The internal experiences of expansion, spaciousness, fluidity, softening, lengthening, allowing, groundedness, and integration are mediated via the consciousness, awareness, responsiveness and communication of every cell in the body. In working with our clients, we know we connect with the whole person on multiple levels. However, the question of how our physical contact with another person affects the cells of myoneurofascial tissues during and after a Structural Integration session has been of great interest to many of us. The melting we feel as the tissues soften and release has been characterized as a thixotropic reaction to the applied pressure and shear. What about the cells in the tissue? This paper presents and discusses evidence suggesting that the kinds of manipulations we employ result in collagen fibers being sheared apart where they interface with the softer watery matrix or ground substance that surrounds them. The cells in these areas are undoubtedly stretched and their fine microscopic processes are broken. This process of cells experiencing shear, compressive and tensional forces, rearranging themselves and restructuring the extracellular matrix is part and parcel of living in gravity, and of experiencing bodywork.
I am blessed to have the pleasure of using state-of-the-art microscopes to peer into the world of tissues, cells and extracellular matrix for hours at a time, and this helps me see how alive, responsive, and fragile our cells really are. Time-lapse imaging is my favorite because it allows us to watch the dynamics of cell movement, as well as how cells interact and respond to manipulations or other perturbations. My early research experience in a connective tissue laboratory at the University of New Mexico felt like a perfect segue for entering the field of Structural Integration. However, even after several years of studying connective tissues in the lab, I had never fully realized how manual manipulation of this tissue (as well as the other levels of interaction that occur within the therapeutic relationship) could lead to such profound changes on so many levels – that is, until I experienced Structural Integration. In this article I would like to share some older and more recent findings regarding the amazing structure and functional plasticity of connective tissue and the resident fibroblasts. I also hope to share some of the thoughts I have, based on my own research and from others in the connective tissue research arena, on what might be happening to the tissue and cells under our fingertips. I am particularly excited about some recent microscopic observations of extremely fine actin-containing cellular processes that form the hyaluronan-rich pericellular matrix. I will also present microscopic images of the immediate cellular response following manipulation of cultured cells. These data point to a potential mechanism of mechanical signaling and cellular responses during and following fascial manipulation (or injury) based primarily on the changes in cell shape that result from externally applied stretching and shearing.
<center>Fibrocartilage in tendon</center>
Another question we have all asked is how do connective tissues become so hard, bunched-up, and tight? Research from the late 1980s and early ?90s investigating the role of mechanics on cellular function may shed some light on this question. This is a period when bioengineers and others were designing all sorts of equipment to apply forces to cells and tissues. Mine was a tendon masher. The equipment was designed to test the idea that cyclic mechanical compression of cultured tendon segments can lead to changes in composition and structure in the tissue. This was used as a model to study how tendons undergo a transition into fibrocartilage in locations where they wrap around a bony pulley and receive compressive forces. Examples of tendon fibrocartilage include the human tibialis posterior tendon, where it wraps around the medial malleolus, and bovine(1,2) and rabbit(3) flexor digitorum longus tendons, which wrap around sesamoid bones in the metatarsal phalangeal joint. I have also seen a similar situation in rat extensor digitorum longus tendons where they wrap around and contact the backside of the extensor retinaculum (Evanko, unpublished observation). Additionally, fibrocartilage can be found at tendon insertions or entheses.(4) Some regard this fibrocartilage transition as pathological and suggest it may predispose a tendon to rupture. We often have our fingers in the transition areas and tendon insertions where the fibrocartilage is forming, so it seems a topic worthy of more consideration by Structural Integrators.
Figure 1. Histology of bovine tendon. A, B, Light microscopy of tensional (A) and compressed (B) regions of fetal tendon. The arrow in A points to a seam of loose tissue and cells between the dense fiber bundles that would yield to manipulation. See figure 2. Note the basket weave arrangement of the collagen fibers and increased ground substance in the compressed region. C, D, Electron microscopy of fetal tendon shows linear collagen fibrils in the tensional region and more proteoglycan rich matrix in the compressed region. Proteoglycans are indicated with arrows. E, F, Electron microscopy of adult tensional and compressed tendon regions. Note how the collagen fibrils increase in size, as do the proteoglycans (arrows) of the compressed region. Adapted from Evanko and Vogel (6).
Figure 1 shows light and electron microscopic histology of the tensional and compressed regions of bovine flexor tendon. In the region of tendon that receives purely tensile forces, collagen fibers and elongated cells are aligned in the direction of force and small collagen-binding proteoglycans are dominant. In the location of tendon that receives compression, the cells are round and chondrocyte-like and the matrix is a basket-weave of collagen fibers and watery spaces filled with large, compression resistant proteoglycans, hyaluronan and other glycoproteins. Other studies have found that tendon cells are connected via gap junctions and these are lost in locations that become fibrocartilaginous.(5) In the wrap around portion of the tendon, these changes happen particularly on the side in direct contact with the bone, with a transition to normal tendon architecture and composition as you move away from the compressed surface. The transition in fiber orientation in the compressed region in bovine flexor tendon begins in utero, coinciding with the onset of limb movement in the fetus, suggesting that non-weight bearing movement initiates some shear in the tissue.(6) The tissue only becomes hard and fibrocartilaginous after birth when the animal begins significant weight-bearing and the tendon is compressed during standing and walking. This textural change results from the accumulation of chondroitin sulfate proteoglycans (aggrecan) and hyaluronan.(7) Proteoglycans and hyaluronan can influence collagen fibril formation and the spacing between fibrils, as well as provide compressive stiffness to the tissue. Fetal tendons do not have as much of the large aggrecan proteoglycans and are much softer in texture. In addition, the diameter of collagen fibrils increases in the adult tendon, particularly in the region that transmits purely tensional forces. Studies have shown that surgical translocation of the wrap around part of the tendon to a place where it no longer experiences compression causes the tendon to change back into the normal tension bearing appearance and composition.(8) The synthesis of the large aggrecans is diminished in tissue explants cultured in the absence of the normal compressive environment in vivo.(2)
We asked whether cyclic compression, applied to full-thickness segments of developing bovine tendon in vitro, can lead to biosynthetic changes in the resident cells that would indicate they may be transitioning into chondrocyte-like cells. These experiments used a frequency of 0.16 Hz, or 1 cycle every 6 seconds. As little as a few hours of cyclic compression at this frequency led to measurable changes in proteoglycan synthesis in the tissue pieces.(9) The response of adult tendon fibrocartilage was similar, but less pronounced.(10) The proteoglycans synthesized by the cells in response to compression, i.e., aggrecan and biglycan, were consistent with differentiation toward a fibrocartilaginous phenotype. Uncompressed control tissue continued to make primarily the small collagen-binding proteoglycan, decorin, which is characteristic of tensional tendon. Figure 2 shows how these compression-induced changes were accompanied by an increase in the amount of staining for transforming growth factor-beta (TGF-beta) in the tendon cells (Fig. 2).(11) TGF-beta is a growth factor known to be important in cartilage formation.(12) In compressed tendon, TGF-beta appears to be acting in an autocrine or paracrine fashion to create a chondrogenic milieu that arises from the mechanical environment. In these experiments, the tissue changes were independent of the nervous system.
Figure 2. Cultured tendon segments were cyclically compressed for 3 days and then stained for TGF-beta1 using immunohistochemistry. A, control tissue. B, compressed tissue. Note the rounder shape and the increase in TGF-beta staining in the cells following compression.
Recent data suggest that there are stem cells in tendons that may be responsive to the loading.(13) TGF-beta is also involved in differentiation of fibroblasts into myofibroblasts.(14,15) In vivo, the transition of fibroblasts into myofibroblasts or fibrochondrocytes will also be dependent on other growth factors, cytokines, and levels of inflammation, which we know can involve nerves as well as inflammatory cells. Although it may be stating the obvious, the mechanical environment seems to be of prime importance in directing tissue composition and fiber arrangement, and therefore, the textures we feel. The influence of the nervous system would be superimposed on the inherent connective tissue responses. One of my clients wears a prosthetic leg, which cyclically compresses the tissues around his ischial tuberosity. The tissues on that side felt like they could be undergoing a transition to fibrocartilage, similar to that seen in the tendons. Regular work there keeps those tissues softer and the nerves free.
<center>Manipulating tissues and cells</center>
What might the cells in the connective tissues experience during compression in vitro, or in a clinical setting, subjected to either light or forceful myoneurofascial manipulations? Helene Langevin and her colleagues have shown how stretching of connective tissue, by twisting acupuncture needles or performing yoga-like stretches, induces changes in cell shape, nuclear shape, and the distribution of the cytoskeletal elements within the cells.(16-18) When cells in the superficial fascia are stretched, they become larger and more highly spread; their nuclei become larger, more spherical, and lose their invaginations and grooves. All of these changes can lead to altered gene regulation and protein synthesis. Stretch and tension also influences TGF-beta production in the tissues and can drive fibrosis.(14,16)
If you take fresh, dense regular connective tissue, such as the bovine gastrocnemius fascia shown in Figure 3A, and manipulate it with your fingertip, i.e., use it as a blunt probe, you will essentially pry apart the collagen fibers as shown in Figure 3B. It is clear that the tissue yields most readily in the places where there are ?seams? of loose ground substance-rich matrix between the dense collagen fibers. The arrows in Figure 1A and Figure 3 point to similar seams within a tendon. These areas have numerous rounded cells that are surrounded by a loose, watery matrix containing proteoglycans and other glycoproteins. Blood vessels and nerve twigs tend to course along these seams. In Figure 3B you can see a small arteriole that was perturbed by the manipulation, which could have resulted in bruising in a living subject. We have all seen such bruising. When we manipulate living tissue, the cells in these seams are certainly going to be stretched and sheared to varying degrees. Mathematical modeling has predicted that palpable tissue release could result from deformation in softer tissues, such as superficial nasal fascia.(19) Yet, according to that study, denser fascial tissues required huge, non-physiologic forces even for small shear or compressive deformations. However, I?m not sure this is always the case.
Figure 3. Manipulation of fresh fascial tissue. Bovine gastrocnemius fascia before (A) and after (B) manipulation with my fingertip. The glistening is due to the watery proteoglycan- and hyaluronan-rich ground substance. The arrows point to a seam between fibers like the seam shown in Figure 1A. The arrowhead indicates a small arteriole that is also between the fibers. The tissue in B was stained with toluidine blue after manipulation to provide better contrast.
Studies have shown that the elongated cells within the dense fiber bundles of tendons are arranged in discrete linear chains, with relatively strong end-to-end connections between cells.(20) This would be in the tensional region of tendon. The cell chains can be isolated in long arrays by mechanical shearing and enzymatic treatment. The cells in the chain are surrounded by matrix containing proteoglycan (versican) and other fibrillar structures. In addition to the end-to-end connections, electron microscopy shows that fibroblasts also have fine lateral processes that encircle the collagen fiber bundles and contact distant cells. Figure 4 shows cross-sectional electron microscopic images of the bundles of collagen fibrils from the densely fibrous tensional tendon (one bundle of fibrils constitutes a fiber). We can see how small cellular processes encircle the fibril bundles like arms, and how the watery proteoglycan-rich areas form the gel-like glue between the fibers. Potentially, these fine processes can easily be broken during fascial manipulations, possibly without destroying the whole cell. The cells would initially retract the broken processes and assume a new, more rounded shape. The cells could potentially re-extend new processes, form new pericellular matrix and new gap junctions or other kind of connections with each other.
Figure 4. Electron microscopy of young tendon. A, cross section showing a bundle of fibrils (indicated by the asterisk) surrounded by cytoplasmic cell processes (arrow). The nucleus of the cell is also indicated (arrow). B, shows another example of the compressed part of tendon with collagen fibril bundles running various directions and more hyaluronan-dependent matrix around the cell (arrows). C, in a young calf tendon, proteoglycans (arrows) are in the seam between two fibers. D, shows an example of extremely fine cell processes (arrows) between collagen bundles.
Cultured cells and confluent cell sheets have relatively few collagen fibers and more proteoglycans in their extracellular matrix, and therefore provide a useful and reasonable model of superficial fascia or the loose connective tissue seam areas between dense collagen fibers. We have used cultured cells to study the effects of manipulations at the cellular level. Thus far, my experiments have focused on single acute stretches in an effort to model what might happen during a single manipulation. Figure 5 shows cultured human lung fibroblasts during manipulation of the monolayer with a micropipette. The instrument had an extremely fine point, approximately 40 micrometers, which is finer than a hair and smaller than a single fibroblast. Even with only about 40 milligrams of force (~1/1000 oz.), you can see that the cell sheet is torn quite easily. The cells are stretched and sheared to varying degrees; some tear completely in half and are destroyed, others have only a few fine processes that are broken. Again, most of the cells immediately affected by the manipulation assume a new shape, which is a known regulator of cell function.(21)
Figure 5. Micromanipulation of cell sheets. Fibroblasts cultured for 1 week were manipulated with a micropipette. Note how cells are stretched (arrow) and sometimes broken. The holes or perforations in the cell layer tend to widen after the manipulation as cells recoil suggesting there is a tensional prestress in the sheet generated by the cells.
If fibroblasts are grown in culture for an extended period, considerable tension will develop in the cell layer due to collective contractile forces exerted by individual fibroblasts. As shown in Figure 5, if holes are poked in the cell sheet with a pipette tip (or perhaps with fingernails), the holes will usually open further due to the prestress in the cell sheet. If the cell layer is released from one end of the culture dish, the sheet will rapidly contract and roll up into a small ball. I think that a similar phenomenon is part of the palpable tissue release in practice. Many of us intuitively know how gently poking into hard, strained tissue with fingernails can be a good way to begin the opening process. Fingernails would be expected to produce high levels of compressive and shear force at the local cell level. These observations of cultured cell sheets provide some rationale for judicious and strategic use of fingernails during myofascial work. In my practice, I find that fingernails are extremely effective at releasing restrictions around nerves in the superficial fascia. Because of the long lever arm between the tip of the nail and the nail bed, I find the fingernails are also extremely sensitive just for palpating the superficial-most layers, i.e., that first layer of availability.
Figure 6. Response of a single cell to stretching with a micropipette. A fibroblast with thin cell processes was stretched at the two locations indicated with the asterisk. A particle exclusion assay was used to reveal hyaluronan pericellular matrix, and is seen as a clear zone that surrounds the cell body (arrowheads). The matrix formed (or was revealed) as the cell recoiled from the trauma. The arrows point to dilations that have formed along the stretched processes. A third process projects downward and was broken immediately during the manipulation.
When individual cells were stretched, we found that a hyaluronan-dependent pericellular coat formed around the cell body or stretched process within as little as 20 minutes following manipulations. Figure 6 shows a cell with three long processes that were stretched with the pipette. The arrows point to distinct swellings that formed along the stretched processes, shortly before they broke. The arrowheads indicate the hyaluronan dependent cell coat that formed as the cell rounded up. We have recently found that these swellings (indicated with arrows) appear to be rich in actin, which presumably has broken and recoiled into a ball inside the cell process (Evanko, unpublished observation). Hyaluronan matrix was also seen around cell fragments. The degree of retraction of the torn edges of the cell layer depended on the level of pre-stress in the cell layer, substrate adhesiveness, the degree of intercellular connection and orientation relative to the direction of force. By 6 hours, hyaluronan-dependent pericellular matrices were seen around the cells migrating from the retracted edges of the cell sheet. As shown in Figure 7, fluorescent staining 24 h later revealed long hyaluronan and versican-rich cables that connected cells, and occasionally completely bridged the wound gap (>1 mm long). These data suggest that alterations in hyaluronan amount and organization may underlie tissue changes following manipulation or stretching. In addition, hyaluronan may be part of an early mode of cellular re-coupling following manipulation. One time-lapse sequence demonstrates how cells can use this matrix as a guide during re-extension of their processes, as they try to reconnect with each other (not shown here).
Figure 7. Hyaluronan cables form across wound gap. Cells were stretched and cleared from a portion of the culture surface with a pipette tip. After 24 hours, the cells were fixed and stained for hyaluronan. Long cables of hyaluronan (arrows) have formed between the retracted edges of the cell layer and appear to bridge the wound.
Hyaluronan is often increased under inflammation and is partly responsible for holding the water in edematous tissue. Hyaluronan and proteoglycans in high concentrations can make for a very viscous and sticky pro-inflammatory matrix.(22,23) Hyaluronan is also increased in achilles tendons following immobilization.(24) How inflammation affects the process of extracellular matrix formation and remodeling is the subject of ongoing research. When cells were treated with inflammatory mediators just after manipulation, increased hyaluronan cables were produced, but cell migration was impaired. It is possible that the composition, and therefore adhesiveness, of the matrix can influence the ability of cells to migrate and recover from stretching or wounding.
<center>Actin containing cell protrusions</center>
Recent studies have discovered the presence of extremely fine, actin-containing cellular protrusions that are instrumental in the formation of the hyaluronan pericellular matrix.(25,26) Figure 8 shows fluorescence microscopy and a scanning electron micrograph of single fibroblasts with several of the fine protrusions. When the cells are stained for hyaluronan it is clear that the protrusions are surrounded by a coat of hyaluronan. We have also found that the proteoglycan versican is associated with the protrusions. These processes are extremely slender – only about 40 nm wide, and contain only one or two actin filaments; and they can be extremely long – up to several cell lengths. The protrusions depend on hyaluronan for strength. The cell will retract the protrusions following digestion of the matrix with an enzyme that degrades hyaluronic acid,(26) and as we have seen, after manipulation in culture. Time-lapse data suggest that the fine protrusions are dynamically extended and retracted and may be used by the cells to form pericellular matrix cables and to traction and remodel the ground substance and matrix fibers. I have been curious as to whether this activity may be part of the basis for palpable tissue motility.
Figure 8. Extremely fine cellular protrusions of fibroblasts. Numerous protrusions coated with hyaluronan extend from the edge of the cells. A, hyaluronan staining around the protrusions. B, scanning electron micrograph showing several long protrusions.
I am currently studying whether formation of the fine protrusions is induced in fibroblasts under various inflammatory conditions. There is evidence that these fine processes exist in real connective tissues, for example, rat tendon(5) and calf tendon (see Figure 4). The small canaliculi connecting the osteocytes within bone contain similar fine processes, and are another real tissue example. In the laboratory, it has been demonstrated that cell protrusions are extremely fragile; they can be easily sheared off just by agitating the fluid or culture medium over the cells. I suspect that in vivo these tiny processes can be broken, even when the lightest of touch is used, or during indirect fascial manipulations. Breaking these tiny processes would lead to cell shape changes and redistribution of actin, similar to those changes observed following acupuncture needle twisting. This could produce a host of changes in cellular biosynthesis and gene expression in the tissue.
In particular I am interested in how the hardening and tightening of tissues, whether they become fibrotic or more fibrocartilaginous (or some combination thereof), can lead to trapping of nerves and neurogenic inflammation. Factors released by irritated nerves or the nerve sheath(27) can further exacerbate myofibroblast formation, matrix stiffening, and fascial restrictions, as well as hyaluronan- and proteoglycan- mediated swelling and edema. The outgrowth of neurites, or small neuronal processes, underlies the phenomenon of terminal arborization of nerves that may occur during neurogenic inflammation. Neurite outgrowth is influenced by hyaluronan and can be blocked and/or guided by deposits of the chondroitin sulfate proteoglycans, such as aggrecan or versican.(28,29)
Don Hazen and Christoph Sommer have been instrumental in fostering the recent interest in nerve manipulation in the SI community. For interesting reading on the subject, I recommend http://www.dhazen.com. In my own practice, the incorporation of neurofascial work has been an incredibly valuable addition to my understanding and my toolset, and provides a greater range of possibility for pain relief and accomplishing the goals of SI. Although there is little or no data, the general physical cellular effects of manipulation on nerves may be similar to the effects on fibroblasts. As Hazen has pointed out (and I find to be true), light finger pad work is extremely effective at freeing the tethered nerves and pumping the inflammatory exudate back into lymphatics. In addition to breaking up the viscous ground substance, it is tempting to speculate that neurofascial manipulations may break the microscopic neurites of the nervi nervorum as they grow out and reach into the tissue surrounding the nerve sheath. This may mediate tissue release and some of the beneficial effects on the nervous system and surrounding tissues.
In conclusion, when we consider what happens at the cellular level, most of the techniques we employ would tend to place a high degree of shear and stretch on tiny cell processes and probably whole cells. The change in cell shape resulting from such shear and stretch is an important mechanism that likely contributes to the improved pliability we associate with healthier connective tissue and happier nerves.
1. Koob, T.J. and K.G. Vogel, Site-related variations in glycosaminoglycan content and swelling properties of bovine flexor tendon. J Orthop Res, 1987. 5:414-24.
2. Vogel, K.G., A. Ordog, G. Pogany, and J. Olah, Proteoglycans in the compressed region of human tibialis posterior tendon and in ligaments. J Orthop Res., 1993. 11:68-77.
3. Gillard, G.C., M.J. Merrilees, P.G. Bell-Booth, H.C. Reilly, and M.H. Flint, The proteoglycan content and the axial periodicity of collagen in tendon. Biochem J, 1977. 163:145-51.
4. Benjamin, M. and J.R. Ralphs, Fibrocartilage in tendons and ligaments–an adaptation to compressive load. J Anat, 1998. 193 ( Pt 4):481-94.
5. Ralphs, J.R., M. Benjamin, A.D. Waggett, D.C. Russell, K. Messner, and J. Gao, Regional differences in cell shape and gap junction expression in rat Achilles tendon: relation to fibrocartilage differentiation. J Anat, 1998. 193 (Pt 2):215-22.
6. Evanko, S.P. and K.G. Vogel, Ultrastructure and proteoglycan composition in the developing fibrocartilaginous region of bovine tendon.
Matrix, 1990. 10:420-36.
7. Koob, T.J., Effects of chondroitinase-ABC on proteoglycans and swelling properties of fibrocartilage in bovine flexor tendon. J Orthop Res, 1989. 7:219-27.
8. Gillard, G.C., H.C. Reilly, P.G. Bell-Booth, and M.H. Flint, The influence of mechanical forces on the glycosaminoglycan content of the rabbit flexor digitorum profundus tendon. Connect Tissue Res, 1979. 7:37-46.
9. Evanko, S.P. and K.G. Vogel, Proteoglycan synthesis in fetal tendon is differentially regulated by cyclic compression in vitro. Arch Biochem Biophys, 1993. 307:153-64.
10. Koob, T.J., P.E. Clark, D.J. Hernandez, F.A. Thurmond, and K.G. Vogel, Compression loading in vitro regulates proteoglycan synthesis by tendon fibrocartilage. Arch Biochem Biophys, 1992. 298:303-12.
11. Robbins, J.R., S.P. Evanko, and K.G. Vogel, Mechanical loading and TGF-beta regulate proteoglycan synthesis in tendon. Arch Biochem Biophys, 1997. 342:203-11.
12. Sanders, E.J., S. Prasad, and N. Hu, The involvement of TGF-beta 1 in early avian development: gastrulation and chondrogenesis. Anat Embryol (Berl), 1993. 187:573-81.
13. Bi, Y., D. Ehirchiou, T.M. Kilts, C.A. Inkson, M.C. Embree, W. Sonoyama, L. Li, A.I. Leet, B.M. Seo, L. Zhang, S. Shi, and M.F. Young, Identification of tendon stem/progenitor cells and the role of the extracellular matrix in their niche. Nat Med, 2007. 13:1219-27.
14. Gabbiani, G., The biology of the myofibroblast. Kidney Int, 1992. 41:530-2.
15. Tomasek, J.J., G. Gabbiani, B. Hinz, C. Chaponnier, and R.A. Brown, Myofibroblasts and mechano-regulation of connective tissue remodelling. Nat Rev Mol Cell Biol, 2002. 3:349-63.
16. Bouffard, N.A., K.R. Cutroneo, G.J. Badger, S.L. White, T.R. Buttolph, H.P. Ehrlich, D. Stevens-Tuttle, and H.M. Langevin, Tissue stretch decreases soluble TGF-beta1 and type-1 procollagen in mouse subcutaneous connective tissue: Evidence from ex vivo and in vivo models. J Cell Physiol, 2007. 214:389-395.
17. Langevin, H.M., N.A. Bouffard, D.L. Churchill, and G.J. Badger, Connective tissue fibroblast response to acupuncture: dose-dependent effect of bidirectional needle rotation. J Altern Complement Med, 2007. 13:355-60.
18. Storch, K.N., D.J. Taatjes, N.A. Bouffard, S. Locknar, N.M. Bishop, and H.M. Langevin, Alpha smooth muscle actin distribution in cytoplasm and nuclear invaginations of connective tissue fibroblasts. Histochem Cell Biol, 2007. 127:523-30.
19. Chaudhry, H., R. Schleip, Z. Ji, B. Bukiet, M. Maney, and T. Findley, Three-dimensional mathematical model for deformation of human fasciae in manual therapy. J Am Osteopath Assoc, 2008. 108:379-90.
20. Ritty, T.M., R. Roth, and J.E. Heuser, Tendon cell array isolation reveals a previously unknown fibrillin-2-containing macromolecular assembly. Structure, 2003. 11:1179-88.
21. Ingber, D.E., Cellular tensegrity: defining new rules of biological design that govern the cytoskeleton. J. Cell Sci., 1993. 104:613-627.
22. de La Motte, C.A., V.C. Hascall, A. Calabro, B. Yen-Lieberman, and S.A. Strong, Mononuclear leukocytes preferentially bind via CD44 to hyaluronan on human intestinal mucosal smooth muscle cells after virus infection or treatment with poly(I.C). J Biol Chem, 1999. 274:30747-55.
23. de la Motte, C.A., V.C. Hascall, J. Drazba, S.K. Bandyopadhyay, and S.A. Strong, Mononuclear leukocytes bind to specific hyaluronan structures on colon mucosal smooth muscle cells treated with polyinosinic acid:polycytidylic acid: inter-alpha-trypsin inhibitor is crucial to structure and function. Am J Pathol, 2003. 163:121-33.
24. Okita, M., T. Yoshimura, J. Nakano, M. Motomura, and K. Eguchi, Effects of reduced joint mobility on sarcomere length, collagen fibril arrangement in the endomysium, and hyaluronan in rat soleus muscle. J Muscle Res Cell Motil, 2004. 25:159-66.
25. Evanko, S.P., M.I. Tammi, R.H. Tammi, and T.N. Wight, Hyaluronan-dependent pericellular matrix. Adv Drug Deliv Rev, 2007.
26. Rilla, K., R. Tiihonen, A. Kultti, M. Tammi, and R. Tammi, Pericellular hyaluronan coat visualized in live cells with a fluorescent probe is scaffolded by plasma membrane protrusions. J Histochem Cytochem, 2008.
27. Sauer, S.K., G.M. Bove, B. Averbeck, and P.W. Reeh, Rat peripheral nerve components release calcitonin gene-related peptide and prostaglandin E2 in response to noxious stimuli: evidence that nervi nervorum are nociceptors. Neuroscience, 1999. 92:319-25.
28. Hynds, D.L. and D.M. Snow, Neurite outgrowth inhibition by chondroitin sulfate proteoglycan: stalling/stopping exceeds turning in human neuroblastoma growth cones. Exp Neurol, 1999. 160:244-55.
29. Yamada, H., B. Fredette, K. Shitara, K. Hagihara, R. Miura, B. Ranscht, W.B. Stallcup, and Y. Yamaguchi, The brain chondroitin sulfate proteoglycan brevican associates with astrocytes ensheathing cerebellar glomeruli and inhibits neurite outgrowth from granule neurons. J Neurosci, 1997. 17:7784-95.