Skip to main content

Transposon mediated transgenesis in a marine invertebrate chordate: Ciona intestinalis


Achievement of transposon mediated germline transgenesis in a basal chordate, Ciona intestinalis, is discussed. A Tc1/mariner superfamily transposon, Minos, has excision and transposition activities in Ciona. Minos enables the creation of stable transgenic lines, enhancer detection, and insertional mutagenesis.


DNA transposons are powerful tools for genetic analyses. Transposons are employed for creation of stable transgenic lines, enhancer detection, gene trapping, and insertional mutagenesis. These transposon-mediated techniques have been facilitated by the discovery and reconstruction of active transposons in several organisms [111]. Despite their utility, transposon technologies are restricted to a few model organisms. Marine invertebrates include most of the phyla whose study is crucial to elucidating the evolutionary molecular mechanisms of diversification in metazoans. To date, transposon technologies have been introduced for only a few marine invertebrate species. Because of the scarcity of refined genetic techniques, research into gene functions in marine invertebrates has remained limited.

Recent achievement of germline transgenesis in a marine invertebrate chordate, Ciona intestinalis, has altered this situation [12, 13]. C. intestinalis has several characteristics that make it amenable for genetics research. In this basal chordate, a Tc1/mariner superfamily transposon, Minos, has the complete activity required for its transposition [14]. Minos introduced into Ciona is excised from a plasmid vector by transposase and is integrated into TA dinucleotides of another DNA molecule. The TA dinucleotides are known as target sequences of Tc1/mariner transposons [15]. Transposition occurs in Ciona germ cells, and Minos is inserted into the chromosomes of germ cells [12]. The insertions are inherited stably by subsequent generations, thereby creating stable transgenic lines. Using this transformation technique, genetic techniques such as enhancer detection and insertional mutagenesis have been introduced into Ciona using Minos [13, 16, 17]. In this article, recent achievements with transposon techniques in Ciona, as well as characteristics of Ciona as a new genetic model, are discussed.

Characteristics of Ciona intestinalisas an experimental system for genetics

Ascidians, or sea squirts, are members of the subphylum Tunicata [18]. Tunicata belong to the phylum Chordata along with Cephalochordata (amphioxus) and Vertebrata (Figure 1a) [19, 20] (Kawashima T, Putnam N, personal communication). As this phylogenetic position suggests, ascidians possess a simplified chordate body plan. This characteristic is most apparent in their larval stage. The larvae of ascidians are typical tadpole larvae and swim like fish (Figure 1b). Each larva has a dorsal hollow neural tube and notochord, both of which represent common characteristics of chordates. In contrast to those apparent similarities, the ascidian larval body is strikingly simple compared with that of vertebrate tadpoles. The typical ascidian larva consists of numerous, but countable, cells [18]. For example, the larvae of C. intestinalis consist of approximately 2,600 cells, of which 40 constitute the notochord, 26 make up the muscle, and about 330 the nervous system [18, 21]. Ascidians develop rapidly; many ascidians complete embryogenesis within 1 day. This simplicity and rapid embryogenesis aid detailed cell-by-cell analyses of the mechanisms of tadpole body formation. In fact, ascidians are the only chordates for which cell lineages have been described [2224].

Figure 1

An ascidian - Ciona intestinalis. (a) Phylogenetic relationships of chordates. Ascidians are included in the subphylum Tunicata. (b) A Ciona intestinalis larva. This photograph was constructed by merging three photographs of the same individual. Scale bar: 100 μm. (c) Ciona intestinalis adults. After metamorphosis, Ciona loses its tail and starts to settle. Most ascidians are filter feeders.

Ascidian larvae change their structure through metamorphosis and become sessile adults (Figure 1c) [25]. After metamorphosis, ascidians start to take in food by filter feeding. During metamorphosis, the larval tail is lost and adult tissues grow rapidly, which include characteristic chordate structures such as pharyngeal gills and an endostyle (the endostyle is homologous to vertebrate thyroid gland [26, 27]). Metamorphosis of typical ascidians is completed within several days. Metamorphosis is a dramatic event in ascidian development and provides a good experimental system in which to uncover the mechanisms of metamorphosis and their conservation among vertebrates.

Ciona intestinalis (hereafter referred to as Ciona) is a cosmopolitan ascidian [2831]. It is hermaphroditic and self-fertile. This characteristic represents a great advantage when performing mutant screens because creating animals that are homozygous with respect to mutation sites is possible without genotyping [32, 33]. An adult Ciona bears thousands of eggs; eggs and sperm can be collected surgically.

Surgically collected eggs can be fertilized with sperm from a different individual. They subsequently exhibit normal, synchronized development. Natural spawning can be induced by simple dark-light adjustment to facilitate self-fertilization. Moreover, sperm can be stored on ice for 1 week without loss of fertility. Cryopreservation of sperm is also established to store mutants or transgenic lines semipermanently [34]. The easy handling of germ cells enables reduction in labor associated with mutant screening and preservation of lines.

The most striking characteristic that distinguishes Ciona from other ascidians is the availability of a draft genome sequence [35]. The Ciona genome size is approximately 166 megabases per haploid, which contains 15,852 protein coding genes [36]. The genome size and gene number are comparable to those of Drosophila melanogaster, and much smaller than those of most vertebrates. In addition, the Ciona genome is less redundant than those of vertebrates [37, 38], which is probably related to the twofold to threefold duplication of genomes during vertebrate evolution [39]. Therefore, Ciona possesses the basic set of genes for a chordate body plan. Because of its compact genome, Ciona provides a simple experimental system in which to uncover genetic mechanisms that specify the chordate body plan as well as mechanisms of chordate evolution.

An unusual aspect of the Ciona genome with respect to transposons is that an extensive search of the Ciona draft genome identified no Tc1/mariner transposon (Table 1). Taking into consideration the global conservation of this transposon family [40], the absence of Tc1/mariner transposon in Ciona genome is curious. Two possibilities are readily apparent. One is that Tc1/mariner transposon has been lost in the Ciona, ascidians, or tunicates branch by accumulation of mutations. Another possibility is that a hypothetical suppressor of this transposon family interfered with the lateral transfer of transposons. This might be related to the weak transposon activity of several Tc1/mariner transposons in Ciona, as discussed below.

Table 1 BLAST search of Tc1/mariner superfamily transposons in five eukaryotic genomes

In Ciona, techniques to support the practice of genetics research have been developed. The Ciona life cycle is about 2 to 3 months. An inland culture system has been established [17, 34]. Settlement after metamorphosis enables retention of several lines in the same aquarium. Introduction of DNA and RNA into eggs by microinjection or electroporation is performed routinely [41, 42]. The latter technique can introduce DNA and RNA into hundreds of eggs within 1 hour, thereby facilitating creation of transgenic lines.

There are three major obstacles to use of Ciona to conduct genetics studies. First, no inbred strain has been created; most experiments are dependent on natural populations. Creating strains had been difficult because of complications in culturing. Recent improvements of inland culture systems are expected to resolve this problem [43]. Second, natural Ciona harbor many single nucleotide polymorphisms. The genome project reported that 1.2% of nucleotide differences were observed between alleles of the single individual [35]. This score is 15 times higher than that in humans, and three times higher than that in pufferfish. Such highly frequent polymorphism would render it difficult to perform systematic fine mapping of point mutations. On the contrary, high polymorphism might allow retention of highly frequent natural mutants, which are a valuable resource for mutant screening. In C. intestinalis, and its related species C. savignyi, several mutants have been isolated through screening of wild populations [34, 44, 45]. The third obstacle to genetics studies is the requirement for seawater for culture. Large-scale culturing requires a considerable amount of seawater, which limits the culturing of Ciona to laboratories that are near to the sea. Recently, Ciona culture with artificial seawater has been achieved [34, 43], which will promote the spread of Ciona studies to inland laboratories.

Activity of Minos transposon in Ciona

Minos is a member of the Tc1/mariner superfamily of transposons isolated from Drosophila hydei [5]. Minos exhibited both excision and transposition activity from protostomes to deuterostomes [4654], suggesting a wide host range. Minos is the only transposon whose activity has been described in Ciona [1214, 55, 56]. Its excision is observed in almost all embryos when Minos is injected into Ciona embryos together with transposase mRNA (Figure 2). Footprint sequences indicate that Minos is excised correctly by transposase. The typical footprint sequences of Minos are 5'-TACTCGTA-3' or 5'-TACGAGTA-3'; both typical and atypical footprint sequences are observed in Ciona [14, 57]. The atypical footprint sequences might be related to the endogenous repair system of Ciona. Neither excision nor transposition occurs without transposases, suggesting that no Ciona protein mimics Minos activity. Interplasmid transposition assay using donor and recipient plasmids (Figure 2) has revealed that Minos has slightly lower transposition activity in Ciona than in insects [14, 49]. The manner of insertion of Minos into the recipient plasmid is identical to that previously reported; the target sequences are TA dinucleotides and the duplication of the TA sequences occurs, which flanks two inverted repeats. The frequency of excision and transposition activity suggests that Minos has sufficient activity to cause germline transgenesis in Ciona, as shown by microinjection of transposase mRNA with recombinant Minos containing a promoter-green fluorescent protein (GFP) cassette. The scheme of screening transgenic lines is shown in Figure 3. About 30% to 36% of Minos-injected Ciona become founders and transmit Minos insertions to progeny. The average insertion number inherited from a founder was estimated at around two (Sasakura Y, unpublished data). This transgenesis frequency is comparable to that of Sleeping Beauty (SB) in zebrafish [58]. Thermal asymmetric interlaced (TAIL)-polymerase chain reaction (PCR) is used to identify Minos insertion sites [12, 59]. Minos was preferably inserted into TA-rich sequences such as introns [12].

Figure 2

Procedures of experiments testing transposon activity in Ciona. (a) Excision assay. A mixture of transposon vector and in vitro transcribed transposase mRNA was injected into Ciona one-cell embryos. After these embryos reach the late tailbud stage, DNA was extracted from embryos and was subjected to polymerase chain reaction analyses to identify the excision. (b, c) An example of excision assay: a Minos vector (pMiFr3dTPOG-B1PM) with an insert longer than 12 kilobases was examined. Excised bands are detected only when the transposase mRNA is co-injected. (d) Interplasmid transposition assay. A donor vector containing a Minos construct and a target vector are introduced into Ciona embryos with transposase mRNA. Plasmids are recovered from Ciona embryos and are used to transform Escherichia coli. The occurrence of transposition was monitored by selection of E. coli using antibiotics. bp, base pairs; ORF, open reading frame.

Figure 3

Creation of stable transgenic lines with Minos and mutant screening. Mixture of a transposon vector containing a promoter-gfp cassette and transposase mRNA is introduced into Ciona one-cell embryos by microinjection or electroporation. These animals are cultured until gamete maturation. Generally, sperm from Minos-introduced individuals are used to fertilize wild-type eggs to obtain F1 family individuals, which are screened to find green fluorescent protein (GFP)-positive animals. The GFP-positive animals are selected for further culture, thereby establishing transgenic lines. The F1 animals with matured eggs and sperm are subjected to self-fertilization to obtain F2 family individuals, which are screened to identify mutant phenotypes. ORF, open reading frame.

Another convenient transgenesis technique of Ciona with Minos was achieved using electroporation [56]. As described above, electroporation enables rapid and reproducible transgenesis of early Ciona embryos. This technique simultaneously electroporates Minos DNA and in vitro synthesized Minos transposase mRNA in Ciona embryos. The transformation frequency by electroporation mediated transgenesis is about 20% to 30%, which is lower than that by microinjection mediated transgenesis, perhaps because of a lower amount of mRNA introduced into embryos by electroporation. By microinjection, 5 to 10 ng/μl of Minos DNA and 50 to 200 ng/μl of transposase mRNA are included in the injected solution. The current electroporation method requires 60 μg of Minos DNA and 60 μg of transposase mRNA, which would correspond to 5 to 10 ng/μl of DNA and RNA in the injection solution. Nevertheless, electroporation mediated transgenesis is now the main strategy of Ciona transformation because of its convenience.

Minos exhibited constant excision and transformation activity, even when the length of insertion is sufficiently long to suppress transposition of another Tc1/mariner transposon, namely SB [60]. So far, an insert size of up to 10 kilobases has been found to have no adverse effect on insertion efficiency (Sasakura Y, unpublished data). Such flexibility of Minos with respect to insert length allows the creation of various transposon constructs that are appropriate for experimental purposes.

Activity of other Tc1/mariner transposons in Ciona

The identification of other active transposons would make transposon technology more versatile in Ciona, because it would be useful to create 'jump starter' lines of Minos. Modifier screens of mutants generated by Minos must be done using a different transposon. Different transposons can be expected to have different insertion site preferences. Therefore, execution of large-scale mutagenesis with two transposons would be effective for saturation mutagenesis. In addition to these technical innovations, description of activity of transposons in various organisms is necessary to elucidate cross-species activity of transposons and the mechanisms that determine transposon activity in nonhost organisms. Such knowledge would be valuable for further improvement of transposon technologies. Transposon activity in marine invertebrates has not been described, except for Minos in Ciona and in a crustacean [12, 53]. Ciona is the pioneer organism of transposon technology among marine invertebrates; testing of various transposons in this organism is important.

The Tc1/mariner superfamily includes many transposons whose consistent activity in several protostomes and vertebrates has been described [6164]. We have tested some of these transposons, including SB, Frog Prince (FP), and Mos1, in Ciona. The former two transposons are resurrected transposons that are derived from vertebrate genomes [8, 11]; Mos1 was isolated from the insect Drosophila mauritiana [3, 4]. All three transposons are active in vertebrates [11, 61, 63, 64]. Excision activity of the three transposons was examined using a PCR-based assay (Figure 2a), and SB, FP, and Mos1 exhibited excision in Ciona (Figure 4). Their excisions have been supported by the presence of footprint sequences (Figure 4). However, the excision efficiency of these transposons was lower than that of Minos. When the excision efficiencies of transposons were compared with the same condition (5 to 10 ng/μl of transposon DNA and 50 to 200 ng/μl of transposase mRNA in the injection solution), Minos showed excision in almost all embryos, whereas Mos1, SB, and FP showed excision in only a few embryos. For example, nine out of 16 embryos exhibited excision in the case of Mos1, two out of 16 in the case of SB, and eight out of 32 in the case of FP. Interplasmid transposition assay and germline transformation of Ciona with Mos1, SB, and FP were also tested, but no transposition was detected (Awazu S, Sasakura Y, unpublished data).

Figure 4

Excision activity of Mos1, SB, and FP transposons in Ciona. (a) (top part) Excision of Mos1. The left panel shows the polymerase chain reaction result of Mos1 transposon and transposase-injected embryos; the right panel shows the results of Mos1 transposon-injected control embryos. The expected sizes of correct excision events are shown by arrowheads. (bottom part) Footprint sequences of Mos1 observed in Ciona and typical footprint sequences reported previously (typical footprint). (b) Excision of Sleeping Beauty (SB). Note that three to six times more transposon DNA was injected in this experiment, which resulted in the detection of excised bands from every embryo. (c) Excision of Frog Prince (FP). bp, base pairs; M, marker lane.

What might restrict Mos1, SB, and FP activity in Ciona? One possibility is that co-factors that are required for transposase activity are incompatible or absent in Ciona. In fact, SB and FP are transposons derived from vertebrates [8, 11], and therefore they retain high activity in vertebrates, indicating that all sets of co-factors required for SB and FP activity are present in vertebrates. Recent studies have revealed necessary co-factors for SB transposases [65, 66]. Although Ciona contains the basic set of genes for the chordate body plan, many genes are specific to vertebrates. The supply of such co-factors may be necessary to make transposons active in Ciona if SB and FP transposases require such vertebrate-specific co-factors. An alternative possibility is that a factor is present that inhibits transposases. Inhibition of the transposase activity has been reported in the Tn5 transposon of Escherichia coli, in which an inhibitor of the transposition protein (a truncated form of Tn5 transposase that does not possess DNA-binding activity) forms a complex with Tn5 transposase and interferes with transposition [67]. The presence of such an inhibitor has not been demonstrated in Tc1/mariner transposons, but the possibility remains that there is a Ciona protein that binds transposases and inhibits their activity.

The inefficiency of Mos1, SB and FP in Ciona implies that activity of transposons must be tested in each animal model to seek an active transposon. Identification of factors that restricts the activity of transposons is necessary to make them more valuable tools for genetics research in various organisms.

Enhancer detection

The compact genome of Ciona is a convenient feature for studying regulatory elements of gene expression [68, 69]. High density of enhancer elements is expected in the Ciona genome, facilitating efficient enhancer detection, which is necessary to identify enhancers that cannot be identified using conventional cis element analyses. This technique is also useful in creating marker lines that express reporter genes in a tissue-specific manner. In Ciona, techniques of germline transgenesis were established recently, but to date only a few marker transgenic lines are available. Enhancer detection will provide useful marker lines for future genetics studies. In addition, novel tissues or subpopulations of tissues are identifiable by enhancer detection that were previously unidentifiable by simple observation or in situ hybridization (Awazu S, Sasakura Y, unpublished data).

In Ciona, enhancer detection by microinjection mediated transgenesis has been reported [16]. An enhancer detection line near the musashi orthologous gene (Ci-musashi [16]) has been identified among 21 Minos injected animals [12]. A recent enhancer detection screen using a promoter of the Ciona thyroid peroxidase orthologous gene (Ci-TPO) yielded six enhancer detection lines from 110 injected animals [70]. The frequency of enhancer detection in Ciona is therefore estimated at 4.7% to 5.4% per injected animal. This frequency is higher than that of SB mediated enhancer detection in zebrafish (2.5% per injected animals [71]), but lower than that of Tol2 mediated enhancer detection (12% [72]).

Thus far, the promoter of Ci-TPO is the only promoter that has been used for enhancer detection in Ciona. It includes 860 base pairs of upstream sequence from the initiation codon of the gene and exhibits weak expression in endodermal tissues [72]. This might not be an ideal promoter for enhancer detection in all tissues. There might be enhancers to which Ci-TPO promoter could not respond, because minimal promoters exhibit different responsiveness to enhancers (Lemaire P, personal communication). In fact, most enhancer detection lines with Ci-TPO promoter showed reporter gene expression in endodermal tissues [70]. Comparing the efficiency of enhancer detection between the Ci-TPO promoter and a basal promoter or a minimal promoter derived from a housekeeping gene may be necessary to identify an ideal promoter for enhancer detection in Ciona.

In the Ci-musashi enhancer detection line, Minos was inserted into an intron [16]. Detailed analysis of the line revealed that expression of Ci-musashi is regulated by many enhancers located at the 5' upstream region and in introns [16]. These enhancers have both redundant and distinct functions for gene expression. Such an enhancer complex is probably necessary to ensure the appropriate spatial and temporal expression of Ci-musashi. Enhancer identification in the context of chromosomes is necessary to understand the in vivo function of these enhancers. Enhancer detection is a viable method for this purpose.

Remobilization of Minos in Cionagenome

Non-autonomous transposons in the genome can be remobilized by providing transposase mRNA (Figure 5). This technique is useful for creating new insertions and 'local hopping', and for creating new mutant alleles by deletions. Remobilization of Minos within the Ciona genome was achieved by microinjection of transposase mRNA into embryos whose respective genomes contain tandem arrays of Minos (Figure 5a[70]). This method has been used for enhancer detection [70].

Figure 5

Remobilization of Minos in the Ciona genome. (a) Remobilization from a tandem array. (b) An example of enhancer trap lines created using remobilization technique. Green fluorescent protein is expressed in the pharyngeal gill. Bar = 100 μm. (c) Remobilization of Minos from a single inserted site. (d) Excision of single Minos insertion in the somatic cells. The polymerase chain reaction bands show Minos excision from the genome (arrowhead). M, marker lane.

We created a transgenic 'mutator' line harboring a tandem array of Minos vector for enhancer detection, which contains a promoter of Ci-TPO [70]. The tandem array in the mutator line was estimated to include as many as 255 transposons. In this study, remobilization of a few copies of Minos copies probably occurred from the concatemer. Screening enhancer detection using the remobilization technique was conducted as follows. Transposase mRNA was injected into unfertilized wild-type eggs. These eggs were fertilized with sperm from the mutator line. Because our enhancer detection vector shows GFP expression in a part of somatic cells, these transposase-introduced Ciona were selected to remain as GFP positive, transposon containing animals. These GFP positive animals were crossed with wild-type individuals; then, the GFP expression pattern in the next generation was monitored to screen families exhibiting altered GFP expression.

The results indicated that 79% of transposase-injected animals transmitted enhancer detection insertions (Figure 5b). This frequency is considerably higher than that seen in the microinjection mediated approach. Although many of the enhancer detection lines showed GFP expression in endodermal tissues, a few lines showed expression in ectodermal or mesodermal tissues. Therefore, this method could be more efficient for large-scale enhancer detection with creation of many valuable lines. The tandem array interferes with detailed analyses of insertions by Southern blot and TAIL-PCR. In fact, Southern blot was done to show the presence of novel insertions created by remobilization. However, the signal was not conspicuous in many individuals, and as a result the number of new insertions is likely to have been underestimated. Identification of new insertion sites by TAIL-PCR was performed after digestion of genomic DNA with restriction enzymes to suppress PCR amplification within the concatemer [73]. Numerous lines have insertion sites that were unidentifiable, even after restriction enzyme treatment. The enhancer detection insertions can be segregated from the original tandem array by passing through several generations. This may result in the establishment of transgenic lines that have a single insertions of enhancer detection in their genome. Characterization of their insertion sites may increase the efficiency of identification of the causal insertions that were obtained using the remobilization technique.

Remobilization of a single Minos insertion might reduce these problems (Figure 5c). Several tests of remobilization of a single insertion have been carried out using microinjection of transposase mRNA into embryos of transgenic lines (Sasakura Y, unpublished data). In somatic cells excision events were observed (Figure 5d). However, the frequency of excision appeared to be low, and evidence of excision or transposition in the germ cells was not obtained. The primordial germ cells of ascidians are suggested to be two small cells, called B7.6, in early embryogenesis [74]. Thus, germ cells are derived from a small number of primordial cells. Less injected transposase mRNA would be delivered to germ cells than to somatic cells. Therefore, the frequency of excision and transposition in the germ cells would be much lower than in the somatic cells. A technical innovation, such as generation of 'jump starter' lines, is necessary to achieve highly frequent jumping of a single Minos copy in germ cells [75].

Insertional mutagenesis

Insertions of Minos can disrupt gene function to create mutants. Insertional mutants are distinguishable from background mutations by the fact that they segregate with the insertions. In Ciona, a small-scale mutagenesis screen was carried out using self-fertilization (Figure 3), and two insertional mutants were isolated from 120 transgenic lines, which are estimated to correspond to 240 insertions; one mutant can be isolated for every 120 insertions. The mutant frequency is lower than with insertional mutagenesis with pseudotyped retrovirus in zebrafish (one mutant per 85 insertions [76, 77]). Taking into consideration the compact genome of Ciona, which has less redundancy, it is curious that insertional mutagenesis in Ciona would be less efficient than in zebrafish. There are two possible explanations. One is that the preference of the insertion sites in the gene, such as 5' end, introns, exons, or 3' end, might be different between Minos and pseudotyped retrovirus. In the zebrafish approach, approximately 60% of the mutagenic insertions reside in the promoter, first exon, or first intron [77]. As mentioned above, Minos is preferably inserted into TA-rich sequences such as introns and intergenic regions. The second possible explanation is that pseudotyped retrovirus would be more mutagenic than Minos. Introduction of a gene trap cassette into the pseudotyped retrovirus vector did not affect the mutation frequency [77, 78]. Pseudotyped retrovirus might interfere with splicing to produce truncated proteins, even without such a cassette. In contrast, the single Minos insertions into introns appeared to be insufficient to cause mutations (Sasakura Y, Awazu S, unpublished data). The mutant frequency would therefore reflect the difference between two vectors.

From mutant screening, one insertional mutant has been characterized in detail [17]. In this mutant, an insertion at the promoter of a gene encoding cellulose synthase (Ci-CesA [79]) disrupts expression of this gene. Animals homozygous for this insertion exhibit abnormalities in the process of metamorphosis. At the larval stage, their trunks show post-metamorphosed states, although they retain tails, which would normally be lost during metamorphosis. The trunk-metamorphosed larvae continue to swim vigorously. This mutant was named swimming juvenile (sj). This mutant showed a novel function of animal cellulose synthase for the process of normal metamorphosis as well as for the biosynthesis of cellulose. As described above, a concatemer of Minos is inserted into the promoter of Ci-CesA. In another insertional mutant (Matsuoka T, Sasakura Y, unpublished data), a concatemer of Minos is inserted into an intron. Such concatemers are very long and may therefore disrupt promoters or introns. However, mutations by a single insertion are superior to concatemers; some refinement of transposon vectors, such as gene trap, is necessary to produce highly mutagenic Minos. Recently, we attempted to introduce a gene trap method into Ciona (Oogai Y, Sasakura Y, unpublished data).

Because insertional mutagenesis with Minos has been achieved, the next step will be saturation mutagenesis using this transposon. Ciona contains a smaller set of genes with less redundancy than in vertebrates. This characteristic renders this ascidian a suitable organism for saturation mutagenesis. It is necessary to estimate the frequency of essential genes for development in order to calculate the number of transgenic lines that are necessary for saturation mutagenesis. For such estimation, isolation of more mutants is necessary. In addition, several obstacles to Ciona genetics must be overcome in order to conduct saturation mutagenesis. One is the need to create a mutagenic Minos construct. Other obstacles are associated with the primitive state of Ciona genetics, resulting from its short history. Although we take only Ciona into consideration here, most of these points also pertain to other marine invertebrates.

In the mutant screen, we used a Minos construct with a GFP reporter. The expression of GFP was used to judge whether mutations are associated with insertions. However, a correlation between a mutation and GFP expression does not always indicate that the mutation is caused by a Minos insertion. A wild population of Ciona was used to create insertional mutants. Wild populations maintain frequent background mutations. Sometimes these natural mutation sites are located very close to Minos insertion sites, and therefore natural mutants, so-called associated mutants, appear to be related to the Minos insertions. These associated mutants must be discriminated from insertional mutants because transposon insertion sites in the associated mutants are close, but not identical, to the actual sites of mutations. In the recent small-scale mutagenesis studies [13], four mutants exhibited strong correlation with GFP expression (>90% of homozygous mutants showed GFP expression). Two of them were associated mutants and two were insertional mutants. Associated mutants are distinguishable from insertional mutants by imperfect correlation between mutations and GFP expression. Reporter gene expression is a good marker for this purpose, because hundreds of mutants can be examined through simple observation. The mutants showing perfect correlation with GFP expression are candidates for insertional mutants. Several experiments must be performed to conclude that they actually are insertional mutants. Identification of the insertion sites is primarily required. It is also necessary to demonstrate perfect homozygosity of mutants with respect to the insertions, which is evidence that recessive insertional mutants have been created. Finally, to establish a causal link, it is necessary to identify those genes that are responsible for mutants; this may be achieved through rescue experiments or knockdown of genes by microinjection of antisense morpholino oligonucleotides or dominant negative forms [17, 42].

The second disadvantage of Ciona, after its suboptimal mutation frequency, is that its embryos sometimes develop poorly compared with those of other model organisms. Typical unhealthy development includes kinked tails at the larval stage. Families showing such unhealthy development are omitted from screens. Such omission might cause the loss of mutants that would show the kinky-tail phenotype. Several insertional mutants might have been lost through this technical limitation. Therefore, the mutation frequency with Minos described above might have been underestimated. Recent improvements in culturing systems will enable continual production of healthy embryos. If a family shows unhealthy development in this setting, then the phenotype is probably derived from mutations.

Forward genetics is a powerful technique in which to identify gene function; it is possible to identify gene functions that are neglected by reverse genetics. This approach has recently been employed in Ciona with the chemical mutagen N-ethyl-N-nitrosourea and Minos transposon [17, 32, 33, 80, 81]. Causal genes have been identified in only a few mutants. Most of the mutants generated in the near future would therefore be novel ones. Insertional mutagenesis provides an ideal system in Ciona because the causal genes are identifiable in a short period of time, without time consuming fine mapping.


In this article we review recent achievements in germline transgenesis with the Minos transposon in Ciona intestinalis. These studies have revealed that Minos is a highly active transposon in this organism, as shown by establishment of techniques such as stable transgenic lines, enhancer detection, and insertional mutagenesis. These technical innovations will be of great value to future genetic analyses in C. intestinalis. Frequent enhancer detection by remobilization will provide useful transgenic lines. Insertional mutagenesis allows the identification of novel functions of genes during development, as shown by the example of cellulose synthase. Taking into consideration the advantages of Ciona as a subject of genetics research, future genetic analyses in this organism will provide unique insights into chordate gene function. In addition to these technical innovations with Minos, we describe several technical hurdles that Ciona researchers must overcome if they are to conduct large-scale mutagenesis studies.

Minos is a valuable transposon, and its activity may be the first to be tested in organisms for which no genetic approach has yet been introduced. We also provide evidence that some other Tc1/mariner superfamily transposons have excision activity in Ciona. However, these transposons have not been found to be efficient in causing germline transgenesis in Ciona. This information may be useful in elucidating the mechanisms that determine transposon activity in different organisms. Resolving these issues would make these transposons further valuable tools in Ciona genetics research.


  1. 1.

    Rubin GM, Kidwell MG, Bingham PM: The molecular basis of P-M hybrid dysgenesis: the nature of induced mutations. Cell. 1982, 29: 987-994. 10.1016/0092-8674(82)90462-7.

    PubMed  CAS  Article  Google Scholar 

  2. 2.

    Emmons SW, Yesner L, Ruan KS, Katzenberg D: Evidence for a transposon in Caenorhabditis elegans. Cell. 1983, 32: 55-65. 10.1016/0092-8674(83)90496-8.

    PubMed  CAS  Article  Google Scholar 

  3. 3.

    Jacobson JW, Medhora MM, Hartl DL: Molecular structure of a somatically unstable transposable element in Drosophila. Proc Natl Acad Sci USA. 1986, 83: 8684-8688. 10.1073/pnas.83.22.8684.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  4. 4.

    Bryan G, Garza D, Hartl D: Insertion and excision of the transposable element mariner in Drosophila. Genetics. 1990, 125: 103-114.

    PubMed  CAS  PubMed Central  Google Scholar 

  5. 5.

    Franz G, Savakis CC: Minos, a new transposable element from Drosophila hydei, is a member of the Tc1-like family of transposons. Nucleic Acid Res. 1991, 19: 6646-10.1093/nar/19.23.6646.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  6. 6.

    Koga A, Suzuki M, Inagaki H, Bessho Y, Hori H: Transposable element in fish. Nature. 1996, 383: 30-10.1038/383030a0.

    PubMed  CAS  Article  Google Scholar 

  7. 7.

    Frazer MJ, Ciszczon T, Elick T, Bauser C: Precise excision of TTAA-specific lepidopteran transposons piggyBac (IFP2) and tagalong (TFP3) from the baculovirus genome in cell lines from two species of Lepidoptera. Insect Mol Biol. 1996, 5: 141-151.

    Article  Google Scholar 

  8. 8.

    Ivics Z, Hackett PB, Plasterk RH, Izsvák Z: Molecular reconstruction of Sleeping Beauty, a Tc1-like transposon from fish, and its transposition in human cells. Cell. 1997, 91: 501-510. 10.1016/S0092-8674(00)80436-5.

    PubMed  CAS  Article  Google Scholar 

  9. 9.

    Lampe DJ, Akerley BJ, Rubin EJ, Mekalanos JJ, Robertson HM: Hyperactive transposase mutants of the Himar1 mariner transposon. Proc Natl Acad Sci USA. 1999, 96: 11428-11433. 10.1073/pnas.96.20.11428.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  10. 10.

    Kawakami K, Shima A, Kawakami N: Identification of a functional transposase of the Tol2 element, an Ac-like element from the Japanese medaka fish, and its transposition in the zebrafish germ lineage. Proc Natl Acad Sci USA. 2000, 97: 11403-11408. 10.1073/pnas.97.21.11403.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  11. 11.

    Miskey C, Izsvak Z, Plasterk RH, Ivics Z: The Frog Prince: a reconstructed transposon from Rana pipiens with high transpositional activity in vertebrate cells. Nucleic Acids Res. 2003, 31: 6873-6881. 10.1093/nar/gkg910.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  12. 12.

    Sasakura Y, Awazu S, Chiba S, Satoh N: Germ-line transgenesis of the Tc1/mariner superfamily transposon Minos in Ciona intestinalis. Proc Natl Acad Sci USA. 2003, 100: 7726-7730. 10.1073/pnas.1230736100.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  13. 13.

    Sasakura Y: Germline transgenesis and insertional mutagenesis in the ascidian Ciona intestinalis. Dev Dyn. 2007, 236: 1758-1767. 10.1002/dvdy.21111.

    PubMed  CAS  Article  Google Scholar 

  14. 14.

    Sasakura Y, Awazu S, Chiba S, Kano S, Satoh N: Application of Minos, one of the Tc1/mariner superfamily transposable elements, to ascidian embryos as a tool for insertional mutagenesis. Gene. 2003, 308: 11-20. 10.1016/S0378-1119(03)00426-8.

    PubMed  CAS  Article  Google Scholar 

  15. 15.

    van Luenen HG, Colloms SD, Plasterk RH: The mechanism of transposition of Tc3 in C. elegans. Cell. 1996, 79: 293-301. 10.1016/0092-8674(94)90198-8.

    Article  Google Scholar 

  16. 16.

    Awazu S, Sasaki A, Matsuoka T, Satoh N, Sasakura Y: An enhancer trap in the ascidian Ciona intestinalis identifies enhancers of its Musashi orthologous gene. Dev Biol. 2004, 275: 459-472. 10.1016/j.ydbio.2004.08.029.

    PubMed  CAS  Article  Google Scholar 

  17. 17.

    Sasakura Y, Nakashima K, Awazu S, Matsuoka T, Nakayama A, Azuma J, Satoh N: Transposon-mediated insertional mutagenesis revealed the functions of animal cellulose synthase in the ascidian Ciona intestinalis. Proc Natl Acad Sci USA. 2005, 102: 15134-15139. 10.1073/pnas.0503640102.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  18. 18.

    Satoh N: Developmental Biology of Ascidians. 1994, New York: Cambridge University Press

    Google Scholar 

  19. 19.

    Delsuc F, Brinkmann H, Chourrout D, Philippe H: Tunicates and not cephalochordates are the closest living relatives of vertebrates. Nature. 2006, 439: 965-968. 10.1038/nature04336.

    PubMed  CAS  Article  Google Scholar 

  20. 20.

    Bourlat SJ, Juliusdottir T, Lowe CJ, Freeman R, Aronowicz J, Kirschner M, Lander ES, Thorndyke M, Nakano H, Kohn AB, et al: Deuterostome phylogeny reveals monophyletic chordates and the new phylum Xenoturbellida. Nature. 2006, 444: 85-88. 10.1038/nature05241.

    PubMed  CAS  Article  Google Scholar 

  21. 21.

    Nicol D, Meinertzhagen IA: Cell counts and maps in the larval central nervous system of the ascidian Ciona intestinalis (L.). J Comp Neurol. 1991, 309: 415-429. 10.1002/cne.903090402.

    PubMed  CAS  Article  Google Scholar 

  22. 22.

    Nishida H, Satoh N: Cell lineage analysis in ascidian embryos by intracellular injection of a tracer enzyme. I. Up to the eight-cell stage. Dev Biol. 1983, 99: 382-394. 10.1016/0012-1606(83)90288-9.

    PubMed  CAS  Article  Google Scholar 

  23. 23.

    Nishida H, Satoh N: Cell lineage analysis in ascidian embryos by intracellular injection of a tracer enzyme. II. The 16- and 32-cell stages. Dev Biol. 1985, 110: 440-454. 10.1016/0012-1606(85)90102-2.

    PubMed  CAS  Article  Google Scholar 

  24. 24.

    Nishida H: Cell lineage analysis in ascidian embryos by intracellular injection of a tracer enzyme. III. Up to the tissue restricted stage. Dev Biol. 1987, 121: 526-541. 10.1016/0012-1606(87)90188-6.

    PubMed  CAS  Article  Google Scholar 

  25. 25.

    Cloney RA: Ascidian larvae and events of metamorphosis. Amer Zool. 1982, 22: 817-826.

    Article  Google Scholar 

  26. 26.

    Ogasawara M, Wada H, Peters H, Satoh N: Developmental expression of Pax1/9 genes in urochordate and hemichor-date gills: insight into function and evolution of the pharyngeal epithelium. Development. 1999, 126: 2539-2550.

    PubMed  CAS  Google Scholar 

  27. 27.

    Ogasawara M, Di Lauro R, Satoh N: Ascidian homologs of mammalian thyroid peroxidase genes are expressed in the thyroid-equivalent region of the endostyle. J Exp Zool. 1999, 285: 158-169. 10.1002/(SICI)1097-010X(19990815)285:2<158::AID-JEZ8>3.0.CO;2-0.

    PubMed  CAS  Article  Google Scholar 

  28. 28.

    Satoh N: The ascidian tadpole larva: comparative molecular development and genomics. Nat Rev Genet. 2003, 4: 285-295. 10.1038/nrg1042.

    PubMed  CAS  Article  Google Scholar 

  29. 29.

    Satoh N, Satou Y, Davidson B, Levine M: Ciona intestinalis: an emerging model for whole-genome analyses. Trends Genet. 2003, 19: 376-381. 10.1016/S0168-9525(03)00144-6.

    PubMed  CAS  Article  Google Scholar 

  30. 30.

    Shi W, Levine M, Davidson B: Unraveling genomic regulatory networks in the simple chordate, Ciona intestinalis. Genome Res. 2005, 15: 1668-1674. 10.1101/gr.3768905.

    PubMed  CAS  Article  Google Scholar 

  31. 31.

    Satoh N, Levine M: Surfing with the tunicates into the post-genome era. Genes Dev. 2005, 19: 2407-2411. 10.1101/gad.1365805.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  32. 32.

    Moody R, Davis SW, Cubes F, Smith WC: Isolation of developmental mutants of the ascidian Ciona savignyi. Mol Gen Genet. 1999, 262: 199-206. 10.1007/s004380051075.

    PubMed  CAS  Article  Google Scholar 

  33. 33.

    Nakatani Y, Moody R, Smith WC: Mutations affecting tail and notochord development in the ascidian Ciona savignyi. Development. 1999, 126: 3293-3301.

    PubMed  CAS  Google Scholar 

  34. 34.

    Hendrickson C, Christiaen L, Deschet K, Jiang D, Joly JS, Legendre L, Nakatani Y, Tresser J, Smith WC: Culture of adult ascidians and ascidian genetics. Methods Cell Biol. 2004, 74: 143-170.

    PubMed  CAS  Article  Google Scholar 

  35. 35.

    Dehal P, Satou Y, Campbell RK, Chapman J, Degnan B, De Tomaso A, Davidson B, Di Gregorio A, Gelpke M, Goodstein DM, et al: The draft genome of Ciona intestinalis: Insight into chordate and vertebrate origins. Science. 2002, 298: 2157-2167. 10.1126/science.1080049.

    PubMed  CAS  Article  Google Scholar 

  36. 36.

    Simmen MW, Leitgeb S, Clark VH, Jones SJ, Bird A: Gene number in an invertebrate chordate, Ciona intestinalis. Proc Natl Acad Sci USA. 1998, 95: 4437-4440. 10.1073/pnas.95.8.4437.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  37. 37.

    Satou Y, Satoh N: Genomewide surveys of developmentally relevant genes in Ciona intestinalis. Dev Genes Evol. 2003, 213: 211-212. 10.1007/s00427-003-0330-z.

    PubMed  Article  Google Scholar 

  38. 38.

    Sasakura Y, Shoguchi E, Takatori N, Wada S, Meinertzhagen IA, Satou Y, Satoh N: A genomewide surveys of developmentally relevant genes in Ciona intestinalis X. Genes for cell junctions and extracellular matrix. Dev Genes Evol. 2003, 213: 303-313. 10.1007/s00427-003-0320-1.

    PubMed  CAS  Article  Google Scholar 

  39. 39.

    Donoghue PCJ, Purnell MA: Genome duplication, extinction and vertebrate evolution. Trends Ecol Evol. 2005, 20: 312-319. 10.1016/j.tree.2005.04.008.

    PubMed  Article  Google Scholar 

  40. 40.

    Plasterk RH, Izsvak Z, Ivics Z: Resident aliens: the Tc1/mariner superfamily of transposable elements. Trends Genet. 1999, 15: 326-332. 10.1016/S0168-9525(99)01777-1.

    PubMed  CAS  Article  Google Scholar 

  41. 41.

    Corbo JC, Levine M, Zeller RW: Characterization of a notochord-specific enhancer from the Brachyury promoter region of the ascidian, Ciona intestinalis. Development. 1997, 124: 589-602.

    PubMed  CAS  Google Scholar 

  42. 42.

    Satou Y, Imai K, Satoh N: Action of morpholinos in Ciona embryos. Genesis. 2001, 30: 103-106. 10.1002/gene.1040.

    PubMed  CAS  Article  Google Scholar 

  43. 43.

    Joly JS, Kano S, Matsuoka T, Auger H, Hirayama K, Satoh N, Awazu S, Legendre L, Sasakura Y: Culture of Ciona intestinalis in closed systems. Dev Dyn. 2007, 236: spc1-10.1002/dvdy.21250.

    Article  Google Scholar 

  44. 44.

    Jiang D, Munro EM, Smith WC: Ascidian prickle regulates both mediolateral and anterior-posterior cell polarity of notochord cells. Curr Biol. 2005, 15: 79-85. 10.1016/j.cub.2004.12.041.

    PubMed  CAS  Article  Google Scholar 

  45. 45.

    Jiang D, Tresser JW, Horie T, Tsuda M, Smith WC: Pigmentation in the sensory organs of the ascidian larva is essential for normal behavior. J Exp Biol. 2005, 208: 433-438. 10.1242/jeb.01420.

    PubMed  Article  Google Scholar 

  46. 46.

    Loukeris TG, Livadaras I, Arca B, Zabulou S, Savakis C: Gene transfer into the Medfly, Ceratitis capitata, with a Drosophila hydei transposable element. Science. 1995, 270: 2002-2005. 10.1126/science.270.5244.2002.

    PubMed  CAS  Article  Google Scholar 

  47. 47.

    Loukeris TG, Arca B, Livadaras I, Dialektaki G, Savakis C: Introduction of the transposable element Minos into the germ line of Drosophila melanogaster. Proc Natl Acad Sci USA. 1995, 92: 9485-9489. 10.1073/pnas.92.21.9485.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  48. 48.

    Klinakis AG, Zagoraiou L, Vassilatis DK, Savakis C: Genome-wide insertional mutagenesis in human cells by the Drosophila mobile element Minos. EMBO Rep. 2000, 1: 416-421. 10.1093/embo-reports/kvd089.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  49. 49.

    Klinakis AG, Loukeris TG, Pavlopoulos A, Savakis C: Mobility assays confirm the broad host-range activity of the Minos transposable element and validate new transformation tools. Insect Mol Biol. 2000, 9: 269-275. 10.1046/j.1365-2583.2000.00183.x.

    PubMed  CAS  Article  Google Scholar 

  50. 50.

    Shimizu K, Kamba M, Sonobe H, Kanda T, Klinakis AG, Savakis C, Tamura T: Extrachromosomal transposition of the transposable element Minos in embryos of the silkworm Bombyx mori. Insect Mol Biol. 2000, 9: 277-281. 10.1046/j.1365-2583.2000.00182.x.

    PubMed  CAS  Article  Google Scholar 

  51. 51.

    Zhang H, Shinmyo Y, Hirose A, Inoue Y, Ohuchi H, Loukeris TG, Eggleston P, Noji S: Extrachromosomal transposition of the transposable element Minos in embryos of the cricket Gryllus bimaculatus. Dev Growth Differ. 2002, 44: 409-417. 10.1046/j.1440-169X.2002.00654.x.

    PubMed  CAS  Article  Google Scholar 

  52. 52.

    Drabek D, Zagoraiou L, deWit T, Langeveld A, Roumpaki C, Mamalaki C, Savakis C, Grosveld F: Transposition of the Drosophila hydei Minos transposon in the mouse germ line. Genomics. 2003, 81: 108-111. 10.1016/S0888-7543(02)00030-7.

    PubMed  CAS  Article  Google Scholar 

  53. 53.

    Pavlopoulos A, Averof M: Establishing genetic transformation for comparative developmental studies in the crustacean Parhyale hawaiensis. Proc Natl Acad Sci USA. 2005, 102: 7888-7893. 10.1073/pnas.0501101102.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  54. 54.

    Metaxakis A, Oehler S, Klinakis A, Savakis C: Minos as a genetic and genomic tool in Drosophila melanogaster. Genetics. 2005, 171: 571-581. 10.1534/genetics.105.041848.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  55. 55.

    Matsuoka T, Awazu S, Satoh N, Sasakura Y: Minos transposon causes germline transgenesis of the ascidian Ciona savignyi. Dev Growth Differ. 2004, 46: 249-255. 10.1111/j.1440-169X.2004.00742.x.

    PubMed  CAS  Article  Google Scholar 

  56. 56.

    Matsuoka T, Awazu S, Shoguchi E, Satoh N, Sasakura Y: Germline transgenesis of the ascidian Ciona intestinalis by electroporation. Genesis. 2005, 41: 61-72. 10.1002/gene.20096.

    Article  Google Scholar 

  57. 57.

    Arca B, Zabalou S, Loukeris T, Savakis C: Mobilization of a Minos transposon in Drosophila melanogaster chromosomes and chromatid repair by heteroduplex formation. Genetics. 1997, 145: 267-279.

    PubMed  CAS  PubMed Central  Google Scholar 

  58. 58.

    Davidson AE, Balciunas D, Mohn D, Shaffer J, Hermanson S, Sivasubbu S, Cliff MP, Hackett PB, Ekker SC: Efficient gene delivery and gene expression in zebrafish using the Sleeping Beauty transposon. Dev Biol. 2003, 263: 191-202. 10.1016/j.ydbio.2003.07.013.

    PubMed  CAS  Article  Google Scholar 

  59. 59.

    Liu YG, Mitsukawa N, Oosumi T, Whittier RF: Efficient isolation and mapping of Arabidopsis thaliana T-DNA insert junctions by thermal asymmetric interlaced PCR. Plant J. 1995, 8: 457-463. 10.1046/j.1365-313X.1995.08030457.x.

    PubMed  CAS  Article  Google Scholar 

  60. 60.

    Karsi A, Moav B, Hackett P, Liu Z: Effects of insert size on transposition efficiency of the Sleeping Beauty transposon in mouse cells. Mar Biotechnol. 2001, 3: 241-245. 10.1007/s101260000072.

    PubMed  CAS  Article  Google Scholar 

  61. 61.

    Fadool JM, Hartl DL, Dowling JE: Transposition of the mariner element from Drosophila mauritiana in zebrafish. Proc Natl Acad Sci USA. 1998, 95: 5182-5186. 10.1073/pnas.95.9.5182.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  62. 62.

    Bessereau JL, Wright A, Williams DC, Schuske K, Davis MW, Jorgensen EM: Mobilization of a Drosophila transposon in the Caenorhabditis elegans germ line. Nature. 2001, 413: 70-74. 10.1038/35092567.

    PubMed  CAS  Article  Google Scholar 

  63. 63.

    Fischer SE, Wienholds E, Plasterk RH: Regulated transposition of a fish transposon in the mouse germ line. Proc Natl Acad Sci USA. 2001, 98: 6759-6764. 10.1073/pnas.121569298.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  64. 64.

    Horie K, Kuroiwa A, Ikawa M, Okabe M, Kondoh G, Matsuda Y, Takeda J: Efficient chromosomal transposition of a Tc1/mariner-like transposon Sleeping Beauty in mice. Proc Natl Acad Sci USA. 2001, 98: 9191-9196. 10.1073/pnas.161071798.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  65. 65.

    Zayed H, Izsvák Z, Khare D, Heinemann U, Ivics Z: The DNA-bending protein HMGB1 is a cellular confactor of Sleeping Beauty transposition. Nucleic Acids Res. 2003, 31: 2313-2322. 10.1093/nar/gkg341.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  66. 66.

    Walisko O, Izsvák Z, Szabo K, Kaufman CD, Herold S, Ivics Z: Sleeping Beauty transposase modulates cell-cycle progression through interaction with Miz-1. Proc Natl Acad Sci USA. 2006, 103: 4062-4067. 10.1073/pnas.0507683103.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  67. 67.

    Cruz NB, Weinreich MD, Wiegand TW, Krebs MP, Reznikoff WS: Characterization of the Tn5 transposase and inhibitor proteins: a model for the inhibition of transposition. J Bact. 1993, 175: 6932-6938.

    PubMed  PubMed Central  Google Scholar 

  68. 68.

    Corbo JC, Di Gregorio A, Levine M: The ascidian as a model organism in developmental and evolutionary biology. Cell. 2001, 106: 535-538. 10.1016/S0092-8674(01)00481-0.

    PubMed  CAS  Article  Google Scholar 

  69. 69.

    Keys DN, Lee BI, Di Gregorio A, Harafuji N, Detter JC, Wang M, Kahsai O, Ahn S, Zhang C, Doyle SA, et al: A saturation screen for cis-acting regulatory DNA in the Hox genes of Ciona intestinalis. Proc Natl Acad Sci USA. 2005, 102: 679-683. 10.1073/pnas.0408952102.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  70. 70.

    Awazu S, Matsuoka T, Satoh N, Inaba K, Sasakura Y: High-throughput enhancer trap by remobilization of transposon Minos in Ciona intestinalis. Genesis. 2007, 45: 307-317. 10.1002/dvg.20290.

    PubMed  CAS  Article  Google Scholar 

  71. 71.

    Balciunas D, Davidson AE, Sivasubbu S, Hermanson SB, Welle Z, Ekker SC: Enhancer trapping in zebrafish using the Sleeping Beauty transposon. BMC Genom. 2004, 5: 62-10.1186/1471-2164-5-62.

    Article  Google Scholar 

  72. 72.

    Parinov S, Kondrichin I, Korzh V, Emelyanov A: Tol2 transposon-mediated enhancer trap to identify developmentally regulated zebrafish genes in vivo. Dev Dyn. 2004, 231: 449-459. 10.1002/dvdy.20157.

    PubMed  CAS  Article  Google Scholar 

  73. 73.

    Dupuy AJ, Fritz S, Largaespada DA: Transposition and gene disruption in the male germline of the mouse. Genesis. 2001, 30: 82-88. 10.1002/gene.1037.

    PubMed  CAS  Article  Google Scholar 

  74. 74.

    Shirae-Kurabayashi M, Nishikata T, Takamura K, Tanaka KJ, Nakamoto C, Nakamura A: Dynamic redistribution of vasa homolog and exclusion of somatic cell determinants during germ cell specification in Ciona intetinalis. Development. 2006, 133: 2683-2693. 10.1242/dev.02446.

    PubMed  CAS  Article  Google Scholar 

  75. 75.

    Robertson HM, Preston CR, Phillis RW, Johnson-Schlitz DM, Benz WK, Engels WR: A stable genomic source of P element transposase in Drosophila melanogaster. Genetics. 1988, 118: 461-470.

    PubMed  CAS  PubMed Central  Google Scholar 

  76. 76.

    Amsterdam A, Burgess S, Golling G, Che W, Sun Z, Townsend K, Farrington S, Haldi M, Hopkins N: A large-scale insertional mutagenesis screen in zebrafish. Genes Dev. 1999, 13: 2713-2724. 10.1101/gad.13.20.2713.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  77. 77.

    Amsterdam A: Insertional mutagenesis in zebrafish. Dev Dyn. 2003, 228: 523-534. 10.1002/dvdy.10381.

    PubMed  CAS  Article  Google Scholar 

  78. 78.

    Chen W, Burgess S, Golling G, Amsterdam A, Hopkins N: High-throughput selection of retrovirus producer cell lines leads to markedly improved efficiency of germ line-transmissible insertions in zebra fish. J Virol. 2002, 76: 2192-2198. 10.1128/jvi.76.5.2192-2198.2002.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  79. 79.

    Nakashima K, Yamada L, Satou Y, Azuma J, Satoh N: The evolutionary origin of animal cellulose synthase. Dev Genes Evol. 2004, 214: 81-88. 10.1007/s00427-003-0379-8.

    PubMed  CAS  Article  Google Scholar 

  80. 80.

    Sordino P, Heisenberg CP, Cirino P, Toscano A, Giuliano P, Marino R, Pinto MR, De Santis R: A mutational approach to the study of development of the protochordate Ciona intestinalis (Tunicata, Chordata). Sarsia. 2000, 85: 173-176.

    Article  Google Scholar 

  81. 81.

    Sordino P, Belluzzi L, De Santis R, Smith WC: Developmental genetics in primitive chordates. Philos Trans R Soc Lond B Biol Sci. 2001, 356: 1573-1582. 10.1098/rstb.2001.0919.

    PubMed  CAS  PubMed Central  Article  Google Scholar 

  82. 82.

    Eukaryotic Genomics. []

Download references


The authors thank Kazuo Inaba, Yoshikazu Okada, Yasuyo Kasuga, Kazuko Hirayama, Aru Konno, Katsutoshi Mizuno, Yasutaka Tsuchiya, Toshihiko Sato, Hideo Shinagawa, Yoshiko Harada, and members of the Shimoda Marine Research Center, University of Tsukuba, for their kind cooperation with our study. Erik Jorgensen, Zoltán Ivics, Zsuzsanna Izsvák, and Charalambos Savakis are acknowledged for their kind provision of Mos1, SB, FP, and Minos. We thank Koichi Kawakami, Hiroshi Wada, Takeshi Kawashima, Nik Putnam, and Patrick Lemaire for useful discussions. This study was supported by Grants-in-Aid for Scientific Research from JSPS and MEXT to YS and NS. This research was also supported by JST, Japan to NS (CREST project). YS was supported by NIG Cooperative Research Program (2006-A72).

This article has been published as part of Genome Biology Volume 8, Supplement 1, 2007: Transposons in vertebrate functional genomics. The full contents of the supplement are available online at

Author information



Corresponding author

Correspondence to Yasunori Sasakura.

Additional information

Competing interests

The authors declare that they have no competing interests.

Rights and permissions

Reprints and Permissions

About this article

Cite this article

Sasakura, Y., Oogai, Y., Matsuoka, T. et al. Transposon mediated transgenesis in a marine invertebrate chordate: Ciona intestinalis. Genome Biol 8, S3 (2007).

Download citation


  • Insertional Mutant
  • Insertional Mutagenesis
  • Sleep Beauty
  • Saturation Mutagenesis
  • Stable Transgenic Line