* Montreal Neurological Institute and Department of Biology, McGill University, Montreal, Quebec, Canada
Biology Department, Rhode Island College, Providence, Rhode Island
Department of Zoology, Graduate School of Science, Kyoto University, Kyoto, Japan
Correspondence: E-mail: ken.hastings{at}mcgill.ca.
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Abstract |
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Key Words: contractile regulatory mechanism muscle chordate evolution Ciona intestinalis gene family evolution
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Introduction |
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An integral aspect of muscle function is the contractile regulatory mechanism. One regulatory mechanism, based on the Ca2+-sensitive troponin complex in cooperation with tropomyosin, is associated with sarcomeric thin (actin) filaments throughout the metazoa (Farah and Reinach 1995; Gergely 1998; Squire and Morris 1998; Gordon, Homsher, and Regnier 2000). Troponin/tropomyosin regulation also occurs in several unusual nonsarcomeric muscles (Toyota, Obinata, and Terakado 1979; Endo and Obinata 1981; Myers et al. 1996), including adult ascidian body-wall muscle (Toyota, Obinata, and Terakado 1979; Endo and Obinata 1981). The troponin complex consists of three subunits: (1) troponin C (TnC), a Ca2+-binding subunit, (2) troponin T (TnT), a tropomyosin-binding subunit, and (3) troponin I (TnI), a subunit that interacts with actin, and with TnC and TnT, and that is the primary contractile regulatory subunit (Farah and Reinach 1995; Perry 1999).
Vertebrates have three TnI isoforms, sharing 60% amino acid sequence identity, each encoded by a distinct gene (Dhoot and Perry 1979; Hastings 1997). Several short amino acid sequence motifs have been evolutionarily conserved across the metazoa (Kobayashi et al. 1989; Barbas et al. 1991), including a functionally critical 12-amino-acid segment near the middle of the molecule that binds actin and TnC and inhibits the actin-myosin contractile interaction (Talbot and Hodges 1981). Vertebrate skeletal muscle TnI isoforms (TnIfast and TnIslow) are characteristically shorter than most TnIs,
180 amino acids long (Wilkinson and Grand 1978). In contrast, the vertebrate heartspecific TnIcardiac isoform is
210240 amino acids in length, similar in size to most protostome invertebrate TnIs (200250 amino acids). Although they are relatively short, vertebrate skeletal muscle TnIs are not C-terminally truncated, as are several short TnI isoforms found in the protostome invertebratese.g., in the arthropod Drosophila (Barbas et al. 1991; Beall and Fyrberg 1991). Rather, the unique shorter chain length of vertebrate skeletal muscle TnIs reflects differences near the N-terminus (Wilkinson and Grand 1978 ), a region that interacts with TnC (Farah and Reinach 1995; Perry 1999).
The evolutionary origins of vertebrate-characteristic features of skeletal muscle, such as short but C-terminally complete TnI, are not well understood. Studies of ascidian larval tail muscle may provide evolutionary insight, but to date they have led to enigmatic findings. Yuasa et al. (1997, 2002) found that embryonic/larval muscle of the ascidian Halocynthia roretzi expressed several closely related TnIs that were markedly truncated at the C-terminus; a distinct gene encoding a C-terminally complete TnI was expressed in adult muscles. In their C-terminal truncation, the larval muscle TnIs of Halocynthia do not resemble any vertebrate TnI, but they are reminiscent of some protostome invertebrate TnIs. Did vertebrate skeletal muscle evolve from an ancestral chordate muscle that, like Halocynthia larval tail muscle, expressed a C-terminally truncated TnI? Or is vertebrate skeletal-muscletype TnI (short but with complete C-terminus) an ancient feature of chordate locomotory muscle, with the Halocynthia larval TnI C-terminal truncation perhaps representing a lineage-specific innovation (Yuasa et al. 2002)?
Here we report studies of the TnI expressed in larval tail muscle of Ciona intestinalis, an ascidian of the Order Enterogona/Suborder Phlebobranchiata, distantly related to Halocynthia (Order Pleurogona/Suborder Stolidobranchiata, or Order Stolidobranchia; see Wada (1998) and Swalla et al. (2000) for recent discussion of tunicate classification). We have previously characterized a vertebrate skeletal-musclelike 182-residue C-terminally complete TnI expressed in Ciona adult body-wall muscle (MacLean, Meedel, and Hastings 1997). We now show that, in contrast to the mutually exclusive expression of specialized larval and adult TnI genes in Halocynthia, the same Ciona TnI isoform and gene that is expressed in adult muscle is also expressed in embryonic/larval tail muscle cells. Thus, unlike Halocynthia, Ciona expresses a TnI in larval tail muscle cells that has the typical features of vertebrate skeletal muscle TnI. Comparisons of amino acid sequences and genetic mechanisms of C-terminal truncation clearly indicate that the C-terminally truncated larval TnIs of Halocynthia represent a derived, rather than an ancestral, feature. Our findings indicate that vertebrate skeletal-musclecharacteristic features of TnI were developed in the locomotory muscle of chordates before the tunicate/vertebrate divergence.
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Materials and Methods |
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RNA Isolation and Analysis
RNA was prepared by phenol extraction/salt precipitation from adult tissues or embryos as described (Meedel and Whittaker 1978; Meedel and Hastings 1993). Northern blot analysis was as described (Meedel and Hastings 1993) using a radiolabeled Ci-TnI antisense riboprobe derived from the cDNA clone pCTp2 (MacLean, Meedel, and Hastings 1997) and RNA size markers from Invitrogen Life Technologies. TnI mRNA 5'-RACE was performed using the Clontech AmpliFinder 5'-RACE kit as described (Vandenberghe, Meedel, and Hastings 2001). Reverse transcription polymerase chain reaction (RT-PCR) amplification of TnI mRNAs was based on primers GCTTAGCAACGCAACAAAA and GCAACATGCCAAAGAAAAATAC in the 5'- and 3'-untranslated regions, respectively. These primers amplify a 733-bp product from adult body-wall muscle RNA, a 874-bp product from heart RNA, and a 2.1-kb product from genomic DNA (MacLean, Meedel, and Hastings 1997). Prior digestion of RNA samples with DNase I (to remove traces of genomic DNA) and conditions for reverse transcription and PCR amplification were as described (Meedel, Lee, and Whittaker 2002).
Gene Structure Analysis
We determined the complete sequence (GenBank U94694) of a previously cloned 2.1-kb genomic DNA PCR product (MacLean, Meedel, and Hastings 1997) and located exons by comparison to body-wall muscle cDNA (GenBank U55261) and heart mRNA RT-PCR product (MacLean, Meedel, and Hastings 1997) sequences. Additional 5'-flanking sequence (GenBank AF237978) was cloned by an inverse polymerase chain reaction (IPCR) procedure (Ochman, Gerber, and Hartl 1988) using TGCGGTAATAAGTGAGGTC and AGCGAGAAATGGAACAAA as primers on circularized BstYI fragments of Ciona genomic DNA (Vandenberghe, Meedel, and Hastings 2001). The resulting 2.8-kb IPCR product contained 2,067 bp of TnI DNA upstream of the ATG initiation codon; in 354 bp of overlapping sequence data, there were no differences between the IPCR product and the cloned 2.1 kb genomic PCR product.
In addition to the above-mentioned sequences, which were derived from Atlantic coast (Cape Cod) Ciona material, we also isolated TnI genomic DNA clones by hybridization screening of a lambda ZAP Express (Stratagene) phage library of partial Sau3A-cut Ciona intestinalis DNA (kindly provided by Dr. Robert Zeller) based on Pacific coast (Southern California) material, using the BstYI IPCR product as probe. Two overlapping phage clones, one of which included 5,429 bp of DNA upstream of the ATG codon, were sequenced, including all exons (GenBank AF237979). Our Pacific TnI gene sequence is >99% identical to that present in the recently released Ciona genome sequence, which was also derived from a Southern California source (Dehal et al. 2002). In a 2.6-kb segment, including all exons and introns and 127 bp upstream of the ATG initiation codon, there were only 17 single-base differences, only one of which was in a protein-coding sequence (a synonymous substitution in alanine codon 176 (body-wall muscle TnI numbering). In the "Pacific" alleles the number, length, and distribution of exons corresponded exactly to those of the sequenced "Atlantic" allele, and they were 96% identical in sequence. There were six amino acid differences, one in exon 4 (the Pacific alleles encode EAKKAEL where the Atlantic sequence has EAEKAEL), and the rest in heart-specific exons 2 and 3. Introns were of similar lengths, although markedly divergent in sequence (<80% identity), and with 24 small (113 bp) indels and one large one (203 bp), in the first intron.
Expressed Sequence Tag Analysis
Expressed sequence tag (EST) analysis of tailbud embryo and larval mRNA populations was as previously described (Satou et al. 2001; Kusakabe et al. 2002; Satou et al. 2002) based on Ciona material collected in Japan. The EST sequence database can be accessed at http://ghost.zool.kyoto-u.ac.jp./indexr1.html. The National Center for Biotechnology Information accession number for EST cluster 00173 is UniGene Cluster Cin.13140.
TnI/ß-gal Reporter Construct
The 1.5-kb TnI/ß-gal reporter construct contained Ci-TnI genomic DNA derived from the BstYI IPCR product (see above) (via a 1.8-kb KpnI fragment, 1454 to +327, subcloned into pBluescriptIISK+ (Clontech)) extending from the KpnI site at 1454 to a Bpu1102I site at 26 (nucleotides numbered with respect to the TnI ATG start codon.) The blunted Bpu1102I site was ligated to the SmaI end of a SmaI-BglII fragment of pSP72-1.27ß-gal (Corbo, Levine, and Zeller 1997) carrying the nuclear ß-gal reporter gene. The BglII end of the nuclear ß-gal DNA fragment was ligated to the BamHI site of the vector, pBluescriptIISK+. Electroporation of plasmid DNA constructs into one-celled embryos was as described by Corbo, Levine, and Zeller (1997). After 12 h, normally developed embryos were sorted and analyzed by ß-gal histochemistry. Embryos were fixed for 30 min in seawater containing 1.5% paraformaldehyde, 0.1% Tween 80; they were then washed in 0.1% Tween 80 in phosphate buffered saline and transferred to staining solution (0.04% X-Gal, 2 mM MgCl2, 0.06 M Na2HPO4, 0.04 M NaH2PO4, 4 mM potassium ferrocyanide, 4 mM potassium ferricyanide) for 1 h at room temperature.
Phylogenetic Analysis
TnI amino acid sequences encoded by DNA sequences corresponding to ancestral core exons II-V were aligned using Clustal W (Thompson, Higgins, and Gibson 1994) within the BioEdit 5.0.9 environment (Hall 1999). Phylogenetic analysis was carried out with MEGA software version 2.1 (Kumar et al. 2001) employing 100 bootstrap resamplings. Distance matrix methods (unweighted pair group method with arithmetic mean (UPGMA) and Neighbor Joining (NJ)) used pairwise gap deletion and Poisson correction. The NJ results were similar to the UPGMA results shown, except that Ciona TnI grouped more consistently with Halocynthia larval TnIs on bootstrap analysis. Maximum parsimony analysis used the close-neighbor interchange search method; PAUP 4.0 (Swofford 2000) generated the same consensus tree as did MEGA.
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Results |
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The Ci-TnI gene comprises seven exons (fig. 1). There is a great similarity of exon organization and sequence to vertebrate TnI genes, particularly in the region encoding the C-terminal half of the protein (fig. 2). Ci-TnI exons 57 correspond precisely or approximately in length to exons 68 of the vertebrate TnI genes and share 50%80% amino acid sequence identity with them. The N-terminal half of the protein is less highly conserved both in amino acid sequence (<50% identity) and in exon organization; e.g., Ci-TnI exons 1 and 4 are each represented in vertebrate TnI genes by two separate exons. Ci-TnI gene organization resembles that of the recently reported Halocynthia adult TnI gene (Yuasa et al. 2002), although the latter has three rather than two serial heart-specific exons, making a total of eight exons.
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In 43 tailbud embryo/larval Ci-TnI ESTs that extended far enough in the 5' direction to include the exon 1 junction, exon 1 was found in every case to be joined to exon 4 (body-wall-muscle splicing pattern) not exon 2 (heart splicing pattern). In 54 ESTs that extended far enough in the 3' direction to include the C-terminal protein-coding region, the complete C-terminal structure encoded by exon 7 was present in every case; there were no novel splicing products encoding C-terminally truncated TnIs similar to the Halocynthia larval TnIs. Thus the Ci-TnI gene is expressed in embryos/larvae, and the splicing pathway followed, and the TnI protein produced, correspond exactly to that used in adult body-wall muscle.
Additional studies independently supported this conclusion and gave further information. Developmental Northern blot studies, using Ci-TnI body-wall muscle cDNA clone pCTp2 (MacLean, Meedel, and Hastings 1997) as the hybridization probe, showed no hybridization with RNA from unfertilized oocytes or from embryos at 1.5, 3, or 4 h post-fertilization; a faint band of hybridization at 6 h (late gastrula); and strong and increasing hybridization at 9 h (neurula/early tailbud), 11 h (mid- tailbud), and 16 h (pre-hatch larva) of development (fig. 4A). By comparison with adult tissue RNA samples (fig. 4B), the embryonic/larval TnI mRNA corresponded in length to the body-wall muscle RNA (900 nt) rather than the longer, alternatively spliced heart RNA (1,050 nt) (MacLean, Meedel, and Hastings 1997). Reverse transcriptase PCR using primers based on Ci-TnI 5'- and 3'-untranslated mRNA sequences amplified a product from 16 h embryo RNA that co-migrated with the 733-bp product amplified from adult body-wall muscle RNA (data not shown); heart TnI mRNA generates a 874-bp product with these primers (MacLean, Meedel, and Hastings 1997). Developmental gene expression analysis by RT-PCR using the same primers confirmed the presence of Ci-TnI mRNA in 6 h embryos, as detected by Northern blot, and showed that very small amounts could be detected as early as 4 h postfertilization (64-cell stage) (data not shown). Similar temporal expression profiles have been reported for other muscle genes whose expression is developmentally activated in the larval tail muscle cell lineages (Meedel and Whittaker 1983; Makabe et al. 1990; Araki et al. 1994; Kusakabe, Hikosaka, and Satoh 1995; Meedel, Farmer, and Lee 1997).
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The foregoing evidence of Ci-TnI gene expression in tailbud embryos/larvae implies the presence of cis-regulatory elements able to drive transcription in these developmental stages. This was confirmed by transfection of a Ci-TnI reporter transgene construct into Ciona embryos by zygote electroporation. We have previously shown that a cytoplasmic ß-gal reporter gene driven by 1.5 kb of DNA upstream of the Ci-TnI ATG start codon was expressed in embryonic tail, apparently in muscle cells (Vandenberghe, Meedel, and Hastings 2001). To further clarify the cellular basis of Ci-TnI transgene expression, we prepared a nuclear-targeted ß-gal reporter driven by the same Ci-TnI upstream DNA. In transfected embryos stained at 12 h of development, nuclear staining of muscle cells was evident in almost all embryos (fig. 5). In some embryos all muscle cells showed strong nuclear ß-gal expression. In the tail, there was no staining of peripheral cells (epidermis) or of central cells (notochord, nerve cord), and there was little staining in the trunk. These transgene studies indicate that the Ci-TnI gene contains promoter/enhancer elements capable of driving tissue-specific expression in embryonic muscle cells.
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Discussion |
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Our findings in Ciona raise the possibility that the C-terminally truncated larval TnIs of Halocynthia could represent a derived, rather than an ancestral chordate, character, and this idea is strongly supported by amino acid sequence comparisons. As shown in figure 6, molecular phylogenetic analysis indicates that the gene duplication that gave rise to adult and larval TnI gene classes occurred in the tunicate TnI lineage after the divergence of the tunicate and vertebrate TnI lineages (see also Yuasa et al. [2002]). This relationship is also supported by gene structure data; all the Halocynthia TnI genes, adult and larval, share with Ci-TnI the exon 1 and exon 4 features that distinguish Ci-TnI from all the vertebrate TnI genes (see fig. 3). The level of sequence divergence between the Halocynthia adult and larval TnIs is very similar to their sequence divergence from Ciona TnI, suggesting that the initial adult/larval gene duplication occurred early in tunicate evolution. This is reflected graphically in the trifurcation of ascidian TnIs in the consensus phylogenetic trees shown in figure 6B and C. The simplest overall interpretation is that the ancestral chordate, like its descendants the vertebrates and the Enterogona ascidian Ciona, produced only C-terminally complete TnI, but that during tunicate evolution a gene duplication occurred in the Halocynthia lineage or its antecedents that permitted the evolution of larva-specific genes encoding C-terminally truncated TnIs. Consistent with its origin as an independent evolutionary innovation, the C-terminal truncation of Halocynthia larval TnIs has a different molecular genetic basis from those found in protostome invertebrates. In Halocynthia the stop codon that defines the C-terminal truncation is not created by a splicing event as in the alternatively spliced Drosophila TnI gene (Barbas et al. 1991; Beall and Fyrberg 1991), nor does it occur within ancestral core exon IV, as is the case for the tni-4 gene of the nematode Caenorhabditis (K.E.M.H., unpublished analysis), but in a downstream exon that may or may not be derived from ancestral core exon V (see Yuasa et al. 1997, 2002). All of the evolutionary analyses indicate that the C-terminally complete larval TnI of Ciona more closely reflects the ancestral ascidian larval TnI than do the larval TnIs of Halocynthia.
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Concerning modern ascidians, the biological implications of C-terminally truncated larval TnI in Halocynthia, but not in Ciona, are not clear. Current knowledge of TnI structure/function suggests that C-terminal truncation is likely to have a functional effect (Ramos 1999; Ferrieres et al. 2000; Murphy et al. 2000; Digel et al. 2001; Jin et al. 2001); however, the precise implications for contractile regulation in vivo are not clear. Indeed, there seem to be no obvious differences in larval tail muscle usage between these ascidian species. It is of interest to note that, parallel to observations on TnI, Halocynthia has separate genes encoding larval and adult isoforms of TnT (Endo et al. 1996), whereas the Ciona genome appears to contain a single TnT gene (Dehal et al. 2002; Chiba et al. 2003). It is plausible that the Halocynthia larval TnT gene could be specialized for function in a troponin complex containing a C-terminally truncated larval-specific TnI, a specialization that would not be relevant to Ciona, which uses the same TnI in larval and in adult (body-wall) muscle. Comparative functional and biochemical/biophysical studies of Halocynthia and Ciona larval tail muscle, although technically difficult because of small size, would be worthwhile and may lead to unique insight into the troponin contractile regulatory mechanism.
Because the Ciona Ci-TnI gene is expressed both in larval tail muscle and in adult body-wall muscle and heart, it presents opportunities for molecular genetic analysis of myogenic gene regulatory mechanisms operating in multiple muscle cell types and during both early embryonic development and subsequent metamorphosis/adult development. Moreover, a broader comparative analysis including the specialized adult and larval TnI genes of Halocynthia may provide insight into the evolutionary changes in gene regulatory elements that underlie the diversification of transcriptional specificities after gene duplication. Thus further studies of ascidian TnI genes are likely to lead to better understanding of the development and evolution of chordate muscle types and of the gene regulatory dynamics of gene family evolution.
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Acknowledgements |
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Footnotes |
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Literature Cited |
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Araki, I., H. Saiga, K. W. Makabe, and N. Satoh. 1994. Expression of AMD1, a gene for a MyoD1-related factor in the ascidian Halocynthia roretzi. Roux's Arch. Dev. Biol. 203:320-327.[ISI]
Ausoni, S., M. Campione, A. Picard, P. Moretti, M. Vitadello, C. De Nardi, and S. Schiaffino. 1994. Structure and regulation of the mouse cardiac troponin I gene. J. Biol. Chem. 269:339-346.
Baldwin, A. S., Jr., E. L. Kittler, and C. P. Emerson, Jr. 1985. Structure, evolution, and regulation of a fast skeletal muscle troponin I gene. Proc. Natl. Acad. Sci. USA 82:8080-8084.[Abstract]
Barbas, J. A., J. Galceran, I. Krah-Jentgens, J. L. de la Pompa, I. Canal, O. Pongs, and A. Ferrus. 1991. Troponin I is encoded in the haplolethal region of the Shaker gene complex of Drosophila. Genes Dev. 5:132-140.[Abstract]
Beall, C. J., and E. Fyrberg. 1991. Muscle abnormalities in Drosophila melanogaster heldup mutants are caused by missing or aberrant troponin-I isoforms. J Cell Biol. 114:941-951.[Abstract]
Berrill, N. J. 1955. The origin of vertebrates. Clarendon Press, Oxford, U. K.
Cameron, C. B., J. R. Garey, and B. J. Swalla. 2000. Evolution of the chordate body plan: new insights from phylogenetic analyses of deuterostome phyla. Proc. Natl. Acad. Sci. USA 97:4469-4474.
Chiba, S., S. Awazu, M. Itoh, S. Chin-Bow, N. Satoh, Y. Satou, and K. E. M. Hastings. 2003. A genomewide survey of developmentally relevant genes in Ciona intestinalis: IX. Genes for muscle structural proteins. Dev. Genes Evol. 213:291-302.[ISI][Medline]
Corbo, J. C., M. Levine, and R. W. Zeller. 1997. Characterization of a notochord-specific enhancer from the Brachyury promoter region of the ascidian, Ciona intestinalis. Development 124:589-602.
Corin, S. J., O. Juhasz, L. Zhu, P. Conley, L. Kedes, and R. Wade. 1994. Structure and expression of the human slow twitch skeletal muscle troponin I gene. J. Biol. Chem. 269:10651-10659.
Dehal, P., Y. Satou, and R. K. Campbell, et al. (87 co-authors). 2002. The draft genome of Ciona intestinalis: insights into chordate and vertebrate origins. Science 298:2157-2167.
Dhoot, G. K., and S. V. Perry. 1979. Distribution of polymorphic forms of troponin components and tropomyosin in skeletal muscle. Nature 278:714-718.[ISI][Medline]
Di Gregorio, A., and M. Levine. 1998. Ascidian embryogenesis and the origins of the chordate body plan. Curr. Opin. Genet. Dev. 8:457-463.[CrossRef][ISI][Medline]
Digel, J., O. Abugo, T. Kobayashi, Z. Gryczynski, J. R. Lakowicz, and J. H. Collins. 2001. Calcium- and magnesium-dependent interactions between the C-terminus of troponin I and the N-terminal, regulatory domain of troponin C. Arch. Biochem. Biophys. 387:243-249.[CrossRef][ISI][Medline]
Endo, T., K. Matsumoto, T. Hama, Y. Ohtsuka, G. Katsura, and T. Obinata. 1996. Distinct troponin T genes are expressed in embryonic/larval tail striated muscle and adult body wall smooth muscle of ascidian. J. Biol. Chem. 271:27855-27862.
Endo, T., and T. Obinata. 1981. Troponin and its components from ascidian smooth muscle. J. Biochem. (Tokyo) 89:1599-1608.[Abstract]
Farah, C. S., and F. C. Reinach. 1995. The troponin complex and regulation of muscle contraction. FASEB J. 9:755-767.
Ferrieres, G., M. Pugniere, J. C. Mani, S. Villard, M. Laprade, P. Doutre, B. Pau, and C. Granier. 2000. Systematic mapping of regions of human cardiac troponin I involved in binding to cardiac troponin C: N- and C-terminal low affinity contributing regions. FEBS Lett. 479:99-105.[CrossRef][ISI][Medline]
Gergely, J. 1998. Molecular switches in troponin. Adv. Exp. Med. Biol. 453:169-176.[ISI][Medline]
Gordon, A. M., E. Homsher, and M. Regnier. 2000. Regulation of contraction in striated muscle. Physiol. Rev. 80:853-924.
Hall, T. A. 1999. BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symp. Ser. 41:95-98.
Hastings, K. E. M. 1997. Molecular evolution of the vertebrate troponin I gene family. Cell Struct. Funct. 22:205-211.[ISI][Medline]
Jin, J. P., F. W. Yang, Z. B. Yu, C. I. Ruse, M. Bond, and A. Chen. 2001. The highly conserved COOH terminus of troponin I forms a Ca2+-modulated allosteric domain in the troponin complex. Biochemistry 40:2623-2631.[CrossRef][ISI][Medline]
Katz, M. J. 1983. Comparative anatomy of the tunicate tadpole, Ciona intestinalis. Biol. Bull. 164:1-27.[ISI]
Kobayashi, T., T. Takagi, K. Konishi, and J. A. Cox. 1989. Amino acid sequence of crayfish troponin I. J. Biol. Chem. 264:1551-1557.
Kumar, S., K. Tamura, I. B. Jakobsen, and M. Nei. 2001. MEGA2: molecular evolutionary genetics analysis software. Bioinformatics 17:1244-1245.
Kusakabe, T., I. Araki, N. Satoh, and W. R. Jeffery. 1997. Evolution of chordate actin genes: evidence from genomic organization and amino acid sequences. J. Mol. Evol. 44:289-298.[ISI][Medline]
Kusakabe, T., A. Hikosaka, and N. Satoh. 1995. Coexpression and promoter function in two muscle actin gene complexes of different structural organization in the ascidian Halocynthia roretzi. Dev. Biol. 169:461-472.[CrossRef][ISI][Medline]
Kusakabe, T., R. Yoshida, and I. Kawakami, et al. (11 co-authors). 2002. Gene expression profiles in tadpole larvae of Ciona intestinalis. Dev. Biol. 242:188-203.[CrossRef][ISI][Medline]
MacLean, D. W., T. H. Meedel, and K. E. M. Hastings. 1997. Tissue-specific alternative splicing of ascidian troponin I isoforms. Redesign of a protein isoform-generating mechanism during chordate evolution. J. Biol. Chem. 272:32115-32120.
Makabe, K. W., S. Fujiwara, H. Saiga, and N. Satoh. 1990. Specific expression of myosin heavy chain gene in muscle lineage cells of the ascidian embryo. Roux's Arch. Dev. Biol. 199:307-313.[ISI]
Meedel, T. H. 1998. Development of ascidian muscles and their evolutionary relationship to other chordate muscle types. Pp. 305330 in K. Adiyodi and R. Adiyodi, eds. Progress in Developmental Biology, Reproductive Biology of Invertebrates. Wiley-Interscience, New York.
Meedel, T. H., S. C. Farmer, and J. J. Lee. 1997. The single MyoD family gene of Ciona intestinalis encodes two differentially expressed proteins: implications for the evolution of chordate muscle gene regulation. Development 124:1711-1721.
Meedel, T. H., and K. E. M. Hastings. 1993. Striated muscle-type tropomyosin in a chordate smooth muscle, ascidian body-wall muscle. J. Biol. Chem. 268:6755-6764.
Meedel, T. H., J. J. Lee, and J. R. Whittaker. 2002. Muscle development and lineage-specific expression of CiMDF, the MyoD- family gene of Ciona intestinalis. Dev. Biol. 241:238-246.[CrossRef][ISI][Medline]
Meedel, T. H., and J. R. Whittaker. 1978. Messenger RNA synthesis during early ascidian development. Dev. Biol. 66:410-421.[ISI][Medline]
1983. Development of translationally active mRNA for larval muscle acetylcholinesterase during ascidian embryogenesis. Proc. Natl. Acad. Sci. USA 80:4761-4765.[Abstract]
Murphy, A. M., H. Kogler, D. Georgakopoulos, J. L. McDonough, D. A. Kass, J. E. Van Eyk, and E. Marban. 2000. Transgenic mouse model of stunned myocardium. Science 287:488-491.
Myers, C. D., P. Y. Goh, T. S. Allen, and E. A. Bucher, T. Bogaert. 1996. Developmental genetic analysis of troponin T mutations in striated and nonstriated muscle cells of Caenorhabditis elegans. J. Cell Biol. 132:1061-1077.[Abstract]
Ochman, H., A. S. Gerber, and D. L. Hartl. 1988. Genetic applications of an inverse polymerase chain reaction. Genetics 120:621-623.
Perry, S. V. 1999. Troponin I: inhibitor or facilitator. Mol. Cell. Biochem. 190:9-32.[CrossRef][ISI][Medline]
Ramos, C. H. 1999. Mapping subdomains in the C-terminal region of troponin I involved in its binding to troponin C and to thin filament. J. Biol. Chem. 274:18189-18195.
Satou, Y., N. Takatori, S. Fujiwara, T. Nishikata, H. Saiga, T. Kusakabe, T. Shin-i, Y. Kohara, and N. Satoh. 2002. Ciona intestinalis cDNA projects: expressed sequence tag analyses and gene expression profiles during embryogenesis. Gene 287:83-96.[ISI][Medline]
Satou, Y., N. Takatori, and L. Yamada, et al. 2001. Gene expression profiles in Ciona intestinalis tailbud embryos. Development 128:2893-2904.
Squire, J. M., and E. P. Morris. 1998. A new look at thin filament regulation in vertebrate skeletal muscle. FASEB J. 12:761-771.
Swalla, B. J., C. B. Cameron, L. S. Corley, and J. R. Garey. 2000. Urochordates are monophyletic within the deuterostomes. Syst. Biol. 49:52-64.[CrossRef][ISI][Medline]
Swofford, D. L. 2000. PAUP 4.0: phylogenetic analysis using parsimony (and other methods). Sinauer Associates, Sunderland, Mass.
Talbot, J. A., and R. S. Hodges. 1981. Synthetic studies on the inhibitory region of rabbit skeletal troponin I. Relationship of amino acid sequence to biological activity. J. Biol. Chem. 256:2798-2802.
Thompson, J. D., D. G. Higgins, and T. J. Gibson. 1994. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res. 22:4673-4680.[Abstract]
Toyota, N., T. Obinata, and K. Terakado. 1979. Isolation of troponin-tropomyosin-containing thin filaments from ascidian smooth muscle. Comp. Biochem. Physiol. 62B:433-441.
Vandekerckhove, J., and K. Weber. 1984. Chordate muscle actins differ distinctly from invertebrate muscle actins. The evolution of the different vertebrate muscle actins. J. Mol. Biol. 179:391-413.[ISI][Medline]
Vandenberghe, A. E., T. H. Meedel, and K. E. M. Hastings. 2001. mRNA 5'-leader trans-splicing in the chordates. Genes Dev. 15:294-303.
Wada, H. 1998. Evolutionary history of free-swimming and sessile lifestyles in urochordates as deduced from 18S rDNA molecular phylogeny. Mol. Biol. Evol. 15:1189-1194.[Abstract]
Wilkinson, J. M., and R. J. Grand. 1978. Comparison of amino acid sequence of troponin I from different striated muscles. Nature 271:31-35.[ISI][Medline]
Yuasa, H. J., K. Kawamura, H. Yamamoto, and T. Takagi. 2002. The structural organization of ascidian Halocynthia roretzi troponin I genes. J. Biochem. (Tokyo) 132:135-141.[Abstract]
Yuasa, H. J., S. Sato, H. Yamamoto, and T. Takagi. 1997. Primary structure of troponin I isoforms from the ascidian Halocynthia roretzi. J. Biochem. (Tokyo) 122:374-380.[Abstract]