visual activity regulates neural progenitor cells in ... files/2010/pmc sharma neuron...

11/7/13 4:01 PMVisual Activity Regulates Neural Progenitor Cells in Developing Xenopus CNS through Musashi1

of 24http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3005332/

Neuron. Author manuscript; available in PMC 2011 November 4.Published in final edited form as:

Neuron. 2010 November 4; 68(3): 442–455.doi: 10.1016/j.neuron.2010.09.028

PMCID: PMC3005332NIHMSID: NIHMS241844

Visual Activity Regulates Neural Progenitor Cells in Developing Xenopus CNSthrough Musashi1

Pranav Sharma and Hollis T. Cline

The Scripps Research Institute, 10550 North Torrey Pines Road, La Jolla, CA 92037, USATo whom correspondence should be addressed. Email: [email protected]

Copyright notice and Disclaimer

Publisher's Disclaimer

Abstract

Regulation of progenitor cell fate determines the numbers of neurons in the developing brain. Whileproliferation of neural progenitors predominates during early CNS development, progenitor cell fateshifts toward differentiation as CNS circuits develop, suggesting that signals from developing circuits mayregulate proliferation and differentiation. We tested whether activity regulates neurogenesis in vivo in thedeveloping visual system of Xenopus tadpoles. Both cell proliferation and the number of musashi1-immunoreactive progenitors in the optic tectum decrease as visual system connections become stronger.Visual deprivation for 2 days increased proliferation of musashi1-immunoreactive radial glial progenitorswhile visual experience increased neuronal differentiation. Morpholino-mediated knockdown and over-expression of musashi1 indicate that musashi1 is necessary and sufficient for neural progenitorproliferation in the CNS. These data demonstrate a novel mechanism by which increased brain activity indeveloping circuits decreases cell proliferation and increases neuronal differentiation through the down-regulation of musashi1 in response to circuit activity.

Keywords: neurogenesis, visual experience, Xenopus, musashi1, radial glial cells, BrdU, progenitorpool, differentiation, cell cycle, N-β-tubulin, nrp1

Introduction

The number of neurons in the brain is largely determined by the regulation of neural progenitor cellproliferation and the survival and differentiation of their progeny. The pool of neural progenitor cells canbe expanded by symmetric divisions that give rise to two neural progenitor cells, maintained byasymmetric divisions that result in one of the progeny remaining a neural progenitor cell while the otherdifferentiates, or depleted by terminal differentiation (Butt et al., 2005; Gotz and Huttner, 2005; Huttnerand Kosodo, 2005; Kriegstein and Alvarez-Buylla, 2009; Noctor et al., 2007). While expansion andmaintenance 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

*

*

http://www.ncbi.nlm.nih.gov/entrez/eutils/elink.fcgi?dbfrom=pubmed&retmode=ref&cmd=prlinks&id=21040846

http://dx.doi.org/10.1016%2Fj.neuron.2010.09.028

http://www.ncbi.nlm.nih.gov/pubmed/?term=Sharma%20P%5Bauth%5D

http://www.ncbi.nlm.nih.gov/pubmed/?term=Cline%20HT%5Bauth%5D

mailto:[email protected]

http://www.ncbi.nlm.nih.gov/pmc/about/copyright.html

http://www.ncbi.nlm.nih.gov/About/disclaimer.html

http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3005332/#FN1

http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3005332/#R7







as CNS circuits develop (Kriegstein and Alvarez-Buylla, 2009), suggesting that signals from developingcircuits may shift the fate of neural progenitors and their progeny. Indeed, recent studies suggest thatneuron-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 activity-dependent regulation of progenitor cells in intact animals. Therefore, a fundamental open question in theregulation of neurogenesis is whether feedback mechanisms from developing neuronal circuits regulateprogenitor fate to expand, maintain or deplete the pool of neural progenitor cells in the developing CNSin vivo.

To address this question we tested whether visual activity affects the rate of ongoing cell proliferation inthe developing visual system of Xenopus laevis tadpoles, where cell proliferation in the optic tectumcontinues over an extended period of development while the visual circuitry is both functional and still inthe process of development (Cline, 2001; Peunova et al., 2001; Straznicky and Gaze, 1972). We find thatcell proliferation in the tectum, detected by BrdU incorporation, decreased as visual circuitry maturedbetween stages 46 and 49. Over the same period, immunoreactivity for MCM7, a marker of cells withproliferative potential (Crevel et al., 2007; Facoetti et al., 2006; Khalili et al., 2003), and musashi1, anRNA 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 inproliferation. These data are consistent with the idea that visual activity in the more mature circuit couldnegatively regulate cell proliferation. Indeed, brief visual deprivation for 2 days increased cellproliferation in the optic tectum compared to animals with visual experience, suggesting that feedbackfrom the developing visual circuit shifts the fate of neural progenitors. We used sequential exposure totwo differentially halogenated thymidine analogs (IdU and CldU, referred to collectively as XdUs) toreveal 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 cellcycle, whereas more cells exit the cell cycle and differentiate into neurons in animals with visualexperience. Interestingly, visually-deprived animals have more musashi1-immunoreactive radial glialprogenitors than animals with visual experience. Morpholino-mediated knockdown and rescueexperiments 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 detectableendogenous musashi1-immunoreactiviey and low proliferative activity, increases their proliferation. Ourstudy suggests that sensory experience plays a role in neurogenesis in the developing CNS in vivo byregulating 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 contralateraloptic tectum where they form synapses with tectal neurons (Fig. 1A-C). Between stages 39 and 49, aperiod of 6-7 days, the visual system of Xenopus tadpoles develops rapidly to accommodate thebehavioral needs of the animal. Retinal ganglion cells first innervate and transmit visual information tothe optic tectum at stage 39 (Holt and Harris, 1983) when the majority of cells in the tectum have radialglial morphology and neurons have very simple dendritic arbors (Wu et al., 1999). An initial topographicretinotectal map is established by stage 45 (O'Rourke and Fraser, 1990) and between stages 46 and 49visual experience drives many aspects of visual circuit development pertaining to the detection and

http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3005332/figure/F1/




















processing of visual inputs (Bestman and Cline, 2008; Chiu et al., 2008; Cline and Haas, 2008; Engert etal., 2002; Pratt and Aizenman, 2007, 2009; Pratt et al., 2008; Tao and Poo, 2005) even as ventricularlayer cells with radial glial morphology persist in the tectum (Tremblay et al., 2009). Although it is wellknown that tectal ventricular layer cells proliferate throughout tadpole stages of development andgenerate neurons within the tectum (Peunova et al., 2001; Straznicky and Gaze, 1972), a potential relationbetween 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 systemdevelopment, we exposed tadpoles at stage 46, 48 and 49 to 2 hr of XdU and either processed the brainsimmediately or allowed the animals to develop in normal rearing solution for another 22 hr beforeprocessing the brains as wholemounts for XdU immunodetection. We delivered XdU by exposingtadpoles to rearing solution containing 10 mM XdU for 2 hr. This method efficiently labels proliferativecells 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 Z-series of images was collected through the midbrain of wholemount brains or cryostat sections. Aspreviously reported, proliferating cells, identified by exposure to H-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 dissectormethod and indexed cell counts to an estimated volume of 20,000 µm (see Fig 1D and methods). Stage46 tadpoles have significantly more XdU-labeled cells (99.6 ± 2.4cells/20,000µm , n=8) than eitherstage 48 (36.1 ± 2.3 cells/20,000µm , n=8; p<0.05) or stage 49 (8.9 ± 0.8 cells/20,000µm , n=7;p<0.05) tadpoles (Fig 1H, Supplementary Movies S1, S2 and S3). The XdU-labeled cells continue todivide over the next 22 hr to approximately double the number of XdU-labeled cells (Fig 1I. Stage 46:180.4±4.9 cells/20,000µm , n=8; Stage 48: 71.2±3.9 cells/20,000µm , n=8; Stage 49: 20.3±0.9cells/20,000µm , n=7). These data suggest that proliferation gradually decreases between stages 46 and49, 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 proliferativecells in other experimental systems, change expression over the developmental period when XdUincorporation decreased. MCM7 is a part of minichromosome maintenance complex (MCM) of proteinsand is expressed in cells with proliferative potential (Crevel et al., 2007; Facoetti et al., 2006; Khalili etal., 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 labelthe 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 liningthe ventricle (Fig. 2 A1,A2 and B1, B2) and partially overlap with the distribution of cells detected byincorporation of XdU following 2h exposure. MCM7 expression was down-regulated between stage 46and 49 (Fig. 2 A1,A2 and B1, B2), consistent with the decrease in XdU-incorporation over this timeperiod.

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 Xenopusand named nrp1 (Richter et al., 1990). In vertebrates, there are 2 genes, musashi1 and musashi2, thelatter of which is homologous to xrp1 in Xenopus. We focused on Musashi1 because it is expressed

3

33

3 3

3 33






















http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3005332/#SD2















exclusively in the CNS and is enriched in CNS progenitor cells across phyla (Amato et al., 2005; Good etal., 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 expressedthroughout the body. Musashi1 protein expression is down-regulated in differentiated neurons (Kanekoet al., 2000; Nakamura et al., 1994; Sakakibara et al., 1996; Sakakibara and Okano, 1997). We find thatmusashi1-immunoreactive cells are present in the proliferative layer lining the ventricle of the optictectum (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±8.2cells/20,000 µm to 53±4.3 cells/20,000 µm (p< 0.05, n=4, 4), consistent with the decrease in XdU-and MCM7- labeled cells. These data suggest that the pool of progenitor cells gradually decreases betweenstages 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, weexposed stage 48 tadpoles to XdU for 2 hr and fixed half of them immediately to analyze the distributionof XdU labeling and musashi1-immunoreactive cells. The remaining tadpoles survived for an additional72 hours in the absence of XdU before fixation at stage 49. The majority of the cells labeled with a 2 hrexposure 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 foran 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 tectalneurons are located (Fig 3B4).

Studies in mammalian cortex indicate that radial glia are neural progenitors (Kriegstein and Alvarez-Buylla, 2009), however the potential role of radial glial cells as neural progenitors in midbrain subcorticalstructures has not been established. We find that musashi1-immunoreactive cells in the tadpole optictectum 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 cellsin stage 47 tadpoles by bulk electroporation of a CMV::eGFP expression plasmid into ventricular layercells (Haas et al., 2002). The day after electroporation, eGFP is expressed in cells with radial glialmorphology lining the ventricle (Haas et al., 2002; Tremblay et al., 2009). The majority of eGFP-expressing radial glial cells in stage 47 optic tectum are musashi1-immunoreactive (Fig 3D,E). Theseresults show that a 2 hr exposure to XdU in vivo labels musashi1-immunoreactive neural progenitors andthat 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 thefunctional visual system in Xenopus tadpoles and suggests that visual circuit function may negativelyregulate progenitor cell activity. To test whether visual experience regulates cell proliferation in thetadpole optic tectum, animals were reared in a 12 hr light/12 dark cycle until stage 46 when rates of cellproliferation are still relatively high (Fig 1). Tadpoles were separated into 3 groups: One group continuedunder the normal 12 hr light/12 dark cycle, called ‘ambient light’. The second group was provided withenhanced visual stimulation (from an array of LEDs flashing on and off at 1Hz (Sin et al., 2002)) during

3 3





























the 12 hr light period of the light/dark cycle and the third group was deprived of visual stimulation (bykeeping them in the dark, see diagram in Fig 4A). All tadpoles were kept in the dark during the 12 hr darkperiod of light/dark cycle.

We determined the division history of proliferating cells in the optic tectum using an assay that is basedon a double-label protocol in which animals were exposed to two deoxyuridine analogs, CldU and IdU, atan 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 immunostainingin 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 deoxyuridineanalog X dU (either CldU or IdU; green in Fig 4B) and allowed tadpoles to grow in the absence of X dUlabel for 24 hr, followed by a 2 hr exposure to the second deoxyuridine analog X dU (either CldU or IdU;red in Fig 4B). We used a 24h interval in between the X dU and X dU exposures based on the observationthat the number of XdU-labeled cells approximately doubled over 24 hr (Fig 1I). The optic tectum wasthen analyzed for the presence of X dU and X dU labeling in 30 µm cryostat sections. Three labelingcombinations are predicted: 1. cells with only X dU (green in Fig 4B) 2. cells with only X dU label (red in fig 4B) and 3. cells labeled with both XdU's (yellow in Fig 4B). The number of cells labeled with X dU(green) relative to the total number of labeled cells is the fraction that were in S phase at the first XdUexposure, but were not in S phase at the time of exposure to X dU. This value gives an estimate of thecells that exit the cell cycle between the times of exposure to X dU and X dU. The number of cells labeledwith X dU (red) relative to the total number of labeled cells is the fraction that was in S phase only at thesecond XdU exposure but not during exposure to X dU. This value would represent a potentialrecruitment 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 XdUexposure periods and therefore provides an estimate of the population of progenitors that remainedproliferative during the observation window. Although this protocol does not identify all cells that areproliferating 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 intadpoles reared in a normal 12 hr light/12 dark cycle (referred to as ambient light) with animals providedwith either enhanced visual stimulation or deprived of visual stimulation.

We counted XdU-labeled cells along the midline ventricular layer in sections through the optic tectumand found that tadpoles subjected to reduced visual experience had more XdU-labeled cells in the samevolume (79 ± 7.6 cells/20,000µm , n=5) compared to animals that were exposed to either ambient light(48.7 ± 5.0 cells/20,000µm , n=5; p<0.05) or enhanced visual stimulation (44 ± 7.3 cells/20,000µm ,n=5; p<0.05; Fig 4 C, D and E). Similar indices of XdU-labeled cells were seen in the optic tecta oftadpoles exposed to either enhanced visual activity or ambient light. These data suggest that visualdeprivation for 2 days increases cell proliferation in the optic tectum. They further suggest that theamount of visual activity provided by ambient light is sufficient to reduce cell proliferation in the tectum.

Next, we determined the fraction of X dU-labeled cells that also incorporated X dU, as an estimate of thecell population that continued to proliferate over the 24h period. Animals deprived of visual experiencehave a significantly larger fraction of X dU and X dU double-labeled cells (82.1 ± 1.9%) compared totadpoles 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 experienceindicates that relatively fewer X dU-labeled cells were in S phase of the cell cycle at the time of exposure

1 12

1 2

1 21 2

1

21 2

21

33 3

1 2

1 2

1


















to the second label, X dU. Furthermore, visual experience results in a higher fraction of cells labeled withonly X dU compared to visual deprivation (Fig 4 G) consistent the idea that visual experience changes thefate of tectal progenitors so they exit the cell cycle or become quiescent. Together, these data indicate thatvisual system activity decreases proliferative activity, while reduced visual experience maintains cells in aproliferative state by decreasing cell cycle exit and maintaining cells in the progenitor pool at the expenseof differentiation.

Visual experience increases neuronal differentiation

The data presented above suggest that visual experience may change the fate of the progeny of tectalprogenitor cells so they differentiate into neurons. To test whether visual experience promotes neuronaldifferentiation of newly generated cells, animals were reared in their normal 12h light/12h darkconditions until stage 47 and exposed to XdU for 2h in rearing solution. Animals were then eitherdeprived 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 experienceincreases the proportion of XdU-labeled cells that differentiated into N-β-tubulin-labeled neurons overthe 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 ofprogeny 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 ofmechanisms in relation to neural progenitor cells. The number of neural progenitor cells could remainconstant while the cell divisions occur less frequently or the size of neural progenitor pool could decreasewith visual system activity. Either mechanism would result in a decreased fraction of X dU and X dUdouble-labeled cells and a reciprocal increase in the fraction of cells that stop dividing after X dUexposure. To differentiate between these two possibilities we tested whether the musashi1-expressingneural progenitor cell population is affected by visual experience. Animals were reared under normalconditions to stage 47 when they were either deprived of visual experience or provided with visualexperience, as shown in Figure 5C. The number of musashi1-expressing cells was significantly lower intecta of animals with visual experience (267.2 ± 15.3 cells/20,000µm , n=7) compared to tecta ofvisually-deprived animals (426.9 ± 32.8 cells/20,000µm , n=7; p<0.05) (Fig 6 A, B and C). These resultsare consistent with a model in which the decrease in cell proliferation seen with visual stimulation resultsfrom a reduction in number of musashi1-expressing neural progenitor cells, whereas visual deprivationincreases the pool of musashi1-expressing neural progenitor cells.

The experiments described above indicate that musashi1 expression correlates with the proliferativeactivity of tectal neural progenitor cells. To test whether musashi1 is required for the increase in cellproliferation in visually-deprived animals, shown in Figure 4, we knocked down nrp1B, the Xenopushomolog of musashi1 using morpholino antisense oligonucleotides to nrp1B. To test the efficacy ofknockdown, morpholinos against nrp1B or control scrambled morpholinos were electroporated into theright tectal lobe. After 48 hours, we compared the intensity of musashi1 immunoreactivity in ventricularlayer cells of the right and left optic tecta. Morpholinos against nrp1B reduced the ratio of musashi1immunoreactivity 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 musashi1immunoreactivity between right and left tecta after unilateral electroporation (Supplementary Data

21

1 21

33










Figure 2). We then electroporated stage 46 tadpoles in both tectal lobes with nrp1B morpholinos orcontrol morpholinos and subjected animals to reduced visual stimulation for 60 hrs followed by 2hrexposure to XdU. Morpholinos against nrp1B significantly reduced the number of XdU-labeled cells(36.93 ± 1.94 cells/20,000µm , 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 cDNAwith morpholinos directed against Xenopus nrp1B mRNA completely rescued the morpholino-inducedreduction in proliferation (62.5 ± 4.0 cells/20,000µm , n= 11; p<0.05; Fig 6 F, G). These data indicatethat musashi1 protein levels are negatively regulated by visual activity in the optic tectum and thatmusashi1 is necessary for the increased cell proliferation seen in visually-deprived animals. Together withdata presented in Figure 5, the data show that progenitors increase musashi expression, increaseproliferative activity and expand the progenitor pool in the absence of visual input but that visualexperience triggers two changes in the system: 1. a decrease in musashi expression and a decrease inproliferative activity in radial glial cells and 2. an increase in the rate at which newly generated progenydifferentiate into neurons.

To test whether musashi1 is sufficient to increase cell proliferation in the CNS we returned to stage 49tadpoles, where musashi1 expression is relatively low in ventricular layer cells (Figure 2). Electroporationof ventricular layer cells in stage 49 tadpoles with a dual promoter plasmid co-expressing mousemusashi1 and eGFP or eGFP alone labeled radial glial cells (Figure 7A,B). In vivo time-lapse images ofeGFP+ cells co-expressing mouse musashi1 show a greater increase in eGFP-expressing cells over 6 daysthan seen in control animals, suggesting that exogenous expression of musashi1 increases proliferation ofradial glia. Furthermore, we tested whether musashi1 expression in stage 49 tadpoles increases XdUincorporation. Stage 49 tadpoles were electroporated with a dual promoter plasmid co-expressing eithereGFP alone or mouse musashi1 and eGFP and after 60 hr, animals were exposed to XdU for 2 hrimmediately before sacrifice. Mouse musashi1 expression doubled the number of XdU-labeled cellscompared to expression of eGFP alone (Musashi1: 21.3±1.3 cells/20,000µm , n=17, eGFP: 10.9±0.9cells/20,000µm , n=18; p< 0.05).

Discussion

Rates of neurogenesis decrease in the developing CNS as neuronal circuits become functional, suggestingthat neuronal activity generated by developing circuits may regulate the addition of new cells to thecircuit. We have used quantitative analysis of cell proliferation in the developing visual system of tadpolesto show that rates of cell proliferation decrease as the visual circuit matures. These results demonstrate atemporal correlation between the establishment of functional circuits and a decrease in cell proliferationand suggest the presence of a negative feedback mechanism from the developing circuit to neuralprogenitors to limit their proliferation. Consistent with this idea, we demonstrate that sensory inputactivity decreases cell proliferation in the optic tectum and increases the rate at which newly generatedcells differentiate into neurons. We provide evidence that immunoreactivity to musashi1, a highlyconserved mRNA binding protein which is required to maintain progenitor activity across phyla (Okanoet al., 2005), is down-regulated during the developmental period when cell proliferation decreases andthe normal developmental down regulation does not occur when animals are deprived of visualexperience for 2 days. We show that knockdown of nrp1B, the Xenopus homolog of musashi1 prevents thevisual-deprivation induced increase in cell proliferation. Finally, we report that expression of mousemusashi1 increases rates of cell proliferation when endogenous protein has been knocked down bymorpholinos or has been developmentally down-regulated. These data indicate that the control of neural

3

3

33











progenitor proliferation and cell fate during CNS development is regulated by neuronal activity such thatincreased activity from developing circuits changes the fate of newly generated cells so they exit the cellcycle and differentiate into neurons. The data further suggest that musashi1 expression is regulated byactivity in the intact CNS and that activity-dependent downregulation of musashi1 expression decreasescell proliferation and increases neuronal differentiation in the developing CNS. Finally, the data indicatethat neural progenitor proliferative activity can be dynamically regulated by increasing or decreasingmusashi1 expression, either by modulating brain activity or by more direct manipulation of musashi1expression levels.

Tectal neural progenitor cells are radial glia

We show that musashi1-expressing neural progenitor cells in tadpole optic tectum are radial glial cellsand that they generate tectal neurons. Neural progenitor cells with radial glial morphology have beenreported in cortical regions of many vertebrates, suggesting that radial glial neural progenitors are highlyconserved 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 beenclearly demonstrated. The decrease in cell proliferation and musashi1 immunoreactivity between stages46 and 49 and with enhanced visual experience suggest that the progenitor pool is depleted in response toactivity-dependent signals. It is interesting to note that electroporation of ventricular layer cells in stage49 tadpoles with a CMV::eGFP expression plasmid labels radial glial cells, showing that radial glial cellspersist at these stages, but they have lower proliferative activity and low musashi1 expression. Expressionof mouse musashi1 in radial glial cells of stage 49 tadpoles increases XdU incorporation and increased thenumber of eGFP-expressing cells over a 6 day period of in vivo imaging. This experiment suggests that thenormal experience-dependent developmental decrease in expression of musashi1 in radial glial cellsdecreases their proliferative activity, but that mechanisms that increase musashi1 expression in radialglia, either through a decrease in sensory input or by exogenous expression of musashi1, can increaseproliferation in these relatively quiescent progenitors.

The increased XdU incorporation seen in stage 46 animals compared to stage 49 animals could indicatethat S phase of the cell cycle is longer in younger animals. Similarly, the differences in XdU-incorporationin animals in which musashi expression is decreased by knockdown or increased by exogenous expressionof mouse musashi, could arise from corresponding changes in the length of S phase. To attempt toaddress this possibility, we labeled tecta with antibodies to phospho-histone 3 (PH3) to determine themitotic index. We found that PH3 labels 2-6 cells/20,000µm along the tectal midline, but the labeling istoo sparse to test for statistically significant differences across stages (data not shown). Although wecannot strictly conclude that the mitotic index does not change with developmental stage and musashi1expression, our data nevertheless provide evidence for regulation of neurogenesis by visual experienceand musashi1 expression in radial glial progenitors.

Control of neurogenesis and brain size by an interplay of ‘intrinsic’ cell autonomous and ‘extrinsic’ non-cell autonomous mechanisms

Regulation of neurogenesis establishes the number, type and distribution of cells in the CNS by regulationof progenitor maintenance and proliferation, and the subsequent survival and differentiation of progenyinto neurons or glia. Controlling the size of the progenitor pool by regulating the fate of progenitors andtheir progeny to either die, differentiate or continue to proliferate is a key regulatory event in braindevelopment and ultimately in the establishment of functional circuits and behavior. For instance,

3




maintaining progenitors in an actively proliferative state enlarges the progenitor pool and can enlarge thecerebral cortex of mice (Chenn and Walsh, 2002) by setting an upper bound on the potential number ofneurons generated (Lehmann et al., 2005). Conversely, premature transition of progenitors from anactive proliferative state to differentiated cells can deplete the progenitor pool, while increasing ordecreasing progenitor survival controls cortical size (Depaepe et al., 2005; Putz et al., 2005). A variety ofextrinsic factors can affect the fate of neural progenitor progeny by either maintaining progenitor fate orpromoting 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 singlesignals or combinations of signals operate downstream of visual system activity to decrease rates of cellproliferation during CNS development is not clear.

Extrinsic signals must be read and interpreted by intrinsic cell autonomous mechanisms to culminate incell fate regulation. Our data indicate that sensory input activity depletes the neural progenitorpopulation by down-regulating musashi1, suggesting that musashi1 is a prominent ‘intrinsic’maintenance/differentiation switch which can respond to non-cell autonomous signals from developingbrain circuits to mediate feedback from the developing circuit and control cell proliferation. Musashi1 isthought to function as a translational repressor. Using genomic and proteomic approaches, musashi1 wasfound to affect the expression of a large number of targets involved in the cell cycle, apoptosis, cellproliferation and cell differentiation (de Sousa Abreu et al., 2009). Musashi1 may promote maintenanceof 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 progenitorcells (Hitoshi et al., 2002; Tokunaga et al., 2004). In addition, musashi1 represses expression of p21, 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 earlystages of neuronal differentiation and is important for neuronal migration, differentiation and plasticity(Horisawa et al., 2009). Although considerable evidence suggests that musashi1 maintains neuralprogenitors in a proliferative state by repressing translation of proteins required for cells to exit the cellcycle and differentiate, it is not clear whether a single gate keeper integrates multiple extrinsic signals tocontrol cell fate (Suh et al., 2009). In addition to musashi1, other proteins including musashi2, Sox2 andPTEN 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 ofmusashi1, 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 ofradial 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 cycleand differentiate. A recent study demonstrated that Xenopus tadpole radial glial cell bodies exhibit slowcalcium transients in response to visual stimulation (Tremblay et al., 2009). The same study used time-lapse in vivo imaging to show that radial glial processes near the pia are dynamic and that visualstimulation increases radial glial structural dynamics by a mechanism that includes NMDA receptor-mediated NO signaling. This is interesting because previous studies in Xenopus tadpoles indicated thatNOS-expressing neurons are positioned immediately adjacent to the radial glial cell bodies, and thatdecreasing 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

WAF-1


































rearrangements when they devote their cytoskeletal resources to divide. Together these studies suggestthat visual system activity negatively regulates radial glial cell proliferation in the tadpole optic tectum bya signaling pathway that includes an NMDA receptor-mediated increase in NO, which decreasesproliferation 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 toenriched environments or exercise increases adult neurogenesis, presumably through activity-inducedincreases in trophic factors or neurotransmitters (Brown et al., 2003; Rochefort et al., 2002; Whitmanand Greer, 2009; Zhao et al., 2008). Further investigations to distinguish whether the increasedneurogenesis in adult CNS is the outcome of increased cell proliferation, differentiation or survival ofdifferentiated progeny suggest that, even though exercise may increase rates of proliferation in thehippocampal dentate gyrus (Brown et al., 2003), it appears that the more prevalent effect of increasedbrain 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 ratesof cell proliferation, increases cell cycle exit and increases neuronal differentiation of newly generatedcells, but tunel staining suggests that cell death is not a significant factor in regulating neurogenesis in thedeveloping optic tectum (Peunova et al., 2001), at least over the time-course of our studies. These dataindicate that visual experience limits self-renewal of neural progenitor cells, thereby depleting theprogenitor pool, and shifts the fate of progeny to terminal neuronal differentiation. These differenceslikely reflect distinct functions of neurogenesis in the adult and developing CNS. During CNSdevelopment the function of neurogenesis is to generate a large population of neurons in a relatively finitetime period so that functional neural circuits can be assembled de novo. By contrast, adult-generatedneurons integrate into pre-existing neural circuits where they are thought to modulate learning andmemory (Whitman and Greer, 2009). Therefore, different responses to activity-dependent signals inneural progenitor cells in the developing and adult CNS likely reflect the requirements of the CNS for thegeneration 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, FortAtkinson, WI), were reared in a 12 hr light/12 hr dark cycle incubator at 24°C. Animals were stagedaccording 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 visualstimulation 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 theexperimental conditions tadpoles were reared in a 24°C incubator. The time-course of the reduced orenhanced visual experience protocol is shown in Figure 4A.

Immunohistochemistry

Xenopus tadpoles were anesthetized with 0.02% MS222 (3-aminobenzoic acid ethyl ester), immersed in4% paraformaldehyde (Electron Microscopy Sciences, Fort Washington, PA) in PBS and microwaved for















8 sec (750 W GE microwave) followed by 2h in fixative at room temperature (Li et al., 2010). After rinsingthe 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 µmhorizontal sections with a vibratome. Sections were blocked in 5% goat serum and 0.3% Tween 20 in PBSfor 1 hr before incubating overnight at 4°C with anti-MCM7 (1:250; Abcam, Cambridge, MA), anti-musashi1 (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 wereprepared, 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 collectivelyas XdU's (MP Biomedicals, Solon, OH) at 10 mM in Steinberg's solution (60 mM NaC1, 0.7 mM KCI, 0.8mM MgSO , 0.3 mM Ca(NO ) and 1.4 mM Tris, pH 7.4) for 2 hr. We found that delivery of XdU byinjection into the pericardial cavity or by 2hr exposure of tadpoles to XdU in rearing solution resulted incomparable labeling of proliferative cells. We used exposure in rearing solution for XdU delivery in ourexperiments 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 todetermine the history of proliferative activity, as described (Encinas and Enikolopov, 2008; Vega andPeterson, 2005). Tadpoles were exposed to IdU in Steinberg's solution for 2hr and transferred to freshSteinberg's solution. After 24 hrs they were exposed to CldU in Steinberg's solution for 2 hr, rinsed infresh Steinberg's solution, anesthetized with 0.02% MS222 and fixed as described above. Forwholemount analysis, dissected brains were treated with −20°C methanol for 1 hr followed by proteinaseK treatment as described (Peunova et al., 2001). For detection of IdU and CldU, wholemount brains orsections were treated with 2N HCl for 1 hr at 37°C, rinsed in PBS and incubated in 5% normal goat serumand 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 anti-BrdU Cat# OBT0030G). The IdU-specific antibody was detected using Alexa 488 labeled goat anti-mousesecondary antibody (Molecular Probes/Invitrogen, Eugene, OR). The CldU-specific antibody signal wasenhanced using biotin tagged goat anti-Rat secondary antibody followed by detection with Cy5-streptavidin (Jackson ImmunoResearch). To reduce crossreactivity, highly cross-adsorbed secondaryantibodies were used. Wholemounts and sections were mounted in ProLong™ Gold (MolecularProbes/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 NAwater objective. No significant crossreactivity was observed for IdU and CldU (Supplementary Fig 1). Totest whether there is a labeling bias in our method for detecting IdU and CldU (i.e. whether we can detecta cell in S-phase by IdU or CldU with equal probability) we exposed tadpoles to equimolar CldU and IdUsimultaneously for 2 hr. All the cells were double-labeled, indicating that we can detect S-phase cells intadpoles using IdU or CldU with equal probability (Supplementary Fig 1). We used CldU and IdUinterchangeably and obtained the similar results.

4 3 2









Data Analysis

Image analysis and cell counting were performed using Metamorph (Universal ImagingCorporation/Molecular Devices, Downingtown, PA) image processing software. Images were backgroundsubtracted and cells were counted using the dissector method with the “manually count object” feature ofMetamorph. In wholemounts of brains, cells in the ventricular layer at the midline of the optic tectum inan area 100 µm along the ventricular layer X 20µm lateral from the ventricular layer were counted in twonon-neighboring optical sections. The first optical section was selected where the two tectal lobes meet atthe dorsal midline and the second optical section was 8-10 µm ventral from the first. The averagediameter of cell nuclei is 5 µm and does not change with the different stages or treatments analyzed. Therostral boundary for counting cells within the tectal ventricular layer was defined by the anterior dorsalcommissure and the caudal boundary was identified as the start of the curvature to the caudolateral edgeof the tectum, as shown in Figure 1. Cell counts from the 2 sections were added and presented as cellnumber per 20,000 µm . We used cryostat sections for high-resolution 40X analysis to determine thepresence of IdU only, CldU only or both IdU and CldU in cells. Two horizontal sections from each braincorresponding to about 60-90 µm and 120-150 µm from the dorsal side respectively were used foranalysis. Cells were counted in the entire z stack through the section collected with 2 µm z-step size. Therostral 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 inXenopus laevis (GeneTools,Philomath, OR) with the sequence GCGCTTCTGTCTCCATTCGGTCTCT, orthe 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 homologueof musashi2, according to prediction algorithms by GeneTools (GeneTools,Philomath, OR). The tadpoleswere provided with visual experience or deprived of visual experience as described above. At specifiedtimes 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 constructwith no construct in the second site. Plasmids were electroporated into the optic tectum of anesthetizedtadpoles 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 diskconfocal microscope (Perkin Elmer) with a 20X water immersion lens (0.9 NA), as described (Ruthazerand Cline, 2004).

Statistical tests

Data were tested with the non-parametric Mann–Whitney–Wilcoxon test to compare between groupsunless stated otherwise. Data are represented as mean ± SEM.

Supplementary Material

3











01

02

03

04

Acknowledgements

this work was supported by the NIH (DP10D000458 to HTC), Dart Neuroscience, LLC (HTC) andfellowships from the Cold Spring Harbor Laboratory Association and CIRM to PS. We thank Drs. Imaiand Okano for generously providing the cDNA for mouse Musashi1 and members of the Cline lab forinsightful discussions.

Footnotes

Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a serviceto our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting,typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during theproduction process errors may be discovered which could affect the content, and all legal disclaimers that apply to thejournal pertain.

References

1. Amato MA, Boy S, Arnault E, Girard M, Della Puppa A, Sharif A, Perron M. Comparison of theexpression patterns of five neural RNA binding proteins in the Xenopus retina. J Comp Neurol.2005;481:331–339. [PubMed: 15593335]

2. Battelli C, Nikopoulos GN, Mitchell JG, Verdi JM. The RNA-binding protein Musashi-1 regulatesneural development through the translational repression of p21WAF-1. Mol Cell Neurosci.2006;31:85–96. [PubMed: 16214366]

3. Bestman JE, Cline HT. The RNA binding protein CPEB regulates dendrite morphogenesis andneuronal circuit assembly in vivo. Proc Natl Acad Sci U S A. 2008;105:20494–20499.[PMCID: PMC2629308] [PubMed: 19074264]

4. Blow JJ, Dutta A. Preventing re-replication of chromosomal DNA. Nat Rev Mol Cell Biol.2005;6:476–486. [PMCID: PMC2688777] [PubMed: 15928711]

5. Botia B, Basille M, Allais A, Raoult E, Falluel-Morel A, Galas L, Jolivel V, Wurtz O, Komuro H,Fournier A, et al. Neurotrophic effects of PACAP in the cerebellar cortex. Peptides.2007;28:1746–1752. [PubMed: 17544170]

6. Brown J, Cooper-Kuhn CM, Kempermann G, Van Praag H, Winkler J, Gage FH, Kuhn HG.Enriched environment and physical activity stimulate hippocampal but not olfactory bulbneurogenesis. Eur J Neurosci. 2003;17:2042–2046. [PubMed: 12786970]

7. Butt SJ, Fuccillo M, Nery S, Noctor S, Kriegstein A, Corbin JG, Fishell G. The temporal andspatial origins of cortical interneurons predict their physiological subtype. Neuron. 2005;48:591–604. [PubMed: 16301176]

8. Chenn A, Walsh CA. Regulation of cerebral cortical size by control of cell cycle exit in neuralprecursors. Science. 2002;297:365–369. [PubMed: 12130776]

9. Chiu SL, Chen CM, Cline HT. Insulin receptor signaling regulates synapse number, dendritic



plasticity, and circuit function in vivo. Neuron. 2008;58:708–719. [PMCID: PMC3057650][PubMed: 18549783]

10. Cline H, Haas K. The regulation of dendritic arbor development and plasticity by glutamatergicsynaptic input: a review of the synaptotrophic hypothesis. J Physiol. 2008;586:1509–1517.[PMCID: PMC2375708] [PubMed: 18202093]

11. Cline HT. Dendritic arbor development and synaptogenesis. Current Opinion in Neurobiology.2001;11:118–126. [PubMed: 11179881]

12. Crevel G, Hashimoto R, Vass S, Sherkow J, Yamaguchi M, Heck MM, Cotterill S. Differentialrequirements for MCM proteins in DNA replication in Drosophila S2 cells. PLoS ONE.2007;2:e833. [PMCID: PMC1950684] [PubMed: 17786205]

13. de Sousa Abreu R, Sanchez-Diaz PC, Vogel C, Burns SC, Ko D, Burton TL, Vo DT,Chennasamudaram S, Le SY, Shapiro BA, Penalva LO. Genomic analyses of musashi1downstream targets show a strong association with cancer-related processes. J Biol Chem.2009;284:12125–12135. [PMCID: PMC2673281] [PubMed: 19258308]

14. Depaepe V, Suarez-Gonzalez N, Dufour A, Passante L, Gorski JA, Jones KR, Ledent C,Vanderhaeghen P. Ephrin signalling controls brain size by regulating apoptosis of neuralprogenitors. Nature. 2005;435:1244–1250. [PubMed: 15902206]

15. Encinas JM, Enikolopov G. Identifying and quantitating neural stem and progenitor cells in theadult brain. Methods Cell Biol. 2008;85:243–272. [PubMed: 18155466]

16. Engert F, Tao HW, Zhang LI, Poo MM. Moving visual stimuli rapidly induce direction sensitivityof developing tectal neurons. Nature. 2002;419:470–475. [PubMed: 12368854]

17. Facoetti A, Ranza E, Benericetti E, Ceroni M, Tedeschi F, Nano R. Minichromosome maintenanceprotein 7: a reliable tool for glioblastoma proliferation index. Anticancer Res. 2006;26:1071–1075. [PubMed: 16619508]

18. Gandhi R, Luk KC, Rymar VV, Sadikot AF. Group I mGluR5 metabotropic glutamate receptorsregulate proliferation of neuronal progenitors in specific forebrain developmental domains. JNeurochem. 2008;104:155–172. [PubMed: 17944877]

19. Glazer RI, Wang XY, Yuan H, Yin Y. Musashi1: a stem cell marker no longer in search of afunction. Cell Cycle. 2008;7:2635–2639. [PMCID: PMC3676649] [PubMed: 18719393]

20. Good PJ, Rebbert ML, Dawid IB. Three new members of the RNP protein family in Xenopus.Nucleic Acids Res. 1993;21:999–1006. [PMCID: PMC309235] [PubMed: 8451200]

21. Gotz M, Huttner WB. The cell biology of neurogenesis. Nat Rev Mol Cell Biol. 2005;6:777–788.[PubMed: 16314867]

22. Graham V, Khudyakov J, Ellis P, Pevny L. SOX2 Functions to Maintain Neural ProgenitorIdentity. Neuron. 2003;39:749–765. [PubMed: 12948443]

23. Groszer M, Erickson R, Scripture-Adams DD, Dougherty JD, Le Belle J, Zack JA, Geschwind DH,Liu X, Kornblum HI, Wu H. PTEN negatively regulates neural stem cell self-renewal bymodulating G0-G1 cell cycle entry. Proc Natl Acad Sci U S A. 2006;103:111–116.[PMCID: PMC1325011] [PubMed: 16373498]

24. Haas K, Jensen K, Sin WC, Foa L, Cline HT. Targeted electroporation in Xenopus tadpoles invivo- from single cells to the entire brain. Differentiation. 2002;70:148–154. [PubMed: 12147134]

25. Hitoshi S, Alexson T, Tropepe V, Donoviel D, Elia AJ, Nye JS, Conlon RA, Mak TW, Bernstein A,van der Kooy D. Notch pathway molecules are essential for the maintenance, but not thegeneration, of mammalian neural stem cells. Genes Dev. 2002;16:846–858.[PMCID: PMC186324] [PubMed: 11937492]



26. Holt CE, Harris WA. Order in the initial retinotectal map in Xenopus: a new technique forlabelling frowing nerve fibres. Nature. 1983;301:150–152. [PubMed: 6823290]

27. Horisawa K, Imai T, Okano H, Yanagawa H. 3′-Untranslated region of doublecortin mRNA is abinding target of the Musashi1 RNA-binding protein. FEBS Lett. 2009;583:2429–2434.[PubMed: 19573529]

28. Huttner WB, Kosodo Y. Symmetric versus asymmetric cell division during neurogenesis in thedeveloping vertebrate central nervous system. Current Opinion in Cell Biology. 2005;17:648–657. [PubMed: 16243506]

29. Imai T, Tokunaga A, Yoshida T, Hashimoto M, Mikoshiba K, Weinmaster G, Nakafuku M, OkanoH. The neural RNA-binding protein Musashi1 translationally regulates mammalian numb geneexpression by interacting with its mRNA. Mol Cell Biol. 2001;21:3888–3900.[PMCID: PMC87052] [PubMed: 11359897]

30. Kaneko Y, Sakakibara S, Imai T, Suzuki A, Nakamura Y, Sawamoto K, Ogawa Y, Toyama Y,Miyata T, Okano H. Musashi1: An Evolutionally Conserved Marker for CNS Progenitor CellsIncluding Neural Stem Cells. Developmental Neuroscience. 2000;22:139–153.[PubMed: 10657706]

31. Khalili K, Del Valle L, Muralidharan V, Gault WJ, Darbinian N, Otte J, Meier E, Johnson EM,Daniel DC, Kinoshita Y, et al. Puralpha is essential for postnatal brain development anddevelopmentally coupled cellular proliferation as revealed by genetic inactivation in the mouse.Mol Cell Biol. 2003;23:6857–6875. [PMCID: PMC193944] [PubMed: 12972605]

32. Kriegstein A, Alvarez-Buylla A. The glial nature of embryonic and adult neural stem cells. AnnuRev Neurosci. 2009;32:149–184. [PMCID: PMC3086722] [PubMed: 19555289]

33. Lehmann K, Butz M, Teuchert-Noodt G. Offer and demand: proliferation and survival of neuronsin the dentate gyrus. Eur J Neurosci. 2005;21:3205–3216. [PubMed: 16026459]

34. Li J, Wang Y, Chiu SL, Cline HT. Membrane targeted horseradish peroxidase as a marker forcorrelative fluorescence and electron microscopy studies. Front Neural Circuits. 2010;4:6.[PMCID: PMC2831632] [PubMed: 20204144]

35. LoTurco JJ, Owens DF, Heath MJ, Davis MB, Kriegstein AR. GABA and glutamate depolarizecortical progenitor cells and inhibit DNA synthesis. Neuron. 1995;15:1287–1298.[PubMed: 8845153]

36. Moody S, M. V, Spanos A, F. A. Developmental expression of a neuron-specific beta-tubulin infrog (Xenopus laevis): a marker for growing axons during the embryonic period. Journal ofComparative Neurology. 1996;364:219–230. [PubMed: 8788246]

37. Nakamura M, Okano H, Blendy JA, Montell C. Musashi, a neural RNA-binding protein requiredfor Drosophila adult external sensory organ development. Neuron. 1994;13:67–81.[PubMed: 8043282]

38. Nieuwkoop PD, Faber J. Normal Table of Xenopus laevis (Daudin) Elsevier-North HollandPublishing Co.; Amsterdam: 1956.

39. Ninkovic J, Gotz M. Signaling in adult neurogenesis: from stem cell niche to neuronal networks.Curr Opin Neurobiol. 2007;17:338–344. [PubMed: 17475475]

40. Noctor SC, Martinez-Cerdeno V, Kriegstein AR. Contribution of Intermediate Progenitor Cells toCortical Histogenesis. Arch Neurol. 2007;64:639–642. [PubMed: 17502462]

41. O'Rourke NA, Fraser S,E. Dynamic changes in optic fiber terminal arbors lead to retinotopic mapformations: An in vivo confocal microscopic study. Neuron. 1990;5:159–171. [PubMed: 2383399]

42. Ohtsuka T, Sakamoto M, Guillemot F, Kageyama R. Roles of the basic helix-loop-helix genes



Hes1 and Hes5 in expansion of neural stem cells of the developing brain. J Biol Chem.2001;276:30467–30474. [PubMed: 11399758]

43. Okano H, Imai T, Okabe M. Musashi: a translational regulator of cell fate. J Cell Sci.2002;115:1355–1359. [PubMed: 11896183]

44. Okano H, Kawahara H, Toriya M, Nakao K, Shibata S, Imai T. Function of RNA-binding proteinMusashi-1 in stem cells. Exp Cell Res. 2005;306:349–356. [PubMed: 15925591]

45. Padmanabhan V, Callas P, Philips G, Trainer TD, Beatty BG. DNA replication regulation proteinMcm7 as a marker of proliferation in prostate cancer. J Clin Pathol. 2004;57:1057–1062.[PMCID: PMC1770448] [PubMed: 15452160]

46. Peunova N, Scheinker V, Cline H, Enikolopov G. Nitric Oxide Is an Essential Negative Regulatorof Cell Proliferation in Xenopus Brain. J. Neurosci. 2001;21:8809–8818. [PubMed: 11698593]

47. Peunova N, Scheinker V, Ravi K, Enikolopov G. Nitric oxide coordinates cell proliferation and cellmovements during early development of Xenopus. Cell Cycle. 2007;6:3132–3144.[PubMed: 18073535]

48. Pratt KG, Aizenman CD. Homeostatic regulation of intrinsic excitability and synaptictransmission in a developing visual circuit. J Neurosci. 2007;27:8268–8277. [PubMed: 17670973]

49. Pratt KG, Aizenman CD. Multisensory integration in mesencephalic trigeminal neurons inXenopus tadpoles. J Neurophysiol. 2009;102:399–412. [PMCID: PMC2712281][PubMed: 19386750]

50. Pratt KG, Dong W, Aizenman CD. Development and spike timing-dependent plasticity ofrecurrent excitation in the Xenopus optic tectum. Nat Neurosci. 2008;11:467–475.[PubMed: 18344990]

51. Putz U, Harwell C, Nedivi E. Soluble CPG15 expressed during early development rescues corticalprogenitors from apoptosis. Nat Neurosci. 2005;8:322–331. [PMCID: PMC3075944][PubMed: 15711540]

52. Qiu R, Wang X, Davy A, Wu C, Murai K, Zhang H, Flanagan JG, Soriano P, Lu Q. Regulation ofneural progenitor cell state by ephrin-B. J Cell Biol. 2008;181:973–983. [PMCID: PMC2426945][PubMed: 18541704]

53. Richter K, Good PJ, Dawid IB. A developmentally regulated, nervous system-specific gene inXenopus encodes a putative RNA-binding protein. New Biol. 1990;2:556–565.[PubMed: 1708282]

54. Rochefort C, Gheusi G, Vincent JD, Lledo PM. Enriched odor exposure increases the number ofnewborn neurons in the adult olfactory bulb and improves odor memory. J Neurosci.2002;22:2679–2689. [PubMed: 11923433]

55. Ruthazer ES, Cline HT. Time-Lapse Imaging of CNS Neuron Development in Vivo. In: Yuste R,Lani F, Konnerth A, editors. Imaging: A Laboratory Manual. Cold Spring Harbor Press; ColdSpring Harbor: 2004. pp. 112–120.

56. Sadikot AF, Burhan AM, Belanger MC, Sasseville R. NMDA receptor antagonists influence earlydevelopment of GABAergic interneurons in the mammalian striatum. Brain Res Dev Brain Res.1998;105:35–42.

57. Sakakibara S, Imai T, Hamaguchi K, Okabe M, Aruga J, Nakajima K, Yasutomi D, Nagata T,Kurihara Y, Uesugi S, et al. Mouse-Musashi-1, a neural RNA-binding protein highly enriched inthe mammalian CNS stem cell. Dev Biol. 1996;176:230–242. [PubMed: 8660864]

58. Sakakibara S, Nakamura Y, Satoh H, Okano H. RNA-binding protein Musashi2: developmentallyregulated expression in neural precursor cells and subpopulations of neurons in mammalian



CNS. J Neurosci. 2001;21:8091–8107. [PubMed: 11588182]59. Sakakibara S, Nakamura Y, Yoshida T, Shibata S, Koike M, Takano H, Ueda S, Uchiyama Y, Noda

T, Okano H. RNA-binding protein Musashi family: roles for CNS stem cells and a subpopulationof ependymal cells revealed by targeted disruption and antisense ablation. Proc Natl Acad Sci U SA. 2002;99:15194–15199. [PMCID: PMC137566] [PubMed: 12407178]

60. Sakakibara S, Okano H. Expression of neural RNA-binding proteins in the postnatal CNS:implications of their roles in neuronal and glial cell development. J Neurosci. 1997;17:8300–8312. [PubMed: 9334405]

61. Sin WC, Haas K, Ruthazer ES, Cline HT. Dendrite growth increased by visual activity requiresNMDA receptor and Rho GTPases. Nature. 2002;419:475–480. [PubMed: 12368855]

62. Spitzer NC. Electrical activity in early neuronal development. Nature. 2006;444:707–712.[PubMed: 17151658]

63. Straznicky K, Gaze R. The development of the tectum in Xenopus laevis: an autoradiographicstudy. Journal of Embryology and Experimental Morphology. 1972;28:87–115.[PubMed: 5074324]

64. Suh H, Deng W, Gage FH. Signaling in adult neurogenesis. Annu Rev Cell Dev Biol.2009;25:253–275. [PubMed: 19575663]

65. Suh J, Lu N, Nicot A, Tatsuno I, DiCicco-Bloom E. PACAP is an anti-mitogenic signal indeveloping cerebral cortex. Nat Neurosci. 2001;4:123–124. [PubMed: 11175869]

66. Tao HW, Poo MM. Activity-dependent matching of excitatory and inhibitory inputs duringrefinement of visual receptive fields. Neuron. 2005;45:829–836. [PubMed: 15797545]

67. Tokunaga A, Kohyama J, Yoshida T, Nakao K, Sawamoto K, Okano H. Mapping spatio-temporalactivation of Notch signaling during neurogenesis and gliogenesis in the developing mouse brain.J Neurochem. 2004;90:142–154. [PubMed: 15198674]

68. Tremblay M, Fugere V, Tsui J, Schohl A, Tavakoli A, Travencolo BA, Costa Lda F, Ruthazer ES.Regulation of radial glial motility by visual experience. J Neurosci. 2009;29:14066–14076.[PubMed: 19906955]

69. Vega CJ, Peterson DA. Stem cell proliferative history in tissue revealed by temporal halogenatedthymidine analog discrimination. Nat Methods. 2005;2:167–169. [PubMed: 15782184]

70. Wexler EM, Paucer A, Kornblum HI, Plamer TD, Geschwind DH. Endogenous Wnt signalingmaintains neural progenitor cell potency. Stem Cells. 2009;27:1130–1141.[PMCID: PMC2782960] [PubMed: 19418460]

71. Whitman MC, Greer CA. Adult neurogenesis and the olfactory system. Prog Neurobiol.2009;89:162–175. [PMCID: PMC2748178] [PubMed: 19615423]

72. Wu GY, Zou DJ, Rajan I, Cline H. Dendritic Dynamics In Vivo Change during NeuronalMaturation. J. Neurosci. 1999;19:4472–4483. [PubMed: 10341248]

73. Zhao C, Deng W, Gage FH. Mechanisms and functional implications of adult neurogenesis. Cell.2008;132:645–660. [PubMed: 18295581]

Figures and Tables

Figure 1



Developmental decrease in proliferative cells in the Xenopus tadpole optic tectum. A-C. The Xenopus optic tectumincludes a functional visual circuit and proliferative ventricular layer cells. A. Phase contrast image of an albino Xenopustadpole 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 lateralneuropil. 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 toXdU and fixed immediately thereafter. D, F. Images of 3-D merge of a z series of horizontal confocal sections through themidbrain. E, G. Images of a series of single optical sections every 10 µm from dorsal to ventral tectum. The proliferatingcells labeled by XdU incorporation line the ventricle and decrease significantly between stage 46 and stage 49. XdU-labeled cells were counted in the region marked by the bracket in D. H, I. Quantitative analysis of numbers of XdU-labeledcells in the midline ventricular layer of stage 46, 48 and 49 tadpoles (n=8,8 and 7 animals respectively) which wereexposed to XdU for 2 hr, then fixed and analyzed either immediately (H) or after further rearing in the absence of XdU for22 hr (I) as shown in the schematic on top of each graph. XdU incorporation decreases significantly from stage 46 to stage49. Scale bar = 50 µm in F, also applies to D. *** p<0.0001.

Figure 2



Developmental decrease in MCM7 and Musashi1 immunoreactivity. Confocal images of 30 µm cryostat sections throughthe optic tectum of stages 46 (A, C) and 49 (B, D) tadpoles labeled with anti-MCM7 antibody (green; A1, B1) or anti-musashi1 antibody (C1, D1) and a nuclear stain SytoxO (blue; A2-D2). Cells located along the ventricular layer are highlyimmunoreactive for MCM7 and musashi1. E. Quantitative analysis of the decrease in number of Musashi1immunoreactive cells between stages 46 and 49. Scale bar = 20 µm. *p<0.05.

Figure 3



Lineage and identity of proliferating neural progenitor cells in optic tectum. A-B. Stage 48 tadpoles were exposed to XdUfor 2h in rearing solution and were either fixed immediately (A) or after 72 hrs of further development in the absence ofXdU (B). Confocal images of 30 µm cryostat sections through the optic tectum of tadpoles labeled with antibodies tomusashi1 (msi1; green) and XdU (red). Nuclei were stained with SytoxO (blue). Musashi1-immunoreactive cells line theventricle and are labeled by incorporation of XdU with a 2h survival time (A). The progeny of XdU-labeled neuralprogenitor cells are distributed throughout the cell body layer of the optic tectum after a 72 hr survival time (B). Note thatthe images of musashi1-immunoreactivity in B1 were taken with longer exposure periods than the images in A1, accordingto the lower levels of musashi1-immunoreactivity in ventricular layer cells at stage 49 (see Fig 2C,D). C. Musashi1-immunoreactive 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 24hrs and analyzed for Musashi1 immunoreactivity in 30 µm vibratome sections. D1-D3. 3-D merge of a z series containinga 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 withMusashi1 immunolabeling (E2, E3). Scale bar in D3 = 10 µm and E3 = 5 µm.

Figure 4




Visual deprivation increases cell proliferation in the optic tectum. A. Timeline of the experimental protocol. The 12dark/12h light rearing conditions are shown as shaded and white bars. During the 12h ‘light’ period animals were eitherexposed to normal ambient light, enhanced visual stimulation or reduced visual stimulation (see methods). Animals wereexposed to X dU for 2h and allowed to develop in the absence of X dU for 24 hours, after which they were exposed toX dU at the times marked by the green and red arrows, respectively. This protocol provides an estimate of the number ofproliferating cells (labeled with X dU and/or X dU), the fraction of X dU-labeled cells that remain in the cell cycle(double labeled with X dU and X dU), and the fraction of X dU-labeled cells that exit the cell cycle (X dU only). B. Acartoon of representative results of the assay to detect the division history of proliferating cells as labeled in A. Cellslabeled in green incorporated X dU only. Cells labeled in red incorporated X dU only. Cells labeled in yellow were in Sphase during exposure to X dU and again during exposure to X dU. C, D. Images of the dorsal midline ventricular layer ofthe optic tectum of tadpoles with enhanced visual experience (C1-C4) or reduced visual experience (D1-D4) labeled withanti-X dU (green; C2, D2) and anti-X dU (red; C3,D3). C1, D1. Images of the X dU and X dU-labeled cells in mergedoptical sections of the complete z-series through the depth of the 30 µm cryostat section. C2-C4 and D2-D4. Magnifiedsingle optical sections from the z-series in C and G showing a fraction of cells labeled with X dU only, X dU only and bothX dU and X dU. 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 oftotal numbers of XdU-labeled cells (E), fraction of X dU- and X dU-labeled cells (F) fraction of X dU-only labeled cells(G). * p<0.05, ** p < 0.001 compared to animals with reduced visual stimulation.

1 1

2

1 2 1

1 2 1 1

1 2

1 2

1 2 1 2

1 2

1 2

1 2 1



Figure 5

Visual experience increases neuronal differentiation. Stage 47 tadpoles were exposed to XdU for 2hr and allowed todevelop in absence of XdU for 48 hrs while subjected to either enhanced visual stimulation or reduced visual stimulationas 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 double-labeled with N- β-tubulin antibody. Enhanced visual stimulation increases the proportion of newly generated cells thatdifferentiate into neurons. Scale bar in A3 = 10 µm. ***p<0.0001

Figure 6



Musashi1 is required for visual experience-dependent regulation of cell proliferation. A-C. Visual deprivation for 48hincreases the number of Musashi1 (Msi1) immunoreactive neural progenitor cells compared to animals with enhancedvisual 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 cellswith detectable musashi1-immunoreactivity compared to visual stimulation (n=5 each). D-G. Morpholino-mediatedknockdown of nrp1B, the Xenopus homolog of musashi1, decreases cell proliferation in the optic tectum while expressionof mouse Musashi1 rescues cell proliferation in animals with nrp1B knockdown. Both tectal lobes of stage 46 tadpoleswere co-electroporated with expression constructs and morpholinos. Animals were deprived of visual stimulation for thenext 60h and then were exposed to XdU for 2h immediately before fixation. D-F. Optical sections showing XdU-labeledcells 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-labeledcells in the ventricular layer compared to control morpholinos (D,E,). The nrp1B morpholino-mediated decrease inproliferation was completely rescued by simultaneous expression of mouse musashi1 (F). G. Quantification of numbers ofXdU-labeled cells in animals treated as in D-F. Scale bar in B3 = 10 µm, E= 20 µm.*** p<0.0001.

Figure 7



Musashi1 expression is sufficient to increase cell proliferation. A.B. In vivo images of eGFP-expressing cells collected over6 days after electroporation of stage 49 tadpoles with a dual promoter plasmid expressing eGFP alone (A) or eGFP andmouse Musashi1 (mMsi1; B). Each image is a 3-D merge of a z-series through the optic tectum taken 2 days (d2, A1, B1), 4days (d4, A2, B2) and 6 days (d6, A3, B3) after electroporation. C,D. Musashi1 expression in stage 49 tadpoles increasesXdU incorporation. Animals were electroporated with plasmid expressing eGFP (C) or eGFP/mMsi1 (D) and 2 days laterwere exposed to XdU for 2hrs. Whole mount brains were analysed for XdU immunolabeling. C,D shows single opticalsections of XdU labeled cells in eGFP- (C) and eGFP/mMsi1-expressing animals (D). E. Quantitative analysis of XdU-labeled cells with or without exogenous expression of mMsi1. Exogenous expression of mMsi1 at stage 49 increasesproliferation compared to control. Scale bar in B3 and D = 20 µm, ***p<0.0001

visual activity regulates neural progenitor cells in ... files/2010/pmc sharma neuron...

Documents