG.2.2.36760.98566
9. Mela I, Vallejo‐Ramirez PP, Makarchuk S, Christie G, Bailey D, Henderson RM, et al. DNA Nanostructures for Targeted Antimicrobial Delivery. Angew Chem Int Ed Engl. 2020 Jul 27;59(31):12698–702.