www.elsevier.com/locate/ymcne

Mol. Cell. Neurosci. 32 (2006) 133 – 142

Reelin is transiently expressed in the peripheral nerve during

development and is upregulated following nerve crush

Roger Panteri,a,* Jorg Mey,b Nina Zhelyaznik,b Anna D’Altocolle,c Aurora Del Fa,c

Carlo Gangitano,c Ramona Marino,a Erika Lorenzetto,d Mario Buffelli,d and Flavio Kellera,*

aLaboratorio di Neuroscienze dello Sviluppo, Universita ‘‘Campus Bio-Medico’’, 00155 Roma, ItalybInstitut fur Biologie II, RWTH Aachen, Kopernikusstr. 16, 52074 Aachen, GermanycIstituto di Anatomia Umana e Biologia Cellulare, Universita Cattolica del Sacro Cuore, 00168 Roma, ItalydDipartimento di Scienze Neurologiche e della Visione, Universita di Verona, Strada Le Grazie n.8, 37134 Verona, Italy

Received 27 June 2005; revised 17 January 2006; accepted 16 March 2006

Available online 11 May 2006

Reelin is an extracellular matrix protein which is critical for the

positioning of migrating post-mitotic neurons and the laminar

organization of several brain structures during development. We

investigated the expression and localization of Reelin in the rodent

peripheral nerve during postnatal development and following crush

injury in the adult stage. As shown with Western blotting, immuno-

cytochemistry and RT-PCR, Schwann cells in the developing periph-

eral nerve and in primary cultures from neonatal nerves produce and

secrete Reelin. While Reelin levels are downregulated in adult stages,

they are again induced following sciatic nerve injury. A morphometric

analysis of sciatic nerve sections of reeler mice suggests that Reelin is

not essential for axonal ensheathment by Schwann cells, however, it

influences the caliber of myelinated axons and the absolute number of

fibers per unit area. This indicates that Reelin may play a role in

peripheral nervous system development and repair by regulating

Schwann cell–axon interactions.

D 2006 Elsevier Inc. All rights reserved.

Keywords: Reelin; Reeler; Schwann cell; Nerve development; Nerve injury;

Morphometry

Introduction

Reelin is a large protein that is secreted into the extracellular

matrix and is critical for the laminar organization of several brain

structures during development (D’Arcangelo et al., 1995; Rice and

Curran, 2001; Tissir and Goffinet, 2003). Reelin is secreted by

Cajal–Retzius cells located in the marginal zone of the developing

cerebral cortex and regulates positioning of post-mitotic migratory

neurons. A well-established signaling pathway of Reelin involves

1044-7431/$ - see front matter D 2006 Elsevier Inc. All rights reserved.

doi:10.1016/j.mcn.2006.03.004

* Corresponding authors. Fax: +39 06 22 54 14 56.

E-mail addresses: R.Panteri@unicampus.it (R. Panteri),

f.keller@unicampus.it (F. Keller).

Available online on ScienceDirect (www.sciencedirect.com).

binding to the lipoprotein receptors VLDLR and ApoER2 followed

by tyrosine phosphorylation of the intracellular adaptor Dab1,

leading to cytoskeletal rearrangements and gene expression

changes in the target neurons (D’Arcangelo et al., 1999; Hiesberger

et al., 1999). Reelin, in addition to its well-known function in

neuronal migration, also appears to have a role in the development

and synaptogenesis of hippocampal connections since its absence

leads to alterations in the entorhino-hippocampal pathway,

including reduced axonal branching, an increase in the number

of misrouted aberrant fibers and fewer entorhino-hippocampal

synapses (Del Rio et al., 1997; Borrell et al., 1999). Although most

studies have focused on Reelin in the control of migration and cell

positioning during central nervous system development, little is

known about the role of Reelin in the development of the

peripheral nervous system (PNS) (Ikeda and Terashima, 1997).

Given its putative role in axonal growth and synaptogenesis, we

were prompted to investigate whether Reelin is expressed in the

peripheral nervous system at the early stages of postnatal

development and during nerve regeneration following injury, i.e.

during two stages characterized by axon growth and extracellular

matrix remodeling (Kury et al., 2001; Corfas et al., 2004). Thus, in

the present study, we investigated with RT-PCR, Western blotting

and immunohistochemistry the expression and localization of

Reelin in the mouse sciatic nerve during the early phases of

postnatal development, in the adult stage and following crush

injury. Our findings were extended by analysis on Schwann cell

primary cultures. To investigate whether Reelin plays a role in

axon–Schwann cell interactions in the peripheral nervous system,

a morphometric analysis was conducted on semithin sciatic nerve

sections of reeler mice. Our studies revealed that Reelin is

expressed in the rodent developing peripheral nerve and in vitro

by primary Schwann cell cultures. Levels of Reelin are down-

regulated in adult stages, however, these are induced again

following crush injury to the sciatic nerve. While the morphometric

study suggests that Reelin is not required for the myelination

process, because its absence does not perturb the axon–Schwann

R. Panteri et al. / Mol. Cell. Neurosci. 32 (2006) 133–142134

cell relationship, Reelin appears to subtly influence the caliber of

myelinated axons and the absolute number of myelinated axons per

unit area.

Results

Reelin is expressed in early postnatal mouse sciatic nerves

To investigate the expression of Reelin mRNA in rodent

peripheral nerves, we conducted an RT-PCR analysis on total

RNA extracted from the sciatic nerves of young postnatal (P9)

wild-type (WT), heterozygous reeler (HZ) and homozygous reeler

(RL) mice. Examination of the amplified product (591 bp Reelin

fragment; Fig. 1A) reveals the presence of Reelin mRNA in sciatic

nerve samples prepared from HZ and WT mice, whereas the signal

is absent in the sample prepared from a RL mouse of the same

age. To confirm expression at the protein level and to study

isoforms of Reelin expressed in the peripheral nerve, we

performed Western blot experiments on sciatic nerve protein

extracts of young postnatal WT and RL mice using the

Fig. 1. (A) Expression of Reelin mRNA in early postnatal mouse sciatic nerve. The

is present in extracts of WT (wild-type) and HZ (heterozygous) mice sciatic nerves

(B) Western blot on WT/RL sciatic nerve protein extracts indicates Reelin expres

expressing cells (CER); lane 2, brain extracts from WT at P9; lane 3, brain extracts

nerve extracts from RL at P9. Arrows indicate the two Reelin isoforms at ¨400 kD

as in the sciatic nerve extracts of WT mice at P9. Molecular weight protein marker

for the brain extracts and 15 Ag for the sciatic extracts. (C) Developmental time co

extracts from two different WT mice at P1 (lanes 1, 2), at P4 (lanes 3, 4), at P7

Approximately 10 Ag of total protein was loaded per lane. (D) The net optical intenC and plotted against developmental time.

monoclonal antibody Ab142 that recognizes an epitope localized

in the N-terminal region of Reelin (De Bergeyck et al., 1998). We

found that addition of 1% SDS and 100 mM h-Mercaptoethanol to

the homogenization buffer was critical for the detection of Reelin

from sciatic nerve preparations. As a positive control, we used the

supernatant of Reelin expressing cells (CER, Niu et al., 2004; Fig.

1B, lane 1). The experiment shows in the lane containing the cell

supernatant a Reelin-specific band recognized by Ab142 at ¨400

kDa (full-length protein), plus two additional bands at ¨300 and

320 kDa and another Reelin isoform at ¨150 kDa. In the lanes

containing the sciatic nerve protein extracts from a WT mouse at

P9 (Fig. 1B, lane 4), the Ab142 recognizes the full-length Reelin

band at ¨400 kDa and the lower isoform band at ¨150 kDa,

which are absent in the lane containing the sciatic nerve protein

extracts from a RL mouse at P9 (Fig. 1B, lane 5). The results also

indicate that the ¨400 kDa and the ¨150 kDa isoforms detected

in the WT P9 sciatic nerve are also present in the extracts of the

WT P9 brain (Fig. 1B, lane 2), but not in the extracts of the RL P9

brain (Fig. 1B, lane 3). The smear observed in the high MW range

of the WT brain extract lane is due perhaps to an insufficient

separation of the full-length glycosylated Reelin isoforms which

expected molecular weight of the RT-PCR product is 591 bp. Reelin mRNA

at postnatal day 9 and absent in extracts of RL (reeler) mice of the same age.

sion in early postnatal mouse sciatic nerves. Lane 1, supernatant of Reelin

from RL at P9; lane 4, sciatic nerve extracts from WT at P9; lane 5, sciatic

a and¨150 kDa present in the supernatant of Reelin expressing cells as well

s are indicated on the left. Approximately 35 Ag of total protein was loaded

urse of Reelin expression in the sciatic nerve. Western blot on sciatic nerve

(lanes 5, 6), at P12 (lanes 7, 8), P15 (lanes 9, 10) and P60 (lanes 11, 12).

sity values were measured for the bands of Reelin isoforms obtained in panel

R. Panteri et al. / Mol. Cell. Neurosci. 32 (2006) 133–142 135

tend to form aggregates (Kubo et al., 2002). In another

experiment, permanence of the Reelin signal was observed after

perfusion with saline prior to removal of the sciatic nerve,

showing that these peripheral Reelin isoforms were not contrib-

uted by circulating plasma (data not shown; Smalheiser et al.,

2000). The results show that early postnatal mouse peripheral

nerves express physiological isoforms of Reelin in a similar way

to those expressed in the central nervous system at the same time

of early postnatal development.

Reelin expression is regulated during development

In order to understand how levels of Reelin are regulated during

development, we performed Western blot analysis of protein

samples obtained from WT mouse sciatic nerves at several

developmental time points. The results indicate that all three major

Reelin isoforms at ¨400 kDa, 300 kDa and 150 kDa are regulated

during development in the mouse sciatic nerve (Fig. 1C). At the

earliest age examined, postnatal day 1 (P1), Reelin is expressed at

low levels, however, Reelin expression increases with age to reach

peak levels around postnatal day 12 (P12) and then decreases

moderately at postnatal day 15 to reach again low levels of

expression in adulthood. The relative densitometric analysis

performed on the Reelin isoform bands indicates that Reelin

expression at P4 and P7 is approximately twice and at P12 four

times that corresponding to the P1 stage, while Reelin expression at

adult stage (P60) is similar to that observed at the P1 stage (Fig.

1D). Interestingly, the highest levels of Reelin expression in the

sciatic nerve are observed at P12–P15 stage when the process of

axon myelination is advanced and almost completed, and

numerous events of axon growth and retraction take place during

synapse elimination of supernumerary motor axons in the

peripheral nervous system (Jessen and Mirsky, 1999; Sanes and

Lichtman, 1999).

Fig. 2. Expression of Reelin in Schwann cell cultures in vitro. (A) RT-PCR analy

sciatic nerves of newborn rats. The expected molecular weight of the RT-PCR prod

showing the Reelin-specific PCR product; lane 5: PCR without reverse transcriptio

by Schwann cells into the supernatant. Lanes 1, 2: supernatant of Schwann cells c

cells. (C) Analysis of Reelin expression in Schwann cell primary cultures by imm

Reelin monoclonal antibody E4; the nuclei were counterstained with DAPI. (Panel

Positive immunoreactivity of Schwann cells for S100. (Panel 4) Phase contrast ima

same magnification in all photographs.

Reelin is expressed by Schwann cells in vitro

We hypothesized that Schwann cells produce Reelin in the PNS.

To test this, we analyzed Reelin expression in purified Schwann cell

cultures prepared from neonatal rats (Fig. 2A). Reelin transcripts

could be detected by RT-PCR in samples of Schwann cell cultures

(Fig. 2A, lanes 1–4), whereas no signal was observed in the absence

of reverse transcription (Fig. 2A, �RT). The expression of Reelin

was confirmed with immunoblots of Schwann cell supernatants

and control media, probed with the monoclonal antibody E4

(recognizing an epitope localized in the N-terminal region of

Reelin, De Bergeyck et al., 1998; Fig. 2B). In the blot, we

observed a strong Reelin band at ¨350 kDa and a less intense

Reelin isoform at ¨150 kDa in the lanes containing supernatants

from Schwann cell cultures maintained for 6 days in DMEM/B27

supplement (Fig. 2B, lanes 1, 2), whereas no Reelin bands were

present in the lane containing the medium alone (Fig. 2B, lane 3).

Reelin protein expression during PNS development was confirmed

by immunocytochemistry in Schwann cell primary cultures from

P1–2 rat sciatic nerves (Fig. 2C). Reelin immunoreactivity,

detected with the mouse monoclonal anti-Reelin E4 antibody,

was present in these cultured Schwann cells, and staining appeared

to be cytosolic and perinuclear (Fig. 2C, panel 1). Control

experiments with no primary antibodies confirmed the specificity

of Reelin immunostaining in the Schwann cells (Fig. 2C, panel 2).

The same Schwann cell cultures showed a typical staining pattern

for the Schwann cell marker S100 (Fig. 2C, panel 3). The

corresponding phase contrast image showed the elongated, spindle

morphology typical of Schwann cells (Fig. 2C, panel 4). A few

fibroblasts that were occasionally observed in these primary

cultures were immunonegative for Reelin and S100. These data

demonstrate that Schwann cells derived from neonatal rat sciatic

nerves and maintained in vitro can produce Reelin mRNA and can

also synthesize and secrete physiological Reelin isoforms.

sis of Reelin mRNA expression in Schwann cells cultures derived from the

uct is 501 bp. Lanes 1–4: RNA extract from different Schwann cell cultures

n of the RNA extract. (B) Western blot showing secretion of Reelin isoforms

ultured with DMEM plus B27 supplement; lane 3: culture medium without

unofluorescence. (Panel 1) Positive staining of Schwann cells by the anti-

2) Control staining of Schwann cells without the primary antibody. (Panel 3)

ge of the Schwann cells shown in panel 3. Scale bar marks 20 Am in panel 1,

Fig. 3. Reelin expression is induced following crush lesion of adult mouse

sciatic nerves. (A) Western blot on adult WT (P34) sciatic nerve extracts at

several time points after crush. Lane 1, adult WT, sham side, 2 days post-

lesion; lanes 2, 3, 2 different adult WT, crushed side, 2 days post-lesion; lane

4, adult WT, sham side, 4 days post-lesion; lanes 5, 6, 2 different adult WT,

crushed side, 4 days post-lesion; lane 7, adult WT, sham side, 7 days post-

lesion; lanes 8, 9, 2 different adult WT, crushed side, 7 days post-lesion; lane

10, adult WT, sham side, 10 days post-lesion; lanes 11, 12, 2 different adult

WT, crushed side, 10 days post-lesion. Approximately 10 Ag of total proteinwere loaded per lane. (B) The net optical intensity values of the Reelin bands

shown in A were measured and plotted against time after lesion.

R. Panteri et al. / Mol. Cell. Neurosci. 32 (2006) 133–142136

Reelin is upregulated following crush injury of adult peripheral

nerve

Since the expression data show that levels of Reelin in the

adult sciatic nerve are low but detectable (Fig. 1C), we were

interested in examining whether the pattern of Reelin expression

would change in regenerating sciatic nerves of adult mice. We

thus performed crush injuries and contralateral sham operations to

the sciatic nerves of adult WT mice and analyzed Reelin levels

through Western blot experiments on sciatic nerve extracts from

the injured and uninjured sides distal to the lesion at several time

points after the crush injury (Fig. 3). At early time points after the

lesion, the process of Wallerian degeneration (WD) causes the

progressive degradation of axons and the breakdown of myelin

sheaths distal to the site of injury. This process implies the

downregulation of genes that encode myelin-related proteins such

as P0 and the re-expression of proteins which are characteristic of

non-myelinating and pre-myelinating Schwann cells such as p75

(Jessen et al., 1990; Corfas et al., 2004). Interestingly, we found

that levels of the three major Reelin isoforms at ¨400 kDa, 300

kDa and 150 kDa are upregulated in the distal portion of the

sciatic nerve as soon as 2 days after nerve injury (Fig. 3A).

Reelin upregulation continues progressively in the crushed nerve

at 4 days post-injury, reaches peak levels at 7 days post-injury

and appears to decrease slightly at 10 days post-injury. The

relative densitometric analysis performed on the Reelin isoform

bands indicates that Reelin expression in the crushed sciatic nerve

at 7 days post-injury is approximately four to five times the

physiological levels of Reelin observed in the contralateral sham

side (Fig. 3B). These data thus demonstrate the upregulation of

Reelin after crush injury of the adult peripheral nerve at a time

when Schwann cells dedifferentiate during the process of WD.

Reelin is expressed by Schwann cells in the peripheral nerve

To investigate the localization of Reelin in the sciatic nerve, we

performed immunocytochemistry experiments on crushed and

sham-lesioned adult mouse WT sciatic nerves. Co-localization

experiments were performed on the same sciatic nerve sections

using anti-Reelin (E4) and anti-myelin basic protein (MBP)

antibodies. The results indicate an almost identical distribution of

Reelin (green) and MBP (red) along intact myelinated nerve fibers

in the non-injured sciatic nerve (Figs. 4A and B); the overlaid

image confirms the co-localization of Reelin and MBP (yellow) in

the compact myelin portion of nerve fibers (Fig. 4C). In the

crushed nerve, 7 days after injury, Reelin, like MBP, shows an

altered distribution in the region distal to the injury site where

patches of Reelin immunoreactivity associate with myelin under-

going fragmentation (Figs. 4D and E); the overlaid image shows

the co-localization of Reelin and MBP in myelin ovoids that result

from myelin degeneration (Fig. 4F). Control experiments with no

primary antibodies were also performed and confirmed the

specificity of Reelin (Fig. 4G) and MBP (Fig. 4H) immunostaining

in the sciatic nerve. As a further control, we incubated adult RL

normal sciatic nerve sections with the E4 anti-Reelin antibody and

obtained a negative staining for Reelin (Fig. 4I). Since the pattern

of Reelin immunoreactivity was different in the crushed and sham-

lesioned sciatic nerves, we could not quantify the Reelin

immunohistochemical signal across the two conditions. To extend

our analysis, we also performed co-localization experiments on

normal adult WT sciatic nerve sections using antibodies against

Reelin and the Schwann cell marker S100. The results indicate

positive immunoreactivities of Reelin (red) and S100 (green)

associated with Schwann cells surrounding the intact myelinated

fibers (Figs. 5A and B); the overlaid image confirms the co-

localization of Reelin and S100 (Fig. 5C). These results indicate

that Reelin is expressed by Schwann cells in the sciatic nerve and

appears to be localized to the compact myelin portion of

myelinated nerve fibers.

Absence of Reelin affects the number and caliber of myelinated

axons

To explore whether the absence of Reelin affects the mye-

lination process and the morphology of axon–Schwann cell units,

we performed a morphometric analysis of myelinated fibers on

semithin transverse sections obtained from WT and RL mice at

P15. At this time, when the process of myelin wrapping around

axons by Schwann cells is advanced although not completed, we

observe prominent Reelin expression in the peripheral nerve (Fig.

1C), and in the absence of Reelin, there is the onset of high

frequency tremors, ataxia and loss of motor coordination typical of

RL mice (Tissir and Goffinet, 2003). Our intention was to

investigate the hypothesis that part of this phenotype may be due

to a peripheral neuropathy and thus search for possible alterations

of myelinated nerve fibers in the PNS of RL mice. The

morphometric study of the sciatic nerves of WT and RL mice

Fig. 4. Reelin and MBP immunofluorescence in adult mouse sciatic nerves. (A and B) Double immunofluorescence with antibodies against Reelin (green) and

MBP (red) on the sham-lesioned sciatic nerve. Reelin and MBP immunoreactivities are associated with the myelin portion of intact nerve fibers. (C) Overlay of

the images in A and B shows co-localization of Reelin and MBP along myelinated fibers. (D–E) Crushed sciatic nerve, 7 days after injury, shows patches of

Reelin and MBP immunoreactivity associated with myelin undergoing fragmentation in the distal region of the nerve (arrows in panel D); (F) overlay of the

images in panels D and E shows co-localization of Reelin and MBP in myelin ovoids. (G and H) Control staining procedures with no anti-Reelin (G) or anti-

MBP (H) primary antibodies. (I) Adult RL sham sciatic nerve sections incubated with anti-Reelin antibodies are Reelin-negative. Scale bars mark 30 Am in

panel A, same magnification in panels B–F; 50 Am in G, same magnification in panels H and I.

R. Panteri et al. / Mol. Cell. Neurosci. 32 (2006) 133–142 137

indicates that the structural organization of the RL mouse nerve is

substantially equivalent to the nerve of the control animals as far as

the myelinated axons and extracellular matrix are concerned (Fig.

6A); the total transverse sectional area of the RL mouse nerve is

not significantly different from that of the control WT animals at

the age examined (data not shown). Nevertheless, the morphomet-

ric analysis indicates that the RL nerves at P15 contain more

myelinated axons (+17%) than the WT control nerves (Fig. 6B).

This is due to the presence of a greater number of nerve fibers of

Fig. 5. Reelin and S100 immunofluorescence in adult mouse sciatic nerves. (A an

S100 (green) on normal sciatic nerve. Reelin and S100 immunoreactivities are asso

images in panels A and B shows co-localization of Reelin and S100. Scale bar m

small diameter (1.5–3 Am) and to a smaller number of nerve fibers

of higher diameter (Fig. 6C). In particular, the RL mice have more

small myelinated nerve fibers than WT mice (Fig. 6C). For both

nerves, the more numerous fiber class is the 3 Am diameter one.

These findings about RL mice are characteristic of developing

immature nerves and suggest that the absence of Reelin in

peripheral nerves could cause a delay in postnatal maturation

events, which are normally characterized by a progressive

reduction of the axon number and an increase of their diameters.

d B) Double immunofluorescence with antibodies against Reelin (red) and

ciated with Schwann cells surrounding intact nerve fibers. (C) Overlay of the

arks 10 Am in panel A, same magnification in panels B and C.

Fig. 6. (A) Transverse semithin sections of wild-type (WT) and reeler (RL) mouse sciatic nerve. The morphological features of the RL sciatic nerve are similar

to those of the WT mouse. In both cases, we observe a high number of small myelinated axons. (B) Distribution of the number of myelinated fibers in WT and

RL sciatic nerves. Values are means T SEM of 6 RL and 7 WT preparations. *Significantly different from WT (P < 0.05). (C) Diameter distribution of

myelinated fibers in WT and RT sciatic nerves. Values are means T SEM of 6 RL and 7 WT preparations. *Significantly different from WT (P < 0.05).

R. Panteri et al. / Mol. Cell. Neurosci. 32 (2006) 133–142138

Reelin absence does not affect the morphology of non-myelinating

Schwann cells

To investigate whether the absence of Reelin affected the

maturation and the morphological features of non-myelinating

Schwann cells, we analyzed the morphology of GFAP-stained non-

myelinating Schwann cell populations derived from WT and RL

mice at postnatal day 21. Our intention was to evaluate whether

Reelin had also an effect on the Schwann cell population

associated with non-myelinated small diameter axons (C-fibers)

(Corfas et al., 2004). The analysis was carried out on two

populations of ¨180 cells selected from teased sciatic nerve fiber

preparations of 3 WT and 3 RL mice and involved the measure of

the area, equivalent diameter, perimeter, length and breadth of the

non-myelinating Schwann cells under investigation. The results,

which are not shown, do not suggest a significant difference in any

of the above morphological parameters between WT and RL mice

and therefore do not implicate Reelin in the development and

maturation of the population of non-myelinating Schwann cells in

the peripheral nerve.

Discussion

The present study provides first evidence that during early

postnatal development Reelin is transiently expressed in the

peripheral nervous system and may play a yet unidentified role in

the cellular processes that underlie peripheral nerve development

and response to injury. The expression of Reelin mRNA in

embryonic dorsal root ganglia and peripheral nerve has been

reported previously (Ikeda and Terashima, 1997; Buchstaller et al.,

2004). Our expression analysis on RNA extracts from young mouse

sciatic nerves reveals the presence of Reelin mRNA at early

postnatal stages; furthermore, it appears that Reelin expression in

the periphery is developmentally regulated since levels of Reelin

isoforms in the sciatic nerve increase progressively from P1 up to

P12–P15 and then decrease again to low levels in adulthood. It is

unlikely that Reelin is axonally transported since recent data show

that spinal motoneurons do not express Reelin (Kubasak et al.,

2004). Reelin appears to be produced in situ in the peripheral nerve

since perfusion of the peripheral nerve tissue with saline did not alter

the levels of Reelin isoforms detected by immunoblot analysis. This,

in addition to the mRNA data, suggests that Reelin isoforms, which

are known to be present in the blood circulation (Smalheiser et al.,

2000), contribute little to the total Reelin extracted from the

peripheral nerves for the biochemical analysis. Such developmen-

tally restricted pattern of expression is common to other proteins

produced and secreted by Schwann cells in the peripheral nerve

compartment, such as the collagen-like heparin-binding adhesive

glycoprotein p200 (Chernousov et al., 1999), which plays an

important role in modulating the proliferation and migration of

premyelinating Schwann cells during the early stages of peripheral

nervous system development, the transmembrane glycoprotein h-dystroglycan and laminin-2 (Masaki et al., 2002), which interact to

form an adhesion apparatus that binds the Schwann cell outer

membrane to the basal lamina during the process of axonal

ensheathment by Schwann cells and the extracellular matrix

glycoprotein Tenascin-R (Probstmeier et al., 2001), which, like

R. Panteri et al. / Mol. Cell. Neurosci. 32 (2006) 133–142 139

Reelin, has been implicated in a variety of cell–matrix interactions

involved in the molecular control of neural cell migration in the

developing central nervous system. As suggested by our studies on

primary cell cultures obtained from rat peripheral nerves, immature

Schwann cells express Reelin mRNA and can also synthesize and

secrete Reelin into the extracellular medium. In accordance with

this, it has been recently reported that Reelin is expressed by glial

cells in the adult human neocortex and the pattern of immunolabel-

ing is confined to the nucleus and structures surrounding the outer

nuclear membrane (Roberts et al., 2005), a pattern of labeling similar

to that observed in our Schwann cells in vitro. In the normal sciatic

nerve, Reelin co-localizes with the Schwann cell marker S100 and is

present along the MBP-positive compact myelin of intact nerve

fibers, whereas in the crushed sciatic nerve Reelin co-localizes with

MBP immunoreactive myelin ovoids, which represent a typical sign

of a nerve undergoing Wallerian degeneration (Stoll and Muller,

1999). From the morphometric analysis and the observation of

semithin sciatic nerve sections of RL mice, it appears however that

the absence of Reelin does not perturb the ability of Schwann cells to

form myelin or to achieve the proper 1:1 promyelinating Schwann

cell–axon relationship, allowing for a normal peripheral nerve

development. However, the absence of Reelin affects the caliber of

myelinated axons in the small myelinated fiber class and also

influences the absolute number of myelinated axons, which is

compatible with a delay in the maturation events that characterize

peripheral nerve development (Rakic and Riley, 1983; Jenq et al.,

1986; Luo and O’Leary, 2005). A more thorough analysis at the

electron microscopic level is however required to discern whether

Reelin is essential for the structural integrity of the Schwann cell–

axon unit. Since themorphometric analysis did not include olderWT

and RLmice, we cannot rule out a possible involvement of Reelin in

the maintenance and stability of normal Schwann cell–axon

interactions, in a similar way as myelin-associated glycoprotein

(MAG) (Yin et al., 1998; Garbay et al., 2000). In this context, it is

interesting to point out that MAG-ko mice do show a normal

myelination of nerve fibers in the first few weeks, however around

P35, and more evidently at 3 months and 9 months of age, MAG-

deficient peripheral nerves show decreased axonal caliber that

correlates with decreased neurofilament spacing leading to a chronic

atrophy of myelinated axons (Yin et al., 1998). The induction of

Reelin expression following crush injury in the mature peripheral

nerves suggests new hypotheses concerning the function of Reelin

outside the central nervous system. Peripheral nerve damage by

crush leads to interruption of axonal integrity with ensuing

degeneration of nerve fibers distal to the site of insult, a process

named Wallerian degeneration (WD) (Waller, 1850; Donat and

Wisniewski, 1973; Stoll and Muller, 1999). WD induces complex

cellular reactions involving both resident and hematogenous cells

aiming at removing degenerating axons and creating a microenvi-

ronment that allows successful regrowth of nerve fibers from the

proximal nerve segment. WD begins with prompt degradation of

axoplasm and axolemma induced by the activation of axonal

proteases and calcium influx (Schlaepfer and Bunge, 1973; George

et al., 1995); Schwann cells respond to loss of axons by extrusion of

their myelin sheaths, downregulation of myelin genes, dedifferen-

tiation and proliferation. They finally align in tubes (Bungner bands)

and express surface molecules that guide regenerating fibers.

Resident endoneurial and hematogenous macrophages are rapidly

recruited to the distal stump and remove the vast majority of myelin

debris (Perry and Brown, 1992; Nishio et al., 2002). This sequence

of morphogenetic events may be inductive to Reelin expression

distally to the lesion site, which we detect as soon as 2 days after

nerve injury. Thus, Reelin may be part of the molecular changes in

the distal stump set in motion byWD which include upregulation of

neurotrophins, neural cell adhesion molecules, cytokines and other

soluble factors and their corresponding receptors (Makwana and

Raivich, 2005). Upon loss of axonal contact, myelinating Schwann

cells downregulate production of the myelin components myelin

basic protein (MBP), myelin associated glycoprotein (MAG),

protein zero (P0), peripheral myelin protein-22 (PMP22) and

periaxin within 2 days after injury. Formerly myelinating Schwann

cells dedifferentiate and acquire the phenotype of pre/non-myelinat-

ing Schwann cells by expression of the low affinity neurotrophin

receptor p75, glial fibrillary acidic protein (GFAP), glial maturation

factor-h, the cell adhesion molecule L1 and neural cell adhesion

molecule (N-CAM) (Jessen et al., 1990; Stoll and Muller, 1999).

Since the induction of Reelin is observed very soon after nerve

injury, it is temporally correlated with the proteins that are induced in

dedifferentiating Schwann cells during the process of WD (Kury et

al., 2001; Jessen and Mirsky, 2002). Given that axonal loss also

activates soon after nerve injury resident endoneurial macrophages

and promotes recruitment of hematogenous macrophages that

phagocytose and remove degenerating myelin in a complement-

dependent manner, we cannot exclude that in our injury model part

of Reelin induction may be contributed by other cells, in addition to

Schwann cells, e.g. activated macrophages, which are known to

secrete an enormous range of products such as factors promoting

neurite outgrowth and elongation (Perry and Brown, 1992; Mueller

et al., 2001). However, our analysis reveals that Reelin is expressed

physiologically in the peripheral nerve during development, thus the

macrophage population should not represent a critical source of

Reelin production in the normal peripheral nerve. The presence of

Reelin in the peripheral nerve may be functionally related to the

cadherin-related neuronal receptor (CNR)/protocadherin (Pcdh) a

family of proteins expressed in developing axons (Morishita et al.,

2004). Reelin signaling through CNR/Pcdha receptors is mediated

by Fyn-tyrosine kinases and stimulates cyclin-dependent kinase

(cdk) 5 activity (Senzaki et al., 1999). Cdk5 in the peripheral nerve

phosphorylates the high- and middle-molecular-weight neurofila-

ments (Shetty et al., 1993; Sun et al., 1996; Terada et al., 1998), a

post-translational modification which controls axon caliber (De

Waegh et al., 1992; Nixon et al., 1994). This pathway would be

consistent with our results showing an alteration in the caliber of

myelinated fibers in RL sciatic nerves. Analysis of cdk5 activity and

of neurofilament phosphorylation state in RL peripheral nerves

could tell us more about the putative role of Reelin in controlling

axon caliber. Furthermore, in vitro antibody perturbation experi-

ments and in vivo analysis of the RL mice will help to unravel the

role of Reelin in processes related to axon regeneration following

nerve injury and perhaps provide insight into whether Reelin-

dependent processes contribute similarly to the assembly of

neuronal connections and synapse formation in the central and

peripheral nervous system.

Experimental methods

Rodent colonies and genotyping

Homozygous reeler mice used in this study were bred from heterozy-

gous B6C3Fe-a/a-rl adults obtained from The Jackson Laboratory. The

colony was maintained by mating heterozygous males to heterozygous

female mutants. Mice were kept on a 14/10 h day/night cycle with food and

R. Panteri et al. / Mol. Cell. Neurosci. 32 (2006) 133–142140

water ad libitum. All mice were treated in accordance with the Italian

Ministry of Health policy on the use of animals in research. The mice were

genotyped by PCR as described (D’Arcangelo et al., 1996) except with

slight modifications. Approximately 1 Ag of DNA was placed in the 10�reaction buffer (sterile ddH2O plus 100 mM Tris–HCl, pH 8.3, 500 mM

KCl, 15 mM MgCl2) containing 10 mM dNTPs, 20 mM oligonucleotide

primers and 5 U/ml recombinant Taq polymerase (Takara). Primers were

synthesized by Invitrogen. The forward primer (GM75) is common to both

wild-type and mutant alleles (sequence 5VTAA. TCT. GTC. CTC. ACT.CTG. CC3V), one reverse primer is specific for the wild-type allele (3W1,

sequence 5VACA. GTT. GAC. ATA. CCT. TAA. TC3V), the other is specificfor the rl allele (3R1, sequence 5VTGT. ATT. AAT. GTG. CAG. TGT. TG3V).DNA amplification was performed in a PTC-100 cycler (MJR), according to

the following program: 1 cycle at 94-C for 5 min, 30 cycles at 94-C for 1

min/55-C for 2 min/72-C for 3 min, 1 cycle at 72-C for 10 min. Reaction

products were separated by electrophoresis on 1.5% agarose gel and

visualized by ethidium bromide staining under UV light. Two DNA

fragments are amplified by PCR, one corresponding to wild-type allele

(280 bp) and the other corresponding to the rl allele (380 bp).

Animals and surgical procedures

Adult wild-type mice were anesthetized by intraperitoneal injection of a

mixture of 2 mg/ml ketamine, 0.2 ml/10 g body weight (Ketavet, Gellini

Farmaceutici, Italy) and 0.23 mg/ml medetomidine, 0.24 ml/10 g body

weight (Domitor, Orion Corp., Espoo, Finland). The right sciatic nerve was

exposed at the mid-thigh level and was crushed for 30 s with a smooth-

bladed hemostat forceps chilled on dry ice. Sham operations of the left

sciatic nerves were made by exposing the nerve without crush. For all

procedures, the wound was sutured in layers and the animals allowed to

recover. At several time points after the operation, the mice were sacrificed

and the sciatic nerves were dissected out, frozen on dry ice and kept at

�80-C until further processing.

Morphometric analysis of sciatic nerve fibers

For the morphometric analysis of sciatic nerves, n = 4 wild-type (WT)

and n = 3 reeler (RL) mice at P15 were used. The animals were anesthetized

as described above and transcardially perfused with saline followed by 4%

paraformaldehyde in 0.1 M phosphate buffer, pH 7.2. The sciatic nerves of

both sides were exposed, a 3-mm-long segment at the mid-thigh level was

dissected and additionally fixed by immersion in 2.5% glutaraldehyde at

4-C for 2 h. The samples were rinsed overnight in 0.1 M phosphate buffer,

postfixed in 1% osmium tetroxide, dehydrated with ethanol and embedded

in Epon 812. Transverse semithin sections (0.75 Am) were cut with glass

knives on a Reichert-Jung 2050 microtome, stained with toluidine blue and

examined with a light microscope. The morphometric analysis of sciatic

nerves was performed by using an Axiophot Zeiss light microscope

equipped with a video camera connected to a Zeiss Axiovision LE image

analyzer. In each nerve, 2 non-overlapping quadrangular areas (100 Am �100 Am) were examined, resulting in a total area of 20,000 Am2, equivalent

to approximately 40% of the total transverse sectional area of the sciatic

nerve under investigation. The analysis was carried out on 7 wild-type and

6 reeler mouse sciatic nerves. Histograms of the number and size

distribution of myelinated axons in nerves were obtained by counting and

measuring the diameter of all myelinated axons present in the selected

areas, as previously described (Pallini et al., 1992). The fiber diameter was

calculated from the imaginary circle corresponding to the fiber area. For

statistical analysis, the data were expressed as means T SEM and tested

using Student’s t test. For the morphometric analysis of non-myelinating

Schwann cells in vivo, teased fiber preparations were prepared from the

sciatic nerves of 3 wild-type and reeler mice at P21 and stained with a

mouse anti-GFAP antibody (1:250, Dako Z0334) followed by a biotinylated

secondary anti-rabbit antibody (1:200, Chemicon) and streptavidin–Alexa

488 (1:200, Molecular Probes), as described elsewhere (Jessen et al., 1990).

Staining was analyzed with a Leitz DMRB fluorescence light microscope

(Leica) equipped with a Hamamatsu Digital CCD Camera, and images of

selected cells were analyzed with Simple PCI image analysis software

(Compix Imaging Systems, Hamamatsu).

Immunostaining of Reelin in sciatic nerves and co-localization with

MBP/S100

For the immunohistochemistry experiments, crush- and sham-lesioned

sciatic nerves were excised from adult wild-type mice (P34) 1 week after the

operation, fixed by immersion in 4% paraformaldehyde in 0.1 M phosphate

buffer, pH 7.2 and embedded in paraffin. A series of 5-Am-thick longitudinal

nerve sections were cut using a microtome and mounted onto lysine-coated

slides. Sections were then deparaffinized and treated for antigen retrieval by

immersion in 0.1 M citrate buffer solution pH 6.0 at 80-C for 30 min. After

cooling down and several washes in PBST (PBS + 0.1% Triton-X), the

sections were incubated with the anti-Reelin mouse monoclonal antibodies

E4 and Ab142 (Chemicon, 1:500) in PBS overnight at 4-C. Then, after

several washes in PBST, sections were incubated with a biotinylated goat

anti-mouse secondary antibody (Chemicon, 1:1000) for 10 min followed by

streptavidin–Alexa 488 (Molecular Probes, 1 Ag/ml) for 10 min. The

sections were then incubated with a polyclonal rabbit anti-MBP antibody

(Chemicon, 1:1000) for 1 h followed by a goat anti-rabbit IgG–Alexa 594

antibody (Molecular Probes, 1 Ag/ml) for 30 min. Then, after several washes

in PBST, the nerve sections were mounted with an antifade solution

(SlowFade Light Kit, Molecular Probes). Staining was analyzed with a Leitz

DMRB light microscope (Leica) equipped with a Hamamatsu Digital CCD

Camera, and images were processed with Simple PCI image analysis

software (Compix Imaging Systems, Hamamatsu). For the control staining

procedures, the sections were either incubated with just the anti-MBP or the

anti-Reelin primary antibody followed by incubation with the two secondary

goat anti-rabbit/anti-mouse antibodies and analyzed under identical con-

ditions. For S100 immunohistochemistry, intact sciatic nerves were excised

from adult wild-type mice and fixed by immersion in 4% paraformaldehyde

in 0.1 M phosphate buffer, pH 7.2 for 1 h. After several washes, nerves were

treated with a solution of 30% sucrose for 8 h. Nerves were permeabilized

overnight with a solution containing 4% bovine serum albumin (BSA), 2%

normal goat serum and 0.5% Triton-X100 in phosphate buffer. After that,

nerves were incubated 48 h with primary antibodies: E4 (IgG1, Chemicon,

1:400) to detect Reelin and anti-S100 (rabbit) (DakoCytomation, 1:1000) to

detect Schwann cells. After several washes, nerves were incubated for 48

h with secondary antibodies anti-IgG1 conjugated with Alexa 568 and anti-

rabbit conjugated with Alexa 488 (both from Molecular Probes). After

several washes, nerves were cleaned under a dissection microscope and

mounted on slides with a solution of glycerol and paraphenylendiamine.

Acquisition was made with Olympus fluorescence microscope (Milan,

Olympus-Italy) and a Q-Imaging camera (Burnaby, Canada). Control

staining procedures were carried out without primary antibodies and

showed negative staining (not shown).

Immunostaining of cultured Schwann cells

For immunocytochemistry on Schwann cells in vitro, cells grown on

polylysin-coated cover slips were washed with PBS, fixed for 15 min in 4%

PFA, washed and stained using mouse E4 anti-Reelin (1:1000, Chemicon

MAB 5364) or rabbit anti-S100 antibodies followed by the appropriate

secondary antibody (goat anti-mouse or goat anti-rabbit Alexa Fluor 594,

1:500, Molecular Probes). When necessary, the cell nuclei were counter-

stained with DAPI (4V,6-diamidino-2-phenylindole dihydrochloride; Sigma

D9564). Staining was analyzed with Axiophot epifluorescence microscope

(Zeiss), 40� objective, digital camera and Axiovision image analysis

system. Control staining procedures without primary antibody were carried

out and analyzed under identical conditions.

Gel electrophoresis and Western blotting

Protein extracts from mouse sciatic nerves were prepared by homog-

enizing the tissue in 100 Al of ice-cold lysis buffer (1% SDS, 50 mM Tris

pH 7.5, Leupeptin 0.1 mg/ml, Aprotinin 0.1 mg/ml, PMSF 0.2 mg/ml, 100

R. Panteri et al. / Mol. Cell. Neurosci. 32 (2006) 133–142 141

mM h-Mercaptoethanol). Extracts were cleared by centrifugation at

44,000�g for 20 min. To control for equal protein loading, protein

concentrations in the supernatants were measured using the Bradford

reagent (Sigma), using BSA as a standard; absorbance readings were

performed with a Tecan GENIOS microplate reader. Samples were

separated with 5% SDS-PAGE and blotted on nitrocellulose membranes

using a wet transfer apparatus (Hoefer TE22 tank transfer unit, Hoefer

Scientific) at 250 mA for 2 h. The blots were blocked for 1 h in Tris-

buffered saline with 0.1% Tween 20 (TBST), containing 5% nonfat dry

milk (Amersham) and then incubated with the anti-Reelin mouse

monoclonal mAb 142 (Chemicon; 1:1000) overnight at 4-C followed by

1:10,000 HRP-tagged anti-mouse antibody for 1 h at RT. Immunoblots

were developed using an enhanced chemiluminescence kit (ECL, Amer-

sham). For immunoblot experiments on Schwann cells in vitro, Schwann

cell culture supernatants were separated with discontinuous SDS-PAGE (T

4–15% gradient gels, Biorad) and transferred by semidry blotting. After

transfer, nitrocellulose membranes were incubated with 1:2000 mAb E4

(Chemicon) followed by incubation with 1:5000 HRP–secondary goat anti-

mouse (Sigma) and development with ECL. Films were developed and

images acquired with a Kodak EDAS 290 imaging system. Densitometric

analysis was performed on the Reelin bands identified on the image with

the Kodak 1D Image Analysis Software package. The net intensity values

were registered for each band, pooled together and plotted against age and

days post-lesion. The fitting of curves was performed by cubic interpolation

of the data using the SPSS statistics package (SPSS Inc.).

RNA extraction and RT-PCR analysis

Reelin expression was also examined by RT-PCR analysis in sciatic

nerves and in Schwann cell cultures. Total RNA was extracted from the

sciatic nerves of young wild-type, heterozygous and reeler mice. One

microgram of total RNA was incubated in a reaction buffer containing 0.5

Ag random primers, AMV reverse transcriptase, reaction buffer and RNase

inhibitor, according to the manufacturer’s specifications (Promega).

Expression of Reelin was examined by PCR amplification with specific

primers: Reelin fwd 5V-GAGGTGTATGCAGTG-3V, Reelin rev 5V-TCTCA-CAGTGGATCC-3V corresponding to nucleotides 8500–9090 of mouse

Reelin mRNA (accession number U24703) and yielding a PCR product of

591 bp. The cycling conditions were: initial denaturation at 95-C for 3 min

followed by 30 cycles of 95-C for 30 s, 55-C for 30 s, 72-C for 1 min, with

a final extension at 72-C for 5 min. For RT-PCR analysis of Reelin in rat

Schwann cell primary cultures, samples of 500 ng total RNA were treated

with DNase and reverse transcribed with oligo-dT primers and M-MLV

reverse transcriptase (Life Technologies) according to the manufacturer’s

instructions. Expression of Reelin was monitored by PCR amplification

with specific primers: Reelin fwd 5V-AAACCTCAGCTTCGTCTGGA-3V,Reelin rev 5V-ACGTTGGAAGGGGCTCTAAT-3V corresponding to nucleo-

tides 423–923 of rat Reelin mRNA (accession number NM_080394) andyielding a PCR product of 501 bp. The cycling conditions were: first

denaturation at 94-C for 3 min, 28 cycles of annealing at 51-C for 1 min,

extension at 72-C for 1 min and 30 s, denaturation at 94-C for 1 min,

followed by 1 min annealing and last elongation at 72-C for 10 min. All

PCR products were separated by 1% agarose gel electrophoresis and stained

with ethidium bromide to verify their expected sizes. Control PCR reactions

were performed without prior RT.

Cell cultures

Primary cultures of Schwann cells were prepared from newborn

Sprague–Dawley rats. In one preparation, 15–20 sciatic nerves were

dissected, freed from blood vessels and fat, cut up in small pieces and

digested for 1 h at 37-C in 10 ml DMEM containing 0.6% collagenase and

2.5% trypsin. To obtain single cells, the nerve pieces were gently triturated

(first 0.7 mm, then 0.4 mm gauge cannulae). The cell suspension was plated

out in uncoated culture flasks (25 cm2) with 5 ml Dulbecco’s Modified

Eagle’s Medium (DMEM) containing 10% fetal calf serum (FCS).

Fibroblast growth was reduced by the addition of 10 AM cytosine

arabinoside (Ara C) to the medium for 4 days. To eliminate remaining

fibroblasts, the cells were incubated 30 min with Thy 1.1 antibody (Sigma)

at 37-C followed by treatment with baby rabbit complement (Linaris) for

another 30 min. Schwann cells were then plated out in a poly-l-lysine-

coated culture flask (25 cm2) with 5 ml DMEM containing 10% FCS, 2 AMforskolin (ICN) and 100 Ag/ml bovine pituitary extract (Life Technologies).

Cultures were kept at 37-C in 5% CO2. After the cells had reached

confluence, complement lysis was repeated and cells were cultured in poly-

l-lysine-coated flasks (75 cm2) with 10 ml DMEM containing 10% FCS

and 2 AM forskolin. The identity of cultured Schwann cells was confirmed

with immunocytochemistry against S100. For the analysis of supernatants,

cells were cultivated in serum-free DMEM with B27 supplement (without

vitamin A) for 6 days. After this period, samples of the supernatant were

collected for biochemical analysis.

Acknowledgments

This work was supported by grants from Consiglio Nazionale

delle Ricerche (Programma ‘‘Biomolecole per la salute umana’’,

grant 01.00127.PF33), a grant from the Ministero dell’Universita e

della Ricerca (FIRB-grant RBNE01H3K5), the COFIN grant

2003057332 and Associazione Amici del Campus Bio-medico.

The authors wish to thank G. D’Arcangelo for the gift of CER

supernatant and F. Michetti for the gift of the GFAP antibody and

for helpful discussion of the manuscript.

References

Borrell, V., Del Rio, J.A., Alcantara, S., Derer, M., Martinez, A.,

D’Arcangelo, G., Nakajima, K., Mikoshiba, K., Derer, P., Curran, T.,

Soriano, E., 1999. Reelin regulates the development and synaptogenesis

of the layer-specific entorhino-hippocampal connections. J. Neurosci.

19, 1345–1358.

Buchstaller, J., Sommer, L., Bodmer, M., Hoffmann, R., Suter, U.,

Mantei, N., 2004. Efficient isolation and gene expression profiling of

small numbers of neural crest stem cells and developing Schwann

cells. J. Neurosci. 24, 2357–2365.

Chernousov, M.A., Scherer, S.S., Stahl, R.C., Carey, D.J., 1999. p200, a

collagen secreted by Schwann cells, is expressed in developing nerves

and in adult nerves following axotomy. J. Neurosci. Res. 56, 284–294.

Corfas, G., Velardez, M.O., Ko, C.P., Ratner, N., Peles, E., 2004.

Mechanisms and roles of axon–Schwann cell interactions. J. Neurosci.

24, 9250–9260.

D’Arcangelo, G., Miao, G.G., Chen, S.C., Soares, H.D., Morgan, J.I.,

Curran, T., 1995. A protein related to extracellular matrix proteins

deleted in the mouse mutant reeler. Nature 374, 719–723.

D’Arcangelo, G., Miao, G.G., Curran, T., 1996. Detection of the reelin

breakpoint in reeler mice. Brain Res. Mol. Brain Res. 39, 234–236.

D’Arcangelo, G., Homayouni, R., Keshvara, L., Rice, D.S., Sheldon, M.,

Curran, T., 1999. Reelin is a ligand for lipoprotein receptors. Neuron 24,

471–479.

De Bergeyck, V., Naerhuyzen, B., Goffinet, A.M., Lambert de Rouvroit, C.,

1998. A panel of monoclonal antibodies against reelin, the extracellular

matrix protein defective in reeler mutant mice. J. Neurosci. Methods 82,

17–24.

De Waegh, S.M., Lee, V.M., Brady, S.T., 1992. Local modulation of

neurofilament phosphorylation, axonal caliber, and slow axonal

transport by myelinating Schwann cells. Cell 68, 451–463.

Del Rio, J.A., Heimrich, B., Borrell, V., Forster, E., Drakew, A., Alcantara,

S., Nakajima, K., Miyata, T., Ogawa, M., Mikoshiba, K., Derer, P.,

Frotscher, M., Soriano, E., 1997. A role for Cajal–Retzius cells and

reelin in the development of hippocampal connections. Nature 385,

70–74.

R. Panteri et al. / Mol. Cell. Neurosci. 32 (2006) 133–142142

Donat, J.R., Wisniewski, H.M., 1973. The spatio-temporal pattern of

Wallerian degeneration in mammalian peripheral nerves. Brain Res. 53,

41–53.

Garbay, B., Heape, A.M., Sargueil, F., Cassagne, C., 2000. Myelin

synthesis in the peripheral nervous system. Prog. Neurobiol. 61,

267–304.

George, E.B., Glass, J.D., Griffin, J.W., 1995. Axotomy-induced axonal

degeneration is mediated by calcium influx through ion-specific

channels. J. Neurosci. 15, 6445–6452.

Hiesberger, T., Trommsdorff, M., Howell, B.W., Goffinet, A., Mumby,

M.C., Cooper, J.A., Herz, J., 1999. Direct binding of reelin to

VLDL receptor and ApoE receptor 2 induces tyrosine phosphory-

lation of disabled-1 and modulates tau phosphorylation. Neuron 24,

481–489.

Ikeda, Y., Terashima, T., 1997. Expression of reelin, the gene responsible

for the reeler mutation, in embryonic development and adulthood in the

mouse. Dev. Dyn. 210, 157–172.

Jenq, C.B., Chung, K., Coggeshall, R.E., 1986. Postnatal loss of axons in

normal rat sciatic nerve. J. Comp. Neurol. 244, 445–450.

Jessen, K.R., Mirsky, R., 1999. Schwann cells and their precursors

emerge as major regulators of nerve development. Trends Neurosci.

22, 402–410.

Jessen, K.R., Mirsky, R., 2002. Signals that determine Schwann cell

identity. J. Anat. 200, 367–376.

Jessen, K.R., Morgan, L., Stewart, H.J., Mirsky, R., 1990. Three markers of

adult non-myelin-forming Schwann cells, 217c(Ran-1), A5E3 and

GFAP: development and regulation by neuron–Schwann cell inter-

actions. Development 109, 91–103.

Kubasak, M.D., Brooks, R., Chen, S., Villeda, S.A., Phelps, P.E.,

2004. Developmental distribution of reelin-positive cells and their

secreted product in the rodent spinal cord. J. Comp. Neurol. 468,

165–178.

Kubo, K., Mikoshiba, K., Nakajima, K., 2002. Secreted Reelin molecules

form homodimers. Neurosci. Res. 43, 381–388.

Kury, P., Stoll, G., Muller, H.W., 2001. Molecular mechanism of cellular

interactions in peripheral nerve regeneration. Curr. Opin. Neurol. 14,

635–639.

Luo, L., O’Leary, D.D., 2005. Axon retraction and degeneration in

development and disease. Annu. Rev. Neurosci. 28, 127–156.

Makwana, M., Raivich, G., 2005. Molecular mechanisms in successful

peripheral regeneration. FEBS J. 272, 2628–2638.

Masaki, T., Matsumura, K., Hirata, A., Yamada, H., Hase, A., Arai,

K., Shimizu, T., Yorifuji, H., Motoyoshi, K., Kamakura, K.,

2002. Expression of dystroglycan and the laminin-alpha 2 chain

in the rat peripheral nerve during development. Exp. Neurol. 174,

109–117.

Morishita, H., Kawaguchi, M., Murata, Y., Seiwa, C., Hamada, S., Asou,

H., Yagi, T., 2004. Myelination triggers local loss of axonal

CNR/protocadherin alpha family protein expression. Eur. J. Neurosci.

20, 2843–2847.

Mueller, M., Wacker, K., Ringelstein, E.B., Hickey, W.F., Imai, Y., Kiefer,

R., 2001. Rapid response of identified resident endoneurial macro-

phages to nerve injury. Am. J. Pathol. 159, 2187–2197.

Nishio, Y., Nishihira, J., Ishibashi, T., Kato, H., Minami, A., 2002.

Role of macrophage migration inhibitory factor (MIF) in peripheral

nerve regeneration: anti-MIF antibody induces delay of nerve

regeneration and the apoptosis of Schwann cells. Mol. Med. 8,

509–520.

Niu, S., Renfro, A., Quattrocchi, C.C., Sheldon, M., D’Arcangelo, G.,

2004. Reelin promotes hippocampal dendrite development through the

VLDLR/ApoER2-Dab1 pathway. Neuron 41, 71–84.

Nixon, R.A., Paskevich, P.A., Sihag, R.K., Thayer, C.Y., 1994. Phosphor-

ylation on carboxyl terminus domains of neurofilament proteins in

retinal ganglion cell neurons in vivo: influences on regional neurofila-

ment accumulation, interneurofilament spacing, and axon caliber. J. Cell

Biol. 126, 1031–1046.

Pallini, R., Fernandez, E., Lauretti, L., Draicchio, F., Pettorossi, V.E.,

Gangitano, C., Del Fa, A., Olivieri-Sangiacomo, C., Sbriccoli, A., 1992.

Experimental repair of the oculomotor nerve: the anatomical paradigms

of functional regeneration. J. Neurosurg. 77, 768–777.

Perry, V.H., Brown, M.C., 1992. Role of macrophages in peripheral nerve

degeneration and repair. Bioessays 14, 401–406.

Probstmeier, R., Nellen, J., Gloor, S., Wernig, A., Pesheva, P., 2001.

Tenascin-R is expressed by Schwann cells in the peripheral nervous

system. J. Neurosci. Res. 64, 70–78.

Rakic, P., Riley, K.P., 1983. Overproduction and elimination of retinal

axons in the fetal rhesus monkey. Science 219, 1441–1444.

Rice, D.S., Curran, T., 2001. Role of the Reelin signalling pathway in

central nervous system development. Annu. Rev. Neurosci. 24,

1005–1039.

Roberts, R.C., Xu, L., Roche, J.K., Kirkpatrick, B., 2005. Ultrastructural

localization of reelin in the cortex in post-mortem human brain. J. Comp.

Neurol. 482, 294–308.

Sanes, J.R., Lichtman, J.W., 1999. Development of the vertebrate

neuromuscular junction. Annu. Rev. Neurosci. 22, 389–442.

Schlaepfer, W.W., Bunge, R.P., 1973. Effects of calcium ion concentration

on the degeneration of amputated axons in tissue culture. J. Cell Biol.

59, 456–470.

Senzaki, K., Ogawa, M., Yagi, T., 1999. Proteins of the CNR family are

multiple receptors for Reelin. Cell 99, 635–647.

Shetty, K.T., Link, W.T., Pant, H.C., 1993. cdc2-like kinase from rat spinal

cord specifically phosphorylates KSPXK motifs in neurofilament

proteins: isolation and characterization. Proc. Natl. Acad. Sci. U. S. A.

90, 6844–6848.

Smalheiser, N.R., Costa, E., Guidotti, A., Impagnatiello, F., Auta, J.,

Lacor, P., Kriho, V., Pappas, G.D., 2000. Expression of reelin in

adult mammalian blood, liver, pituitary pars intermedia, and

adrenal chromaffin cells. Proc. Natl. Acad. Sci. U. S. A. 97,

1281–1286.

Stoll, G., Muller, H.W., 1999. Nerve injury, axonal degeneration and neural

regeneration: basic insights. Brain Pathol. 9, 313–325.

Sun, D., Leung, C.L., Liem, R.K., 1996. Phosphorylation of the high

molecular weight neurofilament protein (NF-H) by Cdk5 and p35.

J. Biol. Chem. 271, 14245–14251.

Terada, M., Yasuda, H., Kogawa, S., Maeda, K., Haneda, M., Hidaka, H.,

Kashiwagi, A., Kikkawa, R., 1998. Expression and activity of cyclin-

dependent kinase 5/p35 in adult rat peripheral nervous system.

J. Neurochem. 71, 2600–2606.

Tissir, F., Goffinet, A.M., 2003. Reelin and brain development. Nat. Rev.

Neurosci. 4, 496–505.

Waller, A.V., 1850. Experiments on the section of the glossopharyngeal and

hypoglossal nerves of the frog, and observations on the alterations

produced thereby in the structure of their primitive fibres. Philos. Trans.

R. Soc. Lond. 140, 423–429.

Yin, X., Crawford, T.O., Griffin, J.W., Tu, P., Lee, V.M., Li, C., Roder, J.,

Trapp, B.D., 1998. Myelin-associated glycoprotein is a myelin signal

that modulates the caliber of myelinated axons. J. Neurosci. 18,

1953–1962.