Abstract

   Human parathyroid hormone (1–34) (PTH) exhibits osteoanabolic and
   osteocatabolic effects, with shorter plasma exposure times favoring
   bone formation. Subcutaneous injection (SCI) is the conventional
   delivery route for PTH but faces low delivery efficiency due to limited
   passive diffusion and the obstruction of the vascular endothelial
   barrier, leading to prolonged drug exposure times and reduced
   osteoanabolic effects. In this work, a microcurrent delivery system
   (MDS) based on multimicrochannel microneedle arrays (MMAs) is proposed,
   achieving high efficiency and safety for PTH transdermal delivery. The
   internal microchannels of the MMAs are fabricated using high‐precision
   3D printing technology, providing a concentrated and safe electric
   field that not only accelerates the movement of PTH but also reversibly
   increases vascular endothelial permeability by regulating the actin
   cytoskeleton and interendothelial junctions through Ca^2+‐dependent
   cAMP signaling, ultimately promoting PTH absorption and shortening
   exposure times. The MDS enhances the osteoanabolic effect of PTH in an
   osteoporosis model by inhibiting osteoclast differentiation on the bone
   surface compared to SCI. Moreover, histopathological analysis of the
   skin and organs demonstrated the good safety of PTH delivered by MDS in
   vivo. In addition to PTH, the MDS shows broad prospects for the
   high‐efficiency transdermal delivery of macromolecular drugs.

   Keywords: microchannels, microcurrent, microneedle, osteoanabolic
   effect, PTH
     __________________________________________________________________

   This study proposes a microcurrent delivery system (MDS) using
   multimicrochannel microneedle arrays (MMAs) to improve human
   parathyroid hormone (1–34) (PTH) transdermal delivery. By accelerating
   PTH movement and reversibly increasing vascular permeability, the MDS
   reduces PTH exposure time and enhances its osteoanabolic effects in
   osteoporosis by inhibiting osteoclast differentiation. The MDS also
   holds significant potential for the efficient transdermal delivery of
   other macromolecular drugs.

   graphic file with name SMTD-9-2401144-g003.jpg

1. Introduction

   Osteoporosis (OP) is characterized by reduced bone mass, damage to bone
   microstructure, and compromised strength, which render elderly
   individuals vulnerable to low‐energy fractures.^[ [44]^1 ^] Although
   osteoanabolic drugs are more effective than anti‐osteocatabolic drugs
   in promoting bone formation, their ability to increase hip bone mineral
   density (BMD) is limited to approximately 5%,^[ [45]^2 ^] which is
   insufficient to meet the needs of middle‐aged and elderly patients with
   severe OP for long‐term bone health. As the longest‐used and most
   clinically evidenced osteoanabolic drug, human parathyroid hormone
   (1–34) (PTH) also has a catabolic effect that promotes bone
   resorption.^[ [46]^3 ^] Previous studies have attempted to weaken or
   eliminate the osteocatabolic action of PTH while maintaining its
   osteoanabolic effect to generate a more potent osteogenic ability,^[
   [47]^4 ^] but only limited progress has been achieved to date.

   Exposure time, rather than C[max], is crucial to the mechanism of PTH's
   anabolic action.^[ [48]^5 ^] Exposure time is defined as the duration
   from appearance in plasma until complete elimination. The exposure time
   of PTH subcutaneous injection (SCI) is prolonged due to the obstruction
   of the vascular endothelial barrier and the molecular weight of PTH
   (4117.77 Da), leading to sustained enhancement of osteoclast
   differentiation even after maximal osteogenic induction is achieved.^[
   [49]^6 , [50]^7 , [51]^8 ^] Although many studies have attempted to
   explore PTH analogues with shorter plasma exposure times, limited
   effects have been achieved.^[ [52]^9 , [53]^10 ^]

   PTH is a positively charged peptide when dissolved at physiological pH
   due to the presence of multiple basic amino acid residues, such as
   lysine and arginine.^[ [54]^11 ^] Therefore, an electric field, in the
   form of electroporation or microcurrent, can provide an additional
   electro‐driving force to accelerate the movement of PTH in subcutaneous
   interstitial fluid. Furthermore, electroporation and microcurrent have
   been proposed to increase vascular endothelial permeability,^[ [55]^12
   , [56]^13 ^] which is thought to facilitate the transendothelial
   transport of subcutaneous PTH. These synergistic effects can promote
   the absorption rate of subcutaneous PTH and reduce exposure time. The
   endothelial barrier, formed by lateral connections at cell–cell
   junctions, is a critical factor in determining vascular permeability.^[
   [57]^14 ^] Central to the regulation of the endothelial barrier is
   transmembrane adhesion proteins, whose backbone of VE‐cadherin
   anchoring together with the actin cytoskeleton forms the barrier
   interface.^[ [58]^15 ^] Various extracellular stimuli, including
   electro‐stimulation, can modulate barrier function by altering
   VE‐cadherin and the cytoskeleton, with Ca^2+ signaling playing a vital
   role. Increased intracellular Ca^2+ triggers various signaling
   pathways, including RhoA (Ras homolog family member A) and its
   downstream effector ROCK (Rho‐associated protein kinase), which promote
   actin stress fibers (SFs) contraction, thereby disrupting VE‐cadherin
   and resulting in decreased stability of the endothelial barrier.^[
   [59]^16 ^] The biosafety of electro‐stimulation is critical for the
   endothelial barrier due to the long‐term and repeated administration of
   PTH, requiring the insult to be sufficiently mild to prevent cell
   necrosis. Electroporation usually causes extensive and irreversible
   disruption to the cell cytoskeleton.^[ [60]^17 ^] Although
   microcurrents have been reported to reversibly increase endothelial
   permeability, these results were based on mathematical simulations, and
   the underlying mechanisms remain unclear.^[ [61]^18 ^]

   Transdermal delivery is superior to conventional SCI in terms of
   efficiency, convenience, and patient preference for osteoporotic
   osteoanabolic therapies.^[ [62]^19 ^] Scholars have attempted to
   deliver PTH transdermally using microcurrents or high‐voltage
   electroporation on the skin surface; however, due to the compact
   packing of the stratum corneum, neither increased drug dosage nor
   electric field intensity has achieved satisfactory results.^[ [63]^20 ,
   [64]^21 ^] Physically breaking the barrier of the stratum corneum by
   creating microchannels in the epidermis is fundamental for ensuring
   effective transdermal drug delivery. Since most drug molecules are
   charged, an external electric field is typically employed to control
   the direction and speed of drug flow. Consequently, the use of
   microneedle (MN) arrays in combination with an external electric field
   has become a widely adopted strategy in the development of
   drug‐delivery devices.^[ [65]^22 ^] Numerous microcurrent‐based MN
   systems (such as iontophoresis) have been proposed for transdermal drug
   delivery, including insulin and even macromolecular proteins.^[ [66]^23
   , [67]^24 ^] Yuan Yang et al. incorporated conductive materials with a
   piezoelectric nanogenerator into microneedles, achieving self‐powered
   and controlled transdermal administration of dexamethasone.^[ [68]^25
   ^] Furthermore, wearable devices based on MN technology have recently
   enabled in situ transdermal detection and delivery of methotrexate
   using iontophoresis.^[ [69]^26 ^] However, previous reports have
   primarily focused on optimizing the electro‐driving force of
   microcurrents to improve the efficiency and convenience of transdermal
   delivery, while neglecting the potential effects and safety of
   microcurrents on subcutaneous blood vessels.

   Herein, we present an integrated microcurrent delivery system (MDS) for
   PTH transdermal delivery. The MDS demonstrates excellent
   biocompatibility and high efficiency for PTH delivery, ultimately
   enhancing the osteoanabolic effect of PTH by reducing PTH exposure time
   compared to SCI. Considering cost‐effectiveness and flexibility, the
   through‐microchannels were constructed using high‐precision 3D printing
   technology in multimicrochannel microneedle arrays (MMAs).^[ [70]^27 ^]
   The syringe positive electrode of MDS pushes the drug solution into the
   subcutaneous interstitial fluid, forming a complete circuit between the
   positive electrode, the drug solution, the subcutaneous interstitial
   fluid, and the negative electrode, which drives positively charged drug
   molecules, such as PTH, to move from the positive electrode toward the
   negative electrode (Figure [71]S1, Supporting Information). The
   non‐conductive property of the MMAs concentrates the electric field for
   dual functions: 1) accelerating the movement of charged PTH in
   subcutaneous interstitial fluid; 2) concentrating low and safe
   microcurrents that regulate the cytoskeleton and VE‐cadherin via the
   Ca^2+/cAMP pathway, causing a reversible increase in endothelial
   permeability and promoting PTH absorption. To prove the clinical value
   of the MDS for PTH delivery, we first confirmed that it improves
   delivery efficiency and shortens exposure time compared to SCI both in
   vitro and in vivo. Furthermore, using OP rat models, we demonstrated
   that PTH delivered by the MDS significantly enhanced the osteoanabolic
   effect by inhibiting osteoclast differentiation compared to SCI. Beyond
   PTH, the proposed MDS shows great promise for the high‐efficiency
   transdermal delivery of macromolecular drugs (Figure [72] 1 ).

Figure 1.

   Figure 1
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   Schematic diagram of MDS improving the osteoanabolic effect of PTH
   compared to subcutaneous injection and the Ca^2+‐dependent cAMP
   signaling mechanism.

2. Results and Discussion

2.1. Design and Fabrication of the Microcurrent Delivery System (MDS)

   MDS is designed to integrate microcurrent for subcutaneous drug
   delivery and consists of MMAs, a syringe positive electrode, counter
   electrode microneedles (MNs) as the negative electrode, and an
   injection cap (Figure [74] 2A). Since PTH is clinically administered
   via a syringe, a silver layer is sputtered onto the flattened rubber
   end of the syringe pusher to serve as the positive electrode, which is
   then connected to the external power source through a silver wire. The
   counter electrode MNs, functioning as the negative electrode, are
   combined with the positive electrode through a matching injection cap
   (Figure [75]2B–F). The counter‐electrode MNs have a base diameter of
   approximately 40 µm, a length of approximately 100 µm, and a spacing of
   approximately 110 µm between adjacent MNs. The counter‐electrode MNs
   are then sputtered with a ≈100 nm Au layer to ensure excellent
   biocompatibility, as validated in our previous studies.^[ [76]^28 ^]

Figure 2.

   Figure 2
   [77]Open in a new tab

   Design and characterization of the MDS. A) Components of the MDS. The
   counter electrode MNs and the pusher head of the syringe act as the
   negative and the positive electrode, respectively. The negative
   electrode, the positive electrode and the MMAs are loaded into the
   injection cap together to assemble the MDS. B,C) Optical micrographs
   showing the top view (B) and the bottom view (C) of the injection cap.
   The red arrows and the blue circle represent the rectangular groove for
   loading counter electrode MNs and the circular groove for loading MMAs,
   respectively. The red triangle represents the drug reservoir. Scale
   bar, 1000 µm. D,E) The silver layer and the silver wire (yellow arrows)
   at the end of the syringe pusher as the positive electrode, and the
   head of syringe can be fitted into the injection cap. F) Optical image
   of the counter electrode MNs. Scale bar, 100 µm. G) Schematic diagram
   of the 3D‐print process of the MMAs (top), and the layer‐by‐layer
   structure of the MMAs shows the internal microchannels through the base
   and the inside of MNs (bottom). H,I) Representative side view and top
   view of MMAs. Scale bar, 100 µm. J) Representative image of a single
   MN. Scale bar, 100 µm. K) Mechanical curve of MMAs with microchannels
   of 100 µm diameter under compressive force. L) Rhodamine B staining of
   MMAs penetration wounds in porcine skin. Scale bar, 200 µm.

   MMAs were fabricated using high‐resolution 3D printing technology with
   biocompatible photosensitive resin (Figure [78]2G; Figure [79]S2,
   Supporting Information). The high precision and flexibility of this
   technology allow for customization of MN size, array density, and
   microchannel diameter to suit the delivery requirements of different
   drugs. The MMAs manufactured in this study have a base diameter of
   8 mm, with 4 × 4 MNs positioned at the center of the base, spaced
   800 µm from center to center. Each pyramidal MN has a base length of
   600 µm and a height of 850 µm, with an average edge length of the tip
   measuring 18.2 ± 2.4 µm (Figure [80]S3, Supporting Information). Each
   of the four sides of the pyramidal MN features a through‐microchannel
   with a 100 µm diameter for drug transport, as visualized by SEM
   micrographs (Figure [81]2H–J). The square base of the pyramidal MN has
   been shown to withstand higher axial and transverse forces compared to
   a circular base.^[ [82]^29 ^] The MN tip was designed as a solid
   structure with microchannels open on the sides to ensure the mechanical
   stability of the MMAs for penetrating skin tissue and reducing reverse
   diffusion.^[ [83]^30 ^]

   The MMAs contain multiple through‐microchannels to ensure the high
   delivery efficiency of the MDS. Since the MMAs are made of
   non‐conductive resin, when the MMAs and the counter electrode MNs
   penetrate the skin and contact the interstitial fluid, a complete
   circuit is established between the positive electrode, the drug
   solution, the subcutaneous interstitial fluid, and the negative
   electrode. This configuration accelerates the movement of charged
   molecules (e.g., positively charged PTH).

2.2. Determination of the Optimum Microcurrent and the Diameter of
Microchannels

   Since PTH requires daily delivery, the biocompatibility of microcurrent
   is particularly important for the MDS. MDS combined with different
   microcurrent densities was used to explore their effects on the
   biocompatibility of human umbilical vein endothelial cells (HUVECs) in
   vitro. The live/dead staining results showed that a microcurrent
   density of 1 mA cm^−2 for 5 min exhibited excellent biocompatibility,
   while a significant percentage of dead cells appeared at 2 mA cm^−2
   (Figure [84]S4A,B, Supporting Information). Further assessment using
   the CCK‐8 assay revealed that a microcurrent density of 2 mA cm^−2 for
   5 min significantly inhibited cell proliferation, whereas a
   microcurrent density of 1 mA cm^− ^2 for 5 min demonstrated good
   proliferation safety at 1 and 3 days (Figure [85]S4C, Supporting
   Information). Given that HUVEC proliferation also declined with longer
   stimulation times (>7 min) at 1 mA cm^−2 (Figure [86]S4D, Supporting
   Information), we employed 7‐min stimulation periods in the remaining
   experiments to achieve optimal delivery effects while maximizing cell
   viability.

   A simulation model of the MDS was established to explore the optimal
   diameter of microchannels. The simulated results demonstrated that the
   microchannel diameter was related to the applied voltage, the flow rate
   of the drug, and the mechanical stability under a microcurrent of
   1 mA cm^−2 (Figure [87]S5A–C, Supporting Information). The
   diameter‐related simulation curves indicated that diameters of 100,
   140, or 180 µm might be optimal for the microchannels of MMAs
   (Figure [88]S5D, Supporting Information). However, the mechanical
   properties, as a fundamental factor for penetrating skin tissue, need
   further verification.^[ [89]^31 ^] The vertical deformations of MMAs
   with microchannel diameters of 100, 140, and 180 µm were observed under
   a compressive force increasing from 0 to 20 N (Figure [90]S6A–I,
   Supporting Information). The tips of the MMAs with a microchannel
   diameter of 140 µm were severely deformed, and those with a diameter of
   180 µm experienced significant collapse. Therefore, MMAs with a
   microchannel diameter of 100 µm were selected for subsequent
   verification. The force–time curve demonstrated that MMAs with a
   microchannel diameter of 100 µm did not reach the fracture point when
   the compressive force was increased from 0 to 40 N (Figure [91]2K),
   indicating high mechanical properties that meet clinical needs.^[
   [92]^32 ^] According to previous reports, the force required to insert
   into the skin ranges from 0.08 to 3.04 N,^[ [93]^33 ^] so our MMAs are
   fully capable of puncturing the skin. Pig skin was used to verify the
   penetration ability of the MMAs, and the needle‐shaped outline stained
   with rhodamine B indicated successful skin piercing (Figure [94]2L).

2.3. MDS Reversibly Increased Endothelial Permeability In Vitro

   HUVEC monolayers were established to explore the effect of MDS with
   different microcurrent densities (7 min) on endothelial permeability.
   The results of the transendothelial electrical resistance (TEER)
   indicated that the disruption of monolayer integrity was positively
   correlated with microcurrent density (Figure [95] 3A). Conversely
   mirroring the transient changes in TEER values, the endothelial
   permeability of HUVEC monolayers to fluorescein isothiocyanate‐labeled
   dextran (FITC‐dextran, 5 kDa, PTH analog) increased with rising
   microcurrent densities (Figure [96]3B). To further understand the
   dynamic effect on endothelial permeability triggered by the optimal
   delivery parameters for maximizing cell viability (7 min at
   1 mA cm^−2), it was found that TEER values decreased first following
   microcurrent stimulation, then increased spontaneously and finally
   returned to baseline. The transient changes in endothelial permeability
   exhibited an inverse relationship with the TEER values
   (Figure [97]3C,D), indicating a rapid and spontaneous recovery of
   endothelial permeability.

Figure 3.

   Figure 3
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   The endothelial permeability increased reversibly triggered by MDS in
   vitro. A,B) The transendothelial electrical resistance (TEER) and the
   permeability to 5 kDa FITC–dextran of HUVEC monolayers were measured
   following MDS exposure with different microcurrent densities. The data
   are presented as mean ± SD (n = 9). P values were analyzed by one‐way
   ANOVA. C,D) The TEER and the permeability to 5 kDa FITC–dextran of
   HUVEC monolayers as function of post‐exposure incubation time, for both
   control and MDS stimulated (7 min at 1 mA cm^−2). The data are
   presented as mean ± SD (n = 9). P values were analyzed by unpaired
   Student's t‐test. E–I) Representative confocal microscopy images of
   HUVEC monolayers 0.5 h after MDS stimulation under varying microcurrent
   densities for 7 min. Scale bar, 50 µm. The middle column showing a
   magnification of the region indicated in the VE‐cadherin channel. The
   white arrows point to the cytoplasm with reduced SFs. The percentage of
   intracellular actin (F), the relative VE‐cadherin enriched in the
   junctional regions (G), the cell size (H) measured by the area enclosed
   by VE‐cadherin, and the linearity index (I) defined as the ratio of the
   actual junction length to the linear junction length of VE‐cadherin
   concerning that of untreated cells (Control). The data are presented as
   mean ± SD (n = 30). P values were analyzed by one‐way ANOVA. J–N)
   Representative confocal microscopy images of HUVEC monolayers as
   function of incubation time after MDS stimulation at 1 mA cm^−2 for
   7 min. Scale bar, 50 µm. The middle column showing a magnification of
   the region indicated in the VE‐cadherin channel. The white arrows point
   to the cytoplasm with reduced SFs. Scale bar, 50 µm. The intracellular
   actin (K), the VE‐cadherin enrichment (L), the cell size (M), and the
   linearity index (N) are described above. The data are presented as mean
   ± SD (n = 30). P values were analyzed by one‐way ANOVA. *p < 0.05, **p
   < 0.01, and ***p < 0.001.

   RNA sequencing was applied to reveal the mechanism of spontaneous
   recovery of endothelial permeability in MDS‐stimulated (7 min at
   1 mA cm^−2) HUVEC monolayers. There were 618 upregulated genes and 279
   downregulated genes following MDS stimulation (Figure [99]S7A,B,
   Supporting Information). The downregulated genes were subjected to
   Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway enrichment
   analysis, which showed that the regulation of the actin cytoskeleton
   was significantly enriched, with the highest gene ratio and the most
   significant p‐value (Figure [100]S7C,D, Supporting Information,
   Table [101]S1). The actin cytoskeleton and VE‐cadherin have been
   reported to be involved in the regulation of endothelial permeability
   induced by electrical stimulation.^[ [102]^34 , [103]^35 ^] We observed
   noticeable differences in the actin cytoskeleton 0.5 h after exposure
   to the MDS with different power settings (Figure [104]3E).
   Specifically, differences in actin fiber orientation and associated
   VE‐cadherin were noted: at lower microcurrent densities (<1 mA cm^−2)
   and in control cells, the intracellular space was dominated by stressed
   fibers (SFs) and serrated VE‐cadherin (SVE), while SFs and SVE were
   increasingly replaced by circumferential bundles (CBs) and smooth
   linear VE‐cadherin (LVE) following exposure at 1 mA cm^−2. However,
   excessive contraction of SFs and destruction of VE‐cadherin were
   observed at 2 mA cm^−2 was confirmed by immunofluorescence analysis of
   intracellular actin and junctional regions where VE‐cadherin and CBs
   are localized (Figure [105]3F,G). We also observed a significant
   increase in cell size and a corresponding decrease in the linearity
   index (the junction length/the distance between vertices) at 1 mA cm^−2
   (Figure [106]3H,I). An increased linearity index is typically
   associated with a decrease in the intercellular gap due to the
   tightening of the endothelial barrier.^[ [107]^36 ^] At 2 mA cm^−2,
   VE‐cadherin disappeared with the contraction of SFs, similar to the
   stressed cell state seen in pathological conditions such as trauma and
   inflammation.^[ [108]^37 , [109]^38 ^]

   The dynamics of actin and VE‐cadherin reconstruction triggered by MDS
   exposure (1 mA cm^−2) were further explored and can be summarized into
   three phases: 1) pre‐stimulated cells (control): the cytoskeleton
   predominantly exists in the form of SFs and SVE; 2) electro‐stimulated
   cells: within 0.1 h post‐exposure, SVE and CBs were almost completely
   disintegrated, with an increase in intracellular actin leading to cell
   contraction, as evidenced by smaller cell size and reduced linearity;
   3) electro‐transformed cells: with cellular relaxation, SVE patterns
   gradually transform into stable LVE at 0.5 h post‐exposure and
   intracellular SFs are progressively replaced by CBs, as indicated by
   the loss of fluorescence of intracellular actin. These changes are also
   indirectly observed through increasing cell sizes and the linearity
   index. After 4 h, SVE reappeared as the cells progressed from a
   challenged state toward recovery. The intracellular SFs and cell
   morphology relaxed back to their original pre‐stimulated state 24 h
   post‐exposure (Figure [110]3J–N). Long Lin et al. proposed a
   multimicrochannel microneedle microporation platform that efficiently
   delivers chemotherapy drugs locally into tumor cells through
   electroporation of the cell membrane, significantly enhancing the
   anti‐tumor effect. However, this method is not suitable for regulating
   vascular endothelial cells to promote the absorption and circulation of
   PTH. Furthermore, the long‐term use of electroporation may pose safety
   risks to endothelial cells. In contrast, our results indicate that the
   reversible increase in endothelial permeability induced by microcurrent
   might offer a safer alternative.^[ [111]^39 ^]

2.4. Ca^2+‐Dependent cAMP Mediates Reversible Increase of Endothelial
Permeability via ROCK Activation and Inhibition

   Figure [112]3 shows the spontaneous cytoskeleton remodeling following
   MDS stimulation. To date, no studies have investigated the dynamic
   rebalancing of actin and VE‐cadherin triggered by electro‐stimulation.
   Pulsed electric fields, in the form of electroporation, typically cause
   extensive and irreversible destruction of the cytoskeleton, leading to
   decreased cell viability,^[ [113]^40 ^] which is unsuitable for
   long‐term drug delivery. Additionally, previous studies on
   microcurrent‐assisted MNs have primarily focused on the driving force
   provided by microcurrent, often overlooking its effect on vascular
   permeability.^[ [114]^41 ^] To elucidate the spontaneous cytoskeleton
   remodeling of HUVEC monolayers triggered by MDS, we further
   investigated the underlying signaling mechanisms.

   VE‐cadherin consists of a long extracellular domain that mediates
   calcium‐dependent cellular adhesion.^[ [115]^42 ^] Previous studies
   have suggested that the interaction between electrical stimulation and
   the cytoskeleton mainly involves transient membrane permeation and
   piezo channel activation, which result in elevated intracellular Ca^2+
   levels.^[ [116]^17 ^] Therefore, we focused on changes in intracellular
   Ca^2+ levels caused by MDS. Intracellular Ca^2+ levels increased
   immediately after MDS stimulation, followed by a rapid return to
   baseline within 0.5 h (Figure [117] 4A). The influx of Ca^2+ is known
   to activate the RhoA/ROCK pathway, leading to the formation of (SFs)
   and actomyosin contraction,^[ [118]^43 ^] This process causes
   reversible invagination of VE‐cadherin into the cell,^[ [119]^44 ^]
   explaining the increase in intracellular SFs and the decrease in
   VE‐cadherin fluorescence (Figure [120]3J) and insoluble VE‐cadherin
   protein levels (Figure [121]4B,C) during the electro‐stimulated phase
   (0–0.1 h). RhoA activity and ROCK protein levels increased
   significantly during the electro‐stimulated phase (Figure [122]4D–F).
   Additionally, piezo channel inhibitor (ruthenium red), intracellular
   Ca^2+ chelator (BAPTA‐AM), and ROCK inhibitor (Y27632) all blocked the
   increase in intracellular SFs and the disappearance of VE‐cadherin
   fluorescence during the electro‐stimulated phase (Figure [123]S8A,
   Supporting Information), further proving the key role of the
   Ca^2+‐triggered RhoA/ROCK pathway in endothelial barrier leakage during
   the electro‐stimulated phase.

Figure 4.

   Figure 4
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   Ca^2+‐dependent cAMP mediates reversible increase of endothelial
   permeability via ROCK activation and inhibition. A) Intracellular Ca^2+
   in HUVEC monolayers as function of post‐stimulus time for both control
   (untreated) and MDS stimulation (1 mA cm^−2 for 7 min) cells. The data
   are presented as mean ± SD (n = 3). P values were analyzed by unpaired
   Student's t‐test. B–E) Western blot analyses of insoluble VE‐cadherin,
   soluble VE‐cadherin (total), ROCK and GAPDH protein samples in HUVEC
   monolayers triggered by MDS (1 mA cm^−2 for 7 min) at different
   post‐stimulus time (B,D). Quantitative analyses of insoluble
   VE‐cadherin (C) and ROCK protein expression (E) relative to soluble
   VE‐cadherin and GAPDH, respectively. Data are presented as mean ± SD (n
   = 6). P values were analyzed by one‐way ANOVA. F) Relative RhoA
   activity in HUVEC monolayers triggered by MDS (1 mA cm^−2 for 7 min) at
   different post‐stimulus times with respect to that of control cells
   (untreated). Data are presented as mean ± SD (n = 6). P values were
   analyzed by one‐way ANOVA. G) Intracellular cAMP levels in HUVEC
   monolayers as function of post‐stimulus time for both control
   (untreated) and MDS stimulation (1 mA cm^−2 for 7 min) cells. The data
   are presented as mean ± SD (n = 3). P values were analyzed by unpaired
   Student's t‐test. H) Relative PKA activity in HUVEC monolayers
   triggered by MDS (7 min at 1 mA cm^−2) at different post‐stimulus times
   with respect to that of control cells (untreated). Data are presented
   as mean ± SD (n = 6). P values were analyzed by one‐way ANOVA. I, J)
   The effect of MDS stimulation (7 min at 1 mA cm^−2) and extracellular
   Ca^2+ on intracellular cAMP levels in HUVEC monolayers with and without
   intracellular Ca^2+, respectively. Intracellular Ca^2+ were chelated by
   BAPTA‐AM. The data are presented as mean ± SD (n = 3). K) Mechanism
   diagram of spontaneous recovery of endothelial permeability. *p < 0.05
   and ***p < 0.001.

   During the electro‐transformed phase (0.5–2 h), the fluorescence signal
   and protein level of VE‐cadherin were restored, leading to a decrease
   in endothelial permeability, indicative of endothelial barrier
   stabilization. It is well established that cyclic adenosine
   monophosphate (cAMP) stabilizes endothelial barrier functions by
   regulating VE‐cadherin.^[ [125]^45 ^] We found that intracellular cAMP
   levels increased as Ca^2⁺ levels decreased during the
   electro‐stimulated phase and remained high during the
   electro‐transformed phase (Figure [126]4G). The barrier‐stabilizing
   effect of cAMP signaling is known to be mediated either by protein
   kinase A (PKA) or by exchange protein 1 activated by cAMP (EPAC1).^[
   [127]^46 ^] We observed a significant increase in PKA activity in
   electro‐transformed cells (Figure [128]4H). Moreover, when cells were
   treated with PKA inhibitor (H‐89) or EPAC1 inhibitor (ESI‐09) before
   MDS stimulation, the linearization of VE‐cadherin and the
   transformation of intracellular SFs to CBs were blocked by the former
   0.5 h post‐stimulation (Figure [129]S8B, Supporting Information). cAMP
   signaling has been reported to activate PKA, which then inhibits actin
   contraction caused by RhoA/ROCK pathway activation.^[ [130]^47 ^] Our
   results also demonstrated a significant down‐regulation of RhoA
   activity and ROCK protein levels during the electro‐transformed phase
   (Figure [131]4E,F).

   Finally, we explored the interaction between Ca^2+ and the second
   messenger cAMP signaling. We noted that the increase of intracellular
   cAMP after MDS stimulation was fairly insignificant in the absence of
   extracellular Ca^2+ (Figure [132]4I). Furthermore, BAPTA‐AM (BP), an
   intracellular Ca^2+ chelator, was used to remove intracellular Ca^2+,
   and found that intracellular cAMP was significantly increased only in
   the presence of extracellular Ca^2+ (Figure [133]4J), alluding to the
   strong dependence of the increase of cAMP on the influx of
   extracellular Ca^2+ after MDS stimulation. Collectively,
   Ca^2+‐dependent cAMP signaling pathway plays a pivotal role in the
   microcurrent‐induced regulation of endothelial permeability.
   Figure [134]4K summarizes that the influx of Ca^2+ activates the
   RhoA/ROCK axis during the electro‐stimulated phase, leading to
   endothelial barrier leakage, and that subsequent Ca^2+‐dependent cAMP
   signaling mediates PKA activation to stabilize the endothelial barrier
   by inhibiting the RhoA/ROCK axis during the electro‐transformed phase.

2.5. MDS‐Mediated PTH Efficient Delivery and Osteogenesis Enhanced In Vivo

   To test the effect of microcurrent in MDS on PTH delivery efficiency,
   the PTH‐preloaded MDS was inserted into a PBS solution. The preloaded
   PTH was delivered into the PBS solution through microchannels, driven
   by both the electric field and free diffusion (see Delivery Efficiency
   Test Section). The results showed that the microcurrent density was
   positively correlated with PTH delivery efficiency, and the delivery
   efficiency at 1 mA cm^−2 was significantly higher than that achieved
   through free diffusion (control) (Figure [135] 5A). Previous reports
   have also indicated that the driving force provided by microcurrent
   could increase drug permeation across the skin by 5–10 times compared
   to MNs alone.^[ [136]^48 ^] The delivery efficiency of MDS was further
   verified in vivo by examining the plasma concentration profiles of PTH
   under different administration methods. Compared to the MMAs‐PTH group
   (PTH delivered by MDS without microcurrent) and the SCI‐PTH group, MDS
   delivery (1 mA cm^−2 for 7 min) significantly reduced the plasma C[max]
   time of PTH (Figure [137]5B). Previous studies proposed that the
   obstruction of subcutaneous capillary walls reduces the absorption rate
   of PTH post‐SCI and causes approximately 20% subcutaneous residue,
   delaying the time to plasma C[max] to 30 min followed by elimination in
   4 h,^[ [138]^9 ^] which is consistent with our results of SCI‐PTH
   group. By providing an electro‐driving force and reversibly increasing
   vascular permeability, MDS efficiently accelerates PTH entry into the
   blood, shortening the PTH exposure time to 2 h (Figure [139]5B).

Figure 5.

   Figure 5
   [140]Open in a new tab

   MDS‐mediated PTH efficient delivery and osteogenic enhancement in vivo.
   A) PTH delivery efficiency of MDS at different microcurrent densities
   in vitro. Data are presented as mean ± SD (n = 9). P values were
   analyzed by one‐way ANOVA. B) Plasma concentration‐time profiles of PTH
   using three different delivery methods. Data are presented as mean ± SD
   (n = 3). C) 3D reconstruction of distal femurs from sham‐operated and
   OVX rats treated with PBS or PTH by different methods for 4 weeks. D–H)
   Quantitative results of micro‐CT: (D) bone mineral density, (E) bone
   volume fraction, (F) trabecular number, (G) trabecular thickness, and
   (H) trabecular separation. Data are presented as mean ± SD (n = 5). P
   values were analyzed by one‐way ANOVA. I) Hematoxylin and eosin (H&E)
   staining of distal femurs. Scale bar, 200 µm. J) Masson staining of
   distal femurs from sham‐operated and OVX rats. Blue marks new bone
   collagen fibers, while red marks mature bone. Scale bar, 200 µm.
   Enlarged scale bar, 50 µm. K,L) Plasma concentration of PINP and CTX‐I.
   Data are presented as mean ± SD (n = 5). P values were analyzed by
   one‐way ANOVA. M,N) Immunohistochemistry analyses for osteoblastic
   makers Col1 (M) and OCN (N). Col^+ and OCN^+ osteoblasts on the bone
   surface are marked by red arrows. Scale bar, 200 µm. Enlarged scale
   bar, 50 µm. O,P) The number of Col^+ and OCN^+ osteoblasts per bone
   surface. Data are presented as mean ± SD (n = 5). P values were
   analyzed by one‐way ANOVA. Q) Immunohistochemistry analyses for
   osteoclastic makers Ctsk. Ctsk^+ osteoclasts on the bone surface are
   marked by red arrows. Scale bar, 200 µm. Enlarged scale bar, 50 µm. R)
   Trap staining for assessment of osteoclast activity. Red arrows
   indicate mature osteoclasts. Scale bar, 200 µm. Enlarged scale bar,
   50 µm. S,T) The number of Ctsk^+ and Trap^+ osteoclasts per bone
   surface. Data are presented as mean ± SD (n = 5). P values were
   analyzed by one‐way ANOVA. *p < 0.05, **p < 0.01, and ***p < 0.001.

   To evaluate the osteoanabolic effect of PTH delivered by MDS,
   ovariectomized (OVX) OP rat models were established and subjected to
   different treatments: 1) SCI‐PBS (sham and OVX); 2) MDS‐PBS
   (MDS‐delivered PBS in OVX); 3) SCI‐PTH (PTH delivered via SCI in OVX);
   4) MMAs‐PTH (MDS‐delivered PTH without microcurrent in OVX); 5) MDS‐PTH
   (MDS‐delivered PTH in OVX). PTH was administered at a concentration of
   40 µg k^−1g once daily for one month. Micro‐CT of the distal femurs
   revealed that the OVX and MDS‐PBS groups experienced severe bone loss
   and destruction of bone microarchitecture compared to the sham group.
   The PTH‐treated groups showed significant improvement in bone mass and
   bone microarchitecture compared to the OVX group, with the MDS‐PTH
   group demonstrating an even greater increase in BMD and bone volume
   fraction (Figure [141]5C–H). Numerous studies have shown that direct
   current (DC) could promote osteogenic differentiation of mesenchymal
   stem cells and increase bone mass.^[ [142]^49 , [143]^50 ^] However, in
   these studies, the electrodes were directly applied to the bone, and
   the required current and voltage levels were significantly higher than
   the 1 mA cm^−2 used in our MDS. Furthermore, there was no significant
   difference in bone mass between the MDS‐PBS group and the OVX group,
   indirectly suggesting that localized transcutaneous microcurrent (1
   mA cm^−2) has no substantial effect on bone‐related cells
   (Figure [144]5C–H). Hematoxylin and eosin (H&E) staining and Masson's
   staining also exhibited marked trabecular microarchitecture
   deterioration in the OVX group, with new bone formation (indicated in
   blue) significantly promoted in the MDS‐PTH group compared to the
   SCI‐PTH group (Figure [145]5I,J). According to these findings, MDS
   enhanced osteogenic efficacy by reducing the PTH exposure time compared
   to conventional SCI.

   The effect of MDS on reducing PTH exposure time and its impact on bone
   homeostasis was further analyzed. Serum enzyme‐linked immunosorbent
   assay (ELISA) results indicated that the bone formation marker PINP
   increased notably in both the SCI‐PTH and MDS‐PTH groups compared to
   the OVX group, but there was no significant difference between the
   SCI‐PTH and MDS‐PTH groups. Although the bone resorption marker CTX‐I
   was significantly increased in each group compared to the sham group,
   it was significantly lower in the MDS‐PTH group than in the SCI‐PTH
   group, suggesting that MDS‐PTH enhanced the PTH osteoanabolic effect by
   inhibiting osteoclast metabolism (Figure [146]5K,L). Numerous studies
   have investigated the effect of PTH exposure time on bone turnover
   markers, concluding that prolonged exposure time via intravenous
   infusion leads to a significant increase in bone resorption markers.^[
   [147]^7 , [148]^8 ^] Sections of distal femurs were further subjected
   to immunohistochemistry (IHC) staining for osteoblastic (Col/OCN) and
   osteoclastic markers (Ctsk/Trap). Compared with the OVX group, there
   were more Col^+ and OCN^+ osteoblasts per bone surface in the SCI‐PTH
   and MDS‐PTH groups, but no significant difference between the two
   (Figure [149]5M–P). However, there were dramatically fewer Ctsk^+ and
   Trap^+ osteoclasts per bone surface in the MDS‐PTH group compared to
   the SCI‐PTH group (Figure [150]5Q–T). Choi et al. found that a 30‐min
   PTH exposure was sufficient to achieve maximal induction of the
   osteogenic gene Nurr1, while exposure for more than 2 h resulted in a
   sharp increase in the osteoclastic gene Rankl, peaking at 4 h at the
   transcriptional level.^[ [151]^6 ^] To summarize, MDS reduced the PTH
   exposure time, attenuating osteoclast differentiation while ensuring
   osteoblast differentiation, ultimately enhancing the PTH osteoanabolic
   effect.

2.6. MDS for Safe Drug Delivery In Vivo

   For the long‐term daily administration of PTH, it is necessary to
   validate the biocompatibility of MDS in the skin and major organs.
   Histopathological analysis of skin biopsies showed negligible
   inflammation within 4 h after treatment with MDS, whether with or
   without microcurrent (MMAs), which was alleviated within 24 h
   (Figure [152] 6A). After one month of PTH treatment, there was no
   significant difference in the appearance or weight of major organs in
   the MDS‐PTH group compared to the SCI‐PBS and SCI‐PTH groups
   (Figure [153]6B,C). Finally, histopathological analysis showed no
   structural disruption in the visceral organs of the MDS‐PTH group
   (Figure [154]6D). These results indicate the good biosafety of MDS for
   long‐term delivery of PTH.

Figure 6.

   Figure 6
   [155]Open in a new tab

   MDS for safe PTH delivery in vivo. A) Representative images of skin
   from the back of hairless rats after the treatment of MMAs (MDS without
   microcurrent) or MDS (1 mA cm^−2) for 7 min. Compared to the untreated
   sites (control), the H & E staining showed more scattered immune cells
   (black arrows) at 4 h after treatments, which alleviated within 24 h.
   Scale bar, 100 µm. B) Representative photographs of the heart, lung,
   liver, kidney, and spleen at the end of the experiment. C) Statistical
   analysis of the final weight of organs. Data are presented as mean ± SD
   (n = 3). P values were analyzed by one‐way ANOVA. D) H&E staining of
   the heart, lung, liver, kidney, and spleen of rats at the end of the
   experiment. Scale bar, 100 µm.

3. Conclusion

   In this work, we developed an integrated and functional MDS to
   facilitate PTH delivery for an enhanced osteoanabolic effect. First, we
   elucidated the effect and mechanism of MDS in reversibly increasing
   vascular permeability in vitro. We then verified the high efficiency of
   MDS for PTH delivery both in vitro and in vivo, which resulted in a
   shorter PTH exposure time compared to SCI. Finally, OP rat models
   demonstrated that the enhanced osteoanabolic effect, due to the
   shortened PTH exposure time achieved by MDS, was accomplished by
   inhibiting osteoclast differentiation. The minimal invasiveness of MMAs
   makes them advantageous for preventing vascular and nerve damage,
   penetrating skin tissue, and delivering drugs in situ, making them
   applicable in a wide range of scenarios, including transdermal drug
   delivery. The built‐in microchannels of MMAs provide a concentrated
   electric field for MDS, which not only accelerates the movement of
   subcutaneous drugs but also facilitates circulatory absorption by
   increasing vascular permeability reversibly. As a general localized
   subcutaneous drug delivery platform, MDS may serve as a more versatile
   tool, broadly applicable to macromolecular drugs in addition to PTH.
   Additionally, MDS allows for self‐administration due to its safety and
   user‐friendliness, offering high practicality.

4. Experimental Section

Fabrication and Characterization of MDS

   The MMAs were printed by a high‐resolution 3D printer (Miro P213,
   Xingsheng Technology Inc., Jiangmen, China) with an accuracy of 3 µm,
   using XS‐BIO resin (35% polyester acrylate, 25% glycidyl methacrylate,
   25% poly (ethylene glycol) dimethacrylate and 15% Polyethylene glycol;
   Xingsheng Technology Inc., Jiangmen, China) and was cured under 405 nm
   UV light. Software SolidWorks 2022 was applied to design the 3D model
   of the MMAs, and then the model was sliced into a series of 2D images
   (5 µm per slice). Figure [156]S2 (Supporting Information) shows
   representative printing layers and corresponding projecting images of
   MMAs. Similarly, the injection cap matching the size of MMAs was
   designed and printed, with the diameter of the circular groove for
   loading MMAs being 8 mm, and the diameter of the drug reservoir being
   5 mm.

   The positive electrode was constructed using the pusher head of the
   syringe: flatten the rubber pusher head with a blade, and then sputter
   a layer of silver onto the flat surface. The silver wire connecting the
   power supply was fixed by the silver paste on the sputtered silver
   electrode. The tip of the syringe can be inserted into the injection
   cap for application.

   As the negative electrode, counter electrode MNs were fabricated by
   laser microetching (Inno Laser Technology Co., Ltd, Shenzhen, China)
   using a stainless steel substrate with a thickness of about 100 µm. An
   Au layer of about 100 nm thickness was coated on the surface of the
   counter electrode MNs to enhance biocompatibility using magnetron
   sputtering (Kangdesheng Mechanical & Electrical Technology Co., Ltd.,
   China), and the counter electrode MNs were then inserted into the
   corresponding slot of the injection cap to construct the MDS.

   Above morphologies were characterized by an optical microscope (Axio
   Zoom V16, Zeiss) and scanning electron microscope (SEM, SUPRA 60,
   Zeiss).

Mechanical Properties of MMAs

   A low‐force mechanical testing system (GCD‐203050 M, Daheng Optics,
   Beijing) was applied to test the mechanical properties of the MMAs.
   Briefly, the needle tips of MMAs pointed up and a compressive load was
   applied with increasing force up to the maximum loading force of 40 N.
   The time sensitivity of the compressive sensor probe is 0.1 s. For the
   deformation test, 20 N compressive force was maintained for 20 s in
   each group before MMAs were taken out for observation under an SEM.

Skin Penetration Test

   The needle tips of MMAs were stained with red fluorescent dye using a
   porous sponge filled with rhodamine B (Sigma Aldrich, Merck). The MMAs
   were pressed into a piece of porcine skin and removed 5 min later. The
   position on the porcine skin treated by MMAs was cut into a 2 cm slice,
   then using the fluorescence microscope (IX83, Olympus) to record the
   penetration effect of the cross‐section.

Cell Culture

   HUVECs were cultured in endothelial cell medium (ECM, ScienCell)
   containing 5% FBS, 1% endothelial cell growth factor supplement, and 1%
   antibiotic and were maintained at 37 °C in a humidified atmosphere
   containing 5% CO[2]. After 80–90% confluency, cells can be detached by
   0.05% trypsin–EDTA and reseeded for passages. Cells between 4 and 6
   passages were used in all experiments.

Measurement of TEER

   The integrity of endothelial barrier was assessed by the TEER across
   the HUVEC monolayer. HUVECs were seeded on polycarbonate membrane
   inserts of a 24‐well Transwell (Corning, USA) coated with 0.1% gelatin
   at a density of 3 × 10^5 cells/insert and cultured for 2 days until
   HUVEC monolayer formation. The Millicell ERS‐2 Electrical Resistance
   System (Millipore, USA) was applied to measure the TEER values. The
   normalized value was corrected for the background TEER of a filter
   without cells and expressed as Ω. cm^2 based on the corresponding
   surface area of filter.

Endothelial Permeability Assay

   The HUVEC monolayer was formed on inserts of a 24‐well transwell using
   the method described above. The endothelial permeability of the HUVEC
   monolayer was determined by the flux per unit time of FITC‐dextran
   (M.W. 5 kDa) from the upper to the lower compartments of the Transwell.
   Briefly, 20 µM FITC‐dextran was added to the upper compartment, and the
   same amount of unlabeled dextran was added to the lower compartment to
   balance the osmotic pressure. Then, the HUVEC monolayer was exposed to
   MDS according to the designed protocol. 10 µl of each sample was
   collected from both compartments at various post‐exposure incubation
   times. Fluorescence intensities of FITC‐dextran (485 ex/520 em nm) were
   assessed using a microplate reader (Thermo Scientific, Varioskan LUX,
   USA). The amount of FITC–dextran transported to the lower compartment
   can be calculated by the standard curve.

Immunofluorescence and Analysis

   HUVEC monolayers were fixed with 4% paraformaldehyde for 10 min at 4
   °C. The fixed cells were permeabilized with 0.1% Triton‐X for 5 min and
   blocked with 3% BSA in PBS for 1 h. The primary antibody
   (anti‐VE‐cadherin, 1:400, CST 2500, USA) was incubated at 4 °C
   overnight, and the secondary antibody (anti‐rabbit, 1:1000, Alexa Fluor
   647 Conjugate, CST, USA) was incubated at room temperature for 1 h.
   Actin filaments were stained with phalloidin (iFluor 488 Conjugate,
   1:1000, AAT Bioquest, USA) and the nuclei were counterstained with DAPI
   (Beyotime, 1:500, Shanghai, China). PBS was used to wash thrice between
   each step. Fluorescence images were recorded by confocal microscopy
   (FV3000, Olympus), and all images were acquired under the same
   acquisition parameters.

   Image J was used for the evaluation of immunofluorescence images.
   Briefly, The VE‐cadherin enrichment was measured by the mean
   fluorescence intensity of VE‐cadherin (red) at the intercellular
   junction of the maximum intensity projected image. The intracellular
   actin was evaluated using a masking approach, which eliminated the
   actin covered by VE‐cadherin at the intercellular junction in the
   merged image. Cell sizes (area) were defined by their VE‐cadherin
   structure. The linearity index was defined as the ratio of the actual
   junction length to the linear junction length of VE‐cadherin.

Intracellular Ca^2+

   The level of free cytoplasmic Ca^2+ in HUVEC monolayers were determined
   by Fura‐2 AM Ca^2+ fluorescence probe. Briefly, the HUVEC monolayers
   were incubated in 5 µmol L^−1 Fura‐2 AM in PBS at 37 °C in the dark for
   40 min. Then extracellular Fura‐2 AM was removed by PBS change, and the
   measurements were carried out after another 20 min incubation period.
   The fluorescence intensity at 510 nm emission wavelength with
   excitation wavelengths of 340 nm and 380 nm were measured respectively
   by a microplate reader. The ratio of fluorescence intensity at 340 nm
   to 380 nm (340/380) represents the concentration of intracellular
   Ca^2+.

Intracellular cAMP

   For cAMP assay, HUVEC monolayers were grown in 24‐well plates treated
   with 100 µM IBMX (phosphodiesterase inhibitor) to prevent the
   hydrolysis of cAMP. HUVEC monolayers were exposed to the MDS followed
   by incubation at different time points. Then cells were lysed by
   repeated freeze/thaw, and the supernatant was supplied with the cAMP
   Parameter Immunoassay Kit (KGE002B, R&D Systems, Minneapolis, USA)
   according to the manufacturer's instructions.

Western Blotting

   RIPA was used to extract proteins from lysed cells, and the proteins
   were then transferred to nitrocellulose membranes separated by SDS‐PAGE
   electrophoresis. The membranes were blocked for 1 h using 5% skim milk
   followed by being incubated with primary antibodies (anti‐VE‐cadherin,
   1:1000, CST 2500; anti‐ROCK, 1:5000, Abcam ab45171) overnight at 4 °C
   and then with appropriate secondary antibodies (anti‐rabbit IgG,
   1:2000, CST 7074) for 1 h at room temperature. GAPDH was used as the
   housekeeping gene. Protein bands were visualized using a
   multifunctional imager (Amersham Imager 600, GE Healthcare, USA). Image
   J software was used for photo analysis.

Measurement of Soluble and Insoluble VE‐Cadherin

   The cell lysate was isolated from the cytoskeleton extraction buffer^[
   [157]^51 ^] via centrifugation at 12000 × g for 10 min. The
   cytoskeleton‐associated insoluble fraction in the pellet was then
   incubated at 100 °C for 10 min. Both the cytoskeleton‐associated
   insoluble and soluble (soluble) fractions were quantified by BCA assay.
   SDS‐PAGE and western blotting were applied to isolate and analyze,
   respectively.

Inhibitor Assay

   The inhibitors were used as follows. ROCK inhibitor (Y‐27632, MCE
   HY‐10071): 15 µM; intracellular Ca^2+ chelator (BAPTA‐AM, MCE
   HY‐100545): 20 µM; piezo channel inhibitor (ruthenium red, MCE
   HY‐103311): 20 µM; Epac1 inhibitor (ESI‐09, MCE HY‐16704): 10 µM; PKA
   inhibitor (H‐89, MCE HY‐15979): 10 µM.

Elisa (PINP, CTX‐I and PKA) and Glisa (RhoA)

   Elisa kits were used to detect the expression of PINP (Huamei, Wuhan,
   China) and CTX‐I (Meilian, Shanghai, China). Plasma samples from
   different groups were obtained by the rat facial veins and then
   separated by centrifuging at 1000 g for 30 min at 4 °C. PKA
   Colorimetric Activity Kit (Arbor Assays, Ann Arbor, USA) was used to
   evaluate the activity of PKA. The HUVEC monolayers at different time
   points after MDS treatment (1 mA cm^−2 for 7 min) were lysed in the
   activated cell lysis buffer provided by the kit, and the supernatant
   was separated for subsequent detection. According to the manufacturer's
   protocols, the related reagents were added to wells of Elisa plates
   according to the instructions and incubated at 37 °C. Following the
   termination of the reaction, the optical densities of the wells were
   recorded by a microplate reader, and the relative concentrations were
   calculated.

   The RhoA Glisa kit (Cytoskeleton, Denver CO, USA) was used to evaluate
   RhoA activity according to manufacturer's recommendations. Briefly,
   cell lysates were harvested at 4 °C and then added to the wells of the
   RhoA binding plates after adjusting the protein concentration. Plates
   were incubated at 4 °C for 1 h and then washed twice with wash buffer
   at room temperature. RhoA primary antibodies (1:200) were added and
   incubated for 1 h at room temperature. Following the HRP‐related
   antibody and detection reagent were added, the results were analyzed
   using a microplate reader.

Delivery Efficiency Test

   All microchannels of the MMAs preloaded with the PTH solution using the
   syringe push rod, and then the MDS and the counter electrode MNs were
   placed into PBS solution. The power supply (HLR‐3660D, Guangdong
   Henghuiyuan Electronics Co., Ltd, China) was turned on to deliver
   different microcurrent densities for 5 min. The control group was
   treated in the same manner but no power supply. The PBS solution from
   each group was collected and their concentration of PTH was determined
   by Elisa kit (High Sensitivity Human PTH EIA, Quidel MicroVue, USA)
   according to manufacturer's recommendations. Delivery efficiency was
   defined as the concentration of PTH in PBS per unit of time.

Pharmacokinetics of PTH

   The abdominal skin of Sprague Dawley rats was shaved while the animals
   were anesthetized by pentobarbital sodium salt (30 mg k^−1g). Twelve
   rats (12 weeks old, female) were randomly divided into four groups: 1)
   SCI‐PBS: SCI with 200 µl PBS; 2) SCI‐PTH: SCI with 200 µl PTH (40
   µg k^−1g); 3) MMAs‐PTH: 200 µl PTH (40 µg k^−1g) subcutaneously
   delivered by MDS using the syringe push rod without microcurrent; 4)
   MDS‐PTH: 200 µl PTH (40 µg k^−1g) was first subcutaneously delivered by
   MDS using the syringe push rod and then 1 mA cm^−2 of microcurrent was
   applied for 7 min. At pre‐determined intervals (1, 5, 10, 15, 30, 45,
   60, 90, 120, 240, 360 min), 100 µl of blood was withdrawn from the
   jugular veins and collected in tubes containing EDTA. The plasma was
   separated from the collected blood and then stored at −80 °C until
   required for determination of PTH concentration. The PTH Elisa kit was
   used to evaluate PTH concentration (see Delivery Efficiency Test
   Section). The time to reach peak PTH plasma concentration (T[max]) was
   determined directly from the plasma concentration‐time profiles.

Animal Model Grouping and Treatment

   All rats used in this study were obtained from the Animal Experimental
   Center of Sun Yat‐sen University, and all procedures were approved by
   the Committee on the Ethics of Animal Experiment Center of Sun Yat‐sen
   University (Approval No. SYSU‐IACUC‐2024‐B0801). Thirty Sprague–Dawley
   rats (12 weeks old, female) were used to establish the postmenopausal
   OP model. All animals were anesthetized by pentobarbital sodium salt
   (30 mg k^−1g). Under sterile conditions, the bilateral ovaries were
   exposed through dorsal midline incision and removed in the ovariectomy
   group (OVX, 25 rats), whereas only two pieces of adipose tissues with
   volumes equivalent to those of the ovaries were removed in the sham
   group (5 rats). The bilateral oophorectomy was confirmed to be
   successful when pap smear screening showed no keratosis. Three months
   after surgery, rats of the OVX group were randomly divided into five
   treatment groups: 1) OVX (SCI with PBS); 2) MDS‐PBS (PBS delivered by
   MDS (1 mA cm^−2 for 7 min)); 3) SCI‐PTH (SCI with PTH (40 µg k^−1g));
   4) MMAs‐PTH: PTH (40 µg k^−1g) was subcutaneously delivered by MDS
   using the syringe push rod without microcurrent; 5) MDS‐PTH: PTH (40
   µg k^−1g) was first delivered subcutaneously by MDS using the syringe
   push rod, followed by the application of a 1 mA cm^− ^2 microcurrent
   for 7 min. The sham group was injected subcutaneously with PBS. All
   treatments were performed once a day for one month.

Micro‐CT Evaluation

   The BMD and trabecular bone parameters of rats were detected after one
   month of treatments. The anesthetized rats were subjected to micro‐CT
   examination using a SkyScan 1276 benchtop scanner (Bruker, Kontich,
   Belgium). The left femurs were analyzed, and the trabecular region of
   interest (ROI) was located 0.5 mm above the growth plate and extended
   1.5 mm upward. All left femurs were scanned at an isotropic voxel
   resolution of 20 µm with a 0.5 mm aluminum filter, 70 kV x‐ray tube
   voltage, 200 µA tube electric current, and 460 ms exposure time. The
   BMD and the trabecular bone parameters were quantitatively analyzed by
   software CTAn (Version 1.17.7.2, SkyScan). According to the ROI, the
   locational trabecular bone was reconstructed at 3D levels using the
   software CTvox (Version 3.2, Skyscan).

Histological Evaluation

   The excised skin, visceral tissue and femurs were dissected and fixed
   in 4% paraformaldehyde for 48 h at room temperature. The femurs needed
   to be decalcified for three weeks. All tissues were embedded in
   paraffin and sectioned continuously (5 mm thick). H&E and Masson
   staining were performed according to previous protocol.^[ [158]^52 ^]
   For tartrate‐resistant acid phosphatase (TRAP) staining, the TRAP
   staining kit (SLBT1113, Sigma, USA) was applied according to the
   manufacturer's instructions. Immunohistochemistry staining was
   performed according to previous protocols.^[ [159]^53 ^] The following
   primary antibodies were used: anti‐Col1 (Collagen I, Abcam ab270993,
   1:100), anti‐Ocn (Osteocalcin, Boster PB1009, 1:200), anti‐Ctsk
   (cathepsin K, Santa Cruz sc‐48353, 1:50). The slides were photographed
   using a microscope (IX83, Olympus). The number of positive cells per
   unit bone surface was quantified by Image J.

Statistical Analysis

   Statistical analyses were performed using GraphPad Prism 10 software.
   Data presented in this study are expressed as the mean ± SD. A minimum
   of three replicates were used in each process for all experiments. The
   data was analyzed using ordinary one‐way analysis of variance (ANOVA)
   or unpaired Student's t‐test. P‐values below 0.05 (*p < 0.05, **p
   < 0.01, and ***p < 0.001) were considered significant differences
   between groups.

Conflict of Interest

   The authors declare no conflict of interest.

Supporting information

   Supporting Information
   [160]SMTD-9-2401144-s001.docx^ (15.2MB, docx)

   Supplemental Table 1
   [161]SMTD-9-2401144-s002.xlsx^ (36.4KB, xlsx)

Acknowledgements