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Schwann cells support extensive axonal growth into skeletal muscle implants in adult mouse brain
Neuroscience Letters, 93 (1988) 127 131 Elsevier ScientificPublishers Ireland Ltd.
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Schwann cells support extensive axonal growth into skeletal muscle implants in adult mouse brain P.N. Anderson, M. Turmaine and P. W o o d h a m Department of Anatomy and DevelopmentalBiology, The University College and Middlesex School of Medicine, London ( U.K.) (Received 12 April 1988; Revised version received 20 June 1988; Accepted 21 June 1988) Key word~." Axonal regeneration; Axotomy; Skeletal muscle; Schwann cell; Brain implant; Mouse The ability of striated muscle to support CNS axonal regeneration was tested by grafting pieces of the lateral rectus muscle of the orbit into the hippocampus or neocortex of adult inbred CBA mice. The mice were perfused with fixative 4-5 weeks after operation and ultrathin sections of the grafts examined by electron microscopy. Many axons were present in the grafts and some were traced into the surrounding brain tissue. Most axons were in contact with Schwann cells, or their processes, and both were often associated with basal lamina material left behind by degenerating muscle cells. A few axons and their accompanying Schwann cells were found in contact with the plasma membrane of muscle cells. Fenestrated capillaries were present in the grafts. It is suggested that Schwann cells form the substratum for axonal extension into muscle implants in the CNS, although other factors may contribute to the extensiveaxonal invasion of the tissue.
I n j u r e d a x o n s in the m a m m a l i a n C N S will regenerate into segments of peripheral nerve i m p l a n t e d into the b r a i n [1]. It is c o n s e q u e n t l y widely believed that peripheral nerves constitute a n e n v i r o n m e n t highly f a v o u r a b l e to axonal regeneration, perhaps because of their c o n t e n t o f S c h w a n n cells. However, previous light microscopical studies have s h o w n that axons will also elongate into other tissues i m p l a n t e d into the b r a i n [8, 9, 14] i n c l u d i n g skeletal muscle. T e r m i n a l sprouts of u n d a m a g e d axons within intact muscles are t h o u g h t to be able to elongate slowly across the surface of skeletal muscle cells [5] b u t the s u b s t r a t u m used by the nerve fibres growing into muscle i m p l a n t e d into b r a i n has n o t been identified. Since knowledge o f the range of cells a n d tissues which are capable o f s u p p o r t i n g axonal regeneration is o f considerable neurobiological interest, an u l t r a s t r u c t u r a i study was m a d e o f skeletal muscle tissue i m p l a n t e d into m o u s e brain. A d u l t female i n b r e d C B A mice were used t h r o u g h o u t . D o n o r a n i m a l s were killed
Correspondence." P.N. Anderson, Department of Anatomy and Developmental Biology, The University College and Middlesex School of Medicine, London WlP 6DB, U.K. 0304-3940/88/$ 03.50 O 1988 ElsevierScientificPublishers Ireland Ltd.
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with ether and the lateral rectus muscle dissected from each orbit. The muscles were placed in sterile H a n k ' s balanced salt solution (Gibco) and any extramuscular peripheral nerve cut off with a scalpel. Host animals were anaesthetised with Sagatal (30 mg/kg) and ether. The scalp was opened with a sagittal incision, and a burr hole approximately 1.5 mm in diameter was made in the right parietal bone. The dura was opened and an isolated lateral rectus muscle was inserted into the brain using a fine tungsten needle. The scalp wound was sutured with 3/0 silk. After 4 (one animal) or 5 weeks (4 animals) the animals were perfused under deep anaesthesia with a fixative containing 4% glutaraldehyde and 2% paraformaldehyde in 0.1 M cacodylate buffer pH 7.4. The brain was sliced with a razor blade and those segments containing the implant were processed for electron microscopy. Ultra-thin sections were examined using a Philips EM300 electron microscope. The implants in all 5 animals were very similar although two were confined to the hippocampus, two extended from the hippocampus into the neocortex and one was entirely in the neocortex. Large mature muscle cells were rare, but many immature forms were found, often with centrally located nuclei. Such cells were easily identifiable by their myofibrillar structure (Fig. 1). Large amounts of basal lamina material left behind by degenerated muscle cells was identified, much of which was highly folded (Fig. 1). The grafts were vascularized and contained fenestrated capillaries, as does intact skeletal muscle. Many axons had invaded the grafts. The majority of axons in the grafts were in contact with Schwann cell processes (Figs. i and 2), which appeared to be more abundant than in intact muscle. Whether all axons were associated with Schwann cells remains uncertain, however, because axons cannot always be differentiated from Schwann cell processes using morphological criteria alone [16]. Most axons were unmyelinated but a few profiles surrounded by PNS myelin were also seen. A few Schwann cells and axons were seen on the surface of muscle cells, without an intervening basal lamina. Some axons were followed into the neuropil of the brain surrounding the grafts. Schwann cells and axons were very widely distributed within the implants, and were not limited to the course of the pre-existing intramuscular nerves, which could be identified because of their fascicular structure. Most axons and their accompanying Schwann cells were in contact with the extensive regions of basal lamina left behind by degenerated muscle cells (Fig. 1). Many more axons were present in the striated muscle implants than were normally found in peroneal nerve implants performed in a similar manner (unpublished observations).
Fig. 1. Electron micrograph of part of a lateral rectus muscle implanted into mouse cerebral cortex [~r 5 weeks. A bundle of axons (arrows) and Schwann cell processes (arrowheads), associated with an exlensive region of basal lamina material (B) may be seen near an immature muscle cell (M). Bar = 0.4 g~m. Fig. 2. An axon (A) may be seen invaginated into the perinuclear region of a Schwann cell within a four week skeletal muscle implant in mouse cortex. B a r - 0 . 6 ~m.
130 The present study has confirmed the ability of nerve fibres to invade pieces of skeletal muscle implanted into the brain, in this case the hippocampus and neocortex. Previous studies have produced varying results as to the numbers of nerve fibres which grew into skeletal muscle implants; although extensive ingrowth was reported by some workers [9, 14], others found that muscle contained fewer axons than did implants of thyroid gland or skin [8]. The present ultrastructural observations extend previous studies by showing that the invading axons were associated with Schwann cells which were situated throughout the implants. These Schwann cells were presumably derived from the population which existed inside the muscle at the time of operation. Although the origin of the axons is not known it is probable that some at least might have arisen in the CNS, particularly those that were traced into the neuropil surrounding the grafts. Axons which invade salivary gland tissue implanted into the spinal cord of young mice are also associated with Schwann cells [10]. It is reasonable to propose that the ready growth of many axons into striated muscle implants in the mammalian brain might be at least in part the result of the presence within the implants of large numbers of Schwann cells. Certainly, peripheral nerve implants, which lack striated muscle cells, also promote axonal regeneration [1]. Regenerating axons in vivo are nearly always found in contact with Schwann cells or other suitable glia [2-4, 7, 10, 13] rather than available non-glial cells or extracellular matrix components. However, it is also possible that other factors also promoted axonal elongation into the skeletal muscle implants. Trophic factors capable of influencing some CNS neurons are produced by skeletal muscle tissue [6]. Production of these or other neuronotrophic or tropic factors might explain the greater abundance of axons in muscle implants than in peroneal nerve implants done in the same manner, although other influences might also be implicated. The absence of a perineurium around muscle might make it easier for regenerating axons to penetrate the graft. Muscle basal laminae have been reported to promote axonal regeneration in the PNS [11]. It may also be germane that the skeletal muscle implants retained their fenestrated capillary bed. This may be the morphological representation of the high degree of vascular permeability found in muscle implants [8]. In the PNS regenerating axons are often associated with fenestrated capillaries [2-4] or regions of high vascular permeability [15] and it has been suggested that highly permeable vessels may be one prerequisite for prolonged axonal regeneration [12]. Although many axons invaded the implants, it is not known how far individual axons extended through the muscle tissue. Furthermore the axons and Schwann cells within the grafts were not arranged in the orderly manner of axons confined within longitudinally orientated bands of von Bungner of peripheral nerve implants. Hence it should not be assumed that muscle tissue is more suitable than peripheral nerve for promoting functional regeneration in the injured CNS. The authors gratefully acknowledge support from Action Research for the Cripoled Child.
131 1 Aguayo, A.J., Axon regeneration from injured neurons in the adult mammalian central nervous system. In C.W. Cotman (Ed.), Synaptic Plasticity, Guilford, New York, 1985, pp. 457-484. 2 Anderson, P.N., Mitchell, J., Mayor, D. and Stauber, V.V., An ultrastructural study of the early stages of axonal regeneration through rat nerve grafts, Neuropathol. Appl.'Neurobiol., 9 (1983) 455-466. 3 Anderson, P.N. and Turmaine, M., Peripheral nerve regeneration through grafts of living and freezedried CNS tisue, Neuropathol. Appl. Neurobiol., 12 (1986) 389-399. 4 Anderson, P.N. and Turmaine, M., Peripheral nerve fibres regenerate through myenteric plexus, Neurosci. Lett., 76 (1987) 129 132. 5 Brown, M.C. and Ironton, R., Sprouting and regression of neuromuscular synapses in partially denervated mammalian muscles, J. Physiol. (Lond.), 278 (1978) 325-348. 6 Doherty, P., Dickson, J.G., Flanigan, T.P. and Walsh, F.S., Human skeletal muscle cells synthesize a neuronotrophic factor reactive with spinal neurons, J. Neurochem., 46 (1986) 133-139. 7 Hall, S.M., Regeneration in cellular and acellular autografts in the peripheral nervous system, Neuropathol. Appl. Neurobiol., 12 (1986) 27-46. 8 Heinicke, E.A., Vascular permeability and axonal regeneration in tissues autotransplanted into the brain, Acta Neuropathol., 49 (1980) 177-185. 9 Horvat, J.-C., R6actions r6g6n6ratives provoqu6es dans le cervelet de la souris par des greffes tissulaires homoplastiques et br6phoplastiques, Arch. Sci. Physiol., 21 (1967) 323-343. 10 Horvat, J.-C., Aspects ultrastructuraux de la r6habitation de fragments de glande sous-maxillaire transplant6s dans la moelle 6pin6re de la souris par des fibres nerveuses d'origine centrale, C.R. Assoc. Anat., 54 (1969) 218-230. 11 Keynes, R.J., Hopkins, W.G. and Huang, L.-H., Regeneration of mouse peripheral nerves in degenerating skeletal muscle: guidance by residual muscle fibre basement membranes, Brain Res., 295 (1984) 275-281. 12 Kiernan, J.A., Hypotheses concerned with axonal regeneration in the mammalian nervous system, Biol. Rev., 54 (1979) 155-197. 13 Nathanial, E.J.H. and Clemente, C.D., Growth of nerve fibres into skin and muscle grafts in rat brain, Exp. Neurol., 1 (1959) 65-81. 14 Smith, G.M., Miller, R.H. and Silver, J., Changing role of forebrain astrocytes during development, regenerative failure and induced regeneration upon transplantation, J. Comp. Neurol., 251 (1986) 23 43. 15 Sparrow, J.R. and Kiernan, J.A., Endoneurial vascular permeability in degenerating and regenerating peripheral nerves, Acta Neuropathol., 53 (1981) 181-188. 16 Tohyama, K., Lieberman, A.R. and lde, C., Immunohistochemical studies of peripheral nerve regeneration. In T. lmura, L.S. Maruse and T. Suzuki (Eds.), Electron Microscopy, Vol. IV, The Japanese Society of Electron Microscopy, Tokyo, 1986, pp. 309-310.
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Schwann cells support extensive axonal growth into skeletal muscle implants in adult mouse brain P.N. Anderson, M. Turmaine and P. W o o d h a m Department of Anatomy and DevelopmentalBiology, The University College and Middlesex School of Medicine, London ( U.K.) (Received 12 April 1988; Revised version received 20 June 1988; Accepted 21 June 1988) Key word~." Axonal regeneration; Axotomy; Skeletal muscle; Schwann cell; Brain implant; Mouse The ability of striated muscle to support CNS axonal regeneration was tested by grafting pieces of the lateral rectus muscle of the orbit into the hippocampus or neocortex of adult inbred CBA mice. The mice were perfused with fixative 4-5 weeks after operation and ultrathin sections of the grafts examined by electron microscopy. Many axons were present in the grafts and some were traced into the surrounding brain tissue. Most axons were in contact with Schwann cells, or their processes, and both were often associated with basal lamina material left behind by degenerating muscle cells. A few axons and their accompanying Schwann cells were found in contact with the plasma membrane of muscle cells. Fenestrated capillaries were present in the grafts. It is suggested that Schwann cells form the substratum for axonal extension into muscle implants in the CNS, although other factors may contribute to the extensiveaxonal invasion of the tissue.
I n j u r e d a x o n s in the m a m m a l i a n C N S will regenerate into segments of peripheral nerve i m p l a n t e d into the b r a i n [1]. It is c o n s e q u e n t l y widely believed that peripheral nerves constitute a n e n v i r o n m e n t highly f a v o u r a b l e to axonal regeneration, perhaps because of their c o n t e n t o f S c h w a n n cells. However, previous light microscopical studies have s h o w n that axons will also elongate into other tissues i m p l a n t e d into the b r a i n [8, 9, 14] i n c l u d i n g skeletal muscle. T e r m i n a l sprouts of u n d a m a g e d axons within intact muscles are t h o u g h t to be able to elongate slowly across the surface of skeletal muscle cells [5] b u t the s u b s t r a t u m used by the nerve fibres growing into muscle i m p l a n t e d into b r a i n has n o t been identified. Since knowledge o f the range of cells a n d tissues which are capable o f s u p p o r t i n g axonal regeneration is o f considerable neurobiological interest, an u l t r a s t r u c t u r a i study was m a d e o f skeletal muscle tissue i m p l a n t e d into m o u s e brain. A d u l t female i n b r e d C B A mice were used t h r o u g h o u t . D o n o r a n i m a l s were killed
Correspondence." P.N. Anderson, Department of Anatomy and Developmental Biology, The University College and Middlesex School of Medicine, London WlP 6DB, U.K. 0304-3940/88/$ 03.50 O 1988 ElsevierScientificPublishers Ireland Ltd.
128
with ether and the lateral rectus muscle dissected from each orbit. The muscles were placed in sterile H a n k ' s balanced salt solution (Gibco) and any extramuscular peripheral nerve cut off with a scalpel. Host animals were anaesthetised with Sagatal (30 mg/kg) and ether. The scalp was opened with a sagittal incision, and a burr hole approximately 1.5 mm in diameter was made in the right parietal bone. The dura was opened and an isolated lateral rectus muscle was inserted into the brain using a fine tungsten needle. The scalp wound was sutured with 3/0 silk. After 4 (one animal) or 5 weeks (4 animals) the animals were perfused under deep anaesthesia with a fixative containing 4% glutaraldehyde and 2% paraformaldehyde in 0.1 M cacodylate buffer pH 7.4. The brain was sliced with a razor blade and those segments containing the implant were processed for electron microscopy. Ultra-thin sections were examined using a Philips EM300 electron microscope. The implants in all 5 animals were very similar although two were confined to the hippocampus, two extended from the hippocampus into the neocortex and one was entirely in the neocortex. Large mature muscle cells were rare, but many immature forms were found, often with centrally located nuclei. Such cells were easily identifiable by their myofibrillar structure (Fig. 1). Large amounts of basal lamina material left behind by degenerated muscle cells was identified, much of which was highly folded (Fig. 1). The grafts were vascularized and contained fenestrated capillaries, as does intact skeletal muscle. Many axons had invaded the grafts. The majority of axons in the grafts were in contact with Schwann cell processes (Figs. i and 2), which appeared to be more abundant than in intact muscle. Whether all axons were associated with Schwann cells remains uncertain, however, because axons cannot always be differentiated from Schwann cell processes using morphological criteria alone [16]. Most axons were unmyelinated but a few profiles surrounded by PNS myelin were also seen. A few Schwann cells and axons were seen on the surface of muscle cells, without an intervening basal lamina. Some axons were followed into the neuropil of the brain surrounding the grafts. Schwann cells and axons were very widely distributed within the implants, and were not limited to the course of the pre-existing intramuscular nerves, which could be identified because of their fascicular structure. Most axons and their accompanying Schwann cells were in contact with the extensive regions of basal lamina left behind by degenerated muscle cells (Fig. 1). Many more axons were present in the striated muscle implants than were normally found in peroneal nerve implants performed in a similar manner (unpublished observations).
Fig. 1. Electron micrograph of part of a lateral rectus muscle implanted into mouse cerebral cortex [~r 5 weeks. A bundle of axons (arrows) and Schwann cell processes (arrowheads), associated with an exlensive region of basal lamina material (B) may be seen near an immature muscle cell (M). Bar = 0.4 g~m. Fig. 2. An axon (A) may be seen invaginated into the perinuclear region of a Schwann cell within a four week skeletal muscle implant in mouse cortex. B a r - 0 . 6 ~m.
130 The present study has confirmed the ability of nerve fibres to invade pieces of skeletal muscle implanted into the brain, in this case the hippocampus and neocortex. Previous studies have produced varying results as to the numbers of nerve fibres which grew into skeletal muscle implants; although extensive ingrowth was reported by some workers [9, 14], others found that muscle contained fewer axons than did implants of thyroid gland or skin [8]. The present ultrastructural observations extend previous studies by showing that the invading axons were associated with Schwann cells which were situated throughout the implants. These Schwann cells were presumably derived from the population which existed inside the muscle at the time of operation. Although the origin of the axons is not known it is probable that some at least might have arisen in the CNS, particularly those that were traced into the neuropil surrounding the grafts. Axons which invade salivary gland tissue implanted into the spinal cord of young mice are also associated with Schwann cells [10]. It is reasonable to propose that the ready growth of many axons into striated muscle implants in the mammalian brain might be at least in part the result of the presence within the implants of large numbers of Schwann cells. Certainly, peripheral nerve implants, which lack striated muscle cells, also promote axonal regeneration [1]. Regenerating axons in vivo are nearly always found in contact with Schwann cells or other suitable glia [2-4, 7, 10, 13] rather than available non-glial cells or extracellular matrix components. However, it is also possible that other factors also promoted axonal elongation into the skeletal muscle implants. Trophic factors capable of influencing some CNS neurons are produced by skeletal muscle tissue [6]. Production of these or other neuronotrophic or tropic factors might explain the greater abundance of axons in muscle implants than in peroneal nerve implants done in the same manner, although other influences might also be implicated. The absence of a perineurium around muscle might make it easier for regenerating axons to penetrate the graft. Muscle basal laminae have been reported to promote axonal regeneration in the PNS [11]. It may also be germane that the skeletal muscle implants retained their fenestrated capillary bed. This may be the morphological representation of the high degree of vascular permeability found in muscle implants [8]. In the PNS regenerating axons are often associated with fenestrated capillaries [2-4] or regions of high vascular permeability [15] and it has been suggested that highly permeable vessels may be one prerequisite for prolonged axonal regeneration [12]. Although many axons invaded the implants, it is not known how far individual axons extended through the muscle tissue. Furthermore the axons and Schwann cells within the grafts were not arranged in the orderly manner of axons confined within longitudinally orientated bands of von Bungner of peripheral nerve implants. Hence it should not be assumed that muscle tissue is more suitable than peripheral nerve for promoting functional regeneration in the injured CNS. The authors gratefully acknowledge support from Action Research for the Cripoled Child.
131 1 Aguayo, A.J., Axon regeneration from injured neurons in the adult mammalian central nervous system. In C.W. Cotman (Ed.), Synaptic Plasticity, Guilford, New York, 1985, pp. 457-484. 2 Anderson, P.N., Mitchell, J., Mayor, D. and Stauber, V.V., An ultrastructural study of the early stages of axonal regeneration through rat nerve grafts, Neuropathol. Appl.'Neurobiol., 9 (1983) 455-466. 3 Anderson, P.N. and Turmaine, M., Peripheral nerve regeneration through grafts of living and freezedried CNS tisue, Neuropathol. Appl. Neurobiol., 12 (1986) 389-399. 4 Anderson, P.N. and Turmaine, M., Peripheral nerve fibres regenerate through myenteric plexus, Neurosci. Lett., 76 (1987) 129 132. 5 Brown, M.C. and Ironton, R., Sprouting and regression of neuromuscular synapses in partially denervated mammalian muscles, J. Physiol. (Lond.), 278 (1978) 325-348. 6 Doherty, P., Dickson, J.G., Flanigan, T.P. and Walsh, F.S., Human skeletal muscle cells synthesize a neuronotrophic factor reactive with spinal neurons, J. Neurochem., 46 (1986) 133-139. 7 Hall, S.M., Regeneration in cellular and acellular autografts in the peripheral nervous system, Neuropathol. Appl. Neurobiol., 12 (1986) 27-46. 8 Heinicke, E.A., Vascular permeability and axonal regeneration in tissues autotransplanted into the brain, Acta Neuropathol., 49 (1980) 177-185. 9 Horvat, J.-C., R6actions r6g6n6ratives provoqu6es dans le cervelet de la souris par des greffes tissulaires homoplastiques et br6phoplastiques, Arch. Sci. Physiol., 21 (1967) 323-343. 10 Horvat, J.-C., Aspects ultrastructuraux de la r6habitation de fragments de glande sous-maxillaire transplant6s dans la moelle 6pin6re de la souris par des fibres nerveuses d'origine centrale, C.R. Assoc. Anat., 54 (1969) 218-230. 11 Keynes, R.J., Hopkins, W.G. and Huang, L.-H., Regeneration of mouse peripheral nerves in degenerating skeletal muscle: guidance by residual muscle fibre basement membranes, Brain Res., 295 (1984) 275-281. 12 Kiernan, J.A., Hypotheses concerned with axonal regeneration in the mammalian nervous system, Biol. Rev., 54 (1979) 155-197. 13 Nathanial, E.J.H. and Clemente, C.D., Growth of nerve fibres into skin and muscle grafts in rat brain, Exp. Neurol., 1 (1959) 65-81. 14 Smith, G.M., Miller, R.H. and Silver, J., Changing role of forebrain astrocytes during development, regenerative failure and induced regeneration upon transplantation, J. Comp. Neurol., 251 (1986) 23 43. 15 Sparrow, J.R. and Kiernan, J.A., Endoneurial vascular permeability in degenerating and regenerating peripheral nerves, Acta Neuropathol., 53 (1981) 181-188. 16 Tohyama, K., Lieberman, A.R. and lde, C., Immunohistochemical studies of peripheral nerve regeneration. In T. lmura, L.S. Maruse and T. Suzuki (Eds.), Electron Microscopy, Vol. IV, The Japanese Society of Electron Microscopy, Tokyo, 1986, pp. 309-310.