New Results

Gsx2 is involved in specification of neurons in the inferior olivary nuclei from Ptf1a-expressing neuronal progenitors in zebrafish

Tsubasa Itoh, Miki Takeuchi, Marina Sakagami, Kazuhide Asakawa, Koichi Kawakami, Takashi Shimizu, Masahiko Hibi
doi: https://doi.org/10.1101/2020.03.16.993402

ABSTRACT

Neurons in the inferior olivary nuclei (IO neurons) send climbing fibers to Purkinje cells to elicit functions of the cerebellum. IO neurons and Purkinje cells are derived from neural progenitors expressing the proneural gene ptf1a. In this study, we found that the homeobox gene gsx2 was co-expressed with ptf1a in IO progenitors in zebrafish. Both gsx2 and ptf1a zebrafish mutants showed a strong reduction or loss of IO neurons. The expression of ptf1a was not affected in gsx2 mutants and vice versa. In IO progenitors, the ptf1a mutation increased apoptosis whereas the gsx2 mutation did not, suggesting that ptf1a and gsx2 are independently regulated and have distinct roles. The fibroblast growth factors (Fgf) 3/8a and retinoic acid signals negatively and positively, respectively, regulated gsx2 expression and thereby the development of IO neurons. mafba and hox genes are at least partly involved in the Fgf- and retinoic acid-dependent regulation of IO neuronal development. Our results indicate that gsx2 mediates the rostro-caudal positional signals to specify the identity of IO neurons from ptf1a-expressing neural progenitors.

Summary The homeobox gene gsx2 mediates rostro-caudal positional signaling to specify the identify of neurons in the inferior olivary nuclei from neural progenitors expressing the proneural gene ptf1a.

INTRODUCTION

In an initial step of the formation of neural circuits, neurons are differentiated from their neural/neuronal progenitors. The fate of neurons is thought to be determined by inductive signals that their progenitors receive depending on the position of the neural/neuronal progenitors. By receiving positional signals, neural progenitors express transcription factors, such as basic helix-loop-helix (bHLH) and homeobox transcription factors, that function in the differentiation and specification of neurons. Elucidation of these transcription factors and their regulation is a prerequisite to understanding the mechanism of neural circuit formation. Cerebellar neural circuits provide a good model for revealing positional signal-dependent neurogenesis.

In both mammals and teleosts, such as zebrafish, cerebellar neural circuits are composed of several types of neurons, including granule cells (GCs) and Purkinje cells (PCs) in the cerebellum, as well as neurons in the inferior olivary nuclei (IO neurons) in the caudal hindbrain. PCs receive axons from GCs (parallel fibers) and IO neurons (climbing fibers: CFs), then integrate the input information to elicit the cerebellum’s functions. PCs then send output to the outside of the cerebellum via efferent neurons, which are eurydendroid cells in zebrafish. In zebrafish, a portion of GCs also project their axons to dendrites of crest cells, which are PC-like cells located in the medial octavolateral nucleus (MON) of the dorsal hindbrain (Bae et al., 2009; Hashimoto and Hibi, 2012; Hibi and Shimizu, 2012). These neurons are derived from the hindbrain, which contains seven compartments collectively referred to as the rhombomere (r). PCs and IO neurons are generated from neural progenitors in the dorsal ventricular zone of the hindbrain. Analyses of mutants and cell lineage tracing for the bHLH-type proneural gene pancreas transcription factor 1a (Ptf1a) in mice revealed that PCs and IO neurons are derived from Ptf1a-expressing progenitors and Ptf1a is required for generation of these neurons in mice (Hoshino et al., 2005; Yamada et al., 2007). Lineage tracing with transgenic (Tg) zebrafish also showed that PCs and IO neurons are derived from ptf1a-expressing (ptf1a+) progenitors in zebrafish (Kani et al., 2010). Inhibitory neurons in the dorsal cochlear nuclei (DCN neurons) were also shown to be derived from Ptf1a+ neural progenitor cells in r2-5 in mice (Fujiyama et al., 2009). These reports indicate that despite being present in the dorsal ventricular zone of all rhombomeres, ptf1a+ progenitors generate different types of neurons depending on their position. At r1, PCs are generated from ptf1a+ progenitors and migrate dorsally whereas in the caudal hindbrain, IO neurons are generated from ptf1a+ progenitors and migrate ventrally (reviewed in (Hashimoto and Hibi, 2012; Hoshino, 2012)). However, it is not clear yet how positional signals are interpreted to specify neurons from ptf1a+ progenitors and what molecules control differentiation and/or specification of IO neurons.

Rostro-caudal patterning of the caudal hindbrain is controlled by gradients of fibroblast growth factor 3/8a (Fgf3/8a) and retinoic acid (RA) signals (Dupe and Lumsden, 2001; Gavalas and Krumlauf, 2000; Marin and Charnay, 2000; Maves et al., 2002; Maves and Kimmel, 2005). During early neurogenesis in zebrafish, fgf3/8a are expressed at r4 and are required for fate determination of r5 and r6 (Maves et al., 2002; Shimizu et al., 2006; Walshe et al., 2002; Wiellette and Sive, 2004). In the aldehyde dehydrogenase 1 family, the A2 gene (aldh1a2) encodes an enzyme which is required for the production of RA and is expressed in forming somites during early neurogenesis (Begemann et al., 2001; Grandel et al., 2002). RA signaling is required for the formation of r5-7 and the anterior spinal cord (Begemann et al., 2001; Grandel et al., 2002; Niederreither et al., 2000; Shimizu et al., 2006). These data suggest that Fgf and RA signal gradients in which Fgf and RA signals are high at r4 and the caudal-most hindbrain, respectively, play important roles in specifying neurons in the caudal hindbrain. Although Fgf and RA signals regulate the expression of mafba, knox20, and hox cluster genes (Ghosh et al., 2018; Marin and Charnay, 2000; Walshe et al., 2002), it is largely unknown which genes function downstream of these signals to regulate neuronal specification.

In mice, the homeobox gene Gsx2 (formerly Gsh2) is expressed in the ventral telencephalon and is involved in fate determination of lateral ganglionic eminence (LGE) progenitor cells, which give rise to striatal neurons and olfactory bulb interneurons (Corbin et al., 2000; Hsieh-Li et al., 1995; Szucsik et al., 1997; Waclaw et al., 2009; Yun et al., 2003). Gsx2, which is also expressed in the ventricular zone of the spinal cord and hindbrain, is involved in the generation of a subset of hindbrain neurons and dorsal interneurons in the spinal cord of mice (Kriks et al., 2005; Mizuguchi et al., 2006; Satou et al., 2013). However, the role of gsx2 in the development of IO neurons has not been elucidated. In this report, we show that gsx2 is co-expressed with ptf1a in neuronal progenitors of IO neurons and is required for the development of IO neurons in zebrafish. gsx2 expression is regulated through transcriptional networks that are activated by Fgf and RA signals in the caudal hindbrain. Our results reveal that gsx2 mediates the rostro-caudal positional signals to control the identity of IO neurons.

RESULTS

gsx2 is co-expressed in IO progenitors that express ptf1a

To reveal the mechanisms that generate IO neurons, we sought genes expressed in IO progenitors in the dorso-caudal hindbrain. A previous study with a BAC transgenic fish TgBAC(gsx2:LOXP-Tomato-LOXP-GFP) (hereafter referred to as Tg(gsx2:RFP)) suggested that gsx2 is expressed in the caudal hindbrain (Satou et al., 2013). Thus, we focused on gsx2 and ptf1a, which is also involved in the generation of IO neurons in mice (Yamada et al., 2007). We found that gsx2 was expressed in the dorsal part of the caudal hindbrain and the ventral telencephalon in an early stage (2, 3 days post fertilization: dpf) of zebrafish larvae (Fig. 1A, B, E, F). As reported previously, ptf1a was expressed in the dorsal ventricular zone of all rhombomeres (Fig. 1C, D, G, H). To examine gsx2-expressing cells in detail, we compared RFP expression in Tg(gsx2:RFP) larvae (gsx2:RFP) with GFP expression in gSAIzGFFM35A;Tg(5xUAS:EGFP) (Fig. 2I-L) and Tg(ptf1a:EGFP) larvae (ptf1a:GFP, Fig. 1M-T). gSAIzGFFM35A is a Gal4 trap line that harbors a Gal4 gene in the exon of mafba (previously known as valentino or kreisler) and can drive UAS-mediated GFP expression at r5 and 6 (referred to as mafbaGFF;UAS:GFP, Fig. 1I-L) (Asakawa and Kawakami, 2018). The gsx2:RFP+ and mafbaGFF;UAS:GFP+ cells did not overlap and were separated adjacently in the caudal hindbrain at an early larval stage (3 dpf, Fig. 1I-L), indicating that gsx2 is specifically expressed in r7 of the hindbrain and the rostral spinal cord. As reported previously (Kani et al., 2010), ptf1a:GFP expression was detected in neural progenitors located in the dorsal ventricular zone of all rhombomeres at an early larval stage (5 dpf, Fig. 1M, N, Q). gsx2:RFP expression was detected in the dorsal ventricular zone of only the caudal hindbrain (Fig. 1M, O, Q). gsx2:RFP and ptf1a:GFP were co-expressed in cells in the dorsal ventricular zone (Fig. 1M, Q). Since GFP and RFP (Tomato) are relatively stable, they can be used for lineage tracing in zebrafish (Kani et al., 2010). Although gsx2 and ptf1a were not detected in the ventral hindbrain (Fig. 1E-H), gsx2:RFP and ptf1a:GFP were detected in IO neurons in the ventral hindbrain in 5-dpf larvae (Fig. 1R, S, T). These results indicate that gsx2 is co-expressed with ptf1a in IO progenitors.

Fig. 1.
Fig. 1. gsx2 and ptf1a are co-expressed in IO progenitors.

(A-B, E-F) Expression of gsx2 in 2- and 3-dpf larvae. (C-D, G-H) Expression of ptf1a in 2- and 3-dpf larvae. (I-L) 3-dpf mafbaGFF;UAS:GFP;gsx2:RFP larvae (n=2) were immunostained with anti-DsRed (RFP, magenta) and anti-GFP (green) antibodies. r5-6: rhombomeres 5 and 6. (M-T) 5-dpf gsx2:RFP;ptf1a:GFP larvae were immunostained with anti-DsRed (magenta) and anti-GFP (green) antibodies. (Q) Highly magnified view of the dotted box in M (dorsal optical section). (R-T) Expression of GFP and/or RFP in the IO region. Arrows and arrowhead indicate CFs and IO neurons, respectively. Yellow arrowheads indicate both GFP+ and RFP+ cells. Scale bars: 100 µm in A (applies to A-H) and I (applies to I to L); 50 µm in M (applies to M-O) and P; 20 µm in Q and R (applies to R-T).

Fig. 2.
Fig. 2. Strong reduction or loss of IO neurons in gsx2 mutants.

(A-H) 5-dpf wild-type (WT, A-D, n=10) and gsx2Δ5/Δ5 (E-H, n=10) hspGFFDMC28C;UAS:GFP (28C;UAS:GFP) larvae which were immunostained with anti-Pvalb7 (magenta) and anti-GFP (green) antibodies. Arrows and arrowhead indicate CFs and IO neurons, respectively. (C, G) Highly magnified views of the boxes in B and F. (I-L) 5-dpf WT (I-J n=6) and gsx2 Δ5/Δ5 (K-L n=7) mnx2b:GFF;UAS:GFP larvae. (J, L) Highly magnified views of the boxes in I and K. (M) Number of 28C;UAS:GFP+ IO neurons in WT and gsx2Δ5/Δ5 larvae. 28C;UAS:GFP+ IO neurons were significantly reduced in gsx2 mutant larvae compared to control larvae. ***p < 0.001 (Student’s t-test). (N-S) Expression of endogenous IO neuronal markers in WT and gsx2Δ5/Δ5. (N-O) Expression of pou4f1 in WT (n=4) and gsx2Δ5/Δ5 (n=4). (P-Q) Expression of grm5a in WT (n=3) and gsx2Δ5/Δ5 (n=2). (R-S) Expression of foxp2 in WT (n=5) and gsx2Δ5/Δ5 (n=5). (T) Illustration of PCs and IO neurons in WT and gsx2 mutant larvae. Scale bars: 100 µm in N (applies to N-S); 50 µm in A (applies to A-B, E-F), D (applies to H), and I (applies to K); 20 µm in C (applies to G) and J (applies to L).

The gsx2 mutation leads to a strong reduction or loss of IO neurons

To reveal the roles of Gsx2 in IO neuronal development, we generated gsx2 mutants by the CRISPR/Cas9 method. The gsx2 mutants harbor a 5-bp (gsx2Δ5) or 8-bp (gsx2Δ8) deletion in exon 1 of the gsx2 gene that introduces a premature stop (Fig. S1). Although gsx2 mRNA was not reduced in the gsx2 mutant larvae (Fig. S2), the Gsx2 proteins of the putative mutants lack the homeodomain (Fig. S1) and the mutations are likely null alleles. To analyze the development of IO neurons in gsx2 mutants, we used the Tg line hspGFFDMC28C;Tg(5xUAS:EGFP) (28C;UAS:GFP) which expresses GFP in IO neurons (Takeuchi et al., 2015). GFP+ somata were located in the ventral hindbrain and they extended their axons (CFs) to PCs, which were marked with anti-parvalbumin7 (Pvalb7) antibody in control 5-dpf Tg larvae (Fig. 2A-D, Fig. S3). In contrast, GFP+ somata were rarely detected and GFP+ CFs were not observed in gsx2 mutants (Fig. 2E-H, M, Fig. S3). In addition to 28C;UAS:GFP, we found that the TgBAC(mnx2b:GFF);Tg(5xUAS:EGFP) line also expressed GFP in a portion of IO neurons in addition to motoneurons (mnx2b:GFF;UAS:GFP) (Asakawa et al., 2013) (Fig. 2I, J). Similar to 28C;UAS:GFP+ cells, a strong reduction or loss of mnx2b:GFF;UAS:GFP+ IO neurons was observed in 5-dpf gsx2Δ5 mutant larvae (Fig. 2K, L). We further examined endogenous markers of IO neurons. pou4f1 (also known as brn3a) and a glutamate receptor, metabotropic 5a (grm5a), are expressed in zebrafish IO neurons (Bae et al., 2009; Haug et al., 2013). foxp2 is expressed in IO neurons of mice (Ferland et al., 2003; Fujita and Sugihara, 2012). Whereas the expression of these markers was detected in IO neurons of wild-type (WT) larvae, it was not observed in gsx2 mutant larvae (Fig. 2N-S, Fig. S3), indicating a strong reduction or absence of IO neurons in gsx2 mutants. These data reveal that Gsx2 is required for IO neuronal development (Fig. 2T).

The ptf1a mutation leads to a strong reduction or loss of IO neurons

We next analyzed the roles of ptf1a in the development of IO neurons by generating ptf1a mutants by the CRISPR/Cas9 method. The ptf1a mutants have a 4-bp (ptf1aΔ4) deletion or 11-bp insertion (ptf1a+11) in exon 1 (Fig. S1). Although the ptf1a mutant larvae retained ptf1a mRNA (Fig. S2), the putative mutant Ptf1a proteins lack the bHLH domain and the mutations were likely null alleles. The ptf1a mutant larvae displayed a complete loss of IO neurons that express GFP in 28C;UAS:GFP or mnx2b:GFF;UAS:GFP Tg lines (Fig. 3E-H, K, L, Fig. S3). Furthermore, the IO markers pou4f1, grm5a, and foxp2 were not detected in ptf1a mutant larvae (Fig. 3M-R, Fig. S3). These results are consistent with the defective development of IO neurons in Ptf1a mutant mice (Yamada et al., 2007). In addition to IO neurons, the ptf1a mutants showed a reduction – but not loss – of PCs and crest cells, which were positive for Pvalb7, as was shown previously (Bae et al., 2009) (Fig. 3E, S4). These data suggest that even though PCs, crest cells, and IO neurons are derived from ptf1a-expressing progenitors, ptf1a functions redundantly with other genes during the development of PCs and crest cells while ptf1a is essential for the development of IO neurons in zebrafish (Fig. 3S).

Fig. 3.
Fig. 3. Loss of IO neurons in ptf1a mutants.

(A-H) 5-dpf WT (A-D, n=5) and ptf1aΔ4/Δ4 (E-H, n=5) 28C;UAS:GFP larvae were immunostained with anti-Pvalb7 (magenta) and anti-GFP (green) antibodies. Arrows and arrowhead indicate CFs and IO neurons, respectively. (C, G) Highly magnified views of the boxes in B and F. (I-L) 5-dpf WT (I-J n=2) and ptf1aΔ4/Δ4 (K-L n=4) mnx2b:GFF;UAS:GFP larvae. (J, L) Highly magnified views of the boxes in I and K. (M-N) Expression of brn3a in 5-dpf WT (n=4) and ptf1aΔ4/Δ4 (n=4) larvae. (O-P) Expression of grm5a in 5-dpf WT (n=4) and ptf1aΔ4/Δ4 (n=3) larvae. (Q-R) Expression of foxp2 in 5-dpf WT (n=5) and ptf1aΔ4/Δ4 (n=5) larvae. (S) Illustration of Purkinje cells and IO neurons in WT and ptf1a mutant larvae. Scale bars: 100 µm in M (applies to M-R); 50 µm in A (applies to A-B, E-F), D (applies to H) and I (applies to K); 20 µm in C (applies to G) and J (applies to L).

Independent regulation and distinct role of gsx2 and ptf1a in IO progenitors

We examined the relationship between gsx2 and ptf1a in IO neuronal development. The expression of ptf1a and gsx2 was not affected in gsx2 and ptf1a mutant larvae, respectively, during the development of IO neurons (Fig. 4). These results indicate that ptf1a and gsx2 are independently regulated in the caudal hindbrain. We further analyzed apoptosis by immunohistochemistry with an anti-cleaved caspase3 antibody since Ptf1a mouse mutants show an increase in apoptotic cells in the caudal hindbrain (Yamada et al., 2007). As in Ptf1a mutant mice, the ptf1a zebrafish mutant larvae showed an increase in the number of cleaved-caspase3+ cells (Fig. 5A-H, Q). In contrast, there was no significant increase in cleaved-caspase3+ cells in gsx2 mutant larvae (Fig. 5I-P, R). Furthermore, gsx2:RFP expression was retained in cells migrating from the dorsal progenitor domain in gsx2 mutant larvae (Fig. S5), suggesting that gsx2+ progenitors remained undifferentiated or differentiated into other neurons in gsx2 mutants. Our findings indicate that ptf1a and gsx2 play different roles in IO neuronal development.

Fig. 4.
Fig. 4. Expression of ptf1a and gsx2 was not affected in gsx2 and ptf1a mutants.

(A-D) Expression of ptf1a in 2-dpf WT (A, C, n=2) and gsx2 mutant (B, D, n=4) larvae. (E-H) Expression of gsx2 in 2-dpf WT (E, G, n=2) and ptf1a mutant (F, H, n=3) larvae. Scale bars: 100 µm in A (applies to A-B, E-F) and C (applies to C-D, G-H).

Fig. 5.
Fig. 5. Apoptosis occurs in ptf1a but not gsx2 mutants.

(A-P) 5-dpf ptf1a+/+ (A-D, n=4), ptf1aΔ4/Δ4 (E-H, n=4), gsx2+/+ (I-L, n=5), and gsx2Δ5/Δ5 (M-P, n=5) ptf1a:GFP larvae were stained with anti-GFP (green) and anti-cleaved caspase3 (magenta) antibodies. (C, G, K, O) Highly magnified views of the boxes in A, E, I and M. (D, H, L, P) Highly magnified views of the boxes in B, F, J and N. Arrows indicate both GFP+ and cleaved caspase3+ cells. (Q-R) Number of GFP+ and cleaved caspase3+ cells in the caudal hindbrain in WT, ptf1a, and gsx2 mutant. ns: not significant; ***p < 0.001 (Student’s t-test for Q and R). Scale bars: 50 µm in A (applies to A-B, E-F, I-J, M-N); 20 µm in C (applies to C-D, G-H, K-L, O-P).

gsx2 expression is negatively regulated by Fgf signaling and Mafba

Gradients of Fgf and RA signals are involved in the formation and patterning of the caudal hindbrain (Dupe and Lumsden, 2001; Gavalas and Krumlauf, 2000; Marin and Charnay, 2000; Maves et al., 2002). We inhibited Fgf and RA signals in order to investigate the regulation of gsx2 expression. Antisense morpholino (MO)-mediated inhibition of fgf3 did not significantly increase gsx2:RFP expression or the number of 28C;UAS:GFP+ IO cells compared to WT (Fig. 6C, D, M, R-T, AA), whereas fgf8a mutant larvae showed a slight increase in gsx2:RFP expression and the number of 28C;UAS:GFP+ IO neurons (Fig. 6E, F, M, U-W, AA). Inhibition of fgf8a and fgf3 (injection of fgf3 MO into fgf8a mutants: fgf8ati282a/ti282a;fgf3MO) further increased gsx2:RFP expression and the number of 28C;UAS:GFP+ IO neurons (Fig. 6G, H, M, X-Z). Consistent with this finding, inhibition of the Fgf signal by SU5402, which is a chemical inhibitor of Fgf receptor tyrosine kinases (Mohammadi et al., 1997), led to an expansion of gsx2:RFP expression (Fig. 6K, L, N). Our findings suggest that the Fgf signal suppressed gsx2 expression and thereby suppressed IO neuronal fate.

Fig. 6.
Fig. 6. Fgf signal suppresses gsx2 expression and IO neuronal development.

(A-H) 3-dpf control (fgf8a+/+) or fgf8ati282a/ti282a mafbaGFF;UAS:GFP;gsx2:RFP larvae that received an injection of control MO or fgf3 MO (fgf3MO) were stained with anti-RFP (magenta) and anti-GFP (green) antibodies. (I-L) mafbaGFF;UAS:GFP;gsx2:RFP were treated with 1% DMSO (I, J, n=4) or 200 µM SU5402 (K, L, n=4) from 6 to 22 hpf, fixed at 3 dpf, and stained with anti-RFP (magenta) and anti-GFP (green) antibodies. (M, N) Length of gsx2:RFP+ hindbrain region (gsx2 expression) in fgf8a+/+ (n=9), fgf8a+/+;fgf3MO (n=9), fgf8a ti282a/ti282a (n=7), and fgf8a ti282a/ti282a; fgf3MO (n=6) larvae (M), and in DMSO- (n=4) or SU5402-treated (n=4) larvae (N). (O-Z) Expression of GFP (green) and Pvalb7 (magenta) in control (fgf8+/+) or fgf8a ti282a/ti282a 28C;UAS:GFP larvae received an injection of control MO or fgf3 MO (fgf3MO). (P, S, V, and Y) Highly magnified views of the boxes in O, R, U, and X. (AA) Length of the 28C;UAS:GFP+ IO region in fgf8a+/+ (n=5), fgf8a+/+;fgf3MO (n=5), fgf8a ti282a/ti282a (n=5), and fgf8a ti282a/ti282a; fgf3MO (n=5) larvae. (AA) Length of GFP+ IO regions. r5-6: rhombomere 5 and 6. ns: not significant; *p < 0.05; **p < 0.01; ***p < 0.001 (Student’s t-test for N and one-way ANOVA with Dunnett’s Multiple Comparison Test for M and AA). Scale bars: 100 µm in A (applies to A-H) and I (applies to I-L); 50 µm in O (applies to O, R, U, and X) and Q (applies to Q, T, W, and Z); 20 µm in P (applies to P, S, V, and Y).

mafbaGFF;UAS:GFP expression was reduced when the Fgf signal was inhibited (Fig. 6G, H, K, L). We further analyzed the roles of mafba in gsx2 expression and in the development of IO neurons (Fig. 7). In mafbaGFF/GFF mutant larvae (Asakawa and Kawakami, 2018), rostral gsx2:RFP expression expanded slightly compared to control heterozygotes, and gsx2:RFP+ cells were observed in the mafbaGFF;UASGFP+ region, corresponding to r5/6 (Fig. 7D-F). Furthermore, 28C;UAS:GFP+ IO neurons increased significantly in mafbab337/b337 mutant larvae (Fig. 7J-L, N, O). These data suggest that Mafba is at least partly involved in the Fgf signal-mediated suppression of gsx2 expression and IO neuronal development.

Fig. 7.
Fig. 7. Mafba negatively controls gsx2 expression and IO neuronal development.

(A-F) 3-dpf mafba+/GFF (A-C, n=2) and mafbaGFF/GFF (D-F, n=5);UAS:GFP;gsx2:RFP larvae were stained with anti-RFP (magenta) and anti-GFP (green) antibodies. Yellow arrows indicate that gsx2 expression is extend to rhombomeres 5 and 6. (G-N) 5-dpf WT and mafbab337/b337 28C;UAS:GFP larvae were stained with anti-GFP (green) and anti-Pvalb7 (magenta) antibodies. Arrows and arrowhead indicate CFs and IO neurons, respectively. (O) Number of GFP+ IO neurons in WT and mafbab337/b337. *p < 0.05 (Student’s t-test). Scale bars: 50 µm in A (applies A to F) and G (applies to G, H, J, and K) and I (applies to I and L); 20 µm in M (applies to M and N).

gsx2 expression is positively regulated by the RA-hox cascade

We inhibited the RA signal by a MO against aldh1a2, which recapitulates the phenotypes of an aldh1a2 mutant (Begemann et al., 2001), or an Aldh1a2 inhibitor diethylaminobezaldehyde (DEAB) (Russo, 1997). Inhibition of the RA signal with both MO and DEAB resulted in reduced gsx2:RFP expression but higher mafbaGFF;UASGFP expression (r5/6, Fig. 8A-J). It also reduced the number of 28C;UAS:GFP+ IO neurons (Fig. 8L, N, P, R, W, X) and expression of the IO neuronal marker pou4f1 (Fig. 8S-V). The reduction was stronger in DEAB-treated larvae than in aldh1a2 morphants, consistent with a previous report that showed that DEAB-treated larvae showed more severe phenotypes than aldh1a2 mutants (Begemann et al., 2004). Our data indicate that the RA signal positively controls gsx2 expression in the caudal hindbrain, and is also involved in IO neuronal development.

Fig. 8.
Fig. 8. RA signal positively controls gsx2 expression and IO neuronal development.

(A-D) 3-dpf control morphant (A-B, n=5) and aldh1a2 morphant (C-D, n=5) mafbaGFF;UAS;GFP;gsx2:RFP larvae were stained with anti-RFP (magenta) and anti-GFP (green) antibodies. (E) Length of gsx2:RFP expression in the hindbrain in control and aldh1a2 morphants. (F-I) mafbaGFF;UAS;GFP;gsx2:RFP larvae were treated with 1% DMSO (F-G, n=5) and 0.25µM DEAB (H-I, n=5) from 6 to 22 hpf, fixed at 3 dpf, and stained with anti-GFP and anti-RFP antibodies. (J) Length of gsx2:RFP expression in the hindbrain of larvae treated with DMSO or DEAB. (K-R) 5-dpf control morphant (K, L, n=5) and aldh1a2 morphant (M, N, n=5), DMSO (O, P, n=5) or DEAB-treated (Q, R, n=5) 28C;UAS:GFP larvae were stained with anti-Pvalb7 (magenta) and anti-GFP (green) antibodies. (L, N, P, R) Highly magnified views of the boxes in K, M, O, and Q. (S-V) Expression of pou4f1 in 5-dpf control morphant (n=5), aldh1a2 morphant (n=5), DMSO- (n=3) or DEAB-treated (n=3). (W) Number of 28C;UAS:GFP+ IO neurons in 5-dpf control morphant and aldh1a2 morphant larvae. (X) Number of 28C;UAS:GFP+ IO neurons in 5-dpf larvae treated with DMSO and DEAB. **p < 0.01; ***p < 0.001 (Student’s t-test). Arrows and arrowhead indicate CFs and inferior olivary nuclei, respectively. Scale bars: 100 µm in A (applies to A-D and F-I) and S (applies to S-V); 50 µm in K (applies to K, M, O, and Q); 20 µm in L (applies to L, N, P, and R).

The RA signal controls the expression of hox genes in the caudal hindbrain (Ghosh et al., 2018; Marin and Charnay, 2000). To address the roles of hox genes in IO neuronal development, we examined gsx2 expression and IO neurons in larvae defective of Pbx2 and Pbx4, which are co-factors in the rostral Hox protein (Waskiewicz et al., 2002). The pbx2/pbx4 morphant larvae recapitulate the phenotypes of maternal zygotic pbx2 mutants in which pbx4 inhibition was mediated by MO, and in which r2-6 acquired the r1 state (Maves et al., 2009). In pbx2/4 morphant larvae, ectopic PCs were observed caudally but crest cells were present normally (Fig. 9G). In contrast, gsx2:RFP expression, the number of 28C;UAS:GFP+ IO neurons, and pou4f1 expression were strongly reduced or absent in pbx2/4-deficent larvae (Fig. 9C, D, G, H, J, K). To address the roles of hox genes, we further ectopically expressed hoxb4a since it is expressed in r7 and in the spinal cord. Injection of hoxb4a mRNA expanded gsx2:RFP expression in the caudal hindbrain (Fig. 9N, O). Although hoxb4a might not be the only hox gene regulating caudal-most hindbrain identify, our data suggest that the RA-hox cascade plays a major role in controlling gsx2 expression and subsequent IO neuronal development.

Fig. 9.
Fig. 9. hox genes are involved in gsx2 expression and IO neuronal development.

(A-D) 3-dpf control morphant (A, B, n=4) and pbx2/4 morphant (C, D, n=4) mafbaGFF;UAS;GFP;gsx2:RFP larvae were stained with anti-RFP (magenta) and anti-GFP (green) antibodies. (E-H) 5-dpf control morphant (E, F, n=4) and pbx2/4 morphant (G, H, n=4) 28C;UAS:GFP larvae stained with anti-Pvalb7 (magenta) and anti-GFP antibodies. (I, J) Expression of pou4f1 in 5-dpf control morphant (I) and pbx2/4 morphant (J) larvae. The pou4f1 signal is marked by arrowheads. (K) Numbers of 28C;UAS:GFP+ IO neurons in control and pbx2/4 morphant larvae. (L-O) 3-dpf control (L, M, n=4) and 25 pg hoxb4a RNA-injected (N, O, n=4) larvae were stained with anti-RFP (magenta) and anti-GFP (green) antibodies. (P) Schematic diagram of the roles of Gsx2 in IO neuronal development. Scale bars: 100 µm in A (applies to A-D), L (applies to L-O) and I (applies to J); 50 µm in E (applies to E and G); 20 μm in F (applies to F and H).

DISCUSSION

Expression of gsx2 and ptf1a in IO progenitors located in rhombomere 7

gsx2 was co-expressed with ptf1a in the dorsal ventricular zone of r7 (Fig. 1). Tracing with the Tg lines revealed that gsx2- and ptf1a-expressing cells (gsx2:RFP+ and ptf1a:GFP+ cells) gave rise to IO neurons (Fig. 1). The gsx2 and ptf1a mutations abrogated IO neuronal development (Fig. 2, 3). These data indicate that gsx2/ptf1a-expressing cells are the neural progenitors of IO neurons. Previous lineage tracing with the Cre-loxP system in mice revealed that IO neurons were derived from Ptf1a-expressing neural progenitors (Yamada et al., 2007). Avian grafting studies suggested that IO neural progenitors were located in r6-8 (Ambrosiani et al., 1996; Cambronero and Puelles, 2000; Kawauchi et al., 2006). Our data clearly indicate that in zebrafish, IO neurons are derived from gsx2/ptf1a-expressing neural progenitors in r7.

Role of Ptf1a in development of hindbrain neurons

ptf1a (Ptf1a in mice) is expressed in neural progenitors in the ventricular zone of the entire dorsal hindbrain in both mice and zebrafish (Fujiyama et al., 2009; Hoshino et al., 2005; Kani et al., 2010; Yamada et al., 2007). Zebrafish ptf1a mutants showed a reduction in the number of PCs and crest cells, and the loss of IO neurons (Fig. 3, S4). In mice, Ptf1a mutants showed a complete loss of PCs (and other cerebellar GABAergic interneurons), DCN neurons, and IO neurons (Fujiyama et al., 2009; Hoshino et al., 2005; Yamada et al., 2007). Although the mice and zebrafish phenotypes are slightly different, our and their data suggest that ptf1a-expressing progenitors give rise to PCs and IO neurons in both species, and also contribute to development of DCN neurons in mice and to crest cells in zebrafish (Fig. 9P).

A milder reduction of PCs in zebrafish suggest that other genes compensate for the lack of Ptf1a. In mice, bHLH-type proneural genes Neurogenin(Neurog)1/2 and Ascl1 are expressed in the ventricular zone of the dorsal hindbrain (Kim et al., 2008; Landsberg et al., 2005; Lundell et al., 2009; Zordan et al., 2008). In mice, Neurog2 mutants show a reduction in number and defective dendritogenesis of PCs (Florio et al., 2012) while the Ascl1 mutation led to a reduction of cerebellar GABAergic interneurons and oligodendrocytes (Grimaldi et al., 2009; Sudarov et al., 2011), suggesting the involvement of other proneural genes in the development of PCs and other interneurons in the cerebellum. Zebrafish has neurog1, ascl1a, and ascl1b but not neurog2 (ZFIN, https://zfin.org). Some of these proneural genes may redundantly function with ptf1a in the development of PCs. Unlike PCs and crest cells, IO neurons were completely absent in ptf1a mutants (Fig. 3), indicating that Ptf1a is indispensable for IO neuronal development in both mice and zebrafish.

In addition to PCs, crest cells in MONs were reduced in ptf1a mutant larvae (Fig. S4). In mice, inhibitory neurons in DCNs are derived from Ptf1a-expressing neural progenitors in the r2-5 ventricular zone (Fujiyama et al., 2009). Although zebrafish do not have DCNs, there is a similarity between DCN and MON. Both DCN and MON receive input from hair cells (in the cochlear organ in mice or via the lateral line system in teleosts) (Yamamoto and Ito, 2005). The DCN and MON have a neural-circuit structure similar to the cerebellum and are thus called cerebellum-like structures (Bell et al., 2008; Oertel and Young, 2004). Although crest cells are excitatory, our data suggest that both crest cells and DCN inhibitory neurons are derived from ptf1a-expressing progenitors. It is tempting to speculate that the DCN and MON were derived from a common ancestral origin during evolution.

Role of Gsx2 in the development of IO neurons

The gsx2 mutants showed a strong reduction or loss of IO neurons (Fig. 2), indicating that Gsx2 is required for IO neuronal development. Previous studies in mice showed that Gsx2 is expressed in the ventricular zone of the ventral telencephalon, hindbrain and spinal cord, and that it is required for the development of striatal neurons in the telencephalon, olfactory interneurons in the olfactory bulb, and dorsal interneurons in the spinal cord (Corbin et al., 2000; Kriks et al., 2005; Szucsik et al., 1997). Therefore, although Gsx2 is involved in the development of different types of neurons, it plays a similar role in the specification of neurons from their progenitors in the ventricular zone of these brain regions.

In some gsx2 mutants, a small number of IO neurons remained, and projected CFs to PCs (Fig. 2), suggesting the presence of genes that redundantly function with gsx2 in IO progenitors. gsx1, which is a homologue of gsx2, is expressed in the hindbrain (r1-7) and spinal cord of zebrafish (Begemann et al., 2001). In mice, Gsx1 and Gsx2 redundantly function in LGE patterning and in specification of dorsal interneurons of the spinal cord (Mizuguchi et al., 2006; Yun et al., 2003). Although the expression domains of gsx1 and gsx2 are different in the spinal cord in zebrafish (Satou et al., 2013), it is possible that gsx1 is co-expressed with gsx2 and cooperatively functions in the development of a portion of IO neurons. Future studies on gsx1 expression and gsx1 mutants will clarify this issue.

Different roles of Ptf1a and Gsx2 in IO neuronal development

The expression of ptf1a and gsx2 in the caudal hindbrain was independent during early neurogenesis (Fig. 4). Whereas the ptf1a mutants showed an increase in apoptosis in the caudal hindbrain, the gsx2 mutants did not (Fig. 5), suggesting different roles of ptf1a and gsx2 in IO neuronal development. The phenotype of zebrafish ptf1a mutants is consistent with that of mouse Ptf1a mutants, which also showed increased apoptosis in the IO region (Yamada et al., 2007). These data imply that a major population of IO progenitors that did not differentiate to neurons died by apoptosis in the absence of ptf1a in both mice and zebrafish. However, some cells derived from Ptf1a-expressing progenitors (Ptf1a-lineage cells) become precerebellar neurons that send mossy fibers in Ptf1a mutant mice (Yamada et al., 2007), suggesting a role of Ptf1a in fate determination of IO neurons in mice. However, in zebrafish, the ptf1a/gsx2-lineage cells (ptf1a:GFP+ and gsx2:RFP+ cells) were present in the dorsal hindbrain and did not migrate ventrally where mossy fiber neurons are supposed to be located in ptf1a mutants (Fig. S4, S5). Therefore, even if some cells derived from IO progenitors undergo a change in fate, they are likely a small population in zebrafish.

In contrast to the ptf1a mutants, gsx2-lineage cells migrated ventrally from the ventricular zone and some of them differentiated to neurons that extended axons to the cerebellum region in gsx2 mutants (Fig. S5). These neurons, which are potentially the nucleus commissure of Wallenberg, are located above the IOs and send mossy fibers projecting to GCs, as reported for other teleost species (Xue et al., 2004). However, these fibers were also reduced in gsx2 mutants (Fig. S5), suggesting that development of these neurons partly depends on gsx2. We also detected commissure axons of gsx2-lineage cells in the caudo-ventral hindbrain of gsx2 mutants but not in control larvae (Fig. S5). The loss of gsx2 may lead to a change in fate from IO neurons to commissure neurons. Our findings suggest that ptf1a is involved in neuronal differentiation while gsx2 is involved in the specification of IO neurons. Ptf1a and Gsx2 cooperatively control the development of IO neurons. Ascl1 and Gsx1/2 coordinately control the specification of dorsal glutamatergic sensory neurons in mice (Mizuguchi et al., 2006). Thus, cooperation between proneural gene(s) and gsx-family genes is a common mechanism to coordinately control the differentiation and specification of a subset of neurons from their progenitors.

To further address the cooperative role between ptf1a and gsx2 in IO neuronal development, we expressed gsx2 in ptf1a-expressing progenitors by using ptf1a:Gal4-VP16 and UAS:gsx2 Tg lines (Fig. S6). However, ectopic co-expression of ptf1a and gsx2 did not induce ectopic IO neurons in the rostral hindbrain. These data indicate that the co-expression of ptf1a and gsx2 was not sufficient to induce IO neurons. Other factors may function with Ptf1a and Gsx2 in IO neuronal development. One of the candidate cofactors is a bHLH-type transcription factor Olig3, which cooperates with Ptf1a in IO neuronal development in mice (Storm et al., 2009). Future studies will reveal a set of transcription factors that sufficiently initiate the genetic program of IO neuronal development.

gsx2 mediates Fgf and RA signals to specify IO neuronal fate

Inhibition of the signal of fgf3 and fgf8a, which are known to be expressed in r4 and are involved in the specification of r5/6 (Maves et al., 2002; Walshe et al., 2002; Wiellette and Sive, 2004), led to reduced mafba expression, increased gsx2 expression, and a reduction in the number of IO neurons (Fig. 6). The mafba mutants showed a slight increase in gsx2 expression and the number of IO neurons (Fig. 7). Fgf signal-mediated mafba expression was previously reported for chicken (Marin and Charnay, 2000) and zebrafish (Ghosh et al., 2018). Collectively, these data indicate that the Fgf signal negatively regulates gsx2 expression and thereby suppress IO neuronal development; Mafba is at least partly involved in this regulation (Fig. 9P). Fgf8a is involved in the formation of the cerebellum (Reifers et al., 1998). In fgf8a- or fgf3/fgf8a-defective larvae, the number of IO neurons increased but PCs were absent (Fig. 6). Intriguingly, IO neurons project their axons rostrally to the optic tectum region even in the absence of PCs, which are the targets of CFs (Fig. 6), indicating the presence of a PC-independent axon guidance mechanism for CFs. Future studies will clarify this mechanism. As MafB directly activates hoxb3 expression in mice (Manzanares et al., 1997), the mafba-hoxb3 cascade may play a role in repressing gsx2 expression and IO neuronal identify.

During early neurogenesis, aldh1a2 is expressed in forming somites in mice, chickens, and zebrafish (Begemann et al., 2001; Berggren et al., 1999; Blentic et al., 2003; Grandel et al., 2002; Niederreither et al., 1997). The treatment of mouse embryos with RA increased IO neurons (Yamamoto et al., 2005) but defective RA signals led to a reduced caudal hindbrain in zebrafish (Begemann et al., 2004; Begemann et al., 2001; Grandel et al., 2002). We showed that the inhibition of RA signals resulted in reduced gsx2 expression and IO neuronal development (Fig. 8). That data indicates that the RA signal positively controls gsx2 expression and, subsequently, IO neuronal development. hox genes function downstream of RA signals (reviewed in (Nolte et al., 2019)). Defects in the RA signal in zebrafish result in a strong reduction or absence of hoxb4a in the caudal hindbrain (Begemann et al., 2004; Begemann et al., 2001; Grandel et al., 2002). hoxb4a enhancer-driven YFP expression was detected in IO neurons (Ma et al., 2009; Punnamoottil et al., 2008). Hoxb4 is a direct target of the RA receptor RAR/RXR complex in chickens (Gould et al., 1998). These data suggest that the RA signal controls caudal hindbrain identity through the expression of hox genes, including hoxb4a, in the caudal hindbrain. Since hox paralogues function redundantly, it is difficult to address the roles of individual hox genes. We demonstrated that the loss of Pbx2 and Pbx4, which function as cofactors for rostral Hox proteins harboring a YPWM motif (Johnson et al., 1995; Mann and Chan, 1996; Shanmugam et al., 1997), led to a strong reduction of gsx2 expression and IO neurons (Fig. 9). Furthermore, misexpression of Hoxb4a, which harbors a YPWM motif, elicited the expansion of gsx2 expression (Fig. 9). Our findings suggest that r7-expressed hox genes, such as hoxb4a, function directly downstream of the RA signal to control gsx2 expression and IO neuronal development (Fig. 9). pbx2/pbx4-deficient larvae show defects in the function of most rostral Hox proteins and the loss of r2-6 identity (Waskiewicz et al., 2002). While the number of PCs increased, the number of crest cells was normal in pbx2/pbx4-deficient larvae (Fig. 9), suggesting that the specification of PCs and crest cells does not rely on Pbx-interacting rostral Hox proteins. The RA-hox cascade controls the specification of IO neurons from ptf1a-expressing progenitors by regulating gsx2 expression (Fig. 9P).

ptf1a-expressing neural progenitors give rise to three different neuron PCs, crest cells, and IO neurons following positional signals. Gsx2 mediates Fgf and RA positional signals to specify IO neurons. Future studies will reveal genetic programs for the specification of PCs and crest cells. The identification of targets for Gsx2 and Ptf1a will clarify the mechanisms underlying the development and functionality of IO neurons.

MATERIALS AND METHODS

Zebrafish strain

The animal work in this study was approved by the Nagoya University Animal Experiment Committee and was conducted in accordance with the Regulations on Animal Experiments at Nagoya University. WT zebrafish with the Oregon AB genetic background were used. For immunohistochemistry and whole mount in situ hybridization, larvae were treated with 0.003% 1-phenyl-2-thiourea (PTU) to inhibit the formation of pigmentation. Tg(UAS:EGFP) (nkuasgfp1aTg), Tg(ptf1a:EGFP), TgBAC(mnx2b:GFF), TgBAC(gsx2:LOXP-Tomato-LOXP-GFP), and TgBAC(ptf1a:GAL4-VP16) were previously reported and characterized (Asakawa et al., 2013; Asakawa et al., 2008; Parsons et al., 2009; Pisharath et al., 2007; Satou et al., 2013). hspGFFDMC28C is a Gal4 trap line that drives UAS-dependent expression in IO neurons (Takeuchi et al., 2015). gSAlzGFFM35A is a mafba mutant in which the mafba exon is disrupted by the Gal4 gene trap, and is also known as mafbankgsaizgffm35aGt (Asakawa and Kawakami, 2018). The Tg(UAS-hsp70l;gsx2BLRP-P2A-BirA-P2A-mCherry) line, which harbors a transgene containing gsx2, biotin ligase recognition peptide (BLRP), 2A peptide sequences of porcine teschovirus-1 (P2A), biotin ligase (BirA), and mCherry genes, will be described elsewhere. The mafbab337 and fgf8ati282a mutants, known previously as valentino and acerebellar, respectively, were described elsewhere (Moens et al., 1998; Reifers et al., 1998). The allele names of the gsx2Δ5, gsx2Δ8, ptf1aΔ4, and ptf1a+11 mutants established in this study were designated as gsx2nub32, gsx2nub33, ptf1anub34, and ptf1anub35, respectively in ZFIN (https://zfin.org). The zebrafish were maintained at 28°C under a 14-h light and 10-h dark cycle. Embryos and larvae were maintained in embryonic medium (EM) (Westerfield, 2000).

Establishment of gsx2 and ptf1a mutants by the CRISPR/Cas9 system

The gRNA targets were designed by the web software ZiFit Targeter 4.2 (http://zifit.partners.org/ZiFiT/) (Hwang et al., 2013; Mali et al., 2013). To generate gRNAs, the following oligonucleotides were used: 5′-TAGGCGGAATTCCACTGCTCAA-3′ and 5′-AAACTTGAGCAGTGGAATTCCG-3′ for gsx2Δ5; 5′-TAGGTCTTCCGGGACGGGCAGA-3′ and 5′-AAACTCTGCCCGTCCCGGAAG-3′ for gsx2Δ8; 5′-TAGGAAGAGGCGGAGGCGCATG-3 and 5′-AAACCATGCGCCTCCGCCTCTT-3′ ptf1aΔ4; 5′- TAGGCGTCAAGCTGCCAACGTC-3′ and 5′-AAACGACGTTGGCAGCTTGACG-3′ for ptf1a+11. gRNA and Cas9 mRNA syntheses were performed as previously reported (Nimura et al., 2019). A solution containing 25 ng/µL gRNA and 100 ng/µL Cas9 mRNA or 1000 ng/µL Cas9 protein (Toolgen) was injected into one-cell-stage embryos using a pneumatic microinjector (PV830, WPI). The insertion and/or deletion (indel) mutations on the target region were detected by a heteroduplex mobility assay (Ota et al., 2013). The mutations were confirmed by sequencing after subcloning the target regions amplified from the mutant genome into pTAC-2 (BioDynamics Laboratory Inc.).

Genotyping

To detect indel mutations, the following primers were used: 5′-GTGCGTATCCTCACACATCCACTCT-3′ and 5′-TGCCATCCTCTGGCAGAACG-3′ to detect the gsx2Δ5 mutation; 5′-CATTGGCATGCACTCTCCCG-3′ and 5′-TGAGGATACGCACAGCGGAC-3′ to detect the gsx2Δ8 mutation; 5′-ACCTCAGAGCTGTCCCCTCACAGA-3′ and 5′-GGCAGCTTGACGCAACTGTT-3′ to detect the ptf1aΔ4 mutation; 5′-GGAGGCGCATGAGGTCTGAAGT-3′ and 5′-GCAGTCCCTCGAAAGCATCG-3′ to detect the ptf1a+11 mutation. To identify gSAIzGFFM35A fish, 5′-CGAGGTAGGAGAAGGGCTGT-3′ and 5′-CTGGAGCGTTTGATGGATACAG-3′ primers were used to amplify about 200-bp PCR products from WT but not gSAIzGFFM35A genomic DNA. Since Tg(gsx2:loxp-DsRed-loxp-GFP) are BAC Tg fish and contain a portion of the gsx2 exon1, we could not distinguish the WT allele of gsx2 from the endogenous locus or the transgene in this fish with the primers described above. To detect WT and mutant alleles of gsx2 in gsx2Δ5;Tg(gsx2: LOXP-Tomato-LOXP-GFP) fish, the following primers were used: 5′-CCATCAGCATCTCGCTCAG-3′ and 5′-CAGTGAAGCCTTGTCCCTCG-3′. After PCR amplification, PCR products were digested with XhoI. The WT gsx2 allele gave rise to two bands, 21- and 257-bp fragments, while the mutant allele displayed a single band since it cannot be digested with XhoI. To detect the fgf8ati282a mutation, 5′-CAGGAGGGGGAAACTGATTGTCTAG-3′ and 5′-CCCTTTCTAGGTGGGATTCTTCTC-3′ primers were used. After PCR amplification, PCR products were digested with XbaI. The WT PCR products could not be digested whereas the mutant PCR products gave rise to 130- and 20-bp DNA fragments.

Injection of antisense morpholino oligonucleotides

fgf3, aldh1a2, pbx2, and pbx4 MOs were translation-blocking antisense MOs that were previously reported (Begemann et al., 2001; Maves et al., 2007; Wiellette and Sive, 2004). One nL of 0.3 mM fgf3 MO, 0.1 mM aldh1a2 MO, 0.03 mM pbx2 MO2, 0.06 mM pbx2 MO3, 0.06 mM pbx4 MO1, or 0.06 mM pbx4 MO2 were injected into one-cell-stage embryos. As a control, we used a standard control MO 5′-CCTCTTACCTCAGTTACAATTTATA-3′. These MOs were obtained from Gene Tool (Philomath, OR, USA).

Treatment with chemical inhibitors

DEAB (Wako) and SU5402 (Calbiochem) were dissolved in DMSO at 100 mM and 20 mM, respectively. Embryos were treated with 0.25 µM DEAB or 200 µM SU5402 in 0.003% PTU/EM from 6 to 22 h post fertilization (hpf).

cDNA cloning

Total RNA was isolated from 5- or 12-dpf zebrafish larvae using TRI Reagent (Molecular Research Center, Inc.). cDNA was generated using ReverTra Ace (Toyobo). DNA fragments for gsx2 or foxp2 were amplified from the 5-dpf cDNA library using the primers 5′-GACTCTTTGATTATCAAGGATCCCG-3′ (F) and 5′-CGTCTTCTGAGCGCGGATAAT-3′ (R) for gsx2, and 5′-CCATGGAGGATAATGGGATG-3 (F) and 5′-TGAGGTAAATTTGGGGGTGA-3′ (R) for foxp2, and subcloned to pGEMT-easy (Promega) and pTA2 (Toyobo), respectively (pGEMTe-gsx2 and pTA2-foxp2). Three DNA fragments for grm5a (grm5a-A, grm5a-B, and grm5a-C) were amplified from zebrafish genomic DNA with the primers 5′-CACTTTTCTCCGTCCACCAT-3′ (F) and 5′-GCGATCTGGGGAATATTGAA-3′ (R) for grm5a-A, 5′-ACCTTCAGTGGGGAGATCCT-3′ (F) and 5′-GATGAAGAGTGCCACGATGA-3′ (R) for grm5a-B, and 5′-CGGCCATTATCAAACCATTC-3′ (F) and 5′-TGACGGTAGGATGGTGAACA-3′ (R) for grm5a-C, and subcloned into pTAC2 (pTAC2-grm5a-A/B/C). Three DNA fragments for pou4f1 (pou4f1-A, pou4f1-B, and pou4f1-C) were amplified from zebrafish genomic DNA with the primers 5′- GCGATGAGCTGAGATGAGAG-3′ (F) and 5′-AGCTCGAGTGCAGAGTTGTG-3′ (R) for pou4f1-A, 5′-GGGTAAGAGTCACCCGTTCA-3′ (F) and 5′-AGTCCGTTGTTGACGAGTCC-3′ (R) for pou4f1-B, 5′- CAATTAACGACTCGGACACG-3′ (F) and 5′-TCAGCTATAGCCGCGATTTT-3′ for pou4f1-C, and subcloned to pTAC2 (pTAC2-pou4f1-A/B/C). The full-length cDNA for hoxb4a was amplified from the 12-hpf cDNA library with the following primers: 5′-GGAATTCATGGCCATGAGTTCCTATTT-3′ (F) and 5′-GCTCTAGACTATAGACTTGGCGGAGGTCC-3′ (R). The DNA fragment was digested with EcoRI and XbaI, and subcloned into pCS2+MT (pCS2+MT-hoxb4a).

In situ hybridization

Whole mount in situ hybridization was performed as previously reported (Bae et al., 2009). Detection of ptf1a and pou4f1 riboprobes was described previously (Bae et al., 2009; Kani et al., 2010). A digoxigenin (DIG)-labeled riboprobe was made from pTAC2 -grm5a-A/B/C, pTA2-foxp2, pGEMTe-gsx2, and pTAC2-pou4f1-A/B/C, using SP6, T3, or T7 RNA polymerase after digestion with restriction enzymes. For grm5a and pou4f1, a mixture of three riboprobes was used (grm5a in Fig. 2, 3; pou4f1 in Fig. S3). To detect transcripts in the IO neurons of 5-dpf larvae, the head regions rostral to the spinal cord of larvae were cut and used for in situ hybridization. To stain larval heads, they were hybridized overnight at 65°C for three nights and incubated overnight with 1/2,000 alkaline-phosphatase conjugated with anti-DIG Fab fragment (Roche) at 4°C for three nights. BM-purple signals were acquired using an AxioPlan-2 microscope equipped with an AxioCam CCD camera (Zeiss).

Immunohistochemistry

For immunohistochemistry, anti-parvalbumin7 (1/1000, mouse monoclonal, ascites) (Bae et al., 2009), anti-GFP (1/1000, rat, Nacalai Tesque, Cat# 04404-84), anti-DsRed (1/1,000, rabbit, Clontech Laboratories, Inc., Cat# 632496), and anti-cleaved caspase3 (1/500, rabbit, Cell Signaling Technology, Cat# 9661) were used. Alexa Fluor 488 goat anti-rat IgG (H + L, Thermo Fisher Scientific Cat# A-11006), CF488A goat anti-rat IgG (H + L, Biotium Cat# 20023), CF488A goat anti-mouse IgG (H + L, Biotium Cat# 20018-1), CF568 goat anti-mouse IgG (H + L, Biotium Cat# 20301-1), and CF568 goat anti-rabbit IgG (H + L, Biotium Cat# 20103) were used as the secondary antibodies. Larvae were immunostained as described previously (Bae et al., 2009). Some fixed samples were optically cleared with SeeDB reagent as previously reported (Ke et al., 2013; Ke and Imai, 2014). An LSM700 confocal laser-scanning microscope was used to obtain fluorescence images.

mRNA injection

To make hoxb4a mRNA for expression, pCS2+MT-hoxb4a was linearized with NotI and transcribed with SP6 RNA polymerase in the presence of a G(5′)ppp(5′)G RNA cap structure analog (New England BioLabs Inc.). 25 pg of hoxb4a mRNA was injected into one-cell-stage embryos.

Statistics

Data were analyzed for statistical significance by Student’s t test and one-way ANOVA using Graphpad PRISM (ver. 5.1).

Competing interest statement

The authors declare no competing financial interests.

Supplementary material

Fig. S1. Structure of WT and mutant Gsx2 and Ptf1a. (A) Structure of WT and mutant Gsx2 and nature of gsx2 mutations generated by the CRISPR/Cas9 method. The position of the CRISPR/Cas9 target is also shown. Gsx2 has a homeodomain. The gsx2Δ5 mutant contains a 5-bp deletion causing a flame shift that results in 82 unrelated amino acids and a stop codon. The gsx2Δ8 mutant contains an 8-bp deletion causing a flame shift that results in 116 amino acids and a stop codon. The putative mutant proteins lack the homeodomain. (B) Structure of WT and mutant Ptf1a and nature of ptf1a mutations generated by the CRISPR/Cas9 method. Ptf1a has a bHLH domain. The ptf1aΔ4 mutant contains a 4-bp deletion causing a frame shift that results in the introduction of 157 unrelated amino acids and a stop codon. The ptf1a+11 mutant contains a 1-bp deletion and 12-bp insertion causing a frame shift that results in the introduction of 162 unrelated amino acids and a stop codon. The putative mutant protein lacks the bHLH domain.

Fig. S2. Expression of gsx2 and ptf1a is not affected in gsx2 and ptf1a mutants. (A-D) Expression of gsx2 in 2-dpf WT (A, C, n=2) and gsx2 mutant (B, D, n=3) larvae. (E-H) Expression of ptf1a in 2-dpf WT (E, G, n=2) and ptf1a mutant (F, H, n=3) larvae. Scale bars: 100 µm in A (applies to A-B,E-F) and C (applies to C-D, G-H).

Fig. S3. Defective development of IO neurons in gsx2Δ8 and ptf1a+11 mutants. (A-D) Expression of pou4f1 in 5-dpf gsx2Δ8/Δ8 (B, n=3), and ptf1a+11/+11 (D, n=3), and control WT sibling larvae (A, C, n=3). (E-H) 5-dpf gsx2Δ8/Δ8 (F, n=5), and ptf1a+11/+11 (H, n=5), and control WT sibling larvae (E, G, n=5) were immunostained with anti-Pvalb7 (magenta) and anti-GFP (green) antibodies. Arrows and arrowhead indicate CFs and IO neurons, respectively. Scale bars: 100 µm in A (applies to A-D); 50 µm in E (applies to E-H).

Fig. S4. Reduction of PCs and crest cells in ptf1a mutant larvae. (A-L) 5-dpf WT (A-D, I-J, n=4) and ptf1aΔ4/Δ4 (E-H, K-L, n=6) Tg(ptf1a:GFP) larvae stained with anti-Pvalb7 (magenta) and anti-GFP (green) antibodies. (I-L) Highly magnified views of the boxes in A and E. Arrowhead shows crest cells. Arrow shows migrating GFP+ cells that are derived from ptf1a-expressing progenitors. (M) Number of Pvalb7+ Purkinje cells. (N) Number of Pvalb7+ crest cells. Scale bars: 50 µm in A (applies to A-C, E-G); 20 µm in D (applies to D, H) and I (applies to I-L). ***p < 0.001 (Student’s t-test).

Fig. S5. Lineage tracing of gsx2-expressing cells in gsx2 mutants. (A-L) 5-dpf WT (A-D, n=13), gsx2Δ5/Δ5 (E-H, n=8) and ptf1aΔ4/Δ4 (I-L, n=3) gsx2:RFP larvae were stained with anti-Pvalb7 (green) and anti-GFP (magenta) antibodies. Arrows and arrowhead indicate CFs and IO neurons, respectively. (C, G, K) Highly magnified views of IO regions. Asterisks indicate axons from gsx2:RFP+ cells, which are potentially the nucleus commissure of Wallenberg. Note that the commissure axons of gsx2:RFP+ cells were observed in the caudo-ventral hindbrain of gsx2 mutants but not in the control or ptf1a mutants (G). Scale bars: 50 µm in A (applies to A-B, E-F, I-J) and D (applies to D, H, L); 20 µm in C (applies to C, G, K).

Fig. S6. Ectopic gsx2 expression alone does not induce IO neurons.(A, B, D, E) Expression of gsx2 in 3-dpf Tg(UAS:gsx2BLRP-P2A-BirA-P2A-mCherry) (control, A, D) and TgBAC(ptf1a:Gal4-VP16);Tg(UAS:gsx2BLRP-P2A-BirA-P2A-mCherry) (B, E) larvae. The Tg(UAS:gsx2BLRP-P2A-BirA-P2A-mCherry) line harbors a transgene containing the open reading frame (ORF) of gsx2, biotin ligase recognition peptide (BLRP), 2A peptide sequences of porcine teschovirus-1 (P2A), biotin ligase (BirA), and mCherry genes. Expression of Gsx2 and mCherry can be driven by Gal4-VP16. (C) Expression of mCherry was induced where ptf1a was expressed in 3-dpf Tg larvae. (F-G) Expression of pou4f1 in 3-dpf control (F, n=3) and Tg(ptf1a:Gal4-VP16);Tg(UAS:gsx2BLRP-P2A-BirA-P2A-mCherry) (G, n=3) larvae. Although gsx2 was ectopically expressed in all ptf1a-expressing cells, it did not induce ectopic expression of pou4f1. IO: inferior olivary nuclei. Scale bars: 100 µm in A (applies to A-G); 50 µm in C.

Acknowledgements

We thank Shin-ichi Higashijima and the National Bioresource Project for providing the transgenic zebrafish, Makoto Kobayashi for the mafbab337 mutant, Masato Kinoshita and Feng Zhang for the hSpCas9 plasmids, Kuniyo Kondoh and Yumiko Takayanagi for managing fish mating and care. We also thank the members of the Hibi Laboratory, Hanako Hagio and Naoyuki Yamamoto, for helpful discussion. This work was supported by JSPS KAKENHI Grant Number JP15H04376, JP18H02448 (to M.H.), JP18K06333 (to T.S.), and JP16K18548 (to. M.T.), and CREST Japan Science and Technology Agency (JST) JPMJCR1753 (to M.H.).

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Gsx2 is involved in specification of neurons in the inferior olivary nuclei from Ptf1a-expressing neuronal progenitors in zebrafish
Tsubasa Itoh, Miki Takeuchi, Marina Sakagami, Kazuhide Asakawa, Koichi Kawakami, Takashi Shimizu, Masahiko Hibi
bioRxiv 2020.03.16.993402; doi: https://doi.org/10.1101/2020.03.16.993402
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