The effect of loading on bone, fibrous tissue, and muscle healing has been a subject of controversy since the establishment of the specialty of orthopaedics.1,2 Some of the most respected physicians and investigators concerned with the musculoskeletal system, including Nicholas André (1659-1742) and Just Lucas-Championnière (1843-1913), taught that early controlled activity promotes healing and accelerates restoration of function. Other respected authorities, including John Hunter (1728-1793), John Hilton (1807-1878), and Hugh Owen Thomas (1834-1878), advanced the opposite view.1 The latter researchers and their followers argued that absolute rest allows healing to proceed at the maximum pace, and that early use of injured tissues increases inflammation and disrupts repair tissue, thereby delaying or preventing healing. Hugh Owen Thomas emphasized that the only way a surgeon could promote healing was by giving an injured part rest, and that an overdose of rest was impossible.
A series of investigations conducted over the past several decades has resolved this controversy and provided the basis for approaches to accelerating restoration of function following musculo-skeletal surgery or injury. In this article, we will attempt to clarify the current understanding of the role of mechanical loading in the healing of musculoskeletal tissues. We will discuss the mechanisms of tissue responses to loading; the effects of loading on bone, fibrous tissue, and skeletal muscle; and the clinical implications of loading on the healing of these tissues.
Mechanisms of Tissue Response to Loading
In 1892, Julius Wolff suggested that changes in the loading of bones cause changes in their structure in accordance with mathematical laws, an observation that has become known as “Wolff's law.”3 Recent investigations have provided insight into the mechanisms responsible for Wolff's law and have shown that connective tissues other than bone also respond to changes in loading. These studies have shown that tissue loading influences cell shape, gene expression, and synthetic and proliferative functions.1,4–10 Bone, tendon, ligament, and joint capsule respond initially to loading by a mechanism of cellular detection of tissue strains, followed by modification of the tissue.
Cells may detect tissue strain directly, through deformation of the cells, or indirectly, through alterations in the matrix due to deformation of the tissue. Stretching or compression of mesenchymal cells alters the alignment of the cytoskeletal elements, changes cell shape, induces gene expression, and alters synthesis of matrix molecules
and prostaglandins.4,8,11–17 Deformation of the matrix can alter macromolecular organization, fluid flow, streaming currents, pressure gradients, or electrical fields. These matrix alterations can influence cell function.4,18–20 Loading of a tissue can also alter the flow of nutrients and metabolites by affecting vascular perfusion and diffusion through the matrix.21 Deformation, in addition to being a temporary stimulus, may also allow tissues to record and average transient strains by altering matrix macromolecular organization, thereby giving cells a more sustained and coherent stimulus. Although much progress has been made, the biologic signal transduction mechanisms that govern cellular response to load are not yet completely understood.
The mechanisms of skeletal muscle response to changes in loading differ in some respects from those of bone and fibrous tissue. The latter tissues consist primarily of mesenchymal cells and an extracellular matrix.22–24 Skeletal muscle consists of innervated muscle cells, or myofibers, and, to a lesser extent, an extracellular matrix.25 The connective tissue component of skeletal muscle has a critical role in maintaining tissue organization and stability. In addition, it contributes to the passive mechanical properties of muscle, including resistance to excessive elongation of the myofibers. Both myofibers and muscle connective tissue cells respond to injury and changes in muscle activity. The muscle connective tissue cells, like the cells in dense fibrous tissues, respond to changes in tissue loading with alterations in cell proliferation and synthetic activity that affect the organization, composition, and mechanical properties of the matrix. Unlike connective tissue cells, myofibers respond primarily to persistent changes in activity with alterations in cell structure, volume, and function. The mechanisms of their response to activity remain unclear, but they respond to both passive stretching
and active contraction.25
Maintenance of normal bone, dense fibrous tissue, and muscle structure requires a minimal level of repetitive loading. Activity below this level causes atrophy and decreases tissue organization and strength. Activity above this level can increase tissue organization and in some instances strength and mass. In general, the tissues of younger individuals are more responsive to increased levels of repetitive loading than the tissues of older individuals.26 Bone and skeletal muscle exhibit more dramatic and obvious responses to increased and decreased use, but dense fibrous tissues also respond to changes in activity level.
Response of Bone
Immobilization of a limb in a cast or in traction for a prolonged period of time causes the rate of bone resorption to exceed that of bone formation.27 Osteoclasts resorb trabecular and cortical bone, and osteoblasts fail to replace bone fast enough to maintain the mass and strength of the tissue. Decreased activity may not produce detectable changes in bone volume and shape, but prolonged immobi- lization of a limb will cause loss of bone mass. Without weight bearing, bone mass declines to less than half the normal value after 12 weeks.27 The resultant changes include decreased density of cancellous bone, loss of trabeculae, and thinning of cortical bone. These alterations decrease bone strength and increase the probability of fracture. Regular contraction of the muscles of an immobilized limb may decrease the rate of loss of bone.28 However, regaining bone density after prolonged disuse, even with vigorous activity, can take many months, even in children. In older individuals, bone density may never return to its previous level.
Persistent increases in cyclic loading of bone cause bone formation to exceed bone resorption and can result in increases in overall bone density, volume, and strength.10,29 In 1864, Sedillot found that removing the tibial diaphysis in dogs caused the fibula to increase in size sufficiently to compensate for the loss of the tibia. A more recent study in pigs provided an equally dramatic example of the ability of bone to adapt to increased loads.29 Resection of the ulnar diaphysis increased the compressive strain on the radius to 2 to 21Ú2 times the normal value. This led to a rapid increase in the diameter of the radius such that within 3 months the cross-sectional area of the radius approached the value for the radius and ulna combined, and the com-pressive strain in the radius decreased to nearly the original value.
Repetitive loading in vigorous physical activities produces similar, although less impressive, effects. For example, professional tennis players develop increased bone density, cross-sectional area, and diaphyseal diameter in the humerus of the dominant arm.30,31
Response of Dense Fibrous Tissues
Decreased loading of dense fibrous tissues that normally resist tension (tendon, ligament, and joint capsule) alters matrix turnover so that with time, matrix degradation exceeds formation. The newly synthesized matrix is less well organized, and both tissue stiffness and strength decline. Prolonged limb immobilization decreases the glycos-aminoglycan and water content, changes the degree of orientation of the matrix collagen fibrils, and may increase collagen cross-linking and decrease collagen mass.32 The duration of decreased loading necessary to produce changes varies among tissues and individuals, but most studies show marked alterations in the tissues after 6 weeks of immobilization.
Decreased loading also affects ligament, tendon, and capsular insertions into bone. The extent and severity of the alterations depend to some extent on the type of insertion. In direct insertions (e.g., those typified by the tibial and femoral insertions of the anterior cruciate ligament), most collagen fibrils pass directly into the bone matrix through a series of well-defined zones that include the substance of the tendon, ligament, or joint capsule; a zone of fibrocartilage; a zone of calcified cartilage; and the bone. In indirect, or peri-osteal, insertions (e.g., the tibial insertion of the medial collateral ligament), many of the collagen fibrils join the periosteum, and relatively few fibrils pass obliquely into the bone matrix.
Decreased ligament loading due to immobilization usually produces more extensive changes in the periosteal type of insertion. In this type of insertion, subperiosteal osteoclasts resorb much of the osseous insertion of ligaments subjected to prolonged immobilization. This leaves the ligament attached primarily to periosteum. In the direct type of insertion, resorp-tion occurs around the insertion, but relatively little resorptive activity occurs within the insertion.
The anterior cruciate and medial collateral ligaments of the knee provide examples of the differences in the response of direct and indirect insertions to immobilization. Prolonged immobilization causes bone resorption around the periphery of the cruciate ligament insertions, but only limited resorptive activity beneath the insertion site and in the zone of mineralized fibrocartilage.33,34 In contrast, prolonged immobilization causes significant diffuse resorption of the osseous part of the tibial insertion of the medial collateral ligament,35 which weakens the bone-ligament junction within 6 to 8 weeks.
After resumption of normal joint use, cells at the insertion site begin to form new bone and restore the structure and mechanical properties of the insertions toward normal. However, after 6 to 8 weeks of immobilization, complete restoration of structure and strength at the insertion site requires a longer period of loading.36–38 In one study, 6 to 8 weeks of activity after immobilization of dog knees left ligament insertions weaker than normal insertions, and the available evidence suggests that complete restoration of normal structure and mechanical properties at the ligament insertion site requires as long as 1 year of activity.33,35,39
Repetitive loading can increase the strength, size, matrix organization, and collagen content of tendons, ligaments, and their insertions into bone. In one study,40 application of tension to cultured tendons increased protein and DNA synthesis. In another study,41 12 months of exercise training increased the strength, collagen concentration, and weight of swine extensor tendons. An in vivo study in rabbits showed that localized increased loading alone causes adaptation of dense fibrous tissue.37 Insertion of a pin underneath the medial collateral ligament increased the load on the ligament by 200% to 350%, and over 12 weeks increased the strength of the boneligament complex.
Response of Skeletal Muscle
Decreased use of skeletal muscle rapidly causes changes in muscle volume, structure, and function.25,42,43 Within weeks of a reduction in frequency or intensity of activity, myofiber and myofibril volumes and oxidative capacity are reduced, causing decreases in muscle mass and strength. A decrease in intramuscular capillary density and an increase in intramuscular connective tissue volume relative to myofiber volume accompany these changes in the myofibers.44 Rigid immobilization produces more rapid and severe loss of muscle structure, volume, and function than a reduction in the frequency or intensity of activity. Muscle protein synthesis slows within 6 hours of cast immobilization of a limb.43 Two weeks of cast immobilization decreases muscle fiber size and causes loss of myofibrils. With increasing length of immobilization, the mitochondria enlarge, lose their cristae, and disintegrate.42 Eventually, the muscle cells contain only amorphous protein, vesicles, and fragments of membranes. As the myofibers degenerate, fibrous tissue and fat progressively constitute a larger proportion of the tissue.
Changes in muscle volume and function accompany these structural alterations. In one study performed in cats,42 6 weeks of cast immobilization caused a decrease in muscle weight of nearly 25%, and 22 weeks of cast immobilization caused a decrease of nearly 70%. Furthermore, the ability of muscles to generate tension decreased as muscle weight decreased. In a clinical study,45 6 weeks of cast immobilization for treatment of forearm fractures had a similar effect on muscle strength: maximal voluntary contraction of the adductor pollicis muscle decreased by 55%, and maximal electrically evoked contraction decreased by 33%.
Persistent increases in use also change the structure, functional capacity, and volume of skeletal muscle. The specific adaptive changes in muscle depend on the pattern of increased use. Patterns that produce different muscle responses include low-tension, high-repetition use, which primarily increases muscle endurance; hightension, low-repetition use, which primarily increases muscle strength; and stretching, which primarily increases muscle strength.25 Initially, muscle may respond to a training program with rapid changes in structure and function, but as adaptation occurs, the rate of change decreases, and eventually the muscle reaches a stable state.
Repetitive low-tension, high-repetition exercise, like walking, running, cycling, or swimming, performed for 30 to 60 minutes at a time, increases the capacity of muscle cells for sustained effort. This type of endurance training increases the number and size of muscle cell mitochondria, the muscle glycogen content, and the proportion of muscle cells identified as having high oxidative capacity. These changes can double the oxidative capacity of a muscle.
Strength training programs usually consist of high-tension, low-repetition exercise. These programs increase muscle strength and usually volume, primarily by causing cell hypertrophyÑthat is, by increasing the number of myofibrils.
Strength training programs generally do not increase muscle oxi-dative capacity. Stretching accelerates muscle protein turnover and can cause hypertrophy and increase strength.25 Passive motion can reduce the atrophic changes in skeletal muscle after an operative proce-dure,46 possibly by the effect of repetitive stretching of the muscle.
The Healing Process
Healing, the tissue response that can restore tissue structure and function after injury, results from a complex, interrelated series of cellular, humoral, and vascular events.47 Tissue damage and hemorrhage caused by injury or surgery initiate a response that includes inflammation (the cellular and vascular response to injury), repair (the replacement of necrotic or damaged tissue by new cells and matrix), and remodeling (the reshaping and reorganizing of repair tissue). This continuous sequence of events begins with the release of inflammatory mediators and ends when remodeling of the repair tissue reaches a homeostatic state.47 Loading can positively affect tissue healing from the repair stage through the remodeling stage.
Excessive loading and motion of a fracture can delay healing or even cause failure of healing,47 and fractures will heal under conditions of rigid immobilization. However, there is now evidence that controlled loading of a healing fracture stimulates callus formation and remodeling and accelerates restoration of bone strength. Work by Sarmiento et al48 (Fig. 1) and by Kenwright, Goodship, and co-workers49–51 has shown that early or even almost immediate loading and movement, including induced “mi-cromotion” at long-bone fracture sites, promotes fracture healing. The latter researchers51–53 have also shown that induced micromotion caused by axial loading of tibial fractures produces more rapid healing than rigid fixation (Fig. 2).
Clinical studies confirm that early controlled loading of long-bone fractures does not impair, and probably promotes, fracture heal-ing.54,55 The available evidence indicates that increasing the volume of fracture callus is one of the primary effects of loading of healing fractures.48 In addition, loading of fracture callus during the repair and remodeling phases of healing almost certainly has important effects on the organization and composition of the matrix.
Experimental work by Akeson, Gelberman, Gomez, Salter, Vailas, Woo, and others has shown that loading applied at the optimal time during repair and remodeling of dense fibrous tissue can promote healing.1,56–59 Tensile loading of tendon repair tissue causes the repair cells and matrix collagen fibrils to line up parallel to the line of tension. Lack of tension leaves the repair tissue cells and fibers disoriented. Loading alters the expression of genes responsible for the production of collagen and proteoglycan and also stimulates expression of novel genes by tendon cells. Thus, loading may affect the types of molecules synthesized during repair of fibrous tissues.60,61
In vivo studies have shown that loading accelerates tendon healing.24 Three weeks after injury, sur- gically repaired canine tendons treated with early loading had twice the strength of repaired tendons treated with immobilization (Fig. 3).57 Twelve weeks after injury, repaired tendons treated with early loading still had greater strength than repaired tendons treated with an initial period of immobilization.
Passive motion also facilitated the healing of partial and complete patellar tendon lacerations in a study in rabbits.56 Relatively little work has been done to define the effects of different loading patterns and intensities on tendon healing, but one study in dogs demonstrated that controlled passive motion of healing flexor tendons at a frequency of 12 cycles per minute for 5 minutes a day resulted in improved tensile properties compared with controlled passive motion at a frequency of 1 cycle per minute for 60 minutes a day.62
Experimental and clinical studies have shown that nonoperative treatment of medial collateral ligament injuries can result in restoration of ligament structure and function,63,64 and that loading soon after injury can accelerate ligament healing by increasing the wet and dry weights of injured ligaments, improving matrix organization, and inducing more rapid return of normal tissue DNA content, collagen synthesis, and strength (Fig. 4).59 Furthermore, a study of medial collateral ligament healing in rabbits showed that increasing the tension on healing tissue by inserting a steel pin under the ligaments stimulated significant (P<0.05) increases in ligament stiffness and strength (Fig. 5).58 This improvement in the properties of the healing ligaments also improved joint stability.
Although multiple studies have shown the beneficial effects of loading on healing of dense fibrous tissue, these effects have been demonstrated in models in which the ends of the disrupted tendon or ligament were approximated. When the ends of injured ligaments are not approximated, healing will usually be less successful in restoring the tissue structure and mechanical properties.65 For this reason, more information is needed concerning the effects of loading on healing of dense fibrous tissue injuries with substantial gaps at the injury site.
Excessive or uncontrolled loading of injured tissues disrupts repair tissue, causes further damage, and may delay or prevent repair. In a study of medial collateral ligament healing in rats, forced exercise increased the strength of ligament repair tissue in stable knees.66 However, in unstable knees, forced exercise not only did not increase the stiffness and strength of the repair tissue but actually increased joint instability. For this reason, loading and motion applied to healing dense fibrous tissue injuries must be carefully controlled.
Despite the importance of restoring muscle structure and function after injury or surgery, the effects of loading on muscle healing have not been extensively studied. The effects of activity on damaged muscle depend partially on the timing of tissue loading after injury. Immediate mobilization of injured muscles may increase scar formation and interfere with orderly regeneration of myofibers.67 Motion after a short period of immobilization produces more rapid disappearance of the hematoma and inflammatory cells; more extensive, rapid, and organized myofiber regeneration; and more rapid increase in tensile strength and stiffness.67,68
In contrast, prolonged immobilization after injury produces muscle atrophy and poor organization of the regenerating myofibers. The atrophy caused by disuse also decreases the tensile strength of the muscle and thereby increases the risk of subsequent injury.69 These results suggest that after a brief period of rest (presumably, the inflammatory and early repair phases of healing), controlled use of an injured muscle produces optimal healing.
Basic scientific investigations have shown that early controlled activity promotes healing of bone, tendon, ligament, and skeletal muscle. Much remains to be learned about the optimal application of controlled tissue loading and the effects of loading under these conditions, but several important general principles are clear. Excessive or premature loading inhibits healing, especially during the inflammatory and early repair phases, but treatment of injuries with prolonged rest delays recovery and adversely affects normal tissues. Controlled early resumption of activity that loads healing tissues during the repair and remodeling phases of healing accelerates restoration of tissue structure and function. This effect appears to result from stimulation of repaircell proliferation and matrix synthesis, as well as from increased organization of the repair-tissue matrix that improves its mechanical properties.
These general principles regarding the effects of loading on healing tissues have specific clinical implications. In particular, they imply that loading of injured bone, tendon, ligament, and skeletal muscle should be minimized during the period of acute inflammatory response that follows tissue injury. However, once this response subsides and the initial repair tissue has been formed, loading should be increased to promote further repair and remodeling.
The time required for resolution of the acute inflammatory response and formation of the initial repair tissue varies with the type of tissue, the nature of the injury, and the individual characteristics of the patient. In some minor injuries and following certain surgical procedures, there is minimal acute inflammation, and the tissues are stable; in that situation, motion and loading can be initiated almost immediately. When there is minimal tissue loss or necrosis, little disruption of the blood supply, and close approximation of the surfaces of the injured tissues, the initial repair tissue forms within 3 to 10 days. Examples of these types of injuries include closed impacted or minimally displaced metaphyseal fractures, repaired flexor tendon lacerations, medial collateral ligament tears, and skeletal muscle strains. Motion and loading of these injured tissues can be started as soon as the acute inflammation and accompanying pain have subsided. In injuries with segmental tissue loss, compromised blood supply, or large gaps between tissue surfaces, formation of the initial repair tissue requires more time. Loading before stable repair tissue has formed may adversely affect the result.
In the clinical setting, loading of healing tissues may be done with various combinations of active and passive motion, with or without resistance, or by isometric muscle contraction. Current research efforts are directed toward determining the effects of loading on specific cells and tissues, so as to provide critical information about the role of mechanical stimuli in promoting healing. However, a considerable gap exists between these studies and improved clinical use of loading to facilitate healing. At the cellular level, there is not yet sufficient understanding of mechanotrans-duction mechanisms to optimize therapeutic loading regimens. At the tissue and organ level, the relationship between motion or muscle contraction and tissue loading has not been well defined, and methods of delivering loads to healing tissue and monitoring these loads have received relatively little attention. These subjects should have central roles in future studies of methods of accelerating healing.
New approaches to promoting healing of musculoskeletal tissues are being developed, but they cannot be considered independently of the mechanical signals that stimulate and guide tissue repair and remodeling. A variety of methods, including use of cytokines, cell transplants, and gene therapy, may facilitate the formation of new tissue that has the potential to restore damaged bones, tendons, ligaments, and possibly even muscles. However, failure to take into consideration the effects of loading on tissue healing may lead to poor clinical results. For these reasons, basic science and clinical investigations of methods intended to facilitate healing should include attention to the effects of mechanical stimuli on the repair and remodeling phases of healing.
1. Buckwalter JA: Activity vs. rest in the treatment of bone, soft tissue and joint injuries. Iowa Orthop J
2. Buckwalter JA: Should bone, soft-tissue, and joint injuries be treated with rest or activity? [editorial]. J Orthop Res
3. Brand RA, Rubin CT: Fracture healing, in Albright JA, Brand RA (eds): The Scientific Basis of Orthopaedics
. Norwalk, Conn: Appleton & Lange, 1987, pp 325-345.
4. Gray ML, Pizzanelli AM, Grodzinsky AJ, Lee RC: Mechanical and physico-chemical determinants of the chondro-cyte biosynthetic response. J Orthop Res
5. Grodzinsky AJ: Electromechanical and physicochemical properties of connective tissue. Crit Rev Biomed Eng
6. Matyas JR, Anton MG, Shrive NG, Frank CB: Stress governs tissue phe-notype at the femoral insertion of the rabbit MCL. J Biomech
7. Tanaka H, Manske PR, Pruitt DL, Larson BJ: Effect of cyclic tension on lacerated flexor tendons in vitro. J Hand Surg [Am]
8. Kawata A, Mikuni-Takagaki Y: Mechanotransduction in stretched osteocytes: Temporal expression of immediate early and other genes. Biochem Biophys Res Commun
9. Frank CB, Hart DA: Cellular response to loading, in Leadbetter WB, Buckwalter JA, Gordon SL (eds): Sports Induced Inflammation: Clinical and Basic Science Concepts
. Park Ridge, Ill: American Academy of Orthopaedic Surgeons, 1990, pp 555-564.
10. Rubin C, Gross T, Qin YX, Fritton S, Guilak F, McLeod K: Differentiation of the bone-tissue remodeling response to axial and torsional loading in the turkey ulna. J Bone Joint Surg Am
11. Ingber DE: Tensegrity: The architectural basis of cellular mechanotransduction. Annu Rev Physiol
12. Toma CD, Ashkar S, Gray ML, Schaffer JL, Gerstenfeld LC: Signal transduction of mechanical stimuli is dependent on microfilament integrity: Identification of osteopontin as a mechanically induced gene in osteoblasts. J Bone Miner Res
13. Guilak F: Compression-induced changes in the shape and volume of the chondrocyte nucleus. J Biomech
14. Kim YJ, Grodzinsky AJ, Plaas AH: Compression of cartilage results in differential effects on biosynthetic pathways for aggrecan, link protein and hyaluronan. Arch Biochem Biophys
15. Buschmann MD, Gluzband YA, Grodzinsky AJ, Hunziker EB: Mechanical compression modulates matrix biosynthesis in chondrocyte/agarose culture. J Cell Sci
16. Quinn TM, Grodzinsky AJ, Buschmann MD, Kim YJ, Hunziker EB: Mechanical compression alters proteoglycan deposition and matrix deformation around individual cells in cartilage explants. J Cell Sci
17. Chen CS, Ingber DE: Tensegrity and mechanoregulation: From skeleton to cytoskeleton. Osteoarthritis Cartilage
18. Smalt R, Mitchell FT, Howard RL, Chambers TJ: Mechanotransduction in bone cells: Induction of nitric oxide and prostaglandin synthesis by fluid shear stress, but not by mechanical strain. Adv Exp Med Biol
19. Owan I, Burr DB, Turner CH, et al: Mechanotransduction in bone: Osteo-blasts are more responsive to fluid forces than mechanical strain. Am J Physiol
1997;273(3 pt 1):C810-C815.
20. Sah RLY, Kim YJ, Doong JYH, Grodzinsky AJ, Plaas AHK, Sandy JD: Biosynthetic response of cartilage explants to dynamic compression. J Orthop Res
21. Garcia AM, Frank EH, Grimshaw PE, Grodzinsky AJ: Contributions of fluid convection and electrical migration to transport in cartilage: Relevance to loading. Arch Biochem Biophys
22. Andriacchi T, Sabiston P, DeHaven K, et al: Ligament: Injury and repair, in Woo SLY, Buckwalter JA (eds): Injury and Repair of the Musculoskeletal Soft Tissues
. Park Ridge, Ill: American Academy of Orthopaedic Surgeons, 1988, pp 103-128.
23. Buckwalter JA, Glimcher MJ, Cooper RR, Recker R: Bone biology: Part I. Structure, blood supply, cells, matrix, and mineralization. J Bone Joint Surg Am
24. Gelberman R, An KN, Banes A, et al: Tendon, in Woo SLY, Buckwalter JA (eds): Injury and Repair of the Musculoskeletal Soft Tissues
. Park Ridge, Ill: American Academy of Orthopaedic Surgeons, 1988, pp 1-40.
25. Caplan A, Carlson B, Faulkner J, Fischman D, Garrett W Jr: Skeletal muscle, in Woo SLY, Buckwalter JA (eds): Injury and Repair of the Musculo-skeletal Soft Tissues
. Park Ridge, Ill: American Academy of Orthopaedic Surgeons, 1988, pp 213-291.
26. Buckwalter JA, Woo SLY, Goldberg VM, et al: Soft-tissue aging and musculoskeletal function. J Bone Joint Surg Am
27. Uhthoff HK, Jaworski ZFG: Bone loss in response to long-term immobilisation. J Bone Joint Surg Br
28. Burr DB, Frederickson RG, Pavlinch C, Sickles M, Burkart S: Intracast muscle stimulation prevents bone and cartilage deterioration in cast-immobilized rabbits. Clin Orthop
29. Goodship AE, Lanyon LE, McFie H: Functional adaptation of bone to increased stress: An experimental study. J Bone Joint Surg Am
30. Jones HH, Priest JD, Hayes WC, Tichenor CC, Nagel DA: Humeral hypertrophy in response to exercise. J Bone Joint Surg Am
31. Dalen N, Laftman P, Ohlsen H, Stromberg L: The effect of athletic activity on the bone mass in human diaphyseal bone. Orthopedics
32. Akeson WH: The response of ligaments to stress modulation and overview of the ligament healing response, in Daniel DM, Akeson WH, O'Connor JJ (eds): Knee Ligaments: Structure, Function, Injury, and Repair
. New York: Raven Press, 1990, pp 315-327.
33. Noyes FR, Torvik PJ, Hyde WB, DeLucas JL: Biomechanics of ligament failure: II. An analysis of immobilization, exercise, and reconditioning effects in primates. J Bone Joint Surg Am
34. Noyes FR, DeLucas JL, Torvik PJ: Biomechanics of anterior cruciate ligament failure: An analysis of strain-rate sensitivity and mechanisms of failure in primates. J Bone Joint Surg Am
35. Laros GS, Tipton CM, Cooper RR: Influence of physical activity on ligament insertions in the knees of dogs. J Bone Joint Surg Am
36. Amiel D, von Schroeder H, Akeson WH: The response of ligaments to stress deprivation and stress enhancement: Biochemical studies, in Daniel DM, Akeson WH, O'Connor JJ (eds): Knee Ligaments: Structure, Function, Injury, and Repair
. New York: Raven Press, 1990, pp 329-336.
37. Woo SLY, Wang CW, Newton PO, Lyon RM: The response of ligaments to stress deprivation and stress enhancement: Biomechanical studies, in Daniel DM, Akeson WH, O'Connor JJ (eds): Knee Ligaments: Structure, Function, Injury, and Repair
. New York, NY, Raven Press, 1990, pp 337-350.
38. Yasuda K, Hayashi K: Changes in biomechanical properties of tendons and ligaments from joint disuse. Osteoar-thritis Cartilage
39. Woo SLY, Gomez MA, Sites TJ, Newton PO, Orlando CA, Akeson WH: The biomechanical and morphological changes in the medial collateral ligament of the rabbit after immobilization and remobilization. J Bone Joint Surg Am
40. Slack C, Flint MH, Thompson BM: The effect of tensional load on isolated embryonic chick tendons in organ culture. Connect Tissue Res
41. Woo SLY, Ritter MA, Amiel D, et al: The biomechanical and biochemical properties of swine tendons: Long term effects of exercise on the digital extensors. Connect Tissue Res
42. Cooper RR: Alterations during immobilization and regeneration of skeletal muscle in cats. J Bone Joint Surg Am
43. Booth FW: Physiologic and biochemical effects of immobilization on muscle. Clin Orthop
44. Jozsa L, Kannus P, Thoring J, Reffy A, Jarvinen M, Kvist M: The effect of tenotomy and immobilisation on intramuscular connective tissue: A morphometric and microscopic study in rat calf muscles. J Bone Joint Surg Br
45. Duchateau J, Hainaut K: Electrical and mechanical changes in immobilized human muscle. J Appl Physiol
46. Salter RB: Muscle atrophy, in Salter RB (ed): Continuous Passive Motion (CPM)
. Baltimore: Williams & Wilkins, 1993, pp 131-135.
47. Buckwalter JA, Einhorn TA, Bolander ME, Cruess RL: Healing of the musculoskeletal tissues, in Rockwood CA Jr, Green DP, Bucholz RW, Heckman JD (eds): Fractures in Adults
, 4th ed. Philadelphia: Lippincott-Raven, 1996, vol 1, pp 261-304.
48. Sarmiento A, Schaeffer JF, Beckerman L, Latta LL, Enis JE: Fracture healing in rat femora as affected by functional weight-bearing. J Bone Joint Surg Am
49. Kenwright J, Richardson JB, Cunningham JL, et al: Axial movement and tibial fractures: A controlled randomised trial of treatment. J Bone Joint Surg Br
50. Kershaw CJ, Cunningham JL, Kenwright J: Tibial external fixation, weight bearing, and fracture movement. Clin Orthop
51. Goodship AE, Kenwright J: The influence of induced micromovement upon the healing of experimental tibial fractures. J Bone Joint Surg Br
52. Kenwright J, Goodship AE: Controlled mechanical stimulation in the treatment of tibial fractures. Clin Orthop
53. Kenwright J, Richardson JB, Goodship AE, et al: Effect of controlled axial micromovement on healing of tibial fractures. Lancet
54. Sarmiento A: A functional below-the-knee cast for tibial fractures. J Bone Joint Surg Am
55. Mooney V, Nickel VL, Harvey JP Jr, Snelson R: Cast-brace treatment for fractures of the distal part of the femur: A prospective controlled study of one hundred and fifty patients. J Bone Joint Surg Am
56. Salter RB: Tendon healing, in Salter RB (ed): Continuous Passive Motion (CPM)
. Baltimore: Williams & Wilkins, 1993, pp 136-148.
57. Gelberman RH, Woo SLY, Lothringer K, Akeson WH, Amiel D: Effects of early intermittent passive mobilization on healing canine flexor tendons. J Hand Surg [Am]
58. Gomez MA, Woo SLY, Amiel D, Harwood F, Kitabayashi L, Matyas JR: The effects of increased tension on healing medial collateral ligaments. Am J Sports Med
59. Vailas AC, Tipton CM, Matthes RD, Gart M: Physical activity and its influence on the repair process of medial collateral ligaments. Connect Tissue Res
60. Perez-Castro AV, Vogel KG: In situ expression of collagen and proteoglycan genes during development of fibrocartilage in bovine deep flexor tendon. J Orthop Res
61. Banes AJ, Horesovsky G, Larson C, et al: Mechanical load stimulates expression of novel genes in vivo and in vitro in avian flexor tendon cells. Osteoarthritis Cartilage
62. Takai S, Woo SL, Horibe S, Tung DK, Gelberman RH: The effects of frequency and duration of controlled passive mobilization on tendon healing. J Orthop Res
63. Yamaji T, Levine RE, Woo SL, Niyibizi C, Kavalkovich KW, Weaver-Green CM: Medial collateral ligament healing one year after a concurrent medial collateral ligament and anterior cruciate ligament injury: An interdisciplinary study in rabbits. J Orthop Res
64. Shelbourne KD, Porter DA: Anterior cruciate ligament-medial collateral ligament injury: Nonoperative management of medial collateral ligament tears with anterior cruciate ligament reconstruction-A preliminary report. Am J Sports Med
65. Loitz-Ramage BJ, Frank CB, Shrive NG: Injury size affects long-term strength of the rabbit medial collateral ligament. Clin Orthop
66. Burroughs P, Dahners LE: The effect of enforced exercise on the healing of ligament injuries. Am J Sports Med
67. Jarvinen MJ, Lehto MU: The effects of early mobilisation and immobilisation on the healing process following muscle injuries. Sports Med
68. Kujala UM, Orava S, Jarvinen M: Hamstring injuries: Current trends in treatment and prevention. Sports Med
69. Kaariainen M, Kaariainen J, Jarvinen TL, Sievanen H, Kalimo H, Jarvinen M: Correlation between biomechanical and structural changes during the regeneration of skeletal muscle after laceration injury. J Orthop Res