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. Author manuscript; available in PMC: 2016 Jan 1.
Published in final edited form as: Gene Expr Patterns. 2014 Dec 23;17(1):38–44. doi: 10.1016/j.gep.2014.12.002

Characterization of tweety gene (ttyh1-3) expression in Xenopus laevis during embryonic development

Andrew D Halleran 1,#, Morgan Sehdev 1,#, Brian A Rabe 1,2, Ryan W Huyck 1, Cheyenne C Williams 1, Margaret S Saha 1,*
PMCID: PMC4402287  NIHMSID: NIHMS665539  PMID: 25541457

Abstract

The tweety family of genes encodes large-conductance chloride channels and has been implicated in a wide array of cellular processes including cell division, cell adhesion, regulation of calcium activity, and tumorigenesis, particularly in neuronal cells. However, their expression patterns during early development remain largely unknown. Here, we describe the spatial and temporal patterning of ttyh1, ttyh2, and ttyh3 in Xenopus laevis during early embryonic development. Ttyh1 and ttyh3 are initially expressed at the late neurula stage are and primarily localized to the developing nervous system; however ttyh1 and ttyh3 both show transient expression in the somites. By swimming tadpole stages, all three genes are expressed in the brain, spinal cord, eye, and cranial ganglia. While ttyh1 is restricted to proliferative, ventricular zones, ttyh3 is primarily localized to postmitotic regions of the developing nervous system. Ttyh2, however, is strongly expressed in cranial ganglia V, VII, IX and X. The differing temporal and spatial expression patterns of ttyh1, ttyh2, and ttyh3 suggest that they may play distinct roles throughout embryonic development.

Keywords: Xenopus, tweety, ttyh1, ttyh2, ttyh3, Nervous system, Embryo

First identified as a transcriptional unit adjacent to the Drosophila flightless I gene, the tweety genes comprise a highly conserved, evolutionarily ancient family that encode large-conductance chlo-ride channels (Campbell et al., 1993, 2000). Displaying a broad phylogenetic distribution indicative of important cellular functions, tweety orthologs have been identified in plants, slime molds, protozoa, and a wide range of animals (Matthews et al., 2007). In vertebrates, the tweety gene family has three members, tweety homolog 1 (ttyh1), tweety homolog 2 (ttyh2), and tweety homolog 3 (ttyh3), all of which are reported to encode large-conductance trans-membrane chloride channels. Predicted to have either five (Ttyh1, Ttyh2) or six (Ttyh3) transmembrane domains, Ttyh1 is activated by cellular swelling while Ttyh2 and Ttyh3 appear to be activated in response to elevated Ca2+ concentrations (He et al., 2008; Suzuki, 2006; Suzuki and Mizuno, 2004).

The association of tweety genes with fundamental cellular processes as well as several pathological conditions has created a heightened interest in the members of this gene family. Primarily localized to neural tissues, ttyh1 encodes a calcium-binding endoplasmic reticulum protein that may be involved in regulating Ca2+ sequestration and concentration during mitosis. Inactivation of both copies of this gene in mice leads to very early embryonic lethality (Kumada et al., 2010). Ectopic expression of ttyh1 in human epithelial kidney cells results in long, branched filopodia while overexpression in cultured rat hippocampal neurons leads to intense neuritogenesis and elaborate dendritic trees (Matthews et al., 2007; Stefaniuk et al., 2010). The latter observation is consistent with the upregulation of ttyh1 in epilepsy and epileptogenesis (Lukasiuk et al., 2003; Stefaniuk and Lukasiuk, 2010; Stefaniuk et al., 2010). Ttyh1 expression is also strongly upregulated in an array of childhood brain tumors; fusion of the ttyh1 promoter to the large microRNA cluster C19MC drives the development of a particularly aggressive form of pediatric cancer (Kleinman et al., 2014; Matthews et al., 2007). While less is known about ttyh2, it is strongly upregulated in both colon and renal cell carcinoma (Rae et al., 2001; Toiyama, 2007), suggesting a possible role in cell division and tumor growth. Although the ttyh3 gene product has been associated with calcium dynamics in excitable tissues such as skeletal muscle, brain, and spinal cord, research has focused almost exclusively on its biochemistry rather than expression or function (He et al., 2008; Suzuki, 2006).

Despite the importance of the tweety genes in development and disease, relatively little is known about their expression outside of select adult human tissue samples, tumor biopsies, a few cell lines, and adult rodent tissues (Al-Jumaily et al., 2007; Matthews et al., 2007; Rae et al., 2001; Suzuki, 2006; Suzuki and Mizuno, 2004; Toiyama et al., 2007). The only information on the expression of any of the tweety genes during embryonic development is Kumada et al. (2010) who analyzed expression of ttyh1 in the E7.5 and E14.5 mouse brain and showed its essential role during embryogenesis. There has been no comprehensive analysis of the expression pattern of this important gene family during embryonic development. Here, we have chosen to characterize the spatial and temporal expression pattern of the three vertebrate tweety homologs throughout embryogenesis in Xenopus laevis, a particularly suitable model system given the ability to access and manipulate all stages of development (Sive et al., 2000). Using Xenopus laevis, we found that each of the tweety genes has a unique temporal and spatial expression pattern localized primarily to the developing nervous system.

1. Results and discussion

1.1. Cloning of Xenopus tweety orthologs

Using available sequence information from GenBank (GenBank accession numbers BC073236.1; BC073236.1; BC073236, respectively), ttyh1, ttyh2, and ttyh3 were cloned from X. laevis tadpole (stage 35) cDNA using reverse transcriptase polymerase chain reaction (RT-PCR) (all stages are according to Nieuwkoop and Faber, 1994). Sequencing confirmed identity (99–100% at the nucleotide level) for all three clones. Pairwise comparisons between each clone and the GenBank sequence for the other two homologs showed minimal identity, the highest being 24%. This ensured that the probe for one family member would not cross-hybridize with endogenous mRNA from another family member under the conditions employed during whole mount in situ hybridization (ISH).

1.2. Expression of Xenopus ttyh1

Using whole mount in situ hybridization from early blastula stages onward, signal for ttyh1 was first detected at neurula stages throughout the developing central nervous system. By late neurula stages (stage 19), ttyh1 transcripts become particularly abundant in the midbrain in the presumptive eye region (Fig. 1A). During tailbud stages mRNA levels increased throughout the nervous system following an anterior (strongest signal) to posterior (weaker signal) gradient, with distinct expression in the somites also present (Fig. 1B–D). Signal increased dramatically at swimming tadpole stages by which point it extended to the posterior end of the spinal cord (Fig. 1E,F). Histological analysis revealed that ttyh1 was distributed along the entire dorsal–ventral axis of the neural tube with the exception of the roofplate in the forebrain, hindbrain and spinal cord and the spinal cord floorplate. Notably, throughout the entire central nervous system ttyh1 is tightly restricted to the ventricular, mitotic zone of the neural tube with little expression in the more lateral zones (Fig. 2A–E). Otic vesicle expression was apparent beginning at early tailbud stages, and continued through the swimming tadpoles stage (Figs 1C–F, 2C). There is strong ttyh1 expression in the inner and outer layers of the retina by swimming tadpole stages (Figs 1E,F, 2B). Ttyh1 transcripts appeared in cranial ganglia VII and IX beginning at the swimming tadpole stages (Figs 1F, 2C).

Fig. 1.

Fig. 1

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Whole mount in situ hybridization showing spatial and temporal expression of tweety homologs in developing X. laevis embryos. Row 1 and 2: dorsal views. Rows 3, 4, 5: both lateral (top embryo, with dorsal side on top) and dorsal (bottom embryo) views. Anterior is oriented to the right for all embryos. Row 6: close up of anterior part of the embryo, with a lateral view (dorsal side on top) and a dorsal view (with anterior directed upwards). (A, G, M) Stages 19–20 (late neurula). (B, H, N) Stages 22–23 (early tailbud). (C, I, O) Stages 25–26 (mid-tailbud). (D, J, P) Stages 29–31 (late tailbud). (E, K, Q) Stages 34–35 (swimming tadpole). (F, L, R) Stages 34–35 (swimming tadpole, close up). (A–F) Ttyh1 expression. (G–L) Ttyh2 expression. (M–R) Ttyh3 expression. Arrowheads indicate regions of expression. (fb, forebrain; e, eye; hb, hindbrain; mb, mid-brain; ot, otic vesicle; pa, pharyngeal arches; sc, spinal cord; nt, neural tube; so, somites; Roman numerals: cranial ganglia.) Scale bars = 1.0 mm.

Fig. 2.

Fig. 2

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Histological analysis of tweety homolog expression in X. laevis stages 35–37 (swimming tadpole) using whole-mount in situ hybridization. Transverse 18 μm sections along the anterior–posterior axis with dorsal to the top. (A–O) 10× magnification images focused on neural regions with signal. (A′–O′) 4× magnification images to show to the absence of signal in other regions. (A, F, K) Sections showing forebrain. (B, G, L) Sections showing midbrain. (C, H, M) Sections showing hindbrain. (D, I, N) Sections showing anterior spinal cord. (E, J, O) Sections showing posterior spinal cord. (A–E) Ttyh1 expression. (F–J) Ttyh2 expression. (K–O) Ttyh3 expression. Arrowheads indicate regions of expression. (asc, anterior spinal cord; mb, midbrain; hb, hindbrain; fb, forebrain; e, eye; nc, neural crest; ot, otic vesicle; psc, posterior spinal cord; sc, spinal cord; Roman numerals: cranial ganglia.) Scale bars = 0.25 mm.

Since gene expression would not be detectable using whole mount in situ hybridization if transcripts were uniformly distributed at low levels, qRT-PCR was employed to determine if any of the tweety genes had earlier, particularly maternal, expression. While very low levels of ttyh1 mRNA were detectable prior to the onset of transcription at mid-blastula stages, these levels were 100-fold less than at neurula stages and over a thousand fold less than at swimming tadpoles stages making it questionable whether the low level of early expression is physiologically relevant (Fig. 3).

Fig. 3.

Fig. 3

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qRT-PCR showing ttyh gene expression from single cell to stage 45. Stage 0 in the figure denotes unfertilized eggs. At each stage gene expression was quantitated using the ΔΔCT method and corrected for expression of the control gene (Drosha). Relative expression is displayed log10 transformed.

1.3. Expression of Xenopus ttyh2

As with ttyh1, qRT-PCR revealed trace amounts of ttyh2 transcripts throughout pre-gastrula stages followed by a hundred fold increase at neurula stages and another dramatic rise in mRNA abundance at swimming tadpole stages (Fig. 3). This result was consistent with whole mount in situ hybridization. Using in situ hybridization, ttyh2 transcripts are first detectable at hatching stages in the cranial ganglia; by swimming tadpoles stages ttyh2 is prominently expressed in cranial ganglia V, VII, IX and X as well as the outer layer of the eye (Figs 1K,L, 2G–I). To further investigate the expression of ttyh2 in the cranial ganglion, co-localization with a known cranial nerve marker xVGlut1 (Gleason et al., 2003) was assayed. Co-expression was evident in cranial ganglia V, VII, and X (Fig. 4B,C). By swimming tadpoles stages, ttyh2 is also expressed weakly and diffusely throughout the neural tube (Fig. 1K,L), but there is strong expression in the dorsal aspect, particularly in the roofplate of the posterior neural tube (Fig. 2J).

Fig. 4.

Fig. 4

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Due to the concentrated expression of ttyh2 in the cranial ganglia, double fluorescent in situ hybridization was carried out to determine its co-localization with another known cranial ganglia marker. Histological analysis of tweety homolog 2 (ttyh2) (Cy3-tyramide) and xVGlut1 (FITC-tyramide) expression in X. laevis stage 35 (swimming tadpole) using whole-mount fluorescent in situ hybridization. Transverse 25 μm sections along the anterior–posterior axis with dorsal to the top. (A) Section showing fore-brain. (B) Section showing midbrain. (C) Section showing hindbrain. (mb, midbrain; hb, hindbrain; fb, forebrain; e, eye; Roman numerals: cranial ganglia.) Scale bars = 0.25 mm.

1.4. Expression of Xenopus ttyh3

As with ttyh1, ttyh3 was detectable using whole mount in situ hybridization at neurula stages when strong, localized signal appeared in the anterior regions of the nervous system, particularly the forebrain and developing eye (Fig. 1M,N). Like ttyh1, ttyh3 also had transient signal in developing somite regions beginning at neurula stages (stage 20) (Fig. 1M), signal that gradually disappeared by the tailbud stage (stage 25) (Fig. 1O). Beginning at hatching stages (stage 30), ttyh3 was also detected in the developing pharyngeal arches (Fig. 1P). By hatching stages, expression remained constant in the anterior nervous system and become stronger in the spinal cord (Fig. 1P). At swimming tadpole stages, ttyh3 continued to be expressed throughout the central nervous system (Fig. 1Q,R). In contrast to ttyh1, thyh3 was located preferentially in the postmitotic, lateral regions of the neural tube throughout its anterior– posterior axis (Fig. 2K–O). Interestingly, the expression patterns of ttyh1 and ttyh3 appear to be somewhat mutually exclusive; ttyh1 is localized medially in highly mitotic domains along the inner edges of the brain ventricle while ttyh3 is expressed throughout the more lateral regions of the neural tube. Through swimming tadpole stages, ttyh3 transcripts are detected in ganglia V, VII, IX, and X (Figs 1R, 2M). Similar to ttyh1 and ttyh2, ttyh3 qRT-PCR showed an extremely low, but nevertheless detectable, level of expression during neurula stages and peaked at tailbud stages.

1.5. Discussion

The tweety gene family displays expression patterns highly suggestive of an important role during embryonic development. While delineation of the specific function of the three genes awaits further investigation, previous studies have suggested they play a critical role in dynamically regulating ion concentrations within the cell, particularly calcium (Kumada et al., 2010; Suzuki, 2006). The regions in which the genes are expressed – the developing nervous system and somites – are known to be highly dependent on tightly regulated calcium activity for proper development during embryogenesis. Ca2+ activity has been widely shown as a governing factor in nearly every aspect of neural development (Leclerc et al., 2006; Moreau et al., 2008; Rosenberg and Spitzer, 2011), from cell division (Li et al., 2008) to specification of neurotransmitter phenotype (Borodinsky et al., 2004, 2012), and also in somite formation and muscle regeneration (Chernoff and Hilfer, 1982; Tu and Borodinsky, 2014; Webb and Miller, 2006).

Although each of the genes displays some overlapping regions of expression during neural development, each possesses a unique temporal and spatial pattern suggesting distinct functions. Ttyh1 for example does not display significant maternal expression, but rather shows onset of expression at late neurula stages with dramatically increased neural-specific expression through swimming tadpole stages, a result also consistent with RNA-Seq results in X. tropicalis transcriptome database (Tan et al., 2013). Ttyh1 is consistently and tightly localized to the ventricular regions of the developing nervous system, where cells are mitotic. The association with proliferative cells is consistent with the role of ttyh1 in highly aggressive pediatric brain tumors (Kleinman et al., 2014; Matthews et al., 2007). Ttyh1 has also been associated with neuritogenesis and dynamic filipodia-like protrusions in rat cell cultures and brain slices (Matthews et al., 2007; Stefaniuk and Lukasiuk, 2010). In Xenopus, the onset and timing of ttyh1 expression coincides approximately with the timing of neurite formation (Hartenstein, 1989).

Like ttyh1, ttyh2 is implicated in tumorigenesis (Rae et al., 2001; Toiyama et al., 2007), and it is almost exclusively restricted to the cranial ganglia during its initial phase of expression. Ttyh3, on the other hand, is expressed in postmitotic regions in a nearly mutually exclusive manner to ttyh1. Ttyh3 is also expressed far more weakly and broadly than ttyh1, although both genes are transiently expressed in the somites. While qRT-PCR results were overall consistent with those from in situ hybridization there were some apparent discrepancies. This discrepancy may also be attributable to splice variants known to occur in the tweety gene family. Ttyh3, for example, showed prominent expression throughout the nervous system at swimming tadpole stages while qRT-PCR revealed peak expression at tailbud stages. It should be noted, however, that the color reaction took considerably longer to produce distinct signal, a result that lends validity to the qRT-PCR data. Notably Ttyh1 and Ttyh2 share more similarity at the amino acid and predicted protein level than with Ttyh3 which actually has a different transmembrane structure. Predicted evolutionary relationships suggest that ttyh3 homologs are more distant, having diverged prior to a duplication event that led to the ttyh1 and ttyh2 genes (Matthews et al., 2007). Although previous research on tweety gene expression has been limited primarily to cell lines and adult tissues, for the one example of embryonic expression for ttyh1 in mice, the results presented here remain consistent with that study. The growing recognition of the key role that ion fluxes play in normal development, regenerative processes, and pathogenesis, make these genes intriguing candidates for further investigation (Borodinsky et al., 2012; Levin, 2014; Rosenberg and Spitzer, 2011). The broad phylogenetic distribution of this gene family, their proposed role in calcium dynamics and the discrete expression patterns of the tweety genes in the developing nervous system presented here suggest an important role for tweety genes in neural proliferation and differentiation.

2. Experimental procedures

2.1. Animal usage

Embryos were obtained from matings between adult albino X. laevis following injection of human chorionic gonadotropin as detailed by Sive et al. (2000). Embryos were staged according to Nieuwkoop and Faber (1994). All animal care and procedures were performed in accordance with the regulations set forth by the Institutional Animal Care and Use Committee (IACUC) at the College of William and Mary.

2.2. Cloning and sequence analysis

Primers for PCR were designed from cDNA clone sequences of X. laevis mRNA from tweety homologs. Sequences were obtained from NCBI Reference Sequences (NCBI accession numbers: ttyh1, NM_001093946; ttyh2, NM_001087123; ttyh3, NM_001092244) and used to design primers that produced PCR products approximately one kilobase in length.

Primer pairs for cloning were designed to be both unique to each homolog and highly conserved to ensure recognition of X. laevis sequences. Primers used for cloning were as follows: ttyh1, (5′-CCAAGAGCCAAAGGGGCTTA-3′) and (5′-AATCACTCAGTCCCACAGCC-3′); ttyh2, (5′-GACACTGCCAATGGGTTCTT-3′) and (5′-AGAGA TTTCCGTCTGCCTCA-3′); ttyh3, (5′-TGATGCTTCTGGCTTGTGTC-3′) and (5′-AACCACGGCAATCTACCAAG-3′).

Total RNA was extracted from X. laevis stage 35 embryos with RNeasy Mini Kit (Qiagen). cDNA for PCR was synthesized using the Qiagen iScript cDNA Synthesis kit. PCR was performed using the Ambion Supertaq Plus DNA Polymerase Kit with annealing temperatures between 55 and 57 °C. PCR products were then cloned into the Agilent pSC-A-amp/kan PCR Cloning Vector and transformed into One Shot DH5α Competent Escherichia coli cells. PCR products within plasmid DNA were purified using the PureYield Plasmid Midiprep kit from Promega followed by sequencing to confirm each clone's identity. Sequencing was performed with ABI 3130 Genetic Analyzer sequences using the BigDye Terminator v3.1 Ready Reaction Cycle Sequencing Kit (Applied Biosystems). Using BLAST, sequence results were compared for both total nucleotide and amino acid similarities across all homologs to ensure stringency of probes. Sequences were aligned among orthologs including X. tropicalis and H. sapiens using ClustalW2 multi sequence alignment tool.

2.3. qRT-PCR

At each of stages 0, 5, 6.5, 7, 8, 10, 12, 15, 20, 25, 30, 35, 40 and 45 10 embryos were frozen in liquid nitrogen. Frozen embryos were homogenized by grinding with a pestle in 700 μL TRIzol (Life Technologies). RNA was extracted from homogenate with 4-bromoanisole and purified with the RNeasy Mini Kit (Qiagen). cDNA was synthesized from total RNA using iScript Reverse Transcription Supermix for RT-qPCR (Bio-Rad). qRT-PCR was performed using the TaqMan Gene Expression Master Mix (Life Technologies) on a StepOne Real-Time PCR System. Cycling conditions were: 95 °C for 10 min; 40 cycles of 95 °C for 15 s and 60 °C for 1 min. TaqMan probes were labeled at the 5′ end with FAM dye and at the 3′ end with the NFQ quencher. Primers used were: ttyh1, (5′-GTGTCACCTGGAGCTGTGT-3′) and (5′-GCTGTTTCCGTAGAAACCAATGC-3′); ttyh2, (5′-CTGGA CTGCTGCTGTGTCT) and (5′-CATTGGTCTCACTGTTCCCATAGAA-3′); ttyh3, (5′-TGGAAGAATCATTACTTGCTGGATGAA-3′) and (5′-CAGAA GCAGGCCCAGGTA-3′); Drosha, (5′-ACCCCGATCGCCTTCATG-3′) and (5′-GGCTTTCAAACTGCACTTACAGAGA-3′). Three technical replicates were performed for each run. Raw data were processed with StepOne Software v2.3. Data were normalized to Drosha RNA (ΔΔCT analysis).

2.4. Expression analysis

Plasmid DNA was linearized with restriction enzymes, and mRNA probes were transcribed while incorporating digoxigenin-labeled uracil using the respective RNA polymerases [ttyh1 sense – HindIII and T7; ttyh1 antisense – Not1 and T3; ttyh2 sense – HindIII and T7; ttyh2 antisense – BamH1 and T3; ttyh3 sense – HindIII and T7; ttyh3 antisense – Not1 and T3]. Probes were then used for in situ hybridization of X. laevis embryos stages 15–40. Sense probes were used to verify the absence of nonspecific binding. Chromagenic in situ hybridization was performed using BCIP/NBT (nitroblue tetrazolium/5-bromo-4-chloro-3-indolyl phosphate) alkaline phosphatase substrates as described by Harland (1991) with slight modifications. Double fluorescent in situ hybridization was performed to determine co-expression of ttyh2 and xVGlut1 (Gleason et al., 2003) in cranial ganglion. The multiplex fluorescent in situ, described by Vize et al. (2009) with some modifications, used ttyh2 Q5 mRNA probes transcribed while incorporating fluorescein-labeled uracil and xVGlut1 mRNA probes transcribed while incorporating digoxigenin-labeled uracil. The reaction was developed in Cy3-tyramide and FITC-tyramide (Harland and Keller, 1999) for ttyh2 and Q6 xVGlut1, respectively.

Whole mount patterning of spatial and temporal expression was visualized following dehydration and clearing of embryos according to Sive et al. (2000). Whole mount photography was obtained using an Olympus SZX7 dissecting scope attached to an Olympus DP71 camera. Histological data were obtained from embryos that were cryoprotected in a 1.6 M sucrose solution for at least 12 hours at 4 °C and subsequently fixed in Tissue Freezing Medium for a minimum of 2 hours at room temperature. Embryos were then embedded in TFM at −20 °C, cryosectioned into 18 μm slices, and transferred to slides for imaging. Transverse histological analysis was performed in accordance to Li et al. (2006). Histological photography was taken using an Olympus BX60 scope attached to a Media Cybernetics QCapture digital camera. For each stage and each gene, in situ hybridization was repeated at least three times with a minimum of 10 embryos for each stage.

Acknowledgements

We thank Lidia Epp for her assistance with sequencing and Sam Clamons for his technical help. We also thank Eileen Ablondi and Catherine Bianchi for their suggestions to the manuscript. Support was provided by grants from the National Science Foundation IOS-1257895 and NIH R15 NS067566-01 and R15HD077624-01 to MSS and by a grant from the Howard Hughes Medical Institute Undergraduate Science Education Program to the College of William and Mary (52005868).

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