Flexible Electromagnetic Coil Patch-enhanced Healing of Full-Thickness Skin Wounds in Rats

Summary

This protocol demonstrates a method for creating full-thickness circular skin wounds in rats and enhancingtheirhealingusing a flexible copper coilpatch that delivers apulsedelectromagnetic field.

Abstract

Chronic and large skin wounds present significant clinical challenges due to delayed healing and high infection risk. Pulsed electromagnetic field (PEMF) therapy is a non-invasive approach that can promote tissue repair. This study describes a reproducible rat wound model protocol using a flexible printed copper coil patch connected to a custom PCB controller to deliver localized PEMF stimulation to full-thickness dorsal wounds. The contralateral wound served as an untreated control. Healing progression was assessed through regular photography and wound area measurements. The protocol provides a practical, adaptable system for evaluating and optimizing PEMF-based therapies in vivo across various conditions, with a focus on localized application and parameter adjustment for regenerative medicine. Parameter selection was based on an orthogonal CCK-8 screen of fibroblast proliferation that tested 0.25-1.3 mT, 10-40 Hz, and 30-90 min/day; main-effect analysis identified 1.3 mT, 30 Hz, and 60 min/day as the optimal levels, which were adopted for in vivo testing.

Introduction

Chronic cutaneous wounds (such as diabetic ulcers) are a growing health concern, often leading to impaired healing, infection, or amputation¹. Traditional treatments can be prolonged and costly, prompting interest in adjunctive therapies that can speed up tissue repair². Electromagnetic field therapy is one such approach, which is non-invasive and has demonstrated pro-healing effects in both preclinical and clinical settings^2,3,4,5. Pulsed electromagnetic fields (PEMFs) in the extremely low-frequency range (~1-100 Hz) can stimulate cellular processes important for repair, including angiogenesis and growth factor release^2,6,7.

For example, PEMF exposure significantly accelerated wound closure in diabetic and healthy rodents by upregulating fibroblast growth factor-2 (FGF-2) and increasing angiogenesis⁸. Similarly, in the impaired healing model (e.g., diabetic rats), PEMF treatment has been shown to improve healing rates and increase the tensile strength of newly formed tissue⁹. Clinically, PEMF therapy has been explored in patients with chronic wounds. A pilot randomized trial in diabetic foot ulcer patients reported the enhanced microcirculatory blood flow and a trend toward faster wound closure under PEMF exposure¹⁰. In general, PEMF has also shown beneficial effects in other wound types, for instance, improved healing was observed in burn wounds treated with pulsed magnetic fields¹¹. These studies collectively underscore the potential of electromagnetic stimulation as a complementary therapy for wound care^8,9,10,11.

Compared with conventional wired stimulators, magnetic actuation eliminates the need for transcutaneous connectors and reduces the risk of hardware-related complications. Wireless magnetic systems have achieved tissue-specific stimulation in other contexts, for example, a conductive nerve guidance conduit can generate induced microcurrents under an alternating magnetic field, with the current amplitude indirectly determined by the field parameters¹². Magnetic strategies can also support vascularization -- a critical bottleneck in skin healing -- because static or dynamic magnetic fields, alone or in combination with magnetically responsive biomaterials, have been shown to modulate endothelial behavior and angiogenic signaling¹³. These precedents motivated our development of a wearable approach: a conformal, removable coil patch that concentrates PEMF at the wound bed while preserving non-invasiveness and external programmability^12,13. In the following protocol, we detail the steps for implementing this PEMF therapy in a rat full-thickness wound model, including device application, stimulation regimen, and outcome assessment.

Protocol

All animal procedures were approved by the Institutional Animal Care and Use Committee (IACUC) of Jiangnan University (JN. No2025-0630S0360-915[408]). We implemented the protocol using male SD rats (6-8 weeks old, approximately 200-250 g). Prior to experimentation, rats were individually housed with free access to food and water, maintained on a 12 h light/dark cycle, in accordance with the Guide for the Care and Use of Laboratory Animals.

1. Creation of full-thickness excisional wounds

1. Prepare the animal and surgical field.
   1. Induce anesthesia with 3-4% isoflurane in an induction chamber. After loss of the righting reflex, maintain anesthesia at ~2% via nose cone.
   2. Place the rat in the prone position on a sterile surgical surface. Shave the dorsal thoracic region (shoulder blades to mid-lumbar area, including flanks) using electric clippers.
   3. NOTE Remove fur from at least a 5 × 5 cm area on each side of the midline to allow clear surgical access and coil placement.
   4. Disinfect the shaved area with 70% ethanol followed by iodophor solution. Repeat the ethanol/iodine scrub three times, ending with iodine.
   5. Drape the surgical field with sterile gauze, leaving only the planned wound sites exposed.
2. Create bilateral full-thickness excisional wounds.
   1. Outline two 2 cm diameter circles on the dorsal flanks using a sterile surgical marker, positioning one on each side of the midline behind the forelimbs.
      NOTE: Ensure both circles are equal in size and symmetrically positioned to enable within-animal comparison^14,15.
   2. Excise the marked circular skin area using sterile scissors or a scalpel. Remove the epidermis, dermis, and panniculus carnosus to create a full-thickness wound (~3.14 cm²).
   3. Achieve hemostasis by applying gentle pressure with sterile gauze. Create an identical 2 cm wound on the contralateral flank using the same technique.
      ​NOTE: The wound bed should consist of exposed fascia or muscle with no residual dermis.
   4. Administer postoperative analgesia immediately after wound creation (e.g., lidocaine 4 mg/kg subcutaneously).
3. Assign experimental groups.
   1. Use a within-subject design in which one wound receives PEMF treatment and the contralateral wound serves as a control.
   2. Include a separate cohort receiving a sham device (identical non-powered coil) to control for device presence.
   3. Maintain untreated or sham-treated controls to distinguish electromagnetic effects from handling or dressing effects.

2. Application of the PEMF device

1. Dress the wound and position the coil.
   1. Place sterile thin gauze or semipermeable film directly over the wound designated for PEMF treatment. Ensure complete coverage of the wound bed.
   2. Position the flexible copper coil patch over the dressed wound (Figure 1). Center the coil so that the wound lies beneath the middle of the coil.
      NOTE: The coil diameter (~5 cm) may extend beyond the wound margins; ensure the wound remains within the effective magnetic field zone.
2. Secure the coil.
   1. Secure the coil using medical-grade adhesive tape wrapped gently around the torso. Maintain close contact between the coil and dressing without compressing the thorax.
   2. Place a flexible aluminum strip over the coil to stabilize positioning if needed. Avoid restricting respiration or local blood flow.
   3. Route lead wires away from the body (e.g., toward the head) to prevent entanglement.
3. Administer PEMF stimulation.
   1. Connect the coil to the PCB controller (Figure 2) and activate the device. Set stimulation parameters to 30 Hz frequency and ~1.3 mT magnetic field strength for 60 min/day.
   2. Perform stimulation sessions at the same time each day (±1 h) to maintain consistent intervals.
      NOTE: These parameters were selected based on orthogonal CCK-8 screening (0.25-1.3 mT, 10-40 Hz, 30-90 min/day, Table 1) identifying 1.3 mT, 30 Hz, and 60 min/day as optimal (Table 2).
   3. Place an identical inactive coil over the control wound for 1 h to provide sham treatment. Clearly mark the sham coil to avoid confusion.
4. Remove the device and inspect the wound.
   1. Turn off the controller after 60 min. Remove the coil and adhesive materials carefully.
   2. Inspect the wound for bleeding, edema, or adverse effects. Replace sterile gauze if necessary.
      ​NOTE: Coil surface temperature remains near ambient during 1 h stimulation; no significant heating was observed.
5. Provide posttreatment care.
   1. Fit an Elizabethan collar to prevent self-trauma. Ensure proper fit without impairing respiration or feeding.
   2. Wrap gauze lightly around the hind limbs to reduce scratching of dorsal wounds.
   3. Monitor the rat until fully recovered from anesthesia. Continue analgesia (e.g., lidocaine 4 mg/kg every 12 h for 2-3 days or per institutional protocol).
   4. Repeat daily PEMF treatments for the study duration (e.g., 14 days).

3. Wound monitoring and data collection

1. Capture wound images.
   1. Photograph wounds at 0, 3, 7, 10, and 14 days post wounding. Lightly anesthetize animals if necessary to prevent movement.
   2. Include a measurement scale in each image and position the camera perpendicular to the wound at a fixed distance^14,15.
2. Measure wound area.
   1. Calibrate image scale in ImageJ using the reference ruler. Trace wound margins to calculate area in mm².
   2. Record wound area at each time point and calculate percent wound closure relative to baseline.
   3. Compare PEMF-treated and control wounds quantitatively.
3. Harvest tissue for histology (optional).
   1. Euthanize animals at the experimental endpoint according to IACUC-approved procedures.
   2. Excise wound tissue samples and fix in formalin for histological processing.
   3. Perform paraffin embedding and staining (e.g., H&E, Masson's trichrome) to assess collagen deposition, re-epithelialization, and angiogenesis^13,16.

4. Data analysis and interpretation

1. Generate wound closure curves.
   1. Plot percent wound closure over time for each group.
   2. Calculate healing rate and time to predefined closure thresholds.
2. Perform statistical analysis.
   1. Compile wound area data for all animals and calculate mean ± SD at each time point.
   2. Confirm normality (Shapiro-Wilk test) and homogeneity of variance (Levene's test).
   3. Analyze differences using two-way ANOVA with factors "Treatment" and "Time." Perform Tukey's post hoc testing if significant interaction is detected.
   4. Define statistical significance as *p < 0.05 and **p < 0.01.

Results

In this full-thickness rat wound model, daily PEMF therapy via a flexible coil patch resulted in significantly faster wound closure compared to controls. To rigorously evaluate the effect, rats were allocated into three groups based on the experimental design outlined in the Protocol section: a within-subject control group (where each rat had one PEMF-treated wound and one contralateral untreated wound, data pooled for analysis), a sham control group (PATCH: wound covered with an unpowered coil), and an untreated control...

Discussion

Critical steps include creating symmetric full-thickness wounds, ensuring the coil patch lies flat over a sterile barrier without gaps, and maintaining consistent daily exposure; deviations can reduce field delivery and increase variability in healing outcomes.

This work has several limitations that should be considered when interpreting the findings and assessing translational potential. First, the protocol was demonstrated in healthy male SD rats (6-8 weeks, 200-250 g) with acute excisional ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
96-well cell culture plates	BaiDi	H908002	For CCK-8 assay
ADHESION MICROSCOPE SLIDES	CITOTESTO	P/N.80312-3161	Equipment for  H&E/Masson staining
Analgesic (Lidocaine Hydrochloride Injection)	HUAMU	70011657	Opioid analgesic, 4 mg/kg (for pain control)
Anesthesia machine	RWD Life Science (Shenzhen, CN)	R520-Iso	Isoflurane vaporizer with oxygen supply
Automated immunohistochemical staining machine	Leica	21.2201	automated immunohistochemical staining
Biopsy punch (20 mm)	Integra Miltex	#33-37	Circular skin punch, 2 cm diameter
Camera, digital	Huawei Mate 70 pro	N/A	≥12 MP for wound imaging
CD31 antibody	Abcam	ab182981	Endothelial marker for IHC (Figure 4B)
CD34 antibody	Abcam	ab81289	Endothelial marker for IHC (Figure 4B)
Cell Counting Kit-8 (CCK-8)	Biosharp	BS350B	Used for in vitro PEMF parameter screening (OD450)
Cell culture dish (10 cm)	BaiDi	H802004	10 cm (100 mm) tissue culture dish for fibroblast culture
Cell culture medium (e.g., DMEM + FBS)	BaiDi	L100-500	For fibroblast culture in CCK-8 screen
CO2 incubator	Thermo	BB150-2TCS-L	For fibroblast culture
Coil, flexible copper	Custom-fabricated (in-house)	N/A	Spiral printed coil (5 cm diam., polyimide)
Controller board (PEMF generator)	Custom-built (in-house design)	N/A	Custom PCB, drives coil at 30 Hz, ~1.3 mT
Electric hair clipper	Wahl or Oster (pet grooming)	Model 9160	Animal grooming clipper for shaving fur
Elizabethan collar (rodent)	MukeBio	MK-EC-01	Prevent wound interference
Embedding Station	Epredia HistoStar	A81000002	Processing for histology (if performed)
face mask	vedeng	V531596	Maintain anesthesia
Fibroblasts (cell line L929)	Procell	CL-0137	Cells used for CCK-8 proliferation screening
Finite element simulation software (e.g., COMSOL Multiphysics)	COMSOL	CM010004	Magnetic field simulation (Figure 5B-C)
Flexible aluminum strip splint	deargo	1592	Support to prevent coil displacement
Forceps	Majestic	01625	For handling skin and dressings during surgery
Hematoxylin & Eosin (H&E) Automatic dyeing and sealing uni	DRS-Prisma-P+FILM	61770249	Consumable for routine histology staining
Image analysis software	National Institutes of Health	Version 1.54	For wound area measurement (e.g., ImageJ)
Image analysis software	National Institutes of Health	Version 1.54	For wound area measurement (e.g., ImageJ)
Induction chamber	RWD Life Science (Shenzhen, CN)	802-00132-00	Isoflurane anesthesia induction
Infrared thermal imaging camera	HIKMICRO	H11	Monitor coil surface temperature (Figure 5A)
Iodine Povidone Disinfectant 10%	HYNAUT	N/A	Skin antiseptic (iodophor)
Isoflurane	RWD Life Science (Shenzhen, CN)	20037015	Inhalation anesthetic (USP grade)
Lidocaine Injection 10%	SANMA	N/A	For postoperative analgesia
Masson's trichrome staining kit	BASO	C250202	Consumable for collagen/ECM visualization
Medical Skin Marker and Ruler	Tondaus	N/A	Include scale in wound photographs
Microplate reader	Thermo	A51119500C	For OD450 measurement in CCK-8 assay
Microtome	Epredia HM 340E	23900670	Sectioning for histology/IHC (if performed)
Polyformaldehyde tissue fixative	share-bio	SB-C009	Fixation of animal tissue specimens
Rat housing cage	FengQiao	CP-JM	Standard rat housing cage for individual housing.
Rats, Sprague Dawley (male, 6–8 weeks)	WeiJingyu SUZHOU	SCXK2024-0004	200–250 g; individually housed; IACUC-approved use
Scalpel	Jinhuan	35W0602	For excisional wound creation
Semipermeable film dressing	ZD	Q20189	Alternative wound barrier
Statistical analysis software (e.g., GraphPad Prism or R)	Software Mackeiv	10.6.9(890)	Two-way ANOVA and post hoc multiple comparisons
Sterile gauze	ZD	3141626	Wound dressing/protective barrier
Sterile surgical drapes	ZD	V271617	To drape the surgical field after skin prep
Surgical scissors	Majestic	01627	For excisional wound creation
Syringes and needles	MINANK	20193141643	For subcutaneous analgesic administration
Whole-slide scanner	3DHISTECH	N/A	Digital slide scanning