Добавил:
Sekretar
kiopkiopkiop18@yandex.ru
t.me/Prokururor I Вовсе не секретарь, но почту проверяю
Опубликованный материал нарушает ваши авторские права? Сообщите нам.
Вуз:
Предмет:
Файл:Ординатура / Хирургия / Библиотека им академика М.И. Перельмана / Книга_531_Библиотеки_им_академика_М_И_Перельмана
.pdf
Figure 2). Mechanical lesions lead to high, localized levels of membrane integrity loss, tension
change and Ca2+ inux. These physical tears of the plasma membrane are often repaired by
targeted exocytosis and endocytosis. Contrastingly, smaller injuries such as those generated
by electroporation and osmotic shock induce low levels of membrane disruption, tension
change and Ca2+ inux across large membrane areas. These in turn facilitate processes such as
cytoskeletal remodeling or caveolae aening. Conversely, membranes disrupted by toxic
pores do not lead to substantial increase in plane tension. As such, they can either be rapidly
shed or degraded following caveolae-mediated endocytosis. Furthermore, it appears that the
wound-healing mechanisms prevalent in a given cell-type fall not only in accordance with the
prevalence of specic injury types (i.e., PFTs vs. tears vs. ablations), but also according to cell
type-specic dierences in cell tensegrity and polarity (e.g., muscle cells vs. epithelial cells).
Similar to the plasma membrane and cytoskeletal elements interact to create tensegrity in the
single-cell scale, adhesive forces of single cells and the extracellular matrix (ECM) provide
structural stiness to tissues [1]. Considering the above, it should be no surprise that successful
single-cell repair inuences the success of tissue repair. Indeed, contrary to tissue repair, single-
cell repair is largely a binary event: it either takes place allowing the cell’s survival, or not,
leading to lysis or apoptotic removal. While relevant to wound healing at the tissue-level, these
events have lile to no relevance for single-cell wound healing outside of the modication of
the environment of other injured cells in the surrounding area (asymmetric binding, change
in ROS, Ca2+ concentration, etc.). Conversely, it seems that successful repair in one cell may
lead to an increased repair potential in surrounding cells [175, 176]. This “potentiated” repair
has been shown to involve purinergic and nitric oxide (NO)/PKG-signaling pathways [175,
176]. Similarly, repeated insults to a cell’s structural and membrane integrity presumably aect
a cell’s ability to undergo subsequent membrane resealing and cytoskeletal repair, which
would be reected in its long-term viability in a given tissue. Indeed, the prominent view of
the origin of the phenotypes associated with muscular dystrophies point toward a heighted
susceptibly to repeated mechanical wounding, leading in turn in a higher rate of single-cell
repair failure (reviewed in [177, 178]).
Another parallel between single-cell and tissue wound-healing mechanisms is their reliance
on contractile arrays. This similarity has been conrmed in multicellular models such as
Xenopus embryos [179], Caco-2 intestinal epithelial monolayers [180] and Madin-Darby canine
kidney (MDCK) epithelial monolayers [181].
Wound closure in epithelial sheets has been demonstrated to be driven by the coupling of
actomyosin contraction and collective cell migration [182–185]. The relative contribution of
each mechanism in overall re-epithelialization depends on numerous biomechanical factors,
including wound geometry [182–184], wound size [182, 186], tissue stiness [186] and ECM
composition [182]. In particular, wounds of cultured bovine corneal endothelial cell monolay-
ers in ECM-deprived conditions were observed to reseal predominantly through actomyosin
activity [182]. This is intriguing since cytoskeletal dynamics greatly inuence single-cell
wound-healing processes (see Section 3.3) and exhibits the ECM and cytoskeleton’s analogous
relationship across biological scales in the context of wound healing. These observations
How Plasma Membrane and Cytoskeletal Dynamics Influence Single-Cell Wound Healing: Mechanotransduction,
Tension and Tensegrity
http://dx.doi.org/10.5772/63765
213

suggest that the importance of tensegrity components in wound repair are conserved across
single-cell and multicellular models.
Considering the single cell’s tensegral context in future wound-healing study will help further
characterize an increasingly complex unied pathway theory of plasma membrane repair.
Author details
Eric Boucher, Tatsuya Kato and Craig A. Mandato
*
*Address all correspondence to: craig.mandato@mcgill.ca
Department of Anatomy and Cell Biology, Faculty of Medicine, McGill University, Montreal,
Quebec, Canada
References
[1] Ingber, D.E., N. Wang, and D. Stamenovic, Tensegrity, cellular biophysics, and the mechan-
ics of living systems. Rep Prog Phys, 2014. 77(4): p. 046603.
[2] Fuller, R.B., Tensegrity. Portfolio Art News Annu, 1961. 4: pp. 112–27, 144, 148.
[3] Wang, N., et al., Mechanical behavior in living cells consistent with the tensegrity model. Proc
Natl Acad Sci U S A, 2001. 98(14): pp. 7765–70.
[4] Mendez, M.G., D. Restle, and P.A. Janmey, Vimentin enhances cell elastic behavior and
protects against compressive stress. Biophys J, 2014. 107(2): pp. 314–23.
[5] Chen, C., et al., Eects of vimentin disruption on the mechanoresponses of articular chondro-
cyte. Biochem Biophys Res Commun, 2016. 469(1): pp. 132–7.
[6] Diz-Munoz, A., D.A. Fletcher, and O.D. Weiner, Use the force: membrane tension as an
organizer of cell shape and motility. Trends Cell Biol, 2013. 23(2): pp. 47–53.
[7] Xue, F., et al., Eect of membrane stiness and cytoskeletal element density on mechanical
stimuli within cells: an analysis of the consequences of ageing in cells. Comput Methods
Biomech Biomed Engin, 2015. 18(5): pp. 468–76.
[8] Knoll, R., A role for membrane shape and information processing in cardiac physiology.
Pugers Arch, 2015. 467(1): pp. 167–73.
[9] Chaerjee, S., et al., Shear stress-related mechanosignaling with lung ischemia: lessons from
basic research can inform lung transplantation. Am J Physiol Lung Cell Mol Physiol, 2014.
307(9): pp. L668–80.
Wound Healing: New insights into Ancient Challenges214

model for congenital diaphragmatic hernia: a randomized study. PLoS One, 2013. 8(7): p.
e69210.
model. Biochem Cell Biol, 2007. 85(5): pp. 543–51.
membrane interactions in phospholipid systems. Ciba Found Symp, 1984. 103: pp. 9–27.
pore size and pore line tension. Biochim Biophys Acta, 1993. 1147(1): pp. 89–104.
326: pp. 1–15.
(Basel), 2011. 11(2): pp. 1744–55.
high temporal resolution. Biophys J, 2005. 88(2): pp. 1143–55.
Journal-Special Topics, 2014. 223(9): pp. 1813–1829.
cholesterol-depleted erythrocyte membrane. Biochim Biophys Acta, 1980. 597(1): pp. 155–
65.
membrane: eects of protein-lipid interactions. Biochemistry, 1984. 23(2): pp. 332–9.
membranes. Can J Biochem Cell Biol, 1984. 62(8): pp. 752–9.
of divalent cations and spectrin release. Biochim Biophys Acta, 1978. 509(1): pp. 58–66.
repair. Proc Natl Acad Sci U S A, 2003. 100(8): pp. 4592–7.
depletion in capacitating sperm cells. Mol Membr Biol, 2007. 24(4): pp. 313–24.
scaolding proteins exhibits high similarities in neurons and mammalian spermatozoa.
Commun Integr Biol, 2010. 3(6): pp. 513–21.
sperm. Reproduction, 2015. 150(2): pp. R65–76.
How Plasma Membrane and Cytoskeletal Dynamics Influence Single-Cell Wound Healing: Mechanotransduction,
http://dx.doi.org/10.5772/63765
Tension and Tensegrity
215

protein insertion into membranes and lamellar body secretion. Biochim Biophys Acta, 2013.
1833(5): pp. 1244–55.
Lung Cell Mol Physiol, 2006. 291(4): pp. L596–601.
prerequisite for content release or kiss-and-run. Ann N Y Acad Sci, 2009. 1152: pp. 43–52.
recruitment, fusion mode and vesicle recycling in neuroendocrine cells. J Neurochem, 2016.
DOI: 10.1111/jnc.13565.
neurotransmission. J Neurochem, 2013. 126(2): pp. 146–54.
calcium sensors in short-term plasticity. Front Cell Neurosci, 2014. 8: p. 356.
75.
hearing. Trends Neurosci, 2012. 35(11): pp. 671–80.
cells. J Neurosci, 2010. 30(40): pp. 13281–90.
cell wound healing. Biochim Biophys Acta, 2015. 1853(10 Pt A): pp. 2649–61.
Cytol, 1955. 1(5): pp. 445–58.
caveolae to the surface membrane of frog skeletal muscle bres at dierent sarcomere lengths. J
Physiol, 1975. 250(3): pp. 513–39.
mechanosensitive I(Cl,swell) channel activation during swelling in the rat ventricular myo-
cyte. PLoS One, 2009. 4(12): p. e8312.
by caveolae in the adult cardiac myocyte. J Mol Cell Cardiol, 2008. 45(1): pp. 88–92.
144(3): pp. 402–13.
pp. 1269–78.
Wound Healing: New insights into Ancient Challenges216

function of caveolae. Curr Opin Cell Biol, 2014. 29: pp. 99–106.
Genet, 2000. 9(20): pp. 3047–54.
cellular defense against bacterial pore-forming toxins. Biochim Biophys Acta, 2015. 1853 (9):
p. 2045-54.
lumen of MVBs for degradation. Trac, 2012. 13(3): pp. 483–94.
pp. 331–71.
electron microscopy-X-ray crystallography study. J Mol Biol, 1994. 238(2): pp. 199–213.
V and lipids in planar supported lipid bilayers. Biochemistry, 1999. 38(9): pp. 2779–86.
repair. Nat Commun, 2011. 2: p. 270.
dependent muscle membrane repair. J Neurosci, 2013. 33(12): pp. 5085–94.
5): pp. 719–31.
transection: evidence for enhancement by Ca(2+)-triggered protease activity and cytoskeletal
disassembly. J Neurosci, 1991. 11(10): pp. 3257–67.
Interface, 2014. 11(95): p. 20140071.
Biorheology, 2009. 46(2): pp. 93–105.
cytoskeletal actin ber structure in osteoblastic cells. J Biomech, 2005. 38(9): pp. 1895–901.
cytoskeletal tapestry. Cell Motil Cytoskeleton, 2002. 52(4): pp. 266–74.
ently of myosin light chain dephosphorylation. Interface Focus, 2011. 1(5): pp. 754–66.
How Plasma Membrane and Cytoskeletal Dynamics Influence Single-Cell Wound Healing: Mechanotransduction,
http://dx.doi.org/10.5772/63765
Tension and Tensegrity
217

Biophys J, 2015. 108(1): pp. 43–52.
mentation of strained actin laments. Biophys J, 2015. 108(9): pp. 2270–81.
adhesion proteins. Commun Integr Biol, 2012. 5(6): pp. 572–7.
plex. Pugers Arch, 2015. 467(1): pp. 141–55.
tension-dependent binding of colin to the lament. J Cell Biol, 2011. 195(5): pp. 721–7.
Cell Biol, 2010. 20(4): pp. 187–95.
polymerization. Proc Natl Acad Sci U S A, 2004. 101(51): pp. 17664–8.
2004. 117(Pt 12): pp. 2449–60.
nonexcitable cells. J Cell Biol, 1995. 128(4): pp. 589–98.
divalent metal chlorides. Acta Biochim Biophys Acad Sci Hung, 1979. 14(1–2): pp. 31–42.
Ca2+. Biochim Biophys Acta, 1983. 742(1): pp. 135–41.
2012. 1824(1): pp. 224–36.
ics. Curr Opin Cell Biol, 2004. 16(1): pp. 94–8.
Cdc42/N-WASP-induced actin laments. Mol Biol Cell, 2004. 15(2): pp. 520–31.
Biol, 1997. 9(1): pp. 18–28.
343(6174): p. 1247136.
resealed by Ca2+-dependent vesicle-vesicle fusion events. J Cell Biol, 1997. 139(1): pp. 63–74.
Wound Healing: New insights into Ancient Challenges218

Cell Sci, 2000. 113 (Pt 11): pp. 1891–902.
occurs through three distinct phases of membrane and cytoskeletal remodeling. J Cell Biol, 2011.
193(3): pp. 455–64.
1963. 18: pp. 135–51.
2007. 8(7): pp. 785–94.
2007. 19(4): pp. 409–16.
plasma membrane repair. Trends Cell Biol, 2014. 24(12): pp. 734–742.
close actomyosin rings around Xenopus oocyte wounds. J Cell Biol, 2001. 154(4): pp. 785–97.
within the organelle. Cell, 2002. 108(3): pp. 357–69.
Rev Mol Cell Biol, 2005. 6(6): pp. 499–505.
Biochem Cell Biol, 2005. 83(6): pp. 711–20.
cell wounds. J Cell Biol, 2005. 168(3): pp. 429–39.
RhoA during cleavage plane specication. J Cell Biol, 2005. 170(1): pp. 91–101.
by Rho-kinase in vivo. J Cell Biol, 1999. 147(5): pp. 1023–38.
2: pp. 241–55.
promotes actin nucleation by Arp2/3 complex and bacterial actin-based motility. J Cell Biol,
1999. 146(6): pp. 1319–32.
dependent signals to actin assembly. Cell, 1999. 97(2): pp. 221–231.
How Plasma Membrane and Cytoskeletal Dynamics Influence Single-Cell Wound Healing: Mechanotransduction,
http://dx.doi.org/10.5772/63765
Tension and Tensegrity
219

[92] Taunton, J., et al., Actin-dependent propulsion of endosomes and lysosomes by recruitment of
N-WASP. J Cell Biol, 2000. 148(3): pp. 519–30.
[93] Burkel, B.M., et al., A Rho GTPase signal treadmill backs a contractile array. Dev Cell, 2012.
23(2): pp. 384–96.
[94] Vaughan, E.M., et al., Lipid domain-dependent regulation of single-cell wound repair. Mol
Biol Cell, 2014. 25(12): pp. 1867–76.
[95] Yu, H.Y. and W.M. Bement, Control of local actin assembly by membrane fusion-dependent
compartment mixing. Nat Cell Biol, 2007. 9(2): pp. 149–59.
[96] Arun, S.N., et al., Cell wounding activates phospholipase D in primary mouse keratinocytes.
J Lipid Res, 2013. 54(3): pp. 581–91.
[97] Abreu-Blanco, M.T., J.M. Verboon, and S.M. Parkhurst, Coordination of Rho family GTPase
activities to orchestrate cytoskeleton responses during cell wound repair. Curr Biol, 2014.
24(2): pp. 144–55.
[98] Steinhardt, R.A., G. Bi, and J.M. Alderton, Cell membrane resealing by a vesicular mecha-
nism similar to neurotransmier release. Science, 1994. 263(5145): pp. 390–3.
[99] Jaiswal, J.K., N.W. Andrews, and S.M. Simon, Membrane proximal lysosomes are the major
vesicles responsible for calcium-dependent exocytosis in nonsecretory cells. J Cell Biol, 2002.
159(4): pp. 625–35.
[100] Reddy, A., E.V. Caler, and N.W. Andrews, Plasma membrane repair is mediated by Ca(2+)-
regulated exocytosis of lysosomes. Cell, 2001. 106(2): pp. 157–69.
[101] Tam, C., et al., Exocytosis of acid sphingomyelinase by wounded cells promotes endocytosis
and plasma membrane repair. J Cell Biol, 2010. 189(6): pp. 1027–38.
[102] Lei, J. and E.T. Kavalali, Ca2+ Dependence of synaptic vesicle endocytosis. Neuroscientist,
2015. DOI: 10.1177/1073858415588265.
[103] Bi, G.Q., J.M. Alderton, and R.A. Steinhardt, Calcium-regulated exocytosis is required for
cell membrane resealing. J Cell Biol, 1995. 131(6 Pt 2): pp. 1747–58.
[104] Rust, M.B., ADF/colin: a crucial regulator of synapse physiology and behavior. Cell Mol Life
Sci, 2015. 72(18): pp. 3521–9.
[105] Bi, G.Q., et al., Kinesin- and myosin-driven steps of vesicle recruitment for Ca2+-regulated
exocytosis. J Cell Biol, 1997. 138(5): pp. 999–1008.
[106] McNeil, P.L., Repairing a torn cell surface: make way, lysosomes to the rescue. J Cell Sci, 2002.
115(Pt 5): pp. 873–9.
[107] Rizo, J. and J. Xu, The synaptic vesicle release machinery. Annu Rev Biophys, 2015. 44: pp.
339–67.
Wound Healing: New insights into Ancient Challenges220

[108] Detrait, E., et al., Axolemmal repair requires proteins that mediate synaptic vesicle fusion. J
Neurobiol, 2000. 44(4): pp. 382–91.
[109] Detrait, E.R., et al., Plasmalemmal repair of severed neurites of PC12 cells requires Ca(2+) and
synaptotagmin. J Neurosci Res, 2000. 62(4): pp. 566–73.
[110] Fox, M.A. and J.R. Sanes, Synaptotagmin I and II are present in distinct subsets of central
synapses. J Comp Neurol, 2007. 503(2): pp. 280–96.
[111] Shen, S.S., et al., Molecular regulation of membrane resealing in 3T3 broblasts. J Biol Chem,
2005. 280(2): pp. 1652–60.
[112] Chakrabarti, S., et al., Impaired membrane resealing and autoimmune myositis in synapto-
tagmin VII-decient mice. J Cell Biol, 2003. 162(4): pp. 543–9.
[113] Toops, K.A. and A. Lakkaraju, Let’s play a game of chutes and ladders: lysosome fusion with
the epithelial plasma membrane. Commun Integr Biol, 2013. 6(4): p. e24474.
[114] Shen, S.S. and R.A. Steinhardt, The mechanisms of cell membrane resealing in rabbit corneal
epithelial cells. Curr Eye Res, 2005. 30(7): pp. 543–54.
[115] Rao, S.K., et al., Identication of SNAREs involved in synaptotagmin VII-regulated lysosomal
exocytosis. J Biol Chem, 2004. 279(19): pp. 20471–9.
[116] Tardieux, I., et al., Lysosome recruitment and fusion are early events required for trypanosome
invasion of mammalian cells. Cell, 1992. 71(7): pp. 1117–30.
[117] Miyake, K. and P.L. McNeil, Vesicle accumulation and exocytosis at sites of plasma membrane
disruption. J Cell Biol, 1995. 131(6 Pt 2): pp. 1737–45.
[118] Togo, T., T.B. Krasieva, and R.A. Steinhardt, A decrease in membrane tension precedes
successful cell-membrane repair. Mol Biol Cell, 2000. 11(12): pp. 4339–46.
[119] Cooper, S.T. and P.L. McNeil, Membrane repair: mechanisms and pathophysiology. Physiol
Rev, 2015. 95(4): pp. 1205–40.
[120] Togo, T. and R.A. Steinhardt, Nonmuscle myosin IIA and IIB have distinct functions
in the exocytosis-dependent process of cell membrane repair. Mol Biol Cell, 2004.
15(2): pp. 688–95.
[121] Andrews, N.W., M. Corroe, and T. Castro-Gomes, Above the fray: surface remodeling by
secreted lysosomal enzymes leads to endocytosis-mediated plasma membrane repair. Semin Cell
Dev Biol, 2015. 45: pp. 10–7.
[122] Stauber, W.T., Eccentric action of muscles: physiology, injury, and adaptation. Exerc Sport
Sci Rev, 1989. 17: pp. 157–85.
[123] Proske, U. and D.L. Morgan, Muscle damage from eccentric exercise: mechanism, mechanical
signs, adaptation and clinical applications. J Physiol, 2001. 537(Pt 2): pp. 333–45.
How Plasma Membrane and Cytoskeletal Dynamics Influence Single-Cell Wound Healing: Mechanotransduction,
Tension and Tensegrity
http://dx.doi.org/10.5772/63765
221

[124] Takekura, H., et al., Eccentric exercise-induced morphological changes in the membrane
systems involved in excitation-contraction coupling in rat skeletal muscle. J Physiol, 2001.
533(Pt 2): pp. 571–83.
[125] Bashir, R., et al., A gene related to Caenorhabditis elegans spermatogenesis factor fer-1 is
mutated in limb-girdle muscular dystrophy type 2B. Nat Genet, 1998. 20(1): pp. 37–42.
[126] Liu, J., et al., Dysferlin, a novel skeletal muscle gene, is mutated in Miyoshi myopathy and limb
girdle muscular dystrophy. Nat Genet, 1998. 20(1): pp. 31–6.
[127] Anderson, L.V., et al., Dysferlin is a plasma membrane protein and is expressed early in human
development. Hum Mol Genet, 1999. 8(5): pp. 855–61.
[128] Bansal, D., et al., Defective membrane repair in dysferlin-decient muscular dystrophy.
Nature, 2003. 423(6936): pp. 168–72.
[129] Pangrsic, T., et al., Hearing requires otoferlin-dependent ecient replenishment of synaptic
vesicles in hair cells. Nat Neurosci, 2010. 13(7): pp. 869–76.
[130] Johnson, C.P. and E.R. Chapman, Otoferlin is a calcium sensor that directly regulates
SNARE-mediated membrane fusion. J Cell Biol, 2010. 191(1): pp. 187–97.
[131] Reisinger, E., et al., Probing the functional equivalence of otoferlin and synaptotagmin 1 in
exocytosis. J Neurosci, 2011. 31(13): pp. 4886–95.
[132] Piccolo, F., et al., Intracellular accumulation and reduced sarcolemmal expression of dysferlin
in limb – girdle muscular dystrophies. Ann Neurol, 2000. 48(6): pp. 902–12.
[133] McDade, J.R., A. Archambeau, and D.E. Michele, Rapid actin-cytoskeleton-dependent
recruitment of plasma membrane-derived dysferlin at wounds is critical for muscle membrane
repair. FASEB J, 2014. 28(8): pp. 3660–70.
[134] Cai, C., et al., Membrane repair defects in muscular dystrophy are linked to altered interaction
between MG53, caveolin-3, and dysferlin. J Biol Chem, 2009. 284(23): pp. 15894–902.
[135] Matsuda, C., et al., The C2A domain in dysferlin is important for association with MG53
(TRIM72). PLoS Curr, 2012. 4: p. e5035add8ca4.
[136] Waddell, L.B., et al., Dysferlin, annexin A1, and mitsugumin 53 are upregulated in muscular
dystrophy and localize to longitudinal tubules of the T-system with stretch. J Neuropathol Exp
Neurol, 2011. 70(4): pp. 302–13.
[137] Caccioolo, M., et al., Reverse engineering gene network identies new dysferlin-interacting
proteins. J Biol Chem, 2011. 286(7): pp. 5404–13.
[138] Redpath, G.M., et al., Calpain cleavage within dysferlin exon 40a releases a synaptotagmin-
like module for membrane repair. Mol Biol Cell, 2014. 25(19): pp. 3037–48.
[139] Fuson, K., et al., Alternate splicing of dysferlin C2A confers Ca(2)(+)-dependent and Ca(2)(+)-
independent binding for membrane repair. Structure, 2014. 22(1): pp. 104–15.
Wound Healing: New insights into Ancient Challenges222
Соседние файлы в папке Библиотека им академика М.И. Перельмана
