Molecular Vision 2006; 12:76-84 <>
Received 27 January 2005 | Accepted 24 January 2006 | Published 3 February 2006

Induction of vitronectin and integrin αv in the retina after optic nerve injury

An-Guor Wang,1,2,3 May-Yung Yen,2,3 Wen-Ming Hsu,2,3 Ming-Ji Fann1,4,5

1Institute of Neuroscience, 2Faculty of Medicine, and 4Department of Life Sciences, National Yang-Ming University, Taipei, Taiwan; 3Department of Ophthalmology, Taipei Veterans General Hospital, Taipei, Taiwan; 5Brain Research Center, University System of Taiwan, Taipei, Taiwan

Correspondence to: Dr. Ming-Ji Fann, Department of Life Sciences, National Yang-Ming University, Taipei, Taiwan 11221, Republic of China; Phone: 886-2-2826-7184; FAX: 886-2-2820-0259; email:


Purpose: Vitronectin is a secreted glycoprotein present in blood plasma and is present in the extracellular matrix of many tissues. It was found in the retinal cDNA library that contains genes whose expression is upregulated after optic nerve injury in a previous study. The purpose of this study was to assess the temporal and spatial changes in expression of vitronectin and integrin αv in the retina following optic nerve injury.

Methods: Adult Balb/c mice underwent crush of the optic nerve in one eye only. RT-PCR was used to determine the temporal expression of vitronectin mRNA in the retina after injury. In addition, expression at the protein level in the retina and the optic nerve of vitronectin and its major receptor subunit, integrin αv, was analyzed by immunohistochemistry.

Results: Upregulation of vitronectin mRNA in the retina was detected at one day after injury, peaking at three days, and maintained up to one week. An elevated expression of vitronectin protein was also observed in the inner retina, optic nerve head, and the optic nerve after nerve crush. In the inner retina, the increased expression of vitronectin was found in retinal ganglion cells (RGCs) and its surrounding extracellular matrix. Expression of integrin αv was also increased in the RGC layer and in the glial cells of the nerve head and the crush site.

Conclusions: As vitronectin is an extracellular protein that can support cell attachment and promote neurite extension, elevated expression of vitronectin and its receptor may facilitate axonal regeneration following injury. We propose that treatment sustaining secretion of endogenous vitronectin or direct application of exogenous vitronectin may be a method to augment regeneration of the severed optic axons.


Vitronectin is a secreted glycoprotein present in blood plasma and is present in the extracellular matrix of many tissues [1]. It is a multiadhesion molecule and interacts with extracellular glycosaminoglycans, collagen, and other extracellular proteins [2]. Moreover, through the RGD sequence, vitronectin can bind integrins on the cell surface, thus connecting cells to matrix proteins [3]. Circulating vitronectin is synthesized in the liver, and its concentration in the blood plasma is upregulated during systemic inflammation [4]. However, significant amounts of vitronectin mRNA have also been detected in many other tissues, such as brain and heart, indicating that vitronectin can be produced locally [4,5].

In addition to being a pivotal regulator in the complement, fibrinolytic, and coagulation systems, vitronectin plays a number of roles in the retina. Vitronectin mRNA is detected in the neural retina, including photoreceptor cells and retinal ganglion cells (RGCs) [6], and in the retinal pigment epithelium (RPE) [7]. The integrin αvβ5 vitronectin receptor is also expressed on the apical membrane of human RPE. This vitronectin receptor participates in the binding of photoreceptor rod outer segment during phagocytosis by cultured human RPE [8]. Moreover, in age-related macular degeneration, vitronectin is a major component of the distinct extracellular deposits that accumulate below the RPE on Bruch's membrane and is suspected as a possible causative factor that results in the compromised central vision [9]. Several lines of evidence also suggest that vitronectin influences neurite outgrowth of developing retinal cells. Vitronectin is expressed in the chick eye from embryonic day 5 (E5) [10]. Expression of vitronectin receptors, αvβ3 and αvβ5, is also detected in developing chick retina [11]. Cultured embryonic chick retinas attach and extend neurites on vitronectin-coated dishes [12]. Furthermore, purified chick RGCs from E10 chick embryo grow better on vitronectin than on laminin [13]. These results show that vitronectin can support survival and neurite outgrowth of differentiating retinal neurons [10]. However, the effects of vitronectin on adult neural retina are still not clear.

In adult mammals, severed axons within the optic nerve may show a transient local sprouting but do not regrow over long distances [14]. This failure of regeneration is usually attributed to the formation of a gliotic scar [15], the presence of extracellular inhibitors [16-20], intrinsic changes in the gene expression of mature neurons [21,22], and massive apoptosis of damaged neurons [23-25]. By neutralizing hostile factors, potentiating neuronal outgrowth ability, and maintaining cell survival, one may be able to facilitate functional recovery of visual pathways [26]. Although many studies have demonstrated how to counteract inhibitory molecules and support neuronal survival [27-29], few investigations have focused on intrinsic gene expression necessary for neuronal regeneration [30,31].

Previously, we looked for genes that may behave in a similar manner as GAP43, which is known to augment neuronal regeneration after injury [32,33], using a subtractive hybridization approach [34]. Several genes that are upregulated in the retina following optic nerve injury at different time points were identified [34]. Among these genes, vitronectin was detected with a high frequency in the subtractive libraries. Here, we used reverse transcription-polymerase chain reaction (RT-PCR) and immunohistochemistry to further explore the expression pattern of vitronectin throughout the injured retina. The expression of one of its receptor subunits, integrin αv [35-38], was also analyzed. We find that after optic nerve crush both proteins are induced in RGCs and in glial cells that are present in the nerve head and at the injured site.


Animals and surgery

Adult Balb/c mice that were at least two months old and weighing over 25 g were obtained from the animal facility of National Yang-Ming University. The experimental protocol was approved by the National Yang-Ming University Animal Care and Use Committee. Institute guidelines were followed on handling of animals. Mice were given food and water ad libitum. For the optic nerve crush, mice were anesthetized by injecting a combination of ketamine (0.1 mg/g body weight) and xylocaine (0.03 mg/g body weight) intraperitonially. Surgery for all mice was conducted by the same surgeon under sterile conditions using a stereomicroscope. A conjunctival incision was made over the dorsal aspect of one eye, which was then gently rotated downward in the orbit. The orbital muscles were teased and deflected aside to expose the optic nerve at its exit from the globe. A jewelers forceps was used to crush the optic nerve twice across the entire width near the back of the eye (within 0.5 mm). Care was taken not to damage the ophthalmic artery and retrobulbar sinus. The eye was then rotated back into position and rinsed with sterile saline. Mice were allowed to recover. The contralateral eye without surgery was used as control. At different time points (one day, three days, and seven days) after surgery, the retina and the attached nerve were dissected out for analysis.


Superscript reverse transcriptase II (RTase) was purchased from Invitrogen (Carlsbad, CA). dATP, dCTP, dGTP, and dTTP were obtained from Roche (Mannheim, Germany). RNase inhibitor (RNasin) and Taq DNA polymerase were purchased from Promega (Madison, WI). Primers were synthesized by MDBiol (Taipei, Taiwan). All other chemicals, unless specified, were purchased from Merck (Darmstadt, Germany).

RT-PCR and quantitative image analysis

Total RNA of five retinas from each group at each time point was prepared using RNeasy Mini kit according to the user manual (Qiagen, Valencia, CA). Total RNA was denatured at 75 °C for 5 min. Reverse transcription was performed in a total volume of 20 μl containing 50 mM Tris-HCl, pH 8.3, 75 mM KCl, 3 mM MgCl2, 10 mM DTT, 0.5 mM dNTP, 0.5 μg of oligo dT, 20 U of RNasin, and 100 U of RTase for 1.5 h at 50 °C. Samples were stored at -20 °C until use. Primers used for PCR amplification were as follows: for GAPDH (NM_008084, from nucleotide position 565 to 1006), ACC ACA GTC CAT GCC ATC AC and TCC ACC ACC CTG TTG CTG TA; for vitronectin (NM_011707, from nucleotide position 114 to 498), TCC CTT GAT CAG TGG TGT CGG and CTG GCT GAC CAA GAG TCA TGC. The expected size of PCR fragment for GAPDH is 452 bp and for vitronectin is 385 bp. Amplification of cDNA was performed in a GeneAmp 9700 (Applied Biosystems, Foster City, CA) with each tube containing a final volume of 50 μl, consisting of 1 μl of cDNA, 1X PCR buffer, 1.25 U of Taq DNA polymerase, 0.25 mM dNTP, and one set of primers (200 nM). Samples were covered with 15 μl of mineral oil and were first denatured at 94 °C for 30 s, then cycles of 94 °C for 5 s, 56 °C for 15 s, 72 °C for 30 s were performed. Reaction was paused at each cycle after cycle numbers 23 to 27, and 8 μl of PCR products were retrieved before resumption of the PCR. Samples were analyzed on a 2% agarose gel, and DNA was visualized with ethidium bromide staining and UV illumination. Gel intensity after cycle 25 was analyzed using the ImageQuant software (version 1.1, Molecular Dynamics, Sunnyvale, CA). A scanning area was chosen around the target band on the gel, and an area of the same size was used for each sample analyzed at the various time points. The local median was selected as the method for background correction. The volume for each target band was measured and expressed as the ratio relative to GAPDH. Statistical analysis was performed with the paired t-test.


The injured eye and control eye were enucleated at one day, three days, and one week postsurgery (four mice for each time points), and maintained in Hank's solution. After the anterior segments were removed, posterior eyecups were fixed in 4% paraformaldehyde solution for 1 h at room temperature. The eyeballs were transferred to 30% sucrose to sit overnight, were then submerged in Tissue-Tek OCT media (Sakura Finetechnical Co., Tokyo, Japan) before being frozen on dry ice. Sections were cut at 12 μm thick. Crushed and control retina sections were collected on the same slides (SuperFrost, Fisher Scientific, Pittsburgh, PA).

For staining, sections were fixed with 4% paraformaldehyde solution for one h at room temperature, washed with phosphate buffer solution (PBS)/0.1% Tween for 15 min at room temperature, followed by PBS for 15 min twice. Blocking was done with 20% normal goat serum or 5% bovine serum albumin (BSA) for 1 h at room temperature, and then the sections were stained with rabbit antivitronectin antiserum (Biotrend, Cologne, Germany) or anti-integrin αv antiserum (Chemicon, Temecula, CA), and antineurofilament monoclonal antibody (heavy, 200 kDa, clone RT 97, Roche) diluted in 3% normal goat serum or 5% BSA at 4 °C overnight. Sections blocking with 20% normal goat serum or 5% BSA gave similar results. The rabbit antivitronectin antiserum recognizes mouse vitronectin but does not recognize goat vitronectin. One set of slides without primary antibody was used as control in each experiment. Sections were then washed with PBS three times and incubated with fluorescein-conjugated goat antirabbit antibody and rhodamine-conjugated goat antimouse antibody (Cappel, Aurora, OH) at room temperature for 2 h. For nuclear staining, 4',6-diamidino-2-phenyindole (DAPI, 0.16 μg/ml) was added for 15 min at room temperature, and washed with PBS three times. Sections were analyzed with a Nikon Diaphot inverted microscope (E300, Nikon, Tokyo, Japan) equipped with a MicroMax cool CCD (Princeton Instrument, Trenton, NJ). Image acquisition was performed using MetaMorph software (version 4.5r6, Universal Image Corporation, Downingtown, PA) with individual filter sets for each channel and were assembled with Adobe PhotoShop (version 7.0, Adobe Systems, San Jose, CA).


Induction of vitronectin mRNA after optic nerve injury

To pursue whether vitronectin mRNA expression in the retina is elevated after damage in the optic nerve, a semiquantitative RT-PCR assay was employed to measure amount of vitronectin mRNA in the retina. A representative gel of this analysis is shown in Figure 1A. The expression levels of vitronectin relative to those of GAPDH were then calculated using the ImageQuant software. The statistical analysis of three independently prepared RNA sets was performed and is shown in Figure 1B. The results reveal that expression of vitronectin had increased by two fold on day one and day three after optic nerve crush in comparison with the control samples. Although the induction of vitronectin mRNA was still visible at seven days after injury, this increase was not statistically significant.

Vitronectin and integrin αv expression in normal eye

To characterize whether the increase in vitronectin mRNA expression is accompanied with synthesis of vitronectin protein and to clarify whether the effect is in the RGCs, we first examined the expression pattern of vitronectin in the normal eye using antivitronectin antiserum. The expression pattern of the vitronectin receptor was also investigated using anti-integrin αv antiserum, since integrin αv is the major α subunit of vitronectin receptors [35-38]. In the normal eye, vitronectin was found in RPE, inner segments of photoreceptors, the outer plexiform layer, and the inner retina (Figure 2D). A diffuse expression pattern of vitronectin was observed, which may indicate the extracellular distribution of secreted vitronectin. In the inner retina, the staining for vitronectin was present in RGCs and in the extracellular matrix that surrounds the RGCs (Figure 2F,G). The retinal ganglion layer was identified by the DAPI nuclear staining (Figure 2G). To outline the location of nerve fiber layer in the retina, we labeled the sections with an antineurofilament monoclonal antibody. Neurofilament staining revealed the axonal fibers in the nerve fiber layer, which represent the projection of the RGCs (Figure 2C, J). The merged picture (Figure 2E) showed the vitronectin expresses over the nerve fiber layer in addition to the RGC layer. We also examined the expression of integrin αv in retina. It was observed in the RGC layer, inner and outer nuclear layers, the outer plexiform layer, inner segments of photoreceptors, and RPE (Figure 2K). Staining of integrin αv was detected on each cell in the RGC layer with a membrane-staining pattern (Figure 2M, arrows). In control, experiments which were performed without adding primary antibodies, no staining was observed in the inner retina (data not shown). As displaced amacrine cells are present in the RGC layer, we can not exclude the possibility that some immunopositive cells in the RGC layer were actually amacrine cells.

The expression patterns of vitronectin and integrin αv in the optic nerve head were investigated. Optic axon fibers originating from the nerve fiber layer converge on the optic nerve head and exit the eyeball through the scleral canal. Within the optic nerve head, the nerve bundles proceed in parallel and were stained with the antineurofilament antibody (Figure 2Q,V). In between the nerve bundles, columns of glial cells could be identified with DAPI staining (Figure 2P,U). The vitronectin expression in exiting axon fibers was much weaker than that of the inner retina layer (Figure 2R), which could be due to less accumulation of vitronectin in the exiting axons. A similar pattern of integrin αv expression was also present in the optic nerve head (Figure 2W).

Increase of vitronectin and integrin αv expression in the retina after nerve crush

When the optic nerve was subjected to crush injury, the vitronectin expression became elevated at the inner retina (Figure 3). The RGC layer and nerve fiber layer can be clearly recognized with antivitronectin antibodies after optic nerve crush. The induction of vitronectin was present on day one after crush, culminated at day three, and was maintained at day seven. Surprisingly, induction of integrin αv was also observed in the inner retina using antibodies against integrin αv (Figure 3). Expression of integrin αv was elevated in RGCs from one day to one week after crush. This increase of expression was unique for integrin αv, as no upregulation of α5 which binds fibronectin but not vitronectin was detected (data not shown). Among three integrin β subunits that bind vitronectin, β5 is upregulated in the RGC layer at day three after optic nerve injury (data not shown), suggesting the possibility that vitronectin can have effects on RGC by interacting with αvβ5.

Expression of vitronectin and integrin αv in the optic nerve

The differences of vitronectin and integrin αv expression between the crushed and control optic nerves were also examined. After optic nerve injury, the expression of vitronectin had increased at the optic nerve head at day one (data not shown) and on day three post-crush (Figure 4F). Similarly, increased expression of integrin αv was observed in the optic nerve head three days after the crush injury (Figure 4N). Induction of vitronectin did not co-localize with glia cells marked by DAPI, and seemed likely to be in the extracellular matrix of optic nerve head. The increased expression of integrin αv was primarily located in glial cells, which are distributed either within the septum or around the central retinal artery tract. In the optic nerve proper, faint immunoreactivities of vitronectin and integrin αv were observed in the normal nerve (Figure 4Q,U). After optic nerve crush, the crush sites (arrows in Figure 4R,V) could be clearly identified by the abrupt interruption of neurofilament staining (data not shown) in the optic nerve. Many glial cells at crush sites showed an induction of vitronectin expression (Figure 4S,T). There was also a higher level of vitronectin expression in extracellular matrix. Within the crush line, several cells with an increase of integrin αv expression were also found (Figure 4W,X).


In a previous study [34], we analyzed the early change in gene expression in the retina after an optic nerve crush by a suppression subtractive hybridization approach. Among the genes detected in the analysis, vitronectin was found in a subtractive retinal cDNA library that contains genes whose expression is upregulated at 24 h after the nerve injury [34]. To verify this observation, we examined whether a rise of vitronectin mRNA and protein could be detected after the nerve injury. Our results demonstrate that there is a 2 fold upregulation of vitronectin mRNA in the retina between one day and three days following optic nerve injury. Moreover, elevated expression of both vitronectin and integrin αv proteins was present in the inner retina, the optic nerve head, and the optic nerve. As the massive death of RGCs does not occur until seven days after injury [23,24], this induction is thus unlikely due to the result of apoptosis of RGCs.

Several lines of evidence have implicated that both extraneuronal factors and intrinsic alterations in aged neurons result in the limited capacity for axonal growth in the adult mammalian RGCs [16,20-22]. It is known that myelin-derived growth inhibitors, such as Nogo [17], MAG [18,19], and OMgp [39]; gliotic scar [15]; and extracellular matrix molecules, like chondroitin sulfate proteoglycans [40-42] and tenascin [43,44] inhibit axonal growth in the adult CNS. Furthermore, adult RGCs show a progressive loss of responsiveness to factors that stimulate axon outgrowth, thus restricting the extent of RGC regrowth after crush injury [21,22]. For example, RGCs from E6 chicks, but not those from E11, extend profuse neurites on laminin [45]; and RGCs from E14 hamsters can regrow into adult tectum under an explant coculture paradigm, but postnatal RGCs cannot innervate tectal targets, even when the target is embryonic [21]. Thus, to facilitate successful regeneration after retinal axonal injury, it is necessary to enhance expression of intrinsic growth-related genes in adult RGC neurons, while neutralizing the extraneuronal inhibitory molecules. Supporting this thought, it has been shown that ectopic expression of GAP43 and CAP 23, two genes involved in the axonal outgrowth, in adult CNS neurons stimulate their regeneration [31].

Vitronectin is present in many different tissues, including brain [1], during development. Vitronectin immunoreactivity has been found in most neuroepithelial cells at E5 in chick retina, and on the nerve fiber and inner plexiform layer at E9 [10]. In adult human, expression of vitronectin is found in the photoreceptor layer and RGC layer of retina and in the septa stroma of optic nerve [6,46]. In our study expression of vitronectin is similarly detected in adult mouse retina, specifically in the RPE, photoreceptor, and RGC (Figure 2C). For expression of vitronectin receptors, it was previously reported that integrin αv is expressed in developing chick retina at least since E6 [11], and on the apical membrane of fetal human RPE and adult primate photoreceptors and RPE [8,47]. We found that, in addition to RPE, the expression of integrin αv in adult mouse retina is present in the RGC layer, inner and outer nuclear layers, the outer plexiform layer, and inner segments of photoreceptors (Figure 2K). Functionally, vitronectin regulates cell differentiation, neuronal survival, and neurite outgrowth in the developing chicken retina [10], and purified E10 chick RGCs grow better on vitronectin than on laminin [13]. These studies implicate vitronectin as an important component in the retinal extracellular matrix for axonal growth. Our data here shows that there is an upregulation of vitronectin in the inner retina, and this may indicate an attempt of RGCs to repair after injury by generating a favorable extracellular environment. The upregulation of integrin αv could also render RGCs responsive to this changing milieu.

An increase in vitronectin and integrin αv is observed in glial cells within the optic nerve head after injury. Changes of extracellular matrix components have been observed following brain injury [48,49], and there is evidence that integrin expression on astrocytes is upregulated in pathological states [50]. Whether this phenomenon represents an overall beneficial or detrimental effect is yet to be fully determined. As different subtypes of integrin expressed on astrocytes affect the permissiveness for neurite outgrowth in a coculture system [51,52], and as vitronectin is known to promote axonal growth [10,12,13], upregulation of integrin αv and vitronectin in the astrocytes within the optic nerve may contribute to RGC axonal regeneration by fostering a growth-promoting environment. On the contrary, another possible scenario after optic nerve injury is that the increase of vitronectin expression in the optic nerve may result from an extravasation of serum vitronectin through damaged vessels at the crush injury. With increased expression of integrin on cell surface, glial cells migrate and proliferate in response to the increased vitronectin, which in turn promotes reactive gliosis and hinders the regeneration of RGC axons in the optic nerve head [53]. In the retina, however, extravasation of plasma vitronectin is less likely to occur, since a local elevation of vitronectin mRNA is detected in the retina, and the injured site lies far away from the retina.

In conclusion, our results show that there is an increase of vitronectin and integrin αv in the retina and the optic nerve after optic nerve injury. Induction of vitronectin and integrin αv may have an important role in the regeneration of adult mammalian RGCs. Thus, it will be interesting to test whether an exogenously applied stimulus, which elevates the amount of vitronectin in the retinal nerve fiber layer and optic nerve and enhances expression of integrin α in RGC, could facilitate the visual recovery.


The authors thank L.-S. Kao for helpful comments and critical reading of the manuscript. This paper was supported by grants from Taipei Veterans General Hospital (VGH 91-217), National Science Council (90-2314-B-075-130), and VGH-UST (VTY91-P1-11) of Tsou's Foundation to A.-G. Wang and M.-J. Fann, and National Health Research Institutes (NHRI-GT-EX89S732C) to M.-J. Fann.


1. Kreis T, Vale R, editors. Guidebook to the extracellular matrix, anchor, and adhesion proteins. 2nd ed. Oxford: Oxford University Press; 1999. p. 496-497.

2. Preissner KT. Structure and biological role of vitronectin. Annu Rev Cell Biol 1991; 7:275-310.

3. Schvartz I, Seger D, Shaltiel S. Vitronectin. Int J Biochem Cell Biol 1999; 31:539-44.

4. Seiffert D, Crain K, Wagner NV, Loskutoff DJ. Vitronectin gene expression in vivo. Evidence for extrahepatic synthesis and acute phase regulation. J Biol Chem 1994; 269:19836-42.

5. Seiffert D. Constitutive and regulated expression of vitronectin. Histol Histopathol 1997; 12:787-97.

6. Anderson DH, Hageman GS, Mullins RF, Neitz M, Neitz J, Ozaki S, Preissner KT, Johnson LV. Vitronectin gene expression in the adult human retina. Invest Ophthalmol Vis Sci 1999; 40:3305-15.

7. Ozaki S, Johnson LV, Mullins RF, Hageman GS, Anderson DH. The human retina and retinal pigment epithelium are abundant sources of vitronectin mRNA. Biochem Biophys Res Commun 1999; 258:524-9.

8. Lin H, Clegg DO. Integrin alphavbeta5 participates in the binding of photoreceptor rod outer segments during phagocytosis by cultured human retinal pigment epithelium. Invest Ophthalmol Vis Sci 1998; 39:1703-12.

9. Crabb JW, Miyagi M, Gu X, Shadrach K, West KA, Sakaguchi H, Kamei M, Hasan A, Yan L, Rayborn ME, Salomon RG, Hollyfield JG. Drusen proteome analysis: an approach to the etiology of age-related macular degeneration. Proc Natl Acad Sci U S A 2002; 99:14682-7.

10. Martinez-Morales JR, Marti E, Frade JM, Rodriguez-Tebar A. Developmentally regulated vitronectin influences cell differentiation, neuron survival and process outgrowth in the developing chicken retina. Neuroscience 1995; 68:245-53.

11. Gervin DB, Cann GM, Clegg DO. Temporal and spatial regulation of integrin vitronectin receptor mRNAs in the embryonic chick retina. Invest Ophthalmol Vis Sci 1996; 37:1084-96.

12. Neugebauer KM, Emmett CJ, Venstrom KA, Reichardt LF. Vitronectin and thrombospondin promote retinal neurite outgrowth: developmental regulation and role of integrins. Neuron 1991; 6:345-58.

13. Brocco MA, Panzetta P. Survival and process regrowth of purified chick retinal ganglion cells cultured in a growth factor lacking medium at low density. Modulation by extracellular matrix proteins. Brain Res Dev Brain Res 1999; 118:23-32.

14. Ramon y Cajal S, DeFelipe J, Jones EG. Cajal's degeneration and regeneration of the nervous system. New York: Oxford University Press; 1991. p. 100-126.

15. Vernadakis A. Neuron-glia interrelations. Int Rev Neurobiol 1988; 30:149-224.

16. Davies SJ, Goucher DR, Doller C, Silver J. Robust regeneration of adult sensory axons in degenerating white matter of the adult rat spinal cord. J Neurosci 1999; 19:5810-22.

17. GrandPre T, Nakamura F, Vartanian T, Strittmatter SM. Identification of the Nogo inhibitor of axon regeneration as a Reticulon protein. Nature 2000; 403:439-44.

18. McKerracher L, David S, Jackson DL, Kottis V, Dunn RJ, Braun PE. Identification of myelin-associated glycoprotein as a major myelin-derived inhibitor of neurite growth. Neuron 1994; 13:805-11.

19. Mukhopadhyay G, Doherty P, Walsh FS, Crocker PR, Filbin MT. A novel role for myelin-associated glycoprotein as an inhibitor of axonal regeneration. Neuron 1994; 13:757-67.

20. Schwab ME, Caroni P. Oligodendrocytes and CNS myelin are nonpermissive substrates for neurite growth and fibroblast spreading in vitro. J Neurosci 1988; 8:2381-93.

21. Chen DF, Jhaveri S, Schneider GE. Intrinsic changes in developing retinal neurons result in regenerative failure of their axons. Proc Natl Acad Sci U S A 1995; 92:7287-91.

22. Shen S, Wiemelt AP, McMorris FA, Barres BA. Retinal ganglion cells lose trophic responsiveness after axotomy. Neuron 1999; 23:285-95.

23. Berkelaar M, Clarke DB, Wang YC, Bray GM, Aguayo AJ. Axotomy results in delayed death and apoptosis of retinal ganglion cells in adult rats. J Neurosci 1994; 14:4368-74.

24. Garcia-Valenzuela E, Gorczyca W, Darzynkiewicz Z, Sharma SC. Apoptosis in adult retinal ganglion cells after axotomy. J Neurobiol 1994; 25:431-8.

25. Villegas-Perez MP, Vidal-Sanz M, Bray GM, Aguayo AJ. Influences of peripheral nerve grafts on the survival and regrowth of axotomized retinal ganglion cells in adult rats. J Neurosci 1988; 8:265-80.

26. Goldberg JL, Barres BA. The relationship between neuronal survival and regeneration. Annu Rev Neurosci 2000; 23:579-612.

27. Schnell L, Schwab ME. Axonal regeneration in the rat spinal cord produced by an antibody against myelin-associated neurite growth inhibitors. Nature 1990; 343:269-72.

28. Bregman BS, Kunkel-Bagden E, Schnell L, Dai HN, Gao D, Schwab ME. Recovery from spinal cord injury mediated by antibodies to neurite growth inhibitors. Nature 1995; 378:498-501.

29. Cai D, Shen Y, De Bellard M, Tang S, Filbin MT. Prior exposure to neurotrophins blocks inhibition of axonal regeneration by MAG and myelin via a cAMP-dependent mechanism. Neuron 1999; 22:89-101.

30. Buffo A, Holtmaat AJ, Savio T, Verbeek JS, Oberdick J, Oestreicher AB, Gispen WH, Verhaagen J, Rossi F, Strata P. Targeted overexpression of the neurite growth-associated protein B-50/GAP-43 in cerebellar Purkinje cells induces sprouting after axotomy but not axon regeneration into growth-permissive transplants. J Neurosci 1997; 17:8778-91.

31. Bomze HM, Bulsara KR, Iskandar BJ, Caroni P, Skene JH. Spinal axon regeneration evoked by replacing two growth cone proteins in adult neurons. Nat Neurosci 2001; 4:38-43.

32. Schaden H, Stuermer CA, Bahr M. GAP-43 immunoreactivity and axon regeneration in retinal ganglion cells of the rat. J Neurobiol 1994; 25:1570-8.

33. Doster SK, Lozano AM, Aguayo AJ, Willard MB. Expression of the growth-associated protein GAP-43 in adult rat retinal ganglion cells following axon injury. Neuron 1991; 6:635-47.

34. Wang AG, Chen CH, Yang CW, Yen MY, Hsu WM, Liu JH, Fann MJ. Change of gene expression profiles in the retina following optic nerve injury. Brain Res Mol Brain Res 2002; 101:82-92.

35. Marshall JF, Rutherford DC, McCartney AC, Mitjans F, Goodman SL, Hart IR. Alpha v beta 1 is a receptor for vitronectin and fibrinogen, and acts with alpha 5 beta 1 to mediate spreading on fibronectin. J Cell Sci 1995; 108:1227-38.

36. Brooks PC, Clark RA, Cheresh DA. Requirement of vascular integrin alpha v beta 3 for angiogenesis. Science 1994; 264:569-71.

37. Smith JW, Vestal DJ, Irwin SV, Burke TA, Cheresh DA. Purification and functional characterization of integrin alpha v beta 5. An adhesion receptor for vitronectin. J Biol Chem 1990; 265:11008-13.

38. Nishimura SL, Sheppard D, Pytela R. Integrin alpha v beta 8. Interaction with vitronectin and functional divergence of the beta 8 cytoplasmic domain. J Biol Chem 1994; 269:28708-15.

39. Wang KC, Koprivica V, Kim JA, Sivasankaran R, Guo Y, Neve RL, He Z. Oligodendrocyte-myelin glycoprotein is a Nogo receptor ligand that inhibits neurite outgrowth. Nature 2002; 417:941-4.

40. Jones LL, Tuszynski MH. Spinal cord injury elicits expression of keratan sulfate proteoglycans by macrophages, reactive microglia, and oligodendrocyte progenitors. J Neurosci 2002; 22:4611-24.

41. Jones LL, Yamaguchi Y, Stallcup WB, Tuszynski MH. NG2 is a major chondroitin sulfate proteoglycan produced after spinal cord injury and is expressed by macrophages and oligodendrocyte progenitors. J Neurosci 2002; 22:2792-803.

42. Lemons ML, Howland DR, Anderson DK. Chondroitin sulfate proteoglycan immunoreactivity increases following spinal cord injury and transplantation. Exp Neurol 1999; 160:51-65.

43. Becker T, Anliker B, Becker CG, Taylor J, Schachner M, Meyer RL, Bartsch U. Tenascin-R inhibits regrowth of optic fibers in vitro and persists in the optic nerve of mice after injury. Glia 2000; 29:330-46.

44. Becker CG, Becker T, Meyer RL, Schachner M. Tenascin-R inhibits the growth of optic fibers in vitro but is rapidly eliminated during nerve regeneration in the salamander Pleurodeles waltl. J Neurosci 1999; 19:813-27.

45. Cohen J, Burne JF, Winter J, Bartlett P. Retinal ganglion cells lose response to laminin with maturation. Nature 1986; 322:465-7.

46. Fukuchi T, Sawaguchi S, Ueda J, Abe H. [Distribution of cell adhesion glycoproteins in the human retrobulbar optic nerve]. Nippon Ganka Gakkai Zasshi 1997; 101:57-63.

47. Anderson DH, Johnson LV, Hageman GS. Vitronectin receptor expression and distribution at the photoreceptor-retinal pigment epithelial interface. J Comp Neurol 1995; 360:1-16.

48. Hertel M, Tretter Y, Alzheimer C, Werner S. Connective tissue growth factor: a novel player in tissue reorganization after brain injury? Eur J Neurosci 2000; 12:376-80.

49. Liesi P, Kaakkola S, Dahl D, Vaheri A. Laminin is induced in astrocytes of adult brain by injury. EMBO J 1984; 3:683-6.

50. Sobel RA, Chen M, Maeda A, Hinojoza JR. Vitronectin and integrin vitronectin receptor localization in multiple sclerosis lesions. J Neuropathol Exp Neurol 1995; 54:202-13.

51. Smith-Thomas LC, Fok-Seang J, Stevens J, Du JS, Muir E, Faissner A, Geller HM, Rogers JH, Fawcett JW. An inhibitor of neurite outgrowth produced by astrocytes. J Cell Sci 1994; 107:1687-95.

52. Milner R, Relvas JB, Fawcett J, ffrench-Constant C. Developmental regulation of alphav integrins produces functional changes in astrocyte behavior. Mol Cell Neurosci 2001; 18:108-18.

53. Gladson CL, Stewart JE, Olman MA, Chang PL, Schnapp LM, Grammer JR, Benveniste EN. Attachment of primary neonatal rat astrocytes to vitronectin is mediated by integrins alphavbeta5 and alpha8beta1: modulation by the type 1 plasminogen activator inhibitor. Neurosci Lett 2000; 283:157-61.

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