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Video Article
Murine Model of Wound Healing
1,2 1,2 1,2 1,2 1,2,3 1,2
Louise Dunn , Hamish C. G Prosser , Joanne T. M. Tan , Laura Z. Vanags , Martin K. C. Ng , Christina A. Bursill
1
The Heart Research Institute
2
Sydney Medical School, University of Sydney
3
Cardiology Department, Royal Prince Alfred Hospital

Correspondence to: Christina A. Bursill at bursillc@hri.org.au

URL: http://www.jove.com/video/50265
DOI: doi:10.3791/50265
Keywords: Medicine, Issue 75, Anatomy, Physiology, Biomedical Engineering, Surgery, Tissue, Lacerations, Soft Tissue Injuries, Wound Infection,
Wounds, Nonpenetrating, Penetrating, Growth Substances, Angiogenesis Modulating Agents, Wounds and Injuries, Wound healing, mouse,
angiogenesis, diabetes mellitus, splint, surgical techniques, animal model
Date Published: 5/28/2013
Citation: Dunn, L., Prosser, H.C.G., Tan, J.T.M., Vanags, L.Z., Ng, M.K.C., Bursill, C.A. Murine Model of Wound Healing. J. Vis. Exp. (75), e50265,
doi:10.3791/50265 (2013).

Abstract
Wound healing and repair are the most complex biological processes that occur in human life. After injury, multiple biological pathways become
activated. Impaired wound healing, which occurs in diabetic patients for example, can lead to severe unfavorable outcomes such as amputation.
There is, therefore, an increasing impetus to develop novel agents that promote wound repair. The testing of these has been limited to large
animal models such as swine, which are often impractical. Mice represent the ideal preclinical model, as they are economical and amenable
to genetic manipulation, which allows for mechanistic investigation. However, wound healing in a mouse is fundamentally different to that of
humans as it primarily occurs via contraction. Our murine model overcomes this by incorporating a splint around the wound. By splinting the
wound, the repair process is then dependent on epithelialization, cellular proliferation and angiogenesis, which closely mirror the biological
processes of human wound healing. Whilst requiring consistency and care, this murine model does not involve complicated surgical techniques
and allows for the robust testing of promising agents that may, for example, promote angiogenesis or inhibit inflammation. Furthermore, each
mouse acts as its own control as two wounds are prepared, enabling the application of both the test compound and the vehicle control on the
same animal. In conclusion, we demonstrate a practical, easy-to-learn, and robust model of wound healing, which is comparable to that of
humans.

Video Link
The video component of this article can be found at http://www.jove.com/video/50265/

Introduction
Impaired wound healing is responsible for significant morbidity and mortality worldwide; this is particularly true for sufferers of diabetes
1,2 3
mellitus . In humans, wound healing is a continuum of processes, in which there is significant overlapping . Immediately following wounding,
inflammatory processes are initiated. Inflammatory cells release factors that encourage the processes of cell proliferation, migration and
angiogenesis. After re-epithelialization and new tissue formation there is a phase of remodeling that entails both apoptosis and re-organization of
matrix proteins such as collagen.

The complexity of wound healing cannot currently be replicated in vitro and this necessitates the use of animal models. To date, wound-healing
studies have been limited to large animal models, such as swine, to ensure that the healing processes are equivalent and comparable to
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humans. However, using large animals for such studies can be difficult to house and are not always practical . The laboratory mouse represents
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an economical animal model that can be easily genetically manipulated for mechanistic investigation . However, murine wounds heal differently
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to human’s, primarily due to the process of contraction . This is in part, due to an extensive subcutaneous striated muscle layer called the
panniculus carnosus that is largely absent in humans. In mice, this muscle layer allows the skin to move independently of the deeper muscles
and is responsible for the rapid contraction of skin following wounding.
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To overcome this limitation, murine wound healing can be adapted to replicate human wound healing by use of a splint (Figure 1) . In this video
we demonstrate the splinted murine wound model that eliminates wound contraction and more closely approximates the human processes of
re-epithelialization and new tissue formation. In this model two full-thickness excisions that include the panniculus carnosus are created on the
dorsum, one on each side of the midline of the mouse. A silicone splint is placed around the wound with the assistance of adhesive and the splint
then secured with interrupted sutures. Each mouse acts as its own control, with one wound receiving treatment and the other vehicle control,
thereby reducing animal numbers. Following topical applications, a transparent occlusive dressing is applied. The dressing can be removed
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when required for further topical applications and/or measurement of the wound area . At the completion of experiments, wound closure,

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morphological architecture and degree of neovascularization can be assessed by immunohistochemistry. This economical and easy to perform
model can also be utilized to assess wound healing in the context of diabetes mellitus or other pathophysiologies.

Protocol

1. Preparation of Splints and Occlusive Dressings

1. Outline 10 mm circles on 0.5 mm thick silicone sheeting and use scissors or a biopsy punch to create silicone disks.
2. Centre a 5 mm biopsy punch in the middle of the 10 mm circle and press firmly to create a hole to form a "donut"-like disc that will be used as
a splint.
3. Outline 10 mm circles on a transparent occlusive dressing such as Opsite and use scissors to create circular dressings.

2. Experimental Animals
1. Obtain Animal Ethics Committee approval for all experiments that will be performed.
2. Use 8-week old (22-26 grams) male C57BL/6J mice from a commercial breeder (e.g. The Jackson Laboratories).
3. Keep mice in standard conditions of 21 °C and a 12 hr light-dark cycle with free access to food and water.
4. (Optional): Diabetes can be induced in 6 to 7 week old mice by a bolus intraperitoneal injection of 165 mg/kg streptozotocin (in citrate buffer,
pH 4.5), with hyperglycemia confirmed by regular blood glucose monitoring (AccuChek glucometer). Diabetic mice may have polyuria and so
their bedding may need to be changed more frequently to eliminate wetness and their weights should be monitored closely.

3. Anesthesia and Operative Preparation

1. Induce general anesthesia using 5% isoflurane in 100% oxygen (flow rate 1 L/min) and maintain anesthesia using 1-3% isoflurane.
2. Ensure the deep pedal reflexes of the mouse are suppressed and place the mouse in the prone position.
3. Prepare the operative region by removing fur with clippers from the base of the neck to 3 cm further down the back and between the two
shoulder blades.
4. A light application of depilatory cream may be applied for no longer than 2 min. Wet gauze swabs can be used to ensure all cream and
remaining fur is removed.
5. Wipe the skin with an alcohol swab and two applications of 10% povidone-iodine (Betadine) and drape the mouse.

4. Excision and Splinting of Wound

1. Diagrammatic representation of the murine wound-healing model is presented in Figure 1.
2. Use a sterile 4 mm biopsy punch to outline two circular patterns for the wound on either side of the mouse's midline at the level of the
shoulders (see Figures 2a-2b).
3. Use serrated forceps to lift the skin in the middle of the outline and iris scissors to create a full-thickness wound that extends through the
subcutaneous tissue (Figures 2c-2d), including the panniculus carnosus (Figure 2e), and excise the circular piece of tissue.
4. Repeat the process for the wound on the other side of the midline (Figure 2f).
5. Remove plastic protective coating from each side of the silicone splint.
6. Apply cyanoacrylate adhesive (e.g. Super Glue or Krazy Glue) to one side of a silicone splint.
7. Centre the splint over the wound (Figure 2g) and anchor the splint with interrupted 6-0 nylon sutures to ensure positioning (Figure 2h).
8. Repeat the splinting process on the other wound.
9. If required at this time-point, apply the therapeutic compound to be tested (up to 100 μl) to one wound (Figure 2i), and the vehicle control to
the other.
10. Cover the wound with a transparent occlusive dressing (for example, OpSite) (Figure 2j-2k).
11. A ruler is placed below the splints and a photomicrograph taken (Figure 2l).

6. Postoperative Management
1. Carprofen (5 mg/kg) is administered once daily via sub-cutaneous injection for post-operative pain relief.
2. After the surgery animals are individually caged and maintained on heat mats until fully recovered.
3. Monitor animals twice daily for manifestations of pain and weight loss. We observed no gross behavioral displays of pain or weight loss.

7. Wound Measurement and Treatment

1. The wound can be measured daily.
2. Induce general anesthesia using 5% isoflurane gas (flow rate 1 L/min), and then ensure the deep sensory reflexes of the mouse are
suppressed using 1-3% isoflurane.
3. Gently peel back the occlusive dressing with forceps.
4. Use surgical calipers to measure the wound diameter. We take the average of three measurements along the X, Y and Z axis (Figures
2m-2o) as well as a photomicrograph for future reference (Figure 2p).
5. Optional: At this time the animal can be assessed for blood perfusion using a laser Doppler imager (Figure 2q).
6. Re-application of the therapeutic compound and the vehicle control can be performed at this point.
7. A clean transparent occlusive dressing is then re-applied and the animals kept warm until fully recovered.

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8. Note: if splints are not secured properly the wound will readily contract (Figure 2r).

8. Histological Analysis
1. Euthanize the mice after 10 days with an overdose of anesthesia.
2. Using forceps and a scalpel blade remove the sutures and carefully peel away the splint.
3. Use iris scissors to create a wide, full excision around and under the wound area and incubate the tissue in 4% paraformaldehyde in
phosphate buffered saline (PBS) at 4 °C overnight.
4. Transfer tissue to 17% sucrose in PBS solution for a further 24 hr at 4 °C then remove excess solution by dabbing carefully on tissue and
place in O.C.T compound and freeze at -80 °C.
5. Hematoxylin and eosin staining can be used to visualize the wound structure and epithelial gap. Neovascularisation can be assessed by
immunohistochemical analyses to determine the number of capillaries (using Von Willebrand factor).

Representative Results
A wound closure curve is determined by calculating the average diameter of the wound and expressing the results as a percentage, i.e. 100 -
(Day 0 diameter/Day X diameter). In this experiment a therapeutic compound (or vehicle control) was applied daily to the wound. The therapeutic
compound greatly accelerated wound closure (Figure 3). It is important to note that the splints must be maintained for the duration of the
experiment, as removal of splints will lead to rapid wound contraction (Figure 2r) and diverge from the pattern of wound healing observed in
humans.

Figure 1. Schematic representation of the murine wound healing model. In this model two full thickness wounds are created on either side
of the midline allowing each mouse to serve as their own control. Silicone splints are adhered and sutured to the wound perimeter to prevent
wound contraction, providing a model replicable to that of humans.

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Figure 2. Wound healing surgery and post-surgical measurements. Following hair removal and preparation of the skin with iodine and
alcohol (a-b) a biopsy punch is gently used to outline two circles on the dorsum, either side of the midline. (c) A small incision is then created
and (d) a circular piece of skin is removed, (e) including the panniculosus carnosus, (f) to create two full-thickness wounds. (g) Adhesive is
then applied to the silicone splints and the splints adhered to the wound perimeter. (h) Splints are then secured with sutures. (i) Treatments
are topically applied and (j-k) an occlusive, transparent dressing is placed over the wound and adhered to the splints (adhesive can be used if
required). (l) Photomicrographs are taken daily and wound area is calculated from the average of three diameter measurements on the (m) y-
axis, (n) x-axis and (o) z-axis. (p) A representative photo of the wounds at day 10, noting the smaller wound on the right that was treated with a
therapeutic compound. (q) Representative laser Doppler image of blood perfusion of the wound at day 6. (r) Example of rapid wound contraction
following removal of the silicone splints.

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Figure 3. Representative wound closure graph. Wound area is calculated from the average of three daily diameter measurements along the x,
y and z-axes. Wound closure is expressed as a percentage of initial wound area at day 0.

Discussion
This is an experimental murine model of cutaneous wound healing. A significant feature of this model is the use of silicone splints to prevent
wound contraction so that re-epithelialization and new tissue formation may occur, making it similar to the process that occurs in humans. This
model is versatile and can be used to assess wound healing in both physiological and pathophysiological (e.g. diabetes mellitus) settings. The
model may also be used to assess potential wound healing or angiogenesis therapeutics in an economical setting. With each mouse acting as its
own control, animal numbers are minimized. The surgical techniques required for this model are not highly sophisticated, thus this model can be
widely used by those with relatively little surgical experience.

To ensure reproducibility and accurate quantification it is imperative that the splint is adequately adhered and anchored to the skin with sutures,
and that there is minimal delay between creating the two wounds. The propensity of murine wounds to rapidly contract following loosening of the
splint or partial removal due to scratching by the mouse requires daily monitoring of the splints. Careful application of adhesive is also required
to minimize irritation of healthy skin around the splint that may promote scratching. It is also very important to follow aseptic techniques and
thoroughly disinfect equipment, particularly the calipers, between mice. The application of the occlusive dressing must also be considered,
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especially if wounds are not going to be treated or dressed daily. Opsite and Tegaderm (3M) dressings are comparable , and it has been shown
that Tegaderm dressings may only remain adhered for 1-2 days. Should longer-term wound dressings be required an alternative approach has
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been described by Chung and colleagues .

Potential weaknesses of this model may include inflammation due to the anchoring of sutures, the diffusion of the treatment or vehicle between
the wounds and the entry of the treatment into the systemic circulation. In regards to the sutures inducing local inflammation, the sutures are
placed relatively far from the wound and as each wound is created the same, therefore any inflammation that may occur should be similar
between wounds. Similarly, the distance between the wounds and a lack of edema between the wounds would minimize diffusion between the
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two beds. There is some evidence that a treatment may enter the systemic circulation, which may accelerate healing of the control wound .
To determine the extent of a treatment entering the systemic circulation, littermate mice could be utilized, in which the two wounds are only
treated with the vehicle. The differences in rates of wound closure between mice treated with vehicle only, and the mice receiving both vehicle
and treatment could then be compared.

In conclusion, we have demonstrated a relatively simple murine model of wound healing that exhibits many of the features observed in human
wound healing.

Disclosures
The authors have nothing to disclose.

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Acknowledgements
The authors would like to acknowledge funding support from the National Health and Medical Research Council (NHMRC) of Australia (Project
Grant ID: 632512). Louise Dunn was supported by an NHMRC Early Career Fellowship and Christina Bursill by a National Heart Foundation
Career Development Fellowship.

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