Adult New Zealand White rabbits were obtained from Bakkon Farms (Red Wing, MN) and housed with Research Animal Resources. All studies were approved by the Institutional Animal Care and Use Committee at the University of Minnesota and were in compliance with the NIH and the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research.
Rabbits were anesthetized by an intramuscular injection of 10 mg/kg ketamine and 2 mg/kg xylazine. IGF-I was injected intramuscularly into one superior rectus, randomized before surgery. Five doses were used: 1, 5, 10, 25, and 50 μg, in 100 μL sterile saline (R&D Systems, Minneapolis, MN); six rabbits were injected with each dose of IGF-I. The contralateral superior rectus muscle received an injection of an identical volume of sterile saline to serve as a control.
One week after injection, the rabbits were anesthetized with ketamine and xylazine. The chest was opened surgically, and the animals were euthanatized by thoracotomy followed by exsanguination. The superior rectus muscles of both orbits were removed from scleral insertion to their origin in the orbital apex. The muscles were placed immediately into oxygenated Ringers solution at 30°C. The muscles were carefully pinned to their in situ length, based on previous measurements in situ, and 4-0 silk was tied to the end of each muscle. Loops of suture were used to suspend the muscles in the in vitro incubation chambers, with the upper loop attached to a lever arm and force transducer. The muscles were continually bathed in oxygenated Ringers at 30°C for the duration of the experiment. Generated force in grams was recorded on a computer (model 1205 Intact Muscle Test System and Dynamic Muscle Control software; Aurora Scientific, Aurora, Ontario, Canada). Force produced by a muscle is directly related to the size of the muscle (i.e., a bigger muscle will produce more force than a smaller muscle) but may also be due to intrinsic changes in the muscle independent of size. To account for force generation due to differences in muscle size, we calculated stress, which is force divided by muscle cross-sectional area. After muscle length and mass were obtained, muscle cross-sectional area was determined by dividing muscle mass (grams) by the product of muscle length (centimeters) times a muscle density of 1.056 g/cm3. This yielded stress in grams per square centimeter, which was then converted to millinewtons per square centimeter. If differences in stress are evident between the injected and noninjected muscles, it suggests that increased force generation was due to factors other than just increased muscle size.
Both control and IGF-treated superior rectus muscles were tested simultaneously. Supramaximum stimulation intensity was determined by increasing voltage until maximum contraction was achieved using square-wave pulses of 0.5-ms duration (model 701B bi-phase current stimulator; Aurora Scientific) and delivered to the muscles via flanking platinum electrodes. Isometric length-tension curves were determined by stimulating each muscle at supramaximum intensity (500 mA, 0.5 ms), while varying the preload (resting length) over a range of 0.5 to 10.0 g. The optimal preload was determined by incrementally increasing the resting muscle length to achieve maximum isometric twitch force, allowing 60 seconds of rest between stimuli. All further tests were performed with supramaximal stimulus intensities at optimal preload. After two stabilizing tetanic stimulations (150 Hz, 500 mA, 0.5 ms) with 2 minutes rest between stimuli and 5 minutes rest after two consecutive stimulations, force development was determined for single, double, and triple pulses (0.5-ms pulse duration) with 2 minutes of rest between stimuli. Muscles were stimulated at frequencies of 10, 20, 40, 100, 150, and 200 Hz at a train duration of 500 ms with a 2-minute rest between each stimulation. After 2 minutes rest, the muscles were subjected to a fatigue protocol as follows: A tetanic stimulus was delivered every 2 seconds, consisting of a 1-second train at 150 Hz. The muscles were stimulated for 600 seconds or until there was a 50% reduction in generated muscle force. Data from treated and control muscles were pooled at each postinjection interval and compared with the paired t-test. P ≤ 0.05 was considered statistically significant.
In a second set of 12 rabbits, 10, 25, or 50 μg IGF-I in 100 μL isotonic saline was injected into one superior rectus muscle, whereas the contralateral muscle was injected with an equal volume of saline only. One week after injection, these rabbits were euthanatized with an overdose of barbiturate anesthesia, and both superior rectus muscles were removed, embedded in tragacanth gum and frozen in methylbutane that had been chilled to a slurry on liquid nitrogen. Muscles were sectioned serially at 12 μm and processed for immunohistochemical visualization of the fast, slow, developmental, and neonatal myosin heavy chain isoforms (MyHC). For pan-fast and -slow MyHC (Novocastra, Newcastle, UK) the unfixed sections were incubated with antibody at a 1:40 dilution. For developmental and neonatal MyHC, the antibody (Novocastra) was used at 1:20 dilution. The sections were blocked for nonspecific binding with horse serum and avidin-biotin blocking reagents (Vector Laboratories, Burlingame, CA), and incubated for 1 hour with the primary antibody. The sections were rinsed and incubated with the ABC reagents (Vectastain Elite; Vector Laboratories, Burlingame, CA). The reacted tissue sections were processed using the heavy metal intensified diaminobenzidine procedure.
At least four cross sections from the midbelly of each muscle were analyzed for individual myofiber cross-sectional area by manual tracing under bright-field microscopy. Between 200 and 400 myofibers from a minimum of four fields were analyzed. Mean cross-sectional areas for the myofibers were determined (Bioquant Nova Prime image analysis system; Bioquant Inc., Nashville, TN). Histograms were constructed from the cross-sectional area measurements of the individual control and IGF-treated muscles. The cross-sectional areas were divided into bins of 200-μm increments. The percentage of myofibers positive for the neonatal and developmental MyHC was determined in muscles treated with either 25 or 50 μg IGF-I. These represent the most effective doses and the doses where force began to decrease experimentally, although still significantly greater than control. Care was taken to analyze sections from midbelly region and at the tendon end, since the percentage of myofibers positive for neonatal and developmental MyHC changes along the tendon-to-tendon muscle length.
13 In a second group of control rabbits, normal superior rectus, and tibialis muscles were obtained as previously described and immunostained with an antibody to insulin-like growth factor receptor (IGF-R; 1:10; Abcam, Cambridge, MA) or double immunostained for IGF-R and laminin (1:40; Vector Laboratories) as published previously.
14 All data were analyzed for statistical significance using analysis of variance (ANOVA) and the Dunn multiple comparison tests (Prism and Statmate software; GraphPad, San Diego, CA, or SigmaStat 2.03; SPSS Science, Chicago, IL). An F-test was used to verify that the variances were not significantly different. Data were considered significantly different if
P ≤ 0.05.