Transcytotic transportation of size-controlled nanocarriers into dystrophic skeletal muscle leads to therapeutic outcome in mice

Domain: neuroscience
Published: 2025-12-11
Editorial outcome: major_revision

Abstract
Duchenne Muscular Dystrophy (DMD) is a lethal muscle-wasting disorder with limited therapeutic options. Although nanodrug delivery offers promise, the biodistribution and access routes to dystrophic muscle remain poorly understood. Here we show that intravenously administered mesoporous silica nanoparticle (MSNP) carriers exhibit striking size-dependent biodistribution in male DMD mice. Small nanocarriers (50~100 nm) efficiently accumulate in skeletal muscle while avoiding hepatic and splenic sequestration, outperforming larger particles (200~300 nm). Importantly, we uncover that endothelial transcytosis, not passive vascular leakage, is the dominant and previously unrecognized route by which nanocarriers access dystrophic muscles. Further, we encapsulate tamoxifen, a repurposed drug, into optimized MSNPs. This intervention increases utrophin expression, reduces fibrosis, and diminishes myofiber necrosis, resulting in improved muscle health and strength. Our results establish size-tuned, transcytosis-enabled nanocarriers as a transformative strategy for targeted drug delivery to dystrophic muscle, paving the way for nanomedicine-based therapies in DMD and potentially other muscle disorders.

Introduction
In the past few decades, different nanocarriers have been developed and tested in experimental animals and patients1,2. Despite many nanocarriers being tested in various diseases such as cancer, the neuromuscular field has not witnessed success with systemically delivered nanoparticles (NPs). For example, in diseases such as Duchenne muscular dystrophy (DMD)3–5, the carriers need to efficiently deliver cargo to all muscles of the body. DMD is an X-linked muscle-wasting disorder characterized by progressive muscular weakness6. Recently, Elevidys, an adeno-associated virus vector-based (AAV) gene therapy, was approved to treat pediatric patients with DMD7. However, the high dose of AAV could be associated with potential side effects, and the generation of neutralizing antibodies may limit repeated administrations of AAV8. Therefore, in addition to the viral carrier, there is considerable interest in developing non-viral carriers for DMD therapy.

The concept of a nano-enabled delivery system is an exciting development that holds promise for treating DMD. This includes the encapsulated delivery of various active pharmaceutical ingredients (APIs), such as small molecules, nucleic acids, and gene editing platforms3,9,10. While various lipid- and polymer-based nanomaterials have been tested, there has been limited success in achieving efficient systemic delivery in preclinical models3,9,10. It was demonstrated that a ~15 nm cationic nano gold-based gene editing platform could successfully deliver Cas9 RNPs and donor DNA through intramuscular administration in a DMD mouse model11. Intramuscular injection of RNPs carrying CRISPR/Cas9 led to detectable editing of the Dmd gene. While these results are promising, achieving systemic delivery to multiple muscles in DMD remains a tremendous challenge to overcome, considering there are over 640 skeletal muscles that comprise 40–50% of body mass12–14.

An intriguing question surfaces: How do nanoparticles, administered systemically, traverse the endothelial barrier to reach the muscle tissues from the bloodstream? Understanding the mechanisms could offer valuable insights to enhance nanoparticle-mediated systemic delivery. Systemically delivered nanocarriers must first interact with the vasculature before entering tissues and reaching the muscle sites. Compared to solid tumors that have been extensively studied15–19, the interactions with blood vessels in muscle tissues remain largely unknown. Our laboratory developed an ultrastructural transmission electron microscopy (TEM) procedure to investigate nanoparticle tumor access. This technique can differentiate between enhanced permeability and retention (EPR) effect versus the transcytosis transport pathway in a Kas-mutated pancreatic cancer model15,20. We also reported that tumor transcytosis is regulated by neuropilin-1, which is a physiological process needed for nutritional supply15. Recently, more evidence demonstrated the important role of transcytosis in the entry of nanoparticles into stroma-rich solid tumors16–18,21–23.

Considering the dynamic environment of dystrophic muscles, DMD shares certain similarities with solid tumors, such as a fibrotic and inflamed microenvironment, high metabolic demand, and increased pericyte coverage of the blood vessels24–26. These resemblances have motivated us to develop imageable nanocarriers for investigating the mechanism of systemic nanoparticle delivery in dystrophic muscles. Given that NP sequestration by the reticuloendothelial system (RES) in DMD animal models has often been overlooked, we also examine the RES uptake of NPs in comparison to the homing efficiency in DMD muscles. A remarkable finding using the TEM-imageable gold-labeled model NP is that IV-injected NP utilizes a vascular transport pathway through endothelial cells (ECs) in muscles. We have conducted transcytosis experiments at multiple levels: intact animal level (IVIS imaging, together with a “Zombie” interference experiment), cellular level using sorted muscle ECs (>8000 sorted cells per sample), and ultrastructural level (73 blood vessels region-of-interests from 3 mice and 3 muscle types). Drawing upon these extensive datasets, we have reached the conclusion that transcytosis plays a critical role in facilitating NP access to dystrophic muscles. This discovery offers valuable insights into the underlying mechanisms and biological processes involved in DMD systemic drug delivery. As a proof-of-concept, we further utilize the transcytotic mechanism for the systemic delivery of tamoxifen (TMX), an example cargo that is originally an anticancer drug but has been repurposed for DMD27–29. In male DMD mice, TMX NPs lead to increased utrophin expression in the gastrocnemius and triceps muscles, reduced fibrosis in the gastrocnemius, triceps, and quadriceps, and decreased necrosis in the gastrocnemius and quadriceps. Additionally, muscle strength function is also improved using TMXNPs.

Results
Synthesis of a biocompatible nanocarrier library identifies size-dependent systemic homing to dystrophic muscles

Key impediments to employing nanocarriers for effective drug delivery in DMD have included a lack of understanding of how the physicochemical characteristics of engineered NPs (e.g., size, colloidal stability, surface chemistry, etc.) impact biodistribution, pharmacokinetic (PK), RES sequestration, and ultimately efficacy and safety. While the effect of size has been intensively studied in non-dystrophic animals, a systematic dissection of the effect of size on biodistribution has not been previously reported in dystrophic mice. Fig. 1A depicts a schematic illustrating the structure of a lipid-coated mesoporous silica nanoparticle (MSNP). This drug delivery platform has been established to facilitate systemic delivery of small molecules, nucleic acids, proteins, peptides, and their combinations15,30–37. Four size-controlled MSNP cores were selected for synthesis at primary particle sizes of ~50 nm, ~100 nm, ~200 nm, and ~300 nm. This was achieved through a seed-growth process by first making 50 nm bare MSNPs using a sol-gel chemistry30,31,38, and then using them as cores to further grow the larger particles39,40. In order to obtain colloidally stable particles, the MSNP surface was further coated by a phospholipid bilayer that contains 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC): cholesterol (Chol): 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy (polyethyleneglycol)−2000] (DSPE-PEG2000) = 60:40:3 molar ratio31. Bare and lipid-coated MSNPs were characterized by TEM (Fig. 1B, upper panel) and cryogenic electron microscopy (cryo-EM) (Fig. 1B, lower panel) to assess the primary particle size, morphology, and surface coating integrity. This allowed us to show MSNP core primary sizes of 47.5±6.6–301.6±11.2 nm (Fig. 1C). Cryo-EM data confirmed that there was uniform lipid coating on all 4 samples; the thickness of the coating was 6–7 nm (Fig. 1B, lower panel). The hydrodynamic size of the lipid-coated version was 93.1±1.6–382.1±4.0 nm (Fig. 1C). All the samples exhibited a slightly negative charge at the physiological pH 7.4, with a ζ-potential value of −11 to −15 mV (Fig. 1C).

To track the biodistribution of MSNPs at the tissue and cellular level in dystrophic muscles, we covalently labeled the MSNPs with a near-infrared (NIR) dye, DyLight 680 (Dy680), with λex/em of 692 nm/712 nm. The fluorescence spectra of the labeled NPs demonstrated similar labeling efficiency at identical concentration (Supplementary Fig. 1), which enabled us to semi-quantitatively compare their in vivo biodistribution. We designated the NIR-labeled lipid-coated nanoparticles with controlled sizes as NP50nm, NP100nm, NP200nm, and NP300nm.

To evaluate NPs’ biodistribution, male dystrophic mice (~3 months) received a single intravenous (IV) injection of 10 mg/mL NIR dye-labeled NPs with different size ranges at 50 mg/kg particle dose through the tail vein. All mice survived IV injections and were sacrificed at 24 h. Skeletal and cardiac muscles were dissected and immediately imaged ex vivo using an IVIS imaging system (Fig. 1D). Contrary to previous reports on the impenetrable anatomy of skeletal muscle to nanoparticles in the wild-type (WT) animals13,41, we observed significant nanocarrier access in the dystrophic muscles. Both 50 nm and 100 nm NPs showed strong NIR fluorescence intensity in upper and lower limb skeletal muscles, including gastrocnemius, quadriceps, and triceps. NP50nm and NP100nm samples were also significantly increased in the diaphragm and tibialis anterior muscles relative to other sizes. Conversely, mice receiving NP200nm and NP300nm showed less abundance in skeletal limb muscles and limited signal in the diaphragm and heart.

In addition to muscles, we also harvested the major organs from the same mice, including the RES organs (such as liver and spleen), kidneys, and lungs. We observed a contrasting trend in that NP200nm and NP300nm nanoparticles were dramatically sequestered in RES organs (Fig. 1E). This was especially true of the spleen, which showed a significant size-dependent accumulation, where the accumulation of NP300nm was about 20-fold and 80-fold higher than the NP200nm and NP50nm treatments, respectively. Given our preference for nanocarriers with high muscle uptake and low RES capture, we have deduced that the size range between NP50nm and NP100nm could be optimal.

Ultrastructural analysis of nanoparticle transportation through a transcytosis process across endothelial cells

As it has been demonstrated numerous times that IV-injected nanocarriers do not abundantly biodistribute to skeletal muscle in healthy WT mice13,14,42–44, we set out to identify the mechanism for enhanced nanoparticle retention in dystrophic muscles. Through our size-dependent studies in the DMD preclinical model, we have pinpointed the optimal size range for biodistribution to be 50–100 nm MSNP core. However, we also acknowledge that smaller particle sizes are usually linked to lower loading capacity. Therefore, we made the decision to utilize an intermediate particle size of approximately 70 nm core (Fig. 2A) for mechanistic NP muscle access investigations (Figs. 2 and 3), followed by a proof-of-concept drug delivery study (Fig. 4).

We synthesized NIR dye-labeled NPs (using ~70 nm MSNP core), whose fluorescence properties enabled us to visualize their localization using confocal microscopy alongside multi-color immunofluorescence (IF) muscle staining in tissue sections (Fig. 2A, left). Moreover, we also synthesized ~70 nm MSNPs with a ~10 nm gold core, which could be in situ visualized due to the unique core/shell nanostructure and the high electron density of gold under tissue TEM (Fig. 2A, right). The hydrodynamic DLS size and ζ-potential value for both imageable particles were ~130 nm and ~−10 mV, respectively.

To validate the findings in Fig. 1 and additionally investigate the time-dependent particle muscle homing, NIR-labeled NPs were IV administered (50 mg/kg) via the tail vein in male DMD mice. The mice were then sacrificed at 1, 4, and 24 h, and muscle samples were immediately harvested. This experiment confirmed an effective biodistribution of NP in dystrophic muscles, revealing a noticeable trend of time-dependent NP accumulation inside various muscles (Fig. 2B). To determine whether the nanoparticles were trapped in muscle vasculature or the interstitial space, muscle tissues were collected for histological analysis. Immunofluorescence (IF) staining was used to label collagen IV (in the basement membrane surrounding muscle fibers) and CD31 (an EC marker revealing the vasculature), enabling us to look at NP distribution (Fig. 2C). Visual analysis of NIR fluorescence distribution using confocal microscopy provided clear evidence of nanocarrier extravasation in dystrophic muscles. Additional ex vivo IVIS and confocal imaging further confirmed the preferential retention of NPs in dystrophic muscles but not in the healthy muscles (Supplementary Fig. 2). Interestingly, the accumulations of NPs in liver and spleen at 24 h post injection were both significantly lower (~50% and ~30% reduction, respectively) in dystrophic mice compared to those in WT animals (Supplementary Fig. 2A). We speculate that one possible reason is the competitive retention of NPs within dystrophic muscles. This may reduce the likelihood of long-circulating NPs being captured by the RES, such as the liver and spleen. Other factors might also contribute. Moreover, to understand the influence of age, we also conducted a comparative analysis of NP distribution between young (8-week-old) and aged (30-week-old) WT and dystrophic mice. This analysis was performed 24 h after the IV injection of same labeled NPs as we described in Fig. 2. The results verified the substantial accumulation of NPs in the skeletal muscles of both young and aged mice, with minimal accumulation observed in WT mice in both ages (Supplementary Fig. 3). This finding aligned with our recent research, which demonstrated the retention of particles in various DMD models45.

While the literature reported that different nanocarriers were injected into DMD mice3,9,46,47, the exact nanocarrier transport mechanism remains largely unknown in skeletal muscle, an organ that usually does not correlate with EC permeability48. Therefore, we conducted a repeat of the IV injection experiment using gold-labeled NPs, and after 24 h of IV injection, muscles such as gastrocnemius (G), quadriceps (Q), and triceps (T) were harvested and used for TEM visualization. Although we have provided semi-quantification for TEM pictures later (see Fig. 3), we were able to occasionally visualize the involvement of a leaky fenestration (resembling EPR) and, surprisingly, the more often identification of nanoparticle transcytosis in dystrophic muscles.

To illustrate these two distinct mechanisms of nanoparticle entry to muscle, we have provided three representative blood vessels (Fig. 2D). The dystrophic muscle microenvironment where nanoparticle transportation occurs is heterogeneous due to chronic degeneration and regeneration. TEM pictures of vessels #1 and #2 were obtained at high magnification (Fig. 2D), where we observed ECs, pericytes (PC), red blood cells (RBCs), collagen deposition, and myofibers (MF). The TEM images allowed us to visualize NPs within the lumen of a blood vessel as well as their deposition in the interstitium. An intriguing and frequent observation was nanoparticle transport via a trans-EC process without the requirement of vessel leakiness. We found that numerous NPs were taken up through a process of EC transcytosis, evident by continuous and stepwise procedures, including the presence of cell membrane extrusion, filopodia interacting with NPs, formation of intra-EC vesicles (that co-localize with the NPs), and the subsequent release of NPs into the muscle interstitium (exocytosis). Specially, in the vessel #1 (Fig. 2D, middle panel), we illustrated the early stage (i.e., particle/filopodia interaction), and in the vessel #2 (Fig. 2D, right panel), we showed the intermediary/late stage (i.e., trans-EC transport and exocytosis). The zoomed-in pictures for individual NP, labeled as “α,” “β,” “γ,” and “μ,” which contained the gold-core labeling, allowed us to identify these NPs at ultra high resolution in the TEM images. We saw similar NP transcytosis features at early time points (1 and 4 h), confirming that this effect occurred as early as 1 h post IV injection (Supplementary Fig. 4).

TEM also revealed that NPs’ access to muscle tissues via leaky blood vessels, through enlarged fenestrations, but this occurred much less frequently. This process is exemplified in the case of vessel #3 (Fig. 2E). From left to right, with increased magnification, we demonstrated the presence of NPs in the lumen of a blood vessel (circulating NPs, such as “ρ” in Fig. 2E) and the deposition of NP in the interstitium (extravasated NPs, such as “π” in Fig. 2E), adjacent to the enlarged fenestrations (“leaky site”) in the EC. Similar to Fig. 2D, we could identify the individual NP through the unique core/shell structure under TEM. Another infrequent but detectable event was NP internalization in myofibers (Fig. 2F). Although this aspect is beyond the scope of the current research, it does open the possibility of utilizing our nanocarrier for gene therapy applications. However, achieving this may require additional iteration. In addition, we also conducted a comparative TEM analysis illustrating the blood vessels in WT mice. As expected, there were significantly fewer morphological features associated with NP transcytosis (e.g., filopodia engulfment, vesiculo-vacuolar organelles (VVOs) formation, and exocytosis) in the blood vessels of WT mice (Fig. 2G). Although some circulating nanoparticles were observed in the vascular lumen of WT mice, their internalization within ECs was minimal. This finding aligns with the minimal muscle uptake observed in WT mice receiving various types of nanoparticles14,44.

TEM is a prevalent approach for studying nanoparticle transport (usually in cancer scenarios)15,18; however, logistical limitations inherent to tissue TEM often lead to relatively small sample sizes18. To address this limitation, we first employed a flow cytometry-based technique to examine NP distribution in ECs from the gastrocnemius muscle of mice, dosed IV with Dy680-labeled nanoparticles; WT animals served as the control group (Fig. 2H). Given that only transcytotic NPs internalize and traffic intracellularly, we anticipate that transcytosis-involved EC should be detectable by flow cytometry. In contrast, NPs that enter tissues by extravasation (i.e., that enter tissues through leaky sites) should not be detected within ECs. Our data showed that the abundance of Dy680 positive EC cells was 5.7-folder higher in dystrophic muscles compared to WT mice (Fig. 2H). The flow cytometry gating strategy and the original data for each mouse are available online (Supplementary Fig. 5). Based on this flow data collected from four mice per group, involving over 32,000 EC counts (n=4, >8000 ECs/mouse), we have strong evidence of robust NP transcytosis in dystrophic blood vessels. This discovery has motivated us to explore the extent of transcytosis in NP transportation, essentially probing whether transcytosis plays a primary role in the dystrophic muscle site.

High frequency of transcytosis was responsible for nanoparticle accumulation in dystrophic muscles

While flow cytometry analysis suggests a role for transcytosis in nanoparticle entry into dystrophic muscles, we aimed to investigate this mechanism in greater ultrastructural detail and quantify it to the maximum extent possible. While challenging, we advocate the use of complementary methodologies to study NP transcytosis using combined TEM analysis at ultrastructural resolution (Fig. 3A) and “Zombie” model at intact animal level (Fig. 3C). Accordingly, we first performed IV injections in 3 dystrophic mice and analyzed 3 different muscle types (gastrocnemius, quadriceps, triceps) 24 h post IV injection. Among the 73 regions-of-interest (ROIs) that were collected, 37 vessels from gastrocnemius (G), 28 vessels from quadriceps (Q), and 8 vessels from the triceps (T) were analyzed (Fig. 3B1). A comprehensive summary of all the vessels, assigned with unique identifiers and muscle type information, along with the original TEM images captured at different magnifications, showcasing the key features involved in the NP transportation process, can be found online (Supplementary Data 1). From a morphological perspective, we examined six distinct features, including (i) circulating NPs inside blood vessels (Fig. 3B2), (ii) NP internalization into EC cells (Fig. 3B3, 2nd column), (iii) NP inside intra-EC vesicles (Fig. 3B3, 3rd column), (iv) NP exocytosis from ECs (Fig. 3B3, 4th column), and (v) extravasated NPs adjacent to large fenestrations (Fig. 3B4). We also examined already-transported NPs outside the blood vessels, as the sixth morphological feature (Fig. 3B5). We counted the number of blood vessels exhibiting these features in Fig. 3B3, namely EC internalization + NPs in EC vesicles + exocytosis, as blood vessels involving transcytosis. Additionally, we calculated the number of blood vessels involved in particle leakage through enlarged fenestrations (Fig. 3B4). In the total 73 vessels examined, 41 out of 73 blood vessels involved transcytosis, whereas 5 out of 73 blood vessels involved leakage (Fig. 3B6). In the gastrocnemius, quadriceps, and triceps, transcytosis-positive vessels account for 54% (20 out of 37), 53% (15 out of 28), and 75% (6 out of 8), respectively, while leaky vessels only represent 3%, 14%, and 0% in these muscle types. Based on this analysis, the chance of identifying transcytosis is 8.2-fold higher than EPR in all three muscle types we observed in dystrophic mice. Another impression was that most NPs were free and not captured by the phagocytic cells inside the bloodstream. This observation contrasts with what was noted in WT animals, where the presence of blood vessels containing nanoparticles with transcytosis features is minimal (see Fig. 2G and Supplementary Data 2).

To further illustrate the predominant role of transcytosis at the intact animal level, we conducted a “Zombie model” experiment. This experimental design, originally intended for tumor transcytosis18, was employed to showcase transcytosis in the DMD mouse model. As the “Zombie” mice were fixed and subjected to NP circulation using a peristaltic pump, the expectation was to impede active transcytotic delivery while retaining NP leakage through blood vessels. Two hours post NIR-labeled NP circulation, we observed a substantial fluorescent signal decrease of 71%, 64%, and 66% in quadriceps, gastrocnemius, and triceps in the “Zombie” dystrophic mice compared to control (Fig. 3C). This data served as additional evidence supporting the predominant role of NP transcytosis in dystrophic mice.

We have demonstrated the pivotal role of NP transcytosis in ECs, which frequently involves various membrane-associated signaling cascades, including actin polymerization and plasma membrane ruffling. In light of the quantitative flow cytometry data in Fig. 2H, we conducted IV nanoparticle administration in dystrophic mice, sacrificed the mice 24 h post injection, and performed cell sorting to distinguish between the CD31+NP+ (transcytosing ECs) and CD31+NP− (“quiescent” ECs) populations. This was followed by an RNA sequencing experiment (Fig. 3D). Differential gene expression in transcytosing ECs compared to “quiescent” ECs was shown in the Volcano plot (Fig. 3D1). We identified 50 up-regulated genes and 199 down-regulated genes. The top 25 most differentially expressed genes in transcytosing ECs were listed in the heatmap (Fig. 3D2), among which we identified several genes related to the activation of guanosine triphosphate (GTP)-binding proteins according to the NCBI information. The Rragd (Ras-related GTP binding D) and Depdc1a (DEP domain containing 1a) are related to GTPase binding activity, and Cadm4 (cell adhesion molecule 4) is able to regulate Rac protein signal transduction. From a functional perspective, these gene targets, which are linked to small GTPase activity, could play important roles during transcytosis, such as membrane ruffling, cytoskeletal changes, and the closure of macropinosomes49. Besides the most differentially expressed genes in the heatmap, we also observed up-regulation of several transcytosis-related genes, including Plvap, Lrp10, and Selp, in NP+ ECs, consistent with findings reported in the transcytosis literature (Fig. 3D3)19. To investigate whether endothelial dystrophin deficiency contributes to the differential nanoparticle uptake observed in DMD mice, we also performed comparative transcriptomic profiling of ECs isolated from DMD and WT mice, in the absence of nanoparticle exposure (Supplementary Fig. 6). In our opinion, nanoparticle transcytotic access in dystrophic muscles results from the combined effects of NP-induced stimulation (NP effect) and the endothelial sensing machinery (EC effect). While the loss of endothelial dystrophin might contribute, we tend to believe that the “NP effect” is likely the dominant driver, as the observed “fold increase” became particularly dramatic for key transcytosis markers such as Rragd, Cadm4, and Depdc1a (Fig. 3D3).

Additionally, we explored the potential involvement of macrophages, a cell type specialized in particulate clearance. To investigate this, we performed a mechanistic experiment in mice, pre-administering clodronate liposomes, which effectively deplete macrophages50,51, prior to injecting NIR-labeled NPs. Our results suggest that macrophage depletion had a low to modest impact on NP accumulation in dystrophic muscles, such as the gastrocnemius and triceps (Fig. 3E). We observed a trend toward reduced particle biodistribution in the quadriceps (~20% reduction) following macrophage depletion, although this difference did not reach statistical significance. In contrast, depletion of liver-resident macrophages (Kupffer cells) led to a marked reduction in liver, underscoring the role of macrophage-mediated nanoparticle uptake in liver. This agreed with the extensive TEM analysis (Fig. 2D, E, Supplementary Data 1) and immunofluorescence staining (Supplementary Fig. 7), which revealed that a significant number of nanoparticles were not sequestered by macrophages inside muscles at 24 h post IV injection. This information collectively supported the discovery that transcytosis plays an important role in dystrophic muscles and prompted us to investigate the potential application of transcytotic-based drug delivery for DMD treatment.

Systemic delivery of tamoxifen using the transcytotic nanocarrier led to potent efficacy in dystrophic mice

To demonstrate the potential of transcytotic drug delivery in DMD, we encapsulated tamoxifen (TMX), a cancer drug, within our nanocarrier (Fig. 4A). TMX has shown promise in promoting endocrine crosstalk between the ER and ErbB1/ErbB2 signaling pathway, leading to ER phosphorylation and transcription of utrophin (Supplementary Fig. 8)27. Additionally, TMX reduces fibrosis through the TGF-β pathway27,52. From the chemical structure aspect, TMX is poorly soluble in water, and as a result, the commercial TMX is only available in an oral form53. Daily oral treatment for 15 months was required to achieve therapeutic outcomes27.

We explored whether using our nanocarrier could overcome TMX’s water solubility issue and achieve in vivo efficacy. We achieved this by soaking empty MSNP particles into the TMX ethanol solution. The hydrophobic nature of TMX, containing a tertiary amine, facilitated efficient drug attachment onto the silica surface, which contains silanol and carries a negative charge. This attachment occurred via combined hydrophobic and electrostatic interactions (Fig. 4B). To stabilize the drug-soaked MSNPs, we employed a lipid coating procedure that effectively sealed the silica pores, similar to Fig. 2A. The resulting formulation, termed “TMX NP,” exhibited a drug loading capacity of 17.5% (w/w, drug/MSNP). The TMX NP’s hydrodynamic size was 114.2 nm, and it carried a zeta potential of −8.83 mV (Fig. 4C). Then, dystrophic mice received IV TMX NP injection, followed by blood collection at different time points and LC-MS analysis of drug concentration in the plasma. This allowed us to determine the circulatory half-life, which was ~6.9 h post IV injection (Fig. 4D).

An efficacy study was performed using a total of 11 IV injections of TMX NP in male mice at drug dose of 10 mg/kg (particle dose: ~57 mg/kg; n=7) (Fig. 4E). While the ideal comparison would have been the IV injection of free tamoxifen at an identical dose and frequency, it was technically infeasible due to the high amount of DMSO required for solubilizing the free drug. A reasonable compromise was made by including the daily oral administration of free TMX as a control by embedding TMX in the rodent diet, which was previously published to yield preclinical success in vivo27.

Since the upregulation of utrophin expression, in this case through the use of free drug or TMX NP, is considered a potential therapeutic strategy to counteract the loss of dystrophin and improve muscle function in DMD27,54, we measured utrophin expression using both immunoblotting and IF staining (Fig. 4F and Supplementary Figs. 9 and 10). We also quantified the results based on immunoblotting results (n=7 per group). Our results demonstrated that treatment with TMX NPs led to increased utrophin expression in the gastrocnemius (p=0.0409, Fig. 4F) and triceps (p=0.0082, Supplementary Fig. 9A) muscles, compared to the free drug. A similar trend was observed in the quadriceps, where both free and TMX NPs outperformed saline, although the p-value did not reach statistical significance between the free drug and TMX NP groups (Supplementary Fig. 9B). To evaluate the anti-fibrotic effects of the indicated therapy, we performed Masson’s trichrome (MT) staining and hydroxyproline (HYP) assay on various muscle samples. The results demonstrated better outcome of collagen reduction with TMX NPs treatment over the free drug, resulting in statistical significance (i.e., p=0.0254 in gastrocnemius in Fig. 4G, p=0.0090 in triceps, and p=0.0095 in quadriceps in Supplementary Fig. 11).

Given the widespread use of hematoxylin-eosin (H&E) staining in clinical assessments, we employed H&E staining to quantitatively evaluate necrotic tissue areas, according to a published TREAT-NMD Network SOP27,55. This method allows for the differentiation between Type 1 damage (necrosis with an inflammatory component) and Type 2 damage (characterized by hypercontracted myofibers and degenerating myofibers). To minimize potential observer-induced bias, our standard procedure includes H&E staining, a full tissue scan (~5×5 mm in size), and ROI identification by the same trained pathologist across the entire tissue cross-section (Fig. 4H and Supplementary Figs. 12 and 13). We demonstrated that TMX NP exerted the most pronounced effect in terms of reducing the total necrotic area specifically in the gastrocnemius and quadriceps muscles (Fig. 4H and Supplementary Fig. 13A). In triceps, TMX NPs significantly reduced necrosis compared to saline control. While TMX NPs showed a trend of improvement over the free drug (Supplementary Fig. 13B), the p-value did not reach statistical significance. Moreover, muscle strength improvement was evaluated using the grip power test, following a published SOP for DMD preclinical studies27,55. Both free TMX and TMX NPs improved grip strength compared to saline. Notably, TMX NPs treatment resulted in further improvement compared to the free drug (***p=0.0002, Fig. 4I).

The safety of MSNP drug delivery is of key importance for DMD treatment. This includes the inherent biocompatibility of the carrier as well as any potential benefits that may accrue from the API. No major toxicity was discovered, evidenced by the overall animal health monitor including body weight change (Supplementary Fig. 14A), blood chemistry (Supplementary Fig. 14B), and the H&E histology evaluation on main organs such as heart, liver, spleen, lung, and kidneys (Supplementary Fig. 14C).

Methods
Materials
Tetraethylorthosilicate (TEOS, Cat# 86578), triethanolamine (TEA-ol, Cat# 90279), cetyltrimethylammonium chloride solution (CTAC, Cat# 292737), (3-aminopropyl)triethoxysilane (APTES, Cat# 440140), gold(III) chloride hydrate (HAuCl4·3H2O, Cat# 520918), trisodium citrate dihydrate (Cat# S4641), and were purchased from Sigma-Aldrich. Lipids, including 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC, Cat# 850365), 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy (polyethylene glycol)-2000] (ammonium salt) (DSPE-PEG2000, Cat# 880120), and cholesterol (Chol, Cat# 700000) were obtained from Avanti Polar Lipids. Tamoxifen citrate (Cat# T101373) was purchased from Aladdin. The following antibodies were used: anti-laminin polyclonal antibody (Sigma-Aldrich, Cat# L9393), anti-CD31 monoclonal antibody (clone 390, BD Pharmingen, Cat# 553708), anti-collagen IV polyclonal antibody (Abcam, Cat# ab19808), anti-alpha smooth muscle actin (α-SMA) monoclonal antibody (clone 1A4, Sigma-Aldrich, Cat# A5228), anti-utrophin monoclonal antibody (Santa Cruz Biotechnology, Cat# sc-33700), and anti-F4/80 monoclonal antibody (BioLegend, Cat# 122601). Secondary antibodies were purchased from Thermo Fisher Scientific, including Alexa Fluor® 488-conjugated goat anti-rat IgG (H+L) (Cat# A11006), Alexa Fluor® 594-conjugated goat anti-rat IgG (H+L) (Cat# A11007), Alexa Fluor® 488-conjugated goat anti-rabbit IgG (H+L) (Cat# A11008), and Alexa Fluor® 594-conjugated goat anti-rabbit IgG (H+L) (Cat# A11012). HRP-linked secondary antibodies, including anti-rabbit IgG (Cat# 7074) and anti-mouse IgG (Cat# 7076), were obtained from Cell Signaling Technology. DyLight 680 NHS ester (Cat# 46418) was acquired from Thermo Fisher Scientific. All chemicals were used as received without further purification.

Nanoparticle synthesis
To investigate the size effect of nanocarriers on biodistribution in the DMD mouse model, monodisperse spherical MSNPs with different primary size range from ~50 to ~300 nm were prepared. For systemic administration, we also introduced the same lipid coating on the silica surface to maintain the colloidal stability in the biological environment. To monitor the particles via IVIS imaging or fluorescence microscopy, the MSNP core was labeled with an NIR dye. Briefly, the differently sized MSNPs were synthesized by a sol-gel chemistry using a seed-growth procedure21,22. Fifty nanometers of MSNP with a pore size of ~3 nm were synthesized with minor modifications as described in our previous reports31. Five milliliters of cetyltrimethylammonium chloride (CTAC, 25 wt% in water) solution was added to 15 mL of DI H2O, followed by stirring at 500 rpm for 10 min at 85°C. This was followed by the addition of 0.8 mL of 10% (w/v) triethanolamine (TEA-ol) for 10 min at the same temperature. Then, 1 mL of the silica precursor, tetraethylorthosilicate (TEOS), was added. The solution was stirred at 500 rpm at 85°C for 1 h, leading to the formation of particles with a primary size of ~50 nm. Synthesis of larger MSNPs was performed in the same manner as making the 50 nm particles (serving as “seed”), but adding 1 mL of the above 50 nm seed particles to the mixture of CTAC and TEA-ol solution before adding the TEOS. The final particle size was controlled by adjusting the amount of TEOS in the growth step, which led to ~100, 200, and 300 nm MSNPs. The particles were extensively washed to remove the surfactant CTAC31. The purified MSNPs were modified with amino groups and further labeled by the NIR dye (DyLight 680 NHS ester, the dye: MSNP ratio is 0.1 wt.%)15,31. For lipid coating, the MSNPs were coated with a lipid bilayer (LB)31,32. A mixture of lipids (32 mg DSPC, 10.8 mg Chol, and 5.4 mg DSPE-PEG2000, yielding a molar ratio of 3:2:0.15) was dissolved in 100 µL pure ethanol at ~65°C. One mL of a preheated (~65°C) solution, containing a 20 mg/mL NIR dye-labeled MSNP suspension, was added to the lipid solution and mixed by pipetting. The mixture was probe sonicated using a 10 s/5 s on/off cycle for 10 min. The particles suspension was purified by centrifugation and washing with PBS three times, which was further filtered through a 0.22 μm filter for sterilization. For detecting particles in tissue with TEM analysis, we also prepared MSNP with a gold core with high TEM contrast. In this case, the lipid-coated MSNPs with an ~10 nm gold core was synthesized by a three-step procedure as shown in our previous report15, i.e., first make gold nanoparticles, then grow the mesoporous silica shell on the gold core to achieve the final size ~70 nm, and finally coat the MSNPs with lipid coating.

Nanoparticle characterization
The particle morphology is characterized using a transmission electron microscope (JEOL 1200-EX), and the coated LB on MSNPs was visualized by cryo-EM (TF20 FEI Tecnai-G2). The fluorescence spectra of labeled particles were measured by a Microplate Reader (M5e, Molecular Device, USA). Particle hydrodynamic size and zeta potential were measured by a ZETAPALS instrument (Brookhaven Instruments Corporation) or Zetasizer instrument (Dynamic Light Scattering, Malvern).

Animal study
All animal work was conducted under protocols approved by the institutional Animal Research Committees at the Chinese Academy of Sciences and UCLA. The mice were maintained under a 12-h dark/light cycle, with an ambient temperature of 20–26°C and air humidity of 30–70%. The basic feed was purchased from SiPeiFu Biotechnology Co., Ltd. The main components include corn, soybean meal, fish meal, vitamins, trace elements, and amino acids, and then sterilized using 60Co irradiation. Tamoxifen-enriched feed is prepared by adding 100 mg/kg of tamoxifen (PSAITONG, cat: T80001) to the above-mentioned basic feed, which was customized and provided by Guangzhou Samo Bio-Tech Co., Ltd. Two DMD mouse models were used in this study: the classic mdx strain (C57BL/10ScSn-Dmdmdx/J, Jackson Laboratory #001801)72–74 and a dystrophin-deficient strain B10-Dmd-KO (C57BL/10ScSnJGpt-Dmdem3Cd4/Gpt, GemPharmatech #T003035). The latter is a functionally comparable MDX model commonly used in DMD research75,76. It was generated via a frameshift mutation that results in loss of dystrophin function. The specific animal strain is indicated in the figure legend. Most DMD studies use male mice because DMD is an X-linked disorder that primarily affects males. Although DMD can occur in females, it is much rarer. Approximately 2.5–7.8% of female carriers exhibit muscle weakness, and symptoms are generally milder compared to those in affected males77. Therefore, this study utilized only male animals.

Biodistribution study
Two sets of experiments were performed to compare (1) the distribution of MSNPs with various sizes and (2) the muscle distribution between dystrophic vs wild-type mice. To demonstrate the biodistribution of various MSNPs, dystrophic or WT mice received IV injection of NPs at 50 mg/kg (n=3–4 mice). At the indicated time points (1, 4, and 24 h) post injection, animals were sacrificed and followed by ex vivo imaging of the excised different muscles and major organs. Immediately following IVIS imaging, gastrocnemius, quadriceps, triceps, and tibialis anterior muscles were embedded in OCT compound. Immunofluorescent staining included TrueBlack® Lipofuscin Autofluorescence Quencher (Biotium) and blocking buffer (0.2% gelatin, 3% BSA, 10% goat serum, 0.1% tween-20 in PBS) to improve signal-to-noise ratio in identifying MSNPs by epi-fluorescent (Zeiss Observer-1) and confocal (Zeiss LSM-780 and Leica SP8) microscopy. When necessary, we also stained blood vessel markers (CD31 and α-SMA), macrophage marker (F4/80), or myofiber structure marker (collagen IV and Laminin) to assist the analysis of the intramuscular distribution of our NPs.

Similar to the aforementioned experiment, we also aimed to investigate whether age influences the entry of NPs into muscles. In this case, WT mice and dystrophic mice, aged 8 weeks or 30 weeks, respectively, received IV injections of NIR dye-labeled nanoparticles at the same particle dose (50 mg/kg). After 24 h, mice were euthanized, and quadriceps, gastrocnemius, and triceps were resected for IVIS imaging.

To examine the potential impact of macrophages on NP accumulation in dystrophic muscles, a biodistribution study was performed with or without macrophage depletion. Specifically, DMD mice were intravenously injected with 200 μL of clodronate liposomes (4 mg/mL; SN-ML-E005, SunLipo NanoTech) 24 h prior to receiving NIR dye-labeled nanoparticles, followed by animal sacrifice for ex vivo IVIS imaging.

Ultrastructural analysis of the intramuscular delivery of nanoparticles through TEM viewing
In order to reveal NP’s transportation mechanism in the dystrophic muscle, we repeated the biodistribution study using a gold-core labeled nanocarrier. Dystrophic mice (~3 months, n=3) received an IV injection of 50 mg/kg of the Au-core marked NPs. At the indicated time points (1, 4, and 24 h) post injection, animals were sacrificed, and gastrocnemius, quadriceps, and triceps muscle biopsies were collected, washed in PBS, and immediately fixed with 2.5% glutaraldehyde. Further sample preparation and sectioning were performed. After fixation in 1% OsO4, the samples were dehydrated in propylene oxide and embedded in resin. Tissue slices of 60–80 nm thick were placed on copper grids and viewed under an electron microscope (JEM-2100plus).

“Zombie” model
To reveal the contribution of transcytosis, a “Zombie” model was established in dystrophic mice, according to a published protocol that was originally designed for cancer drug delivery18. For the Zombie group, mice underwent cardiac perfusion with a washing solution (1000 U/mL heparin sodium in PBS) for 20 min using a peristaltic pump (L100-1S-1, LongerPump, China). The perfusion effluent was preserved at 4°C. Subsequently, the mice were carefully positioned to allow the flow back of the fixation solution (4% formaldehyde and 0.5% glutaraldehyde in PBS). After 1 h, the solution was replaced with a washing solution for an additional 20 min to remove any remaining fixation solution. The collecting tubes were then washed, and a nanoparticle solution (5x volume of NPs in perfusion effluent, i.e., 750 μL) was circulated for 2 h at a physiologically relevant flow rate (5–7 mL/min). For the control group, dystrophic mice received an IV injection of NPs. After 2 h, they underwent cardiac perfusion with a washing solution for 20 min, followed by the fixation solution for 1 h. Quadriceps, gastrocnemius, and triceps were then resected from these mice for IVIS imaging.

Flow cytometry analysis
The gastrocnemius muscle was dissected and processed into a single-cell solution using the Mouse Muscle Tissue Mild Enzymatic Hydrolysis Kit (RWD, DHME-10) following the manufacturer’s instructions. Subsequently, the cells suspension was filtered through a 70 μm cell filter, and any remaining RBCs were removed using RBC Lysis Buffer (Biolegend, 420301). The suspension was then labeled with 5 μL of FITC anti-CD31 Rat Monoclonal Antibody per sample (Biolegend, clone: MEC13.3, 102506) for 30 min. Before performing the flow cytometry analysis, the samples were stained with 7-AAD Viability Staining Solution (Biolegend, 420403).

Cell sorting
To isolate the NP positive (NP+) and negative (NP−) cells, we extracted the cells as mentioned above. Cell sorting was achieved via a Fluorescence Flow Cytometry Sorting System (MoFlo XDP, Beckman Coulter). The muscle ECs were sorted into two populations: CD31+NP+ (Dy680+, APC channel) and CD31+NP− in PBS before immediate transfer into TRIzol and delivered to a core facility (Beijing Genomics Institution, BGI Genomics) for RNA sequencing.

RNA sequencing and analysis
The RNA extraction, RNA library preparation, sequencing, and initial data analysis were done by BGI. CD31+NP+ and CD31+NP− muscle ECs of gastrocnemius from three different mdx mice were used and pooled together, respectively, for the sequencing. RNA extraction was done according to the standard procedure. The gene hit counts were used for differential expression analysis using DESeq2. The test was used to generate log2-fold change and Q-values. Genes that had a log2-fold change > 1 and an adjusted Q value ≤ 0.05 were named as differentially expressed genes. The list of up-regulated genes in NP+ muscle ECs was analyzed for gene ontology enrichment.

Demonstration of the efficacy of transcytotic tamoxifen delivery in vivo
In order to demonstrate the impact of drug delivery through a transcytotic nanocarrier mechanism, we used lipid-coated MSNP for systemic delivery of a highly insoluble drug, TMX, which was recently repurposed for DMD management. Briefly, TMX was soaked into MSNP, followed by lipid coating, purification, and characterization before IV injection.

To prepare tamoxifen-laden nanoparticles (TMX NPs), tamoxifen was first loaded in MSNPs (~70 nm) and then coated with the LB as described above. Briefly, tamoxifen (20 mg, 10 mg/mL dissolved in ethanol) was added to MSNP (40 mg, 63 mg/mL in ethanol), mixed and stirred overnight at room temperature. To remove the free tamoxifen, the mixture was centrifuged at 21,000×g for 15 min, then the tamoxifen-MSNP pellet was resuspended in PBS by bath sonication. The tamoxifen-MSNP suspension was then transferred to a lipid ethanol solution, followed by probe sonication for further lipid coating. The final TMX NPs were purified by washing and centrifugation in PBS and filtered with a 0.22 μm sterile membrane. To determine the tamoxifen concentration in the TMX NPs, the particle solution was diluted 1/100 in pure ethanol before the absorbance was measured at 285 nm for tamoxifen (Spark, TECAN). The lipid coating, hydrodynamic diameter, and zeta potential of TMX NPs were characterized by cryo-EM and dynamic light scattering, respectively.

To characterize the pharmacokinetics of TMX NPs, male dystrophic mice (n=3) received a single IV injection of TMX NPs at a TMX dose of 10 mg/kg. The blood samples were collected at 5 min, 1, 3, 6, and 24 h. After separation of the plasma fraction, drugs were extracted with acetonitrile (plasma: acetonitrile 1:4 v/v) followed by vigorous shaking in a tissue homogenizer (QIAGEN, Tissue Lyser II). The plasma suspensions were then centrifuged at 16000×g at 4°C for 10 min. The supernatant was filtered through a 0.22 μm sterile membrane before being loaded to Liquid Chromatography/Mass Spectrometry (LC/MS). The LC/MS (Agilent, 6495C) analysis condition: LC was performed using a Symmetry C18 column (Waters, WAT045905, 4.6×150 mm) at a flow rate of 1.0 mL/min, injection volume of 10 μL, 280 nm absorbance wavelength, and a temperature of 30°C. The LC mobile phase was grade acetonitrile/H2O (50:50 ratio, isocratic). Concentrations of TMX were calculated by determining the area under the curve, using a calibration curve that was established by known concentrations of the standard Tamoxifen compound. The following transitions were used for MRM acquisition: TMX m/z 372-72 (quantitation). The PK data were fitted in a one-compartment model, using PKSolver software78.

For the efficacy study, male mice (~3 months) received a total of 11 IV injections (n=7 mice) as we outlined in Fig. 4E. The TMX and MSNP doses were 10 mg/kg and 57 mg/kg, respectively. The controls include saline (n=7 mice) or free drug (n=7 mice). The poor TMX water solubility prevents us from including free drug control (IV), as we clarified in the manuscript. Instead, a TMX-containing rodent diet (100 mg drug/kg food) was used for daily oral administration of free tamoxifen (~10 mg/kg), using a published protocol in the literature27. During the treatment, animal health conditions were monitored. At the conclusion stage of the experiment, the muscle strength improvement was evaluated by the grip power test27. Muscle strength was measured by the grip power test using a Mouse Grip Tester (Model YLS-13A, Jinan Yiyan Tech Co., LTD). Briefly, each mouse was gently placed on the instrument, and its tail was pulled at a constant speed in a horizontal direction. The instrument recorded the maximum pull force. This procedure was repeated four times per mouse, and the average value was calculated. Then, the animals were sacrificed, the gastrocnemius, quadriceps, and triceps muscles, and other main organs were collected and fixed in 10% formalin for 48 h before being paraffin-embedded. The 6 μm-thick transverse sections were performed with MT staining or H&E staining dye according to standard procedures via an automatic dying machine (DRS-Prisma-P-JCS & Film-JC2, SAKURA, Japan). To quantify the histopathology in the above-mentioned stained muscle sections, the entire cross-sectional areas of muscles were scanned and analyzed. Histological features were identified manually by the researcher and quantified using image analysis software (SlideViewer, 3DHISTECH Ltd., v2.6.0). To quantitate the abundance of a specific histological feature (e.g., myofiber necrosis of Type 1 damage with an inflammatory component or Type 2 of hypercontracted myofibers and degenerating myofibers) per muscle cross-sectional area, the areas were drawn around to get an absolute value (μm2). The area % was calculated as abnormal area/whole area. Histological features were identified according to the TREAT-NMD protocol DMD_M.1.2.007 (Quantification of histopathology in H&E stained muscle sections)27,55.

For utrophin immunoblotting, muscle tissues were homogenized in RIPA buffer containing protease inhibitors. Equal amounts of total protein (40 μg) from muscle extracts were loaded onto 6% Tris-Acetate gels and transferred to a 0.45 μm PVDF membrane over 6.5 h. Membranes were blocked with 5% milk in TBST for 1 h and incubated with primary antibodies against utrophin (1:300, sc-33700, Santa Cruz) or vinculin (1:100,000, A2752, Abclonal). Protein bands were visualized using a chemiluminescence detection system (Amersham ImageQuant 800, Cytiva) and quantified using ImageJ software. For immunofluorescence staining for utrophin, 6 μm-thick transverse paraffin sections were obtained as mentioned above. After dewaxing and rehydration, 3% H2O2 solution in methanol was used to block endogenous peroxidase activity at room temperature for 10 min. Next, antigen retrieval was performed using a citrate buffer (10 mM citrate buffer, pH=6.0) at 95–100°C for 10 min. The sections were then blocked in 5% BSA in PBS for 60 min at 37°C, followed by incubation with primary antibody (monoclonal antibody, sc-33700, Santa Cruz) overnight and Alexa Fluor 488-conjugated secondary antibody for 60 min. Images were finally viewed and captured under a confocal microscope (A1 HD25, Nikon). Section images were scanned with a research slide scanner (SLIDEVIEW VS200, Olympus).

Muscle fibrosis, indicated by collagen content, was quantified using a hydroxyproline assay kit (A030-2-1, Nanjing Jiancheng Biotechnology Co., Ltd.). Moreover, to further assess the biosafety of TMX NP, the levels of alanine aminotransferase (ALT), aspartate aminotransferase (AST), blood urea nitrogen (BUN), and creatinine (CRE) were measured.

Statistical analysis
Comparative analysis of differences between groups was performed using the two-tailed Student’s t-test (Excel software, Microsoft) for two-group comparison. A One-way ANOVA test was performed for multiple group comparisons on Prism 7 software (GraphPad). Data were expressed as mean ± SD or SEM, as stated in the figure legends. A statistically significant difference was considered at p < 0.05.

Ethics statement
All animal experiments were performed following the ARRIVE guidelines for the reporting of in vivo studies and conducted under protocols approved by the institutional Animal Research Committees at the Chinese Academy of Sciences (2022R0014 and AP2024-08-0097) and the University of California, Los Angeles (ARC 2006-119-31A). All the animal studies utilized only male animals, given the biological relevance to DMD, an X-linked disorder that predominantly affects males.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In this revised manuscript, the authors provided compelling evidence demonstrating transcytosis as a primary mechanism for systemic delivery of mesoporous silica nanoparticles to dystrophic muscle. The authors further showed improved therapeutic outcomes with mesoporous silica nanoparticle-mediated delivery of tamoxifen in the mdx mouse model. I only have a few comments.\n1. The diaphragm and heart are the most affected tissues in muscular dystrophy. However, these tissues are poorly targeted by mesoporous silica nanoparticles. This may represent an important translational limitation and should be discussed.\n2. The dramatic difference in mesoporous silica nanoparticle delivery between wild-type and dystrophic muscle suggests that the loss of dystrophin may have significantly altered endothelial function. Is dystrophin expressed in endothelial cells? Are transcytosis biomarkers significantly augmented in dystrophic muscle?\n3. The detargeting of mesoporous silica nanoparticles from the liver and spleen of dystrophic animals is intriguing. It would be helpful if authors could provide a quantitative comparison and discuss the potential mechanism(s).", "reviewer_4": "The authors have carried out a macrophage depletion experiment to address the concerns of macrophages. The Figure 3E seems to show a fairly substantial reduction in quadriceps - it is marked as non-significant, but this must be due to the low n=3. The authors need to be less emphatic about their statements that macrophages do not contribute - they seem to contribute something to at least quadriceps (and liver)."}, "author_response": "We appreciate the additional comments from the reviewers #1 and #4 and the editorial team. To facilitate the next stage of review, we have provided detailed point-by-point responses below and the modifications have been highlighted in yellow within the marked version of revised manuscript.\nReviewer #1\nComment: In this revised manuscript, the authors provided compelling evidence\ndemonstrating transcytosis as a primary mechanism for systemic delivery of mesoporous\nsilica nanoparticles to dystrophic muscle. The authors further showed improved therapeutic\noutcomes with mesoporous silica nanoparticle-mediated delivery of tamoxifen in the mdx\nmouse model. I only have a few comments.\nResponse: We thank the reviewer for acknowledging the “compelling evidence” we have\nprovided and for recognizing the “improved therapeutic outcomes” demonstrated in our\nstudy. Please find our responses to the remaining comments below.\nComment 1. The diaphragm and heart are the most affected tissues in muscular dystrophy.\nHowever, these tissues are poorly targeted by mesoporous silica nanoparticles. This may\nrepresent an important translational limitation and should be discussed.\nResponse: Thank you. We agree that managing cardiomyopathy in DMD remains a\nsignificant and longstanding challenge in the field.\nIn our study, we observed low nanoparticle uptake in the diaphragm, with particularly minimal\naccumulation in the heart. This phenomenon is likely driven by multiple factors, and we\nsuspect that it is not exclusive to silica-based nanocarriers. One contributing factor could be\nthe rapid blood flow through the heart, which limits the residence time of nanoparticles and\nreduces their opportunity to interact with the vascular endothelium. Additionally,\ncardiomyocytes exhibit low endocytic activity and are not inherently phagocytic, further\nrestricting nanoparticle internalization. These physiological characteristics together pose\nsignificant barriers to effective nanoparticle delivery to cardiac tissue in DMD. One possible\ndesign is to incorporate targeting moieties like cardiomyocyte-targeting T9 peptide (Peptides\n2010, 31, 1873) to enhance biodistribution specifically to the heart. As the reviewer noted the\n“translational implication”, it is important to emphasize that any additional design\nmodifications, such as peptide conjugation, would increase the complexity of the\nnanoparticles, might potentially limiting their translational potential from a Chemistry,\nManufacturing, and Controls (CMC) aspect. Therefore, we\n1\nhave incorporated these points into the limitations section at the end of our manuscript.\nIt reads:\nAnother limitation of our study is the limited capacity to address cardiomyopathy in DMD,\nwhich could be a key focus for future iterations of this direction. In our study, we observed\nlow nanoparticle uptake in the diaphragm, with particularly minimal accumulation in the\nheart. This phenomenon is likely driven by multiple factors, and we suspect that it is not\nexclusive to silica-based nanocarriers. One contributing factor could be the rapid blood\nflow through the heart, which limits the residence time of nanoparticles and reduces their\nopportunity to interact with the vascular endothelium. Additionally, cardiomyocytes\nexhibit low endocytic activity and are not inherently phagocytic, further restricting\nnanoparticle internalization. These physiological characteristics together pose significant\nbarriers to effective nanoparticle delivery to cardiac tissue under normal conditions. One\npossible design was to incorporate targeting moieties like cardiomyocyte-targeting T9\npeptide to enhance biodistribution specifically to the heart. It is also important to balance\ndesign complexity with target efficiency, as excessive manufacturing complexity may\nraise difficulties in translation.\nComment 2. The dramatic difference in mesoporous silica nanoparticle delivery between\nwild-type and dystrophic muscle suggests that the loss of dystrophin may have significantly\naltered endothelial function. Is dystrophin expressed in endothelial cells? Are transcytosis\nbiomarkers significantly augmented in dystrophic muscle?\nResponse: Thank you. There are three questions in this comment, and we would like to\naddress them one by one.\n(1) Question about “the loss of dystrophin may have significantly altered endothelial\nfunction.”\nResponse: This drug delivery investigation is to study the transport mechanisms by which\nnanoparticles access skeletal muscle in mdx mice. This process is influenced by the\nintrinsic characteristics of endothelial cells (ECs) in mdx mice and, more importantly in our\nview, by the stimuli imposed on ECs by intravenously injected nanoparticles. Sequencing\ndata revealed key alterations in transcytotic mdx ECs following nanoparticle exposure (Fig.\n3D). This experimental design (current data in Fig. 3) evaluates the impact of nanoparticle\ntreatment within the context of dystrophic mdx ECs.\nDetermining whether the loss of dystrophin significantly alters EC function is an important\npathophysiological question. However, it introduces a new set of variables to answer\nnanoparticle-related questions. For example, the relative contribution of each potential factor\nneeds to be quantitatively assessed. It may also lead to a potential shift in focus regarding\nthe disease model, as our work centers on drug delivery for DMD. Therefore, addressing\nthis question would likely require a separate experimental design and falls\n2\nbeyond the scope of this already comprehensive study.\nTo accommodate the reviewer’s question, however, we would like to share our unpublished\nsequencing data with reviewer 1, which we hope to answer your question. We conducted\na comparative sequencing analysis of primary ECs isolated from mdx and wild-type (WT)\nmice, without the use of nanoparticles. This analysis identified the top 100 up- and down-\nregulated genes, providing a foundation for formulating hypotheses for future investigations\ninto EC function (see below Figure). Given the scope and complexity of this topic, it is not\nfeasible to address all aspects within a single manuscript, and we plan to explore these\nfindings in a separate study.\nFigure in response letter: RNA sequencing analysis of primary ECs isolated from mdx\nand wild-type (WT) mice.\n3\n(2) Question about “Is dystrophin expressed in endothelial cells?”\nResponse: Yes. Previous studies have reported that dystrophin is expressed not only in\nmuscle cells but also in vascular ECs (Circulation, 2001, 103: 864-870; Arterioscler Thromb\nVasc Biol, 2013, 33: 2867).\nAfter confirming with the mdx vendor that the mdx strain was generated using a frameshift\nmutation strategy, we also observed a marked contrast in dystrophin expression between\nwild-type and mdx ECs in the data above.\n(3) Question about “Are transcytosis biomarkers significantly augmented in dystrophic\nmuscle?”\nResponse: In the context of Comment 2, we assume that the reviewer is asking whether\ntranscytosis biomarkers are elevated in dystrophic muscle of untreated mdx mice compared\nto untreated WT mice, without nanoparticles. We therefore looked at the transcytosis-\nrelated genes identified in this study (Fig. 3).\nFigure in response letter: Comparison of the gene expression related to the activation of\ntranscytosis. Left, Transcytosis-related genes in WT ECs vs mdx ECs. Right, Data redisplay\nof Fig. 3D3 in the main manuscript. Mdx mice received IV injection of lipid-coated MSNPs at\n24 h, followed by EC sorting and sequencing as discussed in main manuscript. Based on this\npreliminary analysis, the expression levels of Rragd, Cadm4, Plvap, Lrp10, and Selp remained\nwithin a ±2-fold range when comparing WT ECs to mdx ECs (left panel). However, the fold\nchanges for these genes became more pronounced when comparing NP+ mdx ECs to NP−\nmdx ECs (right panel, that is Fig. 3D3 in our paper). Interestingly, Depdc1a exhibited an\nincrease of over 28-fold in mdx ECs compared to WT ECs, and this difference became more\ndramatic upon nanoparticles exposure.\nAs our unpublished sequencing data only offer preliminary indications, further confirmatory\nstudies and dedicated experimental designs are still needed to determine how the loss of\ndystrophin alters EC function.\n4\n3. The detargeting of mesoporous silica nanoparticles from the liver and spleen of\ndystrophic animals is intriguing. It would be helpful if authors could provide a quantitative\ncomparison and discuss the potential mechanism(s).\nResponse: Thank you for the comments. As shown in the revised Fig. S2A, we conducted\na quantitative comparison of NP distribution in major organs between mdx and WT mice. At\n24 hours post-injection, NP accumulation in the liver and spleen was significantly lower in\nmdx mice, i.e., by approximately 50% and 30%, respectively, compared to WT controls. One\npossible explanation is the competitive retention of NPs within dystrophic mdx muscles,\nwhich may reduce their availability for uptake by the reticuloendothelial system, including\nthe liver and spleen. Other contributing factors may also be involved, and this complexity\nwarrants further in-depth investigation. These analyses and discussions have been\nincorporated into the revised manuscript.\nRevised Supplementary Fig. 2 (A) Ex vivo NIR fluorescence imaging and quantification\nof muscles and organs at 24 h after C57BL/6 mice or mdx mice received IV injection of\nNIR-labeled lipids coated 70 nm MSNPs (50 mg/kg). Data represent mean ± SEM (n= 4\nmice). Statistical analysis was performed using a 2-tailed Student’s t-test; *p < 0.05, **p <\n0.01, ***p < 0.001.\nReviewer #4:\nComment: The authors have carried out a macrophage depletion experiment to address\nthe concerns of macrophages. The Figure 3E seems to show a fairly substantial reduction\nin quadriceps - it is marked as non-significant, but this must be due to the low n=3. The\nauthors need to be less emphatic about their statements that macrophages do not\ncontribute - they seem to contribute something to at least quadriceps (and liver).\nResponse: Regarding the macrophage depletion experiment suggested by Reviewer 4,\nour original statement was: “ ...macrophage depletion...suggesting that macrophages\nplayed a limited role in NP transport to mdx muscles (Fig. 3E).” In response to the suggestion\nof \"less emphatic about their statements that macrophages do not contribute\", we have further\nrefined the statement to make it more descriptive. It now reads as follows:\nOur results suggest that macrophage depletion had a low to modest impact on NP\naccumulation in dystrophic muscles, such as the gastrocnemius and triceps. We\nobserved a trend toward reduced particle biodistribution in the quadriceps (~20%\nreduction) following macrophage depletion, although this difference did not reach\nstatistical significance. In contrast, depletion of liver-resident macrophages (Kupffer\ncells) led to a marked reduction in liver, underscoring the role of macrophage-mediated\nnanoparticle uptake in liver.\nThank you. We hope the revised manuscript meets the journal’s requirements and is\nsuitable for publication.\nHuan Meng, April Pyle and Mellisa Spencer (on behalf of coauthors)\n6"}, {"round": 2, "reviewer_comments": {"reviewer_1": "1. An important finding of the study is the enhanced nanoparticle uptake in the mdx mouse muscle. Some explanations are\nneeded, given the striking difference between wild-type mice and mdx mice. The authors indeed have data to explain their\nfindings (RNA seq data and gene profile analysis Table in the response letter). These data should be included in this\nmanuscript as Supplementary information to support their finding. This information is important as these changes could be\ndue to the loss of endothelial dystrophin, hence, specific to DMD and not applicable to other types of muscular dystrophies.\n2. The authors explained the poor uptake of their nanoparticles in the heart. They cited \"rapid blood flow through the heart\"\nand \"low endocytic activity\". References should be provided to support these statements. Is blood flow in the heart faster\nthan that in skeletal muscle? Skeletal muscle also has low endocytic activity.\n3. The authors did not explain why the diaphragm also had poor uptake. DMD patients die from diaphragm failure and/or\nheart failure. Hence, the newly developed nanoparticles may have limited translation significance for treating DMD. They\nshould acknowledge these limitations.\n4. In the method section, there is no description of the animals used in the study. In the response letter, the authors indicated\nthat the mdx mouse used in the study was generated using a frameshift mutation strategy. The mdx model was established\nin 1989. It is well-established that this model carries a nonsense mutation. Clearly, the authors did not use mdx mice in their\nstudy; rather, a different dystrophin-deficient mouse was used."}, "author_response": "Response Letter\nThank you for handling our manuscript (NCOMMS-25-30019A), now entitled \"Transcytotic Transportation\nof Size-Controlled Nanocarriers into Dystrophic Skeletal Muscle Leads to Therapeutic Outcome in Mice\".\nWe appreciate the additional comments from the reviewers #1 and the editorial team. To facilitate the next\nstage of review, we have provided detailed point-by-point responses below and the modifications have\nbeen highlighted in yellow within the marked version of revised manuscript.\nReviewer #1\nComment 1. An important finding of the study is the enhanced nanoparticle uptake in the mdx mouse\nmuscle. Some explanations are needed, given the striking difference between wild-type mice and mdx\nmice. The authors indeed have data to explain their findings (RNA seq data and gene profile analysis Table\nin the response letter). These data should be included in this manuscript as Supplementary information to\nsupport their finding. This information is important as these changes could be due to the loss of endothelial\ndystrophin, hence, specific to DMD and not applicable to other types of muscular dystrophies.\nResponse: Thank you. We are pleased that the reviewer views the sequencing data comparing wild-type\nand DMD endothelial cells (ECs) as “information [that] is important.” We also acknowledge the suggestion\nof “these data should be included”. In the revised version, we followed this suggestion and included the\ndata as Supplementary Figure 6.\nIn our opinion, nanoparticle (NP) transcytotic access in dystrophic muscles results from the combined\neffects of NP-induced stimulation (NP effect) and the endothelial sensing machinery (EC effect). The data\npresented in the main manuscript (Fig. 3D) focus on the “combined effects”. Similar experimental designs\nwere frequently used in drug delivery research (Nat. Mater. 2020, 19, 566; ACS Nano 2021, 15, 14080;\nNat. Nanotechnol. 2025, 20, 672). Based on the new sequencing data, while the loss of endothelial\ndystrophin might contribute, we tend to believe that the “NP effect” is likely the dominant driver, as the\nobserved “fold increase” became particularly dramatic for key transcytosis markers such as Rragd, Cadm4,\nand Depdc1a. For the question about “other types of muscular dystrophies”, we do not have a definitive\nanswer at this moment because the focus of this work is DMD.\ntype (WT) mice. AA,, Top 100 up- and down-regulated genes identified by RNA sequencing analysis of\nprimary ECs isolated from DMD and WT mice. BB,, Comparison of the gene expression related to the\nactivation of transcytosis.\nComment 2.The authors explained the poor uptake of their nanoparticles in the heart. They cited \"rapid\nblood flow through the heart\" and \"low endocytic activity\". References should be provided to support these\nstatements. Is blood flow in the heart faster than that in skeletal muscle? Skeletal muscle also has low\nendocytic activity.\nResponse: Thank you.First, we would like to clarify that drug delivery to the heart remains a developing\narea with limited experimental data. In the final paragraph of the discussion, we deliberately used the terms\n“likely” and “could be” to reflect the speculative nature of tthhereasoningabout NP access in heart. Perthe\nreviewer’ssuggestion, we have also improved the reference section accordingly.\nThe poor uptake of intravenously injected nanoparticles in the heart results from a combination of unique\nphysiological, cellular, and hemodynamic barriers. Cardiac blood flow velocity is significantly higher than\nin skeletal muscle due toits continuous workload and limited oxygen extraction reserve (ref: Physiological\nReviews, 2008, 88, 1009). This rapid, pulsatile flow could create a delivery barrier for intravenously\nadministered nanoparticles by reducing endothelial contact time and increasing shear-mediated clearance\n(ref: Nature Reviews Cardiology,2021, 18, 389).Assuming nanoparticles could go through the EC barrier\nin heart, additional barriers do exist. Cardiomyocytes are terminally differentiated with minimal\nendocytic/vesicular activity(refs: Physiological Reviews,2007,87, 521; Biochemical Society Transactions,\n2024, 52, 41). Their dense contractile machinery and mitochondrial/myofibrils volume could further limit\nnanoparticle internalization (ref: Journal of Molecular and Cellular Cardiology, 1992, 24, 669). We have\nupdated these references and information in the revised manuscript.”\ndiaphragm failure and/or heart failure. Hence, the newly developed nanoparticles may have limited\ntranslation significance for treating DMD. They should acknowledge these limitations.\nResponse: Thank you. The reviewer has raised the concern about low uptake in the diaphragm before,\nand we have acknowledged this limitation in our response letters and revised versions. In fact, this\nchallenge highlights a keylimitation in the field (see table below).\nTable: Summary of prior literature on nanoparticle access to the diaphragm.\npreviously cited studies.\nWhile current understanding of how nanoparticles access dystrophic muscle tissue remains limited, our\ndata showed low but detectable nanoparticle accumulation in the diaphragm. We surmise that the poor\naccess in this muscle type may be attributed to its unique physiological features, including high collagen\ndensity, distinct vascular architecture, and continuous mechanical stress (refs: Journal of Histochemistry\n& Cytochemistry 2012, 60, 638; Matrix Biology Plus 2023, 18, 100131). We hypothesize that these factors\nmay restrict nanoparticle extravasation or retention. In the future, phage display technology could be\nleveraged to identify diaphragm-targeting peptides, which may enable further functionalization of\nnanocarriers for enhanced delivery.\nTo collectively address Comments 2 and 3, we have revised the statement in the final paragraph of the\nDiscussion section. It now reads:\nAnother limitation of our study is the limited capacity to address cardiomyopathy and diaphragmatic\nmyopathy in DMD, which could be a key focus for future iterations of this direction. In our study, we\nobserved low nanoparticle uptake in the diaphragm, with particularly minimal accumulation in the heart.\nThis phenomenon is likely driven by multiple factors, and we suspect that it is not exclusive to silica-\nbased nanocarriers. For diaphragm, its unique physiology with high collagen density, distinct vascular\narchitecture, and continuous mechanical stress may limit the nanoparticles accumulation60, 61. For\nheart, one contributing factor could be the rapid cardiac blood flow, which limits the residence time of\nnanoparticles and reduces their opportunity to interact with the vascular endothelium62, 63. Additionally,\ncardiomyocytes are terminally differentiated with minimal endocytic/vesicular activity64, 65 and are not\ninherently phagocytic, further restricting nanoparticle internalization. Their dense contractile\nmachinery and mitochondrial/myofibrils volume further limit nanoparticle internalization66. One\npossible design was to incorporate targeting moieties like cardiomyocyte-targeting T9 peptide67 or\ncell-penetrating peptides (Pip6)68 to enhance biodistribution specifically to the heart. It is also possible\nto use phage display technology to identify diaphragm-targeting peptides, which could be employed\nto further functionalize nanocarriers for improved delivery. At the same time, it is necessary to balance\ndesign complexity with target efficiency, as excessive manufacturing complexity may raise difficulties\nin translation69.\nComment 4. In the method section, there is no description of the animals used in the study. In the response\nletter, the authors indicated that the mdx mouse used in the study was generated using a frameshift\nmutation strategy. The mdx model was established in 1989. It is well-established that this model carries a\nnonsense mutation. Clearly, the authors did not use mdx mice in their study; rather, a different dystrophin-\ndeficient mouse was used.\nResponse: Thank you very much for pointing this out. This project was conducted during the pandemic\nand coincided with the research team's relocation, during which the team sourced animals from different\nvendors. As a result, two DMD mouse models were used in this study: the classic mdx strain\n(C57BL/10ScSn-Dmdmdx/J, JAX Laboratory #001801) and a dystrophin-deficient strain B10-Dmd-KO\n(C57BL/10ScSnJGpt-Dmdem3Cd4/Gpt, GemPharmatech #T003035). The latter is a functionally comparable\n4\nmdx model commonly used in DMD research. It was generated via a frameshift mutation that results in\nloss of dystrophin function. This model is marketed by the vendor as an 'mdx' model and is referred to as\nsuch in preclinical research literature (e.g., Cell Reports 2022, 39, 110939; The EMBO Journal 2025, 44,\n30).\nBoth models yielded highly similar results in terms of nanoparticle biodistribution and ultrastructural\nfeatures of transcytosis. Specifically, Fig. 1E demonstrates size-dependent nanoparticle retention in the\nJAX mdx model, while Fig. 2B confirms this trend in the GemPharmatech DMD model in the context of\ntime-dependent retention. Transcytosis features were clearly observed in both models, as shown in Fig.\n2D (JAX classic mdx) and Fig. S4 (GemPharmatech B10-Dmd-KO). To avoid any confusion, we have\nspecified the animal model used throughout the revised manuscript.\nWe hope the revised manuscript meets the journal’s requirements and is suitable for publication.\nHuan Meng, April Pyle and Mellisa Spencer (on behalf of coauthors)\n5"}, {"round": 3, "reviewer_comments": {"reviewer_1": "The authors did a great job addressing all the concerns. I have no further questions."}, "author_response": "Response Letter\nREVIEWERS' COMMENTS: Reviewer #1 (Remarks to the Author):\nThe authors did a great job addressing all the concerns. I have no further questions.\nResponse: Thank you.\nWe hope the revised manuscript meets the journal’s requirements and is suitable for\npublication.\nHuan Meng, April Pyle and Mellisa Spencer (on behalf of coauthors)\n3"}]

Decision letter