Abstract
Electrical stimulation (ES) is proposed as a therapeutic solution for
managing chronic wounds. However, its widespread clinical adoption is
limited by the requirement of additional extracorporeal devices to
power ES‐based wound dressings. In this study, a novel
sandwich‐structured photovoltaic microcurrent hydrogel dressing (PMH
dressing) is designed for treating diabetic wounds. This innovative
dressing comprises flexible organic photovoltaic (OPV) cells, a
flexible micro–electro–mechanical systems (MEMS) electrode, and a
multifunctional hydrogel serving as an electrode–tissue interface. The
PMH dressing is engineered to administer ES, mimicking the
physiological injury current occurring naturally in wounds when exposed
to light; thus, facilitating wound healing. In vitro experiments are
performed to validate the PMH dressing's exceptional biocompatibility
and robust antibacterial properties. In vivo experiments and proteomic
analysis reveal that the proposed PMH dressing significantly
accelerates the healing of infected diabetic wounds by enhancing
extracellular matrix regeneration, eliminating bacteria, regulating
inflammatory responses, and modulating vascular functions. Therefore,
the PMH dressing is a potent, versatile, and effective solution for
diabetic wound care, paving the way for advancements in wireless ES
wound dressings.
Keywords: conducting hydrogel, electrical stimulation, flexible
photovoltaic cells, proteomics, wound dressing
__________________________________________________________________
A sandwich‐structured photovoltaic microcurrent hydrogel dressing (PMH
dressing) for chronic bacteria‐infected diabetic wounds is developed.
PMH dressing generates a biomimetic current under indoor light, which
is delivered to the wound through the hydrogel interface. PMH dressing
significantly enhances the healing of infected diabetic wounds through
improved extracellular matrix regeneration, bacterial clearance, and
regulation of inflammatory responses and vascular functions.
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1. Introduction
Diabetic wounds represent a profoundly debilitating complication,
afflicting ≈25% of individuals with type II diabetes during their
lifetimes.^[ [64]^1 ^] Unfortunately, statistics suggest that a lower
limb is amputated every 30 s worldwide due to non‐healing diabetic
wounds.^[ [65]^2 ^] Despite the availability of various treatment
alternatives, such as antibiotics, nitric oxide therapy, biomaterial
gels, and nanozymes, effective treatment options are limited.^[ [66]^3
^] Typically, diabetic wounds are caused by severe bacterial infection,
abnormal inflammation, impaired collagen production, and reduced
epidermal development.^[ [67]^3a ^] Intriguingly, diabetic wounds also
exhibit a weakening of the endogenous electric field (EF).^[ [68]^4 ^]
The endogenous EF plays a critical role in wound healing by guiding the
migration of electroactive cells such as fibroblasts and epithelial
cells. Upon wound occurrence, a microcurrent, referred to as the
“injury current,” flows from the wound's edge (the anode) toward the
center (the cathode).^[ [69]^5 ^] Numerous studies have demonstrated
that electrical stimulation (ES) promotes wound healing in diabetic
patients with chronic non‐healing wounds.^[ [70]^4 , [71]^6 ^] A
double‐blind randomized control trial that encompassed administering
home‐based ES for diabetic foot ulcers revealed a significant 22%
reduction in wound area in the intervention group after 4 weeks.^[
[72]^7 ^] Commercialized ES dressings such as PosiFect and Procellera,
based on metallic electrodes, have been developed to address
non‐healing wounds.^[ [73]^8 ^] Recently, wound dressings capable of
delivering a directional electric field to treat acute wounds have been
reported.^[ [74]^9 ^] However, achieving a consistent ES analogous to
the injury current is challenging because of the absence of wearable
power supplies and circuit guidance in bioelectric dressings.^[ [75]^10
^] The human body, consisting of soft, water‐rich tissue, stands in
stark contrast to lab‐made bioelectronic devices constructed using
rigid, dry electronic components. This disparity has hindered the
desired seamless interfacing between biological tissue and
electrodes.^[ [76]^11 ^] Therefore, optimizing the tissue–electrode
interface is crucial for treating diabetic wounds of varying sizes,
shapes, and depths. Therefore, forming a compatible and stable
interface can facilitate electrical communication between neural
tissues and external electronics, notably enhancing the efficiency of
ES therapy.^[ [77]^12 ^]
Wearable and implantable electronics have attracted considerable
attention in the health industry owing to their distinct advantages,
such as facilitating seamless connections between humans and
information, applicability to personal healthcare monitoring systems,
and compatibility with smart textiles.^[ [78]^13 ^] The field of
wearable bioelectronics is particularly interested in advancing
flexible and stretchable sensors and functional circuits on 3D freeform
surfaces.^[ [79]^14 ^] These components are designed to capture and
interpret various bio‐signals from our bodies for medical applications.
Recently, a tactile sensor was developed using a
micro‐pyramid‐patterned double‐network ionic hydrogel, capable of
measuring variations in a triboelectric output signal.^[ [80]^15 ^] In
addition, ultrathin AgNWs/parylene hybrid films with an adjustable 3D
structure shaped through laser cutting have been applied for monitoring
electrocardiogram signals and sweat levels.^[ [81]^16 ^] Various
techniques, such as photopatterning, adaptive 3D printing, and transfer
printing, have been employed to develop functional circuits. A
high‐density transistor array (up to 347 transistors cm^−2) was
developed using dielectric and semiconductor materials through the
photopatterning process, contributing to the development of stretchable
digital circuits for integration into electronic skin.^[ [82]^14b ^]
Despite its potential, photopatterning is limited because of high
hardware requirements and technical complexities. By contrast, adaptive
3D printing is a closed‐loop technique that integrates real‐time
feedback control with the direct ink writing of functional materials to
incorporate devices on moving freeform surfaces.^[ [83]^17 ^] Another
efficient method for fabricating 3D electronic devices is transfer
printing, which employs stamps to lift devices from donor substrates
and imprint them onto receiver substrates.^[ [84]^18 ^] This is a
cost‐effective technique for generating complex geometries with high
precision and at a low cost.
While conventional wearable medical devices rely on chemical batteries
(button batteries) and commercial electricity for power, the
requirement of frequent replacement and maintenance renders them
cumbersome and cost‐ineffective. Self‐powered devices, such as
enzymatic biofuel cells (EBFCs), radio frequency energy harvesting
(RFEH) methods, and organic photovoltaic (OPV) cells, have emerged as
potential solutions for gathering operational power from renewable
energy sources, offering promising avenues for wireless self‐powered
health diagnostics and therapeutics.^[ [85]^19 ^] EBFCs are employed to
harvest energy in biofluid models and function as self‐powered
electrochemical glucose sensors.^[ [86]^20 ^] RFEH methods encompass
using rectennas, which combine an antenna and a rectifier, to convert
radio frequency (RF) energy into useful direct current power,
eliminating the need for batteries or wires.^[ [87]^21 ^] However, the
efficiency of these devices is affected by the stability of enzymes or
interference from other RF sources, limiting their suitability for
high‐power applications. OPV cells, ideal for indoor energy
harvesting,^[ [88]^22 ^] have gained attention as potential power
solutions capable of generating sufficient electrical charge for
wearable medical devices. For example, OPV cells have been utilized for
the ES of peripheral nerves or retinal cells, with the aim of restoring
neural functions.^[ [89]^23 ^] In the context of bioelectrical wound
dressings, OPV cells are superior to other wearable power systems owing
to their distinctive advantages. They efficiently convert light energy
into electrical energy, providing a self‐sustaining power source—an
essential criterion in the design of bioelectrical wound dressings.
This functionality aligns with the broader development trend of
wearable devices capable of generating self‐sustaining,
biologically‐responsive electrical stimulation from natural light
energy. Unlike piezoelectric and triboelectric nanogenerators, OPV
cells are wireless and self‐powered, eliminating the need for external
power supplies and resulting in a more compact and convenient wearable
system.^[ [90]^24 ^] Owing to their durability and stability under
diverse environmental conditions, OPV cells ensure consistent
performance even under bodily conditions.^[ [91]^25 ^] Further, the
biocompatibility of OPV cells ensures their safety for usage in direct
contact with biological tissues.^[ [92]^26 ^] Lastly, non‐fullerene
materials endow OPV cells with advantages such as a broader absorption
spectrum, adjustable energy levels, and enhanced resistance to
degradation.^[ [93]^27 ^] Therefore, OPV cells represent a reliable,
efficient, cost‐effective, and safe power source for bioelectrical
wound dressings.
Hydrogels have been extensively investigated in the field of tissue
engineering and wound dressing owing to their exceptional
biocompatibility.^[ [94]^28 ^] The soft and flexible nature of
hydrogels minimizes mechanical mismatches with biological tissues while
maintaining a wet environment with an abundance of ions that mimic
physiological conditions. Moreover, hydrogels exhibit remarkable
versatility in electrical, mechanical, antimicrobial, and biological
properties, making them particularly suitable for serving as an
interface between wound tissue and electronics.^[ [95]^29 ^] Hydrogels,
such as hyaluronan/chitosan, possess immunomodulatory properties.^[
[96]^30 ^] These natural polymers can facilitate the transition of
macrophages from an inflammatory to a proliferative (M2) phenotype,
thereby accelerating diabetic wound healing. In recent years,
commercially available wound dressings contain silver nanoparticles
(AgNPs) that enhance their bactericidal activity.^[ [97]^31 ^] AgNPs
have demonstrated efficacy against a variety of microorganisms,
including multi‐resistant bacteria, exhibiting great potential in the
treatment of diabetic wounds.^[ [98]^32 ^]
This paper proposes a novel protocol for developing a photovoltaic
wound dressing, referred to as the photovoltaic microcurrent hydrogel
(PMH) dressing, and identifying key protein alterations involved in
diabetic wound healing. The PMH dressing is designed to satisfy the
following criteria: a) the ability to provide biomimetic ES; b)
exhibiting antibacterial properties; c) capable of establishing
seamless contact with irregular wound beds; and d) non‐invasive,
wearable, and user‐friendly. The PMH dressing comprises flexible OPV
cells as a portable battery, a flexible micro–electro–mechanical
systems (MEMS) electrode as a guidance circuit,^[ [99]^33 ^] and a
conductive hydrogel as an electrode–wound interface (Figure [100]1 ).
First, the flexible OPV cells, consisting of Ag‐grid MEMS substrates
and non‐fullerene organic solar cells, are used to support ES under
light conditions.^[ [101]^25 , [102]^34 ^] Second, the MEMS electrode,
fabricated using transfer printing techniques, can guide micro‐currents
analogous to injury currents. Third, the conductive hydrogel is
employed as an interface between the MEMS electrode and the wound site.
In vivo experiments are conducted in diabetic mice with
bacteria‐infected wounds to evaluate the efficacy of the PMH dressing
in accelerating wound healing. Finally, we discern the possible
molecular mechanisms through which the PMH dressing facilitates the
healing of chronic wounds.
Figure 1.
Figure 1
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a) The fabrication process of the PMH dressing involves synthesizing
dopamine‐modified hyaluronic acid methacryloyl (HAMA), followed by the
preparation of conductive HD‐Ag hydrogel onto a flexible MEMS
electrode. The PMH dressing is obtained by connecting flexible OPV
cells to the MEMS electrode, designed to provide biomimetic electrical
stimulation under light conditions. The PMH dressing is then applied to
a bacteria‐infected skin wound on diabetic mice. b) The illustration
below outlines the ways the PMH dressing can accelerate wound healing
processes among diabetics: b‐i) providing biomimetic electrical
stimulation, b‐ii) inhibiting bacteria proliferation, b‐iii)
accelerating reepithelization, and b‐iv) enhancing extracellular matrix
organization and tissue remodeling.
2. Results and Discussion
2.1. Design, Fabrication, and Characterization of the PMH Dressing
The wearable PMH dressing employs a sandwich‐type architecture,
featuring flexible OPV cells at the top and a flexible MEMS electrode
in the middle (Figure [104]2a). At its base, serving as the wound
interface, is a multifunctional composite hydrogel composed of
dopamine‐modified HAMA (HAMA‐DA) and in situ‐formed AgNPs (HD‐Ag
hydrogel, Figure [105]1a). As illustrated in Figure [106]1b, under
light conditions, OPV cells convert light energy into electric power.
The MEMS electrode is connected to the OPV cells, and specially
designed microcircuits on the MEMS electrode guide the direction of the
electric current, achieving a biomimetic injury current that flows from
the wound's edge toward the central region. The HD‐Ag hydrogel in the
PMH dressing maintains conformal contact over time, reducing
interfacial impedance and delivering microcurrent stimulation from the
MEMS electrode to the wound area.
Figure 2.
Figure 2
[107]Open in a new tab
a) The schematic of the sandwich‐structured PMH dressing, composed of
large‐area flexible OPV cells on the top, a flexible MEMS electrode in
the middle, and a hydrogel in contact with skin at the bottom. b)
Stress finite element analysis of the MEMS electrode. c) Finite element
simulation of mechanical properties for organic semiconductor
materials. d) J–V curves of binary and ternary OPV cells. e) The J–V
features of the OPV cells with different areas. f) The external quantum
efficiency (EQE) spectra of the OPV cells.
PM6 and Y6 constitute the active layers of the devices, responsible for
converting light into electricity. Since their introduction in 2019,
PM6 and Y6 materials have become iconic in the field of organic
optoelectronics.^[ [108]^35 ^] Their photovoltaic conversion efficiency
significantly surpasses that of the fullerene system, demonstrating
excellent stability and a broad spectrum of light absorption.^[
[109]^36 ^] In this study, we introduced thermoplastic elastomer (TPE)
to enhance mechanical properties, such as stable interfacial adhesion
and bending durability, further improving optoelectronic properties
(Figure [110]S1a–c, Supporting Information). Small‐area devices were
fabricated using patterned ITO glass; while, large‐area flexible
organic solar cells were printed on a flexible electrode using a
sequential slot die method. Finite element simulation revealed the
superior elasticity of the flexible MEMS electrode and the organic
semiconductor light‐absorbing layer (Figure [111]S2b–d, Supporting
Information). This processing method, employed in previous reports,^[
[112]^22 , [113]^37 ^] has yielded numerous high‐performance
optoelectronic devices. Stress is evenly distributed on the edge of the
silver grid, ensuring apparatus durability and resilience
(Figure [114]2b). Under pressure, the organic semiconductor layer's
stress gradually disperses, demonstrating strong tensile and
compression resistance and making it suitable as a dressing on moving
skin tissues (Figure [115]2c). As depicted in Figure [116]2d, the
open‐circuit voltage and short‐circuit current of the ternary materials
are delineated. The open‐circuit voltage remains stable at ≈1.0 V,
while the short‐circuit current is maintained at the standard of 20 mA
cm^−2, a level consistent with our earlier studies that confirm it does
not lead to significant skin burns.^[ [117]^25 ^] The JV curves of our
single‐device and series‐device modules are also presented
(Figure [118]2e). The ternary system ensures a stable supply of voltage
required for ES.^[ [119]^38 ^] As shown in Figure [120]2f, the EQE
spectra for the verified devices exhibit power conversion efficiency
(PCE) test accuracy. The broad absorption peak beyond 900 nm results
from the Y6 molecule. The overlapping voltage in larger devices
suggests potential use over larger skin areas.
For wounds with a diameter of ≈10 mm, an electrical field of 1.0 V
(equivalent to 100 mV mm^−1) is reported as suitable for eliciting a
biological response.^[ [121]^9 , [122]^39 ^] Under typical indoor light
conditions (2700 K LED array with a luminous intensity of 1000 lux),
OPV cells can generate a stable open‐circuit voltage of 1.07–1.12 V.^[
[123]^22b ^] The 0.04 cm^2 small‐area flexible device achieves an
impressive PCE of up to 16%; while, the 1.25 cm^2 large‐area flexible
device attains a PCE of up to 13%, demonstrating the advanced level of
device technology.^[ [124]^40 ^] Due to the excellent flexibility of
poly ethylene terephthalate (PET) loaded with silver grids (Young's
modulus of 0.45 MPa), the silver grid composite substrate devices
(PET‐solar cells and PET‐electrode) display a reduced bending pattern
and adapt well to the skin surface. Moreover, given that biological
safety is paramount for any wound dressing intended for medical use,
our research assesses the potential toxicity of the bioelectronic
components integrated into the novel dressing. Our findings confirm the
excellent biocompatibility of the OPV cells and MEMS electrodes (Figure
[125]S3, Supporting Information), consistent with our previous study.^[
[126]^25 ^]
An ES therapy for wound healing, capable of providing a biomimetic EF
or injury current from the periphery to the center of the wound, is
often considered the optimal solution. The EF intensity at skin injury
sites typically falls within the range of 100–200 mV mm^−1.^[ [127]^9 ,
[128]^41 ^] For instance, the EF in mouse skin wounds is measured at
122 ± 9 mV mm^−1.^[ [129]^42 ^] Thus, our aim is to achieve an ES
intensity of ≈100 mV mm^−1. Most current portable ES wound dressings
utilize a stimulation intensity equivalent to the physiological EF, and
some innovative products use hollow ring electrodes to generate a
biomimetic EF (refer to Table [130]S2, Supporting Information).
However, due to the variable depth of wound tissues, it is challenging
for regular‐shaped metal or silicone rubber electrodes to produce
uniform ES, potentially leading to localized aggregation of
electrolysis products and subsequent chemical burns.
The development of miniaturized and wireless power supply devices is
crucial to the advancement of ES wound dressings. Traditional power
supply devices for electric stimulation are often characterized by
bulkiness, tethering, and inconvenience for patients, thereby limiting
their applicability and adoption.^[ [131]^43 ^] The advent of compact,
wireless power supply devices has paved the way for the development of
user‐friendly, efficient, and portable ES therapies for wound
healing.^[ [132]^44 ^] Details on the performance and applications of
various energy‐harvesting strategies currently available for wearable
biomedical devices are summarized in Table [133]S3, Supporting
Information. OPV cells fabricated with non‐fullerene acceptor (NFA)
materials offer unique benefits for ES wound dressings. NFA materials
are marked by their potential for structural customization, high
property tunability, intense absorption of visible and near‐infrared
light, and remarkable n‐type semiconducting characteristics.^[ [134]^45
^] Further, these OPV cells offer a range of benefits including
cost‐effectiveness, flexibility, semi‐transparency, high mass power
density, short energy payback time, and compatibility with large‐area
printing processes, making them ideal for scalable production.^[
[135]^46 ^] With the surge of biomedical and health industry, a variety
of organic semiconductors with excellent biocompatibility and reliable
diagnostic/therapeutic functions is developed.^[ [136]^47 ^] The OPV
cells applied in this study are flexible, biologically safe, and
capable of providing sufficient voltage for the PMH dressing, rendering
them highly suitable and promising self‐powering devices for innovative
wearable bioelectronics. In the operating mode of the PMH dressing, OPV
cells and MEMS electrodes consistently generate a biomimetic EF,
stimulating the wound through microcircuits on the MEMS electrode
surface and the conductive hydrogel. This model proves particularly
beneficial for chronic wound healing as the bioactive hydrogel
interface reduces the mechanical strength mismatch between tissue and
electrode, delivering uniform and stable ES to the wound.
2.2. Synthesis and Material Properties of HD‐Ag Hydrogel Interface
Hydrogels possess exceptional biomaterial compatibility, high water
content, softness, and adaptable functionality, rendering them ideal
bridging materials for achieving seamless connections between MEMS
electrodes and wound tissue. In this study, we formulated the HD‐Ag
hydrogel, which serves as an interface between the MEMS electrode and
wound tissues. It is composed of a combination of HAMA‐DA and Ag
nanoparticles. We developed a series of HD‐Ag hydrogels by
progressively increasing the concentration of Ag^+ ions from 1 to 3 mg
mL^−1, which were denoted as HD‐Ag[1], HD‐Ag[2], and HD‐Ag[3],
respectively. HD represents the hydrogel without AgNPs. The fabrication
process and characterizations of the HD‐Ag hydrogel are detailed in
Figures [137]S4–S8, Supporting Information). The HD‐Ag hydrogels were
observed to release Ag^+, with Ag concentrations in the range of
1–5 ppm detected after 12 h (Figure [138]3a). This aligns with the
effective bactericidal concentration of 0.1–5.4 ppm, suggesting that
the HD‐Ag series hydrogels may possess effective antibacterial
capabilities.^[ [139]^48 ^]
Figure 3.
Figure 3
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Studies of the physical properties, cytocompatibility, and
antibacterial activity of the HD‐Ag hydrogels. a) The release profile
of Ag^+ ions, b) swelling ratio, and c) electrical conductivity of the
HD‐Ag hydrogels. d) Representative live/dead stain images of
fibroblasts and keratinocytes co‐cultured with the hydrogels for 3
days; green represents live cells, while red represents dead cells.
Cells cultured on a tissue culture plate (TCP) were used as control.
e,i) Representative photographs and quantification analysis of the
inhibition zone against MRSA or PAO bacteria by different hydrogel
disks. f) Quantitative analysis of the MRSA and PAO colonies incubated
with each hydrogel for 24 h. The initial number for two bacterial types
was 5 × 10^7 in this experiment. g,h) Cell proliferation analysis of
fibroblasts and keratinocytes co‐cultured with the HD‐Ag hydrogels for
1, 3 and 5 days. The data were presented as mean ± SD (n = 3, *p <
0.05).
Ag has long been recognized as an effective treatment for burn wounds,
dating back to ancient times and still in use today.^[ [141]^49 ^]
Recently, Ag has been engineered as a nanomaterial due to its enhanced
efficacy in biomedical applications and antimicrobial properties.^[
[142]^50 ^] AgNPs undergo slow oxidation reactions to generate silver
ions (Ag+), exhibiting antibacterial activity while avoiding adverse
reactions from a sudden local release of high concentrations of Ag^+.^[
[143]^51 ^] Through light‐induced reduction reactions, a rapid and
uniform reduction of Ag^+ in situ can be achieved, enabling the even
distribution of AgNPs within the hydrogel system, and this uniform
distribution of particles ensures good conductivity of the hydrogel.^[
[144]^52 ^] In the present study, while photo‐crosslinking occurred in
the HD hydrogel, Ag^+ ions were converted in situ into evenly
distributed AgNPs within the hydrogel via a reduction reaction mediated
by UV light irradiation and catechol groups.^[ [145]^53 ^] This
formulation ensured that the HD‐Ag hydrogel maintained reliable
conductivity while allowing precise control of Ag^+ concentrations. In
addition, HD‐Ag hydrogels exhibited favorable swelling properties
(Figure [146]3b). Their water absorption capacity enabled effective
absorption of effluents, maintaining a moist wound environment that
prevented bacterial infection and accelerated the healing process.
Further, the excellent electrical conductivity of HD‐Ag hydrogels made
them a suitable interface between biological tissues and bioelectronics
(Figure [147]3c).
2.3. Biocompatibility and Antibacterial Properties of HD‐Ag Hydrogel
Interface
Outstanding biocompatibility is a fundamental property of hydrogels
that makes them suitable as interfaces between tissue and
bioelectrodes. In this study, we assessed the biocompatibility of HD‐Ag
hydrogels using basal layer keratinocytes (HaCaT) and dermal
fibroblasts (NIH/3T3) through live/dead staining (Figure [148]3d;
Figure [149]S9, Supporting Information) and the cell counting kit‐8
(CCK‐8) assay (Figure [150]3g,h). We also conducted subcutaneous
embedding experiments (Figure [151]S10, Supporting Information). Our
results confirmed the good biocompatibility of HD, HD‐Ag[1], and
HD‐Ag[2] hydrogels, whereas significant toxicity was associated with
the HD‐Ag[3] hydrogel. Diabetic wounds were characterized by
hard‐to‐heal skin ulcers and bacterial infections, commonly caused by
Methicillin‐resistant Staphylococcus aureus (MRSA, ATCC 43300) and
Pseudomonas aeruginosa (PAO, ATCC 15962).^[ [152]^54 ^] HD‐Ag hydrogel
exhibits excellent antibacterial properties, primarily attributed to
the presence of AgNPs distributed within the hydrogel
(Figure [153]3f,i). Among these candidates, HD‐Ag[2] emerged as the
most promising due to its impressive conductivity, satisfactory
biocompatibility, and antibacterial properties. Therefore, we chose to
incorporate HD‐Ag[2] into the PMH dressing for further research.
2.4. In Vivo Effect of the PMH Dressing on Bacteria‐Infected Diabetic Wound
Healing
This study aimed to evaluate the effectiveness of PMH dressings on skin
wounds co‐infected with MRSA and PAO. We established type II diabetic
mouse models through a high‐fat diet induction, STZ injection, and
maintenance of a hyperglycemic state (Figure [154]4a). In addition, we
tested photovoltaic microcurrent patches (referred to as PM patches,
which lacked hydrogel compared to PMH dressing) and HD‐Ag[2] hydrogel.
The wounds in the Blank group were treated with a Tegaderm film
dressing, a commercially available wound dressing known for its
outstanding water repellency and moisture vapor penetrability.^[
[155]^55 ^] As shown in Figure [156]4b, a yellowish purulent discharge
was evident around the wounds on day 13 for both the Blank and PM patch
groups. In contrast, the yellow exudate disappeared earlier in the PMH
dressing group and the HD‐Ag[2] group (by day 7). On day 22, wounds
treated with PMH dressing achieved a high closure ratio of 97%; while,
those in the other groups remained unhealed (less than 80%)
(Figure [157]4c,f).
Figure 4.
Figure 4
[158]Open in a new tab
PMH dressing enhanced bacteria‐infected diabetic wound healing in vivo.
a) Timeline of model establishment, dressing treatment, wound
observation, and tissue harvest in the animal experiment. b)
Representative images of the wound at predetermined time points. c)
Wound healing boundaries overlayed on each image. d) HE and e) Masson
staining of tissue sections from the wounded areas and adjacent skin.
f) Analysis of wound closure. g) Analysis of re‐epithelization
percentage. h) Analysis of collagen content. Data were shown as mean ±
SD (n = 3, *p < 0.05 when compared with Blank group; #p < 0.05 when
compared with PM patch group; and &p < 0.05 when compared with HD‐Ag[2]
group).
The analysis of wound area demonstrated that both HD‐Ag[2] and PMH
dressing significantly accelerated the wound healing process, with the
latter exhibiting the most pronounced therapeutic effect. The role of
HD‐Ag[2] hydrogel in promoting wound healing was especially noteworthy.
In environments with ample lighting conditions, OPV cells enabled the
PMH dressing to deliver stable and continuous biomimetic ES to the
wound. Even under low‐light conditions, the HD‐Ag[2] hydrogel interface
continued to offer therapeutic benefits, making the dressing adaptable
for various environments. In addition, we harvested wound tissues at
the endpoint of this study and conducted histological analysis through
H&E and Masson's staining. Compared with the Blank group, wounds
treated with PMH dressing demonstrated a significant reduction in
inflammatory cell infiltration, as indicated by the small blue‐stained
cells (Figure [159]4d,e). Moreover, a fundamental epithelial structure
was identified at the wound sites after PMH dressing treatment
(Figure [160]4d,g). The results from Masson's staining revealed a
significant increase in the regeneration of mature fibrous tissue
(identified by intense blue fibers) in the PMH dressing group, with the
area of collagen deposition scaling up to ≈320% (with the Blank group
normalized to 100%) (Figure [161]4e,h). Compared with the HD‐Ag[2]
group, the PMH dressing delivered biomimetic ES stimulation to the
wound through wearable flexible OPV cells, significantly promoting the
healing of wounds in the repair phase. Unlike the recently reported
bioelectrical dressing,^[ [162]^9 ^] the PMH dressing effectively
supplied biomimetic exogenous ES with appropriate electric field
strength through seamless contact with the wound via the hydrogel
interface, thereby promoting the healing of diabetic wounds.
Skin wounds in diabetic mice, due to their exposure to air and elevated
local glucose concentration, are susceptible to bacterial
proliferation.^[ [163]^56 ^] The HD‐Ag[2] hydrogel, acting as a
tissue–electrode interface, effectively eradicates bacteria present in
diabetic wounds. It accelerates the transition from the inflammatory to
the repair stage, thereby achieving outstanding therapeutic results.
Moreover, chronic inflammation is a hallmark of non‐healing diabetic
wounds.^[ [164]^57 ^] In diabetic wounds, the pro‐inflammatory to
anti‐inflammatory macrophage phenotypic switch is impaired and
contributes to chronic inflammation.^[ [165]^58 ^] In this context,
macrophages mainly exhibit a pro‐inflammatory M1 phenotype and express
high levels of inflammatory cytokines (e.g., TNF‐α and IL‐1β); while,
the number of M2 phenotype macrophages, which produce anti‐inflammatory
and angiogenic cytokines (e.g., TGF‐β[1] and IL‐10), decreases. In this
study, macrophage phenotypes in the wound area were evaluated using
iNOS (M1) and TGF‐β1 (M2) immunofluorescence staining. We discovered
that during the mid‐healing period (15d), wounds treated with HD‐Ag[2]
and PMH dressing had fewer M1 type macrophages and more M2 type
macrophages than the Blank and PM patch groups. Further, we assessed
the transcription levels of inflammatory cytokines (TNF‐α, IL‐1β) and
anti‐inflammatory cytokines (TGF‐β1, IL‐10) in wound tissues via
RT‐qPCR. The results suggest that inflammation levels remained high in
the Blank and PM patch groups; while, the HD‐Ag[2] and PMH dressing
groups started transitioning into the repair phase (details can be
found in Figure [166]S12, Supporting Information). This can be
attributed to the immunomodulatory and anti‐inflammatory activities of
high molecular weight (100–250 kDa) hyaluronic acid.^[ [167]^59 ^]
Hence, the antimicrobial and inflammation‐regulating properties of
HD‐Ag[2] hydrogel in PMH dressing are highly beneficial for the healing
of diabetic wounds.
2.5. Potential Mechanism of PMH Dressing Affecting Diabetic Wound Healing
The differential expression of proteins (DEPs) between the Blank group
and the PMH dressing group was identified through label‐free proteomics
(LFP) (Figure [168]5a). In total, 4066 proteins were identified as
co‐expressed through LFP detection (Figure [169]5b). Principal
component analysis (PCA) demonstrated markedly different component
profiles between the two groups (Figure [170]5c). In the PMH dressing
group, 2280 proteins were upregulated (with a p value < 0.05 and a fold
change value greater than 2); while, 287 proteins were downregulated
(with a p value < 0.05 and a fold change value less than 0.5)
(Figure [171]5d). The results of the GO enrichment analysis showed that
proteins related to extracellular matrix collagen deposition and the
keratinization of epithelial cells were significantly upregulated
(Figure [172]5f). In contrast, downregulated proteins were enriched in
coagulation, inflammatory response, extracellular matrix, and matrix
metalloproteinase (MMP) activity (Figure [173]5g). The functional
enrichment analysis results for DEPs indicated that upregulated
proteins participate in extracellular matrix organization,
intercellular connections, and the formation of the cornified envelope
(Figure [174]5h). Downregulated proteins were enriched in coagulation,
bacterial infection, MMP activity, and degradation of the extracellular
matrix (Figure [175]5i).
Figure 5.
Figure 5
[176]Open in a new tab
a) Schematic diagram of tissue collection and proteomics analysis. b)
The expression levels of identified co‐expressed proteins are
illustrated in a heatmap. c) PCA results. d) DEPs are presented in a
volcano plot. e) Venn diagram comparing DEPs to an extracellular matrix
proteomics dataset. f) Results of the gene ontology (GO) analysis for
the upregulated proteins. g) Results of the GO analysis for the
downregulated proteins. h) Functional analysis (KEGG and Reactome
analysis) results of the upregulated proteins. i) Functional analysis
results of the downregulated proteins.
To further understand the effects of PMH dressing treatment on the
extracellular matrix, we analyzed the ECM‐related components among the
DEPs and identified 36 upregulated and 9 downregulated proteins
(Figure [177]5e). The upregulated proteins included various ECM
components, such as collagens, elastin, fibrillin, and laminin. In
addition, proteins related to intercellular junctions, such as galectin
and spondin, along with proteins associated with keratinocyte
differentiation, such as repetin and transglutaminase, were also found
to have increased expression (Table [178]S4, Supporting Information).
Enzymes associated with ECM degradation, specifically MMP, and proteins
involved in the activation of coagulation, such as von Willebrand
factor and thrombospondin, were observed to be downregulated (Table
[179]S5, Supporting Information). Further, an interaction network
analysis of the 45 differential extracellular matrix proteins suggests
that PMH dressing treatment promotes collagen deposition (Figure
[180]6a,b). Through immunofluorescence techniques, we studied the state
of collagen regeneration and vascular distribution in the tissues of
all groups. We noted that wounds treated with PMH dressing exhibited
markedly higher collagen deposition than the other wounds, evident from
the brighter green fluorescence (Figure [181]6c,d). At the same time,
the quantity of blood vessels (detected through the vascular
endothelial‐specific antigen CD31) in wounds treated with PMH dressing
was significantly diminished (Figure [182]6c,e).
Figure 6.
Figure 6
[183]Open in a new tab
Pathway analysis of the a) highly expressed DEPs and b) low expressed
DEPs correlated with extracellular matrix reveals significant
enrichment of proteins involved in collagen organization and
metallopeptidase activity. GeneMANIA ([184]https://genemania.org/) was
used for the analysis. c) Immunofluorescence observation of collagen‐I
(Col‐I) and CD31 staining. The immunofluorescence intensity of positive
d) Col‐I and e) CD31 cells present at the wound beds in each group was
compared to that of the blank group, which was set as 100%, with other
groups calculated as a percentage. Data were shown as mean ± SD (n = 3,
*p < 0.05 when compared with Blank group; #p < 0.05 when compared with
PM patch group; and &p < 0.05 when compared with HD‐Ag[2] group).
ES has been shown to increase the growth, migration, and extracellular
matrix synthesis of fibroblasts derived from diabetic patients.^[
[185]^60 ^] The combination of ES with wearable devices^[ [186]^25 ^]
or biodegradable, electroactive hydrogels^[ [187]^61 ^] has yielded
promising results in regulating skin cell behaviors and accelerating
the wound healing process. Herein, the PMH dressing integrates the
advantages of wearable devices and electroactive hydrogels. Through the
external supply of biomimetic ES, it actively modulates collagen
deposition and remodeling; thus, accelerating the healing process. It
is also worth noting that proteins related to vascular function are
downregulated in wound tissues treated with PMH dressing. The formation
of new blood vessels during wound repair is indispensable for efficient
tissue repair. Nevertheless, recent studies imply that reducing
angiogenesis experimentally may be advantageous to boost long‐term
healing results.^[ [188]^62 ^] Both fetal skin and oral mucosal wounds
demonstrate fast healing with little scarring.^[ [189]^63 ^] Moreover,
wounds treated with PMH dressing transition to the remodeling phase
earlier than those with Tegaderm. This phase entails the
reorganization, degradation, and resynthesis of the extracellular
matrix; while, granulation tissue is progressively remodeled into
acellular and less vascular scar tissue.^[ [190]^64 ^]
The role of MMPs is to facilitate wound debridement and regulate the
metabolism of the ECM during the healing of diabetic wounds.^[ [191]^65
^] The two primary active MMPs in human diabetic wound tissues, MMP‐8
and MMP‐9, have specific roles in wound healing.^[ [192]^66 ^] A study
by Chang et al. assessed how these two MMPs affect the repair of
diabetic wounds in mice, revealing that supplementary MMP‐8 treatment
facilitated wound closure, whereas MMP‐9 negatively impacted healing.^[
[193]^67 ^] Our study; however, observed a downregulation of both forms
of MMPs, which differs from prior findings. To understand this
inconsistency, we conducted immunofluorescence staining on wound
tissues during the early (7d), middle (15d), and late (22d) healing
stages. We observed a high expression of MMP‐8 in wounds treated with
PMH dressing during the early and middle stages, decreasing in the
later stage, whereas MMP‐9 was mostly expressed early on (Figure
[194]S13, Supporting Information). Conversely, in the Blank group,
MMP‐8 was mainly expressed in the late stage; while, MMP‐9 was abundant
throughout. Our investigation focused on the late healing stage (when
the wound is transitioning from repair to remodeling with a low
expression of both MMPs), unlike Chang et al.’s study, which focused on
the middle stage (high MMP‐8, low MMP‐9). Overall, the proposed PMH
dressing could elevate MMP‐8 levels in diabetic wounds, accelerate the
transition from repair to remodeling, and promote healing.
Overall, these results suggest that the PMH dressing aids in wound
healing by modulating extracellular matrix regeneration, eliminating
bacteria, regulating inflammatory responses, and altering vascular
functions. Therefore, the PMH dressing not only emerges as a
therapeutic intervention but also enhances our understanding and
strategy for diabetic wound care.
3. Conclusion
In this study, we developed the PMH dressing, a novel
sandwich‐structured solution tailored to address the intricate
challenges of diabetic wound care. By synergistically integrating
flexible OPV cells, MEMS electrodes, and the multifunctional HD‐Ag
hydrogel, we designed a wireless bioelectrical wound dressing that not
only delivers biomimetic ES but also ensures optimal tissue–electrode
interfacing. Leveraging ultraviolet light, we synthesized the HD‐Ag
hydrogel, which serves as a tissue‐electrode interface for ES delivery.
Our findings confirmed the hydrogel's superior biocompatibility and
antibacterial activity against common pathogenic bacteria. The dressing
exhibited significant therapeutic effects on diabetic wounds
complicated by bacterial infections. We demonstrated that the PMH
dressing promotes the healing of diabetic wounds by regulating the
regeneration and remodeling of extracellular matrix proteins,
accelerating re‐epithelialization, and modulating chronic inflammation.
As a groundbreaking prototype that integrates organic OPV, MEMS
microcurrent electrodes, and hydrogel, the PMH dressing has the
potential to usher in a new generation of more effective and
patient‐friendly wound care solutions. While our research has confirmed
the benefits of PMH dressing on diabetic wounds, there is insufficient
evidence supporting its effectiveness on other prevalent chronic wounds
such as burns, pressure ulcers, and radiation skin injuries, presenting
a research direction for future studies. In addition, our forthcoming
study will focus on the development of environmentally sustainable and
recyclable wound dressings that are also cost‐effective; thus, setting
the stage for potential industrialization.
4. Experimental Section
Fabrication and Characterization of Flexible OPV Cells
PM6 and Y6, purchased from Chemscitech Inc. (Canada), were used as
received.
Poly(2‐methoxy‐5‐(3′,7′‐dimethyloctyloxy)−1,4‐phenylenevinylene) (PM6),
a conjugated polymer, had been employed to develop efficient, low‐cost,
and lightweight OPV cells. In addition, another conjugated polymer,
known as Y6 Polymer, was also utilized in the development of such OPV
cells. The molecular formulas of PM6 and Y6 are shown in Figure
[195]S1a,b, Supporting Information. In this study, the authors
introduced a (TPE) to enhance the mechanical properties such as stable
interfacial adhesion and bending durability and to further improve the
optoelectronic properties (Figure [196]S1c, Supporting Information).
All supplies were used as received. ITO glass, with a sheet resistance
of 15 Ω sq^−1, was purchased from CSG HOLDING Co., Ltd. (China). ZnO
nanoparticles used in this layer were synthesized following a reported
protocol.^[ [197]^68 ^] Active layer solutions were prepared by mixing
PM6 and Y6 materials with o‐xylene and 1,8‐diiodooctane (DIO) in a
specific ratio (D/A ratio of 1:1.2). These solutions were spin‐coated
onto the substrate in an N[2] glove box to form the active layers of
the OPV cells. After applying the active layers, a thin layer of MoOx
was deposited as an anode interlayer via vacuum deposition. This layer
helped improve the performance of the solar cells by facilitating
electron transfer from the active layers to the Ag top electrode. The
Ag electrode was then deposited onto the device at a thickness of
100 nm, completing the fabrication of the OPV cells.
Fabrication of large‐area and module OPV cells was conducted under
ambient conditions instead of in a controlled environment like an N[2]
glove box. The device fabrication method was adapted from the published
work with appropriate modifications.^[ [198]^69 ^] The bottom electrode
of the cells was made from PET/silver‐grid substrates with a striped
pattern, which had a sheet resistance of 10 Ω sq^−1 and a transmittance
of 95% or higher. The R2R instrument (MiniRoll Coater, FOM
Technologies) was used to fabricate the large‐area flexible OPV cells.
Modules with two and four serial stripes were made using this process.
After optimization, ZnO nanoparticles dispersed in isopropanol were
selected to form the electron transport layer of the devices. The
concentration of the ZnO nanoparticles was maintained at 15 mg mL^−1.
The active layers of the OPV cells were produced by dissolving the
donors and acceptors in o‐xylene, with DIO added as an additive
solvent. The active layer solution was then applied using a slot‐die
coater. After applying the active layers, the devices were placed in a
vacuum evaporation system to deposit a thin layer of MoOx as the anode
interlayer and a layer of Ag as the top electrode. The active areas of
the flexible OPV modules were 1.25 and 2.50 cm^2, respectively. The
anode made contact with the electrode on the MEMS electrode; while, the
cathode was designed to connect with the skin. The PEDOT: PSS material
(Clevios PH1000, Heraeus, Germany) was spin‐coated onto the PET/Ag‐grid
substrates at 800 rpm and then baked in ambient air (120 °C for
15 min). This process enhanced the surface flatness and wettability of
the PET/Ag‐grid substrates, making them suitable for use as the bottom
electrode of the OPV cells.
The fabrication process for the rest of the OPV cells was identical to
that used for the reference cells made on glass/ITO substrates. This
involved applying the ZnO electron transport layer, the active layers,
and the anode interlayer and top electrode using the methods described
above.^[ [199]^35 , [200]^38 ^] These steps completed the fabrication
of the OPV cells on PET/Ag‐grid/PH1000 hybrid electrodes. The OPV cells
were also fabricated with patterned ITO glass, a transparent conductive
material used as the substrate for the active layers. After cleaning
the ITO glass, a ZnO electron transport layer was prepared by
spin‐coating at 3000 rpm. This layer assisted in transporting electrons
generated by the active layers to the electrodes, where they could be
collected and used to generate electricity.
The electrical output of the device was evaluated under AM 1.5G (100 mW
cm^−2) conditions with a Newport Thermal Oriel 91159A solar simulator,
and mechanical modeling was conducted using finite element simulation
tools.
Fabrication and Characterization of Flexible MEMS Electrodes
The fabrication of flexible MEMS electrodes involved several steps. The
first step entailed depositing a layer of UV photoresist onto a glass
substrate and patterning it using photolithography. This process
rendered a specific shape, such as a hexagonal honeycomb structure, on
the substrate. This shape promoted maximum light transmission and
exhibited low optical loss, resulting in a high light transmittance
(80–90%) in the flexible electrode. The subsequent step involved
patterning the PET substrate. This was achieved by applying UV glue to
the PET substrate and imprinting the patterned UV photoresist film atop
the UV glue using a nickel master. This action created a mask‐like
pattern on the grid, reserving a grid thickness of 3 µm. The third step
consisted of filling the grooves in the patterned PET substrate with
silver nano ink and sintering the ink at 150 °C for 15 min. This
process formed a conductive silver grid on the PET substrate. The grid
was then electroplated with copper for ≈5 min, utilizing an
electroplating current of 2 A. This allowed for a dense copper coating
that effectively prevented further oxidation. Lastly, the surface of
the film was smoothed with an aqueous solution containing silica
particles. This action created a dense microstructure on the surface of
the Cu layer, which passivated the electrodes and prevented further
oxidation. These steps concluded the fabrication of the flexible MEMS
electrodes.
The structure of the microcircuit on the MEMS electrode was examined
using a scanning electron microscope (SEM, SU‐70, Hitachi, Japan), and
mechanics modeling was conducted using finite element simulation tools.
Biocompatibility of Flexible OPV Cells and Flexible MEMS Electrodes
HaCaT were employed to test the cellular biosafety of flexible OPV
cells and MEMS electrodes. HaCaT cells were cultivated using a standard
protocol.^[ [201]^70 ^] The electronic devices were disinfected by
immersion in 75% alcohol for 30 min; and then, equilibrated in the
culture medium for 12 h. Cells were seeded in 12‐well plates at a
density of 50 000 cells per well and co‐cultured with the electronic
device samples (25 mg mL^−1). After 1 and 3 days of co‐culturing, cell
morphology was observed under an optical microscope. Cell viability at
predetermined time points was assessed using a Live/Dead staining kit
and a CCK‐8 cell viability assay kit. The stained cells were observed
under an inverted fluorescence microscope (IX53, Olympus).
The extracts of the electronics were prepared for the hemolysis
experiment. Flexible OPV cells and MEMS electrode samples were immersed
in DMEM culture medium (25 mg mL^−1) and incubated at 37 °C for 48 h to
prepare the conditioned medium. Erythrocytes were separated from rat
blood by centrifugation (120 × g, 10 min) and washed three times with
DPBS (resuspending and centrifuging each time) to remove damaged
erythrocytes. The methodology for the hemolysis test was consistent
with the procedure detailed in the published works.^[ [202]^71 ^]
Triton X‐100 (0.1%) was used as the positive control; while, DPBS
served as the blank control. The hemolysis rate (H) was calculated
according to Equation ([203]1):
[MATH: H=Ap−AbAt−Ab×100% :MATH]
(1)
where A [p] represents the absorbance of the test electronic device
sample, A [t] represents the absorbance of the positive control (Triton
X‐100), and A [b] represents the absorbance of the blank control
(DPBS).
Synthesis of HAMA and Dopamine Grafting
Methacrylic anhydride (MA),
2‐hydroxy‐4′‐(2‐hydroxyethoxy)−2‐methylpropiophenone (I2959),
N‐(3‐dimethylaminopropyl)‐N′‐ethylcarbodiimide hydrochloride (EDC),
hydroxyl‐PEG‐NHS ester (NHS), and Irgacure 2959 (I2959) were procured
from Aladdin Chemistry Co., Ltd (Shanghai, China). Sodium hyaluronate
(≈1.8–2.2 MDa) was sourced from Bloomage Biotechnology Co., Ltd
(Shandong, China). The synthesis of HAMA followed a previously reported
method with minor modifications.^[ [204]^72 ^] Briefly, hyaluronate
(HA; 3 g, monomer molar amount = 8 mmol) was dissolved in deionized
water (300 mL) and stirred for 2 h to form a 1 wt% solution.
Methacrylic anhydride (MA, 12 mL, 80 mmol) was then gradually added to
the mixture. The solution was placed in an ice bath and vigorously
stirred using a digital overhead stirrer (Eurostar 20 Digital, IKA).
The pH of the mixture was monitored and, if necessary, adjusted to 8.5
with a 0.1 m NaOH solution. The reaction was allowed to proceed for 24
h. The product was dialyzed for 2 days using a membrane with a
molecular weight cutoff (MWCO) of 14 000 Da against deionized water,
which was refreshed three times daily to remove unreacted reagents and
byproducts. Finally, the product was lyophilized and stored at −20 °C
until use.
Dopamine (DA) was grafted onto HAMA using EDC as a crosslinker,
following a previous study.^[ [205]^73 ^] HAMA (260 mg, monomer molar
amount = 0.5 mmol) was dissolved in 10 mL deionized water and stirred
for 24 h at room temperature. EDC (115 mg, 0.6 mmol) and NHS (69 mg,
0.6 mmol) were added to this solution. After 20 min, DA (114 mg,
0.6 mmol) was added, and the pH of the mixture was adjusted to 5.5 via
the addition of HCl (0.1 m) and NaOH (0.1 m). The mixture reacted
overnight under nitrogen at room temperature. Finally, the solution was
dialyzed against deionized water using a membrane (MWCO < 14 000 Da)
for 7 days under acidic conditions. The product, HAMA‐DA, was then
lyophilized and stored at −20 °C until use.
The chemistry of HD was analyzed using the ^1H‐NMR spectrum, and the
catechol content was quantified by measuring the optical absorbance at
280 nm with a spectrometer (UV‐1800, Shimadzu). A dopamine standard
curve was generated for quantification of DA, which indicated that the
degree of dopamine substitution for HD was ≈10%.
Preparation of HD‐Ag Hydrogels
HD was dissolved in a phosphate buffer solution (PBS) to form a 2 wt%
solution. Silver nitrate (AgNO[3]) was added to the solution at varying
concentrations of 1, 2, and 3 mg mL^−1. The photoinitiator I2959 (0.5
wt%) was then incorporated under vigorous stirring. Subsequently, these
hydrogel precursors were pipetted onto substrates and exposed to
ultraviolet radiation (365 nm, 300 mW cm^−2) for 1 min to complete
gelation using a UV lamp (Lumen Dynamics Group Inc., Canada). The
resulting hydrogels were labeled as HD‐Ag[1], HD‐Ag[2], and HD‐Ag[3],
respectively, where HD denotes hydrogels without silver content.
Characterizations of HD‐Ag Hydrogels
Several tests were conducted to investigate the morphological and
physicochemical characterizations of HD and HD‐Ag hydrogels. These
included observations under SEM and transmission electron microscopy
(TEM), X‐ray photoelectron spectroscopy (XPS) analysis, rheological
tests, Ag^+ ion release studies, conductive tests, and swelling tests.
The hydrogels were dried using a vacuum freeze‐drier (Scientz‐18ND,
Ningbo, China); then, sputter‐coated with a thin gold layer for SEM
observation (S‐4800, Hitachi, Japan). For TEM investigation, the
hydrogel samples were fixed in a mixture of 4% paraformaldehyde and 2%
glutaraldehyde in 0.1 m sodium cacodylate (pH 7.4). After rinsing with
cacodylate (0.1 m), the samples were post‐fixed with 1% osmium
tetroxide to enhance contrast and then dehydrated through a graded
series of ethanol solutions. Subsequently, the samples were embedded in
resin, sectioned, mounted on TEM grids, and stained with uranyl acetate
and lead citrate. Images were acquired at an accelerating voltage of
80 kV using a TEM. XPS analysis was conducted using an ESCALAB 250Xi
spectrometer (ThermoFisher Scientific, USA) in the range of ≈0–1200 eV
to determine the valence state of silver within the hydrogel system.
For this, HD and HD‐Ag[2] were tested as examples of pure hydrogel and
silver‐doped hydrogel, respectively.
The rheological behavior and photocuring kinetics of the samples were
examined using a rheometer (Discovery hybrid rheometer 2, USA) with an
LED accessory emitting light at 365 nm.^[ [206]^74 ^] The release of in
vitro Ag^+ ions was quantified by immersing the hydrogel samples in
phosphate‐buffered saline (PBS, 20 µL µg^−1) at 37 °C. At predetermined
time points (1, 2, 6, 24, 48, 72, and 96 h), the supernatant was
collected and the media were replenished with an equal volume of fresh
PBS. The released Ag^+ ion concentration in the supernatant was
measured by a spectrophotometer (Shimadzu, Japan) based on the
formation of a ternary complex among the Ag^+ ion, phenanthroline, and
eosin Y in acidic aqueous media (pH 6, adjusted by dilute NaOH and
HNO[3] solutions).^[ [207]^75 ^] The absorbance was measured at the
wavelength of 550 nm. The concentration of Ag^+ was calculated based on
a calibration curve prepared by silver ion standard solutions ranging
from 0 to 0.2 mg L^−1. All experiments were carried out in triplicate.
The electrical conductivities (σ) of the hydrogels were determined
using an electrochemical workstation (CHI660e, CH Instruments,
Shanghai, China). The hydrogel samples were prepared as discs with a
bottom area of 66.5 mm^2 and a height of 3 mm. The electrical
conductivities were measured at a frequency of 105 Hz and magnitude of
0.5 V at open circuit voltage in accordance with Equation ([208]2).
[MATH: σ=LR×<
mi>S :MATH]
(2)
where L indicates the height, S indicates the bottom area of the
hydrogel sample, and R denotes its bulk resistance.
To intuitively compare the conductivity differences among the hydrogels
of each group, a parallel circuit was constructed that included an LED
indicator and a test bench for gel samples. The test bench consisted of
1.5 mL EP tubes connected to conductive copper foil onto which the
hydrogel was polymerized using ultraviolet irradiation. Once the
external power supply was activated, the variations in the brightness
of the LED indicated the differences in the electrical conductivities
of the hydrogels. The swelling ratio (SR) of the HD‐Ag hydrogels was
calculated using the mass change between the swollen and dry hydrogel.
Briefly, lyophilized hydrogel samples were immersed (≈30 mg) in PBS
(2 mL) and incubated at 37 °C. At pre‐determined time intervals (0, 1,
6, 12, 24, 36, 48, 60, and 72 h), the samples were removed from the
tubes. The surface buffer was then carefully removed using filter paper
and the swollen samples were weighed. The swelling ratio was calculated
as SR = (W [t] – W [d])/W [d] × 100%, where W [d] represents the weight
of the dry hydrogels and W [t] represents the weight of the swollen
hydrogels at different time points. Three independent samples were
weighed from each group.
Antibacterial Evaluation of the Hydrogels
The antibacterial activities of the HD‐Ag hydrogels were evaluated
using the inhibition zone assay and colony count method. For this
study, MRSA and PAO were cultured in Luria–Bertani (LB) broth (Difco,
Becton Dickinson GmbH, Germany) at 37 °C in an aerobic environment.
Both types of bacteria were quantified by measuring the optical density
at 600 nm with a spectrophotometer, according to the established
standard curve. The antibacterial property of the HD‐Ag hydrogels was
evaluated using the inhibition zone method. Briefly, MRSA or PAO was
sprayed in PBS (1 × 10^8 CFU mL^−1, 50 µL) on 6‐cm‐diameter LB agar
plates. Disc‐shaped hydrogel samples (5 mm in diameter and 0.5 mm
thick) were then placed in the central region of these agar plates.
After 12‐h incubation at 37 °C, the agar plates were photographed, and
the area of the clear zone around the hydrogel samples was measured
using Image J software (National Institute of Health, USA).
The antibacterial activity of the HD‐Ag hydrogels was further
investigated through the colony count assay. In brief, MRSA or PAO
(500 µL of 1 × 10^8 CFU mL^−1) was co‐cultured with HD‐Ag hydrogel
samples (5 mm in diameter and 0.5 mm thick) in a 24‐well plate at 37 °C
under aerobic conditions in LB medium for 12 h. The culture media was
then serially diluted, seeded them on corresponding LB agar plates, and
allowed to grow. Following a further period of growth (12 h), the
plates were photographed, and the antibacterial properties of each
hydrogel sample were assessed by counting the number of colonies
formed.
Cytocompatibility Evaluation of the HD‐Ag Hydrogels
NIH/3T3 and HaCaT cells were selected to study the biocompatibility of
the HD‐Ag hydrogels. Both cell types were purchased from Procell Life
Science&Technology Co., Ltd (Wuhan, China) and cultured following the
same protocol as in the literature.^[ [209]^70 ^] Precursor solutions
(150 µL) from each group were used to fabricate hydrogel disks (8 mm in
diameter and 3 mm in height) through photo‐crosslinking gelation. After
sterilizing the disks in 75% alcohol and equilibrating them in culture
medium, the cells were seeded at a density of 1 × 10^4 cells per well
into 24‐well plates. After an overnight cultivation, the hydrogel disks
were added to each well, ensuring the level of culture medium was
slightly lower than the upper surface of the disk to facilitate
cell‐hydrogel contact. The cells were observed after 1, 3, and 5 days
of incubation. Post incubation, DMEM containing 10% CCK‐8 reagent was
added after removing both the disk and medium from each well. The
absorbance was measured at 450 nm with a microplate reader (Multiscan;
Thermo Scientific, USA) after transferring the medium (100 µL) from
each well into a new 96‐well plate. The tissue culture polystyrene
without the hydrogel served as the control. In addition, the cells were
stained with Calcium AM (4 µm) and EthD‐1 (2 µm) in PBS for 30 min
after their incubation with hydrogels for 3 days, following the
instructions provided in the live/dead viability kit (Life
Technologies, USA). Three random fluorescence images were acquired from
each sample, and this process was repeated for three independent
experiments.
Establishment of Type II Diabetic Mice Model
The protocol of the animal experiment was reviewed and approved by the
Animal Ethics and Welfare Committee (AEWC) of Ningbo University.
Approval from the institutional animal ethics committee was obtained
before conducting the animal experiment (approval number: NBU20230142).
C57BL/J mice (6‐week old, male) were purchased from Beijing Vital River
Laboratory Animal Technology Co., Ltd. and housed in plastic cages with
a 12‐h light/dark cycle in a specific pathogen‐free environment. The
temperature and humidity were maintained at 25 °C and 60%,
respectively. The mice were continuously fed a high‐fat diet (60% of
calories from fat, XTHF60, Jiangsu Xietong Medicine Bioengineering Co.,
Ltd., Jiangsu, China) for 1 month. The mice were rendered diabetic by
injection of streptozotocin (STZ). Specifically, the mice were fasted
for 4 h; and then, intraperitoneally injected with STZ (40 mg kg^−1
dissolved in 50 mm sodium citrate buffer, pH 4.5) for 5 consecutive
days. Glucose solution (5%, 1 mL) was injected intraperitoneally 6 h
after STZ treatment to prevent fatal hypoglycemia. In addition, sucrose
water (10%) was routinely provided. The body weights of the mice were
monitored weekly. Daily blood glucose levels in fasting mice (fasting
starting at 7 a.m., blood drawn between 1 and 3 p.m.) were measured
using a One‐Touch Ultra glucometer (Lifescan Inc.) after STZ
administration. Mice with fasting blood glucose levels above 300 mg dl
^−1 (or 16.7 mm) were identified as diabetic.^[ [210]^76 ^] Diabetes
was maintained for 4 weeks to permit manifestation of disease‐related
changes. During this period, the diabetic mice were given subcutaneous
insulin injections (0.5 U, recombinant human insulin, Beijing Solarbio
Science & Technology Co.) on alternate days to ensure their survival.
Thus far, the type II diabetic animal model was successfully
established.
Creation and Treatment of Bacteria‐Infected Diabetic Wounds
The mice were anesthetized using intraperitoneal injection with a
combination of Zoletil 50 (30 mg kg^−1, Virbac, France) and xylazine
hydrochloride (10 mg kg^−1, Huamu Animal Health Products Co., Ltd.,
China). The mice's dorsal fur was removed with an electric clipper.
Additional hair removal was carried out using a non‐irritating
depilatory cream to fully expose the surgical area. Subsequently, a
circular 10‐mm‐diameter wound was created on the back of each mouse.
Then, a bacterial suspension blend of MRSA (25 µL, 1 × 10^7 CFU mL^−1)
and PAO (25 µL, 1 × 10^7 CFU mL^−1) was sprayed onto the wound and
covered with a TegadermTM film. Four kinds of wound dressings were
prepared: a) Tegaderm film dressing (Blank), b) flexible photovoltaic
MEMS electrode (PM patch), c) HD‐Ag[2] hydrogel, and d) PMH dressing.
Simulated indoor light (2700K, 1000 lux) (Enlitech, Kaohsiung city,
Taiwan) was applied as the illumination for the OPV cells. Mice treated
with PM patch and PMH dressing were exposed to this light for 2 h
daily. All the dressings were replaced, and images of the wounds were
captured every 3 days. At the end of the study, mice were asphyxiation
with CO[2] asphyxiation, and tissue samples were harvested for
subsequent tests.
Histological Analysis
The wound tissues were fixed, dehydrated, paraffin‐embedded, and
sectioned, following the same protocol as in the literature.^[ [211]^77
^] The tissue samples were stained according to the instruction of
hematoxylin & eosin (H&E) and Masson kit (Solarbio Science &
Technology, Beijing, China).
Quantitative Label‐Free Global Proteomic Analysis
A total of 100 mg of general protein was extracted from wound tissue
treated with Tegaderm film (Blank group) and PMH dressing, the
methodology of which followed the previously published work.^[ [212]^78
^] The skin tissue samples were homogenized and centrifuged at 10000 ×
g for 5 min. The supernatant, containing the general protein extract
was then collected, and the protein concentration was measured by BCA
method.^[ [213]^79 ^] Following this, the proteins were denatured at
100 °C for 15 min. Next, the protein sample (100 µg) was diluted in a
solution containing 50 mm ammonium bicarbonate buffer (200 µL). To
reduce any disulfide bonds present within the primary structure of
protein, dithiothreitol (DTT; 5 mm; Solarbio Bioscience & Technology
Co., Ltd; Shanghai China) was added and allowed to react at 37 °C for
2.5 h. This was followed by alkylation with iodoacetamide (IAA, 500 mm)
for 40 min under dark conditions at 37 °C. The resultant mixture was
digested by adding a trypsin solution (25 µL, 1 µg µL^−1,
Sigma‐Aldrich, USA) and incubated at 37 °C for 16 h.
Before conducting a nano‐flow LC‐MS/MS analysis of the digested
peptides, the samples were diluted tenfold with 0.1% formic acid
(Sigma–Aldrich) in an aqueous solution, resulting in an estimated
peptide concentration of 0.2 µg µL^−1. The analysis protocol for
peptide samples was similar to the work of Tang and colleagues.^[
[214]^80 ^] The mass spectrometry proteomics data have been deposited
to the ProteomeXchange Consortium via the iProX partner repository^[
[215]^81 ^] with the dataset identifier PXD044920.
Proteomic analysis was performed using the freely available R
language‐based platform Wu Kong
([216]https://wkomics.omicsolution.com/wkomics/main/). Protein
intensities from samples in the Blank and PMH dressing groups were
extracted from Proteome Discoverer result files to represent their
respective expression levels. Proteins with more than 50% missing LFQ
intensity across all samples were filtered out, and a normalization to
the median along with a k‐nearest neighbors (KNN) imputation was
executed to generate the missing values. For more precise quantitation
results, proteins exhibiting an intensity coefficient of variation
larger than 0.3 were further filtered. A 4065 × 6 protein expression
matrix was then generated for subsequent statistical analysis. An
unpaired Student's t‐test was performed to identify differentially
expressed proteins (DEPs) between the Blank and PMH dressing groups,
using a Benjamini–Hochberg (BH) adjusted p value < 0.05 and a fold
change > 2.0 or < 0.5 for upregulation or downregulation, respectively.
The biological function of the DEPs was analyzed through GO, KEGG, and
Reactome pathway‐enrichment analysis.^[ [217]^82 ^] In addition, a
comparison between DEPs and an extracellular matrix protein dataset
(KW‐0272) downloaded from the Uniprot database
([218]https://www.uniprot.org/) identified overlapping proteins. These
results were illustrated using a Venn diagram.^[ [219]^83 ^] Further,
an interaction network comprising co‐expression, predicted genetic
interactions, shared protein domains, physical interactions, and
co‐localization among those overlapping proteins, was constructed
through GeneMANIA ([220]https://genemania.org/).
Immunofluorescence Staining
Tissue slices were individually stained with primary antibodies
(anti‐Col‐I, anti‐CD31, anti‐iNOS, anti‐TGF‐β1, anti‐MMP‐8, and
anti‐MMP‐9, all obtained from Proteintech, Wuhan, China). After
thorough rinsing, the slices were treated with secondary antibodies
labeled with FITC (green fluorescence) or TRITC (red fluorescence) for
color visualization. The nuclei were stained with a
4′,6‐diamidino‐2‐phenylindole (DAPI) containing mounting solution.
Slides were then observed under an inverted fluorescence microscope
(IX53, Olympus).
Real‐Time Quantitative Polymerase Chain Reaction (RT‐qPCR)
Tissue samples were crushed and ground at low temperatures. The
protocol for total RNA extraction and reverse transcription was the
same as in other literature,^[ [221]^84 ^] where total RNA (2 µg) was
added to a 40 µL reverse transcription reaction system. Relative mRNA
expression levels were detected by RT‐qPCR on the LightCycler 96
instrument (Roche Diagnostics, USA). The primer sequences for RT‐qPCR,
obtained from the Shanghai Shenggong Biotech Co., Ltd, are as follows:
TNF‐α (Fwd: CCTGTAGCCCACGTCGTAG, Rev: GGGAGTAGACAAGGTACAACCC), IL‐1β
(Fwd: GCAACTGTTCCTGAACTCAACT, Rev: ATCTTTTGGGGTCCGTCAACT), TGF‐β1 (Fwd:
CCACCTGCAAGACCATCGAC, Rev: CTGGCGAGCCTTAGTTTGGAC), and IL‐10 (Fwd:
GCTCTTACTGACTGGCATGAG, Rev: CGCAGCTCTAGGAGCATGTG).
Statistical Analysis
All quantitative data were presented as mean ± standard deviation (SD).
For normally distributed data sets with equal variances, one‐way ANOVA
testing followed by a Tukey post‐hoc test was carried out across
groups. Statistical analysis was carried out using GraphPad Prism
Software (GraphPad Prism, CA, USA). In all cases, significance was
defined as p < 0.05. The sample sizes (n) were no less than three.
Conflict of Interest
The authors declare no conflict of interest.
Supporting information
Supporting Information
[222]ADVS-11-2307746-s001.pdf^ (1.9MB, pdf)
Acknowledgements