Development of a Rat Model of Fasciotomy Treatment for Compartment Syndrome

Abstract

Skeletal muscle injuries are a major cause of disability for military and civilian populations. Compartment syndrome (CS) in skeletal muscle results from an edema-induced increase in intracompartmental pressure (ICP) after primary injury. Untreated ICP will occlude the tissue vasculature, tissue necrosis, and potential loss of limb. The current standard of care for CS is surgical fasciotomy, an incision through the muscle fascia to relieve ICP. Early fasciotomy will preserve the limb, but often leaves patients with long-term scarring and reduced muscle function. Our group previously developed and characterized a rat model of CS to explore the pathophysiology of CS and test new therapies. We present an expansion of this CS model, including the fasciotomy, to better simulate clinical treatment. CS was induced on the hind limb of adult male Lewis rats and fasciotomy was performed 24 h later. Less than 20% of the rats that underwent fasciotomy showed detectable force 4 days after injury, compared with the 75% of rats that underwent CS induction without fasciotomy. Muscles undergoing fasciotomy showed a significant increase in fibrosis and an increased number of macrophages, Pax7⁺ satellite cells, and α-smooth muscle actin⁺ myofibroblasts at 7 days postinjury. These data indicate that the use of fasciotomy in a rat model of CS resulted in injury sequelae that reflect the severity of human clinical disease presentation along with current standard of care.

Impact Statement

Current animal models of skeletal muscle injury struggle to accurately reflect the injury sequelae seen in humans, particularly in rats and mice. These animals also recover faster than humans do. More accurate recapitulation of the injury is needed to better study the injury progression, as well as screen for novel therapies. This research combines an existing model of compartment syndrome with its clinical standard of care (fasciotomy), creating a more accurate rat model of injury, and providing for a better treatment screening tool. These results show how our model leads to a sustained skeletal muscle deficit with increased inflammation.

Introduction

Muscle trauma, due to injury, disease, or aging, is a major contributor to decreased mobility and quality of life. Severe injuries, if left untreated, result in loss of limb or death. In the United States, one in two adults are affected by some degree of musculoskeletal dysfunction, accounting for a cumulative cost of more than $800 billion annually.¹ As the American population ages, these costs are expected to rise, creating an increased need for novel therapies for the maintenance and/or restoration of muscle function.

Skeletal muscle is divided into compartments that are tightly bound and separated by fascia. Compartment syndrome (CS) is a condition in which the skeletal muscle intracompartmental pressure (ICP) increases secondary to edema and/or hemorrhage, resulting in compromised local circulation and tissue ischemia.² If left untreated, the consequent ischemia will result in tissue necrosis and long-term nerve damage.³ CS is a major contributor to long-term morbidity and disability as a result of combat or civilian-related injuries to the extremities.⁴

In the clinic, CS, or even suspected CS, is treated by surgical fasciotomy, through which >90% of the binding fascia is separated, releasing the ICP.⁵ For the fasciotomy to be effective, all affected compartments must be released, over the entire length and depth of the muscle group affected, utilizing as many incisions as required.⁴ The incision site is not surgically closed until the excess pressure resolves passively, with an average closure time of more than 7 days.⁶ During this time, the wound remains open and debridement of visible fibrotic scar tissue is required every 48–72 h, leading to constant rewounding of the tissue.² While this alleviates the ICP and saves the limb, frequent debridement is associated with multiple rounds of acute inflammation that adds a further tissue insult to an already injured patient.

Timing of the fasciotomy is critical to prevent irreversible ischemic damage to the muscle and nerves. If performed within the first 12 h of elevated pressure, 68% of patients will recover full function.² However, if the fasciotomy is performed more than 12 h after reaching a threshold for elevated ICP, only 8% of patients will have normal function restored.² Identifying treatment options that can help preserve muscle function or restore it after fasciotomy is critical, particularly for military members who often have long delays waiting for extraction from the battlefield.

In healthy skeletal muscle, fibroblasts reside between the muscle fibers and serve to produce the extracellular matrix (ECM) that provides mechanical stability, structural integrity, and longitudinal and lateral force transmission.⁷ Fibrosis is the result of excessive ECM production after injury, leading to muscle weakness and decreased range of motion.⁷ After injury, satellite cells, the resident muscle stem cell population, proliferate, expand, differentiate, and fuse to form new myofibers at the site of injury.⁸ If this process is impaired or disrupted, as may happen with severe muscle injury or with aging, excess ECM (primarily collagen) accumulates at the site of injury leading to the formation of fibrotic lesions that will impair muscle function, contraction, and regeneration.⁸

CS, and the consequences of fasciotomy surgery and frequent tissue debridement, results in the development of nonresolved tissue fibrosis. Criswell et al. established a rat model of CS in 2011 to study the pathophysiology of CS and to develop novel cellular, biological, and pharmacological treatments.³ To make the model more applicable to the clinical condition, we developed the injury model and reproduced the surgical fasciotomy in a rat with characterization of the pathophysiology.

Silicone spacers were used to separate incised muscle fibers on either side of the fasciotomy site within the injured muscles of the rats, replicating the surgical intervention in human patients. Silicone is a clinically accepted material that is chemically inert, noncarcinogenic, and nonallergenic, with few inflammatory or fibrotic responses associated with its short-term use.⁹ The insertion of the spacer kept the fasciotomy site “open” in the injured muscles and allowed for debridement of the tissue after its removal. This technique provides a more clinically relevant model of CS and fasciotomy for therapeutic testing.

Methods

CS injury model

All animal experiments were reviewed and approved by the Wake Forest Institutional Animal Care and Use Committee (IACUC) and the Department of Defense Animal Care and Use Review Office (ACURO). Adult (10–12 months old) male Lewis rats (Envigo, Huntingdon, UK; Charles River, Wilmington, MA) were habituated for a minimum of 72 h before use in experiments. CS was induced utilizing a previously established model in which a neonatal size 2 blood pressure cuff (Welch Allyn, Skaneateles Falls, NY) was wrapped around the left hind limb of an anesthetized rat, and inflated to 120–140 mmHg for 3 h.³ Animals were euthanized at 7 or 14 days after CS injury.

To perform the fasciotomy, an incision was made in the skin over the tibialis anterior (TA) muscle 24 h after CS muscle injury. Forceps were used to pull back the skin, followed by a thin incision through the midbelly of the TA. Animals were split into two groups: animals that underwent CS injury with or without fasciotomy.

Spacers were created using a pourable platinum-cured silicone (Smooth-Sil 950; Smooth-On, Macungie, PA) and a custom-made, 3D-printed mold, which casted the spacers at 1.5 × 0.3 × 0.3 cm in size. After casting, the silicone was vacuum degassed, and left to cure for at least 24 h. The spacers were sterilized using ethylene oxide gas before implantation in rats. Silicone type was selected based on its cure time and material hardness, with recommendations that implants should be of shore 50 or 55 A.^9,10 Spacers were placed into the incision site after which the skin was stapled closed. Spacers were removed 3 days after fasciotomy. The experimental time line is outlined in Figure 1A.

FIG. 1.

CS injury, fasciotomy and silicone spacer implantation in the TA muscle. (A) Experimental time line. (B) View of the swollen muscle compartment 24 h after CS induction. The blue silicone spacer is clearly seen implanted in the TA muscle. (C) Masson's trichrome stain of sectioned TA muscle 3 days post-CS induction after removal of the spacer (black arrow). White arrows indicate fibrotic lesions lining the gap. Scale bar, 2 mm. CS, compartment syndrome; TA, tibialis anterior. Color images are available online.

Muscle function

In vivo muscle function was performed as previously described.³ Briefly, a pair of needle electrodes (Grass Technologies; West Warwick, RI) were passed across the peroneal nerve to stimulate contraction of the anterior muscle compartment.³ Twitch, summation, and tetanus were determined using increasing shock frequencies. The resulting isometric torque was measured using custom software (provided by the US Army Institute of Surgical Research, Houston, TX). Muscle function was assessed no more than 7 days before injury for baseline values, and 7 and 14 days post-CS injury.

Histology

Injured and control TA muscles were collected and weighed at the time of sacrifice and preserved in formalin for at least 48 h. Tissues were fixed in paraffin and cut at 8 μm slices using a Leica microtome. Slides were fixed and stained with picrosirius red, hematoxylin and eosin (H&E), and Masson's trichrome.

Immunohistochemistry (IHC) staining was performed using the mouse anti-CD68 (Abcam, Cambridge, UK), mouse anti-α-smooth muscle actin (α-SMA) (Santa Cruz Biotechnology, Dallas, TX), and mouse anti-Pax7 (DSHB, Iowa City, IO) antibodies and diaminobenzidine. The secondary antibody used was horse biotinylated anti-mouse IgG (Vector Labs, Burlingame, CA). Adjacent sections were incubated with the secondary antibody only to demonstrate specificity of the primary antibodies. The amount of fibrosis present in the TA muscle was determined visually with trichrome staining and quantitatively with picrosirius red staining.

Myofibers were manually quantified by a blinded researcher for their regeneration or degeneration status and their cross-sectional area (CSA) based on H&E images using Olympus cellSense software (Olympus Corp, Shinjuku, Tokyo, Japan). Mature myofibers were identified by the absence of central nuclei within cross-sectional images. Regenerating myofibers were counted as those containing one to two central nuclei and degenerating myofibers were counted as those with three or more central nuclei. CSA was performed by manually tracing the border of each myofiber, with a minimum of 1000 cells counted per sample. Cell area was binned into 500 μm² segments and plotted as a histogram.

All images were obtained using an Olympus BX63 upright microscope (Olympus Corp) and an Olympus DP80 camera, or a Leica DM 4000B upright microscope (Leica Microsystems, Wetzlar, Germany) and an Olympus DP73 camera. Data were collected using Olympus cellSens software (Olympus Corp). Images were analyzed using Visiopharm software (Visiopharm, Broomfield, CO) for the IHC staining, or ImageJ software (NIH, Bethesda, MD) threshold analysis for quantification of fibrosis.

Fibrosis analysis

Representative whole-tissue slices were taken at 2 × magnification for macroscopic qualitative observation. Representative random images of the tissue were taken at 10 × magnification for fibrosis analysis. Tissue fibrosis was determined using the threshold analysis of the single-color component separation of the images of ImageJ software (NIH). Percent fibrosis was calculated based on the number of pixels from the threshold analysis/total tissue area. A minimum of 10 images were analyzed per muscle. All images were taken and analyzed by the same investigator, with blinding to the treatment groups.

Statistical analysis

All functional and histological comparisons were made using GraphPad Prism Software (GraphPad Software, San Diego, CA) and Microsoft Excel (Redmond, WA). Two-way analysis of variance (ANOVA) was used to determine changes in skeletal muscle function with different treatments over time. One-way ANOVA or Student's t-test was used to determine the significance for fibrosis and IHC analyses. All data are graphed as ± standard error of the mean. The alpha for these data was set at p < 0.05.

Experiment

Skeletal muscle function after CS injury

Skeletal muscle function was measured 7 and 14 days after injury. Compared with baseline muscle function (uninjured), both the CS and the CS with fasciotomy groups showed significantly decreased muscle function at 7 (Fig. 2A) and 14 (Fig. 2B) days after injury, with no significant differences observed between these groups at either time point. Although all rats had measurable muscle function by day 14, not all rats showed measurable function at day 7 postinjury. In the CS group, 75% of the rats had detectable function 7 days postinjury (Table 1). In contrast, only 18.75% of the muscles had detectable function from rats treated with fasciotomy after CS injury. These data suggested that the addition of the fasciotomy procedure resulted in increased severity of the muscle injury.

FIG. 2.

Isometric force production from muscles with or without fasciotomy at 7 and 14 days after injury. Muscle function was assessed at (A) 7 days and (B) 14 days after injury. All animals were assessed for baseline function. CS n = 4 rats. CS+Fasc n = 8 rats. No significance was seen at either time point between treatment groups.

Table 1.

CS with Fasciotomy Reduces the Ability to Record Evoked Contraction

	Baseline (uninjured)	CS (day7)	CS+Fasc (day7)
Percent of rats with detectable force	100	75	18.75

CS, compartment syndrome.

Anatomical changes in skeletal muscle after CS injury

Changes in tissue fibrosis were assessed visually using sections stained with Masson's trichrome and quantitated using ImageJ from sections stained with picrosirius red (data not shown). Gross morphological tissue damage and fibrosis are evident in Figure 1C and shown in more detail in Figure 3A–F. As seen in Figure 1C, the incision site is ∼2.4 mm wide and ∼3.6 mm deep. A visual assessment of the tissue stained with Masson's trichrome showed significant fibrosis on the edges of the incision site (Fig. 1C, white arrows), as well as fibrotic lesions present at the site of incision (Fig. 1C, black arrow).

FIG. 3.

Fibrosis in injured muscles with or without fasciotomy. Masson's trichrome stain was used to visualize fibrotic depositions (blue/purple) in injured muscles. (A, C, E) Scale bar, 500 μm. (B, D, F) Scale bar, 100 μm. (G) The percent fibrosis was calculated using picrosirius red-stained sections and ImageJ software as previously described.¹¹ All injured muscles, except for CS (D14), had significantly higher levels of fibrosis than uninjured. Uninjured control n = 26. CS (D7) n = 3 and CS (D14) n = 4 rats. CS+Fasc (D7) n = 8 and CS+Fasc (D14) n = 8 rats, *indicates p < 0.001. Color images are available online.

Increased collagen deposition can be seen in both the CS injury (Fig. 3C, D) and CS injury with fasciotomy (Fig. 3E, F) groups, compared with the uninjured control (Fig. 3A, B). The data in Figure 3 were quantified and plotted. Injured muscles treated with fasciotomy showed a significantly higher area of persistent fibrosis compared with CS injury alone at 14 days after injury (Fig. 3G).

Cellular and inflammatory responses after CS injury

An increase in Pax7 expression by activated muscle satellite cells is used to mark the early stage of skeletal muscle regeneration.¹¹ The number of Pax7⁺ cells was quantified in injured muscles with or without fasciotomy, showing a significant increase in these cells 14 days after CS injury in TA muscles that received fasciotomy (Fig. 4A, B). These results display uninjured control tissue with Pax7⁺ on the surface of mature myofibers. In contrast, muscles from both injured groups displayed an increased number of Pax7⁺ cells interspersed throughout the tissue, with Pax7⁺ cells also seen at areas of myofiber degradation in CS+Fasc muscle, as determined by the number of centrally located nuclei (>2 per myofiber cross section) and indicated by the black arrows.

FIG. 4.

Quantification of Pax7⁺ cells in injured muscles with or without fasciotomy. Sections were stained for Pax7 (A) and imaged for quantification (B). Scale bar, 100 or 50 μm as indicated. Black arrows indicate Pax7⁺ satellite cells. A minimum of 30 images per slide were used in the analysis. CS n = 4 rats. CS+Fasc n = 8 rats.*p < 0.0001, CS (D14) versus CS+Fasc (D14). Color images are available online.

Macrophages play a vital role in tissue recovery after injury through the removal of necrotic tissue and the promotion of regeneration.¹² The presence of macrophages in a tissue can provide insight on the stage of recovery after injury; moreover, macrophages stimulate fibroblasts to produce collagen during the myoblast fusion stage.¹³ The number of total macrophages, as determined by CD68 staining, in the injured muscles was markedly increased at 14 days postinjury in the muscles with fasciotomy compared with CS alone (Fig. 5A, B). Intact mature myofibers are seen in the uninjured tissue, along with little to no CD68⁺ macrophages residing between the myofibers (Fig. 5A). At 14 days after injury, these cell populations were clearly detected by CD68 IHC, with cells infiltrated between myofibers at sites of injury.

FIG. 5.

Quantification of CD68⁺ cells in injured muscles with or without fasciotomy. Sections were stained for CD68 (A) and imaged for quantification (B). Scale bar, 100 or 50 μm as indicated. Black arrows indicate CD68⁺ macrophages. A minimum of 30 images per slide were used in the analysis. CS n = 4 rats. CS+Fasc n = 8 rats. *p < 0.0001, CS (D14) versus CS+Fasc (D14). Color images are available online.

Myofibroblasts are the tissue-resident cell population responsible for deposition of the ECM and are detected in tissue undergoing remodeling due to injury.³ A significant increase in α-SMA⁺ myofibroblasts was found in the injured muscles with fasciotomy compared with injured muscles alone (Fig. 6A, B) at 14 days after injury. These cells are detected on the periphery of blood vessels (black arrow) and in the endomysial space between myofibers.

FIG. 6.

Quantification of α-SMA⁺ cells in injured muscles with or without fasciotomy. Sections were stained for α-SMA (A) and imaged for quantification (B). Scale bar, 100 or 50 μm as indicated. Black arrows indicate α-SMA⁺ myofibroblasts. A minimum of 30 images per slide were used in the analysis. CS n = 4 rats. CS+Fasc n = 8 rats. *p < 0.0001, CS (D14) versus CS+Fasc (D14). α-SMA, α-smooth muscle actin. Color images are available online.

After injury, macrophages degrade damaged myofibers, while proliferating satellite cells fuse together to form new cells at the location of injury.^13,14 Areas of degeneration and regeneration can be seen within the damaged tissue bed (Fig. 7A) as indicated in the H&E stain by black and white arrows, respectively.

FIG. 7.

Regeneration, degeneration, and area of myofibers in injured muscles with or without fasciotomy. H&E stain was used to visually assess and quantify injury within the myofibers 14 days post-CS (A). White arrows indicate regenerating myofibers, as identified by one to two centrally located nuclei. Black arrows indicate representative degenerating myofibers as identified by three or more centrally located nuclei. Scale bar, 50 μm. The number of regenerating myofibers (B) and degenerating myofibers (C) was quantified as a percent of the total myofibers within each data set. Myofiber CSA was quantified and expressed as a histogram with at least 1000 myofibers counted per sample (D). The data expressed herein were not significantly different. CSA, cross-sectional area; H&E, hematoxylin and eosin. Color images are available online.

Both the CS group and the CS+Fasc group at day 14 show increased numbers of these cell populations compared with the uninjured tissue. Quantification of these data via the numbers of internally located nuclei reveals a trend of increasing numbers of regenerating and degenerating myofibers with the addition of the fasciotomy, although this is not statistically significant (Fig. 7B, C). The overall size distribution of these myofibers shifts to the left of the curve (Fig. 7D) after injury compared with the larger, right-shifted healthy myofibers (Fig. 7A, D) indicating increased smaller myofibers present.

Discussion

Acute CS muscle injury is unique in that it occurs as a result of a primary injury such as a gunshot blast, contusion, or burn. The fascia that binds the muscle compartments prevents swelling after injury, which leads to increased ICP, occlusion of blood vessels, and finally, tissue necrosis. The only effective therapy for the treatment of acute CS is a fasciotomy surgery accompanied by frequent debridement of the necrotic tissue.⁴ Unlike other models of muscle injury, such as volumetric muscle loss or cardiotoxin injury, CS relies on the swelling within the muscle compartments to cause tissue ischemia leading to tissue death.² Therefore, the pathology of CS is unique compared with other types of muscle injury, and animal models of CS need to replicate the ischemia to mimic the human progression of disease.

There are other research models of CS in use, but we believe that our model is a better fit for early preclinical studies. The pig model of CS is accurate to human sequelae, although it is more expensive and time-consuming when compared with smaller animal models.¹⁵ Multiple groups utilizing pig models of CS have incorporated fasciotomy into the model, which is easier to perform in the larger muscle compartments of the swine.^5,16 In addition to swine, successful ischemia/reperfusion injuries were created in rabbits to model CS ranging from a single time point injury induction to multiple ischemic periods per day.^17–19

Rodent models of CS offer a more affordable injury model, but come with the distinct disadvantage of rapid recovery after injury dissimilar to the recovery time in humans. We postulate that there is a need for a more accurate preclinical small animal model of CS, which better mimics the human condition. Thus, we, along with several other groups, have created similar models of CS that utilize pneumatic pumps and inflatable cuffs, similarly to what is described here.^3,20–23 These groups have successfully used these models to investigate experimental treatment for CS, however, these models leave out the current standard of care.

By including the fasciotomy surgery in rats and simulating the delay in wound site closure seen in the clinic, we showed a delay in regeneration in our rat model of CS, which makes it a more useful screening tool for potential new therapies and to gain a better understanding of the sequelae of CS.

We sought to account for the additional tissue injury post-CS caused by the fasciotomy and tissue debridement in patients by creating a similar injury in our rat model. However, it is important to note that our fasciotomy procedure does differ from the surgical procedure performed on patients. First, it is very difficult to ascertain the specific fascicles within injured rat muscles due to their small size. Therefore, our “fasciotomy” procedure in the rats does not separate the muscle compartments, as what would be done for patients, but instead an incision is made in the belly of the TA muscle at the time of maximal swelling (∼24 h postinjury).

Second, it is not possible to keep a wound on a rat “open” for multiple days with debridement of the necrotic tissue until pressure resolves passively. The increased pressure takes an average of 7 days to completely resolve in human patients, which we sought to mimic here.²⁴ Preliminary experiments using the fasciotomy surgery in the rat found that the incised muscle rapidly closed and healed (data not shown), unlike what would occur in patients.

Therefore, silicone spacers were used in the site of the fasciotomy to keep the surgical site open with skin closure. The use of the spacer simulated the extended time that the fasciotomy surgery is left open in the clinic, which is typically at least 1 week.^6,24 Many types of silicone have been shown to have excellent biocompatibility, allowing us to study the effect of the open surgery site, without the confounding effect of a noninert material being introduced to the site.¹⁰

We predicted that the addition of the fasciotomy with the silicone spacers would result in significant and sustained reductions in force compared with CS alone. Initial observations of muscle functional recovery when we paired the fasciotomy surgery with the CS injury seemed to show no differences between injured muscles with or without fasciotomy, up to 2 weeks postinjury (Fig. 2A, B). While our changes to the injury model resulted in no significant differences in overall force production, we did see changes in the number of rats with detectable function on day 7 (Table 1).

Historically, we have detected slight variation in the severity of injury at early time points and occasionally have had rats with no detectable function 7 days postinjury. By incorporating the fasciotomy into our model, we see a more severe injury in the majority of animals, which better mimics the severity of the injury in humans. Therefore, this model is a better reflection of clinical CS, where only 68% of patients will regain full function of their limb after CS injury, even when a fasciotomy is performed within the first 12 h of elevated ICP.² Long-term effects on muscle function recovery after CS and fasciotomy in the rat model still need to be evaluated to determine if there is a change in full functional recovery in these animals.

The data presented herein did demonstrate cellular and morphological differences between the CS alone and CS and fasciotomy models. Chronic unresolved fibrotic deposition results in significant scar tissue formation, requiring frequent debridement before closure of the fasciotomy site. Our model accurately mimics this increase in skeletal muscle fibrosis (Fig. 3). These data agree with our previously published results in which the majority of fibrosis is resolved in adult rodents at 2 weeks post-CS injury.²⁵

Based on our previously published data, adult rats will rapidly develop fibrosis that peaks 7 days postinjury and is completely resolved by 28 days after injury, which is significantly different from what is seen in humans, who do not recover without surgical intervention.²⁵ With the addition of the fasciotomy surgery and the spacer implantation, we observed a change in the time course of fibrotic deposition and extent of injury, with a significant increase in fibrosis at days 7 and 14 after injury, suggesting that the addition of the fasciotomy exacerbated the extent of injury.

These data are correlated with increased α-SMA⁺ cells in the fasciotomy group, suggesting that there is an increased activation of myofibroblasts in response to the more severe injury, although the mechanism of this response remains to be explored. This increased severity of the injury and delayed regenerative response are further evidenced by the increase in CD68⁺ macrophages at 14 days after injury, suggesting a sustained inflammatory response in the CS model with fasciotomy. Increased macrophage activation correlated with increased damage to the skeletal muscle, and lends evidence to the hypothesis that this new CS model induced significantly more tissue injury than the previous CS rat models.

As previously mentioned, this reflects a more severe inflammatory microenvironment for an extended period of time, which more closely mimics the pathology of CS in humans. This extended inflammatory period impacts skeletal muscle regeneration, as well as any potential therapies. It is well established that the M1 macrophage inflammatory cytokine release is important for satellite cell activation and proliferation, while M2 macrophage anti-inflammatory release is critical for differentiation and myotube fusion.¹² Thus, we have created a better tool for investigating the underlying pathology of CS and potential new therapies by creating a small animal model that better matches the inflammatory time course seen in humans.

Coinciding with the increased injury severity was the delayed regenerative response in muscles from rats that underwent CS and fasciotomy. While we previously demonstrated a peak in Pax7⁺ cells 4 days after injury, with a return to baseline control levels by 14 in the original rat model of CS,³ our CS model with fasciotomy showed a delay in peak Pax7⁺ satellite cell expression, which we hypothesize is due to the exacerbated additional injury caused by placement of the spacer. These data are supported by the trend toward increased degenerating myofibers in the CS+Fasc group and the leftward shift in myofiber size distribution.

Smaller myofibers are likely due to active fusion taking place within the damaged tissue bed, although further experiments are needed to confirm this. The delay in proliferating Pax7⁺ satellite cells seen with this model demonstrated the delayed regeneration seen in humans, allowing for new therapies and strategies in regenerative medicine to be explored. Indeed, new strategies to reverse this delay in satellite cell activation, proliferation, differentiation, and fusion may be screened with this new rat model, with an optimal change likely to be seen at 7 days or less.

Collectively, the data presented herein suggest that the use of the described fasciotomy procedure effectively enhanced the rat model of CS to better match the clinical condition. However, there are several other parameters still worth investigating. The characterization of functional and histological changes at later time points is necessary, as is further exploration of the molecular mechanisms of the pathophysiology of CS injury and regeneration.

Our current data provide evidence that silicone spacer insertion into the fasciotomy site in this rat model of CS provided a more severe injury than the previous rat model, which will allow for better screen of treatment options for CS. While physicians are able to save life and limb, complete restoration of function for affected patients remains an elusive goal for CS research. This model provides an additional tool toward attaining that goal.

Authors' Contributions

The authors herein are responsible for the collection of these data and the running of this research. J.M.P. was responsible for the project concept, data collection, analysis, drafting of the article, revisions, and final approval. K.M. and B.P.Y. assisted with data collection and analysis, and final approval. Y.Z., T.L.C., and S.S. were responsible for the project concept, data interpretation, article revision, and final approval.

Disclosure Statement

No competing financial interests exist.

Funding Information

This work was Supported by the Department of Defense (USAMRAA AFIRM W81XWH-08-2-0032) and the National Institute of Biomedical Imaging and Bioengineering (5T32EB014836-05).