A Mouse Model of Mechanotransduction-driven, Human-like Hypertrophic Scarring

Summary

This protocol will explain how to establish a hypertrophic scarring murine model that increases mechanotransduction signaling to simulate human-like scarring. This method involves increasing mechanical tension across a healing incision in a mouse and using a specialized device to create reproducible, excessive scar tissue for detailed histological and bioinformatic analyses.

Abstract

Hypertrophic scarring (HTS) is an abnormal process of wound healing that results in excessive scar tissue formation. Over the past decade, we have demonstrated that mechanotransduction—the conversion of mechanical stimuli into cellular responses—drives excessive fibrotic scar healing. A mouse model to assess human-like hypertrophic scarring would be an essential tool for examining various therapeutics and their ability to reduce scarring and improve healing. Specifically, our laboratory has developed a murine wound model that increases mechanical strain to promote human-like HTS. This protocol utilizes biomechanical loading devices, made from modified 13 mm palatal expanders, whose arms are placed on either side of the incision and distracted incrementally apart in order to apply continuous tension across the wound bed during healing. Over nearly two decades of use, this model has been significantly advanced to improve efficacy and reproducibility. Using the murine HTS model, significant dermal fibrotic scars can be induced to be histologically comparable to human hypertrophic scars. This murine model provides an environment to develop biologics involved in the treatment of HTS and mechanotransduction-related conditions such as foreign body response.

Introduction

Wound healing, the process by which the body attempts to repair damaged tissue and rebuild the skin barrier, can result in atypical healing if its processes of hemostasis, inflammation, proliferation, and remodeling are irregular¹. Hypertrophic scarring (HTS) is an example of irregular wound healing, characterized by excessive deposition of extracellular matrix and connective tissue at the site of injury resulting in the formation of an enlarged scar tissue area^1,2,3. Areas on the body that undergo repeated mechanical stretch stimulations, such as around joints or on the face, are more prone to developing HTS and fibrosis^{4,5,6,7,8,9,10}. We and others have shown that mechanical stretch across a wound bed promotes HTS formation through the activation of mechanotransduction pathways—the conversion of mechanical stimuli into cellular responses^9,11.

HTS not only involves complex biological processes but also carries significant social, medical, and economic challenges for the people affected. Affected individuals can struggle with self-esteem and depression, especially when the scars are in visible areas like the face and hands^1,9,10,12. Scientific review articles indicate that the prevalence of HTS varies between 32% and 72% in the United States^10,13. The severity of these aesthetic concerns, especially in cases of serious burn injuries in the facial region, is underscored by the increasing number of full facial transplantation cases to improve appearance¹⁰. These scars can also cause functional impairments by restricting movement^6,14, and surgical intervention is often required to excise scars and restore mobility¹⁰. The cost of HTS treatment can be substantial, including expenses for surgery, treatments, physical therapy, or even long-term care^1,10. In the United States alone, the annual cost of treating HTS exceeds $4 billion¹⁰.

Considering the pervasiveness of HTS and the extreme measures taken to address its complications, conventional therapies (e.g., surgical excision, corticosteroid injections, and laser therapy) remain highly variable^1,2,15,16,17. While these treatments can offer relief in some cases, they can be insufficient due to the complex nature of scar pathology. Factors such as genetic differences among individuals and an incomplete understanding of the mechanisms driving HTS cause therapeutic strategies to remain clinically unsatisfactory^18,19,20. The future of HTS therapy seems to lie in new innovative approaches that target cell mechanistic drivers of HTS, such as mechanotransduction^11,21, which we have extensively demonstrated to drive excessive fibrotic scar healing^{5,6,7,8,11,21,22,23,24,25}. Specifically, we had previously developed a murine model that increases wound mechanical strain to promote human-like HTS⁹. However, after nearly two decades of use, the model has been significantly advanced to improve efficacy and reproducibility. This protocol will allow researchers to best utilize an updated and optimized HTS mouse model to explore the cell populations and drivers behind excessive scarring. The overall goal of this method is to provide researchers with a protocol designed to produce human-like hypertrophic scarring in mice.

Protocol

Approval from the University of Arizona Institutional Animal Care and Use Committee (IACUC) was obtained for all experiments (control number: 2021-0828). This protocol uses 15-week-old C57BL/6J male mice although it could be applied to other ages and strains^9,26.

1. Creating the HTS biomechanical loading device

NOTE: Modifying the palatal expanders into the HTS device can occur at any point before the experiment.

1. Take the unmodified palatal expander and present to makerspace core or fabrication core with specifications according to Figure 1A,B.
   NOTE: If no makerspace or fabrication core is available, continue to the following steps.
2. Use a Mini Universal Bender (MUB) wire bending tool to alter the palatal expander to conform to the specifications of Figure 1A,B.
3. Place the device flat on a surface as shown in Figure 1A1.
4. Bend the arms 90° up for the upper arms and 90° down for the lower arms when the device lays flat with the MUB as seen in Figure 1A2. See Figure 1A3 for the appearance of the device when resting flat on a surface, where the arms now are perfectly parallel with the body of the device.
5. Bend each arm back into the page 90° along the axis of the dotted line as seen in Figure 1A4. See Figure 1A5 for the device after bending.

2. Hair removal and initial incision on postoperative day 0 (POD 0)

NOTE: Clean and autoclave several sets of surgical instruments before surgery (e.g., dissection scissors, scalpel, Adson forceps, Needle driver). Prepare sterilized 5-0 sutures for use and have a surgical marker on hand.

1. Clean the operating space with 70% ethanol spray.
2. Place a heating pad set at 35 °C on the presurgical prep table. Tape down an absorbent pad over the heating pad. Place another absorbent pad over the previous pad (to collect shaved hair). Tape the anesthesia cone (or cones) on the absorbent pad.
   NOTE: [Optional] If a benchtop fume extractor is available, place the cone near the workspace and turn on the suction.
3. Place two or three mice at a time in the anesthesia induction chamber with 1-3% isoflurane and 2 L/min oxygen flow until the mice breathe calmly (2-3 min).
4. Insert each mouse's nose into the nose cone opening, allowing for inhalation of anesthesia composed of 1-3% isoflurane with 2 L/min of oxygen. Confirm adequate anesthesia by the lack of reaction to a toe pinch. Apply vet ointment to the eyes of the mouse.
5. Use an electric razor to shave the hair on the dorsum in the area as seen in Figure 1C.
6. Apply the depilatory paste with a gloved finger or cotton swab onto the skin, covering the shaved area. After 45 s, wipe away the paste with gauze and then an alcohol swab. Wipe the dorsum with another alcohol swab to remove the depilatory paste, leaving behind a hairless patch (Figure 1C).
   NOTE: It is critical to use a moderate amount of paste, remove the paste promptly, and wipe the mouse after removal of the paste to prevent dermal burns by the depilatory paste. Murine skin is thin, making it susceptible to rapid chemical burns that can have detrimental effects both on mouse health and on experimental quality.
7. Ear tag the mouse for identification.
8. Upon removing the hair, place the mouse back in the cage to prevent excessive isoflurane exposure. Monitor the mouse.
9. Remove the top absorbent pad covered with hair to have a clean presurgical pep work surface.
10. Set up a separate surgical work area. Place a heating pad set at 35 °C on the surgical table. Tape down an absorbent pad over the heating pad. Tape the anesthesia cone (or cones) on the absorbent pad.
11. Place a mouse in the anesthesia induction chamber with 2-4% isoflurane and 2 L/min oxygen flow until the mouse breathes calmly (2-3 min).
12. Take the mouse out of the induction chamber and place it on the operating surface. Insert each mouse's nose into the nose cone opening, allowing for inhalation of anesthesia composed of 1-3% isoflurane with 2 L/min of oxygen. Confirm adequate anesthesia by the lack of reaction to a toe pinch. Apply vet ointment to the eyes of the mouse.
13. Inject 0.05 mg/kg buprenorphine into the shoulder subcutaneously using a 21 G needle for postsurgical pain treatment.
14. Clean the mouse's dorsum with an alcohol swab and let it dry. With the surgical marking pen and a ruler, mark a 2 cm line on the dorsal midline where the full-thickness incision will be made, as shown in Figure 1D.
15. Use a scalpel or dissection scissors (surgical preference) to make the full-thickness dorsal midline incision through the marked area.
    NOTE: Be careful not to cut the underlying tissue (e.g., muscle), as seen in Figure 1D.
16. Using 5-0 sutures in a simple interrupted pattern, close the incision by bisecting the wound as shown in Figure 1D. Use at least 5 evenly spaced sutures.
17. Cut a Telfa gauze into a 3 cm x 1 cm piece. Place it in the center of a foam adhesive dressing and place the adhesive dressing on the mouse dorsum such that the gauze covers the incision as shown in Figure 1D. Place a halved dressing on the abdomen and wrap circumferentially until it meets the dorsal dressing.
18. After the dressing is complete, place the mouse in a separate sterile cage and monitor it until it has fully recovered from the anesthetic.
19. Repeat the procedure with all mice, regardless of the experimental group. Let the incision heal over the next 4 days before the next step.

3. Placement of HTS biomechanical loading device (POD 4)

NOTE: Clean and autoclave the HTS devices and several sets of surgical instruments before surgery (e.g., dissection scissors, scalpel, Adson forceps, Needle driver, skin stapler, skin staples). Prepare sterilized 5-0 sutures for use. [Optional] If a benchtop fume extractor is available, place the cone near workspace and turn on the suction.

1. Prepare the area of surgery as described in steps 2.1 and 2.2.
2. Anesthetize the mouse by following steps 2.11-2.12.
3. Use the forceps or needle driver to separate the dressing from the mouse's abdomen by working the tool side to side against the ventral side. Leaving the tool between the wrap and skin to elevate the dressing off the skin, use scissors to cut the dressing off.
   NOTE: Be careful not to cut the mouse skin, rip the dressing off, or disturb the healing incision.
4. Clean the dorsum with an alcohol swab. Examine the incision for wound dehiscence or signs of infection.
5. Ensure the HTS device is not extended/expanded and is in its most reduced form, as shown in Figure 1B. Lightly coat the arms of the device with medical glue.
6. With one hand, render the skin on the mouse dorsum slightly taut in the transverse direction. With the other hand, place the HTS device on the mouse dorsum such that the incision is equidistant from each arm of the HTS device, thereby centering the device over the incision. Ensure the skin between the arms of the HTS device is uniformly taut. Hold the device in place until the glue has dried (~30 s), as seen in Figure 2A.
   NOTE: Keeping the skin uniformly taut is important to ensure equal amounts of tension are placed on the incision with the device. Leave the sutures intact during this process because the process of placing the device adds mechanical tension to the skin, which can reopen the healing incision. The sutures remain in place until the first day of stretching to ensure that the wound remains closed.
7. Secure four sutures around each arm and through the skin as shown in Figure 2B. While suturing, make sure that the needle exits the skin towards the incision.
   NOTE: Inserting the needle from the incision side of the device can sometimes tear the incision open due to the force necessary to puncture the skin.
8. Now place three skin staples around the arms and through the skin, securing the HTS device to the skin as shown in Figure 2B.
9. Bandage the mouse by following steps 2.17 and 2.18.
10. Repeat the procedure with all mice, regardless of the experimental group.
    NOTE: All mice receive the same preparation; however, they are randomly assigned in each experimental group (e.g., control, stretch) to ensure an unbiased distribution of mice in the experimental design. Specifically, both control and stretch mice will have the device attached to their dorsum. The control mice's devices will remain untouched, while the stretch mice will undergo the next steps.

4. Initial stretch of HTS biomechanical loading device (POD 5)

NOTE: Clean and autoclave several sets of surgical instruments (e.g., dissection scissors, scalpel, Adson forceps, Needle driver) prior to surgery. [Optional] If a benchtop fume extractor is available, place the cone near the workspace and turn on the suction.

1. Randomly assign each mouse via ear tag to the desired group. For control mice, simply remove the sutures on the incision and change the dressings of the wounds (steps 1-6, and then 8-9). If desired, remove the device at the time of stretching and take a photo with a measuring implement for scale to track the scar size.
2. Prepare the surgical area, anesthetize the mouse, and remove the dressing by performing steps 3.1-3.4.
   NOTE: Be careful not to tear the dressing off or disturb the healing incision. If an arm of the HTS device has detached from the skin, lightly glue back into place and add a skin staple or suture depending on the size of the detachment.
3. Remove the sutures from the wound with dissection scissors or other method of choice.
4. Insert the HTS device key into the device and turn to expand the device until the skin is taut but not at risk of tearing the skin, as seen in Figure 2C.
   NOTE: This initial distraction may take around 4-8 full turns of the key since the skin may be loose between the arms of the device.
5. Bandage the mouse by performing steps 2.17 and 2.18.

5. Subsequent stretch of HTS biomechanical loading device (POD 7, 9, 11, 13, 15, 17)

NOTE: Clean and autoclave several sets of surgical instruments (e.g., dissection scissors, scalpel, Adson forceps, Needle driver) prior to surgery. [Optional] If benchtop fume extractor is available, place the cone near the workspace and turn on the suction.

1. For control mice, simply change the dressings of the wounds (steps 1-5, and then steps 13-14). If desired, remove the device at the time of stretching and take a photo of the scar with a measuring implement for scale to track the scar size.
2. Prepare the surgical area, anesthetize the mouse, and remove the dressing by following steps 3.1-3.4.
   NOTE: If an arm of an HTS device has detached from the skin, glue it in place and add a skin staple or suture depending on the size of the detachment.
3. Insert the HTS device key into the device and turn to expand the device until the skin is taut but not at risk of tearing the skin.
   NOTE: This may take approximately 4 turns or ~2 mm of total distraction. If the device has reached maximum extension and cannot be extended further, perform steps 5.4-5.7 to take off and re-attach a new device. Otherwise, bandage the mouse by performing steps 2.17 and 2.18.
4. If the device has reached maximum extension, remove the device from the mouse's dorsum by removing the staples with a staple remover or scissors by prying the prongs of the staples open. Then, remove the sutures and carefully remove the device.
5. Clean the device with a scalpel, alcohol swabs, and paper towels. Soak the device in 70% ethanol for ~20 min to aid in cleaning. Then, use the key to contract the device to its thinnest form.
6. Reattach the HTS device following steps 3.5-3.8.
7. Bandage the mouse by following steps 2.17 and 2.18.

6. Harvesting the HTS tissue (POD 19)

NOTE: Harvesting tissue can take place at any point in the process. We have harvested tissue after only 4 days of stretch to examine early time points; however, tissue is most consistently harvested at POD 19 (2 weeks after strain was initiated). Clean and autoclave several sets of surgical instruments (e.g., dissection scissors, scalpel, Adson forceps) prior to surgery. To get photos of the scar over time, the device can be removed before each stretching step to take a photo of the scar before re-applying the device and re-initiating mechanical strain. [Optional] If a benchtop fume extractor is available, place the cone near the workspace and turn on the suction. The benchtop fume extractor may be turned off when the isoflurane gas is no longer being used.

1. Prepare the surgical area, anesthetize the mouse, and remove the dressing by following steps 3.1-3.4.
2. Sacrifice the mouse via cervical dislocation.
   NOTE: Take care not to pull the skin and tear the mouse's dorsum.
3. Use dissecting scissors or a skin staple remover to remove the skin staples. Cut off the sutures. Gently remove the HTS device, taking care not to tear the skin.
4. Using either a scalpel or scissors, cut the skin surrounding the HTS scar. Preserve the skin for histological analysis in the desired fashion.
   NOTE: If the skin is to be used for transcriptomic or protein analysis (e.g., qPCR, western blot, single-cell analysis), be sure to excise just the HTS scar tissue to minimize the amount of surrounding healthy tissue. This will ensure that the analysis will only capture the scar tissue.
5. Scrape the HTS devices clean with a scalpel. Place devices in a beaker with 70% ethanol to soften any remaining adhesive or tissue.
   NOTE: After soaking in 70% ethanol, the devices may require further wiping, scraping, and cleaning before autoclaving.

7. Measuring average scar width

NOTE: This was accomplished with image analysis software ImageJ, and the information was recorded on a spreadsheet.

1. Open images in ImageJ. Trace the edges of the scar with the polygon tool. Click analyze | measure to measure this area.
   NOTE: The scar can be identified by its discoloration and lack of hair follicles.
2. Measure the length of the scar from end to end using the segment tool. Click analyze | measure to measure this length.
3. Measure the length of 1 cm or any other standardized unit of length in the picture, using the segment tool. Click analyze | measure to measure this length.
4. Using the spreadsheet, take the Area of the scar (in pixels; px) and divide that by the Length of the scar (px length). That result gives the average scar width in pixels.
5. Divide that result by the measured standard unit of length (px length). The result will be the average scar width in the unit of the standard unit of length (e.g., cm) used for the experiment.

Results

To clearly demonstrate the effective use of the HTS protocol and identify successful "positive" results, the model was established as shown in Figure 3A. In the representative study, there were two groups: No Stretch Control (n = 6) and Mechanical Stretch HTS group (n = 6) where human-like levels of mechanical strain were induced across the incision to generate an HTS, seen in Figure 3B,C. Within the experimental plan given in

Discussion

The HTS mouse model is a cost-effective and highly reproducible method for inducing HTS via mechanotransduction and developing potential therapies. While there is an initial learning curve to effectively use the model, the protocol can, with practice, be performed by any researcher without surgical training. Using this model allows researchers to better understand HTS formation and the role of mechanotransduction in wound healing, which may lead to tangible improvements in patient wound care. The video demonstration acco...

Materials

Name	Company	Catalog Number	Comments
100 mL PYREX Griffin beaker	Milipore Signma	CLS1000100
Aesculap Exacta mini trimmer	Aesculap
AutoClip System	Fine Surgical Instruments	12020-00
BD brand isopropyl alcohol swabs	Fisher Scientific	13-680-63
Buprenorphine SR (0.5 mg/mL)	Buprenex, Indivior Inc.	12496-0757-1
C57/BL6 females (6–8 weeks old)	The Jackson Laboratory	000664
Covidien sterile gauze	Fisher Scientific	2187
Covidien TelfaTM non-adherent pads	Fisher Scientific, Covidien	1961
Dental surgical ruler	DoWell Dental Products	S1070
Depilatory cream (Nair Hair Remover Lotion)	Church&Dwight, CVS	339823
Ethanol 70% solution	Fisher Scientific	64-17-5
Excel	Microsoft Cooperation	Microsoft.com	software program
ImageJ	ImageJ, Wayne Rasband	imagej.net	software program
Inhalation anesthesia system	VetEquip	922130
Iris scissors 4½ in. stainless	McKesson	43-2-104
Isoflurane, USP	Dechra Veterinary Products	17033-094-25
Kaka industrial MUB-1	Kaka Industrial	173207	Only necessary if there is no maker space or fabrication shop available
Leone Rapid Palatal Expander- 13 mm	Great Lakes Dental Technologies	125-004	The key necessary to expand and cotnract the device will come with this product in the box
Liquid repellent drape 75 x 90 cm with adhesive hole 6 x 9 cm	Omnia S.p.A.	12.T4362
Medequip Depot Silk Black Braided Sutr 6-0 Rx	Medequip Depot D707N, Fisher Scientific	NCO835822
Needle holder 5 in. with serrated jaws	McKesson	43-2-842
Prism 9	GraphPad Holdings, LLC	graphpad.com	software program
Puralube ophthalmic ointment	Dechra, NDC	17033-211-38
R studio Desktop	RStudio PBC	rstudio.com	software program
Surgical skin marker	McKesson	19-1451_BX
Tegaderm, 3 M	VWR	56222-191	foam adhesive dressing
Thermo-peep heating pad	K&H, Amazon
Tissue forceps 4¾ in. stainless 1 x 2 teeth	Mckesson	43-2-775
Vetbond (3 M)	Saint Paul, MN	1469SB