Visual Activity Regulates Neural Progenitor Cells in Developing Xenopus CNS through Musashi1
Neuron. Author manuscript; available in PMC 2011 November 4.
Published in final edited form as:
11/7/13 4:01 PM
PMCID: PMC3005332
NIHMSID: NIHMS241844
Neuron. 2010 November 4; 68(3): 442–455.
doi: 10.1016/j.neuron.2010.09.028
Visual Activity Regulates Neural Progenitor Cells in Developing Xenopus CNS
through Musashi1
Pranav Sharma and Hollis T. Cline*
The Scripps Research Institute, 10550 North Torrey Pines Road, La Jolla, CA 92037, USA
*To whom correspondence should be addressed. Email: cline@scripps.edu
Copyright notice and Disclaimer
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Abstract
Regulation of progenitor cell fate determines the numbers of neurons in the developing brain. While
proliferation of neural progenitors predominates during early CNS development, progenitor cell fate
shifts toward differentiation as CNS circuits develop, suggesting that signals from developing circuits may
regulate proliferation and differentiation. We tested whether activity regulates neurogenesis in vivo in the
developing visual system of Xenopus tadpoles. Both cell proliferation and the number of musashi1immunoreactive progenitors in the optic tectum decrease as visual system connections become stronger.
Visual deprivation for 2 days increased proliferation of musashi1-immunoreactive radial glial progenitors
while visual experience increased neuronal differentiation. Morpholino-mediated knockdown and overexpression of musashi1 indicate that musashi1 is necessary and sufficient for neural progenitor
proliferation in the CNS. These data demonstrate a novel mechanism by which increased brain activity in
developing circuits decreases cell proliferation and increases neuronal differentiation through the downregulation of musashi1 in response to circuit activity.
Keywords: neurogenesis, visual experience, Xenopus, musashi1, radial glial cells, BrdU, progenitor
pool, differentiation, cell cycle, N-β-tubulin, nrp1
Introduction
The number of neurons in the brain is largely determined by the regulation of neural progenitor cell
proliferation and the survival and differentiation of their progeny. The pool of neural progenitor cells can
be expanded by symmetric divisions that give rise to two neural progenitor cells, maintained by
asymmetric divisions that result in one of the progeny remaining a neural progenitor cell while the other
differentiates, or depleted by terminal differentiation (Butt et al., 2005; Gotz and Huttner, 2005; Huttner
and Kosodo, 2005; Kriegstein and Alvarez-Buylla, 2009; Noctor et al., 2007). While expansion and
maintenance of neural progenitor cells is favored over differentiation during early CNS development,
differentiation progressively dominates progenitor cell fate leading to depletion of the pool of progenitors
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as CNS circuits develop (Kriegstein and Alvarez-Buylla, 2009), suggesting that signals from developing
circuits may shift the fate of neural progenitors and their progeny. Indeed, recent studies suggest that
neuron-derived trophic factors may induce progenitors to stop dividing and differentiate (Botia et al.,
2007; Kriegstein and Alvarez-Buylla, 2009), however there is little evidence of endogenous activitydependent regulation of progenitor cells in intact animals. Therefore, a fundamental open question in the
regulation of neurogenesis is whether feedback mechanisms from developing neuronal circuits regulate
progenitor fate to expand, maintain or deplete the pool of neural progenitor cells in the developing CNS
in vivo.
To address this question we tested whether visual activity affects the rate of ongoing cell proliferation in
the developing visual system of Xenopus laevis tadpoles, where cell proliferation in the optic tectum
continues over an extended period of development while the visual circuitry is both functional and still in
the process of development (Cline, 2001; Peunova et al., 2001; Straznicky and Gaze, 1972). We find that
cell proliferation in the tectum, detected by BrdU incorporation, decreased as visual circuitry matured
between stages 46 and 49. Over the same period, immunoreactivity for MCM7, a marker of cells with
proliferative potential (Crevel et al., 2007; Facoetti et al., 2006; Khalili et al., 2003), and musashi1, an
RNA binding protein that is essential for maintenance of the neural progenitor population (Glazer et al.,
2008; Kaneko et al., 2000; Okano et al., 2005) decreased, correlating with the developmental decrease in
proliferation. These data are consistent with the idea that visual activity in the more mature circuit could
negatively regulate cell proliferation. Indeed, brief visual deprivation for 2 days increased cell
proliferation in the optic tectum compared to animals with visual experience, suggesting that feedback
from the developing visual circuit shifts the fate of neural progenitors. We used sequential exposure to
two differentially halogenated thymidine analogs (IdU and CldU, referred to collectively as XdUs) to
reveal the division history of proliferating cells (Encinas and Enikolopov, 2008; Vega and Peterson,
2005) and found that a larger fraction of cells in animals with brief visual deprivation remain in the cell
cycle, whereas more cells exit the cell cycle and differentiate into neurons in animals with visual
experience. Interestingly, visually-deprived animals have more musashi1-immunoreactive radial glial
progenitors than animals with visual experience. Morpholino-mediated knockdown and rescue
experiments show that musashi1 is required for the increased proliferation seen with visual-deprivation.
Finally, exogenous expression of musashi1 in stage 49 radial glial cells, which have little detectable
endogenous musashi1-immunoreactiviey and low proliferative activity, increases their proliferation. Our
study suggests that sensory experience plays a role in neurogenesis in the developing CNS in vivo by
regulating the fate of progenitors and their progeny.
Results
Cell proliferation in the optic tectum decreases with visual system development
In the visual system of Xenopus laevis tadpoles retinal ganglion cells project axons to the contralateral
optic tectum where they form synapses with tectal neurons (Fig. 1A-C). Between stages 39 and 49, a
period of 6-7 days, the visual system of Xenopus tadpoles develops rapidly to accommodate the
behavioral needs of the animal. Retinal ganglion cells first innervate and transmit visual information to
the optic tectum at stage 39 (Holt and Harris, 1983) when the majority of cells in the tectum have radial
glial morphology and neurons have very simple dendritic arbors (Wu et al., 1999). An initial topographic
retinotectal map is established by stage 45 (O'Rourke and Fraser, 1990) and between stages 46 and 49
visual experience drives many aspects of visual circuit development pertaining to the detection and
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processing of visual inputs (Bestman and Cline, 2008; Chiu et al., 2008; Cline and Haas, 2008; Engert et
al., 2002; Pratt and Aizenman, 2007, 2009; Pratt et al., 2008; Tao and Poo, 2005) even as ventricular
layer cells with radial glial morphology persist in the tectum (Tremblay et al., 2009). Although it is well
known that tectal ventricular layer cells proliferate throughout tadpole stages of development and
generate neurons within the tectum (Peunova et al., 2001; Straznicky and Gaze, 1972), a potential relation
between development of the functional visual circuit and cell proliferation has not been explored.
To test whether rates of cell proliferation in the optic tectum change over this period of visual system
development, we exposed tadpoles at stage 46, 48 and 49 to 2 hr of XdU and either processed the brains
immediately or allowed the animals to develop in normal rearing solution for another 22 hr before
processing the brains as wholemounts for XdU immunodetection. We delivered XdU by exposing
tadpoles to rearing solution containing 10 mM XdU for 2 hr. This method efficiently labels proliferative
cells in the brain and allows greater control over XdU exposure time than standard injection methods
(Peunova et al., 2001). Brains were processed to detect XdU with antibodies and a complete confocal Zseries of images was collected through the midbrain of wholemount brains or cryostat sections. As
previously reported, proliferating cells, identified by exposure to 3H-thymidine (Straznicky and Gaze,
1972) or BrdU-labeling (Peunova et al., 2001) with a short survival time, line the ventricle (Figs 1D,F and
3A,B). We counted XdU-labeled cells in the ventricular layer at the tectal midline using the dissector
method and indexed cell counts to an estimated volume of 20,000 µm3 (see Fig 1D and methods). Stage
46 tadpoles have significantly more XdU-labeled cells (99.6 ± 2.4cells/20,000µm3, n=8) than either
stage 48 (36.1 ± 2.3 cells/20,000µm3, n=8; p<0.05) or stage 49 (8.9 ± 0.8 cells/20,000µm3, n=7;
p<0.05) tadpoles (Fig 1H, Supplementary Movies S1, S2 and S3). The XdU-labeled cells continue to
divide over the next 22 hr to approximately double the number of XdU-labeled cells (Fig 1I. Stage 46:
180.4±4.9 cells/20,000µm3, n=8; Stage 48: 71.2±3.9 cells/20,000µm3, n=8; Stage 49: 20.3±0.9
cells/20,000µm3, n=7). These data suggest that proliferation gradually decreases between stages 46 and
49, a time interval during which the visual circuit matures.
Cells expressing MCM7 or Musashi decrease with development
We next tested whether MCM7 and Musashi1, which have been characterized as markers of proliferative
cells in other experimental systems, change expression over the developmental period when XdU
incorporation decreased. MCM7 is a part of minichromosome maintenance complex (MCM) of proteins
and is expressed in cells with proliferative potential (Crevel et al., 2007; Facoetti et al., 2006; Khalili et
al., 2003). Proteins in the MCM complex are down-regulated when cells become quiescent, differentiated,
or senescent (Facoetti et al., 2006; Padmanabhan et al., 2004). Antibodies to MCM7 are thought to label
the population of cells that is not yet differentiated and is capable of proliferation (Blow and Dutta,
2005). In the optic tectum, we find that MCM7-immunoreactive cells are present in the cell layers lining
the ventricle (Fig. 2 A1,A2 and B1, B2) and partially overlap with the distribution of cells detected by
incorporation of XdU following 2h exposure. MCM7 expression was down-regulated between stage 46
and 49 (Fig. 2 A1,A2 and B1, B2), consistent with the decrease in XdU-incorporation over this time
period.
The Musashi proteins are highly conserved RNA binding proteins (Good et al., 1993; Nakamura et al.,
1994; Sakakibara et al., 2001), whose founding member, Musashi1, was originally discovered in Xenopus
and named nrp1 (Richter et al., 1990). In vertebrates, there are 2 genes, musashi1 and musashi2, the
latter of which is homologous to xrp1 in Xenopus. We focused on Musashi1 because it is expressed
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exclusively in the CNS and is enriched in CNS progenitor cells across phyla (Amato et al., 2005; Good et
al., 1993; Kaneko et al., 2000; Nakamura et al., 1994; Sakakibara et al., 1996; Sakakibara and Okano,
1997) and because it is required for maintenance of neural stem cells (Okano et al., 2002; Okano et al.,
2005; Sakakibara et al., 2001; Sakakibara et al., 2002), whereas musashi2/xrp1 is widely expressed
throughout the body. Musashi1 protein expression is down-regulated in differentiated neurons (Kaneko
et al., 2000; Nakamura et al., 1994; Sakakibara et al., 1996; Sakakibara and Okano, 1997). We find that
musashi1-immunoreactive cells are present in the proliferative layer lining the ventricle of the optic
tectum (Fig 2 C1, C2 and D1,D2) and overlap with the distribution of MCM7-immunolabeled cells.
Musashi1-immunoreactive cells decrease in number between stages 46 and 49 from 158.3±8.2
cells/20,0003 µm to 53±4.3 cells/20,0003 µm (p< 0.05, n=4, 4), consistent with the decrease in XdUand MCM7- labeled cells. These data suggest that the pool of progenitor cells gradually decreases between
stages 46 and 49.
Musashi-expressing cells are radial glial progenitor cells
To test whether the musashi1-immunoreactive cells are neural progenitors in the tadpole CNS, we
exposed stage 48 tadpoles to XdU for 2 hr and fixed half of them immediately to analyze the distribution
of XdU labeling and musashi1-immunoreactive cells. The remaining tadpoles survived for an additional
72 hours in the absence of XdU before fixation at stage 49. The majority of the cells labeled with a 2 hr
exposure to XdU are located in the ventricular layer and label with antibodies to musashi1 (Fig 3 A1-4),
indicating that musashi1-immunoreactive cells are progenitors. When XdU-labeled animals survived for
an additional 72 hr (in the absence of further XdU exposure) the number of XdU-labeled cells increased (
Fig 3 B1-4), indicating that the XdU-labeled neural progenitor cells continue to proliferate. Furthermore,
after 72 hr, the XdU-labeled cells were distributed within the tectal cell body layer where mature tectal
neurons are located (Fig 3B4).
Studies in mammalian cortex indicate that radial glia are neural progenitors (Kriegstein and AlvarezBuylla, 2009), however the potential role of radial glial cells as neural progenitors in midbrain subcortical
structures has not been established. We find that musashi1-immunoreactive cells in the tadpole optic
tectum extend a radial process to the pia, indicating that they have radial glial morphology (
Fig 3 C1 and C3). To further characterize the musashi1-immunoreactive cells, we labeled radial glial cells
in stage 47 tadpoles by bulk electroporation of a CMV::eGFP expression plasmid into ventricular layer
cells (Haas et al., 2002). The day after electroporation, eGFP is expressed in cells with radial glial
morphology lining the ventricle (Haas et al., 2002; Tremblay et al., 2009). The majority of eGFPexpressing radial glial cells in stage 47 optic tectum are musashi1-immunoreactive (Fig 3D,E). These
results show that a 2 hr exposure to XdU in vivo labels musashi1-immunoreactive neural progenitors and
that musashi1-immunoreactive cells are radial glia.
Visual deprivation increases cell proliferation in the optic tectum
The developmental decrease in progenitor cell proliferation correlates with the maturation of the
functional visual system in Xenopus tadpoles and suggests that visual circuit function may negatively
regulate progenitor cell activity. To test whether visual experience regulates cell proliferation in the
tadpole optic tectum, animals were reared in a 12 hr light/12 dark cycle until stage 46 when rates of cell
proliferation are still relatively high (Fig 1). Tadpoles were separated into 3 groups: One group continued
under the normal 12 hr light/12 dark cycle, called ‘ambient light’. The second group was provided with
enhanced visual stimulation (from an array of LEDs flashing on and off at 1Hz (Sin et al., 2002)) during
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the 12 hr light period of the light/dark cycle and the third group was deprived of visual stimulation (by
keeping them in the dark, see diagram in Fig 4A). All tadpoles were kept in the dark during the 12 hr dark
period of light/dark cycle.
We determined the division history of proliferating cells in the optic tectum using an assay that is based
on a double-label protocol in which animals were exposed to two deoxyuridine analogs, CldU and IdU, at
an interval longer than the cell cycle to identify cells which remain proliferative (Encinas and Enikolopov,
2008; Vega and Peterson, 2005) as shown in Fig 4B. CldU and IdU can be identified by immunostaining
in Xenopus optic tectum without significant cross-reactivity or labeling bias (Supplemental Figure S1).
We refer to CldU and IdU generically as XdU. We provided a 2-hour exposure to the first deoxyuridine
analog X1dU (either CldU or IdU; green in Fig 4B) and allowed tadpoles to grow in the absence of X1dU
label for 24 hr, followed by a 2 hr exposure to the second deoxyuridine analog X2dU (either CldU or IdU;
red in Fig 4B). We used a 24h interval in between the X1dU and X2dU exposures based on the observation
that the number of XdU-labeled cells approximately doubled over 24 hr (Fig 1I). The optic tectum was
then analyzed for the presence of X1dU and X2dU labeling in 30 µm cryostat sections. Three labeling
combinations are predicted: 1. cells with only X1dU (green in Fig 4B) 2. cells with only X2dU label (red in
fig 4B) and 3. cells labeled with both XdU's (yellow in Fig 4B). The number of cells labeled with X1dU
(green) relative to the total number of labeled cells is the fraction that were in S phase at the first XdU
exposure, but were not in S phase at the time of exposure to X2dU. This value gives an estimate of the
cells that exit the cell cycle between the times of exposure to X1dU and X2dU. The number of cells labeled
with X2dU (red) relative to the total number of labeled cells is the fraction that was in S phase only at the
second XdU exposure but not during exposure to X1dU. This value would represent a potential
recruitment of quiescent progenitors to proliferate. The number of cells labeled with both XdUs (yellow)
relative to the total number of labeled cells is the fraction of cells that were in S phase during both XdU
exposure periods and therefore provides an estimate of the population of progenitors that remained
proliferative during the observation window. Although this protocol does not identify all cells that are
proliferating or all cells that remain in the cell cycle over the 24 hr interval (Encinas and Enikolopov,
2008; Vega and Peterson, 2005), it allows us to compare cell proliferation and cell cycle parameters in
tadpoles reared in a normal 12 hr light/12 dark cycle (referred to as ambient light) with animals provided
with either enhanced visual stimulation or deprived of visual stimulation.
We counted XdU-labeled cells along the midline ventricular layer in sections through the optic tectum
and found that tadpoles subjected to reduced visual experience had more XdU-labeled cells in the same
volume (79 ± 7.6 cells/20,000µm3, n=5) compared to animals that were exposed to either ambient light
(48.7 ± 5.0 cells/20,000µm3, n=5; p<0.05) or enhanced visual stimulation (44 ± 7.3 cells/20,000µm3,
n=5; p<0.05; Fig 4 C, D and E). Similar indices of XdU-labeled cells were seen in the optic tecta of
tadpoles exposed to either enhanced visual activity or ambient light. These data suggest that visual
deprivation for 2 days increases cell proliferation in the optic tectum. They further suggest that the
amount of visual activity provided by ambient light is sufficient to reduce cell proliferation in the tectum.
Next, we determined the fraction of X1dU-labeled cells that also incorporated X2dU, as an estimate of the
cell population that continued to proliferate over the 24h period. Animals deprived of visual experience
have a significantly larger fraction of X1dU and X2dU double-labeled cells (82.1 ± 1.9%) compared to
tadpoles exposed to either ambient light (62.4 ± 1.6%, p<0.05) or enhanced visual stimulation (70.9 ±
3.5%; p<0.05; Fig 4 F). A smaller proportion of double-labeled cells in animals with visual experience
indicates that relatively fewer X1dU-labeled cells were in S phase of the cell cycle at the time of exposure
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to the second label, X2dU. Furthermore, visual experience results in a higher fraction of cells labeled with
only X1dU compared to visual deprivation (Fig 4 G) consistent the idea that visual experience changes the
fate of tectal progenitors so they exit the cell cycle or become quiescent. Together, these data indicate that
visual system activity decreases proliferative activity, while reduced visual experience maintains cells in a
proliferative state by decreasing cell cycle exit and maintaining cells in the progenitor pool at the expense
of differentiation.
Visual experience increases neuronal differentiation
The data presented above suggest that visual experience may change the fate of the progeny of tectal
progenitor cells so they differentiate into neurons. To test whether visual experience promotes neuronal
differentiation of newly generated cells, animals were reared in their normal 12h light/12h dark
conditions until stage 47 and exposed to XdU for 2h in rearing solution. Animals were then either
deprived of visual experience for 48h or exposed to enhanced visual stimulation as described in Fig 5C.
We used N-β-tubulin antibodies to label differentiated neurons (Moody et al., 1996). Visual experience
increases the proportion of XdU-labeled cells that differentiated into N-β-tubulin-labeled neurons over
the intervening 48h compared to that seen in visually-deprived animals (61.7 ± 1.5% vs 40.2 ± 1.1%,
p<0.05, n=11,11 animals, respectively). These results indicate that visual experience changes the fate of
progeny to exit the cell cycle and differentiate into neurons.
Visual deprivation expands the neural progenitor cell population by increasing Musashi1 expression
The reduced proliferation seen in animals with visual experience could result from two types of
mechanisms in relation to neural progenitor cells. The number of neural progenitor cells could remain
constant while the cell divisions occur less frequently or the size of neural progenitor pool could decrease
with visual system activity. Either mechanism would result in a decreased fraction of X1dU and X2dU
double-labeled cells and a reciprocal increase in the fraction of cells that stop dividing after X1dU
exposure. To differentiate between these two possibilities we tested whether the musashi1-expressing
neural progenitor cell population is affected by visual experience. Animals were reared under normal
conditions to stage 47 when they were either deprived of visual experience or provided with visual
experience, as shown in Figure 5C. The number of musashi1-expressing cells was significantly lower in
tecta of animals with visual experience (267.2 ± 15.3 cells/20,000µm3, n=7) compared to tecta of
visually-deprived animals (426.9 ± 32.8 cells/20,000µm3, n=7; p<0.05) (Fig 6 A, B and C). These results
are consistent with a model in which the decrease in cell proliferation seen with visual stimulation results
from a reduction in number of musashi1-expressing neural progenitor cells, whereas visual deprivation
increases the pool of musashi1-expressing neural progenitor cells.
The experiments described above indicate that musashi1 expression correlates with the proliferative
activity of tectal neural progenitor cells. To test whether musashi1 is required for the increase in cell
proliferation in visually-deprived animals, shown in Figure 4, we knocked down nrp1B, the Xenopus
homolog of musashi1 using morpholino antisense oligonucleotides to nrp1B. To test the efficacy of
knockdown, morpholinos against nrp1B or control scrambled morpholinos were electroporated into the
right tectal lobe. After 48 hours, we compared the intensity of musashi1 immunoreactivity in ventricular
layer cells of the right and left optic tecta. Morpholinos against nrp1B reduced the ratio of musashi1
immunoreactivity in the electroporated right tectum to the unelectroporated left tectum to 57.2 ± 0.03%
(p<0.05), whereas control morpholinos did not show any significant difference in musashi1
immunoreactivity between right and left tecta after unilateral electroporation (Supplementary Data
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Figure 2). We then electroporated stage 46 tadpoles in both tectal lobes with nrp1B morpholinos or
control morpholinos and subjected animals to reduced visual stimulation for 60 hrs followed by 2hr
exposure to XdU. Morpholinos against nrp1B significantly reduced the number of XdU-labeled cells
(36.93 ± 1.94 cells/20,000µm3, n= 14; Fig 6 E,G;) compared to control morpholinos (52.8 ± 1.7 cells, n=
16; p<0.05; Fig 6 D,G). Furthermore, coelectroporation of plasmid containing mouse musashi1 cDNA
with morpholinos directed against Xenopus nrp1B mRNA completely rescued the morpholino-induced
reduction in proliferation (62.5 ± 4.0 cells/20,000µm3, n= 11; p<0.05; Fig 6 F, G). These data indicate
that musashi1 protein levels are negatively regulated by visual activity in the optic tectum and that
musashi1 is necessary for the increased cell proliferation seen in visually-deprived animals. Together with
data presented in Figure 5, the data show that progenitors increase musashi expression, increase
proliferative activity and expand the progenitor pool in the absence of visual input but that visual
experience triggers two changes in the system: 1. a decrease in musashi expression and a decrease in
proliferative activity in radial glial cells and 2. an increase in the rate at which newly generated progeny
differentiate into neurons.
To test whether musashi1 is sufficient to increase cell proliferation in the CNS we returned to stage 49
tadpoles, where musashi1 expression is relatively low in ventricular layer cells (Figure 2). Electroporation
of ventricular layer cells in stage 49 tadpoles with a dual promoter plasmid co-expressing mouse
musashi1 and eGFP or eGFP alone labeled radial glial cells (Figure 7A,B). In vivo time-lapse images of
eGFP+ cells co-expressing mouse musashi1 show a greater increase in eGFP-expressing cells over 6 days
than seen in control animals, suggesting that exogenous expression of musashi1 increases proliferation of
radial glia. Furthermore, we tested whether musashi1 expression in stage 49 tadpoles increases XdU
incorporation. Stage 49 tadpoles were electroporated with a dual promoter plasmid co-expressing either
eGFP alone or mouse musashi1 and eGFP and after 60 hr, animals were exposed to XdU for 2 hr
immediately before sacrifice. Mouse musashi1 expression doubled the number of XdU-labeled cells
compared to expression of eGFP alone (Musashi1: 21.3±1.3 cells/20,000µm3, n=17, eGFP: 10.9±0.9
cells/20,000µm3, n=18; p< 0.05).
Discussion
Rates of neurogenesis decrease in the developing CNS as neuronal circuits become functional, suggesting
that neuronal activity generated by developing circuits may regulate the addition of new cells to the
circuit. We have used quantitative analysis of cell proliferation in the developing visual system of tadpoles
to show that rates of cell proliferation decrease as the visual circuit matures. These results demonstrate a
temporal correlation between the establishment of functional circuits and a decrease in cell proliferation
and suggest the presence of a negative feedback mechanism from the developing circuit to neural
progenitors to limit their proliferation. Consistent with this idea, we demonstrate that sensory input
activity decreases cell proliferation in the optic tectum and increases the rate at which newly generated
cells differentiate into neurons. We provide evidence that immunoreactivity to musashi1, a highly
conserved mRNA binding protein which is required to maintain progenitor activity across phyla (Okano
et al., 2005), is down-regulated during the developmental period when cell proliferation decreases and
the normal developmental down regulation does not occur when animals are deprived of visual
experience for 2 days. We show that knockdown of nrp1B, the Xenopus homolog of musashi1 prevents the
visual-deprivation induced increase in cell proliferation. Finally, we report that expression of mouse
musashi1 increases rates of cell proliferation when endogenous protein has been knocked down by
morpholinos or has been developmentally down-regulated. These data indicate that the control of neural
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progenitor proliferation and cell fate during CNS development is regulated by neuronal activity such that
increased activity from developing circuits changes the fate of newly generated cells so they exit the cell
cycle and differentiate into neurons. The data further suggest that musashi1 expression is regulated by
activity in the intact CNS and that activity-dependent downregulation of musashi1 expression decreases
cell proliferation and increases neuronal differentiation in the developing CNS. Finally, the data indicate
that neural progenitor proliferative activity can be dynamically regulated by increasing or decreasing
musashi1 expression, either by modulating brain activity or by more direct manipulation of musashi1
expression levels.
Tectal neural progenitor cells are radial glia
We show that musashi1-expressing neural progenitor cells in tadpole optic tectum are radial glial cells
and that they generate tectal neurons. Neural progenitor cells with radial glial morphology have been
reported in cortical regions of many vertebrates, suggesting that radial glial neural progenitors are highly
conserved morphologically and functionally in cortical development (Kriegstein and Alvarez-Buylla,
2009), however the role of radial glial cells as neural progenitors in subcortical brain regions has not been
clearly demonstrated. The decrease in cell proliferation and musashi1 immunoreactivity between stages
46 and 49 and with enhanced visual experience suggest that the progenitor pool is depleted in response to
activity-dependent signals. It is interesting to note that electroporation of ventricular layer cells in stage
49 tadpoles with a CMV::eGFP expression plasmid labels radial glial cells, showing that radial glial cells
persist at these stages, but they have lower proliferative activity and low musashi1 expression. Expression
of mouse musashi1 in radial glial cells of stage 49 tadpoles increases XdU incorporation and increased the
number of eGFP-expressing cells over a 6 day period of in vivo imaging. This experiment suggests that the
normal experience-dependent developmental decrease in expression of musashi1 in radial glial cells
decreases their proliferative activity, but that mechanisms that increase musashi1 expression in radial
glia, either through a decrease in sensory input or by exogenous expression of musashi1, can increase
proliferation in these relatively quiescent progenitors.
The increased XdU incorporation seen in stage 46 animals compared to stage 49 animals could indicate
that S phase of the cell cycle is longer in younger animals. Similarly, the differences in XdU-incorporation
in animals in which musashi expression is decreased by knockdown or increased by exogenous expression
of mouse musashi, could arise from corresponding changes in the length of S phase. To attempt to
address this possibility, we labeled tecta with antibodies to phospho-histone 3 (PH3) to determine the
mitotic index. We found that PH3 labels 2-6 cells/20,000µm3 along the tectal midline, but the labeling is
too sparse to test for statistically significant differences across stages (data not shown). Although we
cannot strictly conclude that the mitotic index does not change with developmental stage and musashi1
expression, our data nevertheless provide evidence for regulation of neurogenesis by visual experience
and musashi1 expression in radial glial progenitors.
Control of neurogenesis and brain size by an interplay of ‘intrinsic’ cell autonomous and ‘extrinsic’ noncell autonomous mechanisms
Regulation of neurogenesis establishes the number, type and distribution of cells in the CNS by regulation
of progenitor maintenance and proliferation, and the subsequent survival and differentiation of progeny
into neurons or glia. Controlling the size of the progenitor pool by regulating the fate of progenitors and
their progeny to either die, differentiate or continue to proliferate is a key regulatory event in brain
development and ultimately in the establishment of functional circuits and behavior. For instance,
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maintaining progenitors in an actively proliferative state enlarges the progenitor pool and can enlarge the
cerebral cortex of mice (Chenn and Walsh, 2002) by setting an upper bound on the potential number of
neurons generated (Lehmann et al., 2005). Conversely, premature transition of progenitors from an
active proliferative state to differentiated cells can deplete the progenitor pool, while increasing or
decreasing progenitor survival controls cortical size (Depaepe et al., 2005; Putz et al., 2005). A variety of
extrinsic factors can affect the fate of neural progenitor progeny by either maintaining progenitor fate or
promoting differentiation (Ninkovic and Gotz, 2007), including neurotransmitters (Gandhi et al., 2008;
LoTurco et al., 1995; Sadikot et al., 1998; Spitzer, 2006), secreted peptides/proteins (Botia et al., 2007;
Suh et al., 2001; Wexler et al., 2009), and transmembrane or GPI-linked proteins (Depaepe et al., 2005;
Putz et al., 2005; Qiu et al., 2008). Despite the plethora of possible extrinsic signals, whether single
signals or combinations of signals operate downstream of visual system activity to decrease rates of cell
proliferation during CNS development is not clear.
Extrinsic signals must be read and interpreted by intrinsic cell autonomous mechanisms to culminate in
cell fate regulation. Our data indicate that sensory input activity depletes the neural progenitor
population by down-regulating musashi1, suggesting that musashi1 is a prominent ‘intrinsic’
maintenance/differentiation switch which can respond to non-cell autonomous signals from developing
brain circuits to mediate feedback from the developing circuit and control cell proliferation. Musashi1 is
thought to function as a translational repressor. Using genomic and proteomic approaches, musashi1 was
found to affect the expression of a large number of targets involved in the cell cycle, apoptosis, cell
proliferation and cell differentiation (de Sousa Abreu et al., 2009). Musashi1 may promote maintenance
of neural progenitor cells by blocking translation of Numb, a repressor of Notch signaling (Imai et al.,
2001). Activation of the Notch signaling pathway in turn promotes the maintenance of neural progenitor
cells (Hitoshi et al., 2002; Tokunaga et al., 2004). In addition, musashi1 represses expression of p21WAF1, a cyclin-dependent kinase inhibitor, which causes exit from the cell cycle (Battelli et al., 2006). A third
target of musashi1 repression is doublecortin, a microtubule associated protein that is expressed at early
stages of neuronal differentiation and is important for neuronal migration, differentiation and plasticity
(Horisawa et al., 2009). Although considerable evidence suggests that musashi1 maintains neural
progenitors in a proliferative state by repressing translation of proteins required for cells to exit the cell
cycle and differentiate, it is not clear whether a single gate keeper integrates multiple extrinsic signals to
control cell fate (Suh et al., 2009). In addition to musashi1, other proteins including musashi2, Sox2 and
PTEN promote neuronal progenitor cell self-renewal and decrease differentiation (Graham et al., 2003;
Groszer et al., 2006; Ohtsuka et al., 2001; Sakakibara et al., 2002). Activity-dependent regulation of
musashi1, musashi2, Sox2 or PTEN has not been reported and would be an interesting topic to pursue.
How signals from developing circuits regulate intracellular signaling pathways controlling cell fate of
radial glial progenitors is not clear. Calcium transients downstream of neurotransmitter receptor activity
(LoTurco et al., 1995) or bioactive peptides (Botia et al., 2007) may cause progenitors to exit the cell cycle
and differentiate. A recent study demonstrated that Xenopus tadpole radial glial cell bodies exhibit slow
calcium transients in response to visual stimulation (Tremblay et al., 2009). The same study used timelapse in vivo imaging to show that radial glial processes near the pia are dynamic and that visual
stimulation increases radial glial structural dynamics by a mechanism that includes NMDA receptormediated NO signaling. This is interesting because previous studies in Xenopus tadpoles indicated that
NOS-expressing neurons are positioned immediately adjacent to the radial glial cell bodies, and that
decreasing NO signaling increases cell proliferation in the tadpole CNS and deceases cell motility
(Peunova et al., 2001; Peunova et al., 2007), suggesting that cells temporarily stop exploratory structural
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rearrangements when they devote their cytoskeletal resources to divide. Together these studies suggest
that visual system activity negatively regulates radial glial cell proliferation in the tadpole optic tectum by
a signaling pathway that includes an NMDA receptor-mediated increase in NO, which decreases
proliferation in tectal progenitors.
Role of activity in neurogenesis in the developing and adult CNS
It is widely recognized that neurogenesis continues in the adult brain and that exposing animals to
enriched environments or exercise increases adult neurogenesis, presumably through activity-induced
increases in trophic factors or neurotransmitters (Brown et al., 2003; Rochefort et al., 2002; Whitman
and Greer, 2009; Zhao et al., 2008). Further investigations to distinguish whether the increased
neurogenesis in adult CNS is the outcome of increased cell proliferation, differentiation or survival of
differentiated progeny suggest that, even though exercise may increase rates of proliferation in the
hippocampal dentate gyrus (Brown et al., 2003), it appears that the more prevalent effect of increased
brain activity in adult animals is to increase the survival of differentiated neurons in target brain regions
(Whitman and Greer, 2009; Zhao et al., 2008). By contrast, we find that visual experience reduces rates
of cell proliferation, increases cell cycle exit and increases neuronal differentiation of newly generated
cells, but tunel staining suggests that cell death is not a significant factor in regulating neurogenesis in the
developing optic tectum (Peunova et al., 2001), at least over the time-course of our studies. These data
indicate that visual experience limits self-renewal of neural progenitor cells, thereby depleting the
progenitor pool, and shifts the fate of progeny to terminal neuronal differentiation. These differences
likely reflect distinct functions of neurogenesis in the adult and developing CNS. During CNS
development the function of neurogenesis is to generate a large population of neurons in a relatively finite
time period so that functional neural circuits can be assembled de novo. By contrast, adult-generated
neurons integrate into pre-existing neural circuits where they are thought to modulate learning and
memory (Whitman and Greer, 2009). Therefore, different responses to activity-dependent signals in
neural progenitor cells in the developing and adult CNS likely reflect the requirements of the CNS for the
generation and maintenance of new neurons at different stages of life.
Material and Methods
Animals
Albino Xenopus laevis tadpoles, obtained from our lab colony or commercial sources (Nasco, Fort
Atkinson, WI), were reared in a 12 hr light/12 hr dark cycle incubator at 24°C. Animals were staged
according to Nieuwkoop and Faber (Nieuwkoop and Faber, 1956).
Reduced and enhanced visual stimulation protocol
Tadpoles were deprived of visual experience by placing them in a black plastic box. Enhanced visual
stimulation was applied by placing tadpoles in a box containing an array of light emitting diodes (LEDs)
flashing on and off at 1 Hz to create a simulated motion stimulus, as described (Sin et al., 2002). In all the
experimental conditions tadpoles were reared in a 24°C incubator. The time-course of the reduced or
enhanced visual experience protocol is shown in Figure 4A.
Immunohistochemistry
Xenopus tadpoles were anesthetized with 0.02% MS222 (3-aminobenzoic acid ethyl ester), immersed in
4% paraformaldehyde (Electron Microscopy Sciences, Fort Washington, PA) in PBS and microwaved for
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8 sec (750 W GE microwave) followed by 2h in fixative at room temperature (Li et al., 2010). After rinsing
the animals, their brains were dissected and either analyzed in wholemount, were cryoprotected in 30%
sucrose and cut into 30 µm horizontal sections with a cryostat, or embedded in gelatin and cut into 30 µm
horizontal sections with a vibratome. Sections were blocked in 5% goat serum and 0.3% Tween 20 in PBS
for 1 hr before incubating overnight at 4°C with anti-MCM7 (1:250; Abcam, Cambridge, MA), antimusashi1 (1:250; Abcam, Cambridge, MA) or anti-β-tubulin I+II (1:200; Sigma-Aldrich, St. Louis, MO)
antibodies. Detection was performed using appropriate fluorophore-tagged secondary antibodies
(Molecular Probes/Invitrogen, Eugene, OR). Sections or brains were mounted in ProLong™ Gold
(Molecular Probes/Invitrogen, Eugene, OR) and imaged with a Zeiss LSM 510 Meta confocal microscope
(Carl Zeiss, Jena, Germany). When levels of labeling were compared across samples, all samples were
prepared, photographed and analyzed in parallel using the same acquisition and analysis settings.
Labeling with XdU's
Tadpoles were exposed to the halogenated thymidine analogs, BrdU, IdU or CldU, referred to collectively
as XdU's (MP Biomedicals, Solon, OH) at 10 mM in Steinberg's solution (60 mM NaC1, 0.7 mM KCI, 0.8
mM MgSO4, 0.3 mM Ca(NO3)2 and 1.4 mM Tris, pH 7.4) for 2 hr. We found that delivery of XdU by
injection into the pericardial cavity or by 2hr exposure of tadpoles to XdU in rearing solution resulted in
comparable labeling of proliferative cells. We used exposure in rearing solution for XdU delivery in our
experiments because this method allows greater control of XdU delivery.
Analysis of history of cell division
Sequential exposure to the differentially halogenated thymidine analogs CldU and IdU was used to
determine the history of proliferative activity, as described (Encinas and Enikolopov, 2008; Vega and
Peterson, 2005). Tadpoles were exposed to IdU in Steinberg's solution for 2hr and transferred to fresh
Steinberg's solution. After 24 hrs they were exposed to CldU in Steinberg's solution for 2 hr, rinsed in
fresh Steinberg's solution, anesthetized with 0.02% MS222 and fixed as described above. For
wholemount analysis, dissected brains were treated with −20°C methanol for 1 hr followed by proteinase
K treatment as described (Peunova et al., 2001). For detection of IdU and CldU, wholemount brains or
sections were treated with 2N HCl for 1 hr at 37°C, rinsed in PBS and incubated in 5% normal goat serum
and 0.3% Tween 20 in PBS for 1 hr before incubating overnight at 4°C with 1:400 IdU specific antibody
(BD Biosciences mouse anti-BrdU Cat # 347580) and 1:400 CldU-specific antibody (Accurate rat antiBrdU Cat# OBT0030G). The IdU-specific antibody was detected using Alexa 488 labeled goat anti-mouse
secondary antibody (Molecular Probes/Invitrogen, Eugene, OR). The CldU-specific antibody signal was
enhanced using biotin tagged goat anti-Rat secondary antibody followed by detection with Cy5streptavidin (Jackson ImmunoResearch). To reduce crossreactivity, highly cross-adsorbed secondary
antibodies were used. Wholemounts and sections were mounted in ProLong™ Gold (Molecular
Probes/Invitrogen, Eugene, OR) and imaged with a Zeiss LSM 510 Meta confocal microscope (Carl Zeiss,
Jena, Germany). Images were collected using a Zeiss Fluor 20X 0.75 NA or CApochromat 40X 1.2 NA
water objective. No significant crossreactivity was observed for IdU and CldU (Supplementary Fig 1). To
test whether there is a labeling bias in our method for detecting IdU and CldU (i.e. whether we can detect
a cell in S-phase by IdU or CldU with equal probability) we exposed tadpoles to equimolar CldU and IdU
simultaneously for 2 hr. All the cells were double-labeled, indicating that we can detect S-phase cells in
tadpoles using IdU or CldU with equal probability (Supplementary Fig 1). We used CldU and IdU
interchangeably and obtained the similar results.
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Data Analysis
Image analysis and cell counting were performed using Metamorph (Universal Imaging
Corporation/Molecular Devices, Downingtown, PA) image processing software. Images were background
subtracted and cells were counted using the dissector method with the “manually count object” feature of
Metamorph. In wholemounts of brains, cells in the ventricular layer at the midline of the optic tectum in
an area 100 µm along the ventricular layer X 20µm lateral from the ventricular layer were counted in two
non-neighboring optical sections. The first optical section was selected where the two tectal lobes meet at
the dorsal midline and the second optical section was 8-10 µm ventral from the first. The average
diameter of cell nuclei is 5 µm and does not change with the different stages or treatments analyzed. The
rostral boundary for counting cells within the tectal ventricular layer was defined by the anterior dorsal
commissure and the caudal boundary was identified as the start of the curvature to the caudolateral edge
of the tectum, as shown in Figure 1. Cell counts from the 2 sections were added and presented as cell
number per 20,000 µm3. We used cryostat sections for high-resolution 40X analysis to determine the
presence of IdU only, CldU only or both IdU and CldU in cells. Two horizontal sections from each brain
corresponding to about 60-90 µm and 120-150 µm from the dorsal side respectively were used for
analysis. Cells were counted in the entire z stack through the section collected with 2 µm z-step size. The
rostral and caudal boundaries for counting cells were the same as for wholemounts.
Knockdown and over-expression of Musashi1
Lissamine-tagged morpholino antisense oligonucleotides against nrp-1B, the musashi1 homologue in
Xenopus laevis (GeneTools,Philomath, OR) with the sequence GCGCTTCTGTCTCCATTCGGTCTCT, or
the five basepair mismatched oligonucleotide with the sequence GCCCTTGTGTGTCCAATCCGTGTCT,
were electroporated into the optic tectum of stage 46 tadpoles as described (Bestman and Cline, 2008;
Chiu et al., 2008). The morpholino against nrp-1B does not recognize the xrp1, the Xenopus homologue
of musashi2, according to prediction algorithms by GeneTools (GeneTools,Philomath, OR). The tadpoles
were provided with visual experience or deprived of visual experience as described above. At specified
times after electroporation of morpholinos, tadpoles were exposed to XdU for 2hr in Steinberg's solution,
fixed and analyzed in wholemounts as described above.
Mouse musashi1 cDNA was generously provided by Dr. Imai and Dr. Okano as pcDNA-Flag-Musashi1
(Imai et al., 2001). Mouse musashi1 was subcloned into a dual CMV promoter vector (Bestman and Cline,
2008) to co-express eGFP and musashi1. Control eGFP expression was from the dual promoter construct
with no construct in the second site. Plasmids were electroporated into the optic tectum of anesthetized
tadpoles as described (Bestman and Cline, 2008; Chiu et al., 2008).
In vivo time-lapse imaging
Animals were screened for eGFP expression on a fluorescence microscope and imaged on a spinning disk
confocal microscope (Perkin Elmer) with a 20X water immersion lens (0.9 NA), as described (Ruthazer
and Cline, 2004).
Statistical tests
Data were tested with the non-parametric Mann–Whitney–Wilcoxon test to compare between groups
unless stated otherwise. Data are represented as mean ± SEM.
Supplementary Material
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01
02
03
04
Acknowledgements
this work was supported by the NIH (DP10D000458 to HTC), Dart Neuroscience, LLC (HTC) and
fellowships from the Cold Spring Harbor Laboratory Association and CIRM to PS. We thank Drs. Imai
and Okano for generously providing the cDNA for mouse Musashi1 and members of the Cline lab for
insightful discussions.
Footnotes
Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service
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Figures and Tables
Figure 1
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Developmental decrease in proliferative cells in the Xenopus tadpole optic tectum. A-C. The Xenopus optic tectum
includes a functional visual circuit and proliferative ventricular layer cells. A. Phase contrast image of an albino Xenopus
tadpole head. Retinal ganglion cells axons project via the optic nerve (in red) to the contralateral optic tectum. B.
Midbrain of Xenopus laevis showing the optic tectal lobes (ot). C. Cartoon of the cell types in Xenopus optic tectum.
Afferent axons from retinal ganglion cells (red) form synapses with dendrites of tectal neurons (green) in the lateral
neuropil. Tectal cell bodies are located medially from the retinotectal neuropil. Radial glial cells line the ventricle (blue).
D-G. Proliferation in the optic tectum. Stage 46 (D and E) and 49 (F and G) tadpoles were labeled with a 2-hr exposure to
XdU and fixed immediately thereafter. D, F. Images of 3-D merge of a z series of horizontal confocal sections through the
midbrain. E, G. Images of a series of single optical sections every 10 µm from dorsal to ventral tectum. The proliferating
cells labeled by XdU incorporation line the ventricle and decrease significantly between stage 46 and stage 49. XdUlabeled cells were counted in the region marked by the bracket in D. H, I. Quantitative analysis of numbers of XdU-labeled
cells in the midline ventricular layer of stage 46, 48 and 49 tadpoles (n=8,8 and 7 animals respectively) which were
exposed to XdU for 2 hr, then fixed and analyzed either immediately (H) or after further rearing in the absence of XdU for
22 hr (I) as shown in the schematic on top of each graph. XdU incorporation decreases significantly from stage 46 to stage
49. Scale bar = 50 µm in F, also applies to D. *** p<0.0001.
Figure 2
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Developmental decrease in MCM7 and Musashi1 immunoreactivity. Confocal images of 30 µm cryostat sections through
the optic tectum of stages 46 (A, C) and 49 (B, D) tadpoles labeled with anti-MCM7 antibody (green; A1, B1) or antimusashi1 antibody (C1, D1) and a nuclear stain SytoxO (blue; A2-D2). Cells located along the ventricular layer are highly
immunoreactive for MCM7 and musashi1. E. Quantitative analysis of the decrease in number of Musashi1
immunoreactive cells between stages 46 and 49. Scale bar = 20 µm. *p<0.05.
Figure 3
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Lineage and identity of proliferating neural progenitor cells in optic tectum. A-B. Stage 48 tadpoles were exposed to XdU
for 2h in rearing solution and were either fixed immediately (A) or after 72 hrs of further development in the absence of
XdU (B). Confocal images of 30 µm cryostat sections through the optic tectum of tadpoles labeled with antibodies to
musashi1 (msi1; green) and XdU (red). Nuclei were stained with SytoxO (blue). Musashi1-immunoreactive cells line the
ventricle and are labeled by incorporation of XdU with a 2h survival time (A). The progeny of XdU-labeled neural
progenitor cells are distributed throughout the cell body layer of the optic tectum after a 72 hr survival time (B). Note that
the images of musashi1-immunoreactivity in B1 were taken with longer exposure periods than the images in A1, according
to the lower levels of musashi1-immunoreactivity in ventricular layer cells at stage 49 (see Fig 2C,D). C. Musashi1immunoreactive cells have radial processes extending to the pia. D,E. Radial glial cells, visualized by expression of eGFP,
are immunoreactive for musashi1. Stage 47 tadpoles were electroporated with an eGFP expression plasmid, fixed after 24
hrs and analyzed for Musashi1 immunoreactivity in 30 µm vibratome sections. D1-D3. 3-D merge of a z series containing
a complete eGFP-expressing cell with radial glial morphology (green; D1,D3) and Musashi1 immunolabeling (red; D2,
D3). E1-E3. A single optical section through the ventricular cell region showing eGFP (E1, E3) colocalization with
Musashi1 immunolabeling (E2, E3). Scale bar in D3 = 10 µm and E3 = 5 µm.
Figure 4
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Visual deprivation increases cell proliferation in the optic tectum. A. Timeline of the experimental protocol. The 12
dark/12h light rearing conditions are shown as shaded and white bars. During the 12h ‘light’ period animals were either
exposed to normal ambient light, enhanced visual stimulation or reduced visual stimulation (see methods). Animals were
exposed to X1dU for 2h and allowed to develop in the absence of X1dU for 24 hours, after which they were exposed to
X2dU at the times marked by the green and red arrows, respectively. This protocol provides an estimate of the number of
proliferating cells (labeled with X1dU and/or X2dU), the fraction of X1dU-labeled cells that remain in the cell cycle
(double labeled with X1dU and X2dU), and the fraction of X1dU-labeled cells that exit the cell cycle (X1dU only). B. A
cartoon of representative results of the assay to detect the division history of proliferating cells as labeled in A. Cells
labeled in green incorporated X1dU only. Cells labeled in red incorporated X2dU only. Cells labeled in yellow were in S
phase during exposure to X1dU and again during exposure to X2dU. C, D. Images of the dorsal midline ventricular layer of
the optic tectum of tadpoles with enhanced visual experience (C1-C4) or reduced visual experience (D1-D4) labeled with
anti-X1dU (green; C2, D2) and anti-X2dU (red; C3,D3). C1, D1. Images of the X1dU and X2dU-labeled cells in merged
optical sections of the complete z-series through the depth of the 30 µm cryostat section. C2-C4 and D2-D4. Magnified
single optical sections from the z-series in C and G showing a fraction of cells labeled with X1dU only, X2dU only and both
X1dU and X2dU. Scale bar = 10 µm for C2-4 and D2-4. Scale bar in D4 corresponds to 50 µm in C1 and D1 .E-G. Graphs of
total numbers of XdU-labeled cells (E), fraction of X1dU- and X2dU-labeled cells (F) fraction of X1dU-only labeled cells
(G). * p<0.05, ** p < 0.001 compared to animals with reduced visual stimulation.
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Figure 5
Visual experience increases neuronal differentiation. Stage 47 tadpoles were exposed to XdU for 2hr and allowed to
develop in absence of XdU for 48 hrs while subjected to either enhanced visual stimulation or reduced visual stimulation
as described in panel C. A, B. Images of single optical sections of N-β-tubulin (green; A1, A3, B1 and B3) and XdU (red;
A2, A3, B2 and B3) immunoreactivity in 30 µm vibratome sections from animals subjected to enhanced visual stimulation
(A) or reduced visual stimulation (B). C. Timeline of the experimental protocol. D. Percent of XdU+ cells that are doublelabeled with N- β-tubulin antibody. Enhanced visual stimulation increases the proportion of newly generated cells that
differentiate into neurons. Scale bar in A3 = 10 µm. ***p<0.0001
Figure 6
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Musashi1 is required for visual experience-dependent regulation of cell proliferation. A-C. Visual deprivation for 48h
increases the number of Musashi1 (Msi1) immunoreactive neural progenitor cells compared to animals with enhanced
visual experience. Images of Musashi1 immunoreactivity in 30 µm cryostat sections (A1,B1; green in A3, B3) and SytoxO
(A2, B2; blue in A3, B3) and merged images (A3, B3). C. Visual deprivation significantly increases the number of cells
with detectable musashi1-immunoreactivity compared to visual stimulation (n=5 each). D-G. Morpholino-mediated
knockdown of nrp1B, the Xenopus homolog of musashi1, decreases cell proliferation in the optic tectum while expression
of mouse Musashi1 rescues cell proliferation in animals with nrp1B knockdown. Both tectal lobes of stage 46 tadpoles
were co-electroporated with expression constructs and morpholinos. Animals were deprived of visual stimulation for the
next 60h and then were exposed to XdU for 2h immediately before fixation. D-F. Optical sections showing XdU-labeled
cells in animals coelectroporated with the control eGFP expression plasmid (ContVec) and control scrambled morpholino
(ContMO) (D), the control plasmid and morpholinos to nrp1B (Msi1MO) (E), or a plasmid expressing mouse musashi1
(mMsi1) and morpholinos to nrp1B (F). Morpholinos against nrp1B significantly decreased the number of XdU-labeled
cells in the ventricular layer compared to control morpholinos (D,E,). The nrp1B morpholino-mediated decrease in
proliferation was completely rescued by simultaneous expression of mouse musashi1 (F). G. Quantification of numbers of
XdU-labeled cells in animals treated as in D-F. Scale bar in B3 = 10 µm, E= 20 µm.*** p<0.0001.
Figure 7
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Musashi1 expression is sufficient to increase cell proliferation. A.B. In vivo images of eGFP-expressing cells collected over
6 days after electroporation of stage 49 tadpoles with a dual promoter plasmid expressing eGFP alone (A) or eGFP and
mouse Musashi1 (mMsi1; B). Each image is a 3-D merge of a z-series through the optic tectum taken 2 days (d2, A1, B1), 4
days (d4, A2, B2) and 6 days (d6, A3, B3) after electroporation. C,D. Musashi1 expression in stage 49 tadpoles increases
XdU incorporation. Animals were electroporated with plasmid expressing eGFP (C) or eGFP/mMsi1 (D) and 2 days later
were exposed to XdU for 2hrs. Whole mount brains were analysed for XdU immunolabeling. C,D shows single optical
sections of XdU labeled cells in eGFP- (C) and eGFP/mMsi1-expressing animals (D). E. Quantitative analysis of XdUlabeled cells with or without exogenous expression of mMsi1. Exogenous expression of mMsi1 at stage 49 increases
proliferation compared to control. Scale bar in B3 and D = 20 µm, ***p<0.0001
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