Experiment

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Fabrication and Yield Improvement of GUV-in-GUV Structures

Purpose of this experiment

In constructing GIFTs, the key structural element is the GUV-in-GUV architecture. We hypothesized that a critical factor for forming this structure is that the inner liposomes must be significantly smaller in size than the outer ones. Based on this hypothesis, we first examined whether this structure could be successfully fabricated. Furthermore, we optimized the preparation conditions to improve the formation yield of GUV-in-GUV structures. Specifically, we investigated experimental parameters such as lipid composition, buffer composition, and centrifugation conditions to determine the optimal settings.

Materials and Methods

The following protocol represents the optimized procedure that maximizes the yield of GUV-in-GUV structures.

Dissolving lipids in mineral oil

DOPC, POPG, and cholesterol were mixed at a molar ratio of 3:1:6 and prepared with 0.0178 mol% NBD-PE for green fluorescence labeling.
A total of 1.0 μmol of lipids in chloroform was evaporated under a nitrogen stream to form a lipid film on a glass tube.
The lipid film was placed under vacuum in a desiccator overnight.
A 100 μL aliquot of mineral oil (Sigma-Aldrich, Cat. No. 330760-1L, heavy) was added to the lipid film on the glass tube.
The glass tube was sonicated at 50–60°C for 60 minutes to dissolve lipids in mineral oil.

Preparation of Inner Liposomes

A total of 6.7 μL of liposome internal buffer (0.5× TAE, 300 mM trehalose) was added to the sonicated glass tube.
The mixture was vortexed until it became cloudy, forming a water-in-oil (W/O) emulsion.
The emulsion was gently layered on top of 100 μL of liposome external buffer A (0.5× TAE, 5 mM KCl, 225 mM glucose, 75 mM trehalose).
The sample solution was centrifuged at 10,000 × g for 5 minutes, forming a pellet on the bottom of the tube.
The formed pellet was carefully collected with a pipette tip inserted from the top of the tube. To minimize contamination from mineral oil, residual oil on the pipette tip was carefully removed using laboratory tissue.
Liposomes were observed and analyzed using a confocal laser scanning microscope (Nikon AX, Nikon Solutions, Tokyo, Japan).

Preparation of GUV-in-GUV Structures

The recovered small vesicle solution was thoroughly mixed by pipetting, and 10 μL of it was added to the sonicated lipid film as the liposome internal buffer.
The mixture was vortexed until it became cloudy, forming a W/O emulsion.
The emulsion was gently layered on top of 100 μL of liposome external buffer B (0.5× TAE, 240 mM glucose, 60 mM trehalose).
The sample was centrifuged stepwise at 3,000 × g for 5 minutes, 6,000 × g for 5 minutes, and 8,000 × g for 5 minutes.
The liposome pellet was carefully collected with a pipette tip, as described in the previous section.
The formed GUV-in-GUV structures were observed and evaluated using a confocal laser scanning microscope.

Results

The green fluorescence corresponding to the inner vesicle membrane was observed inside the red fluorescent outer membrane, suggesting that the use of a small-vesicle-containing internal buffer successfully led to the formation of GUV-in-GUV structures (Fig.1).

Fig.1 Confocal fluorescence images of inner liposomes and a representative image of the GUV-in-GUV structure.

(a) Confocal image of inner liposomes labeled with NBD (green fluorescence). Scale bar = 100 µm. (b) Confocal microscope image of a double liposome structure. From left to right: a merged image of NBD and rhodamine (Rhodamine) fluorescence, channel images of the outer membrane (rhodamine-labeled, red), and inner membrane (NBD-labeled, green). The outer and inner membranes are observed with different fluorescent colors, clearly demonstrating that the inner liposome is significantly smaller than the outer liposome. Scale bar = 10 µm.

Through the optimization of preparation conditions, we ultimately achieved the GUV-in-GUV formation yield of 24.2% (Fig.2 a, b).

Fig.2 (a) Confocal image of GUV-in-GUV obtained under optimized conditions. Scale bar = 50 µm. (b) Yield of GUV-in-GUV structures relative to outer liposomes under optimized conditions (N = 5 independent experiments, n = 6 images per experiment). (c) Line profile analysis of a GUV-in-GUV. Scale bar = 10 µm. The white line indicates the position of fluorescence intensity measurement.

Among the obtained structures, GUVs containing multiple inner vesicles accounted for 32.4% (Fig. 3b).

Fig.3 (a) Confocal fluorescence images of inner liposomes and a representative image of the GUVs-in-GUV structure. Scale bar = 10 µm. (b) Yield of GUVs-in-GUV structures relative to GUV-in-GUV structures under optimized conditions (N = 5 independent experiments, n = 6 images per experiment).

Sectional Conclution

We confirmed that GUV-in-GUV structures can be formed by using GUVs as the internal solution for outer GUV fabrication, and that the optimization of preparation conditions improved the formation yield to 24.2%.

Introducing stepwise and sequential centrifugation enhanced the formation efficiency of GUV-in-GUV structures. The highest formation rate was achieved when both inner and outer vesicles had the lipid composition DOPC : POPG : Cholesterol = 3:1:6, which includes negatively charged POPG. Furthermore, using an outer solution containing 25% trehalose produced the best results. The KCl concentration was kept low (5 mM) to prevent excessive electrostatic screening between membranes.

We also succeeded in obtaining GUV-in-GUV structures containing multiple inner vesicles, with a formation yield of 32.4%. This structure is the target architecture envisioned for GIFTs construction.

DNA Origami

Purpose of this experiment

The key factors for properly folding DNA origami are the annealing conditions and MgCl₂ concentration. In this section, we used electrophoresis to confirm whether the DNA origami nanopores were correctly formed. After optimizing the annealing conditions, each sample was purified using a molecular weight cut-off filter to remove excess staple strands.

Materials and Methods

We used agarose gel electrophoresis to optimize the MgCl₂ concentration and annealing conditions for DNA origami folding.

1. Scaffold strands (M13mp18), staple strands, 1×TAE buffer, and MgCl₂ were mixed in a 0.2 mL tube. The MgCl₂ concentration was tested at 0, 2.5, 5.0, 7.5, 10.0, 12.5, 15.0, 17.5, and 20.0 mM.

2. Two annealing protocols were tested:

- The protocol is based on previous studies from Iwabuchi et al.

- A shorter, overnight protocol that allows folding in a single night. This protocol was slightly modified based on a previous study(Suzuki, Y et al).

### Annealing Protocols

Ⅰ. Protocol from Previous Study

- Heat at 90 °C for 20 s.

- Cool from 70 °C to 45 °C at a rate of 0.8 °C/h.

- Cool from 45 °C to 24 °C at a rate of 2.0 °C/h.

- Maintain and store at 24 °C.

Ⅱ. Short Protocol

- Heat at 65 °C for 15 min.

- Cool from 60 °C to 42 °C at a rate of 1.0 °C/h.

- Maintain and store at 15 °C.

3. Preparation of Samples

Each sample was prepared in 1× TAE buffer containing 10 nM scaffold and 50 nM staple strands.For the MgCl₂ concentration, samples were prepared under nine different conditions: 0, 2.5, 5.0, 7.5, 10.0, 12.5, 15.0, 17.5, and 20.0 mM. For each condition, the annealing samples shown in Table. 1 were prepared, and 1 μL of each sample was applied to the gel.

4. Agarose Gel Electrophoresis

Electrophoresis was performed using 1% agarose gels with 0.5× TBE buffer containing 5 mM MgCl₂. The electrophoresis was performed at 70 V for 60 minutes.

5. Staining and Imaging

The gels were stained with SYBR Gold and imaged using a gel imager ( Luminograph ).

Table.1 Annealing Sample

Table.2 Agarose Gel Electrophoresis (a) 1% Agarose gel recipe (b) Loading Sample

Samples prepared with the optimized MgCl₂ concentration and annealing conditions were purified using a column, and the purification was confirmed by agarose gel electrophoresis.

1. A total of 40 μL of the DNA origami solution and 460 μL of the centrifugation buffer (1x TAE, 5 mM MgCl2) were added to the filter and centrifuged at 4,500 × g for 5 min. After centrifugation, 400 μL of centrifugation buffer was added to the filter and centrifuged again at 4,500 × g for 5 min. This centrifugation cycle was repeated three times. Finally, the filter unit was inverted and centrifuged at 1,000 × g for 2 min to collect the purified DNA origami nanostructures. The Mg²⁺ concentration was then restored by adding a 250 mM MgCl₂ solution.

2. Agarose gel electrophoresis was carried out using 1% agarose gels with 0.5× TBE buffer containing 5 mM MgCl₂, at 70 V for 60 minutes.

3. The gels were stained with SYBR Gold and imaged using a gel imager ( Luminograph ).

Results

Optimization of MgCl₂ Concentration and Annealing Conditions

Regarding the annealing temperature and time, the protocol on the left represents the one-night method, while the protocol on the right follows the previous study.

The samples with MgCl₂ concentrations between 15.0 and 20.0 mM showed that the bands of misfolded or aggregated DNA disappeared, suggesting that the structures were correctly assembled (Fig. 4). In particular, the band position shifted to the lowest at 15 mM MgCl₂. Therefore, we decided to perform annealing under the condition of 15.0 mM MgCl₂.

Fig.4 Electrophoresis results of agarose gel for examining annealing conditions

After the purification, the bands corresponding to the staple strands were removed (Fig. 5).

Fig.5 Purification results of the DNA origami nanopore

Verification of the Docking–Undocking System

Purpose of this experiment

To confirm that the Docking–Undocking system functions as designed, we first performed polyacrylamide gel electrophoresis (PAGE) using only the DNA strands used in the strand displacement reactions for each step.

Materials and Methods

We tested whether the Docking–Undocking system could proceed using only the SNARE part and the DNA Signal, without using DNA origami.

1. The DNA samples were prepared as shown in Table. 3

2. Polyacrylamide gel electrophoresis (PAGE) was performed using 10% acrylamide gels with 1× TBE buffer containing 12.5 mM MgCl₂, at 140 V for 60 minutes at 25°C.

3. The gels were stained with SYBR Gold and imaged using a gel imager ( Luminograph ).

Table.3 Strand Displacement Reaction sample

Table. 4 Recipe for 10% polyacrylamide gel

Table.5 Loading samples for the strand displacement reaction

Next, we tested the nanopore selectivity of the Docking–Undocking system using DNA origami nanopores. Origami selectivity refers to the ability to determine which iSNARE participates in the reaction. This is achieved by using two sets of Docking Blockers and Docking Signals with different sequences.

This demonstrates that multiple Docking–Undocking systems can operate without interfering with each other, simply by redesigning the Docking Signals and Blockers.

In this experiment, in addition to Docking Blockers 1 and 2 and Docking Signals 1 and 2, we designed another set: Docking Blockers a1 and a2 and Docking Signals a1 and a2.

Fig.6 the Docking–Undocking system with origami selectivity

Using only the SNARE part, Docking Blockers, and Docking Signals, we carried out Step 1 of the reaction.

Two experiments were performed as follows:

1. We confirmed that Docking Blockers a1 and a2 could bind to iSNARE 1 and 2, in the same way as Docking Blockers 1 and 2.

2. We showed that iSNAREs with Docking Blockers 1 and 2 reacted only with Docking Signals 1 and 2, and Docking Blockers a1 and a2 reacted only with Docking Signals a1 and a2, resulting in the release of iSNAREs.

Experiment 1: To show that Docking Blockers a1 and a2 bind in the same way as Docking Blockers 1 and 2

Docking Blockers 1 and 2 were added to iSNAREs 1 and 2 with Lids, and the reaction was allowed to proceed for more than 15 minutes.
The electrophoresis samples were prepared as shown in Table. 5. The samples were run on a 10% polyacrylamide gel at 140 V for 60 minutes, and the bands were analyzed.
The gels were stained with SYBR Gold and imaged using a gel imager ( Luminograph ).

Experiment 2 : To verify origami selectivity

iSNAREs were protected with Docking Blockers (1 and 2 or a1 and a2), and then the corresponding Docking Signals were added.

The complexes of Lid, iSNAREs 1 and 2, and Docking Blockers (1 and 2 or a1 and a2) were incubated with the Docking Signals for 15 minutes at room temperature.

The electrophoresis samples were prepared as shown in Table. 5.

The samples were run on a 10% polyacrylamide gel at 140 V for 60 minutes, and the bands were analyzed.

The gels were stained with SYBR Gold and imaged using a gel imager ( Luminograph ).

Four experimental conditions were set up as follows:

Table.6 Experimental conditions for origami selectivity

Results

Experiment 1: To show that Docking Blockers a1 and a2 bind in the same way as Docking Blockers 1 and 2

In Fig. 7, the hybridization between iSNARE 1, iSNARE 2, and lid were evaluated.

The bands of the complex of i SNARE 1 + Lid + i SNARE 2 showed the highest band position. This result suggests the complex was formed as designed.

Fig.7 iSNARE 1, Lid, iSNARE2 complex

Fig. 8, Docking Blockers 1 and 2 bound to the complex of iSNARE 1 and 2 with the Lid, so a band appeared at a higher position.

They successfully formed a bound complex.

Fig.8 Confirmation of Docking Blocker binding

In Fig.9 , the lane marked with a check mark shows the sample in which Docking Signals 1 and 2 were added to the complex of iSNAREs 1 and 2, Lid, and Docking Blockers 1 and 2.

In this lane, two bands are observed: one corresponding to the iSNARE 1 and 2 + Lid complex, and the other corresponding to the Docking Blocker + Docking Signal complex.

From the electrophoresis results, it is suggested that Docking Signal 1 binds to Docking Blocker 1, while Docking Signal 2 binds to Docking Blocker 2.

As a result, it is also suggested that the Docking Blockers are released from the iSNAREs, exposing the regions of the iSNAREs that can bind to oSNAREs.

Fig.9 Removal of Docking Blockers by the addition of Docking Signals

From Fig.10, adding oSNARE 1 and 2 caused them to bind to iSNARE 1 and 2, resulting in the removal of the Lid. In the checked lane, the top band corresponds to the Lid, the second band from the top corresponds to the iSNARE + oSNARE complex, and the bottom band represents the unreacted oSNAREs.

Fig.10 Binding of iSNARE and oSNARE

In Fig. 11a, the iSNARE 1+ oSNARE 1 complex was separated by the addition of Undocking Signal 1, resulting in the formation of a complex between oSNARE 1 and Undocking Signal 1.

In the checked lane, the top band corresponds to the oSNARE 1 + Undocking Signal 1 complex, while the bottom band corresponds to iSNARE 1.

In Fig. 11b, the bound iSNARE 2 and oSNARE 2 were separated by Undocking Signal 2, resulting in the formation of a complex between oSNARE 2 and Undocking Signal 2. In the checked lane, the top band corresponds to the oSNARE 2 + Undocking Signal 2 complex, and the bottom band corresponds to iSNARE 2.

Fig.11 Unbinding of iSNARE and oSNARE. (a) Unbinding of iSNARE1 and oSNARE1 (b) Unbinding of iSNARE2 and oSNARE2

In Fig. 12a, the complex of oSNARE1 and Undocking Signal1 was dissociated by Undocking Signal Canceller1, resulting in the formation of a complex between Undocking Signal1 and Undocking Signal Canceller1. In the checked lane, the top band corresponds to the Undocking Signal1 + Undocking Signal Canceller1 complex, while the lower band corresponds to oSNARE1.

In Fig.12b, the complex of oSNARE2 and Undocking Signal2 was dissociated by Undocking Signal Canceller2, resulting in the formation of a complex between Undocking Signal2 and Undocking Signal Canceller2. In the checked lane, the upper band corresponds to the Undocking Signal2 + Undocking Signal Canceller2 complex, while the lower band corresponds to oSNARE2.

Fig.12 Removal of the Undocking Signals (a) Removal of the Undocking Signal 1 (b) Removal of the Undocking Signal 2

The electrophoresis in Fig.13 evaluated whether the dissociated iSNARE could be reassembled into the complex by adding Lid and Docking Blocker 1 and 2 again.

In the checked lane, the band pattern is identical to that of the iSNARE1 and 2 + Lid + Docking Blocker1 and 2 complex observed in Fig.8 (third lane from the right).

This indicates that the iSNARE + Lid + Docking Blocker complex was successfully reformed.

Fig.13 Reintroduction of the Lid and Docking Blockers

Experiment 2 : To verify origami selectivity

Regarding the parallelization of the Docking–Undocking system, Fig. 14 suggests that the complex consisting of iSNARE1 and 2, Lid, and Docking Blocker a1 and a2 was successfully formed.

Fig. 14 Confirmation of the functionality of Docking Blockers a1 and a2

Fig. 15 shows the electrophoresis results for the four tested conditions.
From the right, the fifth lane and those to its right correspond to conditions (A), (B), (C), and (D), respectively.
As expected, reactions occurred only under conditions (A) — where iSNARE1 and 2, Lid, and Docking Blocker1 and 2 were mixed with Docking Signal1 and 2 — and (C) — where iSNARE1 and 2, Lid, and Docking Blocker a1 and a2 were mixed with Docking Signal a1 and a2.

In these two cases, unlike in (B) and (D), the band corresponding to the iSNARE1 and 2 + Lid + Docking Blocker (1,2 or a1,a2) complex disappeared, while a new band corresponding to the iSNARE1 and 2–Lid complex appeared.

Furthermore, in both (A) and (C), an additional band was observed around the 20-bp position, indicating the formation of the Docking Blocker + Docking Signal complex.

Taken together, these results demonstrate that the Docking Signal forms a specific hybridization only in the presence of its corresponding Docking Blocker, thereby switching the iSNARE into a docking-active state.

Fig.15 Results showing that Docking Blockers are selectively removed by Docking Signals a1 and a2

Table. 7 Origami selectivity results

Regarding the selectivity of the Docking–Undocking system for DNA origami nanopores, the system was found to operate only under conditions (A) and (C). This result confirmed that the reaction proceeds only when a Docking Signal with a complementary sequence to the corresponding Docking Blocker is present.

From the electrophoresis results, it was confirmed that this strand displacement reaction functions as intended.

Insertion of DNA Origami Nanopores into Liposomal Membranes

Purpose of this experiment

DNA origami nanopores have recently attracted attention as a technology for introducing artificial permeation pathways into lipid bilayers. In particular, membrane anchoring via cholesterol-modified DNA is an effective method for stably immobilizing DNA structures on the surface of liposomes [2].To evaluate the binding of DNA origami nanopores to liposomal membranes, we introduced DNA origami nanopores into liposomal membranes.

Materials and Methods

To incorporate DNA origami nanopores into liposomes, we prepared a mixture containing liposomes, cholesterol-modified ssDNA, and DNA origami nanopores as follows:

Incubate the cholesterol-DNA at 55°C for 15 minutes.
Cholesterol-modified ssDNA was added to the liposome solution.
The mixture was incubated at room temperature for 10 minutes.
Fluorescently labeled DNA origami nanopores containing complementary strands to the ssDNA were added.
The mixture was incubated again at room temperature for 10 minutes.
Fluorescence localization was observed using a confocal laser scanning microscope.

Table 8. Composition of the sample

Results

The addition of DNA origami nanopores to liposomes with cholesterol-modified ssDNA led to the FAM fluorescence localizaion on the liposomal membrane surface modified with Rhodamine (Fig.16).

As shown in Fig. 16, it was observed that when cholesterol-modified ssDNA was introduced into liposomes (red), the addition of fluorescently labeled DNA origami nanopores containing complementary strands resulted in a clear fluorescence signal (green) localized on the liposomal membrane surface. In contrast, no FAM fluorescence was detected on the membrane without cholesterol modification. These results suggest that DNA origami nanopores were successfully attached on the membrane and the anchoring events depend on cholesterol-mediated interactions.

Fig 16. Confocal laser microscopy images showing the insertion of DNA origami nanopores into liposomal membranes.
(a) Confocal fluorescence images showing the localization of DNA origami nanopores (FAM-labeled, green) to the lipid membrane of liposomes (rhodamine-labeled, red). From left to right: merged image, FAM channel, rhodamine channel. The overlap of green and red signals on the membrane indicates the successful localization of DNA nanopores to the lipid bilayer. bar = 10 µm. (b) Control experiment using single-stranded DNA (ssDNA) labeled with FAM. The FAM signal does not colocalize with the liposome membrane, indicating that ssDNA does not bind to the membrane. From left to right: merged image, FAM channel, rhodamine channel. Scale bar = 10 µm. (c) Fluorescence intensity profiles obtained along the lines shown in the confocal images. In both the upper and lower panels, the FAM (green) and rhodamine (red) signals exhibit peaks at the same positions, quantitatively demonstrating colocalization of the DNA origami nanopores with the lipid membrane. Black arrows indicate the positions of the peaks.

Sectional Conclusion

The DNA origami nanopores were stably anchored onto liposomal membranes through hybridization with cholesterol-modified ssDNA.

Reference

[1]Y. Suzuki, I. Kawamata, K. Mizuno, and S. Murata, “Large Deformation of a DNA-Origami Nanoarm Induced by the Cumulative Actuation of Tension-Adjustable Modules,” Angew. Chem. Int. Ed., vol. 59, no. 15, pp. 6230-6234, Apr. 2020, doi:10.1002/anie.201916233.

[2]M. Langecker, V. Arnaut, T. G. Martin, et al., "Synthetic lipid membrane channels formed by designed DNA nanostructures," Science, vol. 338, no. 6109, pp. 932–936, Nov. 2012, doi: 10.1126/science.1225624.

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