Visualization of the Plasmid DNA Delivery System by Complementary Fluorescence Labeling of Arginine-Rich Peptides

Abstract

Cell-penetrating peptides, such as arginine-rich peptides, encapsulate nucleic acid drugs and deliver them to intracellular compartments. Comprehensive tracking of drug delivery systems (DDSs) provides information about the behavior of the drug as well as the fate of the drug carrier after drug release, which is crucial for minimizing side effects. In this study, we labeled peptides designed to carry plasmid DNA with two types of dyes, traditional dye fluorescein and aggregation-induced emission (AIE) dye tetraphenylethylene, and subsequently tracked the DDS through the complementary ON and OFF fluorescence behaviors of the dyes. Traditional fluorescent dyes are susceptible to aggregation-caused quenching during bioimaging, a problem that is mitigated by using AIE dyes. However, by using both of these contrasting fluorescent labels, we were able to clearly visualize the DDS at different stages of its deployment, from drug transport and delivery to carrier dissociation and migration, demonstrating the feasibility of accurate DDS visualization by complementary fluorescence labeling.

Introduction

Tetraphenylethylene (TPE) is a representative aggregation-induced emission (AIE) dye exhibiting unique fluorescence properties in the aggregate state, and thus it has been actively investigated for medical applications.^1–10) While performing photodynamic therapy (PDT) at high drug concentrations, AIE luminogens (AIEgens) promote the production reactive oxygen species (ROS) owing to the reduced energy gap between the singlet and triplet states (ΔE_S-T),^1,2) which increases therapeutic efficacy, thereby driving extensive research in this area.

AIEgens have also garnered attention for labeling drug delivery carriers.^11–14) These dyes become emissive when the carriers encapsulate drugs and reach high concentrations, making them useful for visualizing drug delivery systems (DDSs). Although studies on the behavior of drugs are numerous, studies on the comprehensive tracking of DDSs through different stages of their deployment—including the fate of carriers after drug release—remain limited.^15–18) To minimize side effects, it is essential to monitor both the drug and the carrier, from cellular uptake through to post-drug release.

We previously found that arginine (Arg)-rich peptides,¹⁹⁾ which are cell-penetrating peptides (CPPs), can interact with plasmid DNA (pDNA) through electrostatic interactions, deliver their cargo to the intracellular compartment, and achieve efficient transfection.^20,21) In this study, we aimed to clearly visualize Arg-rich peptides designed to carry pDNA by labeling Gly-(L-Arg)₉-NH₂ (R9) with both traditional fluorescent dye fluorescein (FLCN) and prominent AIEgen TPE and comparing their contrasting emission behaviors.

A few gene and nucleic acid DDSs have been labeled with AIEgens.^22–25) In this study, we hypothesized that aggregation-induced emission of TPE would allow visualization of peptide/pDNA complexes, while aggregation-caused quenching (ACQ) of FLCN would allow the detection of dissociated peptides, not peptide/pDNA complexes (Fig. 1A). By using both dyes, we tracked the DDS through different stages of its deployment, from cellular uptake and intracellular distribution to peptide dissociation, through the distinct emission behaviors of the two dye-labeled peptides (Fig. 1B).

Fig. 1. (A) Complementary Emission of Peptide Carriers (R9) Labeled with Prominent AIE Dye Tetraphenylethylene (TPE) and Traditional Fluorescent Dye Fluorescein (FLCN); (B) Schematic Illustration of Plasmid DNA (pDNA) Delivery System with Dye-Labeled Peptide Carrier

Results and Discussion

We used the solid-phase approach to synthesize peptide TPE-R9 labeled with carboxylated tetraphenylethylene²⁶⁾ (Fig. 2). We utilized the previously synthesized FLCN-R9²⁷⁾ in the subsequent experiments.

Fig. 2. Structures of Dye-Labeled Arginine-Rich Peptides

TPE-R9 was mixed with pDNA (6477 bp) at various N/P ratios to prepare peptide/pDNA complexes. The N/P ratio was defined as the molar ratio of guanidino groups in the peptide to phosphate groups in pDNA. The fluorescence intensity increased as N/P increased from 0 to 2 and saturated at N/P of 1.4 (Fig. 3). Therefore, the peptide binds to pDNA at N/P of 0 to 1.4, and excess peptide remains unbound and free in solution at N/P of 1.4 or higher (Fig. 1A). Additionally, the fluorescence intensity of the TPE-R9/pDNA complex (33.3 µg pDNA/mL) was similar at N/P of 2, 4, and 8 (Supplementary Fig. S2A). In contrast, free TPE-R9 at the same concentrations (22.2, 44.4, 88.8 µM) as TPE-R9/pDNA complexes (N/P = 2, 4, 8) was non-emissive (Supplementary Fig. S2B). Therefore, TPE-R9 as a peptide/pDNA complex is visible, while TPE-R9 as the free peptide is undetectable.

Fig. 3. (A) Fluorescence Spectra and (B) Intensity of the Fluorescence Peak at 475 nm of TPE-R9/pDNA Complexes at N/P Ratios of 0 to 2

In our previous study²⁰⁾ on FLCN-R9, the free peptide exhibits fluorescence. In particular, at N/P of 1.25, FLCN-R9/pDNA complexes show background-level emission, while at N/P of 1.5 or higher, the fluorescence intensity increases as the peptide concentration increases. This indicates that FLCN-R9 exists as the free peptide at N/P of 1.5 or higher rather than at N/P of 0 to 1.25. These results are consistent with the results obtained for TPE-R9, as shown in Fig. 3. Therefore, TPE-R9 can be visualized when the peptide binds pDNA, while FLCN-R9 can be visualized when the peptide is free, not bound to pDNA.

The size of TPE-R9/pDNA complexes decreased as the N/P ratio increased from 2 through 4 to 8, reaching the dimension of a nano DDS at N/P of 8, with an appropriate size for intracellular penetration (423 ± 36 nm) and moderate polydispersity index (PDI) (0.317 ± 0.041) (Table 1). The zeta potential also increased as N/P increased. Therefore, TPE-R9/pDNA complexes at N/P of 8 were chosen for cellular uptake experiments.

Table 1. Size, Polydispersity Index (PDI), and Zeta Potential of TPE-R9/pDNA Complexes Prepared at N/P Ratios of 2, 4, and 8

N/P	Size (nm)	PDI	Zeta-potentials
2	3412 ± 250	0.342 ± 0.068	+16.4 ± 0.3
4	1148 ± 105	0.510 ± 0.071	+18.4 ± 0.3
8	423 ± 36	0.317 ± 0.041	+21.9 ± 1.8

Peptide/pDNA complexes (TPE-R9/pDNA or FLCN-R9/pDNA) at N/P of 8 were incubated with Huh-7 human hepatocellular carcinoma cells for 2, 6, 24, and 48 h and then observed using confocal laser scanning microscopy (CLSM) (Fig. 4A, Supplementary Figs. S3, S4). Peptides were visualized by detecting TPE or FLCN (green), while pDNA was visualized by detecting labeled cyanine5 (Cy5, magenta), and nuclei were visualized by staining with Hoechst33342 (blue).

Fig. 4. (A) CLSM of Intracellular Distribution of Dye-Labeled Peptides TPE-R9 and FLCN-R9 (Green) Merged with Cy5-pDNA (Magenta) and Cell Nuclei (Blue); (B) Evaluation of Peptide and pDNA Colocalization in Cells; (C) Fluorescence Intensity of Peptides in Cells

As shown in Fig. 4A, TPE-R9 appeared as punctate green spots in cells after 2 to 24 h of incubation, and these spots diminished in intensity after 48 h of incubation, with the fluorescence signals dispersed throughout the cytoplasm. These results suggested that peptide/pDNA complexes were internalized in cells via endocytosis and dissociated in endosomes/lysosomes or migrated to the cytoplasm. In contrast, FLCN-R9 appeared as both punctate and widely distributed green signals in the cytoplasm after 2 h of incubation, which were more extensively distributed in the cytoplasm after 24 h of incubation and localized in nuclei after 48 h of incubation. These results indicate that free FLCN-R9 either directly translocate across the membrane or enter the cell via endocytosis along with the peptide/pDNA complex. After 6 h of incubation, peptides that dissociated from peptide/pDNA complexes merged with free peptides, which then migrated to the nucleus.

We tracked the peptides and pDNA in cells over time (Fig. 4B) and calculated the colocalization ratio, which was defined as the number of white pixels (overlap of green signals from peptides and magenta signals from pDNA) divided by the number of green pixels in a single cell. The results showed that the colocalization ratio was significantly higher for TPE-R9 than for FLCN-R9 at all time points. Because Cy5 is a traditional fluorescent dye that is susceptible to ACQ, its fluorescence decreases under high-concentration conditions, such as in the presence of peptide/pDNA complexes. Consequently, the colocalization ratio of TPE-R9/pDNA complexes might be underestimated. The significant decrease in the colocalization ratio of TPE-R9 after 48 h probably originates in the dissociation of TPE-R9/pDNA complexes, thereby releasing pDNA and increasing the relative fluorescence intensity of pDNA.

The fluorescence of TPE-R9 in cells was unchanged over time, whereas that of FLCN-R9 in cells increased after 6 h (Fig. 4C). The unchanging fluorescence intensity of TPE-R9 suggests that the cellular uptake of peptide/pDNA complexes and the decomposition of the complexes occur at similar rates. In contrast, the increasing fluorescence intensity of FLCN-R9 indicates that, in addition to the uptake of free peptides, the decomposition of peptide/pDNA complexes occurs after 6 h, resulting in the emergence of new free peptides, which increases the concentration of free peptides in cells.

The cell viabilities of Huh-7 cells treated with TPE-R9 and FLCN-R9 were evaluated using the Cell Counting Kit-8 (Supplementary Fig. S5). TPE-R9 demonstrated mild cytotoxicity at higher concentrations compared to FLCN-R9, suggesting that the incorporation of highly hydrophobic TPE into Arg-rich peptides may increase their cytotoxic effects.

Generally, FLCN in low-pH organelles, such as endosomes/lysosomes, exhibit weak fluorescence. Despite this, the fluorescence signals of TPE-R9 and FLCN-R9 were clearly distinct. Labeling peptides serving as drug carriers with both dyes enables the tracking of not only drug-encapsulated complexes but also free peptides surrounding the complexes, which provides information about the fate of peptides after drug release.

Conclusion

Peptide/pDNA complexes were visualized using TPE-R9, and free peptides were visualized using FLCN-R9. The contrasting fluorescence behaviors allowed us to track the DDS through different stages of its deployment, from drug transport and distribution to the dissociation of peptide/pDNA complexes in cells. Peptide/pDNA complexes were internalized in cells via endocytosis and transported to endosomes/lysosomes. Over time, most of the complexes were decomposed in endosomes/lysosomes, with some complexes breaking down in the cytoplasm, thereby releasing pDNA, which then reached the nucleus. Moreover, peptides that dissociated from peptide/pDNA complexes migrated to the cytoplasm and nucleus.

Comprehensive tracking of the CPPs of DDSs is valuable for reducing side effects. Although the ACQ of traditional dyes is considered a problem, using both traditional dyes and AIEgens together enables accurate visualization and precise localization of the constituents of DDSs.

Experimental

Peptide Synthesis

The peptides were synthesized on a solid supported by 9-fluorenylmethyloxycarbonyl (Fmoc)-based solid-phase methods using Rink Amide resin and Fmoc-protected amino acid as a scale of 50 µmol, as described previously.²¹⁾ The Rink Amide resin was soaked in N,N-dimethylformamide (DMF). After washing the resin with DMF, Fmoc-amino acid (3 equivalents (equiv.)) in DMF, and a mixture of 2-(1H-benzotriazole-1-yl)-1,1,3,3-tetramethyl-uronium hexafluorophosphate (HBTU) (2.9 equiv.), 1-hydroxybenzotriazole (HOBt) (3 equiv.) and N,N-diisopropylethylamine (DIPEA) (6 equiv.) dissolved in DMF were added to the resin. The Fmoc protective groups were removed using 20% piperidine in DMF. This cycle was repeated ten times. Then 4-(1,2,2-triphenylvinyl)benzoic acid (3 equiv.) or 5(6)-carboxyfluorescein (3 equiv.) in DMF, and a mixture of HBTU (2.9 equiv.), HOBt (3 equiv.) and DIPEA (6 equiv.) dissolved in DMF were added to the peptide. The resulting peptide was then suspended in a cleavage cocktail (94% trifluoroacetic acid (TFA), 2.5% water, 2.5% 1,2-ethanedithiol, 1% triisopropylsilane) at room temperature for 2 h to facilitate cleavage from the resin. TFA was evaporated under a stream of N₂ reduced to a small volume, after which the solution was dripped into cold ether to precipitate the peptide.

The synthesized peptides were purified using reverse-phase (RP)-HPLC using a Discovery^® BIO Wide Pore C18 column (25 cm ×21.2 mm solvent A: 0.1% TFA/water, solvent B: 0.1% TFA/MeCN, flow rate: 10.0 mL/min). After purification, the peptide solutions were lyophilized, and peptide purity was assessed using UPLC (Waters, Milford, MA, U.S.A.) and a ACQUITY UPLC^® BEH C18 1.7 µm column (2.1 × 50 mm; solvent A: 0.1% TFA/water, solvent B: 0.1% TFA/MeCN, flow rate: 0.5 mL/min, gradient: 10–90% gradient of solvent B over 4 min) and characterized by ion trap/time-of-flight mass spectrometry with LC (LC-IT-TOF-MS).

Preparation of Peptide/pDNA Complex

Each peptide and pDNA were dissolved separately in 10 mM 2-[4-(2-hydroxyethyl)-1-piperazinyl]ethanesulfonic acid (HEPES) buffer (pH 7.3). Peptide solution was added to the pDNA solution to form peptide/pDNA complexes in HEPES buffer. Complex solutions were stored at room temperature for 15 min prior to use. The N/P ratio was defined as the molar ratio of the guanidino groups in peptides to the phosphate groups in pDNA.

Fluorescence Measurements

The fluorescence intensities of peptide solutions and peptide/pDNA complex solutions were measured using a spectrofluorometer (ND-3300, N. anoDrop, Wilmington, DE, U.S.A.).

Dynamic Light Scattering (DLS) Measurements

The sizes of the peptide/pDNA complexes were evaluated by DLS using Nano ZS (ZEN3600, Malvern Instruments, Ltd., U.K.). A He–Ne ion laser (633 nm) was used as the incident beam. Light scattering data were obtained at a detection angle of 173° and a temperature of 25 °C, and were subsequently analyzed using the cumulant method to determine the hydrodynamic diameters and polydispersity index (PDI) (µ/Γ²) of the complexes. Results were presented as the mean and standard deviation derived from three measurements.

Zeta-Potential Measurements

The zeta-potentials of the peptide/pDNA complexes were evaluated using the laser-Doppler electrophoresis method with Nano ZS with a He–Ne ion laser (633 nm). Zeta-potential measurements were performed at 25 °C, utilizing a scattering angle of 173° for the measurements. Results were presented as the mean and standard deviation derived from three measurements.

Confocal Laser Scanning Microscopy (CLSM) Observation

Huh-7 cells were seeded onto 8-well glass-based plate (20000 cells/well) and incubated in 200 µL of Dulbecco’s modified Eagle’s medium (DMEM) containing 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin (PS). After exchanging the medium, peptide/Cy5-pDNA complexes prepared at an N/P ratio of 8 containing 0.5 µg pDNA were added to each well, and the cells were then incubated for each time. The medium was subsequently removed, and the cell were washed 3 times with phosphate-buffered saline (PBS) before being fixed with 10% formalin. The intracellular uptake of peptides and Cy5-pDNA was visualized using an LSM900 Airyscan (Carl Zeiss, Oberkochen, Germany) after staining the nuclei with Hoechst 33342. Observations were performed using a 63×objective lens at an excitation wavelength of 405 nm (UV laser) for Hoechst 33342, 488 nm (Ar laser) for peptides, and 633 nm (He–Ne laser) for Cy5-pDNA. The colocalization of peptides with Cy5-pDNA was quantified as follows:

  

where peptide pixels_{colocalization} represents the number of peptide pixels colocalizing with Cy5-pDNA in the cell, and peptide pixels_total represents the total number of pixels in the cell (n = 20).

Cell Viability

Huh-7 cells were seeded onto 96-well culture plates and incubated in 100 µL of DMEM supplemented with 10% FBS and 1% PS. The culture media were then replaced with fresh media containing the peptides at various concentrations. After 24 h of incubation, cell viability was assessed using the Cell Counting Kit-8, following the manufacture’s protocol.

Acknowledgments

We thank S. Ibuki for technical assistance. This study was supported in part by Japan Society for the Promotion of Science (JSPS) KAKENHI (No. 24K18254), Grant-in-Aid from the Naito Foundation, Japan Science and Technology Agency COI-NEXT (Grant Number: JPMJPF2114), Japan Science and Technology Agency ACT-X (Grant Number: JPMJAX222L), and Kyoto Prefectural Public University Corporation Program for Inter-University Collaboration and Joint Research Support.

Conflict of Interest

The authors declare no conflict of interest.

Supplementary Materials

This article contains supplementary materials.

References

• 1) Kang M., Zhang Z., Song N., Li M., Sun P., Chen X., Wang D., Tang B. Z., Aggregate (Hoboken), 1, 80–106 (2020).
• 2) Zhu C., Kwok R. T. K., Lam J. W. Y., Tang B. Z., ACS Appl. Bio Mater., 1, 1768–1786 (2018).
• 3) Wang H., Li Q., Alam P., et al., ACS Nano, 17, 14347–14405 (2023).
• 4) Song B.-L., Zhang X.-H., Qiao Z.-Y., Hao W., CCS Chem., 4, 437–455 (2022).
• 5) Chowdhury P., Banerjee A., Saha B., Bauri K., De P., ACS Biomater. Sci. Eng., 8, 4207–4229 (2022).
• 6) Hu J.-J., Jiang W., Yuan L., Duan C., Yuan Q., Long Z., Lou X., Xia F., Aggregate (Hoboken), 2, 48–65 (2021).
• 7) Cai X., Liu B., Angew. Chem. Int. Ed., 59, 9868–9886 (2020).
• 8) Li J., Wang J., Li H., Song N., Wang D., Tang B. Z., Chem. Soc. Rev., 49, 1144–1172 (2020).
• 9) He X., Xiong L.-H., Huang Y., Zhao Z., Wang Z., Lam J. W. Y., Kwok R. T. K., Tang B. Z., Trends Analyt. Chem., 122, 115743 (2020).
• 10) Yang S., Yu H., Xu X., Yang T., Wei Y., Zan R., Zhang X., Ma Q., Shum H. C., Song Y., ACS Nano, 17, 8195–8203 (2023).
• 11) Wang Y., Zhao J., Dong Z., Wang C., Meng H., Li Y., Jin H., Wang C., Mater. Today Chem., 21, 100537 (2021).
• 12) Pei Y., Wang Z., Wang C., Mol. Pharm., 18, 3951–3965 (2021).
• 13) Dong Y., Liu B., Yuan Y., J. Control. Release, 290, 129–137 (2018).
• 14) Yuan Y., Liu B., Chem. Sci., 8, 2537–2546 (2017).
• 15) Song X., Yang F., Ji R., Yang B., Lv Y., Wei Z., New J. Chem., 48, 5186–5198 (2024).
• 16) Zhong J., Quan Y., Zhao X., Li S., He Z., Ye G., Sun M., Miao Y., Ma C., Yang H., Chen X., Huang Y., Biomater. Adv., 134, 112711 (2022).
• 17) Dong Z., Bi Y., Cui H., Wang Y., Wang C., Li Y., Jin H., Wang C., ACS Appl. Mater. Interfaces, 11, 23840–23847 (2019).
• 18) Xue X., Zhao Y., Dai L., Zhang X., Hao X., Zhang C., Huo S., Liu J., Liu C., Kumar A., Chen W.-Q., Zou G., Liang X.-J., Adv. Mater., 26, 712–717 (2014).
• 19) Kato T., Kita Y., Iwanari K., Asano A., Oba M., Tanaka M., Doi M., Bioorg. Med. Chem., 38, 116111 (2021).
• 20) Oba M., Demizu Y., Yamashita H., Kurihara M., Tanaka M., Bioorg. Med. Chem., 23, 4911–4918 (2015).
• 21) Yokoo H., Dirisala A., Uchida S., Oba M., ACS Biomater. Sci. Eng., 10, 890–896 (2024).
• 22) Tang F., Liu J.-Y., Wu C.-Y., Liang Y.-X., Lu Z.-L., Ding A.-X., Xu M.-D., ACS Appl. Mater. Interfaces, 13, 23384–23395 (2021).
• 23) Cheng Y., Sun C., Liu R., Yang J., Dai J., Zhai T., Lou X., Xia F., Angew. Chem. Int. Ed., 58, 5049–5053 (2019).
• 24) Jin G., Feng G., Qin W., Tang B. Z., Liu B., Li K., Chem. Commun., 52, 2752–2755 (2016).
• 25) Yuan Y., Zhang C.-J., Liu B., Angew. Chem. Int. Ed., 54, 11419–11423 (2015).
• 26) Yamashita H., Demizu Y., Shoda T., Sato Y., Oba M., Tanaka M., Kurihara M., Bioorg. Med. Chem., 22, 2403–2408 (2014).
• 27) Oba M., Kunitake M., Kato T., Ueda A., Tanaka M., Bioconjug. Chem., 28, 1801–1806 (2017).