The Effects of Engineered Skeletal Muscle on Volumetric Muscle Loss in The Tibialis Anterior Of Rat After Three Months In Vivo

Abstract

Volumetric muscle loss (VML) is traumatic, degenerative, or surgical loss of skeletal muscle that exceeds the regenerative capacity of the remaining muscle, thus resulting in impaired muscle function. In humans, the loss of 30% or more mass of any one muscle will result in permanent structural and functional loss. Current VML repair treatments are limited by donor site morbidity and graft tissue availability, necessitating alternative muscle graft sources. To address this need, our lab has fabricated tissue-engineered skeletal muscle units (SMUs) for implantation into a 30 % VML model in the tibialis anterior (TA) muscle of rat. Previous results showed that after 28 days in vivo, muscle with a 30% VML repaired with our SMUs produced significantly more force than muscle with acute VML. But repair with our SMU did not fully restore muscle force production to that of native muscle. Thus, we hypothesized that more time for in vivo tissue regeneration would allow for greater force recovery. Therefore, the purpose of this study was to examine the long-term (3-month) effects of our SMUs on a 30% VML repair. We also assessed the effects of reinnervation by redirecting a branch of the peroneal nerve to the repair site. Thirty-nine, 2-month old female F344 rats were separated into a nonsurgical control group (n=5) and four surgical experimental groups (VML Only, n=5; VML+Nerve Redirect, n=6; VML+SMU, n=5; VML+SMU+ Nerve Redirect, n=8). Experimental rats were allowed a 3-month recovery period post-surgery before undergoing in situ force testing of the surgical (left) TA. The left TA of the control animals also underwent in situ force testing. Finally, the surgical (left) and contralateral (right) TAs of the experimental animals, as well as the left TA of the control animals, were explanted for histological analysis. Results for specific force showed that while all groups recovered specific forces similar to that of native muscle, the two SMU groups had significantly higher specific forces, on average, compared to the uninjured control group. Histological staining showed small muscle fibers in the repair site in animals that received an SMU. The average cross-sectional area of the native fibers just outside the area of repair (or the equivalent area in control animals) was not significantly different between groups, indicating that hypertrophy of remaining fibers did not contribute to the recovery of force following the VML. Our results suggest that following a 30% VML of the TA muscle, all surgical groups were able to recover TA mass, maximum tetanic and specific force production. Thus, creating a 30% VML in the TA in a rat model is not enough a sufficient VML to produce the sustained VML seen in humans following similar 30% loss of muscle volume.

Lay Summary

Volumetric muscle loss (VML) or the loss of muscle that exceeds the body’s ability for self-repair is a devastating clinical issue with no reliable options for repair. Previous work from our laboratory that used our engineered skeletal muscle to repair a VML for 28 days suggested that longer recovery times were necessary for optimal repair of a 30% VML. Therefore, the purpose of this study was to examine the effects of longer recovery times on the ability of our SMUs to repair a 30% VML injury by allowing a 3-month recovery instead of 28 days. We hypothesized that a longer recovery time would show continued improvement of both structure and contractile function of the damaged muscle. Our results showed that the longer recovery time did enhance recovery of VML regardless of the repair modality used.

Introduction

Volumetric muscle loss (VML) is the traumatic, degenerative, or surgical loss of skeletal muscle that exceeds the body’s regenerative capacity, thus resulting in chronic impaired muscle function and physical deformity [1]. Current treatment strategies for VML include autologous free muscle transfer and muscle flap transposition [2]. Muscle transfer and transposition procedures have both shown a degree of success but are limited by lack of available graft tissue and donor site morbidity. Thus, a demand exists for exogenous muscle grafts, prompting regenerative medicine approaches to be increasingly investigated as potential strategies for VML treatment.

While no laboratory has successfully engineered tissue that is structurally or functionally equivalent to adult skeletal muscle, our lab has developed a technique for engineering scaffold-less 3-D skeletal muscle constructs similar to native muscle [3, 4]. These constructs, or skeletal muscle units (SMUs), are multiphasic in that they have tendon-like endings and develop functional myotendinous junctions in vitro [5]. This provides a means of anchoring the construct to native tendon and bone and allows contractile force production to be translated into movement. The SMUs are also capable of contracting and when stimulated electrically, producing an average maximum isometric force of 279 ± 51 μN in vitro [3, 4, 6]. Previously, we have shown that following one week of implantation in an ectopic site in the hindlimb of rat, the maximum isometric force of the SMUs increased to an average of 549 ± 103 μN. Also, within one week of implantation, our SMUs achieved vascularization and began to develop innervation. This is crucial because, as a highly complex and metabolically demanding tissue, skeletal muscle relies on advanced vascular networks for blood perfusion and neurotization for maintenance of structure and contractile control.

Our laboratory has developed a rat model for creating and repairing acute VML by removing a longitudinal section, approximately one third of the total width, from the lateral aspect of the tibialis anterior (TA) muscle [4]. In a previous experiment, we used this model and replaced approximately 10% of the lost tissue volume with an SMU, sutured one tendon end to the remains of the native proximal tendon and the other into a bone tunnel on the distal end of the tibia, then allowed 28 days of recovery before explantation [4]. After 28 days, the SMUs developed small muscle fibers aligned along the axis of the muscle with an average cross-sectional area 8% that of native fibers. In addition, the SMUs exhibited advanced sarcomere organization, vascularization, and innervation, and had developed an enthesis at the tendon-bone interface. After 28 days, the TAs repaired with SMUs failed to fully recover muscle mass or maximum force production to that of native muscle, however, they did recover significantly more force than muscles with acute VML with no repair.

Therefore, the purpose of this study was to examine the long-term effects of our SMUs on the repair of a 30% VML injury by allowing 3 months of recovery instead of 28 days. We hypothesized that a longer recovery time would show continued improvement of both structure and contractile function of the damaged muscle. In addition, we hypothesized that muscles with VML that received an SMU would recover better than those with VML and no intervention. We also investigated the ability to enhance recovery via direct reinnervation of the surgical site by routing a branch of the peroneal nerve to the injury site and hypothesized that muscles that received a rerouted nerve would recover better than those whose native nerve was simply cut.

Materials and Methods

Animal model and animal care

The study was conducted using 120 to 150 g female Fischer 344 rats obtained from Charles River Laboratories Inc. (Wilmington, MA) and Harlan Laboratories (Haslett, MI). Briefly, thirty-nine rats were separated into a control and four experimental groups (control, n=5; VML Only, n=5; VML+Nerve Redirect, n=6; VML+SMU, n=5; VML+SMU+Nerve Redirect, n=8). There were initially 5 rats per group, but additional animals were added to the VML+Nerve Redirect and VML+SMU+Nerve Redirect groups due to the difficulty of the nerve redirect surgeries. Following the VML procedure, all rats were allowed to recover for 3 months and then underwent in situ force testing of the repaired muscle. Next, the repaired muscle was explanted for histological analysis. All animals were acclimated to our colony conditions, i.e. light cycle and temperature, for one week prior to any procedure. The animals were fed Purina Rodent Chow 5001 and water ad libitum. Harvested tissues taken from donor animals were used as an allogenic cell source of muscle satellite cells and bone marrow-derived stem cells for production of the tissue-engineered muscle and tendon tissues. All surgical procedures were performed in an aseptic environment, with animals in a deep plane of anesthesia induced by isoflurane (non-terminal procedures only) or intraperitoneal injections of sodium pentobarbital at 65 mg/kg. Supplemental doses of pentobarbital were administered as required to maintain an adequate depth of anesthesia, and animals were euthanized via pneumothorax. All animal care and animal surgery procedures were in accordance with The Guide for Care and Use of Laboratory Animals [7] and the protocol was approved by the University of Michigan’s Institutional Animal Care & Use Committee.

Preparation of media

All media was prepared and stored at 4°C prior to use and then warmed to 37°C in a heated bead bath immediately before use. Transport medium (TM), consisting of Dulbecco’s phosphate-buffered saline (DPBS; cat. No. 14190144; Gibco, Carlsbad, CA) with 2% antibiotic-antimycotic (ABAM; cat. No. 15240062; Gibco), was used to transfer freshly isolated muscle from the surgical suite to the tissue culture facility for isolation. Bone growth medium (B-GM) contained 79% Dulbecco’s modified eagle medium (DMEM; cat. No. 11995065; Gibco), 20% fetal bovine serum (FBS; cat. No. 10437028; Gibco), 1% ABAM, 6 ng/mL recombinant human FGF-basic (FGFb; cat. No. 100–18B; PeproTech, Rocky Hill, NJ), and 5 ng/mL dexamethasone (SKU D4902; Sigma-Aldrich). Bone differentiation medium (B-DM) contained 92% DMEM, 7% FBS, 1% ABAM, 0.13mg/mL L-ascorbic acid 2-phosphate (SKU A8960; Sigma-Aldrich), 5ng/mL L-proline (SKU P0380; Sigma-Aldrich), and 5ng/mL dexamethasone. Muscle growth medium (M-GM) contained 60% F-12K nutrient mixture (cat. No. 21127022; Gibco), 24% DMEM, 15% FBS, 1% ABAM, and 2.4 ng/mL FGFb. Muscle differentiation medium (M-DM) was composed of 70% medium 199 (cat. No. 11150059; Gibco), 23% DMEM, 6% FBS, 1% ABAM, 0.01% insulin-transferrin-selenium-X (SKU I1884; Sigma-Aldrich, St. Louis, MO), and 10.5 ng/mL L-ascorbic acid 2-phosphate.

Preparation of muscle construct dishes

As previously described, muscle constructs were fabricated in individual 60-mm cell culture dishes (product No. 353002; Corning, Inc., Corning, NY) [4]. Each plate was coated with 5 mL Sylgard 184 silicone elastomer (material No. 4019862; Dow Consumer Solutions, Midland, MI) and allowed to cure for one week. One to five days prior to use, each Sylgard-coated plate was coated with 1μg/cm² natural mouse laminin (cat. No. 23017–015, Gibco), suspended in 4 mL DPBS, and left to dry for 24 to 48 hours. Residual salt crystals were dissolved and removed by rinsing the dishes with 4 mL DPBS. The dishes were then filled with 3 mL M-GM and decontaminated with UV light (wavelength 253.7nm) for one hour and placed in a 37°C/5% CO₂ incubator for up to three days prior to plating the muscle cells.

Preparation of tissue engineered bone anchors

Bone marrow was removed under aseptic conditions and bone marrow-derived stem cells were isolated with Ficoll-Paque PLUS density gradient media (product No. 17144002; GE Healthcare, Pittsburg, PA). Isolated bone marrow cells were plated in 100-mm tissue culture dishes (product No. 353003; Corning, Inc.) in B-GM for two days and then passaged and re-plated. After four such passages, the cells were plated in 100-mm tissue culture dishes at a density of 1.3 million cells per plate and left to grow in B-GM. On the second day, the cells became confluent and formed a monolayer in each dish. Following five days on B-GM, the cells were switched to B-DM for two days, after which time the monolayers delaminated. Delaminated monolayers were collected and pinned in cylindrical form in Sylgard-coated dishes and fed B-DM. Five to ten days after delamination and 3-D construct formation, bone constructs were cut into 5-mm sections to be used as engineered bone anchors and subsequently pinned onto developing muscle monolayers.

Dissection of muscle and isolation of muscle satellite cells

Both soleus muscles from donor rats were removed under aseptic conditions and sterilized in 70% ethanol, then placed in 5 mL TM for five minutes. The muscles were then minced using a razor blade, placed under UV light for 15 minutes in 15 mL Ham’s F-12 nutrient mixture (cat. No. 11765–047; Gibco), and then added to a dissociation media consisting of 40U dispase II (0.5U/mg; cat. No. 17105–041; Gibco) and 1600U type IV collagenase (160U/mg; cat. No. 17104–019; Gibco) in 20 mL Ham’s F-12. The mixture was kept at 37°C with agitation for 90 minutes to allow the minced muscle to dissociate, and then poured through a 100-μm filter and centrifuged. Finally, the dissociation media was aspirated off and the cells were resuspended in M-GM.

Construct formation

The cells from the muscle isolation procedure were plated in M-GM at a density of 400,000 cells per laminin- and Sylgard-coated 60-mm cell culture dish. To facilitate attachment to the dish surface, the cells were left undisturbed for five days following initial plating. The cells were then fed M-GM every two days until they became ~70% confluent, at which point they were passaged and re-plated. The cells were then fed M-GM every two days until they were 100% confluent and a network of elongated myotubes began to form. At this point, the 5-mm tissue-engineered bone anchors were pinned onto the cell monolayers 1.5 cm apart and the media was changed to M-DM, which the plates were fed every two days. After approximately a week on M-DM, the monolayers delaminated from the dishes and rolled into cylindrical muscle constructs held at length by the bone anchors.

SMU contractile measurements

Contractile properties of the SMU constructs were measured prior to implantation into host animals. The protocol for measuring contractility of engineered muscle constructs has been described previously [3, 4, 6]. Briefly, the pin on one end of the construct was freed from the Sylgard and attached to a force transducer with canning wax. For field stimulation of the entire construct, platinum wire electrodes were placed along either side of the SMU. The temperature of the construct was maintained at 37° C, using a heated aluminum platform. Passive baseline force was measured as the average baseline passive force preceding the onset of stimulation. Twitches were elicited using a single 2.5-ms pulse at 10, 30, 60 and 90 mA, whereas maximum tetanic force was determined using a 1-s train of 2.5-ms pulses at 90 mA and 10, 20, 40, 60 and 80 Hz. Data files for each peak twitch force and peak tetanic force trace were recorded and analyzed using LabVIEW 2009 data acquisition software.

Surgical procedure

As previously described, an incision was made along the left lower hindlimb exposing the TA, and a longitudinal cut was made to remove ~30% of the TA volume [4]. A 0.9-mm tunnel was drilled into the proximal tendon insertion site on the tibia. In all surgical groups, the peroneal nerve distal to the innervation of the extensor digitorum longus was transected, along with its associated vasculature. The peroneal nerve is not the nerve innervating the TA muscle. At the anatomical site of peroneal nerve cut, we only denervated the big toe and that does not produce any no measurable effect on ambulation. The tibial nerve innervating the TA muscle is left untouched. In the VML Only and the VML+SMU groups the peroneal nerve was cut, directed away from the VML site and left unrepaired. In the, VML+Nerve Redirect and VML+SMU+Nerve Redirect groups, the distal portion of the peroneal nerve, along with its associated vasculature, was placed into the VML site for enhanced innervation. In the groups repaired with and SMU, a single SMU was then placed in the repair site, with one of its bone ends pulled into the tibial bone tunnel and sutured in place on the periosteum, and the other end sutured to the remaining distal tendon of the TA. The transected nerve was then sutured to the SMU using 9–0 suture (Figure). The surgical site was closed with surgical staples and Carprofen was administered at a dose of 5mg/kg every 12 hours for 48 hours post-surgery. The staples were removed after 10 days.

TA contractile measurements

Following 3 months of recovery, in situ contractile properties of the TA muscles were measured as described by [8]. Briefly, the animal was anesthetized and placed on a temperature-controlled platform to maintain body temperature at 37°C. The whole TA muscle was isolated from surrounding muscle and connective tissue using great care not to damage the nerves or blood vessels. 2–0 silk suture was tied around the distal tendon and the tendon was severed. The hindlimb was securely tied to a fixed post with wire at the knee. The distal tendon of the TA muscle was then tied to the lever arm of a servomotor (model 305B; Aurora Scientific, Aurora, ON). A continual drip of saline warmed to 37°C was administered to the TA muscle to maintain its temperature and to prevent desiccation. The muscle was activated by stimulation of the sciatic nerve using a bipolar platinum wire electrode. The voltage of single 0.2-ms stimulation pulses was adjusted to give a maximum isometric twitch. Subsequently, muscle length was adjusted to the optimal length (L_o) at which twitch force was maximal. With the muscle held at L_o, 300-ms trains of stimulus pulses were applied at increasing stimulation frequencies until the maximum isometric tetanic force (P_o) was achieved.

After all force measurements were completed, the animal was euthanized and both TA muscles were removed, trimmed of their tendons, blotted, and weighed. Muscle fiber length was calculated by multiplying L_o by the ratio of fiber length to TA muscle length [9]. Functional muscle CSA was calculated by dividing the muscle mass by the product of muscle fiber length and the density of mammalian skeletal muscle (1.06 g/cm³). Specific force was calculated by dividing P_o by functional CSA for each muscle. Immediately after muscle mass was measured, muscles were coated in tissue freezing medium (cat. No. TFM5; General Data Company, Inc., Cincinnati, OH), frozen in 2-methylbutane cooled by dry ice, and stored at −80°C until analyzed for histology.

Histological analysis

Frozen TA samples were sectioned at 12 μm and mounted on Superfrost Plus microscopy slides for histological analysis. Sections were stained with H&E for observations of general morphology and immunofluorescent antibodies for identification purposes. For immunohistochemical analysis, frozen sections were fixed with −20°C methanol for 10 minutes and rinsed with DPBS. The sections were then submerged for 15 minutes in 0.05% Triton X-100 (Sigma-Aldrich) in DPBS (PBST) and blocked with PBST containing 3% bovine serum albumin (BSA; cat. No. A2153–10g; Sigma-Aldrich) at room temperature for 30 minutes. Next, sections were incubated overnight at 4°C with primary antibodies diluted in PBST with BSA. The primary antibodies used detect the presence of myosin heavy chains (mouse monoclonal antibody, 1:20 dilution, cat. No. ALD-58; Developmental Studies Hybridoma Bank, Iowa City, IA) and laminin (rabbit polyclonal antibody, 1:200; cat. No. ab30320, Abcam, Cambridge, MA). Following 3 washes in PBST, the sections were incubated in 1:500 dilutions of Alexa Fluor antimouse or anti-rabbit antibodies (Invitrogen, Carlsbad, CA) for three hours at room temperature. Following another three washes in PBST, the sections were fixed in Prolong Gold with DAPI (cat. No. P36935; Invitrogen) and coverslipped. Stained sections were examined and photographed with an Olympus BX51 microscope. Images were merged using PhotoshopCS6 and analyzed using Fiji image processing software.

Statistical analysis

Values are presented as mean ± standard error and were calculated using Microsoft Excel 2016. Calculations of significant differences between means were performed using GraphPad Prism 8 statistical analysis software. Means were compared using one-way analysis of variance tests with Tukey post-hoc analyses. Differences were considered significant at p<0.05.

Results

Body and Muscle Mass

Body mass was measured at the time of implantation and explantation to determine the effect of SMUs on the animals’ overall health and growth, while surgical and contralateral TA muscle mass at the time of explantation were measured and compared to determine the effect of VML and SMU repairs on TA muscle mass. The body mass (grams) during recovery period of the Control, VML Only, VML+Nerve Redirect, VML+SMU, and VML+SMU+Nerve Redirect groups were 163.3 ± 1.2, 169.4 ± 3.9, 170.0 ± 2.1, 167.0 ± 2.0, and 183.8 ± 5.2, respectively and there were no significant differences in growth between the surgical groups. The percent increase in body mass during recovery period of the VML Only, VML+Nerve Redirect, VML+SMU, and VML+SMU+Nerve Redirect groups were 14.20 ± 3.2%, 18.49 ± 2.5%, 18.75 ± 1.8%, and 24.48 ± 5.3% respectively, and there were no significant differences in growth between the surgical groups. This indicates that neither the implantation of an SMU nor the redirecting of the peroneal nerve had a negative impact on the overall health or growth of the animals. Note that this metric is not available for the control group because this group was age-matched to the surgical groups at the time of explant and thus body mass was only recorded at one time point. The TA muscle of the surgical leg in the VML Only, VML+Nerve Redirect, VML+SMU, and VML+SMU+Nerve Redirect groups had an average mass deficit of 7.16 ± 2.4%, 8.36 ± 3.2%, 10.29 ± 3.8%, 12.19 ± 2.5%, respectively, compared to the contralateral leg. However, the percent mass recovery of the surgical TAs compared to the mass of the contralateral uninjured TA were not significantly different between the experimental groups (Figure 1). Thus, while all experimental groups showed significant regeneration of TA muscle mass, compared to the nonsurgical contralateral TAs the experimental TAs still had non- significant mass deficits ranging from 7 – 12%.

Figure 1:

Following 3 months of recovery from VML, both the surgical and contralateral (non-surgical) TAs were dissected and weighed. Five experimental groups were created to test the efficacy of using our SMUs to repair VML: Control – No VML (n=5); VML Only (n=5); VML+Nerve Redirect (NR; n=6); VML+SMU (n=5); and VML+SMU+Nerve Redirect (NR; n=8). Since there was not a significant difference in mass between the TA of the right leg compared to the left leg in the control (No VML group), the mass of each experimental animal’s surgical TA was compared to its contralateral TA as a measure of muscle mass recovery. There was no significant difference in muscle mass recovery between groups. NR = Nerve Redirect. Data represents means ± SEM.

Contractile Properties

In vitro SMUs had a mean pre-implantation force of 279 ± 49 μN. Following 3 months of recovery, the constructs were integrated into the host TAs and could not be separated and force-tested independent of the host muscle. The average maximum tetanic force of the TAs of the uninjured control group was 4894 ± 176 mN, while the average maximum tetanic force of the surgical TAs of the VML Only, VML+Nerve Redirect, VML+SMU, and VML+SMU+Nerve Redirect groups were 4511 ± 165 mN, 4893 ± 128 mN, 4931 ± 141 mN, and 5004 ± 107 mN, respectively, and were not significantly different from the control group or from each other (Figure 2). The average specific force of the TAs of the uninjured control group was 22.39 ± 1.4 N/cm², while the average specific force of the surgical TAs of the VML Only, VML+Nerve Redirect, VML+SMU, and VML+SMU+Nerve Redirect groups were, 23.58 ± 1.0 N/cm², 25.34 ± 0.8 N/cm², 29.32 ± 1.7 N/cm², and 27.26 ± 0.7 N/cm², respectively (Figure 3). On average, the specific forces of the VML+SMU group and the VML+SMU+Nerve Redirect group were significantly greater than the control (p=0.0034 and p=0.0287, respectively). In addition, the specific forces of the VML+SMU group were significantly greater, on average, than the VML Only group (p=0.0182).

Figure 2:

Following 3 months of recovery from VML, both the grafted and contralateral TAs were assessed for maximum force production. Five experimental groups were created to test the efficacy of using our SMUs to repair VML: Control – No VML (n=5); VML Only (n=5); VML+Nerve Redirect (n=6); VML+SMU (n=5); and VML+SMU+Nerve Redirect (n=8). Maximum force production was not significantly different between groups. NR = Nerve Redirect. Data represents means ± SEM.

Figure 3:

Following 3 months of recovery from VML, both the surgical and contralateral TAs were assessed for specific force production (maximum force normalized by muscle cross-sectional area). Five experimental groups were created to test the efficacy of using our SMUs to repair VML: Control – No VML (n=5); VML Only (n=5); VML+Nerve Redirect (n=6); VML+SMU (n=5); and VML+SMU+Nerve Redirect (n=8). Specific force production was significantly higher for both SMU groups compared to the control group, and for the VML+SMU group compared to the VML Only. NR = Nerve Redirect. Data represents means ± SEM. * Significance from control group, p<0.05. ** Significance from control group, p<0.01. + Significance from VML & cut group, p<0.05.

Morphology and Histology

Following 3 months of recovery and measures of contractile properties, both the surgical and contralateral TA muscles were dissected from each animal and observed for gross morphology. At 3 months, the repaired TA’s were indistinguishable from the contralateral TA with respect to whole muscle cell morphology, except in the nerve redirected muscles the presence of the suture used for repair was visible and used as an indicator for the redirected nerve (Figure 4A &B). H&E staining of the repaired and control TA sections was performed to analyze the general morphology of the TA muscles after 3 months of recovery (Figure 4). Following 3 months of recovery, SMUs explanted from the VML model exhibited small but distinct muscle fibers (Figure 4B). These fibers were aligned along the axis of the muscle. Such fibers were not visible in any of the animals without an SMU repair. Immunostaining for myosin heavy chain content confirmed the identity of these structures as muscle fibers, while immunostaining for laminin indicated that each fiber had a laminin-rich extracellular matrix like the endomysium of native muscle fibers. After 3 months of recovery, there was no significant difference in the average cross-sectional area of the native muscle fibers surrounding the VML repair site between the control, VML Only, VML+Nerve Redirect, VML+SMU, and VML+SMU+Nerve Redirect groups (p=0.4837; Figure 5).

Figure 4:

Following 3 months of recovery from VML, the TA muscles were isolated and gross morphology of the repair site assessed (A). The site of innervation and vascularization was confirmed intact (B). IHC staining (RED-MF20, GREEN-LAMININ) on cross-sections through the mid-belly of the regenerated tissue showed regenerated muscle fibers (C).

Figure 5.

Average cross-sectional area of native TA muscle fibers surrounding the VML repair site. After three months of recovery, there was no significant difference in the average cross-sectional area of the native muscle fibers surrounding the VML repair site between the Control, VML Only, VML+Nerve Redirect, VML+SMU, and VML+SMU+Nerve Redirect. NR = Nerve Redirect. Data represents means ± SEM.

Discussion

The purpose of this study was to determine the effect of three months in vivo recovery on our tissue-engineered skeletal muscle constructs following a 30% VML in the tibialis anterior of rat. We hypothesized that after three months we would see improved structure and function of the TA muscles compared to our previous the 28-day experiment [4], that muscles repaired with SMUs would recover better than TAs with VML and not repaired, and that muscles that had the peroneal nerve rerouted to the repair site would recover better than those without.

After three months of recovery, the TA muscles from all four surgical groups still had mass deficits ranging from 7 – 12% compared to the contralateral TA. However, as there were no significant differences in mass deficit between any of the surgical groups regardless of whether they were repaired with an SMU or not repaired, neither the SMUs nor the direct nerve reinnervation appear to have affected muscle mass recovery. In corroboration with the mass data, the maximum tetanic forces were not significantly different between any of the five groups, including the control, meaning all of the surgical groups were able to recover maximum tetanic force production. While this data indicates that neither the SMU nor the nerve redirect interventions aided in the recovery of maximum force, the specific forces in the groups repaired with an SMU did result in an increase in the specific force compared to control. Specific force, or force per unit area is an indicator of muscle health. An increase in specific force in the groups with SMUs suggests that the SMU did have a significant role in skeletal muscle tissue regeneration. One can observe in Figure 4, that the SMU does add new skeletal muscle fibers to the repair site and could be the reason for the significant increase in force. Published studies on the SMU also show that prior to implantation the SMU is highly aligned with muscle fibers and extracellular matrix and that this structure is maintained during regeneration and would aid in the longitudinal transmission of force. In fact, when comparing the repair site of a VML with or without an SMU [10] one sees that the repair site without an SMU has an amorphous infiltration of scar tissue versus the highly organized extracellular matrix and muscle observed following repair with an SMU. Albeit, the forces that are developed by the small muscle fibers of the SMU in this study account for only a small increase in maximum tetanic force of the whole TA muscle. Future experiments are planned to implant larger volumes of SMU tissue to increase the regenerative potential and subsequent force production.

Our results show that following a 30% VML of the TA muscle in a rat model, all surgical groups were able to recover TA mass and maximum tetanic force production. The complete recovery of the maximum tetanic forces in the VML Only group indicates that a 30% VML in a rodent model does produce the long-lasting functional deficits observed in humans with the same VML injury. Upon observing the regeneration of both mass and maximum tetanic force following the 30% loss of mass, we hypothesized that the recovery of the mass and maximum tetanic force production could be explained by hypertrophy of endogenous (native) muscle fibers. However, quantification of the cross-sectional area of the native muscle fibers surrounding the graft site showed that there were no significant differences in cross-sectional areas between the groups. This data suggest that removing more than 30 % of the muscle in the TA in a rat model is necessary to produce sustained VML seen in humans following similar loss of muscle volume. Future studies will involve larger VML defects in order to ascertain the amount of tissue removal necessary to cause a permanent functional deficit.