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. 2011 Nov 10;195(4):191–200. doi: 10.1016/j.neuroscience.2011.08.035

Cytoarchitectural disruption of the superior colliculus and an enlarged acoustic startle response in the Tuba1a mutant mouse

A Edwards a, CD Treiber b, M Breuss b, R Pidsley a, G-J Huang a, J Cleak a, PL Oliver c, J Flint a, DA Keays c,⁎
PMCID: PMC3188702  PMID: 21875651

Abstract

The Jenna mutant mouse harbours an S140G mutation in Tuba1a that impairs tubulin heterodimer formation resulting in defective neuronal migration during development. The consequence of decreased neuronal motility is a fractured pyramidal cell layer in the hippocampus and wave-like perturbations in the cerebral cortex. Here, we extend our characterisation of this mouse investigating the laminar architecture of the superior colliculus (SC). Our results reveal that the structure of the SC in mutant animals is intact; however, it is significantly thinner with an apparent fusion of the intermediate grey and white layers. Birthdate labelling at E12.5 and E13.5 showed that the S140G mutation impairs the radial migration of neurons in the SC. A quantitative assessment of neuronal number in adulthood reveals a massive reduction in postmitotic neurons in mutant animals, which we attribute to increased apoptotic cell death. Consistent with the role of the SC in modulating sensorimotor gating, and the circuitry that modulates this behaviour, we find that Jenna mutants exhibit an exaggerated acoustic startle response. Our results highlight the importance of Tuba1a for correct neuronal migration and implicate postnatal apoptotic cell death in the pathophysiological mechanisms underlying the tubulinopathies.

Key words: tubulin lissencephaly, superior colliculus, acoustic startle response

Abbreviations: BrdU, 5-bromo-2-deoxyuridine; DAPI, 4',6-diamidino-2-phenylindole; DpG, deep grey layer; DpW, deep white layer; InG, intermediate grey layer; InW, intermediate white layer; Jenna, Jna/+; Op, optic layer; PAG, periaqueductal grey; SC, superior colliculus; SuG, superficial grey layer; Tpd52L1, tumour protein D52-like-1; Zn, zonal layer

Highlights

▶We investigate the superior colliculus and behaviour of a mouse model of lissencephaly. ▶We find that there is abnormal lamination and impairment in neuronal migration. ▶This is accompanied by a massive increase in postnatal neuronal cell death. ▶This is associated with an exaggerated acoustic startle response. ▶This study aids our understanding of disease mechanisms.

The Jenna (Jna/+) mouse, generated from an N-ethyl-N-nitrosourea mutagenesis screen, has a dominantly inherited mutation in exon four of the α-tubulin gene, Tuba1a (Keays et al., 2007). This mutation, an S140G substitution, impairs tubulin heterodimer formation, which results in defects in neuronal migration during development. Consequently, mutant animals have a fractured pyramidal cell layer in the hippocampus, and laminar abnormalities in the cerebral cortex, predominantly in layers III and IV. Human studies have found that this mouse is a model for lissencephaly, a disorder characterised by simplified gyration of the cortex, mental retardation and epilepsy (Guerrini and Marini, 2006). Mutations in a number of genes cause lissencephaly, including DCX, LIS1, VLDLR and the reelin gene (des Portes et al., 1998; Gleeson et al., 1998; Hong et al., 2000; Boycott et al., 2005). In the later case, homozygous mutations in reelin result in gyral simplification, a thickened cortex and cerebellar hypoplasia (Hong et al., 2000).

Reelin, a large extracellular protein, was first implicated in neuronal migration after the identification of deletions in the reeler mouse, which is noted for its inverted cortex and disorganised hippocampus and cerebellum (D'Arcangelo et al., 1995). More recently, the catalogue of neuroanatomical abnormalities in the reeler mouse has been extended to the superior colliculus (SC). In vertebrates, the SC consists of seven layers that are both anatomically and functionally organised. The superficial SC consists of the three uppermost layers: the zonal (Zn), superficial grey (SuG) and the optic layer (Op), and the deep SC contains four layers: the intermediate grey (InG), intermediate white (InW), deep grey (DpG) and deep white (DpW). In the reeler mouse, it has been reported that the superficial layers of this structure are cytoarchitectually and myeloarchitectually disorganised (Baba et al., 2007). Similarly, disruption of the laminar patterning in the SC has been observed in the reelin-deficient shaking rat Kawasaki (Sakakibara et al., 2003).

Given that both mutations in reelin and TUBA1A cause similar phenotypes in humans, in this article, we set out to investigate the cytoarchitecture of the SC in the Jna/+ mice. Using histological tools, we found that the laminar structure of the SC in mutant animals was intact; however, it was significantly thinner with an apparent fusion of the InG and InW. Using birthdate labelling, we showed that the neuronal migration defect that is observable in the cortex and the hippocampus of affected animals is also apparent during the formation of the SC. Additionally, an elevated rate of cell death leads to a significant loss of neurons in the SC of the Jna/+ mouse between postnatal day 21 (P21) and 12 weeks of age, with the majority of the loss occurring in the deep layers. Consistent with the role of the SC in modulating sensorimotor gating, we observed an exaggeration of the acoustic startle response in mutant animals.

Experimental procedures

Animals

Mice were maintained on a C3H/HeH (Harlan, UK) background and housed on a 12:12 light:dark cycle at a temperature of 22±1 °C and humidity of 60%–70%. Males and females were separated at weaning (P21) and housed separately in groups of five where possible. The genotype of animals was determined by polymerase chain reaction analysis, as previously described (Keays et al., 2007), and only littermates were selected for experiments. Cages were environmentally enriched with cardboard tubing, and mice were permitted ad libitum access to food. Experiments were performed in accordance with the UK Animals (Scientific Procedures) Act 1986.

5-bromo-2-deoxyuridine (BrdU) labelling

For birthdate labelling experiments, pregnant C3H females were injected with BrdU (50 μg/g of body weight), 12.5 and 13.5 days after copulation. All resultant P0 pups were killed and genotyped. P0 brains were extracted and drop-fixed in 4% paraformaldehyde for 4 h before being placed in sucrose overnight. Brains were then embedded in OCT and sectioned at 14 μm using a cryostat, then mounted onto electrostatic slides and stored at −20 °C. Prior to staining, antigen retrieval was performed in citrate buffer at 90 °C (0.01 M, pH 6). To quench peroxidase activity, slides were placed in 3% H2O2 solution for 10 min, followed by three washes in PBS. Sections were then digested in trypsin (0.0125%) for 10 min at 37 °C, washed three times in PBS, followed by a 20-min incubation in 2 N HCl at 37 °C. After washing, slides were incubated in a 0.3% Triton/PBS with 2% rabbit serum (Rat Elite ABC Kit, Vector Labs, Peterborough, UK) and 1:100 rat anti-human BrdU antibody (Accurate Chemical & Scientific, Westbury, NY, USA) in a humidity chamber overnight. Slides were then washed and stained as directed in the Elite ABC Kit, using DAB permanent staining.

Gallyas staining

Twelve-week-old Jna/+ mice and wild-type (WT) littermates were perfused with 0.9% saline and 10% formalin solution. Brains were extracted and left in 10% formalin solution for 2 weeks. Subsequently, brains were cryoprotected in 30% sucrose/formalin and sectioned on a freezing microtome (25 μm) before storing in 5% formalin solution at 4 °C for a further 2 weeks. Sections were matched, mounted on electrostatically charged slides and stained using a Gallyas silver staining method based on study by Pistorio et al. (2006).

In situ hybridisation

A 429-base pair probe targeting the tumour protein D52-like-1 (Tpd52L1) gene was generated by PCR using the following primers: Tpd521l_F GAAAGTAGGTGGGACAAACCAC and Tpd521l_R GGAGACCAAGTCAAAACCAAAG and cloned into a pCR2.1-TOPO vector (Invitrogen, CA, USA). Digoxigenin-labelled riboprobes were then synthesized using appropriate RNA polymerases (Roche Applied Science, Burgess Hill, UK), and hybridisation was performed, as previously described (Isaacs et al., 2003). All hybridisations were performed on 14-μm brain sections prepared from Jna/+ mice and WT littermates aged 12 weeks.

Juvenile and adult immunohistochemistry

Cohorts of P21 and 12-week-old Jna/+ and WT littermate mice (n=4) were perfused with 0.9% NaCl and 4% paraformaldehyde, and brains were postfixed for 6 h, followed by dehydration in 30% sucrose solution. Sections (40 μm) were prepared on a freezing microtome and stored in antifreeze solution at −20 °C. For each P21 and 12 week brain, three matched SC sections were selected for staining. Sections were incubated with the primary antibody overnight in 0.3% Triton/PBS with 2% of the appropriate serum at the following concentrations: NeuN (1:500) (Millipore, Billerica, MA, USA); Calbindin (1:500) (Millipore, Billerica, MA, USA); Er81 (1:4000). After three washes in PBS (5 min each), sections were incubated with a biotinylated secondary antibody (1:500) in 0.3% Triton/PBS for 2 h. After a series of washes in PBS, sections were incubated for 1 h with fluorescein-conjugated streptavidin (1:500) (Vector Labs, Peterborough, UK). Sections were mounted on electrostatic slides andcoverslipped with 4',6-diamidino-2-phenylindole (DAPI) containing mounting media (Vector Labs, Peterborough, UK).

Apoptosis study

Seven-week-old mice Jna/+ and WT littermate mice were perfused and sectioned in the aforementioned manner (n=5). Every 8th section from the beginning of the SC through to the start of the inferior colliculus was mounted on electrostatic slides. Prior to staining, slides underwent antigen retrieval, peroxidase quenching and PBS washing. Sections were incubated with the caspase 3 primary antibody (1:400) (Cell Signaling Technology, Boston, MA, USA) overnight in 0.3% Triton/PBS with 2% goat serum. Slides were then washed and stained, as directed in the Elite ABC Kit, using DAB permanent staining. Total cell counts were then obtained by multiplying the number of positive cells observed by a factor of 8.

Quantification and statistics

The SC was analysed through the creation of 10 zonal analysis boxes in which cells could be counted. Boxes were drawn on SC images using ImageJ freeware, were positioned 100 μm from the midline and were a consistent 250 μm in width. The boxes were of equal height within an image, with the height of each box being a 10th of the distance between the surface of the Zn to a depth in the periaqueductal grey (PAG) level with the cerebellar aqueduct. Once the analysis boxes were formatted, labelled cells within each box were counted manually. The percentage of neurons within each box was calculated from the number of NeuN staining cells as a proportion of DAPI cells. Statistics were performed in GraphPad Prism. A 1-way ANOVA with a Bonferroni correction for multiple comparisons was used for the analysis of behavioural measurements and for NeuN, BrdU and DAPI cell density zonal measurements.

Behavioural testing

The acoustic startle response was measured using a commercially available behavioural testing system (San Diego Instruments, San Diego, CA, USA). This apparatus consisted of a sound-proof box with a 6-cm speaker to deliver the acoustic stimuli, coupled to an accelerometer to monitor the animal's movement. The behavioural testing was performed in a separate room, and the cages were moved into the room at least 2 h before the start of the experiment. Prior to testing, each mouse was placed inside the startle apparatus and allowed to acclimatise for 5 min. To test the acoustic startle response, acoustic stimuli were presented as 40-ms impulses of white noise with four different intensities (90, 100, 110 and 120 dB) for a total of 20 times, each in a pseudorandom order and spaced at random intervals between 10 and 20 s. Responses were expressed in arbitrary units and averaged for each type of trial.

Results

Structure of the SC in the Jna/+ mouse

When dissecting brains from adult Jna/+ mutants, it is immediately apparent that the SC is more exposed in comparison to control animals (Fig. 1A). A sagittal Nissl stain reveals that it is also elongated in the rostrocaudal plane in comparison with WT controls (Figs. 1F, G), and a coronal Nissl stain (Fig. 1B) demonstrates that it is thinner both at P0 (F=18.16, P<0.001) and 12 weeks of age (F=102.41, P<0.0001) (Fig. 1C). To investigate the laminar structure of the SC, we began by searching for markers that selectively label different cellular populations. We tested commonly used cortical makers (Cux-1, Foxp2, ER81, calbindin, calretinin) and searched the Allen Brain Atlas for genes that showed layer-specific expression (Lein et al., 2007). We found that calretinin, which labels GABAergic interneurons in layers 2/3 of the mouse cortex labels the superficial OP layers (Park et al., 2002); TPD52L1, a cell cycle-regulated protein labels cells in the proximity of the InG (Boutros and Byrne, 2005) and Er81, a transcription factor which labels layer V neurons in the cortex, localised slightly deeper to InG/InW (Watakabe et al., 2007). Nissl staining, immunohistochemistry (Er81 and calretinin) and in situ hybridisation (TPD52L1) revealed that, despite some dispersion of Er81 and calbindin-positive cells, the laminar structure of the SC in Jna/+ mutants is essentially intact (Fig. 1E). Next, we used a Gallyas silver stain to investigate the myeloarchitecture of the colliculus (Gallyas, 1979) (Fig. 2A). In WT animals, a clear distinction could be made between the various grey and white layers, the dark silver stain labelling the white matter, particularly the Op layer, and the large myelinated cells in the InW (Fig. 2C). In mutants, the definition between superficial and intermediate layers was less defined, exemplified by the fusion of the InG layer with the InW layers (Fig. 2D). The SC commissure that is observable in the DpW was observed to cross the midline in both WT controls and Jna/+ mutants (Figs. 2E, F).

Fig. 1.

Fig. 1

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The structure of the SC in Jna/+ mutants. (A) Dorsal view of wild-type (WT) and Jna/+ brains. The red arrow highlights the exposed SC in mutant animals. (B) Coronal Nissl stain of WT and mutant animals illustrating the thinner SC in Jna/+ mutants. (C) Quantification of the thickness of the SC in WT and mutant animals at P0 and 12 wk reveals a significant difference between controls and Jna/+ animals (F=18.1, P<0.001, F=102.4, P<0.0001). (D) Diagram illustrating the laminar structure of the SC adapted from Paxinos and Watson's Brain Atlas (Paxinos et al., 2007). The SC consists of seven layers: the zonal layer (Zn), the superficial grey layer (SuG), the optic layer (Op), the intermediate grey layer (InG), the intermediate white layer (InW), the deep grey layer (DpG) and the deep white (DpW) layer. Other abbreviation: PAG, periaqueductal grey. (E) Calretinin, TPD52I1 and ER81 staining (top to bottom) reveal that the laminar structure of the SC is intact in Jna/+ mutants. (F, G) Sagittal sections of the SC in WT littermates and mutant animals reveal that it is elongated in Jna/+ mutants (G). Red arrow between dotted black lines shows the SC. Scale bars 5 mm (A), 1 mm (B, F, G) and 250 μm (E).

Fig. 2.

Fig. 2

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Myeloarchitectural investigation of the SC in Jna/+ mutants. (A) Diagram illustrating the laminar structure of the SC adapted from Paxinos and Watson's Brain Atlas (Paxinos et al., 2007). (B) Diagram showing the layers of the SC aligned with Gallyas-stained WT and mutant sections. Equally proportioned analysis boxes (shown in white) were drawn from the aqueduct to the dorsal surface of Zn layer. These boxes map to similar anatomical regions. Boxes 1–5 align with the superficial and intermediate layers (Zn, SuG, Op InG, InW), Boxes 6, 7 the deep layers (DpG, DpW) and Boxes 8–10 the PAG. (C, D) Enlarged images of the superficial and intermediate layers of Gallyas-stained sections in WT and mutant animals. The superficial SC of the mutant appears compacted and less defined with a fusion of the InG and InW (white arrow). (E, F) Enlarged images of the deep layers (DpG and DpW) of Gallyas-stained sections in WT and mutant animals. Sections appear similar, with crossing of the collicular commissure observed in both mutants and WT animals. Scale bars show 250 μm.

Defective neuronal migration in the Jna/+ SC

We then examined whether the anatomical abnormalities observed in the SC might be associated with a defect in neuronal migration. The laminar structure of the SC differs from the neocortex, as most GABAergic interneurons migrate radially, and there is a temporal overlap in neurogenesis between deep and superficial layer neurons (Tan et al., 2002; Tsunekawa et al., 2005). The peak generation time for deep layer neurons (which migrate in an inside-out manner) is E11 to E13, and for superficial layer neurons (which migrate in a more complex pattern) is E12 to E13 (Altman and Bayer, 1981; Edwards et al., 1986). We labelled newly born cells with BrdU at E12.5 and E13.5. Harvesting mice at their day of birth, we counted labelled cells, having divided the SC into 10 equal zones extending from the aqueduct to the surface of the colliculus (Figs. 3A, B). We tested for an interaction between genotype and the mean percentage of cells in each zone. When labelling at E12.5, we found that the percentage of labelled cells present in the two most superficial analysis boxes was significantly reduced in Jna/+ mutants (n=4) (F=20.3, P<0.0001; F=26.96, P<0.0001 [zones 1 and 2, respectively]). This was complemented by a significantly higher percentage of BrdU-labelled cells present in zone 6 (a region that includes the DpG) in Jna/+ mutants (F=11.78, P<0.01) (Figs. 3A, E). When labelling at E13.5, which labels a higher percentage of superficial neurons, we found a significant reduction in the portions of BrdU-labelled cells in zones 2 (F=10.56, P<0.05) and 3 (F=18.32, P<0.001) and a higher portion in zones 6 (F=20.9, P<0.0001) and 7 (F=15.39, P<0.01) in Jna/+ animals (n=3) (Figs. 3B, F). These results are indicative of a defect in neuronal migration in Jna/+ mutants. Overall, we observed no significant difference in the total number of BrdU-labelled cells when comparing WT and Jna/+ mutants when labelling at E12.5 (F=0.56, P>0.5) or E13.5 (F=1.19, P>0.1) suggesting that the rate at which neurons are generated is not affected by the S140G mutation (Figs. 3C, D).

Fig. 3.

Fig. 3

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Impaired neuronal migration in the SC of Jna/+ mutants. (A, B) Staining for BrdU in the SC of littermate controls and Jna/+ mutants harvested at P0 after injection of BrdU at E12.5 (n=4) and E13.5 (n=3). The SC was divided into 10 equal bins (shown in white) extending from the aqueduct to the dorsal surface of Zn layer and BrdU-positive cells were counted. (C, D) There was no significant difference in the average number of cells labelled per section between WT littermates and Jna/+ mutants. (E, F) The distribution of BrdU-positive cells per zone in WT controls and Jna/+ mutants after injections at E12.5 (E) and E13.5 (F). The percentage of BrdU cells in the upper two zones was significantly reduced in Jna/+ mutants when injected at E12.5 (layer 1 [F=20.3, P<0.001], layer 2 [F=27, P<0.001]), and higher in layer 6 (F=11.78, P<0.01). Similarly, when injected at E13.5, the percent of BrdU-labelled cells in zones 2 (F=10.56, P<0.05) and 3 (F=18.32, P<0.001) was reduced in Jna/+ animals (n=3), complemented by a higher portion observed in zones 6 (F=20.9, P<0.0001) and 7 (F=15.39, P<0.01) (n=3). (G) Staining of postmitotic neurons with the marker NeuN reveals no significant difference in the percentage of neurons per a zone at P21. Scale bars in A and B show 250 μm.

Next, we examined the distribution of neurons in the SC in juvenile animals by staining serial sections with the postmitotic marker NeuN at P21 (n=4), again dividing the SC into 10 zones. We tested for an interaction between genotype and the mean percentage of NeuN-positive cells in each zone. Although we observed a higher percentage of neurons in the deep layers in mutant animals, this difference was not significant (Fig. 3G). These results suggest that either the neurons delayed during development catch up between P0 and P21, or alternatively, the SC in mutant animals is remodelled by differential rates of apoptosis in the deep, intermediate and superficial zones (Finlay et al., 1982; Cowan et al., 1984).

Loss of neurons in the adult SC in Jna/+ mutants

Next, we assessed how the S140G mutation affects the survival of neurons within the layers of the SC at P21 and 12 weeks of age. We once again divided the colliculus into 10 equal bins extending from the aqueduct to the surface, and counted the number of NeuN-positive and DAPI-labelled cells (n=4) (Figs. 4A, B). At P21, we found that both mutant and littermate control animals had similar percentages of neurons (50%) in each zone (Fig. 4C). This contrasts with our observations at 12 weeks of age, where we found a large reduction in the percentage of neurons in all zones in mutant animals (Fig. 4D). The greatest loss of neurons occurred in zones 6 and 7 in the vicinity of the DpG and DpW (F=33.07, P<0.0001; F=41.57, P<0.0001). Additionally, zones 8–10, which cover the PAG showed a significant drop in the percentage of neurons (F=32.58, P<0.0001; F=18.87, P<0.001; F=28.2, P<0.0001). To demonstrate that this loss of neurons is due to cell death, we immunostained coronal sections of the SC with an antibody specific for caspase-3, an activated protease that labels apoptotic cells (Porter and Janicke, 1999). We performed this experiment on mice aged 7 weeks of age (n=5), a time point midway between P21 and 12 weeks (Figs. 4E, F). Staining of serial sections revealed that there was a significant increase in both the number (F=21.2, P<0.01) and density (F=72.1, P<0.0001) of caspase-3-positive cells in Jna/+ mutants (Figs. 4G, H).

Fig. 4.

Fig. 4

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Loss of neurons and increased apoptosis in the SC of Jna/+ mutants. (A, B) The SC was divided into 10 equal bins and DAPI (blue) and NeuN (green)-labelled cells counted at P21 and 12 wk of age for both WT littermates and Jna/+ mutants (n=4). This enabled the percentage of neurons to be calculated for each zone at each time point (C, D). We compared the percentage of neurons in WT littermates and Jna/+ mutants and found no significant difference between genotypes for any zone in the P21 cohort (C). In contrast, there was a massive reduction in the percentage of neurons throughout the SC in 12-wk-old Jna/+ mutants (D). The loss of neurons reached significance in all zonal boxes, particularly those mapping to the DpG/DpW layers (zone 6 [F=33.07, P<0.001], zone 7 [F=41.57, P<0.001]). (E, F) Immunostaining of the SC of 7-wk-old animals revealed a significant increase in the total number (F=21.2, P<0.01) and density (F=72.1, P<0.0001) of caspase-positive cells (black arrows) in Jna/+ mutants (n=5) when compared with WT littermates (G, H). Scale bars show 250 μm.

Exaggerated acoustic startle response in Jna/+ mutants

Inhibitory cells in the deep layers of the SC have been implicated in mediation of the acoustic startle response (Meloni and Davis, 2000), a behaviour that is characterised by the involuntary contraction of muscles in response to a sudden acoustic stimulus (Koch, 1999). Therefore, we tested the acoustic startle response in Jna/+ mutants by exposing them to 40-ms impulses of white noise with four different intensities (90, 100, 110 and 120 dB) in a pseudorandom order and spaced at random intervals between 10 and 20 s. We found that in the absence of a weight correction, the acoustic startle response is significantly higher in both males and female Jna/+ mutants when exposed to a 120 dB stimulus (male [F=56.1, P<0.0001], female [F=48.08, P<0.0001]) (Figs. 5A, B). When normalising for the reduced mass of Jna/+ mutants, we found that mutant animals of both sexes display a significantly larger acoustic startle response to both 110 dB (male [F=17.85, P<0.001], female [F=15.5, P<0.001]) and 120 dB stimuli (male [F=134.8, P<0.0001], female [F=96.8, P<0.0001]) (Figs. 5C, D).

Fig. 5.

Fig. 5

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Exaggerated acoustic startle response in Jna/+ mutants. (A, B) Assessment of the acoustic startle response in male (n=7) and female Jna/+ mutants (n=8) in response to randomised 40 ms bursts of 90 dB, 100 db, 110 db and 120 db white noise. In the absence of a weight correction, the acoustic startle response is significantly higher in both males and female Jna/+ mutants when exposed to a 120 dB stimulus (male [F=56.1, P<0.0001], female [F=48.08, P<0.0001]). (C, D) When these data are normalised to take into account the differences in weight between genotypes, mutant animals of both sexes display a significantly larger acoustic startle response to 110 dB (male [F=17.85, P<0.001], female [F=15.5, P<0.001]) and 120 dB stimuli (male [F=134.8, P<0.0001], female [F=96.8, P<0.0001]).

Discussion

In this article, we have shown that the S140G mutation in Tuba1a mutant animals results in a cytoarchitecturally disrupted SC. Although the general lamination of the SC is intact, our studies have revealed a marked thinning of the SC, with a fusion of the InG layer with the InW layers. Birthdate labelling at E12.5 and E13.5, followed by examination at P0, revealed a severe impairment in the radial migration of neurons. Cell counting using the neuronal marker NeuN demonstrated that the distribution of neurons at P21 in mutant animals was comparable with WT controls suggesting postnatal rectification of migration, or alternatively, remodelling of the SC coincident with axonal innervation (Finlay et al., 1982; Cowan et al., 1984). A quantitative assessment of neuronal number in adulthood showed that there was a massive reduction in the percentage of postmitotic neurons in mutant animals, which we attribute to an increase in neuronal apoptosis between P21 and 12 weeks of age in Jna/+ mutants. Consistent with the role of the SC in modulating sensorimotor gating, and the circuitry that modulates this behaviour, we find that Jna/+ mutants exhibit an exaggerated acoustic startle response.

What underlying cellular defect might cause these phenotypes? Our results suggest that the thinning of the SC, as well as the reduction of neurons in adulthood, is unlikely to be a result of a defect in mitotic division. We observe comparable numbers of labelled cells in WT controls and Jna/+ mutants after injections of BrdU at E12.5 and E13.5, and all currently available evidence suggests that Tuba1a is limited in its expression to postmitotic neurons (Bamji and Miller, 1996; Gloster et al., 1999; Keays et al., 2010). Instead, we implicate increased cell death in the pathophysiology of the tubulinopathies showing a massive increase in apoptosis in the adult SC in Jna/+ mutants. Our mouse data are consistent with observations made in human patients with mutations in TUBA1A. Bahi-Buisson and colleagues have reported five patients with mutations in TUBA1A who presented with severe microcephaly (R264C, L397P, R422C, G436R, R422H) despite normal biparietal diameters in the third trimester (Bahi-Buisson et al., 2008), and Morris-Rosendahl and colleagues have identified two patients with mutations in TUBA1A (R402L, E55K) who again exhibit progressive microcephaly (Morris-Rosendahl et al., 2008). Taken together with our mouse data, we conclude that mutations in Tuba1a can lead to an increase in neuronal apoptosis.

How might a mutation in Tuba1a cause an increase in neuronal apoptosis? The molecular cascade that regulates cell death involves a host of proteins that include members of the caspase family, Fas, Bcl, Pten and Bax (Merry et al., 1994; Vekrellis et al., 1997; Cheema et al., 1999; Groszer et al., 2006). An attractive candidate for mediation of apoptosis in the Tuba1a mutant mice is the GTPase, RhoA. RhoA, along with a host of other small GTP-binding proteins that include Rnd2, Rac1 and Cdc42, acts within multiple signalling pathways that converge on the actin and microtubule cytoskeletons to regulate neuronal morphology, adhesion and migration (Luo, 2000; Govek et al., 2005). Tucker and colleagues have extended the functional repertoire of RhoA, recently implicating it in the control of postnatal apoptosis in the vertebrate brain (Sanno et al., 2010). Using a dominant-negative genetic inhibitor of the Rho GTPases, they showed that inhibition of Rho GTPases results in a large increase in the number and density of neurons in the somatosensory cortex of mice, a consequence of decreased apoptosis. Cellular studies attributed this affect to RhoA, which was found to cause increased apoptosis when overexpressed in cultured cortical neurons. Sitting at a molecular intersection between the cytoskeleton and an apoptotic signalling pathway, it is possible that in response to microtubule dysfunction, activated RhoA may mediate programmed cell death in Jna/+ mutants (Wittmann and Waterman-Storer, 2001).

We have shown that the loss of neurons, as well as the cytoarchitectural abnormalities apparent in the SC, is accompanied by an increase in the acoustic startle response in Jna/+ mutant mice. The acoustic startle response is mediated by a well-characterised circuit that consists of the auditory nerve, the ventral cochlear nucleus, the dorsal nucleus of the lateral lemniscus, the caudal pontine reticular nucleus, spinal interneurons and spinal motor neurons (Davis et al., 1982; Koch, 1999). Neurons located deep in the SC, where we observe the greatest loss of neurons in Jna/+ mutants, receive GABAnergic input from the substantia nigra and synapse directly with those in the caudal pontine reticular nucleus modulating the startle response (Shammah-Lagnado et al., 1987; Meloni and Davis, 1999, 2000). Although we cannot exclude the presence of anatomical or functional defects associated with other elements of this circuit, the cell loss and cytoarchitectural disruption of the SC we observe in the Jna/+ mouse are consistent with the abnormal acoustic startle response we have described in this article.

This study has shown that the S140G mutation in the Jna/+ mouse results in an impairment of neuronal migration in the developing SC, an increase in adult neuronal apoptosis and an enlarged acoustic startle response. These observations raise several interesting questions. First, it provides additional evidence for the importance of Tubala1a for neuronal migration, prompting the question: is Tuba1a vital for all forms of neuronal migration? Might Tuba1a also be required for the correct migration of peripheral neurons and those that form the precise lamination in the retina? Second, is the increased neuronal apoptosis we observe in the SC, a more global phenomenon? Might we also observe increased cell death in the adult telencephalon in Jna/+ mutants? Third, one is compelled to ask whether the acoustic startle response and SC are also abnormal in human patients who harbour mutations in Tuba1a?

Acknowledgments

We are grateful to Dr. Thomas Jessel for his gift of ER81 antibody. This work is supported by a grant from Le Fondation Jérôme Lejeune and the Wellcome Trust. DAK is supported by an FWF grant (P21092).

References

  1. Altman J., Bayer S.A. Time of origin of neurons of the rat superior colliculus in relation to other components of the visual and visuomotor pathways. Exp Brain Res. 1981;42:424–434. doi: 10.1007/BF00237507. [DOI] [PubMed] [Google Scholar]
  2. Baba K., Sakakibara S., Setsu T., Terashima T. The superficial layers of the superior colliculus are cytoarchitectually and myeloarchitectually disorganized in the reelin-deficient mouse, reeler. Brain Res. 2007;1140:205–215. doi: 10.1016/j.brainres.2006.09.113. [DOI] [PubMed] [Google Scholar]
  3. Bahi-Buisson N., Poirier K., Boddaert N., Saillour Y., Castelnau L., Philip N., Buyse G., Villard L., Joriot S., Marret S., Bourgeois M., Van Esch H., Lagae L., Amiel J., Hertz-Pannier L., Roubertie A., Rivier F., Pinard J.M., Beldjord C., Chelly J. Refinement of cortical dysgeneses spectrum associated with TUBA1A mutations. J Med Genet. 2008;45:647–653. doi: 10.1136/jmg.2008.058073. [DOI] [PubMed] [Google Scholar]
  4. Bamji S.X., Miller F.D. Comparison of the expression of a T alpha 1: nlacZ transgene and T alpha 1 alpha-tubulin mRNA in the mature central nervous system. J Comp Neurol. 1996;374:52–69. doi: 10.1002/(SICI)1096-9861(19961007)374:1<52::AID-CNE4>3.0.CO;2-M. [DOI] [PubMed] [Google Scholar]
  5. Boutros R., Byrne J.A. D53 (TPD52L1) is a cell cycle-regulated protein maximally expressed at the G2-M transition in breast cancer cells. Exp Cell Res. 2005;310:152–165. doi: 10.1016/j.yexcr.2005.07.009. [DOI] [PubMed] [Google Scholar]
  6. Boycott K.M., Flavelle S., Bureau A., Glass H.C., Fujiwara T.M., Wirrell E., Davey K., Chudley A.E., Scott J.N., McLeod D.R., Parboosingh J.S. Homozygous deletion of the very low density lipoprotein receptor gene causes autosomal recessive cerebellar hypoplasia with cerebral gyral simplification. Am J Hum Genet. 2005;77:477–483. doi: 10.1086/444400. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Cheema Z.F., Wade S.B., Sata M., Walsh K., Sohrabji F., Miranda R.C. Fas/Apo [apoptosis]-1 and associated proteins in the differentiating cerebral cortex: induction of caspase-dependent cell death and activation of NF-kappaB. J Neurosci. 1999;19:1754–1770. doi: 10.1523/JNEUROSCI.19-05-01754.1999. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Cowan W.M., Fawcett J.W., O'Leary D.D., Stanfield B.B. Regressive events in neurogenesis. Science. 1984;225:1258–1265. doi: 10.1126/science.6474175. [DOI] [PubMed] [Google Scholar]
  9. D'Arcangelo G., Miao G.G., Chen S.C., Soares H.D., Morgan J.I., Curran T. A protein related to extracellular matrix proteins deleted in the mouse mutant reeler. Nature. 1995;374:719–723. doi: 10.1038/374719a0. [DOI] [PubMed] [Google Scholar]
  10. Davis M., Gendelman D.S., Tischler M.D., Gendelman P.M. A primary acoustic startle circuit: lesion and stimulation studies. J Neurosci. 1982;2:791–805. doi: 10.1523/JNEUROSCI.02-06-00791.1982. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. des Portes V., Pinard J.M., Billuart P., Vinet M.C., Koulakoff A., Carrie A., Gelot A., Dupuis E., Motte J., Berwald-Netter Y., Catala M., Kahn A., Beldjord C., Chelly J. A novel CNS gene required for neuronal migration and involved in X-linked subcortical laminar heterotopia and lissencephaly syndrome. Cell. 1998;92:51–61. doi: 10.1016/s0092-8674(00)80898-3. [DOI] [PubMed] [Google Scholar]
  12. Edwards M.A., Caviness V.S., Jr, Schneider G.E. Development of cell and fiber lamination in the mouse superior colliculus. J Comp Neurol. 1986;248:395–409. doi: 10.1002/cne.902480308. [DOI] [PubMed] [Google Scholar]
  13. Finlay B.L., Berg A.T., Sengelaub D.R. Cell death in the mammalian visual system during normal development: II: Superior colliculus. J Comp Neurol. 1982;204:318–324. doi: 10.1002/cne.902040403. [DOI] [PubMed] [Google Scholar]
  14. Gallyas F. Silver staining of myelin by means of physical development. Neurol Res. 1979;1:203–209. doi: 10.1080/01616412.1979.11739553. [DOI] [PubMed] [Google Scholar]
  15. Gleeson J.G., Allen K.M., Fox J.W., Lamperti E.D., Berkovic S., Scheffer I., Cooper E.C., Dobyns W.B., Minnerath S.R., Ross M.E., Walsh C.A. Doublecortin, a brain-specific gene mutated in human X-linked lissencephaly and double cortex syndrome, encodes a putative signaling protein. Cell. 1998;92:63–72. doi: 10.1016/s0092-8674(00)80899-5. [DOI] [PubMed] [Google Scholar]
  16. Gloster A., El-Bizri H., Bamji S.X., Rogers D., Miller F.D. Early induction of Talpha1 alpha-tubulin transcription in neurons of the developing nervous system. J Comp Neurol. 1999;405:45–60. doi: 10.1002/(sici)1096-9861(19990301)405:1<45::aid-cne4>3.0.co;2-m. [DOI] [PubMed] [Google Scholar]
  17. Govek E.E., Newey S.E., Van Aelst L. The role of the Rho GTPases in neuronal development. Genes Dev. 2005;19:1–49. doi: 10.1101/gad.1256405. [DOI] [PubMed] [Google Scholar]
  18. Groszer M., Erickson R., Scripture-Adams D.D., Dougherty J.D., Le Belle J., Zack J.A., Geschwind D.H., Liu X., Kornblum H.I., Wu H. PTEN negatively regulates neural stem cell self-renewal by modulating G0-G1 cell cycle entry. Proc Natl Acad Sci U S A. 2006;103:111–116. doi: 10.1073/pnas.0509939103. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Guerrini R., Marini C. Genetic malformations of cortical development. Exp Brain Res. 2006;173:322–333. doi: 10.1007/s00221-006-0501-z. [DOI] [PubMed] [Google Scholar]
  20. Hong S.E., Shugart Y.Y., Huang D.T., Shahwan S.A., Grant P.E., Hourihane J.O., Martin N.D., Walsh C.A. Autosomal recessive lissencephaly with cerebellar hypoplasia is associated with human RELN mutations. Nat Genet. 2000;26:93–96. doi: 10.1038/79246. [DOI] [PubMed] [Google Scholar]
  21. Isaacs A.M., Oliver P.L., Jones E.L., Jeans A., Potter A., Hovik B.H., Nolan P.M., Vizor L., Glenister P., Simon A.K., Gray I.C., Spurr N.K., Brown S.D., Hunter A.J., Davies K.E. A mutation in Af4 is predicted to cause cerebellar ataxia and cataracts in the robotic mouse. J Neurosci. 2003;23:1631–1637. doi: 10.1523/JNEUROSCI.23-05-01631.2003. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Keays D.A., Cleak J., Huang G.J., Edwards A., Braun A., Treiber C.D., Pidsley R., Flint J. The role of Tuba1a in adult hippocampal neurogenesis and the formation of the dentate gyrus. Dev Neurosci. 2010;32:268–277. doi: 10.1159/000319663. [DOI] [PubMed] [Google Scholar]
  23. Keays D.A., Tian G., Poirier K., Huang G.J., Siebold C., Cleak J., Oliver P.L., Fray M., Harvey R.J., Molnar Z., Pinon M.C., Dear N., Valdar W., Brown S.D., Davies K.E., Rawlins J.N., Cowan N.J., Nolan P., Chelly J., Flint J. Mutations in alpha-tubulin cause abnormal neuronal migration in mice and lissencephaly in humans. Cell. 2007;128:45–57. doi: 10.1016/j.cell.2006.12.017. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Koch M. The neurobiology of startle. Prog Neurobiol. 1999;59:107–128. doi: 10.1016/s0301-0082(98)00098-7. [DOI] [PubMed] [Google Scholar]
  25. Lein E.S., Hawrylycz M.J., Ao N., Ayres M., Bensinger A., Bernard A., Boe A.F., Boguski M.S., Brockway K.S., Byrnes E.J., Chen L., Chen L., Chen T.M., Chin M.C., Chong J., Crook B.E., Czaplinska A., Dang C.N., Datta S., Dee N.R., Desaki A.L., Desta T., Diep E., Dolbeare T.A., Donelan M.J., Dong H.W., Dougherty J.G., Duncan B.J., Ebbert A.J., Eichele G., Estin L.K., Faber C., Facer B.A., Fields R., Fischer S.R., Fliss T.P., Frensley C., Gates S.N., Glattfelder K.J., Halverson K.R., Hart M.R., Hohmann J.G., Howell M.P., Jeung D.P., Johnson R.A., Karr P.T., Kawal R., Kidney J.M., Knapik R.H., Kuan C.L., Lake J.H., Laramee A.R., Larsen K.D., Lau C., Lemon T.A., Liang A.J., Liu Y., Luong L.T., Michaels J., Morgan J.J., Morgan R.J., Mortrud M.T., Mosqueda N.F., Ng L.L., Ng R., Orta G.J., Overly C.C., Pak T.H., Parry S.E., Pathak S.D., Pearson O.C., Puchalski R.B., Riley Z.L., Rockett H.R., Rowland S.A., Royall J.J., Ruiz M.J., Sarno N.R., Schaffnit K., Shapovalova N.V., Sivisay T., Slaughterbeck C.R., Smith S.C., Smith K.A., Smith B.I., Sodt A.J., Stewart N.N., Stumpf K.R., Sunkin S.M., Sutram M., Tam A., Teemer C.D., Thaller C., Thompson C.L., Varnam L.R., Visel A., Whitlock R.M., Wohnoutka P.E., Wolkey C.K., Wong V.Y. Genome-wide atlas of gene expression in the adult mouse brain. Nature. 2007;445:168–176. doi: 10.1038/nature05453. [DOI] [PubMed] [Google Scholar]
  26. Luo L. Rho GTPases in neuronal morphogenesis. Nat Rev Neurosci. 2000;1:173–180. doi: 10.1038/35044547. [DOI] [PubMed] [Google Scholar]
  27. Meloni E.G., Davis M. Muscimol in the deep layers of the superior colliculus/mesencephalic reticular formation blocks expression but not acquisition of fear-potentiated startle in rats. Behav Neurosci. 1999;113:1152–1160. doi: 10.1037//0735-7044.113.6.1152. [DOI] [PubMed] [Google Scholar]
  28. Meloni E.G., Davis M. GABA in the deep layers of the superior Colliculus/Mesencephalic reticular formation mediates the enhancement of startle by the dopamine D1 receptor agonist SKF 82958 in rats. J Neurosci. 2000;20:5374–5381. doi: 10.1523/JNEUROSCI.20-14-05374.2000. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Merry D.E., Veis D.J., Hickey W.F., Korsmeyer S.J. bcl-2 protein expression is widespread in the developing nervous system and retained in the adult PNS. Development. 1994;120:301–311. doi: 10.1242/dev.120.2.301. [DOI] [PubMed] [Google Scholar]
  30. Morris-Rosendahl D.J., Najm J., Lachmeijer A.M., Sztriha L., Martins M., Kuechler A., Haug V., Zeschnigk C., Martin P., Santos M., Vasconcelos C., Omran H., Kraus U., Van der Knaap M.S., Schuierer G., Kutsche K., Uyanik G. Refining the phenotype of alpha-1a Tubulin (TUBA1A) mutation in patients with classical lissencephaly. Clin Genet. 2008;74:425–433. doi: 10.1111/j.1399-0004.2008.01093.x. [DOI] [PubMed] [Google Scholar]
  31. Park H.J., Kong J.H., Kang Y.S., Park W.M., Jeong S.A., Park S.M., Lim J.K., Jeon C.J. The distribution and morphology of calbindin D28K- and calretinin-immunoreactive neurons in the visual cortex of mouse. Mol Cells. 2002;14:143–149. [PubMed] [Google Scholar]
  32. Paxinos G., Halliday G.M., Watson C., Koutcherov Y., Wang H. Academic Press; London: 2007. Atlas of the developing mouse brain. [Google Scholar]
  33. Pistorio A.L., Hendry S.H., Wang X. A modified technique for high-resolution staining of myelin. J Neurosci Methods. 2006;153:135–146. doi: 10.1016/j.jneumeth.2005.10.014. [DOI] [PubMed] [Google Scholar]
  34. Porter A.G., Janicke R.U. Emerging roles of caspase-3 in apoptosis. Cell Death Differ. 1999;6:99–104. doi: 10.1038/sj.cdd.4400476. [DOI] [PubMed] [Google Scholar]
  35. Sakakibara S., Misaki K., Terashima T. Cytoarchitecture and fiber pattern of the superior colliculus are disrupted in the Shaking Rat Kawasaki. Brain Res Dev Brain Res. 2003;141:1–13. doi: 10.1016/s0165-3806(02)00618-1. [DOI] [PubMed] [Google Scholar]
  36. Sanno H., Shen X., Kuru N., Bormuth I., Bobsin K., Gardner H.A., Komljenovic D., Tarabykin V., Erzurumlu R.S., Tucker K.L. Control of postnatal apoptosis in the neocortex by RhoA-subfamily GTPases determines neuronal density. J Neurosci. 2010;30:4221–4231. doi: 10.1523/JNEUROSCI.3318-09.2010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Shammah-Lagnado S.J., Negrao N., Silva B.A., Ricardo J.A. Afferent connections of the nuclei reticularis pontis oralis and caudalis: a horseradish peroxidase study in the rat. Neuroscience. 1987;20:961–989. doi: 10.1016/0306-4522(87)90256-9. [DOI] [PubMed] [Google Scholar]
  38. Tan S.S., Valcanis H., Kalloniatis M., Harvey A. Cellular dispersion patterns and phenotypes in the developing mouse superior colliculus. Dev Biol. 2002;241:117–131. doi: 10.1006/dbio.2001.0505. [DOI] [PubMed] [Google Scholar]
  39. Tsunekawa N., Yanagawa Y., Obata K. Development of GABAergic neurons from the ventricular zone in the superior colliculus of the mouse. Neurosci Res. 2005;51:243–251. doi: 10.1016/j.neures.2004.11.011. [DOI] [PubMed] [Google Scholar]
  40. Vekrellis K., McCarthy M.J., Watson A., Whitfield J., Rubin L.L., Ham J. Bax promotes neuronal cell death and is downregulated during the development of the nervous system. Development. 1997;124:1239–1249. doi: 10.1242/dev.124.6.1239. [DOI] [PubMed] [Google Scholar]
  41. Watakabe A., Ichinohe N., Ohsawa S., Hashikawa T., Komatsu Y., Rockland K.S., Yamamori T. Comparative analysis of layer-specific genes in Mammalian neocortex. Cereb Cortex. 2007;17:1918–1933. doi: 10.1093/cercor/bhl102. [DOI] [PubMed] [Google Scholar]
  42. Wittmann T., Waterman-Storer C.M. Cell motility: can Rho GTPases and microtubules point the way? J Cell Sci. 2001;114:3795–3803. doi: 10.1242/jcs.114.21.3795. [DOI] [PubMed] [Google Scholar]

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