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Article

Effects of Artificial Achilles Tendon on Hindlimb Movement Biomechanics and Muscle Morphology in Rabbits

by
Obinna P. Fidelis
1,*,
Katrina L. Easton
1,
Madison Smith
1,
Gabriela Bastos
2,
Kristin Bowers
2,
David E. Anderson
2 and
Dustin L. Crouch
1
1
College of Engineering, University of Tennessee, Knoxville, TN 37996, USA
2
College of Veterinary Medicine, University of Tennessee, Knoxville, TN 37996, USA
*
Author to whom correspondence should be addressed.
Biomechanics 2025, 5(3), 47; https://doi.org/10.3390/biomechanics5030047
Submission received: 19 May 2025 / Revised: 18 June 2025 / Accepted: 25 June 2025 / Published: 1 July 2025
(This article belongs to the Section Injury Biomechanics and Rehabilitation)
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Abstract

Background/Objectives: Artificial tendons offer an alternative to biological tendon grafts and may restore normative biomechanical functions in humans and animals suffering segmental or complete tendon loss. The aim of this study was to quantify movement biomechanics during hopping gait and muscle properties of New Zealand White rabbits with a polyester silicone-coated (PET-SI) artificial tendon. Methods: In five rabbits, the biological Achilles tendon of the left hindlimb was surgically replaced with a PET-SI artificial tendon; five operated control rabbits underwent complete surgical excision of the biological Achilles tendon in the left hindlimb with no replacement (TE). Results: Across both groups at 2 and 8 weeks post-surgery compared to baseline, the maximum ankle angle during stance and swing phases of stride was significantly lower (i.e., more dorsiflexed) (p < 0.001), the peak vertical force was significantly higher (p < 0.001), and the average ground contact area was significantly lower (p < 0.001). At 8 weeks post-surgery, the muscle cross-sectional area of the lateral gastrocnemius was significantly higher in the PET-SI group than in the TE group (p = 0.006). Muscle mass and length were lower in the operated limb compared to the non-operated limb across the two groups (TE and PET-SI), with no significant differences between groups. Conclusions: The artificial Achilles tendon did not appear to provide superior biomechanical support during hopping compared to the TE group. However, the artificial tendon preserved muscle structural properties that correspond to the muscle’s capacity to generate force. Future studies should optimize the tendon–tissue interface.

1. Introduction

Tendon injuries resulting in gaps that are “critically sized,” defined here as being too large to heal spontaneously or repair directly (e.g., by suturing the ends together), are challenging to manage [1]. The current gold-standard treatment for critically sized tendon gaps is a tendon graft from the same individual (autograft) or a different individual (allograft) [2,3]. Tendon grafts are used in tendon rupture treatments; for example, between 2005 and 2011, about 5.5% of Achilles tendon ruptures in the United States were treated with tendon grafts [4]. However, grafts have serious limitations that may affect clinical outcomes. For example, autograft harvesting causes significant donor site morbidity [5,6]. Drawbacks of allografts include risk of disease transmission, limited availability, and potential mismatch of graft length and size [7,8].
Replacement of the damaged tendon with an artificial tendon is an alternative to biological tendon grafts [9,10]. Artificial tendons could overcome the key limitations of biological tendon grafts listed above [11]. Additionally, the size of the artificial tendon can be customized for each patient [11]. Artificial tendons, investigated since the early 1900s, have been made from diverse materials such as silk, rubber, braided tantalum, nylon monofilament covered with polyethylene, woven Teflon, Dacron, carbon fiber, and polyester [12]. A recent, promising artificial tendon design consisted of braided strands of polyester microfiber suture [9,10,13]. The polyester, silicone-coated (PET-SI) artificial tendon was tested for replacement of the quadriceps tendon in goats for up to 180 days; the artificial tendon integrated with the muscle without evidence of microencapsulation, forming a myotendinous junction that was stronger than the biological junction [13].
A major limitation of the previous in vivo studies on PET-SI tendons is that they did not investigate the effect of artificial tendons on movement biomechanics or muscle properties [9,10,13,14]. Tendons play key biomechanical roles during movement, including storing and releasing elastic energy and transmitting forces from muscles to bones [15,16,17]. Due to their biomechanical roles, tendons influence muscle forces and length changes, which regulate muscle properties [18,19]. In a recent study by our research group, we quantified the movement biomechanics of New Zealand White (NZW) rabbits in which the biological tibialis cranialis tendons were replaced with a PET-SI artificial tendon [11]. Rabbits with the artificial tendon exhibited near-normal gait biomechanics. However, to increase the translational value of the animal model, there is a need to test the PET-SI artificial tendon in a clinically representative higher load application (e.g., Achilles tendon replacement). The Achilles tendon accounts for up to 20 percent of all large tendon ruptures and occurs in both athletes and the general population [20,21,22]. Therefore, a reliable artificial Achilles tendon that restores normative limb biomechanics represents a novel treatment of Achilles tendon injuries.
The purpose of the current study was to quantify movement biomechanics and muscle properties of NZW rabbits with a PET-SI artificial Achilles tendon. Since the Achilles tendon is an extensor of the hock, we hypothesized that rabbits with the PET-SI artificial Achilles tendon would have greater ankle range of motion and maximum ankle plantar flexion angle during the swing phase of gait, compared to rabbits with Achilles’ tendon excision only. We also hypothesized that muscle cross-sectional area, muscle mass, and muscle length would be greater in the rabbits with an artificial Achilles tendon compared to rabbits with tendon excision only.

2. Materials and Methods

2.1. Artificial Tendon

The PET-SI artificial Achilles tendon was based on a previously published design [13]. Three strands of custom double-armed size 0 braided polyester suture (RK Manufacturing, Danbury, CT, USA) were folded in half to form a loop at the mid-point, then braided to the desired length (Figure 1A). The artificial tendon length selected for each rabbit at surgery was based on the length of the biological tendon being replaced (see below); therefore, we fabricated several artificial tendons that varied in length in ~2 mm increments to permit animal-specific matching. Medical-grade biocompatible silicone (LSR-4301 silicone elastomer, Factor II, Inc., Lakeside, AZ, USA) was applied to the braided and looped portion of the artificial tendon to discourage tissue adhesion. Prior to surgery, the artificial tendons were cleaned in an ultrasonic cleaner (Model: JPS-08A) and sterilized with hydrogen peroxide gas.

2.2. Animal Handling and Surgery

Ten female NZW rabbits (Envigo RMS, Inc., Indianapolis, IN, USA, and Charles River Laboratories, Wilmington, MA, USA, age = 18.18 ± 1.89 weeks, body mass = 3.34 ± 0.21 kg) were randomly assigned to either the PET-SI (n = 5) group or the tendon excision (TE, n = 5) group. The Institutional Animal Care and Use Committee at the University of Tennessee Knoxville approved all animal procedures (protocol #2726). The rabbits were either pair-housed or individually housed at some point or for the entire duration of the study. The rabbits acclimated to the animal facility for a minimum of 2 weeks prior to surgery and were fed ad libitum with a standard laboratory diet and Timothy hay. Additionally, each rabbit had free access to the playpen via a ramp from their cage during the day on weekdays, prior to surgery. Post-surgery, playpen time was initially limited and then gradually increased until they again had free access.
Prior to surgery, the rabbits were given hydromorphone (0.1 mg/kg IM) as a pre-operative analgesic, sedated with midazolam (1 mg/kg IM), and induced into general anesthesia with isoflurane via a face mask. Then, they were intubated, and general anesthesia was maintained with isoflurane gas vaporized in 100% oxygen. A loading dose of lidocaine (2 mg/kg IV) was administered, followed by a lidocaine CRI (50 mcg/kg/min IV) with isotonic fluids at a rate of 30 mL/h IV throughout the procedure. After shaving, the left hind limb was suspended and aseptically prepared for surgery with 70% isopropyl alcohol and 2% chlorhexidine scrub. Immediately prior to the start of surgery, a second dose of hydromorphone (0.05 mg/kg IM) was administered.
The rabbits were positioned in right lateral dorsal oblique recumbency for surgery. In the TE group, a caudolateral longitudinal skin incision was made over the Achilles tendon using a #10 scalpel blade, extending from mid-tibia to the calcaneus. The skin was retracted to expose the tendon and the calcaneus. The Achilles tendon (fused tendons of the gastrocnemius and soleus muscles) was released by cutting it at its insertion, taking care to preserve the overlying tendon of the superficial digital flexor muscle. The Achilles tendon was then excised by cutting it at the musculotendinous junction. The subcutaneous layer was closed with a simple continuous pattern with 4-0 PDS (Ethicon Inc., Cornelia, GA, USA). The skin was then closed with an intradermal pattern with 4-0 PDS.
In the PET-SI group, a caudolateral longitudinal skin incision was made over the Achilles tendon, extending from mid-tibia to the calcaneus. The skin was retracted to expose the tendon and the calcaneus. The gap length between the musculotendinous junction and the tendon point of insertion was measured intraoperatively with the ankle in full plantarflexion. An artificial tendon that was about 15% shorter than the biological tendon was selected for implantation; a shorter artificial tendon was selected to account for length added by the bone anchor and possible plastic deformation of the artificial tendon in vivo. The biological Achilles tendon (fused tendons of the gastrocnemius and soleus muscles) was released by cutting it at its insertion, taking care to preserve the overlying tendon of the superficial digital flexor muscle. A guide hole was pre-drilled with a 1.7 mm drill bit into the dorsal aspect of the calcaneus, angled in a proximal–caudal to distal–cranial direction, at the insertion of the Achilles tendon. A Parcus Anika bone anchor (2.5 mm MiTi, 2.5 mm × 5.7 mm, Parcus Medical, LLC, Sarasota, FL, USA, Figure 1B), with pre-threaded #2 suture, was screwed into the hole until fully inserted. The looped, distal end of the selected artificial tendon was sutured to the anchor using the pre-threaded suture. The proximal, armed ends of the artificial tendon were inserted through the distal end of the gastrocnemius and soleus muscles using a single suture loop pattern so that they exited from the sides of the muscles about 1 cm proximal to the distal ends [23]. Adjacent suture strings were tied together using six throws. The excess suture was cut and removed. The distal ends of the muscles were sutured around the proximal end of the braided portion of the artificial tendon with 4-0 PDS. The subcutaneous layer was closed with a simple continuous pattern with 4-0 PDS. The skin was then closed with an intradermal pattern with 4-0 PDS (Figure 2).
Post-surgical laser therapy was performed with a MultiRadiance ACTIVet Pro: 1000 Hz for 1 min, 50 Hz for 1 min, and 1000–3000 Hz for 1 min. Hindlimb radiographs were taken immediately and every 2 weeks post-surgery. The operated limb was bandaged for three days post-surgery, and silver sulfadiazine topical cream was applied to the surgical incision. The rabbits were administered meloxicam (1 mg/kg SC) and enrofloxacin (5 mg/kg SC, diluted in 6 mL of sterile saline) immediately following surgery. Post-operative care included hydromorphone (1 mg/kg IM, every 6 h for 3 days), enrofloxacin (5 mg/kg PO, every 12 h for 7 days), and meloxicam (1 mg/kg PO, every 24 h for 7 days). Additionally, lactated Ringer’s solution (150 mL SC) was administered twice daily, beginning the day after surgery and continuing for a total of 5 doses.

2.3. Data Collection

Hindlimb kinematics and ground contact pressure were measured once pre-surgery and every other week post-surgery. Prior to surgery and data recording, the rabbits were trained to hop along a 2.6 m long walkway for approximately 2 weeks. For each data collection session, we shaved the lateral side of the left (operated) hindlimb, then placed reflective, spherical markers (6.4 mm diameter) on the hip (greater trochanter), knee, ankle (lateral malleolus), and 5th metatarsophalangeal (MTP) joints (Figure 3). The markers were placed on the skin using double-sided marker stickers, with one side of the sticker attached to the marker and the other side attached to the skin of the rabbit. Seven high-speed cameras (Prime 13, OptiTrack, NaturalPoint, Inc, Corvallis, OR, USA) were placed in a semi-circle along one side of a pressure-sensitive walkway (2-Tile High-Resolution Strideway System, 1.3 m2 sensing area, Tekscan, Norwood, MA, USA). Hindlimb motion and ground contact pressures were recorded synchronously at 240 Hz by the cameras and pressure mat, respectively, while each rabbit hopped from one end of the walkway to the other.
We recorded ultrasound images of the tibialis cranialis and lateral gastrocnemius muscles in the left (operated) hindlimb using an ultrasound diagnostic system (ECO 3, Chison Medical Technologies Co. Ltd., Wuxi, China). The muscles were imaged four days before surgery (baseline) and approximately every other week post-surgery. The hindlimb was shaved prior to imaging and ultrasound gel was applied to the probe to improve image quality. The tibialis cranialis muscles were imaged by positioning the ultrasound probe on the cranial aspect of the limb, just distal to the end of the tibial tuberosity, and the lateral gastrocnemius muscles were imaged by positioning the probe on the lateral aspect of the limb at the level of the distal end of the tibial tuberosity (Figure 4). For each muscle and timepoint, at least three images that were clear and captured the entire muscle cross-section were analyzed.
At the end of the study (8 weeks post-surgery), the rabbits were humanely euthanized by intravenous overdose of pentobarbital (390 mg/mL, minimum 1 mL/10 lbs). The hindlimbs were collected via hip disarticulation, fixed in 10% phosphate-buffered formalin for at least 5 days, and then stored in 70% ethanol. Following fixation, the hindlimbs were dissected to permit visual inspection of the suture anchor and confirm the integrity of the proximal and distal tendon–tissue attachments. Two groups of muscles were dissected—the involved muscles (with direct connection to the Achilles tendon) and the uninvolved muscles (without a direct connection to the Achilles tendon but flexes the ankle joint). The involved muscles, the lateral gastrocnemius (LG), medial gastrocnemius (MG), and soleus (Sol), and select uninvolved muscles, the tibialis cranialis (TC), extensor digitorum (ED), and superficial flexor digitorum (FD), crossing the ankle were dissected from their origin to the myotendinous junction (Figure 5). For each muscle, mass and length were normalized by body mass and tibia length, respectively.

2.4. Data Analysis

The data were processed using pressure analysis software (Strideway 7.80, Tekscan, Norwood, MA, USA) and custom-written MATLAB scripts (MATLAB 2024a, MathWorks, Natick, MA, USA). The videos were exported from the motion capture software (Motive: Tracker 1.9, OptiTrack, NaturalPoint, Inc. Corvallis, OR, USA) and converted into Tekscan-readable format using a custom-written Python script. The purpose of this conversion was to synchronize the videos with the paw pressure map in Tekscan. Three-dimensional coordinates of the markers were exported from Motive software, and Matlab was used to calculate the joint angles. Joint angles for the knee, ankle in the stance and swing, and MTP (in stance only) phase of gait were calculated from joint angle timeseries data, measured for each gait cycle and averaged across cycles. The joint angles are presented as maximum angle (i.e., maximum joint angle at stance/swing), minimum angle (i.e., minimum joint angle at stance/swing), and range of motion (ROM, the difference between the maximum and minimum angles).
Statistical analysis software (SPSS IBM Statistics v.28) was used to perform a two-factor (group, timepoint, and their interaction) analysis of variance (ANOVA). A p-value < 0.05 was used to determine any significant differences. Groups included “PET-SI” and “TE”, and timepoint included “baseline” (pre-surgery), “2 weeks post-surgery”, and “8 weeks post-surgery”. Tukey’s HSD test was used for post hoc pairwise comparisons. The Kolmogorov–Smirnov test was used to determine if data for the kinematics, kinetics, and muscle properties were normally distributed; data were considered normally distributed if p > 0.05. All comparisons were made between groups and between timepoints.
Muscle cross-sectional area (CSA) was measured from the ultrasound image using image processing software (ImageJ, NIH) and analyzed using a two-way ANOVA with the factors ‘group’ and ‘timepoint’ and their interactions. Muscle CSAs were measured from a minimum of three image samples for each muscle, rabbit, and timepoint. The data were averaged across samples, then normalized by the rabbit’s body mass measured at each respective timepoint.
Muscle mass and length data were also analyzed using a 3-way ANOVA with the factors ‘side’ (operated vs. non-operated), ‘group’ (PET-SI vs. TE), and ‘muscle’ (LG, MG, Sol, TC, ED, and FD); the Tukey HSD test was used for post hoc pairwise comparison, with p < 0.05 considered significant.

3. Results

One rabbit in the TE group died during surgery due to anesthetic complications; all remaining rabbits in the study (n = 9) were included in the data analysis. There were no significant differences in age, body mass at time of surgery or euthanasia, or length of the biological tendon between the two groups (Table 1). The length of the artificial tendons at surgery ranged from 84% to 87% (mean = 85.75 ± 1.09%) of the length of the biological tendon (Table 1).

3.1. Biomechanics: Kinematics

There was no significant effect of group (TE and PET-SI) on the knee (p = 0.704), ankle (p = 0.651), and MTP (p = 0.550) joint kinematics during the stance phase of gait. However, there was a significant effect of timepoint on the knee angle (p < 0.001), ankle angle (p < 0.001) (i.e., ankle was more dorsiflexed), and MTP angle (p < 0.001). In the swing phase of gait, we found no significant effect of group on knee angle (p = 0.969) and ankle angle (p = 0.564) but a significant effect of timepoint on the knee angle (p < 0.001) and ankle angle (p < 0.001) during the swing phase of gait. Pairwise comparisons of kinematic variables across groups and timepoints are presented in Table 2. Table 3 presents kinematic variables, showing how the joint angles changed across timepoints within each group, as well as data normality of kinematic variables.
During the stance phase of gait, for both groups, maximum and minimum knee angles were significantly greater (i.e., the knee was more extended) at 2 weeks and 8 weeks post-surgery compared to baseline (p < 0.001). The knee ROM was significantly higher at baseline compared to 2 weeks post-surgery (p < 0.001) and at 8 weeks post-surgery compared to 2 weeks post-surgery (p = 0.005) (Figure 6, Table 2). In the swing phase, the maximum knee angle was significantly greater (i.e., the knee was more extended) at 8 weeks post-surgery compared to baseline (p = 0.015). The minimum knee angle was significantly greater (i.e., the knee was more extended) at 2 and 8 weeks post-surgery compared to baseline (p < 0.001). The ROM was significantly greater at baseline compared to 2 weeks post-surgery (p = 0.024).
For both groups, the maximum ankle angle during stance was significantly smaller (i.e., the ankle was more dorsiflexed) at 2 and 8 weeks post-surgery compared to baseline (p < 0.001). The minimum ankle angle was also significantly smaller (i.e., the ankle was more dorsiflexed) at 2 and 8 weeks post-surgery compared to baseline (p < 0.001) (Figure 7, Table 2). Ankle ROM was significantly smaller at 2 weeks post- surgery compared to baseline (p < 0.001) and 8 weeks post-surgery (p = 0.001). In the swing phase of gait, the maximum and minimum ankle angles were significantly smaller (i.e., more dorsiflexed) at 2 and 8 weeks post-surgery compared to baseline (p < 0.001). There were no significant differences in ankle ROM across timepoints during swing.
Results for normality of data using the Kolmogorov–Smirnov test are also shown.
In stance, the maximum MTP angle was significantly greater at 2 weeks (p = 0.040) and 8 weeks (p < 0.001) post-surgery compared to baseline and greater at 8 weeks post-surgery compared to 2 weeks post-surgery (p = 0.012) (Figure 8). The ROM was significantly greater at baseline compared to 2 weeks post-surgery (p < 0.001) and compared to 8 weeks post-surgery (p = 0.039) and significantly greater at 8 weeks post-surgery compared to 2 weeks post-surgery (p = 0.002).

3.2. Biomechanics: Pressure Data

A significant effect of group (p < 0.001) and timepoint (p < 0.001) on the peak vertical force (Figure 9A) was detected. At 2 weeks post-surgery, the peak vertical force was significantly greater in the TE group compared to the PET-SI group (p = 0.003). The peak vertical force was significantly greater at 8 weeks post-surgery compared to baseline (p < 0.001) and 2 weeks post-surgery (p < 0.001), but not significantly different at 2 weeks post-surgery compared to baseline (p = 0.711).
A significant effect of group (p = 0.004) and timepoint (p < 0.001) on vertical impulse was detected (Figure 9B). At 2 weeks post-surgery, the vertical impulse was significantly greater in the TE group compared to the PET-SI group (p = 0.001). Vertical impulse was not significantly different at 2 weeks post-surgery compared to baseline (p = 0.439) but was significantly greater at 8 weeks post-surgery compared to baseline (p < 0.001) and 2 weeks post-surgery (p < 0.001).
No significant effect of group (p = 0.641) was detected, but a significant effect of timepoint (p < 0.001) on the average ground contact area was noted (Figure 9C). The average ground contact area at 2 weeks post-surgery was significantly smaller compared to baseline (p < 0.001) and 8 weeks post-surgery (p < 0.001).

3.3. Muscle Cross-Sectional Area (CSA)

At 8 weeks post-surgery, the CSA of the LG was significantly higher in the PET-SI group than in the TE group (p = 0.006) (Figure 10). There were no significant differences in TC CSA between the TE and PET-SI groups at any timepoint. However, there were several significant differences in CSA between timepoints. For example, in the TE group, TC CSA was significantly greater at baseline compared to 8 weeks post-surgery (p = 0.022) as well as greater at 2 weeks post-surgery compared to 8 weeks post-surgery (p < 0.05). There were no significant differences in the CSAs of the TC within the PET-SI group at any timepoint (p < 0.05). Within the TE group, CSA of the LG was significantly greater at baseline compared to 2 weeks post-surgery (p = 0.010) and compared to 8 weeks post-surgery (p < 0.001), and significantly greater at 2 weeks compared to 8 weeks post-surgery (p = 0.003). Compared to baseline, the CSA of the LG was significantly smaller at 2 weeks post-surgery (p = 0.007) and at 8 weeks post-surgery (p = 0.003) in the PET-SI group.

3.4. Muscle Mass and Length

Muscle mass and length were smaller in the operated limb compared to the non-operated limb for most of the muscles across the two groups (TE and PET-SI), but with no significant between-group differences in muscle mass or length for any of the muscles. In the PET-SI group, even with the artificial tendon, the muscle mass of LG, MG, and Sol was 16.7 ± 21.0% (p = 0.503), 39.2 ± 9.1% (p < 0.001), and 53.9 ± 15.7% (p = 0.333) smaller, respectively, in the operated limb compared to the non-operated limb (Figure 11). By comparison, in the TE group, bilateral differences in muscle mass were even greater; masses of the LG, MG, and Sol in the operated limb were 36.3 ± 11.8% (p < 0.001), 50.9 ± 8.7% (p < 0.001), and 72.5 ± 7.5% (p < 0.001) smaller, respectively, compared to the non-operated limb. There was a trend of greater between-limb differences in muscle mass and length of the involved muscles (LG, MG, and Sol) in the PET-SI group compared to the TE group (Figure 11). The mass of the FD was greater in the operated limb compared to the non-operated limb in the two groups; the between-group difference in FD mass was greater in the TE group than in the PET-SI group (p < 0.05).

4. Discussion

The Achilles tendon plantarflexes the ankle joint and helps to maintain joint angle during gait because it transfers forces from the calf muscles to the foot, promoting ankle stability and effective movement [24]. It controls the rate of ankle dorsiflexion during the stance phase of locomotion, thereby maintaining proper joint angles and avoiding excessive forward tibial translation [25]. In both groups (TE and PET-SI), the ankle was more dorsiflexed in both the stance and swing phases of gait at 2 weeks and 8 weeks post-surgery compared to baseline (Figure 7). These kinematic changes in ankle angle suggest that the surgical interventions induced lower ankle plantarflexion torque, leading to a more dorsiflexed ankle angle during gait. This is not surprising since the Achilles tendon transmits forces of the triceps surae muscles, which are key contributors to ankle plantarflexion torque [26,27]. Plantarflexion weakness was apparently not overcome by the artificial Achilles tendon combined with post-surgical rehabilitation in the PET-SI group.
The muscle mass and cross-sectional area data suggest some biomechanical benefits of the artificial Achilles tendon in the PET-SI group compared to tenectomy only. Specifically, muscle cross-sectional area for the lateral gastrocnemius was significantly greater in the PET-SI group compared to the TE group (p = 0.006) (Figure 10). Additionally, the loss in muscle mass in the operated limb (compared to the non-operated limb) was quantitatively less, though not significantly so, in the PET-SI group compared to the TE group (Figure 11). These between-group differences and observations were likely due to greater loading, presumably through the artificial Achilles tendon, of the involved muscles in the PET-SI group.
Interestingly, in both groups, the average flexor digitorum (FD) muscle mass was greater in the operated limb than in the non-operated limb. This suggests that the flexor digitorum, which contributes to ankle plantarflexion based on its moment arm, compensated for weakness of the involved triceps surae muscles. Notably, the bilateral difference in FD muscle mass was significantly greater (p < 0.05) in the TE group (30.9 ± 3.3% increase) than in the PET-SI group (9.5 ± 11.7% increase); this provides further support that the artificial Achilles tendon provided some biomechanical contribution to ankle plantarflexion.
The functional benefits of the artificial Achilles tendon in this study were less than expected, and may be partly due to foreign body reaction and the accumulation of scar tissue around the implant or muscle–tendon junction [28]. The silicone coating around the braided portion of the artificial tendon was used in the original PET-SI tendon design to prevent tissue adhesion [9,10,13]. In a previous study, we observed the formation of a functional pseudo-sheath that allowed an artificial tibialis cranialis tendon to slide relative to surrounding tissues [11]. However, these results were not replicated in this study and the pseudo-sheath effect was not observed. Additionally, there was qualitatively more scar tissue around the artificial Achilles tendon and muscle–tendon junction than observed in the previous study. Scarring may have limited the transmission of muscle force and, hence, the movement of the ankle joint in the PET-SI group. The reason for the difference in pseudo-sheath and scar tissue formation between studies is unclear and should be investigated in future studies. However, features such as anatomical location, surrounding tissues, and mechanical demands distinguish the Achilles and tibialis cranialis tendons. The Achilles tendon, which courses from the gastrocnemius to the calcaneus, experiences higher tensile loads and strains due to the contribution of the triceps surae muscles to body weight support and propulsion. Consequently, the Achilles tendon is more prone to mechanical failure. In contrast, the tibialis cranialis tendon originates from the tibialis cranialis muscle, passes through a retinaculum, and inserts into the fifth metatarsal bone. Because the tibialis cranialis muscle does not contribute to body weight support and propulsion, its tendon experiences lower loads and strains and, therefore, is less prone to failure [29,30,31].
A major limitation of this study is that the mechanical properties of the PET-SI tendon have not been exhaustively quantified. Tendon properties significantly influence muscle mechanics and the muscle’s capacity to generate force [18]. Therefore, to seamlessly replace the form and function of a biological tendon with an artificial tendon, it may be important to match their mechanical properties. For instance, the stiffness of the Achilles tendon contributes to the regulation of ankle dorsiflexion during the stance phase of gait, with higher tendon stiffness being associated with improved plantar pressure distribution and enhanced balance during walking [26]. Although muscles and tendons can be studied in isolation, their unique properties interact to govern the function of the muscle–tendon actuator. Changes in tendon characteristics, such as the stress–strain relationship and slack length, are critical for fully understanding the performance of tendons.
Because the mechanical properties of the artificial Achilles tendon have not been quantified, it is difficult to interpret which specific characteristics of the implant may have contributed to the observed biomechanical or morphological outcomes. For example, under the same muscle force, a more compliant (less stiff) Achilles tendon will stretch more, which might delay ankle plantarflexion and change joint angles at toe-off, as well as the timing and magnitude of knee extension or MTP dorsiflexion [32,33]. Additionally, during the stance phase, the Achilles tendon stores and returns energy as a function of its stiffness and strain [34]. Therefore, quantifying the mechanical properties of the artificial tendon is essential and will be completed in a future study.
The study had other limitations. First, the length of the implanted artificial tendon was selected intraoperatively to be about 15% shorter than the biological tendon, with the expectation that the length of the tendon plus suture anchor would closely approximate the length of the biological tendon. However, iatrogenic tendon lengthening due to, for example, plastic deformation of the artificial tendon at first loading may have occurred. We observed such plastic deformation in some of the preliminary mechanical tensile tests, indicating that artificial tendons may need to be pre-loaded before implantation. Absolute elongation of the muscle–tendon–bone (MTB) unit (e.g., after Achilles tendon rupture or repair) is often associated with decreased plantarflexion strength, impaired heel-rise performance, and altered ankle mechanics during walking and running [35,36,37]. Patients with elongated tendons may experience deficits in strength and function, particularly in tasks that require high force generation or range of motion [35,36]. Elongation of the tendon can also cause shortening of the muscle fascicles, which may further reduce the muscle’s ability to generate force at optimal joint angles [37]. Chronically, muscles may be able to adapt to compensate for changes in the MTB unit length to preserve joint torque output [38]. Also, a small sample size may have underpowered the statistical analysis to detect small but meaningful effects. Lastly, the 8-week duration of the study was relatively short in terms of biological tendon healing. The healing of the Achilles tendon is a gradual process, and after up to 3 months, a repaired Achilles tendon typically achieves only about 65% of its pre-injury stiffness [39]. Though patients may regain sufficient range of motion for basic walking within a few months, they often continue to experience limitations in strength, endurance, and the ability to perform strenuous activities. Full recovery of tendon structure and function requires several months, with significant improvements continuing up to 6–12 months post-surgery [39,40]. In this study, we replaced the entire tendon, spanning the length between the muscle and bone insertion. As such, this study allowed investigation of functional adaptation of the muscle and the effect on limb biomechanics and kinematics. The artificial tendon would not be expected to change its material characteristics over time. Muscle contraction is essential to preservation of muscle mass and restoration of function. A longer period of study may reveal more extensive neuromuscular and functional recovery in the PET-SI group and greater between-group differences. Long-term studies would also provide insight into expected clinical outcomes in patients undergoing complete tendon replacement [41,42].
In conclusion, although the artificial Achilles tendon did not improve hindlimb kinematics in the rabbits, it preserved muscle cross-sectional area and may have mitigated the loss of muscle mass. This preservation of muscle properties is critical because it maintains the muscle’s capacity to generate force and movement. For efficient tendon force transmission and optimal recovery of biomechanical function, further improvements in the integration of artificial tendons with surrounding tissues are needed. Specifically, minimizing scar formation at the interface between the artificial tendon and the native tissue may play a pivotal role in preventing mechanical complications. With further refinement of the design, surgical implantation, tissue integration, and rehabilitation, polyester-based, silicone-coated artificial tendons may become an effective treatment option for large tendon defects.

Author Contributions

O.P.F.—animal surgery and care; data acquisition, analysis, and interpretation; manuscript drafting and revision. K.L.E.—research design; animal surgery and care; data acquisition, analysis, and interpretation. M.S.—data acquisition, analysis, and interpretation. G.B.—animal surgery and care. K.B.—animal surgery. D.E.A.—research design; animal surgery and care. D.L.C.—research design, animal surgery and care; data acquisition, analysis, and interpretation; manuscript drafting and revision. All authors have read and agreed to the published version of the manuscript.

Funding

This study was funded by the National Institutes of Health (R61AR078096, R33AR078096).

Institutional Review Board Statement

This study was conducted according to the guidelines of the Declaration of Helsinki and approved by the Institutional Animal Care and Use Committee, University of Tennessee Knoxville (protocol code #2726, 02/03/2023).

Informed Consent Statement

Not applicable.

Data Availability Statement

The original contributions presented in this study are included in the article. Further inquiries can be directed to the corresponding author(s).

Acknowledgments

We wish to acknowledge the contributions of Elizabeth Croy, Fisher Adkisson, and the staff of the Office of Laboratory Animal Care and Mossman animal housing facility at the University of Tennessee, Knoxville.

Conflicts of Interest

The authors declare no conflicts of interest.

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Figure 1. (A) Polyester silicone-coated artificial Achilles tendon prior to implantation. (B) Anika anchor used to attach artificial tendon to calcaneus.
Figure 1. (A) Polyester silicone-coated artificial Achilles tendon prior to implantation. (B) Anika anchor used to attach artificial tendon to calcaneus.
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Figure 2. Artificial Achilles tendon implanted in the left hindlimb of a rabbit.
Figure 2. Artificial Achilles tendon implanted in the left hindlimb of a rabbit.
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Figure 3. (A) Approximate locations of markers at the hip, knee, ankle, and metatarsophalangeal (MTP) joints and respective joint angles during hopping. The walkway grids surrounding the rabbit did not interfere with the capture of the reflective spherical markers as shown in (B).
Figure 3. (A) Approximate locations of markers at the hip, knee, ankle, and metatarsophalangeal (MTP) joints and respective joint angles during hopping. The walkway grids surrounding the rabbit did not interfere with the capture of the reflective spherical markers as shown in (B).
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Figure 4. Ultrasound images of the lateral gastrocnemius (left) and tibialis cranialis (right). The area bounded by the white dots represents the cross-sectional area (CSA) of each muscle.
Figure 4. Ultrasound images of the lateral gastrocnemius (left) and tibialis cranialis (right). The area bounded by the white dots represents the cross-sectional area (CSA) of each muscle.
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Figure 5. Muscles dissected from the rabbit’s hindlimbs for which mass and length were measured, including involved muscles (with direct connection to the Achilles tendon) and uninvolved muscles (without a direct connection to the Achilles tendon but flexes the ankle joint).
Figure 5. Muscles dissected from the rabbit’s hindlimbs for which mass and length were measured, including involved muscles (with direct connection to the Achilles tendon) and uninvolved muscles (without a direct connection to the Achilles tendon but flexes the ankle joint).
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Figure 6. (A,B). Maximum and minimum knee flexion (Flex) and extension (Ext) angles and range of motion (ROM) during the stance (A) and swing (B) phases of gait. * indicates a significant difference between timepoints. # indicates a significant difference between groups at specific timepoints. Letters a–e represent significant differences within groups. Differences for which p < 0.05 are considered significant. Error bars represent the standard deviation.
Figure 6. (A,B). Maximum and minimum knee flexion (Flex) and extension (Ext) angles and range of motion (ROM) during the stance (A) and swing (B) phases of gait. * indicates a significant difference between timepoints. # indicates a significant difference between groups at specific timepoints. Letters a–e represent significant differences within groups. Differences for which p < 0.05 are considered significant. Error bars represent the standard deviation.
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Figure 7. (A,B) Maximum and minimum ankle angles in plantarflexion (PF) and dorsiflexion (DF) and ankle angle range of motion (ROM) during the stance (A) and swing (B) phases of gait. * indicates a significant difference between timepoints. # indicates a significant difference between groups at specific timepoints. Letters a–e represent significant differences within groups. Differences for which p < 0.05 are considered significant. Error bars represent the standard deviation.
Figure 7. (A,B) Maximum and minimum ankle angles in plantarflexion (PF) and dorsiflexion (DF) and ankle angle range of motion (ROM) during the stance (A) and swing (B) phases of gait. * indicates a significant difference between timepoints. # indicates a significant difference between groups at specific timepoints. Letters a–e represent significant differences within groups. Differences for which p < 0.05 are considered significant. Error bars represent the standard deviation.
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Figure 8. Maximum and minimum metatarsophalangeal (MTP) joint angles in plantarflexion (PF) and dorsiflexion (DF) and MPT joint range of motion during the stance phase of gait. * indicates a significant difference between timepoints. # indicates a significant difference between groups at specific timepoints. Letters a–d represent significant differences within groups. Differences for which p < 0.05 are considered significant. Error bars represent the standard deviation.
Figure 8. Maximum and minimum metatarsophalangeal (MTP) joint angles in plantarflexion (PF) and dorsiflexion (DF) and MPT joint range of motion during the stance phase of gait. * indicates a significant difference between timepoints. # indicates a significant difference between groups at specific timepoints. Letters a–d represent significant differences within groups. Differences for which p < 0.05 are considered significant. Error bars represent the standard deviation.
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Figure 9. Mean values at baseline, B (pre-surgery) and 2 weeks and 8 weeks post-surgery for the operated limb for the TE and PET-SI groups for (A) peak vertical force (kg/BW) at maximum, minimum, and average stance, (B) vertical impulse (kg*s/BW), and (C) average ground contact area (cm2) at maximum, minimum, and average stance. Error bars represent the standard deviation. * represents significant differences between timepoints and # represents significant differences between groups within the same timepoint.
Figure 9. Mean values at baseline, B (pre-surgery) and 2 weeks and 8 weeks post-surgery for the operated limb for the TE and PET-SI groups for (A) peak vertical force (kg/BW) at maximum, minimum, and average stance, (B) vertical impulse (kg*s/BW), and (C) average ground contact area (cm2) at maximum, minimum, and average stance. Error bars represent the standard deviation. * represents significant differences between timepoints and # represents significant differences between groups within the same timepoint.
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Figure 10. Cross-sectional areas (CSAs), measured from ultrasound images, of the lateral gastrocnemius (top) and tibialis cranialis (bottom) muscles at baseline (B), 2 weeks post-surgery, and 8 weeks post-surgery. Letters a–e represent significant differences within groups (p < 0.05). Error bars represent standard deviations.
Figure 10. Cross-sectional areas (CSAs), measured from ultrasound images, of the lateral gastrocnemius (top) and tibialis cranialis (bottom) muscles at baseline (B), 2 weeks post-surgery, and 8 weeks post-surgery. Letters a–e represent significant differences within groups (p < 0.05). Error bars represent standard deviations.
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Figure 11. Mass (top) and length (bottom) of muscles in the operated limb as a percentage of values in the non-operated limb. * Bilateral differences between sides were significant (p < 0.05). LG = lateral gastrocnemius; MG = medial gastrocnemius; Sol= soleus; FD = superficial flexor digitorum; TC = tibialis cranialis; ED = extensor digitorum. Error bars represent standard deviations.
Figure 11. Mass (top) and length (bottom) of muscles in the operated limb as a percentage of values in the non-operated limb. * Bilateral differences between sides were significant (p < 0.05). LG = lateral gastrocnemius; MG = medial gastrocnemius; Sol= soleus; FD = superficial flexor digitorum; TC = tibialis cranialis; ED = extensor digitorum. Error bars represent standard deviations.
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Table 1. Rabbit demographics and biological and artificial tendon lengths for the tendon excision and excision with replacement groups. NA = not applicable; std = standard deviation.
Table 1. Rabbit demographics and biological and artificial tendon lengths for the tendon excision and excision with replacement groups. NA = not applicable; std = standard deviation.
* HousingAge (Weeks)Body Mass at Surgery (kg)Body Mass at Euthanasia (kg)Biological Tendon Length (mm)Artificial Tendon Length (mm)Percent of Biological Tendon (%)
Tendon excision (TE)-only group
TE 1P17.32.853.1540NANA
TE 2P17.42.843.3633NANA
TE 3P17.42.833.4537NANA
TE 4P, I17.43.133.7144NANA
Mean (std) 17.38 (0.04)2.91 (0.13)3.42 (0.2)38.50 (4.03)NANA
Tendon excision and replacement with polyester silicone-coated (PET-SI) tendon group
PET-SI 1I23.42.763.05353086%
PET-SI 2I18.72.783.03--27--
PET-SI 3# P, I17.32.973.44383387%
PET-SI 4P17.32.883.41353086%
PET-SI 5P, I17.43.003.45322784%
Mean (std) 18.83 (2.35)2.87 (0.10)3.28 (0.2)35 (2.12)29.40 (2.24)85.75 (1.09)
* P = pair-housed; I = individually housed; P, I = initially pair-housed, then housed individually because housemate became territorial or # due to death of housemate from anesthetic complication during surgery.
Table 2. Results (p-values) of 2-factor ANOVA with repeated measures for each kinematic variable for knee, ankle, and MTP joint angles (in degrees), during the stance and swing phases of gait, at baseline (B) and 2 and 8 weeks post-surgery.
Table 2. Results (p-values) of 2-factor ANOVA with repeated measures for each kinematic variable for knee, ankle, and MTP joint angles (in degrees), during the stance and swing phases of gait, at baseline (B) and 2 and 8 weeks post-surgery.
VariableGait phase GroupTimepoint GxT B vs. 2 B vs. 82 vs. 8
Knee
Max angleStance0.588<0.001<0.0010.007<0.0010.231
Swing0.1930.017<0.0010.2890.0150.380
Min angleStance0.525<0.001<0.001<0.001<0.0010.989
Swing0.035<0.001<0.001<0.001<0.0010.556
Avg angleStance0.704<0.001<0.001<0.001<0.0010.873
Swing0.969<0.001<0.0010.016<0.0010.532
Ankle
Max angleStance0.271<0.001<0.001<0.001<0.0010.177
Swing0.147<0.001<0.001<0.001<0.0010.912
Min angleStance0.403<0.001<0.001<0.001<0.0010.390
Swing0.822<0.001<0.001<0.001<0.0010.603
Avg angleStance0.651<0.001<0.001<0.001<0.0010.984
Swing0.564<0.001<0.001<0.001<0.0010.992
MTP
Max angleStance0.148<0.001<0.0010.040<0.0010.012
Swing------
Min angleStance0.123<0.001<0.001<0.001<0.0010.297
Swing------
Avg angleStance0.550<0.001<0.001<0.001<0.0010.486
Swing------
The results show the effect of group, timepoints, and their interaction (GxT) for each variable and each joint across the groups.
Table 3. Statistical results (p-values) for post hoc pairwise comparisons for maximum, minimum, and average joint angles for knee, ankle, and MTP joint angles (in degrees) during the stance and swing phases of gait at baseline (B) and 2 and 8 weeks post-surgery.
Table 3. Statistical results (p-values) for post hoc pairwise comparisons for maximum, minimum, and average joint angles for knee, ankle, and MTP joint angles (in degrees) during the stance and swing phases of gait at baseline (B) and 2 and 8 weeks post-surgery.
KneeAnkleMTP
GroupsVariableGait PhaseB vs. 2 B vs. 82 vs. 8 Normality B vs. 2B vs. 82 vs. 8NormalityB vs. 2B vs. 82 vs. 8Normality
TEMax angleStance0.0320.0080.1640.080<0.001<0.0010.9100.0220.0330.0620.4590.192
Swing0.1870.0620.3540.069<0.001<0.0010.8690.200----
Min angleStance<0.001<0.0010.1450.064<0.001<0.0010.7550.080<0.001<0.0010.8500.200
Swing0.008<0.0010.1320.200<0.001<0.0010.0910.012----
Avg angleStance<0.001<0.0010.1720.104<0.001<0.0010.9720.041<0.001<0.0010.5270.200
Swing0.0820.0090.1630.080<0.001<0.0010.7630.049----
PET-SIMax angleStance0.0570.0060.1270.004<0.001<0.0010.0340.1950.033<0.001<0.0010.200
Swing0.5320.0890.0520.132<0.001<0.0010.5300.200----
Min angleStance<0.0010.0050.0120.002<0.001<0.0010.0990.200<0.001<0.0010.0750.077
Swing0.0070.0020.9870.004<0.001<0.0010.6700.200----
Avg angleStance0.0090.0030.466<0.001<0.001<0.0010.7400.200<0.001<0.0010.3990.200
Swing0.0960.0360.4360.061<0.001<0.0010.8160.025----
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Fidelis, O.P.; Easton, K.L.; Smith, M.; Bastos, G.; Bowers, K.; Anderson, D.E.; Crouch, D.L. Effects of Artificial Achilles Tendon on Hindlimb Movement Biomechanics and Muscle Morphology in Rabbits. Biomechanics 2025, 5, 47. https://doi.org/10.3390/biomechanics5030047

AMA Style

Fidelis OP, Easton KL, Smith M, Bastos G, Bowers K, Anderson DE, Crouch DL. Effects of Artificial Achilles Tendon on Hindlimb Movement Biomechanics and Muscle Morphology in Rabbits. Biomechanics. 2025; 5(3):47. https://doi.org/10.3390/biomechanics5030047

Chicago/Turabian Style

Fidelis, Obinna P., Katrina L. Easton, Madison Smith, Gabriela Bastos, Kristin Bowers, David E. Anderson, and Dustin L. Crouch. 2025. "Effects of Artificial Achilles Tendon on Hindlimb Movement Biomechanics and Muscle Morphology in Rabbits" Biomechanics 5, no. 3: 47. https://doi.org/10.3390/biomechanics5030047

APA Style

Fidelis, O. P., Easton, K. L., Smith, M., Bastos, G., Bowers, K., Anderson, D. E., & Crouch, D. L. (2025). Effects of Artificial Achilles Tendon on Hindlimb Movement Biomechanics and Muscle Morphology in Rabbits. Biomechanics, 5(3), 47. https://doi.org/10.3390/biomechanics5030047

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