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. 2026 Feb 7;11(7):12799–12809. doi: 10.1021/acsomega.5c13609

Internal Loading of Environmentally Friendly InP/ZnSe/ZnS QDs in Bovine Milk Exosomes

Minji Ko †, Hansol Lee †, Heejin Jung †, Keyong Nam Lee †, Sekyoo Jeong §, Young Kwang Lee ‡, Young Rag Do †,*
PMCID: PMC12947225  PMID: 41768635

Abstract

To enable stable and reliable fluorescence tracking in biological environments, this study presents a highly efficient, cadmium-free, and eco-friendly platform for exosome labeling using InP/ZnSe/ZnS quantum dots (QDs). QDs, known for high efficiency, were synthesized using a hot-injection method. To ensure water dispersibility and biocompatibility, the QD oleic acid (OA) surface ligands were sequentially exchanged with 11-mercaptoundecanoic acid (MUA) and polyethylene glycol (PEG) to internalize of QDs within exosomes. Structural unchangeable bovine milk-derived exosomes were purified for exosome internalization using electrophoretic oscillation-assisted tangent flow filtration (EPOTF) with 200 nm SiN x nanofilters. Ultrasonic treatment was optimized to internalize QDs into exosomes without causing structural deformation, which was confirmed by TEM imaging. The resulting QD-exosomes retained 89% of their initial photoluminescent intensity under continuous laser exposure, in contrast to a significant drop to 10% for conventional PKH67-labeled exosomes. Confocal microscopy demonstrated efficient uptake of QD-exosomes into HaCaT cells and distinct localization around the nucleus. This approach, which integrates eco-friendly QD synthesis, exosome-compatible surface modification, structurally unchangeable ultrasonic embedding, and dual-stage purification, provides a robust and scalable strategy for bioimaging and is suitable for real-time imaging, targeted drug delivery, and advanced nanomedicine applications.

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1. Introduction

With the recent advancements in bionanotechnology, the demand for high-performance fluorescent nanoparticles for biomolecule detection and imaging has significantly increased. − Quantum dots (QDs) have gained attention for their exceptional optical properties and stability, which make them highly suitable for various biomedical tracking applications such as bioimaging, sensing, drug delivery, phototherapy, and photodynamic therapy. − Compared to traditional organic fluorescent materials, QDs provide superior features, including high photoluminescence efficiency, a broad absorption spectrum, long-term stability, a narrow emission spectrum, and ease of control of emission peak wavelength, allowing precise and sustained imaging in biological environments. , Despite the limitations of highly efficient and well-developed Cd-based QDs, the future of QD technology is promising with the emergence of InP-based QDs as nontoxic, eco-friendly, and narrow emission alternatives. − In parallel, carbon dots (CDs) have attracted attention as eco-friendly fluorescent nanomaterials owing to their carbon-based composition and good biocompatibility. However, their excitation-dependent emission, limited spectral tunability, and batch-to-batch variability often restrict their use in quantitative and long-term bioimaging applications. , In particular, the InP/ZnSe/ZnS core/shell/shell QD structure offers enhanced stability and improved luminescence properties. The dual protective shell layers of ZnSe and ZnS effectively reduce surface defects in the QDs and improve their optical stability such that they are reliable for application in biological environments and suitable for advanced bioimaging of exosomes.

Exosomes, nanometer-sized extracellular vesicles (ranging from 30 to 150 nm), play crucial roles in intercellular communication and physiological regulation. − Owing to their unique characteristics, such as the ability to cross the blood–brain barrier (BBB) and evade detection by the immune system they are promising candidates as diagnostic tools and therapeutic delivery vehicles. − To maximize these properties for real-time tracking and targeted imaging, highly efficient, nontoxic, and biocompatible QD markers should be integrated into exosomes. Such attachment or internalization allows for a more profound understanding of exosome behavior in cancer diagnosis, drug delivery, and physiological regulation while also demonstrating the potential of eco-friendly QDs as a real-time bioimaging platform (Table ). −

1. Summary of Exosome-Based Applications Enabled by Eco-Friendly QDs for Bioimaging and Drug Delivery.

  exosome type quantum dot labeling method efficiency cytotoxicity metrics refs
external loading A549 cell cultures derived exosomes green QD nanospheres, and red QD nanospheres DNA-assisted QD fluorescent nanospheres (QNs) assembly - -
  EVs within the epithelial cell layer isolated from the human vaginal mucosa QDs functionalized with PEG chemical linker - -
  exosomes highly positive for HER2 obtained from serum samples of breast cancer patients CdSe/ZnS (QD655) binding using antibodies - -
  human pancreatic cancer cells derived exosomes folic acid-InP QDs MSA coated ligand exchanged QDs conjugated with antibodies using carbodiimide chemistry 25–30% compared to Rhodamine 6G low
  human breast cancer cells (SKBR3 cells) derived exosome silicon QDs formation of binding between CD63-conjugated Si QD and exosomal CD63 - -
  exosomes secreted by SKBR3 cells gold–carbon quantum dots tumor-specific antibodies attached to GCDs form exosome-specific nanoprobes capable of labeling exosomes via immune reactions - -
  human pancreatic cancer cell line, and nonmalignant human pancreatic cell line derived exosomes Stavidin-coated Qdot 605, Qdot 655 hybridized with antibody-conjugated QD probes for specific EV surface biomarkers - -
  exosomes derived from a monocytic cell line InP/ZnS conjugate the antibody and QD using carbodiimide bonding after targeting exosomes with a CD63 antibody InP/ZnS-MSA 17.82% -
  circulating exosomes quantum dot nanospheres signal amplification for assembling quantum dot nanospheres   -
  A549, H1975, H446, HUVEC carboxylated QD (antibody-linked QDs) QD probes were employed to label tumor markers (immunoaffinity-based labeling) - -
  HL-60 cells amino water-soluble quantum dots (525 and 625 nm) interaction between exosomal proteins and aptamers - -
  MCF-7 cells-derived exosomes CdTe quantum dots CdTe quantum dots act as signal reporters, interacting with Ag+ through cation exchange reactions - -
internal loading B16F10 cells (a murine melanoma cell line) derived small and large exosomes Ag2Se and Ag2Te NIR-II QDs electroporation - no observable
  MCF-7 cells derived exosomes V2C-PEG electroporation at 200 V, 100 μF - -
  Macrophages (J774A.1) and tumor cells (HeLa) derived exosomes ZnCdSe@ZnS QDs QDs were internalized by cells and then released to the QD-exosomes (incubation) - no observable cytotoxicity ≤100 nM
  HepG2, 3T3, and HeLa cells D-GQDs D-GQDs were incubated with exosomes at room temperature with protection from light (incubation) - -
  exosomes isolated from MCF-7 cancer cells FA-CQDs (MTX into FA-based CQDs) extrusion/sonication technique FA-CQDs: 0.54 Ex@MTX-CQDs: 0.21 -
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This study is focused on internalizing eco-friendly InP/ZnSe/ZnS QDs within bovine exosomes to enable dynamic tracking and real-time monitoring at specific target sites in biological systems, as shown in Figure . To this end, a high-efficiency QD synthesis method and a high-purity, large-scale exosome purification method must be developed in advance. Our research team reported the development of a high-efficiency, eco-friendly InP/ZnSe/ZnS QD synthesis method and a new exosome purification method called the electrophoretic oscillation tangent flow (EPOTF) method. − A biocompatible surface ligand, such as 11-mercaptoundecanoic acid (MUA) followed by polyethylene glycol (PEG), realized by substituting the oleic acid ligand decorating QDs, is critical to provide biocompatibility and allow internalization into exosomes. , Among various strategies for embedding QDs into exosomes, we introduce a viable approach using ultrasonic microbubble technology. This method is effective in temporarily punching the exosome bilayer to allow the incorporation of QDs within the vesicle. The study also develops a method to isolate QD-exosomes from unintegrated QDs while maintaining the superior optical properties of InP/ZnSe/ZnS QD-exosomes using an EPOTF process with both 200 and 50 nm nanopores. By optimizing the surface exchange process, developing an embedding methodology, and purifying QD-exosomes, we significantly enhance the tracking capabilities of QD-exosomes. Above all, in this study each method is improved and the three approaches are effectively integrated. This concerted approach is expected to substantially contribute to exosome diagnostics, treatment, and drug delivery and offers a viable approach for targeted imaging and monitoring with high precision and minimal toxicity of InP/ZnSe/ZnS QD embedded exosomes.

1.

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Schematic diagram of (a) exosome isolation process from bovine milk using EPOTF, (b) ligand exchange on InP/ZnSe/ZnS QDs from OA to MUA, and PEG and (c) the QD loading process into exosomes via ultrasonication.

2. Results and Discussion

Obtaining a significant amount of high-quality InP/ZnSe/ZnS QDs and bovine exosomes is essential for effectively studying a practical approach to load QDs into exosomes. High-efficiency InP/ZnSe/ZnS QDs are synthesized using the hot-injection method previously reported in our research papers. The photoluminescence quantum yield (PLQY) of the InP/ZnSe/ZnS QDs used in this study reached approximately 80%. The emission peak wavelength is located at 523 nm, and the full width at half-maximum (FWHM) is 37 nm, as shown in Table S1. To synthesize exosome-compatible QDs, as shown in Figure a, exosomes are purified from bovine milk by the EPOTF isolation method using a SiN x membrane filter with 200 nm pores. On the other hand, as shown in Figure b, the hydrophobic OA ligand bound to the QD surface is exchanged with hydrophilic MUA. More exosome-compatible PEG is then attached to the MUA terminal functional group. Eco-friendly QDs with this hydrophilic surface are exosome-friendly and can enter and remain stably within the exosome. After mixing surface-modified InP/ZnSe/ZnS-MUA-PEG QDs and purified bovine milk exosomes, ultrasound is applied to allow exosome-compatible QDs to enter the exosomes in Figure c. The ultrasound relaxes the bilayer shell of the exosome, thereby creating pores and allowing QDs to enter the exosomes. To purify the obtained QD-exosome, the EPOTF method is applied twice, utilizing SiN x nanofilters with pore sizes of 200 and 50 nm, respectively, to effectively remove aggregated biomaterials and unreacted QDs. It is therefore necessary to first secure high-performance QDs by surface-modifying InP/ZnSe/ZnS-MUA-PEG QDs.

As shown in the schemes of Figure S1, when OA, which is used as a hydrophobic ligand during the synthesis of InP/ZnSe/ZnS QDs, is replaced with hydrophilic MUA through the ligand exchange process, it moves from the chloroform solvent in the lower layer to the methanol solvent in the upper layer. This process is crucial for making the QDs compatible with exosomes. Since the PLQY and optical properties of the MUA-substituted QDs are measured according to the change in QD concentration, the MUA concentration is optimized through the exchange process, as shown in Figure S2a–c. Similarly, the reaction to attach PEG to MUA was optimized by evaluating the changes in PL and visible absorption characteristics according to the reaction time and MUA concentration, as shown in Figure S2d–f. Additionally, the changes in optical characteristics were evaluated according to the MUA concentration, as shown in Figure S2g–i. The optimization of these processes is a key part of our research methodology that ensures the high performance and stability of the QDs. The PL, photoabsorption, and PLQY characteristics of the optimal MUA and the optimal MUA-PEG-substituted QDs are compared, as shown in Figure a–c. Although the PLQY of pristine QDs decreases from 80.8% to 64.1% for QD-MUA and 54.8% for QD-MUA-PEG, the efficiency is more stably maintained compared to previous results of biocompatible QD synthesis reported to date (Table ). The FT-IR spectra indirectly confirmed MUA ligand substitution by showing peaks corresponding to –OH, –CH stretching, and CO bonds when OA was substituted. In addition, peaks corresponding to –OH, –CH stretching, and CO bonds indirectly verified that PEG formed bonds and peaks indicating C–O bonds demonstrated PEG was bound, as shown in Figure d–f. In addition, as shown in the TEM images in Figure g–i, the size, shape, and crystallinity of the QDs obtained through the ligand substitution of MUA and PEG are similar to those of pristine QDs, showing that they do not change significantly. Though there is some reduction in efficiency, the QD-MUA-PEG exhibits significantly enhanced UV stability compared to pristine QDs or surface-modified QDs (QD-MUA and QD-MUA-PEG), as shown in Figure . These characteristics demonstrate the advantage of InP QDs over traditional organic dyes for bioimaging applications due to their strong resistance to photobleaching.

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(a) PL spectra, (b) absorbance spectra, and (c) PLQY and peak wavelength of stepwise-optimized InP-MUA-PEG. FT-IR spectra of (d) InP/ZnSe/ZnS QDs, (e) ligand-exchanged InP-MUA QDs, (f) ligand-exchanged InP-MUA-PEG QDs. TEM images of (g) InP/ZnSe/ZnS QDs, (h) ligand exchanged InP-MUA, (i) ligand exchanged InP-MUA-PEG.

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(a) Relative degradation of PLQY of InP, ligand exchanged InP-MUA, InP-MUA-PEG QD solutions under UV (λ = 365 nm) exposure stability for 1 h and (b) long-term stability under UV (λ = 365 nm) exposure stability for 24 h.

The exosomes capable of loading QDs are obtained from bovine milk. SiN x nanofilters with 200 nm pores are fabricated using nanoimprinting and dual-side photolithography techniques reported in our previous publication, as seen in Figure S3a,b. To fabricate pores in 200 nm pore arrays precisely, the hole size of the nanofilters must be reduced after fabrication. To reduce the hole size, Al2O3 is conformally coated by the ALD method to yield pores of the desired size (Figure S3c). Large amounts of exosomes are purified using a 200 nm filter in a high-voltage-operated housing with separated electrodes, combining electrophoretic oscillation reported in our previous report with tangent flow filtration, shown in Figure S4. As previously reported, the purified exosomes were analyzed by SEM and TEM electron microscopy to confirm the morphology.

The SEM images in Figure a,b clearly show that the exosomes isolated by this purification method have a spherical shape with a diameter of approximately 100 nm. In addition, as shown in Figure c, the cryo-TEM image clearly shows the spherical exosome morphology and that the exosomes obtained through this purification method retain their original morphology without any deformation. The NTA characterization results in Figure d show a distinct peak in the exosome size distribution below 200 nm. This confirms that high-concentration exosomes of 1.24 × 1011 particles/mL with an average size of 106.8 nm are successfully isolated. Additionally, the zeta potential measurement of the exosomes, as shown in Figure e, indicates a surface charge of −12.56 mV. This suggests that the exosomes can effectively respond to electrophoretic oscillation and are well dispersed during filtration under the EPOTF process, as previously reported. As shown in Figure f, Western blot analysis also demonstrates effective exosome purification by the EPOTF method, as seen by the presence of exosome markers such as ALIX, HSP70, TSG-101, and CD63 in high-concentration exosomes. Various morphological, size, and surface analyses confirm that high-concentration exosomes can be obtained using the EPOTF isolation method without morphological changes.

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(a) High magnification SEM image, (b) SEM image, and (c) cryo-TEM image of isolated bovine milk exosomes. (d) NTA spectra of the average particle size, (e) zeta potential, and (f) the Western blot analysis of isolated bovine milk exosomes.

To internalize eco-friendly InP/ZnSe/ZnS QD-MUA-PEG into exosomes, a mixture of QD-MUA-PEG solution and exosome solution is sonicated for 5 min. In Figure , the SEM images (Figure a–d) and NTA results (Figure e–h) are analyzed based on the number of ultrasonic treatments applied. The optimal loading condition is determined as the point at which exosome shape alterations and concentration changes are minimal. Consequently, this study established three cycles of 5-min ultrasonic treatments as the ideal conditions for QD loading into exosomes. The produced QD-exosomes are purified using the EPOTF process, which separates distinct QD-exosomes from free QDs. This separation process allows us to distinguish between QD-exosomes and QDs that are not inside the exosomes, thereby providing a clear understanding of the QD-exosome internalization. As seen Figure S4, when filtered through a 50 nm nanofilter, the PL peak intensity of the QD-exosome solution is almost the same as that of the nonloaded, filtered QD solution. This suggest two possibilities: either no internalization occurred or only some QDs dispersed in the solvent were loaded.

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SEM images of exosomes loaded with QDs via ultrasonication, depending on the number of loading cycles; (a) unloaded exosomes, (b) 1 cycle, (c) 3 cycles, and (d) 5 cycles. NTA spectra of the average particle sizes of exosomes loaded with QDs via ultrasonication, depending on the number of loading cycles; (e) unloaded exosomes, (f) 1 cycle, (g) 3 cycles, and (h) 5 cycles.

To more effectively evaluate whether the QDs are internalized within the exosomes, FE-TEM analysis was performed as shown in Figure a. The TEM images show that the QDs are successfully loaded inside the exosomes, providing strong evidence for the internalization process. Additionally, we evaluate the PL emission and confocal images to determine the viability of using InP/ZnSe/ZnS QD-based nanoprobes for bioimaging. The peak wavelength, FWHM, and peak shape of the emission spectrum of the QD-exosomes are nearly identical to those of pristine QDs and surface-modified QDs (QD-MUA and QD-MUA-PEG) (Figure b). While the PLQY of QD-MUA-PEG decreases slightly to about 56.8%, it is sufficient to replace traditional dyes, as shown in Table S1. We compared the photostability of InP/ZnSe/ZnS QD-exosomes with that of exosomes labeled with PKH 67, a commonly used hydrophobic green fluorescent dye (PKH-exosomes). Both types of labeled exosomes are imaged under continuous laser exposure for 120 s, simulating prolonged or repeated imaging conditions in vitro or in vivo. The results show that the fluorescence intensity of the QD-exosomes retained 89% of their initial brightness after 120 s. In contrast, the intensity of the PKH-exosomes decreases to about 10% during the same time (Figure c). Furthermore, confocal images show that while the fluorescence of QD-exosomes is largely maintained (Figure d), PKH-exosomes demonstrate photobleaching, with nearly all fluorescence fading after 120 s (Figures e and S5). These findings indicate that the InP/ZnSe/ZnS-MUA-PEG-based QD-exosome conjugates developed in this study are suitable for in vitro or in vivo studies that require prolonged or repeated imaging.

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(a) TEM image of the QD-loaded exosome. (b) PL spectra of the stepwise-optimized InP-MUA-PEG. (c) Fluorescence intensity over exposure time for exosomes loaded with InP-MUA-PEG and PKH67. Time-dependent fluorescence images of exosomes loaded with (d) PKH67 and (e) InP-MUA-PEG, recorded before and after 120 s of exposure.

To verify the feasibility of the labeling process of QD-exosome conjugates, the intracellular uptake pathway of QD-exosome conjugates in HaCaT cells was traced through fluorescence imaging. First, the nucleus of HaCaT cells was labeled with a blue fluorescent dye (DAPI) before exosome uptake (Figure a). The bovine exosomes labeled with InP/ZnSe/ZnS-MUA-PEG-based QDs were uptaken into HaCaT cells. The green cell image (Figure b) and the blue-labeled nucleus image (Figure d) were merged to confirm that the bovine exosomes were internalized inside the cells. To further verify that the QD-exosome conjugates were properly uptaken and distributed inside the cells, the membrane of milk exosomes was fluorescently labeled with a red dye (Exo-S) using an ExoGlow-Membrane labeling kit. Subsequently, the intracellular distribution of QD-exosomes and Exo-S-labeled exosomes in HaCaT cells was compared. The red image of the Exo-S-labeled exosomes (Figure c) and the green image of the QD-labeled exosomes (Figure d) confirmed that the distributions of Exo-S and QD-labeled exosomes were almost identical. In particular, the merged image shows orange or white in the area where exosomes were abundant (Figure d). This indicates that QDs are suitable for exosome labeling, similar to the existing Exo-S dye. Finally, the merged image revealed that QD-exosome conjugates accumulated in the cytoplasmic region, particularly around the nucleus, and this phenomenon was consistently observed in all treated cells. Therefore, the feasibility of using InP/ZnSe/ZnS-MUA-PEG-based QDs as a labeling material for studying the biological function of QD by internalizing them into exosomes was proven. QD-exosome conjugates not only effectively label exosomes but also exhibit a stable distribution within cells, providing a reliable platform for future biological studies.

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Fluorescent microscopy images of InP QD-loaded exosomes (labeled with ExoGlow-MembraneTM labeling red dye) uptake by epidermal keratinocyte cells and (a) nuclei stained with DAPI blue dye in keratinocyte cells, (b) InP QD, (c) exosome protein with Exo-S, and (d) merged fluorescent microscopy images of the combined blue-labeled nucleus, red-labeled exosome, green-labeled InP QD uptake in the same keratinocyte cells.

3. Conclusion

In this study, we established an exosome labeling platform using cadmium-free and eco-friendly InP/ZnSe/ZnS quantum dots (QDs) functionalized with MUA and PEG. The optimized surface modification process yielded water-dispersible QDs while maintaining strong photoluminescence properties, with PLQY values of 64.1% for QD–MUA and 54.8% for QD–MUA–PEG, compared to 80.8% for pristine QDs. TEM and FT-IR analyses confirmed that the ligand exchange process did not induce noticeable changes in the QD morphology and verified that MUA and PEG ligands were successfully attached. Efficient association of QDs with exosomes was achieved using an ultrasonic-assisted loading process (three cycles of 5 min), which enabled QD loading while preserving the structural integrity and concentration of exosomes. SEM, cryo-TEM, and NTA analyses showed that the spherical morphology of the exosomes was well maintained after loading, with an average particle size of 106.8 nm and a concentration of 1.24 × 1011 particles mL–1. In addition, zeta potential measurements revealed a surface charge of −12.56 mV, indicating suitable electrophoretic responsiveness during EPOTF-based purification.

A photostability analysis demonstrated that QD-loaded exosomes retained approximately 89% of their initial fluorescence intensity after 120 s of continuous laser exposure, whereas PKH67-labeled exosomes showed a rapid decrease to about 10% under the same conditions. Furthermore, confocal fluorescence imaging confirmed successful cellular uptake of QD-exosomes in HaCaT cells, with clear colocalization of QD signals, exosome membrane markers, and cell nuclei. Overall, these results demonstrate that InP/ZnSe/ZnS–MUA–PEG-based QDs provide a stable, nontoxic, and optically reliable platform for exosome labeling and tracking. In particular, this approach offers a promising strategy for targeted imaging and monitoring with high precision and minimal toxicity and is expected to contribute meaningfully to the advancement of exosome-based diagnostics, therapeutics, and delivery systems.

4. Method

4.1. Fabrication of SiN x Membrane Nanofilters

To fabricate the SiN x nanofilters, 6-in. silicon wafers with double-sided polishing were used as a base substrate. A 200 nm thick SiO2 layer was first deposited by thermal oxidation, followed by the deposition of a 300 nm thick low-stress silicon nitride (LSN) layer using low-pressure chemical vapor deposition (LPCVD). A spin-on-glass (SOG) resin was patterned with nanoscale holes, each with a diameter of 330 nm, using nanoimprinting lithography. Any remaining SOG layer was etched by reactive ion etching (RIE; PlasmaPro 80, Oxford Instruments). Next, a window structure was formed using a positive photoresist, and the LSN layer was selectively etched using RIE. After stripping the photoresist, the wafer’s backside was coated with a 2 μm SiO2 protection layer. A 200 nm Cr layer was deposited and patterned through double-sided alignment photolithography. The Cr hard mask was etched using ETCR-400 (APCT), and the SiO2 and LSN layers on the backside were etched by inductively coupled plasma (ICP; PlasmaPro 100 Cobra). To form the SiN x nanofilter, the silicon layer was etched to a depth of 330 μm using deep reactive ion etching (DRIE), and the wafer was then diced into 21 square chips (2.5 × 2.5 cm2). Residual silicon and SiO2 layers were removed using tetramethylammonium hydroxide (TMAH) and buffered oxide etchant (BOE). To form 200 nm pores for exosome separation, the pore size of the SiN x chips was adjusted by depositing Al2O3 using atomic layer deposition (ALD; Lucida-D-100, NCD).

4.2. Isolation and Purification of Milk Exosomes

Fresh bovine milk was centrifuged at 5000g for 30 min and subsequently at 12,000g for 1 h. After each centrifugation, the supernatant was carefully collected by decantation, and the pellets, along with any visible fat, were removed. This procedure yielded defatted and skimmed milk, which was used for exosome isolation. The resulting liquids were passed through SiN x nanofilter chips (200 nm pore size) using an EPOTF system. The purification process was conducted at 4 °C using a square waveform with an 80% duty cycle and a frequency range at 1 Hz. A + 5 V potential was applied to drive negatively charged exosomes through the filter, while a −1 V signal was used to remove surface-bound particles. The purified exosomes were stored at – 20 °C for subsequent use.

4.3. Synthesis and Ligand Exchange of InP/ZnSe/ZnS QDs

4.3.1. Synthesis of InP Cores

Green-emitting InP cores were synthesized by combining 1.8 mmol of indium iodide (InI3) and 4.4 mmol of zinc chloride (ZnCl2) in 12 mL of oleylamine (OLA) within a 50 mL three-neck flask. The mixture was degassed under vacuum and maintained at 120 °C for 1 h. After purging the flask with nitrogen, 1.05 mL of tris­(dimethylamino)­phosphine ((DMA)3P) and 1.95 mL of trioctylphosphine (TOP) were swiftly added. The system was then heated to 107 °C and maintained for 30 min. Subsequently, the reaction temperature was elevated to 167 °C and held for 5 min before being quenched by rapid cooling to room temperature and immersion in a water bath.

To isolate cores of uniform size, 16 mL of n-hexane was added to the crude reaction mixture. The larger precipitates were removed by centrifugation at 12,000 rpm for 5 min. The remaining supernatant was treated with anhydrous ethanol and centrifuged again at 9000 rpm for 5 min to further purify the product. The final InP cores were redispersed in toluene for subsequent shell growth.

4.3.2. Shelling of InP Cores

For shelling, a solution containing 4.4 mmol of ZnCl2, 6 mL of OLA, and 6 mL of 1-octadecene (ODE) was loaded into a 100 mL three-neck flask and degassed. The mixture was then heated to 120 °C for 6 min. After nitrogen purging, the purified InP cores dispersed in toluene were introduced, followed by a brief degassing step to evaporate residual solvent.

To form ZnSe shell, a selenium precursor solution (3 mmol Se powder dissolved in 3 mL of TOP) and a zinc stearate solution (3.76 g of Zn­(st)2 in 15 mL ODE) were prepared. Initially, 0.5 mL of the Se solution and 4.4 mL of the Zn­(st)2 solution were injected, and the mixture was heated to 200 °C for 30 min. This step was repeated once more under identical conditions. The temperature was then increased to 290 °C and maintained for 30 min to finalize the ZnSe shell.

Subsequently, to form the ZnS intermediate layer, 1 mL of sulfur solution (6 mmol S in 3 mL TOP) and 4.4 mL of Zn­(st)2 solution were added, followed by heating at 300 °C for 1 h. The outermost ZnS shell was grown by injecting 2 mL of 1-dodecanethiol (DDT) at 225 °C and reacting for 1 h. Finally, 2 mL of a zinc acetate solution (3 mmol Zn­(ac)2 in 3 mL of oleic acid (OA)) was added on a 200 °C hot plate, and the reaction was allowed to proceed at 190 °C for 2 h. After completion, the mixture was cooled to room temperature to yield InP/ZnSe/ZnS quantum dots.

For purification, the QD solution was diluted with anhydrous hexane at a 1:2 ratio and allowed to settle for 30 min at ambient temperature. The QDs were then collected by centrifugation at 12,000 rpm for 5 min. This washing procedure was repeated until no impurities remained. In the final step, the QDs dispersed in chloroform were mixed with anhydrous ethanol in a 3:4 ratio and centrifuged at 9000 rpm for 5 min to complete the purification.

4.3.3. Ligand Exchange and PEGylation of InP/ZnSe/ZnS QDs

To achieve water dispersibility, the oleic acid ligands originally present on the surface of the synthesized quantum dots were replaced with 11-mercaptoundecanoic acid, followed by a PEGylation process. For ligand exchange, 0.1 g of 11-mercaptoundecanoic acid (MUA) was dissolved in a solution composed of 1.2 mL of methanol and 0.3 mL of distilled water. The pH of the solution was adjusted to 12 using a sodium hydroxide solution. Subsequently, 800 μL of chloroform-dispersed quantum dots was added, and the mixture was stirred for 3 h. The quantum dot concentrations were varied under four experimental conditions: 0.02, 0.04, 0.06, and 0.08 g/mL. After the reaction, the aqueous phase containing the ligand-exchanged QDs was collected, washed with ethanol, and redispersed in distilled water.

For the PEGylation step, 200 μL of the MUA-treated quantum dot solution was added dropwise into a one-to-one volumetric mixture of PEG 200 and distilled water, and the mixture was stirred at 40 °C. The PEGylation was performed under five different reaction durations of 1, 2, 3, 4, and 5 h. The volume of the PEG solution was also varied under four conditions: 50, 100, 150, and 200 μL.

4.4. QD Loading into Exosomes

A total of 20 μL of PEGylated quantum dots was added to 3 mL of exosome solution containing 4 × 1010 particles/mL. The mixture underwent sonication in cycles of 30 s at 40 kHz, followed by cooling on ice for 90 s. Experiments were conducted with one, three, and five sonication cycles to evaluate the effect of treatment frequency. After sonication, the samples were incubated at 37 °C for 30 min to allow QDs to be internalized into the exosomes.

4.5. Characterization

To observe the surface morphology of the exosomes, field emission scanning electron microscopy (FE-SEM; JSM-7610F, JEOL Ltd.) was used. Zeta potential measurements were obtained using an ELSZ-2000 (Otsuka). Nanoparticle tracking analysis (NTA; Nanosight NS300, Malvern) was employed to determine size and concentration. Absorbance spectra of QDs and exosomes were recorded using a UV–vis spectrophotometer (S-3100, SINCO). Photoluminescence (PL) spectra were collected using a xenon lamp and spectrometer (Darsa, PSI Trading). Quantum yield (PLQY) was calculated using Coumarin153 (QY = 0.58 in ethanol) as a reference dye. Transmission electron microscopy (TEM; JEM-2100F, JEOL) was used to confirm QD size and shape. FT-IR spectroscopy (Nicolet iS50, Thermo Fisher) was employed to verify chemical bonding changes after ligand exchange. Exosome labeling and cellular uptake were analyzed by fluorescence microscopy (Nikon Eclipse Ci-L Plus) and confocal microscopy (Stellaris 5, Leica).

4.6. Western Blot Analysis

Samples were mixed with SDS sample buffer containing 0.05% β-mercaptoethanol, followed by sonication for 5 min and heating at 95 °C for 10 min. Equal amounts of protein (100 μg) were loaded onto tris-glycine gels (Invitrogen) and transferred to nitrocellulose membranes. After blocking with a reagent (Cell Signaling Technology), membranes were incubated overnight at 4 °C with primary antibodies. After washing, HRP-conjugated secondary antibodies were applied for 1 h. Signals were developed using enhanced chemiluminescence (ECL; Thermo Fisher) and visualized using an Amersham ImageQuant 800 (Cytiva).

4.7. Fluorescent Labeling of Exosomes

For lipid labeling, PKH67 dye was diluted 1:500 in Diluent C (1 mL) and mixed with 10 μL of exosome solution in PBS. The mixture was incubated at room temperature for 30 min in the dark, and excess dye was removed via centrifugal filtration. For colabeling of exosome proteins, 2 μL of ExoSparkler was added to 100 μL of exosome solution (1 × 1010 particles/mL), incubated at 37 °C for 30 min, and cleaned by two rounds of PBS washing and filtration (3000g, 5 min each).

4.8. Cellular Uptake Assay

HaCaT cells (0.5 × 105 cells/well) were seeded onto a dual-chambered slide and incubated overnight. After washing with PBS, 25 μL of labeled exosomes were added to each well and incubated for 6 h at 37 °C in 5% CO2. Cells were washed three times with PBS, fixed with 4% paraformaldehyde for 10 min, and stained with DAPI (Sigma-Aldrich) to visualize nuclei.

Supplementary Material

ao5c13609_si_001.pdf (1.3MB, pdf)

Acknowledgments

This work was supported by a National Research Foundation of Korea (NRF) grant funded by the Korea government (MSIT) (RS-2021-NR059326) and the “Regional Innovation System & Education (RISE)” through the Seoul RISE Center, funded by the Ministry of Education (MOE) and the Seoul Metropolitan Government. (2025-RISE-01-005-04).

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsomega.5c13609.

  • The detailed scheme and photographs of ligand exchange on InP/ZnSe/ZnS QDs, along with the relative absorption and photoluminescence spectra of the synthesized InP QDs, are presented. Furthermore, fluorescence images of InP-MUA-PEG- and PKH67-loaded exosomes at different time intervals provided (PDF)

The authors declare no competing financial interest.

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ao5c13609_si_001.pdf (1.3MB, pdf)

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