Chapter 11Neuroproteomics in the Neocortex of Mammals

Molecular Fingerprints of Cortical Plasticity

Cnops L, Hu TT, Van den Bergh G, et al.

Publication Details

11.1. Introduction

With the accumulation of vast amounts of DNA sequences in databases, researchers are realizing more and more that having complete sequences of genomes is not sufficient to elucidate biological function, not even if complemented with detailed information on the dynamics of the transcriptome. A cell or tissue is dependent on a huge number of metabolic and regulatory pathways for its normal functioning and there is no strict linear relationship between gene expressions and the protein complements or “proteome.” Proteomics is therefore complementary to genomics and transcriptomics because it focuses on the gene products, the true active agents in a cell or tissue at any given time during a given physiological state. The advent of proteomics techniques has been enthusiastically accepted in most areas of biology and medicine. In neuroscience a host of applications was proposed ranging from neurotoxicology, neurometabolism, and determination of the proteomes of individual brain regions in health and disease, to name a few. We have implemented functional proteomics to help unravel the molecular pathways of brain plasticity that drive the response of the mammalian sensory neocortex to stimulus deprivation during development and adulthood, with a special emphasis on the visual system of the cat.

11.2. FACTS ABOUT CAT VISUAL SYSTEM

The visual cortex of mammals is immature at birth, both anatomically and physiologically, and develops gradually in the first weeks and months of postnatal life. During this period, spontaneous brain activity and visual stimulation induce specific patterns of neuronal activity in the central visual system that contribute to the establishment of visual perception and visually guided behavior. Vision-associated neurons will eventually be tuned by the quantity and the quality of the visual stimuli through both eyes to adequately interpret the information encoded in environmental stimulation patterns. To achieve this, a substantial anatomical organization takes place. Strengthening, remodeling, and eliminating synapses help create the adult-specific neuronal circuitry.

In cat, primary visual cortex alternating bands, called ocular dominance columns, contain neurons that are preferentially activated by either the left or the right eye (13). One of the seminal discoveries in developmental neuroscience is that monocular deprivation during the critical period (around postnatal day 30 [P30], for example, by mask rearing or eyelid suture) can modify both the typical physiology of the visual cortex and the anatomical representation of the two eyes in the cortex. The closure or damage of one eye leads to expansion of the columns serving the open eye at the expense of those responding to the deprived eye, which become reduced in size and afferent complexity (1,3–5). This results in poor visual acuity (amblyopia) and contrast sensitivity and loss of depth perception (57). Apparently, such cortical structural rearrangements induced by monocular deprivation are restricted to a certain developmental stage, the critical period that starts around eye opening and ends well before adulthood (79).

Visual acuity is poor at birth and cannot improve in the absence of patterned visual input as established in congenital cataract patients. Early binocular deprivation appears to prevent the normal organization of the cortical neural architecture necessary for the later development of sensitivity to fine detail (10). Thus, equal presentation of visual stimulation to both eyes is in itself insufficient to accurately shape the primary visual cortex. Binocular mask rearing from eye opening in cats, the animal models for congenital cataract, has been documented to lead also to behavioral changes in adulthood, such as deficits in global motion and to a lesser extent global form processing, just like in congenital cataract patients (11,12). Binocular deprivation of patterned vision by mask rearing undeniably affects the functionality of the visual system later in life, emphasizing the importance of the quality of the visual stimulation in shaping the cortex during development, and highlighting the extremely fragile nature of developing binocular circuits soon after birth.

Two decades of research have further shown that even beyond the critical period the visual cortex of the animal does not become a static entity but instead remains malleable throughout its life. Bilateral retinal lesions initially lead to functional deficits. Immediately after causing small lesions on homonymous parts of the retina in both eyes, neurons in the primary visual cortex with a receptive field situated in the center of the damaged region in the retina no longer respond to stimulation within their original receptive field (13–21). Yet neurons at the edges of the cortical lesion projection zone (LPZ) remain responsive but display enlarged receptive fields that are slightly shifted toward the intact retina surrounding the lesion (18). In ensuing weeks and months, recovery of the lost functions occurs. Much larger receptive field shifts take place, which can ultimately result in the complete filling in of the LPZ depending on the size of the retinal lesions (14,18,22,23). This extensive remodeling of cortical topography thus involves an enlargement of the representation of peri-lesion retina at the expense of the cortical area previously dedicated to the lesioned retina (18). The cortical long-range horizontal connections are recognized as the structural mediators of topographic map reorganization in visual cortex, first through the “unmasking” of existing sub-threshold connections, in a second phase by experience-dependent strengthening of their synapses and finally through sprouting of new collaterals (13,15,22,24–29).

How these rearrangements in strength and anatomy of cortical connections in response to abnormal visual stimulation, either during the critical period or in adulthood, depend on changes in the genome and the proteome expressed by the relevant brain cells, has been the subject of intensive investigations and the first molecular models of brain plasticity are now emerging.

11.3. NEUROPROTEOMICS OF CAT VISUAL CORTEX PLASTICITY

We performed large-scale proteomics studies by comparing protein expression profiles of the primary visual cortex of cats during early postnatal development with those of fully mature adult animals (30–32). This started from the rationale that proteins involved in visual cortex plasticity will demonstrate an increase or decrease in expression level with age in order to assist the refinement of visual cortical synaptic connectivity (7,8). To identify as many putative molecular determinants as possible a two-dimensional difference gel electrophoresis (2D-DIGE) approach was applied directly or in combination with a third separation dimension (hydrophobicity) allowing for the selective enrichment and identification of low-abundance proteins using a reversed-phase chromatography pre-fractionation step (see Chapters 3 and 4 for discussion of these techniques) (Figure 11.1). Together these large-scale screenings provided a list of proteins with roles in metabolism, neurite growth and guidance, synapse formation, cytoskeleton stabilization, and neurotransmitter release (30–32).

FIGURE 11.1. Work-flow of the proteomic screening experiments performed on cat visual cortex samples.

FIGURE 11.1

Work-flow of the proteomic screening experiments performed on cat visual cortex samples. The consecutive steps depicted: collection of the brain; production of cryostat sections; sample preparation: manual isolation of specific regions of visual area (more...)

For a detailed understanding of the specific role of each of the proteins in the process of cortical plasticity, a deep analysis is needed about their expression under different physiological conditions using complementary technologies. Because neurite growth and guidance, and formation and maturation of synapses underlies cortical plasticity we specifically investigated the plasticity-regulated expression levels of molecules with a relevant meaning in such structural and cellular processes (33–36). We considered two members of the collapsin response mediator protein family (CRMP2 and CRMP4) and two molecules that are involved in the exocy-tosis-endocytosis machinery at the neuronal synapse, namely dynamin I (Dyn I) and synaptotagmin I (Syt I). CRMPs are phosphoproteins implicated in neuronal differentiation, axon guidance, and growth cone collapse through the signal cascade pathway of Sema3A/collapsin-l (37–39). Dyn I plays a role in synaptic vesicle recycling by clathrin-mediated endocytosis (40), while Syt I functions as a calcium sensor that triggers an essential step in exocytosis, namely the fusion of vesicle and plasma membrane during neurotransmission (41).

Based on the 2D-DIGE spot patterns, multiple isoforms of CRMP2 and CRMP4 were detected as differentially expressed between kitten and adult cat area 17 (Figures 11.2 and 11.3). All CRMP4 isoforms showed a similar developmental expression, while the CRMP2 isoforms displayed a more diverse age-dependent pattern (Figure 11.3A and 11.3B). Dynamin I and synaptotagmin I were each detected in one differential spot and showed an opposite expression (Figures 11.2, 11.3C and 11.3D). Dynamin I levels increased with age while synaptotagmin I levels decreased with age. We thus selected four proteins with a clearly different expression profile for further analysis.

FIGURE 11.3. Detailed visualization of developmental expression differences for CRMP2, CRMP4, Dynl, and Syt I as revealed by 2D-DIGE.

FIGURE 11.3

Detailed visualization of developmental expression differences for CRMP2, CRMP4, Dynl, and Syt I as revealed by 2D-DIGE. Mean relative normalized spot volume ratios of postnatal day 10 (P10) or adult levels to postnatal day 30 (P30) protein levels of (more...)

11.4. CRMPS AND CORTICAL PLASTICITY

Using Western blotting we demonstrated that at least the 64 kDa isoform of CRMP2 and all CRMP4 levels are high early in life (P10, P30) when anatomical and structural rearrangements are necessary for the normal build-up of the visual cortex, in line with a role for these molecules in neuronal differentiation and dendritic and axonal guidance [Figure 11.4; references (42–44)]. Their expression can, however, not be strictly regulated by visual experience since it was already high at eye opening (P10). Also the formation of ocular dominance columns, in contrast to what was believed for a long time, starts experience-independently since clustering of lateral geniculate nucleus (LGN) arbors occurs at least a week before the critical period as recently demonstrated by optical imaging techniques (45,46). Indeed, between postnatal day 10 and 30, the so-called “pre-critical period” (46), spontaneous activity already shapes the visual cortex connectivity and visual experience has then little influence on the cortical organization (47,48). CRMPs may thus, together with other genes, form a molecular cascade that drives the growth of axons and dendrites in the developing cortex during this experience-independent time window.

FIGURE 11.4. Expression patterns of CRMP2, CRMP4, Dyn I, and Syt I during normal visual cortex development.

FIGURE 11.4

Expression patterns of CRMP2, CRMP4, Dyn I, and Syt I during normal visual cortex development. Western blot analysis for CRMP2, CRMP4, Dyn I, and Syt I was performed on visual cortex samples (area 17) of kittens of postnatal day 10 (P10) and 30 (P30), (more...)

The strong downregulation of CRMP4 with visual cortex maturation (Figures 11.3 and 4) correlates with the fact that the closure of the critical period is accompanied by the formation of perineuronal nets through extracellular matrix proteins like chondroitin sulfate proteoglycans that surround predominantly parvalbumin (PV)-containing interneurons (49–53). CRMP4 is known to react with chondroitin sulfates (54) and is co-localized with PV in adult cat visual cortex (55). The perineuronal nets form a mesh, which holds secreted proteins like Sema3A (47,49), possibly resulting in a selective downregulation of CRMP4, one of the intracellular mediators of the Sema3A signaling cascade. Moreover, exactly these PV-containing interneurons seem to be the most important players in regulating the critical period closure (47,51,56). Thus, reduced CRMP4 expression in normal juvenile and adult subjects correlates well with the stabilization of the network connectivity in the maturing visual cortex by extracellular matrix proteins. Moreover, the fact that CRMP4, in contrast to CRMP2, was not affected by monocular deprivation (Figure 11.5) could probably be related to the cell-type specific expression of CRMP2 in large pyramidal neurons and of CRMP4 in those small PV-positive interneurons (55).

FIGURE 11.5. Manipulation-specific molecular expression changes during monocular and binocular deprivation in kittens.

FIGURE 11.5

Manipulation-specific molecular expression changes during monocular and binocular deprivation in kittens. The protein expression levels of CRMP2, CRMP4, Syt I, and Dyn I were determined by Western blotting on normal kittens around the age that the visual (more...)

Monocular deprivation influenced the post-translational modification of CRMP2 as only the 62 kDa CRMP2 protein band was significantly affected by this manipulation (Figure 11.5). These observations probably reflect modifications in the activity status of CRMP2, for instance, by dephosphorylation or deglycosylation. CRMP2 glycosylation has indeed been reported to block CRMP2 phosphorylation, thereby regulating its activity (57). Since the expression of CRMP2 is also modulated during regeneration of the olfactory nerve after axotomy and bulbectomy (58), we suggest that during development CRMP2 could induce rearrangements to LGN axonal arbors and intracortical afferents, like retraction of branches and ingrowth of other axonal branches—structural changes that have been implicated in ocular dominance plasticity. On the other hand, binocular deprivation did not affect CRMP2 or CRMP4, indicating that not the loss of quality of vision through visual deprivation, but the disruption of normal binocular visual experience is crucial to induce the observed changes after monocular deprivation. Opposite to our observations in kittens, both CRMP2 and CRMP4 were clearly affected 14 days after binocular retinal lesion-ing (Figure 11.6). Also, other CRMP studies established the same time-dependent alterations after nervous system injury in adults. By analyzing several post-operative times, a peak of CRMP2 expression was detected in motor neurons of the rat hypoglossal nucleus 14 days after nerve transection (59), while CRMP4 levels were apparent around one or two weeks after ischemia in the striatum of adult rats (60). In addition, many other molecules showed major alterations in expression two weeks to one month after retinal lesions (61–66).

FIGURE 11.6. Western blot analysis to monitor the effect of adult plasticity.

FIGURE 11.6

Western blot analysis to monitor the effect of adult plasticity. The expression of the four molecules of interest was measured in the visual cortex of normal adult cat (N) and adult retinal lesion cats (RL) of different post-lesion survival times: 3 days (more...)

FIGURE 11.2. False-colored two-dimensional difference gel electrophoresis (2D-DIGE) images of the comparative analysis of area 17 protein expression patterns between kitten and adult cat.

FIGURE 11.2

False-colored two-dimensional difference gel electrophoresis (2D-DIGE) images of the comparative analysis of area 17 protein expression patterns between kitten and adult cat. False-colored overlay images of a 2D-DIGE experiment with adult cat, propyl-Cy3-labeled (more...)

CRMP2 and CRMP4 may possibly be involved in a molecular cascade that orchestrates the topographical reorganization of area 17. Indeed, CRMP2 can be phospho-rylated by CaMKII (67), a well-known participant in long term potentiation (LTP) processes and already demonstrated to change its phosphorylated state in retinal lesion cats, especially 14 days post-lesion (66). Another factor with a brain-plasticity-related and post-lesion survival time-dependent expression, MEF2C (63), regulates CRMP4 expression (68). Furthermore, also in the adult, both CRMPs may react with structural molecules that coordinate cytoskeleton rearrangements and synaptic plasticity in the adult cortex, like actin or microtubuli bundles and chondroitin-sulfate proteoglycans of the extracellular matrix (54,56,69–71).

CRMPs were implicated in axonal guidance by mediating the chemorepulsive Sema3A signal (72). As suggested for the olfactory system of adult rat, Sema3A could create a molecular barrier to restrict ingrowing olfactory axons (34,58). High CRMP expression in the LPZ of visually deafferented adult cats after two weeks of recovery thus possibly prevents major ingrowth of neuronal collaterals directly into the most inner part of the LPZ. In addition, overexpression of CRMP2 also stimulates the formation of multiple branches on neuronal processes (69,73) and CRMPs may promote increasing spine dynamics and growth cone formation on existing connections in the LPZ. Indeed, it is known that extending dendritic filopodia may scan the environment for new synaptic partners after sensory deprivation (47,74), while growth cone formation after injury is essential to aid subsequent axonal growth during neuronal regeneration (75). At the same time, CRMP4 possibly plays a neuroprotective role in the LPZ by preventing demyelinization of neurons (34). Later on when CRMP levels decline and thus no longer repel the ingrowth of neuronal connections into the central portion of the LPZ, reorganization by axonal sprouting and synaptogenesis could take place (26,27).

In summary, in the adult, both CRMP2 and CRMP4 may thus promote dendritic spine dynamics or growth cone formation in the LPZ and regulate axonal sprouting in the context of cortical map remodeling in adult area 17. During postnatal development, both CRMPs are involved in structural refinements of the connectivity of the visual cortex, but only changes in CRMP2 activity are necessary for ocular dominance column rearrangements after monocular deprivation.

11.5. DYNAMIN I AND MATURE CELL SHAPE

For Dynamin I a steady increase in expression with age was observed (Figures 11.3 and 11.4). The expression profile of Dyn I thus showed a clear relationship with its function in the maintenance of the adult cell shape (76–78) in full agreement with previous results obtained in rodent studies. Dyn I is not expressed in prenatal rat cortex, cerebellum, or hippocampus and is preferentially upregulated from P7 with a principal increase between P7-P15 and P23 that is maintained into adulthood (76,77). Its developmental course thus parallels neuronal maturation (40) at a time when axonal growth and synapse formation are already established (76,79). Dyn I seems not involved in developmental or plasticity-related synapse rearrangements in cat visual cortex, as demonstrated here in the monocular and binocular deprived kitten models as well as in the adult retinal lesion model (Figures 11.5 and 11.6). Since Dyn I is suggested to be a microtubule-associated motor protein (80), it is more reasonable to believe that Dyn I stabilizes the microtubule cytoskeleton and modulates the dynamics of actin filaments during endocytosis of mature neurons (76,81). Indeed, increasing levels of Dyn I during visual cortex development correspond to the maturation of the neurotransmission system. Once stable synaptic contacts are established, Dyn I-regulated clathrin-mediated endocytosis and vesicle recycling (8284) would then be required for continuous cell-to-cell-communication.

11.6. SYNAPTOTAGMIN I AND CORTICAL PLASTICITY

Syt I is the only molecule whose developmental expression profile readily followed the time course of the critical period (Figure 11.4). In the first phase of the postnatal development, Syt I expression was clearly experience-dependently regulated. Around eye opening (P10), very low levels were detected but at the height of the critical period for monocular deprivation (P30), Syt I levels augmented enormously. Syt I protein expression did not immediately decrease afterwards. In juvenile cats, Syt I demonstrated an even more abundant expression while in adults the expression had declined below P30 levels. Since Syt I is an abundant synaptic vesicle protein and essential in mediating calcium-triggered neurotransmitter release (41,85,86), it could play a role in membrane fusion events that contribute to the outgrowth and remodeling of neuronal dendrites (36). Since the postnatal visual cortex still undergoes structural changes, upregulation of Syt I protein expression by visual experience possibly engages refinements in synaptic contacts. Furthermore, Syt I triggers synchronous and suppresses asynchronous neurotransmitter release (87,88). This could be important for the segregation of the LGN afferents in the eye-specific ocular dominance columns within the primary visual cortex. The high Syt I expression levels at juvenile ages have never been reported before. Even if experience-dependent ocular dominance plasticity at that age is already reduced, it is known that around five months of age, extragranular cortical layers are still susceptible to changes in visual input, while layer IV is not plastic any more (48,8991). Furthermore, at four to six months of age, the kitten visual cortex is still susceptible to effects of monocular deprivation (92). In addition, Winfield (1983) described that the number of synapses increases during development of the cat visual cortex with a maximal density of synapses between P70 and P110 where after the synapse density decreases toward adulthood (93). Also between P8 and P37, an increase in the number of synapses is observed (7) which in turn correlates to the initial increase of Syt I in the visual cortex. Thus, increased Syt I levels correspond well with the boost of synaptic contacts and the correlated increase of neurotransmitter release by Syt I regulated exocytosis.

Upon monocular deprivation, diminished connectivity is detected by reduced size and number of presynaptic terminals, mitochondria, and spines (94). Syt I levels that showed a clear experience-dependent expression pattern during the first stages of normal cat visual cortex development were decreased after monocular deprivation (Figure 11.5), probably in correlation with the reduction of synaptic contacts. As a calcium sensor (41,85,86) it is not unlikely that Syt I detects activity changes in the presynaptic compartment.

However, from our results in adult retinal lesioned cats, we cannot conclude that Syt I was responsible for changes in neurotransmission as observed in retinal lesioned cats [Figure 11.6; references (62,64,65)]. Instead, Syt I may be related to synaptic plasticity only in young animals, in contrast to synapsin, another synaptic vesicle protein that plays a role in both developmental and adult plasticity (6,95). Moreover, some studies doubt the role of Syt I as a Ca2+ sensor and indicate only a regulatory role in the membrane fusion of synaptic vesicles (87,96). Furthermore, the observed modifications in GABAergic and glutamatergic neurotransmission as reported after partial visual deafferentation in adult cats (62,64,65) could be explained by changes in amount of neurotransmitter per vesicle, rather than in the number of vesicles that were released under influence of Syt I.

11.7. CONCLUSION

Taken together, specific molecular fingerprints begin to emerge related to the diverse structural and functional response of the visual cortex when coping with alterations in visual input throughout life. Comparative analysis of protein expression patterns in normal, monocular, and binocular deprived animals seems to hold great promise for dissecting the molecular cascades that specifically encode the quality of vision versus binocular competition to guide cortical maturation. Furthermore, comparison of developmental and adult plasticity models will help elucidate to what extent comparable molecular machinery is exploited by visual neurons in reshaping the cortex in response to lesions later in life. Characterization of manipulation-specific molecular sets may hold the key toward understanding the molecular basis of vision. Extracting insights into physiological and pathological mechanisms from complex sets of neuroproteomic experiments is a fascinating challenge. Future integration of genome, transcriptome, proteome, lipidome, and glycome studies with electrophysiological, anatomical, and behavioral investigations should boost our understanding of vision-guided behavior in response to our daily environment.

Identifying key mediators and defining their relationships should be a major goal in the field and will greatly enhance our understanding of the mechanism by which molecular programs regulate cortical plasticity and underlie visual disorders. Finding the keys to cortical plasticity will enable the development of new therapeutic strategies for recovery from sensory loss and brain damage and for goal-directed improvement of post-lesional recovery of brain function throughout life.

11.8. FUTURE CHALLENGES

The many proteomic and genomic data sets that become available are becoming harder to handle and there is a growing need to collate data from an increasing number of studies. Generating lists alone provides little biological insight. Complementing these lists with bioinformatics, protein interaction studies, and functional studies will be crucial in boosting our understanding of the exact relationship between molecular organization and brain function. There is a need for sharing data among laboratories that should be made available freely as was achieved with genome sequences.

ACKNOWLEDGMENTS

We thank past and present members of our laboratory for their contributions and helpful discussions. Work in our laboratory is supported by grants from the FWO Flanders and the Research Council of the K.U. Leuven.

REFERENCES

1.
LeVay S., Stryker M. P., Shatz C. J. Ocular dominance columns and their development in layer IV of the cat’s visual cortex: A quantitative study. J Comp Neurol. 1978;179:223–44. [PubMed: 8980725]
2.
Payne B. R., Peters A. The cat primary visual cortex. San Diego: Academic Press; 2002.
3.
Shatz C. J., Stryker M. P. Ocular dominance in layer IV of the cat’s visual cortex and the effects of monocular deprivation. J Physiol. 1978;281:267–83. [PMC free article: PMC1282696] [PubMed: 702379]
4.
Hubel D. H., Wiesel T. N. Receptive fields of cells in striate cortex of very young, visually inexperienced kittens. J Neurophysiol. 1963;26:994–1002. [PubMed: 14084171]
5.
Wiesel T. N., Hubel D. H. Comparison of the effects of unilateral and bilateral eye closure on cortical unit responses in kittens. J Neurophysiol. 1965;28:1029–40. [PubMed: 5883730]
6.
Berardi N., Pizzorusso T., Ratto G. M., Maffei L. Molecular basis of plasticity in the visual cortex. Trends Neurosci. 2003;26:369–78. [PubMed: 12850433]
7.
Daw N. W. Visual development. New York: Plenum Press; 1995.
8.
Hubel D. H., Wiesel T. N. The period of susceptibility to the physiological effects of unilateral eye closure in kittens. J Physiol. 1970;206:419–36. [PMC free article: PMC1348655] [PubMed: 5498493]
9.
Shatz C. J., Luskin M. B. The relationship between the geniculocortical afferents and their cortical target cells during development of the cat’s primary visual cortex. J Neurosci. 1986;6:3655–68. [PubMed: 3794795]
10.
Lewis T. L., Maurer D. Multiple sensitive periods in human visual development: Evidence from visually deprived children. Dev Psychobiol. 2005;46:163–83. [PubMed: 15772974]
11.
Burnat K., Stiers P., Arckens L., Vandenbussche E., Zernicki B. Global form perception in cats early deprived of pattern vision. Neuroreport. 2005;16:751–4. [PubMed: 15858419]
12.
Burnat K., Vandenbussche E., Zernicki B. Global motion detection is impaired in cats deprived early of pattern vision. Behav Brain Res. 2002;134:59–65. [PubMed: 12191792]
13.
Chino Y. M. Adult plasticity in the visual system. Can J Physiol Pharmacol. 1995;73:1323–38. [PubMed: 8748982]
14.
Chino Y. M., Kaas J. H., Smith E. L. 3rd, Langston A. L., Cheng H. Rapid reorganization of cortical maps in adult cats following restricted deafferentation in retina. Vision Res. 1992;32:789–96. [PubMed: 1604848]
15.
Chino Y. M., Smith E. L. 3rd, Kaas J. H., Sasaki Y., Cheng H. Receptivefield properties of deafferentated visual cortical neurons after topographic map reorganization in adult cats. J Neurosci. 1995;15:2417–33. [PubMed: 7891177]
16.
Dreher B., Burke W., Calford M. B. Cortical plasticity revealed by circumscribed retinal lesions or artificial scotomas. Prog Brain Res. 2001;134:217–46. [PubMed: 11702546]
17.
Eysel U. T. Cortical plasticity: Remodeling of sensor fields in receptive cortices. Current Biology. 1992;2:389–91.
18.
Gilbert C. D., Wiesel T. N. Receptive field dynamics in adult primary visual cortex. Nature. 1992;356:150–2. [PubMed: 1545866]
19.
Kaas J. H. Plasticity of sensory and motor maps in adult mammals. Annu Rev Neurosci. 1991;14:137–67. [PubMed: 2031570]
20.
Kaas J. H., Krubitzer L. A., Chino Y. M., et al. Reorganization of retinotopic cortical maps in adult mammals after lesions of the retina. Science. 1990;248:229–31. [PubMed: 2326637]
21.
Schmid L. M., Rosa M. G., Calford M. B., Ambler J. S. Visuotopic reorganization in the primary visual cortex of adult cats following monocular and binocular retinal lesions. Cereb Cortex. 1996;6:388–405. [PubMed: 8670666]
22.
Chino Y. M. The role of visual experience in the cortical topographic map reorganization following retinal lesions. Restor Neurol Neurosci. 1999;15:165–76. [PubMed: 12671231]
23.
Giannikopoulos D. V., Eysel U. T. Dynamics and specificity of cortical map reorganization after retinal lesions. Proc Natl Acad Sci U S A. 2006;103:10805–10. [PMC free article: PMC1487171] [PubMed: 16818873]
24.
Calford M. B. Mechanisms for acute changes in sensory maps. Adv Exp Med Biol. 2002;508:451–60. [PubMed: 12171142]
25.
Calford M. B., Wright L. L., Metha A. B., Taglianetti V. Topographic plasticity in primary visual cortex is mediated by local corticocortical connections. J Neurosci. 2003;23:6434–42. [PubMed: 12878683]
26.
Darian-Smith C., Gilbert C. D. Axonal sprouting accompanies functional reorganization in adult cat striate cortex. Nature. 1994;368:737–40. [PubMed: 8152484]
27.
Darian-Smith C., Gilbert C. D. Topographic reorganization in the striate cortex of the adult cat and monkey is cortically mediated. J Neurosci. 1995;15:1631–47. [PubMed: 7891124]
28.
Das A., Gilbert C. D. Long-range horizontal connections and their role in cortical reorganization revealed by optical recording of cat primary visual cortex. Nature. 1995;375:780–4. [PubMed: 7596409]
29.
Das A., Gilbert C. D. Receptive field expansion in adult visual cortex is linked to dynamic changes in strength of cortical connections. J Neurophysiol. 1995;74:779–92. [PubMed: 7472382]
30.
Van den Bergh G., Clerens S., Cnops L., Vandesande F., Arckens L. Fluorescent two-dimensional difference gel electrophoresis and mass spectrometry identify age-related protein expression differences for the primary visual cortex of kitten and adult cat. J Neurochem. 2003;85:193–205. [PubMed: 12641741]
31.
Van den Bergh G., Clerens S., Firestein B. L., Burnat K., Arckens L. Development and plasticity-related changes in protein expression patterns in cat visual cortex: A fluorescent two-dimensional difference gel electrophoresis approach. Proteomics. 2006;6:3821–32. [PubMed: 16739136]
32.
Van den Bergh G., Clerens S., Vandesande F., Arckens L. Reversed-phase high-performance liquid chromatography prefractionation prior to two-dimensional difference gel electrophoresis and mass spectrometry identifies new differentially expressed proteins between striate cortex of kitten and adult cat. Electrophoresis. 2003;24:1471–81. [PubMed: 12731035]
33.
Byk T., Ozon S., Sobel A. The Ulip family phosphoproteins—Common and specific properties. Eur J Biochem. 1998;254:14–24. [PubMed: 9652388]
34.
Charrier E., Reibel S., Rogemond V., et al. Collapsin response mediator proteins (CRMPs): Involvement in nervous system development and adult neurodegenerative disorders. Mol Neurobiol. 2003;28:51–64. [PubMed: 14514985]
35.
Napolitano M., Marfia G. A., Vacca A., et al. Modulation of gene expression following long-term synaptic depression in the striatum. Brain Res Mol Brain Res. 1999;72:89–96. [PubMed: 10521602]
36.
Schwab Y., Mouton J., Chasserot-Golaz S., et al. Calcium-dependent translocation of synaptotagmin to the plasma membrane in the dendrites of developing neurones. Brain Res Mol Brain Res. 2001;96:1–13. [PubMed: 11731003]
37.
Goshima Y., Sasaki Y., Nakayama T., Ito T., Kimura T. Functions of semaphorins in axon guidance and neuronal regeneration. Jpn J Pharmacol. 2000;82:273–9. [PubMed: 10875745]
38.
Minturn J. E., Fryer H. J., Geschwind D. H., Hockfield S. TOAD-64, a gene expressed early in neuronal differentiation in the rat, is related to unc-33, a C. elegans gene involved in axon outgrowth. J Neurosci. 1995;15:6757–66. [PubMed: 7472434]
39.
Wang L. H., Strittmatter S. M. A family of rat CRMP genes is differentially expressed in the nervous system. J Neurosci. 1996;16:6197–6207. [PubMed: 8815901]
40.
Liu J. P., Robinson P. J. Dynamin and endocytosis. Endocr Rev. 1995;16:590–607. [PubMed: 8529573]
41.
Sudhof T. C., Rizo J. Synaptotagmins: C2-domain proteins that regulate membrane traffic. Neuron. 1996;17:379–88. [PubMed: 8816702]
42.
Byk T., Dobransky T., Cifuentes-Diaz C., Sobel A. Identification and molecular characterization of Unc-33-like phosphoprotein (Ulip), a putative mammalian homolog of the axonal guidance-associated unc-33 gene product. J Neurosci. 1996;16:688–701. [PubMed: 8551352]
43.
Kamata T., Subleski M., Hara Y., et al. Isolation and characterization of a bovine neural specific protein (CRMP-2) cDNA homologous to unc-33, a C. elegans gene implicated in axonal outgrowth and guidance. Brain Res Mol Brain Res. 1998;54:219–36. [PubMed: 9555025]
44.
Polleux F., Morrow T., Ghosh A. Semaphorin 3A is a chemoattractant for cortical apical dendrites. Nature. 2000;404:567–73. [PubMed: 10766232]
45.
Crair M. C., Horton J. C., Antonini A., Stryker M. P. Emergence of ocular dominance columns in cat visual cortex by 2 weeks of age. J Comp Neurol. 2001;430:235–49. [PMC free article: PMC2412906] [PubMed: 11135259]
46.
Feller M. B., Scanziani M. A precritical period for plasticity in visual cortex. Curr Opin Neurobiol. 2005;15:94–100. [PubMed: 15721750]
47.
Bence M., Levelt C. N. Structural plasticity in the developing visual system. Prog Brain Res. 2005;147:125–39. [PubMed: 15581702]
48.
Katz L. C., Shatz C. J. Synaptic activity and the construction of cortical circuits. Science. 1996;274:1133–8. [PubMed: 8895456]
49.
Dityatev A., Schachner M. Extracellular matrix molecules and synaptic plasticity. Nat Rev Neurosci. 2003;4:456–68. [PubMed: 12778118]
50.
Guimaraes A., Zaremba S., Hockfield S. Molecular and morphological changes in the cat lateral geniculate nucleus and visual cortex induced by visual deprivation are revealed by monoclonal antibodies Cat-304 and Cat-301. J Neurosci. 1990;10:3014–24. [PubMed: 1697900]
51.
Hensch T. K. Critical period plasticity in local cortical circuits. Nat Rev Neurosci. 2005;6:877–88. [PubMed: 16261181]
52.
Hockfield S., Kalb R. G., Zaremba S., Fryer H. Expression of neural proteoglycans correlates with the acquisition of mature neuronal properties in the mammalian brain. Cold Spring Harb Symp Quant Biol. 1990;55:505–14. [PubMed: 2132834]
53.
Lander C., Kind P., Maleski M., Hockfield S. A family of activity-dependent neuronal cell-surface chondroitin sulfate proteoglycans in cat visual cortex. J Neurosci. 1997;17:1928–39. [PubMed: 9045722]
54.
Franken S., Junghans U., Rosslenbroich V., et al. Collapsin response mediator proteins of neonatal rat brain interact with chondroitin sulfate. J Biol Chem. 2003;278:3241–50. [PubMed: 12444086]
55.
Cnops L., Hu T. T., Burnat K., Van der Gucht E., Arckens L. Age-dependent alterations in CRMP2 and CRMP4 protein expression profiles in cat visual cortex. Brain Res. 2006;1088:109–19. [PubMed: 16630590]
56.
Berardi N., Pizzorusso T., Maffei L. Extracellular matrix and visual cortical plasticity: Freeing the synapse. Neuron. 2004;44:905–8. [PubMed: 15603733]
57.
Cole R. N., Hart G. W. Cytosolic O-glycosylation is abundant in nerve terminals. J Neurochem. 2001;79:1080–9. [PubMed: 11739622]
58.
Pasterkamp R. J., De Winter F., Holtmaat A. J., Verhaagen J. Evidence for a role of the chemorepellent semaphorin III and its receptor neuropilin-1 in the regeneration of primary olfactory axons. J Neurosci. 1998;18:9962–76. [PubMed: 9822752]
59.
Suzuki Y., Nakagomi S., Namikawa K., et al. Collapsin response mediator protein-2 accelerates axon regeneration of nerve-injured motor neurons of rat. J Neurochem. 2003;86:1042–50. [PubMed: 12887701]
60.
Liu P. C., Yang Z. J., Qiu M. H., Zhang L. M., Sun F. Y. Induction of CRMP-4 in striatum of adult rat after transient brain ischemia. Acta Pharmacol Sin. 2003;24:1205–11. [PubMed: 14653945]
61.
Arckens L. The molecular biology of sensory map plasticity in adult mammals. Pinaud R., Tremere L. A., De Weerd P., editors. Springer Science + Business Media: Plasticity in the visual system: From genes to circuits. 2006:181–203.
62.
Arckens L., Schweigart G., Qu Y., et al. Cooperative changes in GABA, glutamate and activity levels: The missing link in cortical plasticity. Eur J Neurosci. 2000;12:4222–32. [PubMed: 11122334]
63.
Leysen I., Van der Gucht E., Eysel U. T., et al. Time-dependent changes in the expression of the MEF2 transcription factor family during topographic map reorganization in mammalian visual cortex. Eur J Neurosci. 2004;20:769–80. [PubMed: 15255987]
64.
Massie A., Cnops L., Jacobs S., et al. Glutamate levels and transport in cat (Felis catus) area 17 during cortical reorganization following binocular retinal lesions. J Neurochem. 2003;84:1387–97. [PubMed: 12614339]
65.
Massie A., Cnops L., Smolders I., et al. Extracellular GABA concentrations in area 17 of cat visual cortex during topographic map reorganization following binocular central retinal lesioning. Brain Res. 2003;976:100–8. [PubMed: 12763627]
66.
Van den Bergh G., Eysel U. T., Vandenbussche E., Vandesande F., Arckens L. Retinotopic map plasticity in adult cat visual cortex is accompanied by changes in Ca2+/calmodulin-dependent protein kinase II alpha autophosphorylation. Neuroscience. 2003;120:133–42. [PubMed: 12849747]
67.
Yoshimura Y., Shinkawa T., Taoka M., et al. Identification of protein substrates of Ca(2+)/calmodulin-dependent protein kinase II in the postsynaptic density by protein sequencing and mass spectrometry. Biochem Biophys Res Commun. 2002;290:948–54. [PubMed: 11798165]
68.
Matsuo T., Stauffer J. K., Walker R. L., Meltzer P., Thiele C. J. Structure and promoter analysis of the human unc-33-like phosphoprotein gene. E-box required for maximal expression in neuroblastoma and myoblasts. J Biol Chem. 2000;275:16560–8. [PubMed: 10748015]
69.
Fukata Y., Kimura T., Kaibuchi K. Axon specification in hippocampal neurons. Neurosci Res. 2002;43:305–15. [PubMed: 12135774]
70.
Gu Y., Ihara Y. Evidence that collapsin response mediator protein-2 is involved in the dynamics of microtubules. J Biol Chem. 2000;275:17917–20. [PubMed: 10770920]
71.
Quinn C. C., Gray G. E., Hockfield S. A family of proteins implicated in axon guidance and outgrowth. J Neurobiol. 1999;41:158–64. [PubMed: 10504203]
72.
Goshima Y., Nakamura F., Strittmatter P., Strittmatter S. M. Collapsininduced growth cone collapse mediated by an intracellular protein related to UNC-33. Nature. 1995;376:509–14. [PubMed: 7637782]
73.
Inagaki N., Chihara K., Arimura N., et al. CRMP-2 induces axons in cultured hippocampal neurons. Nat Neurosci. 2001;4:781–2. [PubMed: 11477421]
74.
Lee W. C., Huang H., Feng G., et al. Dynamic remodeling of dendritic arbors in GABAergic interneurons of adult visual cortex. PLoS Biol. 2006;4:e29. [PMC free article: PMC1318477] [PubMed: 16366735]
75.
Geddis M. S., Rehder V. The phosphorylation state of neuronal processes determines growth cone formation after neuronal injury. J Neurosci Res. 2003;74:210–20. [PubMed: 14515350]
76.
Faire K., Trent F., Tepper J. M., Bonder E. M. Analysis of dynamin isoforms in mammalian brain: Dynamin-1 expression is spatially and temporally regulated during postnatal development. Proc Natl Acad Sci U S A. 1992;89:8376–80. [PMC free article: PMC49921] [PubMed: 1387713]
77.
Nakata T., Iwamoto A., Noda Y., et al. Predominant and developmentally regulated expression of dynamin in neurons. Neuron. 1991;7:461–9. [PubMed: 1832879]
78.
Noda Y., Nakata T., Hirokawa N. Localization of dynamin: Widespread distribution in mature neurons and association with membranous organelles. Neuroscience. 1993;55:113–27. [PubMed: 8350983]
79.
Powell K. A., Robinson P. J. Dephosphin/dynamin is a neuronal phosphoprotein concentrated in nerve terminals: evidence from rat cerebellum. Neuroscience. 1995;64:821–33. [PubMed: 7715790]
80.
Shpetner H. S., Vallee R. B. Identification of dynamin, a novel mechanochemical enzyme that mediates interactions between microtubules. Cell. 1989;59:421–32. [PubMed: 2529977]
81.
Schafer D. A. Coupling actin dynamics and membrane dynamics during endocytosis. Curr Opin Cell Biol. 2002;14:76–81. [PubMed: 11792548]
82.
Sontag J. M., Fykse E. M., Ushkaryov Y., et al. Differential expression and regulation of multiple dynamins. J Biol Chem. 1994;269:4547–54. [PubMed: 8308025]
83.
Urrutia R., Henley J. R., Cook T., McNiven M. A. The dynamins: Redundant or distinct functions for an expanding family of related GTPases? Proc Natl Acad Sci U S A. 1997;94:377–84. [PMC free article: PMC34135] [PubMed: 9012790]
84.
van der Bliek A. M., Redelmeier T. E., Damke H., et al. Mutations in human dynamin block an intermediate stage in coated vesicle formation. J Cell Biol. 1993;122:553–63. [PMC free article: PMC2119674] [PubMed: 8101525]
85.
Littleton J. T., Bellen H. J. Synaptotagmin controls and modulates synapticvesicle fusion in a Ca(2+)-dependent manner. Trends Neurosci. 1995;18:177–83. [PubMed: 7778189]
86.
Ullrich B., Sudhof T. C. Differential distributions of novel synaptotagmins: Comparison to synapsins. Neuropharmacology. 1995;34:1371–7. [PubMed: 8606786]
87.
Yoshihara M., Adolfsen B., Littleton J. T. Is synaptotagmin the calcium sensor? Curr Opin Neurobiol. 2003;13:315–23. [PubMed: 12850216]
88.
Yoshihara M., Littleton J. T. Synaptotagmin I functions as a calcium sensor to synchronize neurotransmitter release. Neuron. 2002;36:897–908. [PubMed: 12467593]
89.
Beaver C. J., Ji Q., Daw N. W. Layer differences in the effect of monocular vision in light- and dark-reared kittens. Vis Neurosci. 2001;18:811–20. [PubMed: 11925016]
90.
Daw N. W. Mechanisms of plasticity in the visual cortex. The Friedenwald Lecture. Invest Ophthalmol Vis Sci. 1994;35:4168–79. [PubMed: 8002237]
91.
Daw N. W., Fox K., Sato H., Czepita D. Critical period for monocular deprivation in the cat visual cortex. J Neurophysiol. 1992;67:197–202. [PubMed: 1552319]
92.
Cynader M., Timney B. N., Mitchell D. E. Period of susceptibility of kitten visual cortex to the effects of monocular deprivation extends beyond six months of age. Brain Res. 1980;191:545–50. [PubMed: 7378770]
93.
Winfield D. A. The postnatal development of synapses in the different laminae of the visual cortex in the normal kitten and in kittens with eyelid suture. Brain Res. 1983;285:155–69. [PubMed: 6616262]
94.
Tieman S. B. Morphological changes in the geniculocortical pathway associated with monocular deprivation. Ann N Y Acad Sci. 1991;627:212–30. [PubMed: 1679310]
95.
Obata S., Obata J., Das A., Gilbert C. D. Molecular correlates of topographic reorganization in primary visual cortex following retinal lesions. Cereb Cortex. 1999;9:238–48. [PubMed: 10355904]
96.
Koh T. W., Bellen H. J. Synaptotagmin I, a Ca2+ sensor for neurotransmitter release. Trends Neurosci. 2003;26:413–22. [PubMed: 12900172]
97.
Van den Bergh G., Arckens L. Fluorescent two-dimensional difference gel electrophoresis unveils the potential of gel-based proteomics. Curr Opin Biotechnol. 2004;15:38–43. [PubMed: 15102464]