Experience-Dependent Transfer of Otx2 Homeoprotein into the Visual Cortex Activates Postnatal...

Experience-Dependent Transfer ofOtx2 Homeoprotein into the VisualCortex Activates Postnatal PlasticitySayaka Sugiyama,1,6 Ariel A. Di Nardo,2 Shinichi Aizawa,3 Isao Matsuo,4 Michel Volovitch,2 Alain Prochiantz,2,5,*and Takao K. Hensch1,6,7,*1Lab for Neuronal Circuit Development, RIKEN Brain Science Institute, 2-1 Hirosawa, Wako-shi, Saitama 351-0198, Japan2CNRS UMR 8542, Ecole Normale Superieure, 46 rue d’Ulm, 75230 Paris Cedex 05, France3RIKEN Center for Developmental Biology, 2-2-3 Minatojima-minamimachi, Chuo-ku, Kobe 650-0047, Japan4Department of Molecular Embryology, Osaka Medical Center and Research Institute for Maternal and Child Health, 840 Murodo-cho, Izumi,

Osaka 594-1101, Japan5College de France, Paris, France6Department of Neurology, Children’s Hospital Boston, Harvard Medical School, 300 Longwood Avenue Boston, MA 02115, USA7Department of Molecular & Cellular Biology, Harvard University, 7 Divinity Avenue, Cambridge, MA 02138, USA

*Correspondence: [email protected] (T.K.H.), [email protected] (A.P.)

DOI 10.1016/j.cell.2008.05.054

SUMMARY

Neural circuits are shaped by experience in earlypostnatal life. Distinct GABAergic connections withinvisual cortex determine the timing of the criticalperiod for rewiring ocular dominance to establishvisual acuity. We find that maturation of the parvalbu-min (PV)-cell network that controls plasticity onset isregulated by a selective re-expression of the embry-onic Otx2 homeoprotein. Visual experience pro-moted the accumulation of non-cell-autonomousOtx2 in PV-cells, and cortical infusion of exogenousOtx2 accelerated both PV-cell development and crit-ical period timing. Conversely, conditional removal ofOtx2 from non-PV cells or from the visual pathwayabolished plasticity. Thus, the experience-depen-dent transfer of a homeoprotein may establish thephysiological milieu for postnatal plasticity of a neuralcircuit.

INTRODUCTION

Neural circuits are shaped by experience in postnatal life. For ex-

ample, inputs from the two eyes first converge in the primary vi-

sual cortex (Wiesel and Hubel, 1963), where competitive interac-

tions determine which eye will eventually dominate both

functionally and anatomically. From mouse to man, early monoc-

ular occlusion permanently reduces visual acuity (amblyopia)

through the deprived eye (Daw, 1995; Prusky and Douglas,

2003). Unbalanced sensory experience leads to a rapid shift of

neuronal spiking response in favor of the open eye (ocular dom-

inance) concomitant with a rapid pruning of dendritic spines and

later rewiring of thalamo-cortical afferents (Wiesel and Hubel,

1963; Antonini et al., 1999; Hensch, 2005).

508 Cell 134, 508–520, August 8, 2008 ª2008 Elsevier Inc.

Recently, the maturation of particular inhibitory connections

within the neocortex has been shown to drive this sensitivity

(Hensch, 2005). Mice lacking the synaptic isoform of GABA-syn-

thetic enzyme, glutamic acid decarboxylase (GAD65), fail to shift

visual responsiveness (Hensch et al., 1998) or to lose dendritic

spines (Mataga et al., 2004) upon brief monocular deprivation

(MD). Both can be rescued by concurrent benzodiazepine treat-

ment (to enhance GABAergic transmission). Raising animals in

complete darkness from birth (dark-rearing) naturally delays the

maturation of intracortical inhibition and ocular dominance plas-

ticity (Mower, 1991; Morales et al., 2002; Iwai et al., 2003; Katagiri

et al., 2007). Conversely, benzodiazepines (BDZ) induce a prema-

ture plasticity in young wild-type mice just after eye-opening

(postnatal day, P13) (Fagiolini and Hensch, 2000; Fagiolini

et al., 2004).

Inhibition triggers plasticity through GABAA receptors con-

taining the a1-subunit (Fagiolini et al., 2004; Katagiri et al.,

2007). These are enriched at somatic synapses on pyramidal

cells made by large-basket cells containing the calcium-bind-

ing protein parvalbumin (PV) (Klausberger et al., 2002). Unlike

other GABAergic interneurons, PV-cells emerge with a late

postnatal timecourse in anticipation of critical period onset

(del Rio et al., 1994). Overexpression of brain-derived neurotro-

phic factor (BDNF) accelerates both PV-cell development and

critical period timecourse (Huang et al., 1999; Hanover et al.,

1999).

Gradually with age, peri-neuronal nets (PNNs) of extracellular

matrix molecules condense around PV-cells in parallel with the

decline of critical period plasticity (Celio et al., 1998; Pizzorusso

et al., 2002). Importantly, chondroitinase treatment to destroy

these nets reactivates visual cortical plasticity in adult animals

(Pizzorusso et al., 2002). Moreover, the wide basket-cell axon

plexus stretches horizontally across ocular dominance columns

in the cat and is ideally situated to shape their final spacing

through competitive mechanisms in development (Buzas et al.,

2001; Hensch and Stryker, 2004).

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Figure 1. Otx2 Homeoprotein in Parvalbumin (PV)-Cells of Postnatal Mouse Visual Cortex

(A–E) Distribution of Otx2 (red) and PV (green) protein in supragranular layers of the binocular region of mouse visual cortex before (pre-CP, P18-19) and during the

critical period (CP, P28-30) and into adulthood (Ad, > P50).

(F–J) Colocalization of Otx2 and GAD67- (F), calretinin CR- (G) or PV-positive cells (H and I) at high power in supragranular layers of the binocular zone. The scale

bars represent 50 mm. (J) Quantification of GAD67+/Otx2+, PV+/Otx2+, Otx2+/PV+ and CR+/Otx2+ co-localization (245, 455, 413 and 263 cells, respectively;

600x350 mm area; 6 slices for GAD67+ or CR+, 12 slices for PV+/Otx2+ from 3 mice). Error bars represent SEM.

Due to these convergent lines of evidence, we explored tran-

scription factors as a major determinant of cellular identity that

might specify PV-cell function in particular as a trigger for plasticity

in visual cortex (Butt et al., 2007). Transcription factors have further

been shown to mark distinct neuronal pathways, such as Phox2a/

2b in the visceral nervous circuit (Brunet and Pattyn, 2002). We fo-

cused on the homeodomain protein Otx2, which is normally re-

quired forembryonic head formation (Acamporaetal., 2001).Strik-

ingly, the protein is restricted to relay centers along the primary

visual pathway at birth before PV-cells are evident (Nothias et al.,

1998), including retina, lateral geniculate nucleus (LGN) and cortex

(V1). This is intriguing given the possibility that homeoproteins may

be secreted and internalized (Joliot and Prochiantz, 2004).

Indeed, PV-cells receive the most potent direct thalamo-corti-

cal input (Erisir and Dreusicke, 2005; Cruikshank et al., 2007). Al-

though the expression of Otx2 in regions of the dorsal forebrain in

rats is transient and lost by P13 (Frantz et al., 1994), later expres-

sion with respect to the critical period for circuit plasticity (>P23)

has not been examined (Fagiolini et al., 1994; Gordon and Stryker,

1996). We find that non-cell-autonomous Otx2 coordinates post-

natal PV-cell maturation and activates visual cortical plasticity.

RESULTS

Postnatal Otx2 Regulates PV-Cell MaturationTo address a postnatal role for homeoprotein transcription fac-

tors, we first identified the distribution of Otx2 protein in mouse

visual cortex. Consistent with earlier reports in the rat (Frantz

et al., 1994), little Otx2 protein was detected even at P19 when

weak PV staining was visible prior to critical period onset (Fig-

ures 1A and 1B). Otx2 protein was increasingly concentrated

within PV-cells by the peak of the critical period (P28-30) as PV

expression intensified (Figures 1C and 1D), and persisted into

adulthood (Figure 1E). Similar results were obtained with 6 anti-

bodies recognizing different epitopes of Otx2. Otx2-positive

cells were primarily GABAergic interneurons (Figures 1F and

Cell 134, 508–520, August 8, 2008 ª2008 Elsevier Inc. 509

Figure 2. Experience-Dependent Otx2 Regulation and PV-Cell Maturation in Visual Cortex

(A–D) Decreased expression of both Otx2 (A,B) and PV (C,D) at P24 after binocular enucleation at eye-opening.

(E–J) Decreased expression of Otx2 (E), PV (G), and WFA-stained perineuronal nets (I) by dark-rearing to P30 is rescued in the opposite hemisphere (H and J) by

direct Otx2 infusion (F) in the dark. The scale bars represent 100 mm.

(K) Decreased number of Otx2-positive cells at critical period in supragranular layers of the binocular zone after binocular enucleation or dark-rearing (DR) from

birth (600x350 mm area; three intact, four enucleated mice at P24, p < 0.05, t test; four light-reared (LR), seven DR mice at P28-30, p < 0.0001, t test).

(L) Number of PV-cells or perineuronal net-enwrapped cells is increased by Otx2 infusion in the dark (control versus infused side; 600 3 350 mm area; seven mice

at P28-30, p < 0.005, paired t test). Error bars represent SEM.

1J; 93.1 ± 1.5%), a minority of which contained Calretinin (CR)

(Figures 1G and 1J; 16.6 ± 1.0%). At high resolution, most

Otx2-positive cells were PV-positive and vice versa (Figure 1J;

71.4 ± 0.5% and 78.9 ± 1.7%, respectively) exhibiting nuclear lo-

calization (Figures 1H and 1I).

We examined the influence of visual experience by counting

Otx2-positive cells at the natural critical period peak after two

types of sensory deprivation: removal of the two eyes (enucleation)

from eye-opening (P14) or dark-rearing from birth. Cortical Otx2

protein levels were reduced by P24 in binocularly-enucleated

mice compared to those of intact animals (Figures 2A and 2B).

This was matched by a weakened PV expression (Figures 2C

and 2D). A robust loss of Otx2-positive cells (and reduced PV sig-

nal)was similarly observedatP28 simplybydark-rearing from birth

(Figures 2E, 2G, and 2K; number of Otx2-positive cells/field: intact,

39.7 ± 4.7 versus enucleated, 14.0 ± 5.6, p < 0.05, t test; light-

reared, 35.3 ± 3.5 versus dark-reared, 6.1 ± 1.8, p < 0.0001,

t test). Consistent with previous reports (Tropea et al., 2006; Kata-

giri et al., 2007), dark-rearing thus delayed maturation of PV-posi-

tive inhibitory circuits.

We therefore tested whether exogenous Otx2 would rescue

PV-cell maturation in complete darkness by delivering human

recombinant protein directly into the visual cortex. Laterally-di-

rected flow from osmotic minipumps to treat one cortical hemi-


sphere allowed the other side to serve as an internal control

(Figure S1A available online). After 7 days of infusion in the

dark (P21-23 to P28-30), Otx2 signal was found within PV-cells

throughout the treated visual cortex (Figures 2F and 2H). Inter-

nalization of exogenous Otx2 into cortical neurons is consistent

with previous observations of homeoprotein passage into cells

in vitro (Joliot and Prochiantz, 2004).

Exogenous Otx2 increased the number of PV-positive cells in

the treated side compared to that of the noninfused side in the

same brain (Figures 2H and 2L; number of PV-cells/field: nonin-

fused, 21.7 ± 3.1 versus infused, 33.6 ± 3.2 cells/field, p < 0.005,

paired t test). Enrichment of PNNs enwrapping mature PV-cells

is also delayed by dark-rearing (Pizzorusso et al., 2002). Otx2 in-

fusion increased the number of PNN-bearing cells compared to

that of the noninfused side (Figures 2I, 2J, and 2L; number of net-

bearing cells/field: noninfused, 33.9 ± 1.3 versus infused, 46.1 ±

2.1, p < 0.0005, paired t test).

Thus, endogenous Otx2 may influence late PV-cell maturation

by gradually re-emerging in response to sensory experience after

eye-opening. We verified this idea by exogenous Otx2 infusion

into immature visual cortex. After 5 days of infusion at P19

when endogenous Otx2 levels are low (Figure 1A), robust Otx2

signal was found within PV-cells throughout the treated visual

cortex (Figure 3F). The premature localization of Otx2 accelerated

Figure 3. Acceleration of PV-Cell Maturation by Early Otx2 Infusion

(A–D) Expression of PV within cell somata (D) is intensified at P24 by 5 days of Otx2 infusion (B) compared to the opposite hemisphere (A and C).

(E) Quantitative analysis of PV intensity within somata (open symbols, control side; filled symbols, Otx2-infused) in supragranular layers of the binocular zone (600

3 350 mm area; K-S test, p < 0.001; inset, p < 0.05, paired t test for average intensities).

(F) Exogenous Otx2 protein (B, red) is found specifically in PV-cells (D, green) when images are merged. The scale bar represents 100 mm.

PV expression in the treated side compared to that of the nonin-

fused side in the same brain (Figures 3A–3D). The effects of

Otx2 gain-of-function were PV-cell specific, as Calretinin expres-

sion was not affected by Otx2 infusion (Figure S1C).

Quantification of signal intensity revealed a robust and statisti-

cally significant enhancement across multiple PV-cell markers:

the Kv3.1 class of potassium channel that contributes to fast-spik-

ing behavior (Rudy and McBain, 2001), GAD65 that synthesizes

GABA, and the a1-subunit of the GABAA receptor (Gao and Frit-

schy,1994). Immunofluorescence forPV or Kv3.1withinPV-cell so-

mata was intensified (Figures 3E and S1D; p < 0.05 pair-wise com-

parison across hemispheres of the same brain). Diffuse labeling of

GAD65 and GABAAR a1-subunits was also increased (Figures

S1E–S1G; p < 0.05 pair-wise comparison across hemispheres of

the same brain). No change in cell number indicated an enhance-

ment of pre-existing but immature PV-cells by exogenous Otx2

(Figure S1B; noninfused versus infused cells/field: 39.6 ± 3.3 and

36.8 ± 5.1 for PV; 40.0 ± 2.8 and 43.0 ± 1.6 for Kv3.1; 28.4 ± 4.7

and 29.2 ± 4.5 for Calretinin, respectively; p > 0.3, t test).

Postnatal Otx2 Activates Cortical PlasticityExogenous Otx2 thus promoted the maturation of a constellation

of PV-cell molecular constituents. Pharmacological enhance-

ment of GABAergic function by benzodiazepine agonists is suf-

ficient to trigger plasticity onset (Hensch et al., 1998; Iwai et al.,

2003; Hensch, 2005). We, therefore, examined whether the pre-

mature appearance of Otx2 in the visual cortex could also accel-

erate physiological ocular dominance plasticity using electro-

physiological techniques. The spiking response to visual

stimuli was measured in the binocular zone of anesthetized

mice by extracellular single-unit recording and classified accord-

ing to a traditional scale from group 1 (purely contralateral eye re-

sponse) to group 7 (purely ipsilateral eye response) (Wiesel and

Hubel, 1963; Gordon and Stryker, 1996; Hensch et al., 1998).

Unlike the contralaterally-biased distribution of normal mice, oc-

ular dominance scores typically shift toward the open, ipsilateral

eye following a brief monocular deprivation (MD) of the contralat-

eral eye at P25. This is reflected in a reduction of contralateral

bias index (CBI) from 0.7 to a more balanced value near 0.5

(Gordon and Stryker, 1996; Hensch et al., 1998).

Recombinant Otx2 protein or vehicle solution was infused

early into the visual cortex from P17. Upon vehicle treatment, lit-

tle plasticity occurred with brief MD just after eye-opening, and

visual responses from the closed, contralateral eye remained

strong (Figure 4A, top) as in nondeprived mice (Figure 4B). In

contrast, during early Otx2 infusion, responsiveness shifted pre-

maturely toward the open, ipsilateral eye after brief MD

(Figure 4A, bottom). Moreover, the decline of critical period plas-

ticity with age was also accelerated. At the natural peak of the

critical period, vehicle-treated animals shifted responsiveness

as expected toward the open, ipsilateral eye following brief MD

(from P25), but mice treated earlier (at P17) with Otx2 no longer

showed this plasticity (Figure 4C). The earlier profile of amblyopic

effects was clearly seen in a scatter plot of CBI values from indi-

vidual animals (Figure 4D). Overexpression of BDNF (Hanover

et al., 1999) or early injection of benzodiazepines (Iwai et al.,

2003) also triggers a rapid decline of critical period plasticity

consistent with these Otx2 results.

A well-documented measure of weak inhibition in vivo is the

excessive spike firing by single cells after a visual stimulus has


Figure 4. Critical Period Acceleration by Early Otx2 Infusion

(A) Little ocular dominance shift (toward group 4, dashed line) is observed at P22-23 after monocular deprivation (MD) from P18 (top) but is induced prematurely

by Otx2 infusion (bottom) (p < 0.0001, c2-test).

(B) Ocular dominance distributions at P28-30 are similar for nondeprived vehicle (top) or Otx2-treated (bottom) mice (p > 0.5, c2-test).

(C) Brief MD from P25 shifts ocular dominance in the vehicle control group (top) but no longer does so in animals receiving Otx2 earlier at P17 (bottom)(p < 0.0001,

c2-test). Number of mice in (D).

(D) Accelerated plasticity profile for Otx2. High CBI values of individual animals normally shift when MD begins at P25-26 (vehicle) but shift prematurely at P18

when Otx2 is infused (*** p < 0.0001, t test).

(E) Prolonged neuronal discharge is reduced at P22-23 by Otx2 infusion (vehicle, 68.5 ± 2.7% versus Otx2, 55.8 ± 4.3% of cells, p < 0.05, t test), then drops off

normally by P28 in both groups (vehicle, 6.3 ± 2.1% versus Otx2, 6.7 ± 2.5%, p > 0.9, t test).

(F) Orientation-biased cells emerge slowly after eye-opening regardless of early Otx2 infusion (vehicle, 18.2 ± 2.5% versus Otx2, 15.2 ± 0.8% of cells at P22-23,

p > 0.2, t test; vehicle, 29.8 ± 1.0% versus Otx2, 30.1 ± 1.4% at P28-30, p > 0.8, t test). Error bars represent SEM.


left the receptive field (prolonged discharge). This is typically

prominent in GAD65 knockout (Hensch et al., 1998; Iwai et al.,

2003) or young wild-type mice before the critical period

(Iwai et al., 2003; Fagiolini et al., 2004). Like benzodiazepine

treatment (Hensch et al., 1998), early Otx2 infusion significantly

reduced this immature property in vivo, indicating a rapid func-

tional maturation of inhibitory circuits by this homeoprotein

(Figure 4E). Other receptive field properties, such as low orienta-

tion bias at young ages, were not affected by early Otx2 infusion

(Figure 4F and Table S1), consistent with a molecular dissocia-

tion of ocular dominance and orientation development as re-

ported previously (Hensch et al., 1998; Fagiolini et al., 2003).

Taken together, Otx2 gain-of-function promoted critical period

onset in conjunction with PV-cell maturation.

To confirm whether Otx2 is required for PV-cell maturation and

critical period plasticity, we next carried out Otx2 loss-of-func-

tion experiments. Due to the embryonic lethality of constitutive

Otx2 deletion (Acampora et al., 2001), we turned to a conditional

approach. Otx2 protein was detected within PV-cells at P28 in

Otx2 flox/flox mice in which the Otx2 gene was flanked by LoxP

sites (Tian et al., 2002) (Figure 5A). In contrast, when one Otx2

allele was excised by Cre recombinase under the control of

a postnatal a-calcium/calmodulin-dependent protein kinase II

(aCaMKII) promoter, cortical Otx2 signal was strongly reduced

(Figure 5B). Strikingly, PV expression was also reduced when en-

dogenous Otx2 was disrupted (Figures 5C and 5D), retaining an

immature appearance (see Figure 1B). Immunofluorescence

intensity within PV-cell somata was significantly decreased in

the cumulative distribution or comparisons of average intensity

for individual mice (Figure 5G).

Loss of Otx2 also delayed PNN formation and reduced the

number of cells labeled by WFA (Figures 5F and 5H; control,

43.9 ± 1.8 versus hetKO, 32.0 ± 1.8 cells/field at P28-30; control,

59.9 ± 1.7 versus hetKO, 44.6 ± 1.7 for adults, > P60, respec-

tively; p < 0.0005, t test) or Cat-301-antigen (Figures S2F and

S2G; control, 30.1 ± 1.9 versus hetKO, 15.6 ± 1.8 cells/field at

P28-30; control, 56.8 ± 3.4 versus hetKO, 37.2 ± 2.7 for adults, >

P60, respectively; p < 0.0005, t test). Moreover, PV-expressing

cell number was also reduced by the loss of endogenous Otx2

(Figure S2D; control, 38.0 ± 4.4 versus hetKO, 28.6 ± 1.0 cells/

field, p < 0.05, t test). These effects were specific to PV-cells

since Calretinin-cells were not affected in expression level (Fig-

ures S2A–S2C) or number (Figure S2D; control, 18.5 ± 1.3 versus

hetKO, 20.3 ± 2.3 cells/field, p > 0.5, t test).

Postnatal Otx2 loss-of-function would then be expected to

prevent plasticity if somatic inhibition is compromised (Katagiri

et al., 2007), as for global GAD65 disruption (Hensch et al.,

1998; Hensch, 2005). The frequency of miniature IPSCs, arising

primarily from peri-somatic inputs (Soltesz et al., 1995), was sig-

nificantly reduced in slices from Otx2 KO mice (A. Scheuber and

T.K.H., unpublished data). Ocular dominance plasticity failed to

occur concomitant with immature PV expression (Figure 5D) in

mice lacking even one Otx2 allele (Figures 5I, 5J, and 5L), which

had no noticeable effect on other receptive field properties (re-

sponsiveness, orientation bias, habituation; Table S1). We then

attempted to rescue plasticity in vivo using benzodiazepine ag-

onists to enhance GABAA receptor currents. Ocular dominance

shifts were restored by diazepam treatment during brief MD (Fig-

ures 5K and 5L) as in GAD65 KO mice (Hensch et al., 1998). En-

dogenous Otx2 may thus activate critical period plasticity

through the functional maturation of inhibitory circuits.

Non-Cell-Autonomous Otx2 in PV-CellsRemarkably, inhibitory interneurons in the cortex do not express

aCaMKII (Liu and Jones, 1996; Sik et al., 1998). Crossing

Otx2flox/flox mice with those expressing a Cre-recombinase

driven by the aCaMKII promoter above (Minichiello et al., 1999)

thus results in a conditional disruption of Otx2 synthesis outside

cortical GABA cells. The impairment of PV-cell maturation and

critical period plasticity (Figure 5) must therefore reflect a non-

cell-autonomous action of Otx2.

Consistent with this idea, Otx2 transcripts were not detected in

the visual cortex by in situ hybridization analyses (Figure 6A) or

RT-PCR (Figure S3A) during the critical period. Green fluores-

cent protein (GFP) transcribed from the Otx2 locus in Otx2-

GFP knock-in mice (c/o Drs. D. Acampora and A. Simeone)

also failed to show any cortical signal (Figure 6B). Instead,

both Otx2 mRNA and Otx2-GFP expression were observed as

expected at sub-cortical stages of the visual pathway in the lat-

eral geniculate nucleus (LGN, Figures 6D and 6E), superior colli-

culus (Figures S3A and S3B) and retina (Figures 6G, 6H, S4A,

and S4C) (Bovolenta et al., 1997; Baas et al., 2000; Frantz

et al., 1994; Nothias et al., 1998). These observations confirm

that Otx2 is not synthesized in cortical PV-cells themselves.

Homeoprotein family members have been shown to contain

secretion and internalization signals in the homeodomain (Joliot

and Prochiantz, 2004) and are found in the secretory compart-

ment in vivo (Joliot et al., 1997). This would allow potential trans-

fer from cell to cell as observed in vitro (Joliot and Prochiantz,

2004), raising the possibility that Otx2 homeoprotein may be

propagated into the visual cortex along the visual pathway

(Figure 6C). Even within the retina Otx2 protein was found in gan-

glion cells (Figure 6I), downstream of the bipolar cells containing

its mRNA in the inner nuclear layer (Figure 6G). In the LGN, Otx2

protein was diffusely spread over multiple cells (Figure 6F) unlike

Otx2 transcripts found within discrete GABAergic interneurons

(Figure 6D; GABA+/Otx2+, 93.7 ± 1.4%, 283 cells; Otx2+/

GABA+, 92.0 ± 2.4%, 287 cells; 4 mice; Sugino et al., 2006).

Otx2 along the Visual PathwayTo demonstrate a potential route of transport for Otx2 protein

in vivo along the visual pathway (Figure 6C), we introduced

a tagged protein into the eyes. After an intraocular injection of

biotinylated Otx2, labeled protein was detected in the visual

cortex, where the biotin tag was internalized into cortical cells

(Figure 6J). Notably, the proportion of cells that were both

Otx2-labeled and PV-positive (Figures 6K and 6L; 77.7 ± 3.6%)

was identical to the endogenous ratio (Figure 1J). Consistent

with its non-cell-autonomous distribution in the visual cortex,

neither uninjected nor BSA-biotin injected animals exhibited

such a cortical label (Figures 6M–6O).

Otx2 expression at sub-cortical stages persists from early

postnatal days before eye-opening (Frantz et al., 1994; Baas

et al., 2000). In our conditional deletion mice, Otx2 transcripts

were reduced in the inner nuclear layer of the retina by aCaMKII

promoter-mediated Cre expression (Figures S4A and S4B; Liu


Figure 5. Otx2 Loss-of-Function Prevents PV-Cell Maturation and Visual Cortical Plasticity

(A–F) Both expression of PV (D) and WFA-stained perineuronal nets (F) is reduced at P28 by Otx2 disruption (B) compared to control mice (A, C, and E). The scale

bar represents 50 mm.

(G) Quantitative analysis of PV intensity within somata (open symbols, control; filled symbols, KO) in supragranular layers of the binocular zone (600 3 350 mm area

at P28-30; K-S test, p < 0.001; inset, p < 0.05, unpaired t test for average intensities).

(H) Number of perineuronal net-enwrapped cells (WFA) is reduced throughout life by Otx2 disruption (4-7 mice, 600x350 mm area; CP, P28-30 versus adult, > P60

for control or hetKO, respectively; p < 0.0001, t test; control versus hetKO for CP or adult, respectively; p < 0.0005, t test). Error bars represent SEM.

(I–K) Conditional deletion of Otx2 from non-PV cells is sufficient to block ocular dominance plasticity (I versus J, p < 0.0001, c2-test), which is rescued by diaz-

epam (DZ) injection concurrent with monocular deprivation from P25 (K). Number of mice in (L).

(L) CBI values of nondeprived mice are typically high regardless of genotype (0.72 ± 0.01). Reduction of CBI values by MD for individual animals (open symbols,

control; filled symbols, KO) fails to occur after conditional Otx2 deletion and is rescued by diazepam (control MD, 0.54 ± 0.02; homoKO, 0.66 ± 0.01; hetKO, 0.67

± 0.01; diazepam injected hetKO, 0.57 ± 0.01; ** p < 0.001, * p < 0.01, ANOVA).


Figure 6. Otx2 Transfer along the Visual Path-

way

(A and B) Neither Otx2 mRNA ([A] at P28) nor

Otx2-GFP expression in knock-in mice ([B] at P35)

is detected in visual cortex (VC).

(C) Proposed propagation of Otx2 homeoprotein

along the mouse visual pathway.

(D–F) Restricted expression of Otx2 transcripts (D)

and Otx2-GFP (E) within LGN interneurons. In con-

trast, Otx2 protein (F, at P28) is detected diffusely

across multiple cells of the LGN.

(G–I) Similar localization of Otx2 mRNA (G) and Otx2-

GFP (green [H]) but differential distribution of Otx2

protein (arrowheads [I]) in the retina: ganglion cells,

GC; inner plexiform layer, IPL; inner nuclear layer,

INL; and outer nuclear layer ONL.

(J–L) Passage of biotinylated Otx2 (green) into visual

cortex after intraocular injection (J) and internali-

zation by PV-cells (red) (K, L; proportion of biotin-

labeled cells containing PV, 77.7 ± 3.6%, 186 cells,

600x350 mm area, eight slices from four mice;

arrows, PV-negative cells).

(M–O) Absence of biotinylated BSA (green) in visual

cortex (M) or PV-cells (red) (N, O) after similar intraoc-

ular injection. The scale bars represent 250 mm (A

and B), 100 mm (D–F and J–O), 25 mm (G–I).


et al., 2000). Although visual experience through the eyes was re-

quired for cortical Otx2 appearance, retinal Otx2 synthesis was

unaffected by dark-rearing (Figures S3C and S3D). The emer-

gence of endogenous Otx2 within visual cortex may then include

an experience-dependent transport or transfer, which might

explain homeoprotein accumulation within cortical PV-cells

only after eye-opening and the onset of vision.

We directly addressed the putative passage of Otx2 by local

infusion of antagonists either into the eye or cortex (Figure 7A).

To spatio-temporally intercept the extracellular transfer of Otx2

we used a specific antibody. Seven-day infusion of antibody

into visual cortex (P21-23 to P28-30) was sufficient to capture

and clear Otx2 in the extracellular space (Figures S4E and

S4F). To knock-down the synthesis of homeoprotein we gener-

ated short-interfering RNA (siRNA) constructs coupled to pene-

tratin (Joliot and Prochiantz, 2004). siRNA efficacy was verified in

HeLa cells expressing Otx2 in vitro and in the inner nuclear layer

of the mouse retina in vivo (Figures 7E, S4C, and S4D). These

treatments showed no general effect upon receptive field prop-

erties in visual cortex (Table S1).

Instead, both early intraocular injection and intracortical infu-

sion of anti-Otx2 antibody successfully prevented plasticity (Fig-

ures 7B and 7C). Ocular dominance remained skewed in favor of

the contralateral eye (left of group 4) even after brief MD at P25

(Figure 7F ‘‘a-Otx2’’). In contrast, intracortical siRNA injection

did not block critical period plasticity (Figure 7D), consistent

with the absence of Otx2 mRNA in the cortex (Figure 6A). To pre-

vent plasticity, siRNA was effective only in the eye (Figure 7F

‘‘siRNA’’). Taken together, these results support the idea that

Otx2 is synthesized and transferred from outside the visual

cortex to enable visual cortical plasticity.

DISCUSSION

Our results demonstrate a novel postnatal role for an embryonic

homeoprotein, Otx2, in critical period brain development. Its

mechanism of action is entirely consistent with recent under-

standing of plasticity in the visual cortex based upon the activa-

tion of particular GABAergic connections (Hensch, 2005). Endur-

ing behavioral amblyopia (loss of visual acuity after MD) reflects

a shift in spiking output from V1 during a restricted critical period

(Gordon & Stryker, 1996; Prusky and Douglas, 2003; Tohmi et al.,

2006; Morishita and Hensch, 2008). Spike timing is dictated by

the maturation of GABAergic transmission at somatic synapses

containing the GABAA a1- receptor subunit (Fagiolini et al., 2004;

Katagiri et al., 2007), typically innervated by large-basket PV-

cells which mature postnatally in anticipation of the critical pe-

riod (del Rio et al., 1994; Huang et al., 1999; Di Cristo et al., 2007).

Direct Otx2 gain- or loss-of-function, respectively, promoted

or hindered both the expression of PV-cell molecular constitu-

ents and ocular dominance plasticity. Surprisingly, these bidirec-

tional actions were non-cell-autonomous. Selective deletion of

the Otx2 gene outside cortical interneurons successfully re-

moved Otx2 protein from PV-cells. Moreover, intercepting

Otx2 by infusion of antibodies or blocking its synthesis by siRNA

injection into the eye prevented critical period plasticity. It is in-

triguing that passage of extra-cortical factors may establish

the milieu for cortical plasticity in vivo. Robust Otx2 mRNA ex-


pression is observed in the retina, LGN and superior colliculus,

demarcating a possible conduit for propagation along the visual

pathway (Bovolenta et al., 1997; Baas et al., 2000; Frantz et al.,

1994; Nothias et al., 1998; Sugino et al., 2006; Rath et al.,

2007). The mechanisms of transport and transfer in vivo will be

an important line of investigation.

Homeoprotein family members contain secretion and internal-

ization sequences in the homeodomain, allowing potential trans-

fer from cell to cell as observed in vitro (Joliot and Prochiantz,

2004; Brunet et al., 2007). Recent studies have demonstrated

that embryonic morphogens like WNTs, FGFs and TGFb can

be re-used as anterograde or retrograde factors like neurotro-

phins for neuronal outgrowth and juxtaposition of synaptic com-

ponents in the formation of mature synaptic connections

(Marshak et al., 2007; Mandolesi et al., 2005; Packard et al.,

2003; Salinas, 2005). Thus, the conventional strategy to search

for ‘plasticity genes’ from cortical cDNA libraries (Majdan and

Shatz, 2006; Tropea et al., 2006) may need to be expanded to

include early embryonic factors that originate sub-cortically.

It is generally thought that the number of interneurons and their

diversification has increased with evolution to regulate cortical

function. Cortical interneurons originate within the subcortical

telencephalon (for PV-cells the medial ganglionic eminence)

and then migrate tangentially into the overlying cortex but never

express Otx2 (Butt et al., 2007; Wonders and Anderson, 2006).

Otx2 signals anterior ‘‘head’’ position in all metazoans, starting

with the hydra. Like many homeoproteins (Brunet et al., 2007),

its role has preceded that of classical signaling molecules and

mediators of Otx2 expression in the cortex (e.g., activity–depen-

dent BDNF is absent even in arthropods).

Homeoproteins may act both as transcription factors and reg-

ulators of local protein synthesis (Dubnau and Struhl, 1996;

Nedelec et al., 2004; Brunet et al., 2005), perhaps through direct

interaction with eukaryotic translation initiation factor 4E (eIF4E)

(Nedelec et al., 2004; Topisirovic and Borden, 2005). Once inter-

nalized by PV cells, Otx2 has direct access to the cytoplasm,

where it can engage the protein translation apparatus as shown

for Engrailed in migrating growth cones (Brunet et al., 2005).

A dual action at either the transcriptional or translational level

could accelerate PV-cell maturation and critical period onset

simply by enhancing their known sensitivity to ambient neurotro-

phins such as BDNF (e.g., by upregulation of TrkB receptor)

(Huang et al., 1999; Marty et al., 1997).

Interestingly, PNN structures consisting of cell-adhesion mol-

ecules, hyaluronic acids and chondroitin sulfate proteoglycans

(CSPGs) tightly enwrap mature PV-cells specifically (Celio

et al., 1998). Our gain-of-function and loss-of-function results

show that Otx2 accumulation can induce the maturation of

PNNs. Several putative Otx2 binding sites are found in the

promoter region for Tenascin (Briata et al., 1999), a major extra-

cellular matrix component which may directly regulate PNN

development (Nguyen Ba-Charvet et al., 1998). Indeed, Tenas-

cin-R KO mice exhibit severe defects in PNN formation on

PV-cells as well as weakened GABA function (Bruckner et al.,

2000; Nikonenko et al., 2003; Saghatelyan et al., 2000).

The emergence of Otx2 within PV-cells may further serve as

a positive feedback loop: promoting PNN maturation that conse-

quently promotes Otx2 uptake and the strengthening of somatic

Figure 7. Disruption of Otx2 Propagation Prevents Visual Cortical Plasticity

(A) Experience-dependent passage of Otx2 protein along the visual pathway may establish the cortical milieu for critical period plasticity.

(B–D) Ocular dominance shift (low CBI) is induced after monocular deprivation (MD) from P25 (B) but is not observed following cortical infusion of inhibitory a-Otx2

antibody (C) (p < 0.0001, c2-test). Similarly infused siRNA constructs into visual cortex have no effect on plasticity (D). See methods for injection details. Number of

mice as in (F).

(E) Expression of Otx2-HA is efficiently inhibited in HeLa cells co-transfected with siRNA against Otx2 but not for a control gene. Error bars represent SD.

(F) Reduction of CBI values for individual animals after MD (open symbols, antibody vehicle; gray symbols, control siRNA) is significantly prevented by intraocular

injection of antibody (Veh, 0.52 ± 0.02; a-Otx2, 0.65 ± 0.01) or siRNA (Veh, 0.53 ± 0.01, Otx2 siRNA, 0.64 ± 0.01) (** p < 0.001 versus respective controls, t test). In

contrast, only cortical antibody injection (Veh, 0.53 ± 0.01; a-Otx2, 0.68 ± 0.01; Otx2 siRNA, 0.55 ± 0.01) is similarly effective (** p < 0.001 versus Veh, ANOVA).


inhibition at critical period onset. Negatively charged CSPG

sugars in the PNN bind plant lectins such as WFA with high

affinity and may do the same with positively-charged, homeodo-

main-containing proteins to facilitate internalization. The endog-

enous preference of Otx2 for PV-cells was unexpected from

in vitro studies (Joliot and Prochiantz, 2004). We found that other

infused homeoproteins (Engrailed-2) fail to accumulate in corti-

cal PV-cells (data not shown). Specific sugar epitopes may be

recognized by distinct homeoproteins according to a putative

‘sugar code’ for future study (Holt and Dickson, 2005).

Importantly, the removal of PNNs with chondroitinase injection

reactivates critical period plasticity in adulthood (Pizzorusso

et al., 2002). We found that similar treatment depleted adult

PV-cells of their Otx2 content, resetting them to an immature

state (M. Beurdeley and S.S., data not shown; Saghatelyan

et al., 2000). Instead, high polysialic acid expression during the

pre-critical period would nonspecifically trap Otx2 (Joliot et al.,

1991), leading to a reduction of homeoprotein internalization by

PV-cells, delaying their maturation and visual cortical plasticity

(Di Cristo et al., 2007). Plasticity onset may therefore reflect the

emergence of PV-cell function from a weakened state rather

than as a consequence of PNN removal per se. A dynamic, bidi-

rectional plasticity of afferent input onto PV-cells appears to

underlie critical period plasticity then becomes fixed with age

(Y. Yazaki-Sugiyama and T.K.H., unpublished data). It will be in-

teresting to map the detailed time-course of Otx2 accumulation

and critical period reactivation specifically in PV-cells when

PNNs are dissolved by chondroitinase treatment in the adult.

Persistent passage of Otx2 may further explain two visual phe-

nomena across the life spectrum. Dark-rearing from birth classi-

cally delays critical period onset (Cynader and Mitchell, 1980;

Fagiolini et al., 1994, 2003) by a specific reduction of somatic in-

hibition (Iwai et al., 2003; Katagiri et al., 2007). Here, we find that

Otx2 transport or accumulation into PV-cells is also experience-

dependent and appears only gradually in visual cortex after eye

opening with the onset of natural vision. Critical period time-

course may thus shift with impoverished or enriched environ-

ments (Cancedda et al., 2004) perhaps by regulating delivery

of morphogenic factors from the periphery.

Conversely, intracortical inhibition deteriorates upon aging,

degrading vision but paradoxically improving motion detection

(Leventhal et al., 2003; Betts et al., 2005). Failing eyesight with

senescence could reduce the visually-driven, sustained passage

of Otx2 into cortical interneurons leading to these effects.

Indeed, the dosage effect of Otx2 (one allele is not enough to

activate plasticity) confirms that variations in activity, paralleled

by variation in Otx2 transport, may have graded effects on the re-

finement of cortical architecture. The distribution of factors could

be variable among individual animals exposed to distinct sen-

sory experiences.

Modality-specific transport of a molecular messenger by ac-

tivity from the periphery into the respective cortical area may

be a generally safe way to signal when the sensory epithelium

is fully functional (e.g., whisking before eye opening) prior to en-

gaging local cortical plastic machinery. This would give rise to

differential critical period timecourse across brain regions and

may contribute to functional decline with aging when the sensory

periphery begins to deteriorate. Access to the cortical milieu for


plasticity from the periphery may in turn offer a powerful thera-

peutic tool for recovery of function. It will be of great interest to

confirm whether postnatal plasticity in each sensory modality re-

lies on a unique homeoprotein.

EXPERIMENTAL PROCEDURES

Animals

Conventionally raised C57Bl/6 mice (12:12 hours light:dark cycle) were pur-

chased from Japan SLC. Otx2flox/flox mice (Tian et al., 2002) were crossed

with aCaMKII-Cre transgenic mice (Minichiello et al., 1999) to generate an

Otx2 conditional knockout mouse line. GFP knock-in mice were kindly pro-

vided by Drs. D. Acampora and A. Simeone (MRC, King’s College London,

UK).

cDNA Amplification

Mouse Otx2 and GABAA receptor a2 subunit cDNA was amplified by one-step

RT-PCR (Invitrogen) with degenerate oligonucleotides (see Supplemental

Data). Each cDNA was inserted into pBluescript to confirm the sequence

and to generate the RNA probe.

In Situ Hybridization and Immunohistochemistry

In situ hybridization on floating sections was performed according to methods

described previously (Sugiyama and Nakamura, 2003). Mice were perfused

and post-fixed with 4% paraformaldehyde (PFA). Coronal sections (20 mm)

were hybridized at 65�C with a Digoxigenin (DIG)-labeled RNA probe for

Otx2, then detected by alkaline phosphate (AP)-conjugated anti-DIG antibody.

Primary antibodies (see Supplemental Data) or Biotin-labeled lectin from

Wisteria Floribunda agglutinin (WFA, Sigma) were incubated with floating cor-

onal sections (20 mm) at 4�C over night. Specificity of anti-Otx2 antibodies was

tested by preincubation with human Otx2 protein (2 mg/ml, produced in bacu-

lovirus) for 30 min before immunoreaction with tissues.

Quantification

For quantitative analysis of fluorescence intensity, immunostaining was per-

formed concurrently with the same solutions, and images were photographed

in one sitting with the same gain and exposure time.

The number of cells or intensity of immunofluorescence was measured in

a 600 3 350 mm area covering the supragranular layers of the binocular

zone. The intensity per cell was plotted on a cumulative histogram for compar-

ison across groups (Kolmogorov-Smirnov test). Averaged intensity per hemi-

sphere or for each mouse was compared pair-wise across the same brain or

unpaired between two groups of animals, respectively (t test).

The intensity of Otx2 immunofluorescence in retina was measured per pixel

across a 200 3 20 mm area, covering the inner nuclear layer. Averaged intensity

per mouse was compared between the two groups (t test).

Drug Infusion

Otx2 protein, inhibitory antibody against Otx2 (a-Otx2), vehicle solution or pen-

etratin-coupled siRNA against Otx2 or control (Muratovska and Eccles, 2004)

was infused into the right visual cortex. Cortical infusion was performed by os-

motic minipumps (Alzet1007D, Alza) connected to cannulae stereotaxically

implanted in mice at ages indicated in the text (Hensch et al., 1998). Diazepam

(DZ) or vehicle solution was injected daily into both lateral ventricles, starting

one day before monocular deprivation until the day before recording (Hensch

et al., 1998).

Biotinylated human Otx2 protein or bovine serum albumin (BSA), a-Otx2, ve-

hicle solution or penetratin-coupled siRNA against Otx2 or control was

injected into both eyes by micropipette (at P21 and P23 for proteins; at P16,

P19 and P22 for a-Otx2; at P18, P19, P21, and P22 for siRNA).

Single-Unit Electrophysiology In Vivo

Electrophysiological recordings were performed using standard techniques

(Hensch et al., 1998). Five to seven single units were recorded for each pene-

tration (>3 spaced at 200 mm intervals) across the medio-lateral axis of the bin-

ocular zone to avoid sampling bias. Multiple receptive field properties

(prolonged discharge, orientation bias, response strength, habituation and ret-

inotopy) were evaluated for each mouse (see Supplemental Data; Gordon and

Stryker, 1996; Hensch et al., 1998; Taha et al., 2002). Responses from single

cells were assigned ocular dominance scores using a seven-point classifica-

tion scheme (Wiesel and Hubel, 1963; Hensch et al., 1998). Ocular dominance

in the binocular zone from each mouse was calculated as a contralateral bias

index (CBI, see Supplemental Data). Visual responses after brief (3–5 days)

monocular deprivation (MD) were recorded from the hemisphere contralateral

to the deprived eye.

SUPPLEMENTAL DATA

Supplemental Data include Supplemental Experimental Procedures, Sup-

plemental References, four figures, and one table and can be found with this

article online at http://www.cell.com/cgi/content/full/134/3/508/DC1/.

ACKNOWLEDGMENTS

We thank D. Acampora and A. Simeone for graciously providing the Otx2-GFP

mouse line and the original Otx2 plasmids; A. Ruiz i Altaba and M. Wassef for

Otx2 antibody; S. Dupas for recombinant Otx2 protein; G. Corte for recombi-

nant baculovirus; K. Moya, M. Beurdeley, and A. Scheuber for help in some ex-

periments and discussion. Funded by the Human Frontiers Science Program

(HFSP, RGP0026/2002), Fondation pour La Recherche Medicale (Longevite

Cognitive et Sensorielle), and in part by the Special Postdoctoral Research

Program (RIKEN) and Japanese Ministry of Science, Education and Technol-

ogy (MEXT, IBR 17021047). S.S. is a Research Fellow of the Japan Society

for the Promotion of Science (JSPS). A.A.D. is a fellow of the Canadian Insti-

tutes of Health Research (CIHR).

Received: June 22, 2007

Revised: April 22, 2008

Accepted: May 5, 2008

Published: August 7, 2008

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