Carbon Quantum Dots Assisted Virus Tracking: From Skin to Brain

Abstract

Incurable infection by herpes simplex virus 1 (HSV‐1) can cause severe encephalitis and neurodegenerative diseases, e.g., Alzheimer's disease (AD) and amyotrophic lateral sclerosis. How HSV‐1 reaches the brain from the initial infection site remains inconclusive. Here, an innovative approach combining carbon quantum dots (CQDs) with dissolving microneedles (dMN) for real‐time tracking of HSV‐1 from skin to brain is presented. Upon application, CQDs‐HSV‐1 is released from the dMN through the swelling of interstitial fluid (ISF) in skin and subsequently monitored by living imaging. Remarkably, it is observed that HSV‐1 preferentially infects peripheral skin nerves, almost all viruses directly enter to brain via the spinal cord within 10–30 min, while few viruses enter the brain through the bloodstream via tail vein injection at the same time. Spinal cord injury (SCI) significantly delays the HSV‐1 transport from skin to brain but has no effect on the virus's travel from blood to brain. In a microfluid system, HSV‐1 shows preferential neurite infection, then transports to the cell body of differentiated SH‐SY5Y cells, highlighting the viral traffic process in neurons. The integration of CQDs‐virus labelling technology and dMN delivery model presents a promising tool for investigating the in vivo transport routes of neurotropic viruses with initial skin infections.

A novel carbon quantum dots‐dissolvable microneedle, CQDs‐dMN system, enables painless delivery and real‐time tracking of HSV‐1. The HSV‐1 is labeled with fluorescent CQDs and delivered to the skin dermis, minimizing damage compared to traditional methods. Real‐time imaging shows that the dMN swells with interstitial fluid, releasing HSV‐1, which rapidly infects peripheral nerves and spreads from the spinal cord to brain.

1. Introduction

Many viruses infect the skin and quickly cause severe brain damages and even death. However, the mechanisms by which they infiltrate the central nervous system (CNS) are not entirely conclusive.^{[ 1 ]} HSV‐1, a highly prevalent neuropathic pathogen, commonly infection of the lips, eyes, and genital regions, serves as an ideal model to trace the viral traffic from skin to brain. More importantly, HSV‐1 infection potentially causes incurable neurodegenerative diseases, as evidenced by the frequent presence of HSV‐1 in the brain tissues of Alzheimer's disease (AD) patients.^{[ 2} , ³ , ^{4 ]} HSV‐1 can rapidly reach the brain without manifesting clinical symptoms,^{[ 5 ]} either remaining latent or becoming active, which can induce severe acute necrotizing encephalitis and subsequent neurodegenerative disease.^{[ 6 ]}

Tracking the natural path of viruses in vivo, from the initial skin infection site to the brain, has been a prolonged challenge.^{[ 7} , ⁸ , ^{12 ]} Traditional methods, such as transmission electron microscopy (TEM) or antibody‐based imaging (ABI), only providing static images of infected cells or tissues, making them incapable of capturing the in vivo viral traffic process.^{[ 9} , ^{13 ]} Although HSV‐1 is known to travel from the skin to the brain through either peripheral neurons, the bloodstream, or both routes, the dominant path remains unclear.^{[ 10} , ^{11 ]} The lack of dynamic imaging techniques that can track viral movement in real‐time has significantly hindered our understanding of these mechanisms.

To address these gaps, we develop a novel approach here to track HSV‐1 in vivo by combining carbon quantum dots (CQDs) with skin‐dissolving microneedles (dMN) technologies. CQDs, carbon nanoparticle ranging from 1 to 20 nm with surface functional groups, offer superior photostability and biocompatibility, making them ideal for real‐time tracking of viral transport in living organisms.^{[ 12} , ^{13 ]} The use of dMN as a non‐invasive and painless transdermal delivery system allows for efficient viral directly into the skin, avoiding direct contact with blood vessels or nerves. This eliminates the need for specialized handling skills required for traditional injection methods, such as intravenous injection, intramuscular injection, or skin‐wound infection.^{[ 14} , ^{15 ]} This represents a significant advancement over conventional techniques.^{[ 16 ]}

We developed a CQDs‐dMN system to trace viral dynamics in a living animal. By enabling real‐time imaging of HSV‐1 trafficking, the integration of CQDs‐based viral labeling with the innovative dMN delivery model presents a promising and powerful tool for investigating the intricate routes of viral transport in vivo. This research not only enhances our understanding of how HSV‐1 pathogenesis spreads to and within the CNS but also opens new avenues for exploring therapeutic interventions targeting viral dissemination in the nervous system.

2. Result

2.1. Construction of CQDs‐HSV‐1 and CQDs‐HSV‐1 dMN

We employed CQDs as fluorescent tags to label HSV‐1 (Figure 1a). The CQDs demonstrated strong photostability, remaining stably even after prolonged irradiation for 3 h (Figure S1, Supporting Information). To facilitate the conjugation process, the carboxyl groups on the CQDs were chemically activated. This activation allows for a condensation reaction with the amine groups present on the surface proteins of HSV‐1. The resulting conjugate, referred to as CQDs‐HSV‐1, exhibits a distinct blue fluorescence signal (Figures S2 and S3, Supporting Information), which is crucial for visualization. The successful labeling of HSV‐1 with CQDs was confirmed through various assays. First, we conducted a particle diameter test to verify the size distribution of the CQDs‐HSV‐1 (Figure 1b; Figures S4,S5, Video S1, Supporting Information). Next, a fluorescence absorption test was performed to confirm the fluorescent properties of the labeled virus (Figure 1c; Figures S4d and S6, Supporting Information). Furthermore, cytopathic effect (CPE) and virus titration assays were carried out (Figure S8, Supporting Information), showing that the CQDs‐HSV‐1 maintained comparable infectivity to untagged HSV‐1. Importantly, these assays also demonstrated that the CQDs‐HSV‐1 maintained good virulence compared to the unmodified HSV‐1, indicating that the conjugation process did not adversely affect the viral biological properties. CQDs, with optimal absorption wavelength at 450 and 605 nm are also used to meet our experiments (dMN demonstrations and in vivo imaging) (Figure S7, Supporting Information). To fabricate the CQDs‐HSV‐1 dMN, we design a master mold using Auto CAD, creating a negative poly PDMS mold that preserves the conical shapes (Figure S9, Supporting Information). Using cold‐drying to maintain CQDs‐HSV‐1 activity,^{[ 17} , ^{18 ]} the dMNs we produced exhibit mechanical strength comparable to that of HA dMN with a 10 × 10 array configuration, consisting of 100 independent needles (Figure 1d–g; Figures S10 and S11, Supporting Information). Upon application to isolated mice skin, the dMN effectively releases CQDs‐HSV‐1 into the dermis, as confirmed by blue fluorescence and the preservation of conical pores, highlighting the excellent penetration and release capabilities of this system (Figure 1h).

Figure 1.

Construction of CQDs‐HSV‐1 and CQDs‐HSV‐1 dMN. a) Topography of CQDs‐HSV‐1 characterized by TEM (Scale bar, 50 nm). b) Histogram of the particle diameter distribution of CQDs‐HSV‐1. c) Photoluminescence (PL) emission of CQDs‐HSV‐1. d) A diagram of the preparation process of CQDs‐HSV‐1 dMN. e) Optical image of CQDs‐HSV‐1 dMN (Scale bar, 1000 µm). f,g) The exhibition of CQDs‐HSV‐1 dMN and the single needle by SEM images (Scale bar, 500 µm). h) The puncture and release of CQDs‐HSV‐1 dMN with the skin exhibition (H&E staining section and typical UV fluorescence images).

2.2. Selecting and Tracking of CQDs‐HSV‐1 dMN In Vitro

To select the most appropriate dMN to delivery CQDs‐HSV‐1, we make three distinct sizes of dMNs with base diameters 280, 450, and 550 µm and lengths of 500, 1000, and 1500 µm, respectively (Figure S12, Supporting Information). We characterize dMNs using a ChemiDoc imaging system (Video2). Each needle packages 8.3 × 10⁵, 1.6 × 10⁶ and 2.5 × 10⁶ CQDs‐HSV‐1 virions, respectively (Figure S13, Supporting Information). Both dMN 1000 and dMN 1500 show excellent puncturing ability with over 95% integrity in mice skin, while dMN 500 maintains only ≈77% integrity (Figure S12c, Supporting Information). To quantify the CQDs‐HSV‐1 virions released from dMNs, we dissolve the dMNs in culture medium (Figure S14a, Supporting Information). Virus titration assays reveal ≈90% virus (>1 × 10⁸ virions) released from dMN 1000 and dMN 1500, whereas only ≈10% (≈1 × 10⁶ virions) was released from dMN 500 (Figure 2a). Surprisingly, the dMNs maintain viral infectivity for at least 30 days at 4 °C, exhibiting nearly consistent viral amounts and only a slight decrease in the release rate. In contrast, the virus typically degrades within a week when stored at 4 °C in the Eppendorf tube (Figure 2b).^{[ 19 ]} Similar results are also obtained by viral DNA copy assays (Figure S14b–d, Supporting Information). The recovery of mice skin is recorded with the digital camera after penetrated by dMNs (Figure 2c) The mice skin recovers dramatically within 30 min without visible pores after dMN 1000 puncture (Figure 2d); while dMN 1500 causes wound with bleeding at the penetration site with ≈40% of remain pinholes after 3 h (Figures S15 and S17, Supporting Information). These results are further validated by H&E‐staining of skin sections (Figure 2e; Figure S16, Supporting Information), demonstrating dMN 1000 as an optimal tool for this study.

Figure 2.

Selecting and tracking of CQDs‐HSV‐1 dMN. a) The amounts of CQDs‐HSV‐1 released from the dMNs instant or kept for 30 days at 4 °C. b) The release rate of the dMNs. c) The schematic illustration of the mice skin recovery after dMNs penetration. Figure 2c was created with BioRender.com and is reproduced under an academic subscription license (2025). d) Skin performance after dMN 1000 penetration within 30 min and recovery rate. e) The images of H&E staining skin section of dMN 1000 at 30 min. f) CQDs‐HSV‐1 distribution released from dMN at different times. g) The relative fluorescence intensity of confocal images. All data are representative of three independent experiments. Statistical differences were analyzed using a two‐tailed Student's t‐test and a one‐way analysis of variance (ANOVA), giving p‐values of ns: p ≥ 0.05; ^*, p < 0.05; ^**, p < 0.01; ^***, p < 0.001.

We evaluate the tracking capability of the CQDs‐HSV‐1 released from the optimal dMN in vitro by agar overlay assay in cultured cells (Figure S18a, Supporting Information). Similar to untagged HSV‐1, CQDs‐HSV‐1 is rapidly released within 30 mins (1 × 10⁸ virions) and caused significant CPEs 24 h post infection (p.i.) (Figure S18b,c, Supporting Information). The released CQDs‐HSV‐1 initially localizes in cytoplasm (30 min), then accumulates in the perinuclear region (1 h) and enters the nucleus (till 6 h, Figure 2f). Fluorescence of CQDs‐HSV‐1 in the cytoplasm initially increases, followed by a decrease, while fluorescence in the nucleus keeps increasing from 1 to 6 h (Figure 2g). The continuously declining cell viability also demonstrates the good infectivity of CQDs‐HSV‐1 released from dMN (Figure S19, Supporting Information).

2.3. Distribution of CQDs‐HSV‐1 In Vivo

Nude mice are injected with CQDs‐HSV‐1 via dMN injection (MN), intravenous injection (IV), and subcutaneous injection (SC) (Figure S20, Supporting Information), followed by monitoring of the fluorescence distribution in vivo using IVIS animal imaging system (Figure 3a). Both CQDs‐HSV‐1 solution and CQDs‐HSV‐1 dMN show a strong fluorescence signal (Figure S21, Supporting Information). Notably, after being applied to the mice, only a minimal signal is detected in the CQDs‐HSV‐1 dMN, indicating high releasing efficiency of CQDs‐HSV‐1 from the dMN (Figure S21b, Supporting Information). Except brain, CQDs gradually spread throughout the body within 3 h (Figures S22 and S23, Supporting Information). We first detect the CQDs‐HSV‐1 signal at 5 min and observe that the signal was slightly attenuated at the injection site, but essentially no signal transportation was detected (Figure S23, Supporting Information). Surprisingly, CQDs‐HSV‐1 reaches the spinal cord within just 10 min and the brain at 15 min from dMN injection site of skin. In contrast, when injected via the tail vein, CQDs‐HSV‐1 initially reaches the heart at 10 min, with the brain being reached only after 30 mins (Figure 3b,c). Notably, CQDs‐HSV‐1 colonizes the spinal cord at ≈30 min post‐dMN injection and rapidly concentrates in the brain thereafter. Moreover, we find that the blood viral level in MN group is less than 25% of that in IV group (Figure 3d), demonstrating rare CQDs‐HSV‐1 into the bloodstream following MN injection.

Figure 3.

Live animal IVIS imaging tracking the routes of CQDs‐HSV‐1. a) Schematic illustration of live animal IVIS imaging in nude mice. Figure 3c was created with BioRender.com and is reproduced under an academic subscription license (2025). b) Living images of nude mice at different times. c) Relative fluorescence intensity of mice body at different times according to Figure 3b. d) The CQDs‐HSV‐1 level in the blood from different groups tested by ELISA. e) Living images of representative organs injected with IV and MN. f) Relative fluorescence intensity in different organs at 3 h. g) Viral load in the infected organs. Total genomic DNA is extracted, and the viral loads are assessed by qPCR. h) The temporal changes in viral signal intensity within the spinal cord and brain via IV injection. i) The temporal changes in viral signal intensity within the spinal cord and brain via MN injection. j) Percentage of hemolysis induced by CQDs and dMN matrix (HA). All data are representative of three independent experiments. Statistical differences were analyzed using a one‐way analysis of variance (ANOVA) and indicated as asterisks: ^*, ns: p ≥ 0.05; p < 0.05; ^**, p < 0.01; ^***, p < 0.001.

We sacrificed the mice to examine the distribution of CQDs‐HSV‐1 in various organs (Figure 3e; Figures S24 and S25, Supporting Information). A large amount of CQDs‐HSV‐1 accumulates in the brain of the MN group, but only a little in the IV group. CQDs‐HSV‐1 also localizes in the spine of the MN group but is absent in the IV group (Figure 3f). As expected, CQDs‐HSV‐1 also appears in liver and heart in the IV group but has little impact on the growth or organ weights of mice across all groups (Figure S26, Supporting Information). Moreover, we observe the highest viral load in the brain by MN injection, followed by SC and IV injection (Figure 3g). We evaluate temporal changes of viral signal intensity in the spinal cord and brain. A sustained increase in viral signals in both regions after MN injection within 1 h (Figure 3h), followed by a diminish of viral signals in the spinal cord and a greater viruses transport to the brain. A similar pattern occurred via SC injection, but with significantly lower signal intensity (Figure S25c, Supporting Information). Conversely, a markedly different trend is observed via IV injection, with challenges in detecting viral signals in the spinal cord and a slow increase in brain viral signals (Figure 3i). The favorable blood compatibility of CQDs and CQDs‐HSV‐1 facilitates successful viral signal detection (Figure 3j; Figure S27, Supporting Information). Additionally, we confirm that the dMN matrix also exhibits good blood compatibility, demonstrating extremely low hemolysis in comparison to the positive control. This highlights the safety and testability of our dissolvable dMN matrix (HA).

We apply histological analysis to further confirm the distribution of CQDs‐HSV‐1. No organ damage is observed following MN injection, while apparent tissue damage is evident the lungs after IV injection (Figure S28, Supporting Information) Some inflammatory cell infiltration and alveolar septal capillary congestion also appear in lung by IV and SC injection, comparing to the NC and CQDs injection (Figure S29, Supporting Information). We also check the viral fluorescence distribution at different time points in tissue sections. CQDs‐HSV‐1 also displays in the heart and liver both in the IV and SC groups, while not in the MN group (Figures S30 and S31, Supporting Information). Excitingly, CQDs‐HSV‐1 signals are only detectable in the brain and spine as early as 10 min post‐MN injection, with a markable increase at 30 min, while weakly appearing in the heart, liver, and lungs 3 h later (Figure S32, Supporting Information). Between 30 min and 3 h, ≈40–80% of CQDs‐HSV‐1 concentrates in the brain (Figure S33, Supporting Information). Notably, CQDs‐HSV‐1 signals diminish between 0.5 and 3 h post‐MN injection in the skin (Figure S34, Supporting Information). However, signals appear only in the heart at 10 min and weakly in the lung and liver at 30 min post‐IV injection. Except for a weak signal in the brain, liver, lung, and kidney, no signal is observed in the spine even 3 h post‐IV injection (Figure S33c, Supporting Information). Moreover, only 5% HSV‐1 reaches the brain after 30 min and merely 30% after 3 h. These findings highlight distinct transport routes for HSV‐1 via skin and systemic infections. HSV‐1 preferentially infects peripheral nerves and rapidly transports to brain through spinal cord.

2.4. CQDs‐HSV‐1 Transport Routes Verification in SCI Mice

To further confirm our findings, we observed the HSV‐1 transportation in the SCI mice model with a T9‐T10 spinal cord injury (Figure 4a). Obviously, as expected, the spinal cord injury significantly delays CQDs‐HSV‐1 transport to the brain via MN injection, extending the transport time from the original 15 min to 2 h (Figure 4b,d). In contrast, SCI does not affect CQDs‐HSV‐1 transport via the bloodstream following IV injection. Interestingly, CQDs‐HSV‐1 localizes to the spine, excluding the injured area via MN injection, whereas only minimal or no CQDs‐HSV‐1 signal is detected in the spine via SC or IV injection (Figure 4c,e; Figure S35, Supporting Information). Taken together, our data suggest that once HSV‐1 interferes with venous blood, it initially reaches the heart, then the lungs, liver, and kidneys, and only a small amount crosses the blood‐brain barrier (BBB), finally to reach the brain. In the absence of contact with either nerves or blood vessels, HSV‐1 preferentially infects peripheral nerves and extremely fast transport to the brain through the spinal cord (Figure 4f).

Figure 4.

SCI mice injected with the CQDs‐HSV‐1. a) The preparation process of the SCI mice model. Figure 4c was created with BioRender.com and is reproduced under an academic subscription license (2025). b) Living images of SCI mice at different times via IV and MN injection. c) Living images of SCI mice organs via IV and MN injections. d) Relative fluorescence intensity in SCI mice at different times. e) Relative fluorescence intensity in different organs. f) Schematic diagram of CQDs‐HSV‐1 routes via IV and MN injection. All data are representative of three independent experiments. Statistical differences were analyzed using a one‐way analysis of variance (ANOVA) and indicated as asterisks: ^*, p < 0.05; ^**, p < 0.01; ^***, p < 0.001.

2.5. The Mechanism of CQDs‐HSV‐1 Transport Routes

To further illustrate the mechanism of CQDs‐HSV‐1 transport routes, we induce the neural SH‐SY5Y cells differentiation with cell bodies and neurites, to mimic the relationship between the brain and spinal cord. Spherical SH‐SY5Y cells are differentiated into neurons through continuous serum starvation and the addition of neural factors (Figure 5a; Figure S36, Supporting Information). The differentiated SH‐SY5Y cells can be maintained up to 20 days in Diff3 medium (Figure S37, Supporting Information). Upon CQDs‐HSV‐1 infection, the differentiated cells exhibit CPEs and cell death at 9 h, with only 30% of the cells surviving at 24 h (Figure 5b; Figure S38, Supporting Information). In contrast, CQDs alone do not affect these cells. We observe that the CQDs‐HSV‐1 predominantly localizes in neurites within 10 min after infection, subsequently appearing in both neurites and cell bodies over time (Figure 5c,d). To further dissect the detailed process, we differentiate SH‐SY5Y cells in a microfluidic device (Xnoa) to separate the neurites and cell bodies (Figure 5e). Notably, CQDs‐HSV‐1 exhibits green fluorescence and rapidly infects the neurites in the microfluidic device. Localization occurs as early as 5 s after addition, with increased accumulation and transport along the neurites. By 14 s, numerous CQDs‐HSV‐1 are distributed throughout the microchannels and actively moving toward the cell body. By 20 s, some CQDs‐HSV‐1 reach the cell body, while others continue their transport, demonstrating the virus's ability to quickly access the brain from the spinal cord within just 20 s (Figure 5f; Video S3, Supporting Information).

Figure 5.

In vitro validation of CQDs‐HSV‐1 transportation in neurons. a) Confocal images of SH‐SY5Y cells and differentiation SH‐SY5Y cells. Anti‐MAP2 (red) labels microtubule‐associated protein 2, revealing the neuronal soma and neurites. b) CCK‐8 test of cell viability at different times. c) Confocal images of differentiated SH‐SY5Y cells infected with CQDs‐HSV‐1 at different times. d) The relatively high florescence intensity of the confocal images. e) Schematic diagram of the infection routes of CQDs‐HSV‐1 on differentiated SH‐SY5Y cells in the microfluidic device. Figure 5e was created with BioRender.com and is reproduced under an academic subscription license (2025). f) Video images of differentiated SH‐SY5Y cells infected with CQDs‐HSV‐1 at 0, 5, 14‐and 20‐s postinfection (spi). All data are representative of three independent experiments. Statistical differences were analyzed using a two‐tailed Student's t‐test and a one‐way analysis of variance (ANOVA), giving p‐values of ns: p ≥ 0.05; ^*, p < 0.05; ^**, p < 0.01; ^***, p < 0.001.

3. Discussion

The route of neurotropic virus (e.g., HSV‐1, Dengue virus) to brain from the infection site in skin has been a long‐term argument thus far.^{[ 20 ]} Recent studies have consistently shown that HSV‐1 could cause serious neurodegenerative disease, such as AD, emphasizing the urgent need to identify the paths through which HSV‐1 reaches brain for better intervention.^{[ 21 ]} One prevailing hypothesis suggests that virus primarily goes through the bloodstream and crosses BBB to enter brain.^{[ 22 ]} However, it cannot explain such effective CNS infections (64% prevalence in individuals under 50 worldwide), as such a little virus at the initial infection site (such as skin) hardly overcomes the tremendous challenges to reach brain, including the immune attack at the local tissues and in the bloodstream, BBB and other barriers.^{[ 23} , ^{24 ]} Here, we present a novel technology to track HSV‐1 infection from skin to brain.

We construct the CQDs‐HSV‐1 to track HSV‐1, utilizing CQDs for their excellent water solubility, biocompatibility, and low toxicity, making them ideal for biomedical applications.^{[ 25 ]} Their small size and high surface area facilitate effective labeling while minimizing potential side effects.^{[ 26 ]} Additionally, employing CQDs as a carrier may enhance the stability and delivery of the virus while reducing the exposure of host tissues to free viral particles. Our results indicate that CQDs‐HSV‐1 successfully infects cells, with only a slight reduction in viral titer compared to unmodified HSV‐1 (Figure S8b, Supporting Information). After 24 h postinfection, CPE was observed, suggesting that CQDs labeling minimally affects the viral virulence and infectivity (Figure S8a, Supporting Information), likely due to the small size of CQDs or lack of binding to key virulence‐related proteins.^{[ 7 ]}

DMN offers the advantage of painless and efficient delivery, enabling enhanced bioavailability while minimizing skin irritation.^{[ 27 ]} We fabricate dMN using hyaluronic acid (HA), a well‐known biocompatible material recognized for its solubility, biodegradability, and non‐toxic properties.^{[ 28 ]} HA naturally exists in the extracellular matrix with excellent hydrophilicity, allowing effective dissolution in ISF and releasing therapeutic agents.^{[ 29} , ^{30 ]} We pioneer the loading of CQDs‐HSV‐1 into dMNs with adjustable needle length to establish a skin infection model. Upon exposure to ISF, the dMN gradually dissolves,^{[ 31} , ^{32 ]} releasing CQDs‐HSV‐1 into the skin tissue. The CQDs‐HSV‐1 dMN leverages painless, minimally invasive, and safe properties. In our study, we observe minimal side effects associated with CQDs‐HSV‐1 in the dMN model, primarily linked to HSV‐1 capsid proteins that can trigger immune responses. To mitigate these effects, we optimized the dosage and timing of virus administration to minimize immune activation.

Our dMN 1000 precisely and effectively delivers the virus into the skin without contacting nerves or blood vessels. This dMN, characterized with large viral load capacity, effective puncture ability, and virus release, exactly pierces into the dermis layer with minimal skin damage. The hands‐on procedure is simple and safe. The selected microneedle penetration depth of 1000 µm is optimal for human application, effectively reaching the dermis layer where most peripheral nerves are located, without significantly damaging the epidermis or deeper tissues.^{[ 33 ]} Human skin consists of an outer epidermis (100–150 µm thick) and a dermis (1–4 mm deep).^{[ 34 ]} By targeting the dermis, our microneedle efficiently delivers the virus while ensuring patient safety and comfort, making it suitable for applications in humans. Excitingly, the dMN retains the viral infectivity for at least 30 days, likely because of storing the virus in a solidified form.^{[ 35 ]} The dry state and excipient may enhance viral stability, providing an alternative method for long‐term storage of viruses or other active biomaterials.

It is reported that a virus takes ≈5–30 min to enter host cells and several hours to initiate viral genome replication, and a much longer time to reproduce new virions.^{[ 36 ]} The excellent optical stability of CQDs facilitates successful viral tracking. We take advantage of a confocal microscope and living animal imaging to track HSV‐1 transport both in vitro and in vivo. Given the 3D nature of the mouse body, the organ fluorescence distribution observed through the living animal imaging system may not be entirely accurate. However, fluorescence microscopy of tissue sections compensates for this limitation. For example, while the living animal imaging system shows only a minimal CQDs‐HSV‐1 signal in the brain at 10 min, fluorescence microscopy reveals stronger signals of CQDs‐HSV‐1 in the brain section. However, these results demonstrate that HSV‐1 directly goes to the brain through intracellular neuron traffic from skin after entry, a novel and amusing findings which has not been revealed by any other approaches. Also, our findings also indicate how smart of HSV‐1 to survival in the brain by escaping innate and adaptive immunities. The combination of these two imaging techniques allows for accurate tracking of HSV‐1 from skin to brain.

Our data reveal that HSV‐1 preferentially infects peripheral nerves and rapidly reaches brain through spinal cord without causing immediate damage. This may explain why HSV‐1 can silently invade the brain, remaining latent and undetectable via neurotransmission.^{[ 10 ]} We surprisingly find that SCI significantly slows HSV‐1 transport, further corroborating that the dominant HSV‐1 transport route in vivo is through spinal cord to brain (Figure 4). Interestingly, we observe a partially shared transport route between SC injection (which induces a prominent blister bulge, Figure S20, Supporting Information) and both IV and MN injection (Figures S28 and S29, Supporting Information). We further validate our findings in human neuroblastoma SH‐SY5Y cells, a widely used cell line for studying neural diseases such as AD, neurotoxicity, ischemia, and amyotrophic lateral sclerosis,^{[ 37 ]} as well as for cell differentiation. Notably, we demonstrate that HSV‐1 infects neurites and quickly transports to the cell body within 20 s, further supporting our in vivo findings. We propose that HSV‐1 exploits intrinsic neuronal transport mechanisms, utilizing retrograde transport along microtubules to reach the neuronal cell body.^{[ 38 ]} The virus may hijack motor proteins such as kinesins and dyneins that facilitate cargo movement along the axon.^{[ 39 ]} By leveraging these cellular processes, HSV‐1 may efficiently access the cell body, ensuring its survival and potential reactivation in a latent state.

Our study provides a novel, visible virus infection model that combines dMN with CQDs labelling. The combination of CQDs and dMN represents a transformative advancement in the study of viral infections, enabling dynamic in vivo imaging and a deeper understanding of viral interactions with the CNS. We demonstrate that HSV‐1 preferentially infects peripheral nerves in skin, rapidly and effectively reaching the spinal cord to brain, while showing limited and slow transport from the tail vein to the brain. The developed dMN system is designed to track virus transmission from the skin to the brain, advancing our understanding of neurotropic viruses. Although our current focus is on HSV‐1, whose transmission mechanisms remain a long‐standing scientific question with no definitive conclusions.^{[ 6 ]} This principle can be easily adapted to investigate the transmission pathways of other neuroinvasive viruses, such as flaviviruses: Dengue virus (DENV), Zika virus (ZIKV), and Japanese encephalitis virus (JEV). Additionally, it can be utilized in various animal models, including hamsters, ferrets, camels, rabbits, pigs, and horses.^{[ 40 ]} This versatility holds great promise for developing therapeutic strategies to combat neuroinvasive infections. Furthermore, this technology may be further applied to deliver drugs to brain to bypass BBB or specialized drugs for local treatment such as easily degradable and oral uncapable polypeptides, using its solidified storage capacity.

Our technology could be further applied to clinical detection of viral transmission pathways in humans, enhancing our understanding of neuroinvasive diseases such as AD and enabling early intervention against virus‐induced neurodegeneration. However, anatomical and immunological differences between mouse and human skin and nerve architecture must be considered, as these could impact the translation of our microneedle‐virus system. Mouse skin is thinner and differs in epidermal and dermal composition to human skin,^{[ 41 ]} affecting microneedle penetration and distribution. Additionally, the thinner stratum corneum in mice (≈5 µm) compared to human skin (10–20 µm) may affect insertion ease.^{[ 42 ]} Notably, human skin has more complex nerve networks that could alter viral uptake dynamics,^{[ 43 ]} while mice have a higher density of sweat glands. This allows the microneedles to avoid the influence of multiple sweat glands of mouse skin and thus contact more nerves in the human skin. Immunologically, human skin contains a more diverse array of immune cells that may trigger a stronger local response to viral entry, potentially impacting the microneedle delivery efficacy.^{[ 44 ]}

4. Experimental Section

All chemicals and solvents were obtained from commercial resources without further purification. All the reactions were carried out under normal atmospheric conditions.

Materials

Sylgard 184 polydimethylsiloxane (PDMS) was purchased from Dow Corning Corporation Ltd. Hyaluronic acid (HA, MW = 30–50 kDa), Poly (methyl methacrylate (PMMA), N‐hydroxysuccinimide (NHS) and Carbodiimide hydrochloride (EDC) were obtained from Shanghai Macklin Biochemical Co., Ltd. Carbon Quantum Dots (CQDs) were acquired from Nanjing Mukenano Co., Ltd. Retinoic acid (RA) was sourced from Merck Co., Ltd. N6,2′‐O‐Dibutyryladenosine 3′,5′ ‐cyclic monophosphate sodium salt (Dibutyryl cAMP) and Potassium chloride (KCl) were procured from Shanghai Aladdin Biochemical Technology Co., Ltd. All chemicals were of analytical grade and used without further purification. Fetal bovine serum (FBS), Minimum Essential Medium (MEM) were purchased from the American Type Culture Collection (ATCC, USA). Neurobasal Medium and GlutamaxJ were obtained (100X) from Thermo Fisher Scientific. Penicillin‐Streptomycin Solution was sourced from Wuhan Pricella Biotechnology Co., Ltd. Recombinant Human/Murine/Rat Brain‐Derived Neurotrophic Factor (BDNF) was procured from PeproTech.

Cell Culture and Virus Production

Human Neuroblastoma SH‐SY5Y cells (ATCC) were maintained in MEM supplemented with 10% FBS, and 10 U mL⁻¹ penicillin–streptomycin (Gibco) in a humidified 5% CO2 incubator at 37 °C. The SH‐SY5Y cells were then infected with HSV‐1 strain SM44 (ATCC) at a MOI of 1, 10, 50 for different experiment needs. After infection, the cells were incubated for 24 h at 37 °C under 5% CO2 to allow the virus to replicate and propagate within the SH‐SY5Y cells. During this time, HSV‐1 would undergo its full replication cycle, resulting in the production of new viral progeny. The culture supernatant containing newly produced virus particles was collected. The collected supernatant was typically clarified by centrifugation (4000 x g for up to 30 min) to remove any remaining cellular debris. The clarified supernatant was then transferred to fresh tubes for storage at −80 °C until further use, as described previously.^{[ 45} , ^{46 ]} The differentiated SH‐SY5Y cells were cultured in diff3 medium (neurobasal medium + 0% FBS + 50 ng mL⁻¹ BDNF + 1 m db‐CAMP + 100U/mL of penicillin, and 100 µg mL⁻¹ of streptomycin + 10 µm RA). The same process was used for the differentiated SH‐SY5Y cells infected with HSV‐1, VR‐1493).

Synthesis and Purification of the CQDs‐HSV‐1

SH‐SY5Y cells were cultured up to 70% confluence in MEM medium supplemented with 10% FBS in a 10 cm culture dish. Cells were then changed to 2% FBS medium and infected with HSV‐1 at a multiplicity of infection (MOI) of = 1 for 12 h. CQDs, rich in carboxyl groups on the surface, were previously activated using EDC and NHS at room temperature and then added to the culture dish to incubate for 12 h. The cell supernatant was collected and centrifuged at 4500 rpm for 30 min to remove cell debris, followed by transferring to Centrifugal Filter Devices (Amicon Ultra‐15, MILLIPORE) and centrifuged again to filter out free CQDs. 10 mL MEM medium with 2% FBS was added to the filter tube to centrifuge again. This process was repeated 3 times until the free CQDs were completely removed, achieving a purification fold of at least 1500 times (15 × 10 × 10). Finally, the filtered solution was observed under an ultraviolet lamp at a wavelength of 480 nm, showing blue‐violet fluorescence. The purified CQDs‐HSV‐1 solution was transferred into Eppendorf tubes and stored at −80 °C for subsequent use.

Transmission Electron Microscopy (TEM, Japan Hitachi HT7700) was used to measure the structure and size of CQDs and CQDs‐HSV‐1. The size of CQDs‐HSV‐1 was also confirmed by Malvern NanoSight NS300 (488 nm Laser). A 1 mL syringe was used to draw the purified CQDs‐HSV‐1 into the injection chamber, and its particle size was measured at an excitation wavelength of 488 nm.

Fabrication and Puncture Ability Test of CQDs‐HSV‐1 dMN

The master mold was designed using AutoCAD software 2021 and printed with gray resin using a 3D printer (Phrozen Sonic Mini 8K). A PDMS mold (negative) was fabricated from the master mold by mixing two‐part resin systems containing vinyl groups (part A) and hydrosizloxane groups (part B). It was then dried until fully cured. The 5% pure HA solution and the HA solution containing CQDs‐HSV‐1 were poured into the PDMS mold, respectively, drying both in a vacuum oven (Hi‐Temp 3625A) at 4 °C. The PDMS mold was taken out to replenish the liquid every 3 h and repeated for three times, followed by pouring the 8% PMMA solution into the mold and drying to form a support layer for the entire dMN. The dMNs were obtained by carefully demolding, and their surface morphology was directly observed under a 3D digital camera (Mshot, China).

The isolated mice skin was applied to evaluate the piercing ability of the dMNs. The dMNs were manually penetrated to the mice's skin, and the penetration holes and depth were observed under a stereomicroscope (Olympus, Japan). The obtained images were analyzed using ImageJ software.^{[ 32 ]}

Release and Infectivity of CQDs‐HSV‐1 dMN

The Agar Overlay was first prepared. 2X MEM stock solution was first prepared by dissolving a 1‐liter packet of powdered MEM in 450 mL Milli‐Q‐dH₂O, followed by filter sterilization using a 0.2 µm bottle‐top filter and storing at 4 °C. Then, a 2X MEM working solution was prepared. To prepare 100 mL 2X MEM working solution, add 5 mL 20X NaHCO₃ to 85 mL 2X MEM stock and then add 10 mL FBS (Final FBS = 10%). Finally, for the agar overlay, we melt 1.8 g agar to 100 mL Milli‐Q‐dH₂O in a microwave oven, then transfer to a sterile bottle and cool to 45 °C. Mix equal volumes of pre‐warmed (to 37 °C) 2X MEM working solution with the molten 1.8% agar to obtain the Agar Overlay mixture.^{[ 46 ]}

Replace the medium (MEM with 10% FBS) of SH‐SY5Y cells with the agar overlayer mixture and let it solidify. Then the CQDs‐HSV‐1 dMN was pressed on top of the agar overlay, and only the water‐insoluble PMMA protective layer remained after the needles dissolved. The PMMA layer was then removed, and the agar overlay was washed with PBS three times. The cells were cultured in a 37 °C, 5% CO2 incubator for 24 h, and the CPE was observed under a microscope.

In Vivo Fluorescence Biodistribution

Twelve immunocompetent Nude mice, aged 6–8 weeks, were used to observe the viral distribution in the mice's bodies. Briefly, CQDs‐HSV‐1 dMN (packaged ≈1 × 10⁸ HSV‐1 virions), subcutaneous (SC) injection and intravenous (IV) injection with the same amounts of virions were inserted into the mice skin, subcutaneous thigh, and tail vein to release CQDs‐HSV‐1 into the mice body. The IVIS animal imaging system (IVIS Lumina Series III) was used to observe viral signals of mice at different time points (10, 15, 30 min, and 1, 2, 3 h after injection). The injection was repeated 4 times every 5 days. A negative group with no virus injection was set to be compared with the other three groups. After 20 days, mice were sacrificed, and the heart, liver, spleen, lung, kidney, and tumor were collected and weighed. Then the collected organs were used for bioimaging, and the fluorescence signal was recorded by the animal imaging system; the excitation and emission wavelengths were 470 and 630 nm, respectively. Images were analyzed by Image J software for fluorescence intensity quantification.

DNA Extraction and qPCR

Viral DNA was extracted from the cell lysates using the QIAamp DNA mini kit (Qiagen) according to the manufacturer's protocol.^{[ 47 ]} SYBR Premix Ex Taq (Takara) was used to analyze DNA levels on a Step OnePlus Real‐Time PCR system (Applied Biosystems). The program is shown in the following, 95 °C for 30 s followed by 40 cycles of 95 °C for 5 s and 60 °C for 30 s. Target DNA level was normalized to glyceraldehyde‐3‐phosphate dehydrogenase (GAPDH) in the same sample, and results were calculated using 2–ΔΔCt method.^{[ 48 ]} The primers used in this study show as follows,

HSV‐1: sense (5′‐3′) CAACTACCCCGATCATCAGTTA and antisense (5′‐3′) ACAGTTGCCTCCCATCCGAAACCAA.

GAPDH: sense (5′‐3′) GATTCCACCCATGGCAAATTCCA and antisense (5′‐3′) TGGTGATGGGATTTCCATTGATGA.

Hemolysis Assay

The blood compatibility of CQDs, CQDs‐HSV‐1, and dMN matrix was assessed by a hemolysis assay. Fresh blood was collected from healthy 4‐week‐old mice. A glass rod was used to stir the blood to remove fibrinogen. Next, ≈10 times the volume of 0.9% sodium chloride solution was added to the defibrinated blood and mixed well. The solution was then centrifuged at 1000 rpm for 15 min. After centrifugation, the supernatant was discarded, and the pellet of red blood cells was resuspended in 0.9% sodium chloride solution. The washing steps were repeated 3 times until the supernatant became colorless. Finally, a 4% red blood cell suspension was prepared using 0.9% sodium chloride solution for experimental use. For sample preparation, CQDs were diluted to concentrations of 1, 0.1, and 0.01 mg mL⁻¹. CQDs‐HSV‐1 was prepared at concentrations of 10^7, 10^6, and 10^5 virions/ mL. The dMN matrix (HA solution) was prepared at concentrations of 5%, 1%, and 0.2%. To perform the hemolysis assay, 500 µL of each prepared sample was mixed with 100 µL of the prepared red blood cell suspension. The mixtures were incubated at 37 °C with shaking at 300 rpm for 30 min. After incubation, the mixtures were centrifuged at 1500 rpm for 15 min. The supernatant was then analyzed for hemoglobin release by measuring the optical density (OD) at 540 nm. The state of the red blood cells in the incubated mixtures was documented using a 3D digital camera (Mshot, China). The percentage of hemolysis was calculated using the following formula: (Sample OD – Negative Control OD)/ (Positive Control OD – Negative Control OD) x 100.^{[ 49 ]}

Preparation of Tissue Sections and Histopathological Analysis

The collected organs (brain, heart, liver, lung, spleen, kidney, and spine) were fixed with 10% paraformaldehyde (PFA) at room temperature for 24 h, washed with PBS twice (10 min for each time), and then washed with Milli‐Q water twice (10 min for each time). The skins of the mice in the injection location were also collected and fixed. After that, tissues were dehydrated with ethanol at different concentrations, and tissues were washed with 50%, 70%, 80%, 95%, and 100% ethanol in the Excelsior TM AS Tissue Processor (Thermo). Subsequently, tissues were cleared with xylene and embedded in paraffin. The embedded tissues were cut into 5 µm‐ thick sections, stained with hematoxylin and eosin (H&E), mounted on a glass slip, and then observed under the optical microscope.^{[ 50 ]}

Blood Viral Analysis

Collect blood from mice (serum and plasma) and dilute the serum samples as needed according to the kit instructions. Use an ELISA microplate pre‐coated with the specific virus antigen or antibody. Block with blocking buffer and incubate for 1 h. Add 50–100 µL of diluted mouse serum to each well and incubate for 1–2 h at room temperature. Wash the plate 3–5 times with wash buffer. Add primary antibody, incubate for 1 h, followed by the secondary antibody (enzyme‐conjugated). Incubate for 30 min. Add substrate solution, incubate until color develops. Stop the reaction with the stop solution and measure absorbance at 450 nm. Calculate virus concentration based on a standard curve.

Virus Titration

SH‐SY5Y cells were seeded into 96‐well plates and incubated for 24 h. The cells were then infected with 100 µL per well of serially diluted virus supernatant, with each dilution tested in quintuplicate. The serial dilutions of the virus supernatant were prepared as tenfold dilutions in the appropriate cell culture medium. After 120 h of incubation post‐infection, the 50% tissue culture infectious dose (TCID₅₀) was calculated using the Kärber method, as described previously.^{[ 46} , ^{51 ]}

Differentiation of SH‐SY5Y Cells

The 100k SH‐SY5Y cells were cultured in MEM medium supplemented with 10% FBS. To differentiate the SH‐SY5Y cells, the ratio of FBS in the MEM medium was gradually decreased while simultaneously adding RA. A series of mediums with varying FBS ratios was prepared: diff1 (MEM medium + 2.5% FBS), diff2 (MEM medium + 1% FBS), and diff3 (neurobasal medium + 0% FBS). The medium was changed from diff1 to diff3 on different culture days, and finally cultured well in the diff3 for at least 20 days. The cells incubated in the microfluidics follow the same differentiation process. With the extension of the neurites, they passed through the microchannels, which size only allows neurites pass, and the cell bodies were left in the culture tanks.

Preparation of the Diff1: This medium consists of blank MEM, which provides essential amino acids, vitamins, and glucose necessary for cell survival. To this base, 2.5% hiFBS (heat at 56 °C for 30 min) was added to support cell proliferation by providing growth factors, hormones, and other proteins essential for cellular functions. Warming the mixture at a 37 °C incubator for 30 min. After that, 10 µm RA was added to the warm mixture. This formulation is typically used for experiments where moderate serum supplementation is needed to maintain cell health and encourage cellular differentiation or growth without overwhelming the culture with excess nutrients.

Preparation of the Diff2: This formulation uses MEM medium, similar to Diff1, but with a reduced concentration of 1% FBS. This lower FBS ratio can be used to examine cell behavior under more nutrient‐restricted conditions or to induce differentiation. The 1% FBS offers fewer growth factors, allowing for the study of cellular responses to lower serum levels, which might be more akin to physiological conditions where cells are not exposed to high levels of growth factors.

Preparation of the Diff3: Neurobasal medium, a specialized medium designed for the culture of neural cells, is used as the base for Diff3. This medium provides essential nutrients for their maintenance and differentiation. No FBS is added to this formulation, creating a serum‐free environment. Additionally, BDNF (50 ng/mL), 1X B27 (10 µg/mL), 1 M KCL, 1 M dB‐CAMP and 10 µM RA were added. This type of medium is typically used for studying neural cell differentiation, maturation, or the effects of specific growth factors added to the culture.

CCK‐8 Assay

SH‐SY5Y cells were seeded in 96‐well cell plates at a density of 1 × 10⁴ cells per well. The cells were then placed in a 5% CO₂ incubator for 24 h. Then the cells were treated with the peptides at the desired concentrations for 2 days. 90 µL of fresh cell culture medium and 10 µL of CCK‐8 (BioSharp, BS350B) were added to each well of the 96‐well plate, which were incubated at 37 °C for 1 h.^{[ 51 ]} The absorbance of each well was measured using a microplate reader (BioTek, Synergy H1). The absorbance was recorded at a wavelength of 450 nm.

SCI Mouse Model

The mice spinal cord of T9‐T10 was exposed by laminectomy, and a spinal cord impactor was used to apply a force of 90 kilodynes to cause contusion‐type SCI in the exposed spinal cord. This model was provided by Beijing GeneLine Biotechnology Co., Ltd.

Animal Ethics

All mice were obtained from the City University of Hong Kong and cultured under pathogen‐free conditions. All animal‐related experiments were carried out following the guidelines for the care and use of laboratory animals of the City University of Hong Kong. The study was approved by the Ethics Committee of the City University of Hong Kong with the license from the Department of Health, Hong Kong (18‐69 in DH/SHS/8/2/5 Pt.3).

Statistical Analysis

The experimental results were expressed as the mean ± standard deviation (SD) for each experimental group. All statistical analyses were carried out using the GraphPad Prism 9.0 software (GraphPad Inc., USA). For comparisons between two experimental groups, a two‐tailed Student's t‐test was applied. For comparisons more than two experimental groups, a one‐way ANOVA was applied. A p‐value less than 0.05 (p < 0.05) was considered statistically significant. Exact p values are reported in figures (ns: p ≥ 0.05; ^*: p < 0.05; ^**: p < 0.01; ^***: p < 0.001) All data are representative of three independent experiments.

Conflict of Interest

The authors declare no conflict of interest.

Author Contributions

X.Y. and M.L.H. designed and supervised the study. Y.X.F., X.W., and D.W. conducted experiments, Y.X.F. analyzed data, Y.X.F., C.S.H., C.E.C., Y.R.W., H.H., P.R.C., L.Y.Q., and Q.Y.W. checked the manuscript. Y.X.F., X.Y., and M.L.H. drafted the manuscript.

Supporting information

Supporting Information

Supplemental Video 1

Supplemental Video 2

Supplemental Video 3

Feng Y., Wang X., Chen C., et al. “Carbon Quantum Dots Assisted Virus Tracking: From Skin to Brain.” Adv. Mater. 38, no. 6 (2026): 2508464. 10.1002/adma.202508464

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.