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Keywords:

  • tunicates;
  • transposon;
  • Tc1;
  • mariner;
  • remobilization

Abstract

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

Transgenesis with transposons is an important technique for studying genetic functions. In the ascidian Ciona intestinalis, methods for germline transformation with the Tc1/mariner transposon Minos have been established. A system to remobilize a single Minos copy in the genome is needed to refine this transgenic technique. In this study, such an experimental system was established with a transgenic line expressing Minos transposase in eggs. In the eggs of a double transgenic animal from a cross between the egg transposase line and a transgenic line having a single Minos insertion, the transposon was transposed into new positions of the Ciona genome, thus creating new insertions. Some of the new insertions caused enhancer detection. The majority of the new insertion sites were mapped on different chromosomes from that of the transposon donor. This characteristic of Minos is in contrast to that of the Sleeping Beauty transposon, which causes frequent intrachromosomal transposition. Developmental Dynamics 239:1076–1088, 2010. © 2010 Wiley-Liss, Inc.


INTRODUCTION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

Transgenic technologies with DNA transposons are extremely useful for studying genetic functions in animals (Miskey et al., 2005). Active transposons can be used as transformation vectors that carry the exogenous, artificial DNA into the genomes. For example, a fusion of a tissue-specific promoter and a reporter gene can be introduced into genomes with transposons to create stable transgenic “marker” lines that express reporter genes in a tissue-specific manner. A transposon vector with a minimal promoter and a reporter gene can be used to entrap endogenous enhancers, and such “enhancer detection” technology is another useful way to produce marker transgenic lines as well as to identify enhancers. Transposons are sometimes inserted into the critical regions of genes to disrupt their functions. The insertional mutagenesis is a reliable way to investigate gene functions, because the mutated sites can be identified in a short period by using the transposon sequence as a tag. With these applications, transformation technologies with transposons are powerful ways to perform genetic analyses in various model organisms in which genetic approaches have been established.

The ascidian Ciona intestinalis (hereafter referred to as “Ciona”) is a useful material for genetic study. Its life cycle is relatively short, approximately 2–3 months, and inland Ciona culture systems have been established (Joly et al., 2007). A draft genome sequence of Ciona that allows us to know the sequences of almost all genes of this organism has been determined (Dehal et al., 2002). In Ciona, the Tc1/mariner superfamily transposon Minos has both excision and integration activity (Sasakura et al., 2003a). Germline transformation of Ciona has been achieved with Minos, and approximately 30% of Minos-introduced Ciona become founders that transmit transposon insertions to subsequent generations (Sasakura et al., 2003b). With Minos, genetic techniques such as the creation of stable transgenic lines, enhancer detection and insertional mutagenesis have been introduced in Ciona (Awazu et al., 2004; Sasakura et al., 2005).

An important factor in transposon-mediated transgenesis is the efficiency with which new insertions are created. There are several methods to create new insertions of Minos in Ciona (Sasakura, 2007), the simplest of which is the simultaneous introduction of a Minos vector and transposase mRNA into eggs or embryos by microinjection or electroporation (Sasakura et al., 2003b; Matsuoka et al., 2005). Although this method can easily be carried out soon after the vector is prepared, it has several disadvantages. For example, microinjection is a difficult technique to acquire, and electroporation-mediated transformation is costly because it requires a large amount of in vitro-transcribed transposase mRNA. Sometimes Ciona shows abnormal development following the introduction of these components, which results in death or slow growth. Taking enhancer detection into consideration, the efficiency of creating enhancer detection lines by these microinjection- or electroporation-mediated transformation methods is low. For these reasons, another system to create new insertions was desired.

We recently established a transgenic line expressing transposase in the male germ cells (Sasakura et al., 2008). This “transposase line” provides us with a useful way to create new transposon insertions by crossing the line with a transposon donor line that has transposon insertions in its genome. Transposon and transposase meet in the male germ cells of a double transgenic animal produced from a cross between the two lines, and transposase can cut and paste transposons in the chromosomes of the cells. Progeny derived from such germ cells inherit new insertion sites that are different from those in their parents, and thus new transgenic lines can be established. An advantage of this method is that time-consuming experiments are not necessary; the crossing procedures are the only required steps for this method. By using the transposase line, efficient enhancer detection was achieved in Ciona (Sasakura et al., 2008).

In our earlier study (Sasakura et al., 2008), we used a transgenic line with a multiple, tandem-arrayed transposon insertion as a transposon donor. Having many transposon copies increases the chances for transposase to cut and paste transposons, which should result in a greater likelihood of creating new insertions. However, multiple transposon copies sometimes make further analyses difficult. First, the original tandem transposon array usually interferes with polymerase chain reaction (PCR) analyses to identify transposon insertion sites. Therefore, the tandem-arrayed insertion has to be separated from the insertion sites-of-interest by segregation analysis. Second, any progeny created with this system frequently has several independent insertions. Even though a transposon insertion site is determined from a progeny having several insertion sites, further time-consuming experiments are necessary to investigate whether the identified insertion is responsible for the mutation or enhancer detection. To simplify the analysis, it is better to remobilize a single transposon copy (Urasaki et al., 2008). We tried to remobilize a single transposon copy by the microinjection of transposase into transposon donor lines or by using transposase lines of male germ cells. Although we detected transposon excision in the somatic cells by the microinjection of transposase (Sasakura et al., 2007), we so far have not detected any remobilization event of a single transposon copy in the germ cells. A new system is necessary to achieve remobilization of a single transposon copy.

One possible way to accomplish the remobilization of a single transposon copy is to use a transposase line expressing transposase in eggs, and we have created such a line in the present study. Using the egg transposase line, the remobilization of a single transposon copy and enhancer detection were achieved. With this method, we investigated the tendency of Minos to remobilize in the Ciona genome. Minos showed frequent interchromosomal transposition.

RESULTS

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

Creation of a Transposase Line Expressing Transposase in Eggs

To express transposase in eggs, we used the cis element of Ci-Nut. This gene shows both maternal and zygotic expression, and the zygotic expression is observed in the neural tissues (Kitaura et al., 2007). The vector pSBFr3dTPORCiNutMiTP was created (Fig. 1A). This vector has a Ci-Nut promoter and the cDNA of a Minos transposase fusion cassette, a marker cassette that expresses red fluorescent protein (RFP) in the somatic cells at the juvenile stage, and inverted repeats of a Sleeping Beauty (SB) transposon (SB10 in Ivics et al., 1997). Because the SB transposon has only weak activity in Ciona (Sasakura et al., 2007), a transgenic line having an insertion of this vector was created by DNA electroporation (Matsuoka et al., 2005), which can create transgenic lines by nonhomologous recombination. This transgenic line (named Ju[SBFr3dTPORCiNutMiTP]1; abbreviated Ju[CiNutMiTP]1 in this manuscript) expressed RFP in the endostyle, retropharyngeal band, peripharyngeal band, and muscle cells at the juvenile stage (Fig. 1B). This RFP expression was used as a selective marker for this line. Expression of Minos transposase mRNA in the eggs of Ju[CiNutMiTP]1 was detected by reverse transcription (RT)-PCR (Fig. 1C). Eggs of this transgenic line were fertilized normally with wild-type sperm. The embryos developed to be normal larvae, and subsequent development after metamorphosis was also normal until the reproductive stage (data not shown). These results suggest that the expression of Minos transposase in eggs does not have major effects on embryogenesis or development.

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Figure 1. A transgenic line expressing Minos transposase in eggs. A: The structure of pSBFr3dTPORCiNutMiTP. Inverted terminal repeats (ITRs) of Sleeping Beauty (SB) are shown by black triangles. pBluescript backbone is not included in this diagram. NLS, nuclear localization signal; Ter, SV40 transcription termination sequence; MiTP, Minos transposase cDNA. A Fr3 enhancer from an intron of Ci-musashi (Awazu et al., 2004) is included in this construct to enhance expression of DsRed from the Ci-TPO promoter. B: A juvenile of Ju[CiNutMiTP]1. Left: Red fluorescent protein (RFP) expression. RFP is expressed in the endostyle (En), peripharyngeal band (PB), retropharyngeal band (RB), body wall muscle (BM), and oral siphon muscle (OS). Right: The bright field image. C: Expression of Minos transposase in the eggs of Ju[CiNutMiTP]1. Total RNA isolated from the eggs of wild-type animals (wt) and Ju[CiNutMiTP]1 (NutMiTP) was subjected to reverse transcriptase-polymerase chain reaction (RT-PCR) analysis. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as the positive control. RT, lanes are negative controls without reverse transcription. M, marker lane. MiTP, Minos transposase.

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To determine the activity of Minos transposase in eggs of Ju[CiNutMiTP]1, we first examined whether the transposase can catalyze excision of Minos from the plasmid DNA. A Minos vector, pMiCiAKRK, was created (Fig. 2A). This vector includes a cis regulatory element of Ci-AKR (Tokuoka et al., 2004) for driving the Kaede reporter gene (Ando et al., 2002) in the mesenchyme of embryos and larvae. This plasmid vector was introduced into the eggs of Ju[CiNutMiTP]1. The embryos were cultured until the late tailbud stage, then PCR was performed using DNA isolated from these embryos (Fig. 2B). If excision occurs in Ciona embryos, the plasmids become shorter as a result of the removal of transposons, and PCR bands of approximately 169 bp are amplified. In the present study, such PCR bands were obtained from embryos derived from eggs of Ju[CiNutMiTP]1 into which pMiCiAKRK had been introduced (Fig. 2C, arrow), while no such bands were obtained from pMiCiAKRK-introduced wild-type embryos. The DNA sequences of the bands suggesting excision were determined. The bands contained a footprint sequence characteristic of excision events caused by transposase (Fig. 2D). A footprint sequence is a specific DNA sequence created during and after the excision of transposons (van Luenen et al., 1994). Therefore, we concluded that the Minos transposase expressed in Ju[CiNutMiTP]1 has excision activity.

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Figure 2. Minos transposase is active in eggs of Ju[CiNutMiTP]1. A: A Minos vector pMiCiAKRK used in this assay. Inverted terminal repeats (ITRs) of Minos are shown by black triangles. pBluescript backbone is not included in this diagram. Ter, SV40 transcription termination sequence. B: A diagram showing the excision assay procedure. pMiCiAKRK was introduced into 1-cell embryos of Ju[CiNutMiTP]1 by electroporation, and these DNA-introduced embryos were cultured for overnight. At the late tailbud stage, the plasmid DNA was extracted from the embryos and was subjected to polymerase chain reaction (PCR) analysis with primers. The primer sites are shown by small arrows. If excision occurs, the plasmid becomes shorter and bands of approximately 169 bp are amplified. C: The result of excision assay. Lanes 1–8: the PCR products from wild-type embryos into which pMiCiAKRK was electroporated. Lanes 9–16: the PCR products from embryos of Ju[CiNutMiTP]1 into which pMiCiAKRK was electroporated. Bands of 169 bp (arrow) were amplified in lanes 10, 11, 13–16. Some DNAs were subjected to rearrangement in the Ciona embryos, and PCR bands derived from such rearranged DNAs were also amplified in each lane. D: Footprint sequences of Minos excision, obtained from the PCR bands in C. The upper two footprints are typical footprint sequences of Minos, and the bottom one is an atypical sequence. E: Expression of Kaede in a transgenic line of pMiCiAKRK at the larval stage. This line was created by electroporation of pMiCiAKRK in one-cell embryos of Ju[CiNutMiTP]1. Kaede fluorescence is observed in the mesenchyme (Me).

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The animals developed from pMiCiAKRK-electroporated eggs of Ju[CiNutMiTP]1were cultured until the reproductive stage, and they were crossed with wild-types to screen transgenic lines expressing Kaede in the mesenchyme. Twenty-one animals were screened, and one animal became a founder. A transgenic line, Tg[MiCiAKRK]1, was established from this founder (Fig. 2E). The insertion site of Minos in this transgenic line was determined by thermal asymmetric interlaced (TAIL)-PCR (Liu et al., 1995). Two insertion sites were identified from the transgenic line (Table 1). Minos was inserted into the TA dinucleotides, and the targeted TA dinucleotides were duplicated by the integration of Minos. These insertion characteristics coincided with the reported characteristics of Tc1/mariner superfamily transposons (van Luenen et al., 1994; Arca et al., 1997), suggesting the transposition of Minos from plasmid DNA to genomes of Ciona. However, the efficiency of these insertions was lower than that of Ciona transformation by the simultaneous introduction of a Minos vector and transposase mRNA (Sasakura et al., 2003b; Matsuoka et al., 2005).

Table 1. Sequences of the Insertion Sites of Tg[MiCiAKRK]1
Sequence of the insertion siteaScaffold number in the JGI genome browserNearest gene modelbPosition in the geneChromosome
TATTGTCTGTTTGATCAGCTGCGTTCAATCAGTCGCCTGAAATTCGACCA102 in the version 1 browserKH.C9.476Intron9
TACATTTAAGCCTTGCCTGTAAACG CTAAATTTAGTTAAGCCCACGTTTA316 in the version 2 browserNo gene model  

Remobilization of a Single Minos Insertion in the Genome of Ciona With Ju[CiNutMiTP]1

To analyze the remobilization of a single Minos copy in the Ciona genome, we selected a green fluorescent protein (GFP) -enhancer detection line expressing GFP in the epidermis at the ampulla (E[MiTSAdTPOG]18) as a donor for transposon insertion (Fig. 3A; Awazu et al., 2007). E[MiTSAdTPOG]18 has a single transposon insertion in the upstream region of Ci-meta2 (Fig. 6B). An enhancer detection line was chosen as a transposon donor because the reporter gene (gfp) expression pattern of an enhancer detection line is dependent on its insertion site. Progeny having remobilized transposon insertions will show different GFP expression patterns from the original one, and these can be recognized by observing the GFP expression patterns under fluorescent microscopy. Our enhancer detection vector, pMiTSAdTPOG, contains a promoter of Ci-TPO (Ogasawara et al., 1999). This vector shows constitutive GFP expression at the ends of the endostyle (Sasakura et al., 2003b), which helps with the screening of progeny having remobilized transposon insertions, even though the new insertions do not cause enhancer detection. E[MiTSAdTPOG]18 was crossed with Ju[CiNutMiTP]1 to obtain double transgenic animals (hereafter the term “double transgenic animal” refers to animals having this genotype; Fig. 3A–C). The double transgenic animals were cultured until the reproductive stage, and eggs from the animals were fertilized with sperm from wild-types to generate progeny. The GFP expression of the progeny was observed to determine whether remobilization of Minos occurred.

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Figure 3. Green fluorescent protein (GFP) and red fluorescent protein (RFP) expression of transposon donors, Ju[CiNutMiTP]1 and progeny from eggs of double transgenic animals between them. A: GFP expression of E[MiTSAdTPOG]18 at the juvenile stage. GFP fluorescence is seen at the ampulla (Am). B: A juvenile of Ju[CiNutMiTP]1. C: A double transgenic animal of E[MiTSAdTPOG]18 and Ju[CiNutMiTP]1. D: A progeny derived from an egg of a double transgenic animal between E[MiTSAdTPOG]18 and Ju[CiNutMiTP]1. GFP fluorescence is observed at the end of the endostyle (arrow). This GFP expression is the constitutive expression from Ci-TPO promoter and named “TPO pattern” in this manuscript. E,F: Two examples of progeny with enhancer detection patterns of GFP expression, derived from an egg of a double transgenic animal between E[MiTSAdTPOG]18 and Ju[CiNutMiTP]1. E: GFP is expressed in the pharyngeal gill (Gi) and oral siphon (OS). F: GFP is expressed in the endostyle (En) and blood cells (Bl). G: A juvenile of Tg[MiCiTnIG]2. GFP is expressed in muscle cells. H,I: Two examples of progeny with enhancer detection patterns of GFP expression, derived from an egg of a double transgenic animal between Tg[MiCiTnIG]2 and Ju[CiNutMiTP]1. H: GFP is expressed in the whole body. I: GFP is expressed in the endostyle in addition to the muscle tissues. Scale bars = 100 μm in A, 100 μm in B–I.

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As shown in Table 2, in total only 33.2% of progeny from the eggs of double transgenic animals showed the GFP expression pattern of E[MiTSAdTPOG]18, which is a lower frequency than that expected from Mendel's law regarding a single heterozygous locus. If no remobilization occurred, approximately 50% of the progeny should show the E[MiTSAdTPOG]18 GFP expression pattern (Table 3). The frequency of RFP expression in these progeny, which is the marker of the transposase locus and thus independent of the gfp locus, was in accordance with Mendel's law. These data suggest that the insertion derived from E[MiTSAdTPOG]18 was lost in the eggs of double transgenic animals. The loss of the Minos insertion may have been caused by excision of the transposon. In addition to this observed phenomenon, 3.8% of the progeny from eggs of double transgenic animals showed GFP expression at the ends of the endostyle similar to the GFP expression pattern of pMiTSAdTPOG (the TPO pattern in Table 2 and Fig. 3D). Furthermore, 0.6% of the progeny showed enhancer detection of GFP expression (Table 2; Fig. 3E,F). These results suggest that the Minos insertion of E[MiTSAdTPOG]18 was cut from the genome and that some of the insertions were integrated again in different positions in the genome to create new insertions during oogenesis of the double transgenic animals. As a negative control, sperm of double transgenic lines were used to fertilize wild-type eggs to obtain progeny, and the GFP expression of these progeny at the juvenile stage was observed. E[MiTSAdTPOG]18-patterned GFP expression was in accordance with Mendel's law (Table 3). In addition, no progeny showed GFP expression patterns different from E[MiTSAdTPOG]18 in the control family (Table 3).

Table 2. GFP Expression Patterns of the Egg-Derived Progeny of the Double Transgenic Animals Between E[MiTSAdTPOG]18 and Ju[CiNutMiTP]1a
ID of the double transgenic animalGFP expression patternRFP
Original patternTPO patternbEnhancer detectionGFP negative
  • a

    GFP, green fluorescent protein; RFP, red fluorescent protein.

  • b

    The constitutive GFP expression pattern of pMiTSAdTPOG

  • c

    Not examined.

1555/14872/207 (1%)4/1491 (0.3%)932/1487NEc
1 (31 days later)67/1782/121 (1.7%)1/178 (0.6%)NE57/116
2165/4034/120 (3.3%)4/407 (1%)238/403NE
2 (21 days later)13/531/53 (1.9%)0/53 (0%)39/5337/50
365/26415/264 (5.7%)3/264 (1.1%)181/26463/131
3 (28 days later)19/1187/118 (6%)2/118 (1.7%)90/11866/123
473/2899/289 (3.1%)1/289 (0.3%)206/28988/152
511/1159/115 (7.8%)3/115 (2.6%)92/11565/121
Total968/2907 (33.2%)49/1287 (3.8%)18/2915 (0.61%)1778/2729 (65.1%)376/693 (54.2%)
Table 3. GFP Expression Patterns of the Sperm-Derived Progeny of the Double Transgenic Animals Between E[MiTSAdTPOG]18 and Ju[CiNutMiTP]1
ID of the double transgenic animalGFP expression patternRFP
Original patternGFP negativeTPO patternEnhancer detection
  1. aGFP, green fluorescent protein; RFP, red fluorescent protein.

1289/552263/5520/2210/221 
1 (31 days later)179/344165/3440/3440/34459/111
2131/306175/3060/3060/306 
2 (21 days later)149/286137/2860/2860/28673/149
3188/359171/3590/3590/35981/161
3 (28 days later)80/16686/1660/1660/16666/119
4127/252125/2520/2520/25273/142
566/13771/1370/1370/13766/140
Total1209/2402 (50.3%)1193/2402 (49.7%)00418/822 (50.8%)

The frequency of the progeny showing the remobilized GFP expression pattern (the TPO pattern and enhancer detection pattern in Table 2) was compared among double transgenic animals. The frequency of the progeny showing the TPO pattern varied from 1% to 7.8%, and enhancer detection varied from 0% to 2.6%. There is a tendency for the family showing a higher frequency of progeny with the TPO pattern to also show a higher frequency of progeny with the enhancer detection pattern. The different frequencies among double transgenic animals suggest that the frequency of the remobilization of Minos differs among double transgenic animals. To investigate whether the age of the double transgenic animals and the timing of gamete isolation affect the remobilization frequency, we isolated two generations of progeny from double transgenic animals and compared the frequency of GFP-expressing juveniles between the families derived from the same double transgenic animals. No major difference was observed (Table 2), suggesting that the age of double transgenic animals and the timing of the isolation of progeny do not affect the remobilization frequency.

Data suggesting excision and remobilization of Minos single insertions were obtained when another transgenic line, the muscle-specific marker line Tg[MiCiTnIG]2, was used as the transposon donor (Table 4; Fig. 3G). The transposon vector for Tg[MiCiTnIG]2 contains an upstream sequence of the muscle-specific gene Ci-TnI, which shows constitutive expression of GFP in the muscle tissues. A few progeny derived from the eggs of a double transgenic line between Tg[MiCiTnIG]2 and the egg transposase line showed GFP expression in other tissues in addition to the muscles (Fig. 3G–I), suggesting that the transposon vector was remobilized to the genomic sites under the effect of strong enhancers. These results suggest that remobilization of a single transposon copy by the transposase line Ju[CiNutMiTP]1 is not specific to the E[MiTSAdTPOG]18 donor, but the transposase line can remobilize Minos insertions at various loci.

Table 4. Frequency of GFP-Positive Animals of the Progeny of the Double Transgenic Animals Between Tg[MiCiTnIG]2 and Ju[CiNutMiTP]1a
ID of the double transgenic animalEgg or spermbGFP expression patternRFP
GFP positiveGFP negative
  • a

    The number of juveniles showing enhancer detection GFP expression pattern was not included because the frequency was too low. GFP, green fluorescent protein; RFP, red fluorescent protein.

  • b

    The resutls of the progeny derived from eggs or sperm of the double transgenic animals are shown.

1Egg26/167 (16%)141/167 (84%)85/167 (51%)
Sperm110/203 (54%)93/203 (46%)114/203 (56%)
2Egg37/149 (25%)112/149 (75%)75/149 (50%)
Sperm85/193 (44%)108/193 (56%)95/193 (49%)

Excision of a Single Minos Copy by an Egg Transposase Line

By using E[MiTSAdTPOG]18, we investigated the remobilization of a single Minos insertion in detail. First, we investigated whether excision events occurred in progeny from the eggs of double transgenic animals. As mentioned in the previous section, some of the progeny that did not show the E[MiTSAdTPOG]18-pattern of GFP expression should have the chromosome from which the E[MiTSAdTPOG]18 insertion was excised. If this is the case, their genomes would contain the footprint sequence at the excised site. PCR primers were designed to specifically amplify the typical footprint sequence of Minos, and genomic DNA of GFP-negative animals derived from a cross of eggs of double transgenic animals and sperm of wild types was subjected to PCR analysis with the primers. In PCR analysis, 18.7% (n = 96) of animals showed PCR bands (Fig. 4A), suggesting the presence of the footprint sequence in their genome. As a negative control, the progeny derived from the sperm of double transgenic animals were subjected to the same analyses. They did not show such bands, which suggests that excision is specific to the progeny derived from the eggs of double transgenic animals. The intensity of the amplified bands varied among individuals (Fig. 4A). A previous study examining the extrachromosomal excision activity of Minos in Ciona showed that the footprint sequences of Minos tend to be atypical patterns in Ciona (Sasakura et al., 2003a), and the different band intensity in the above PCR analyses may reflect typical and atypical footprint patterns.

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Figure 4. Excision and transposition of a single transposon insertion in the Ciona genome. A: Excision of the insertion of E[MiTSAdTPOG]18. The upper column represents the detection of excision observed in the progeny derived from eggs of a double transgenic animal between E[MiTSAdTPOG]18 and Ju[CiNutMiTP]1. Bands showing the presence of the footprint sequence (arrow) were detected in lanes 3, 4, 6, 14, 16, and 23. The lower column represents a negative control showing no polymerase chain reaction (PCR) bands in the progeny derived from sperm of a double transgenic animal between E[MiTSAdTPOG]18 and Ju[CiNutMiTP]1. B: Footprint sequences detected in the progeny derived from eggs of a double transgenic animal between E[MiTSAdTPOG]18 and Ju[CiNutMiTP]1. The wild-type genome sequence at this site, Minos insertion site of E[MiTSAdTPOG]18 and footprint sequences are shown. The targeted TA dinucleotide in E[MiTSAdTPOG]18 and footprint sequences are capitalized. The numbers in the right-hand column are the frequencies of the footprint patterns obtained in this study. Primer site used in Figure 4A is shown by an arrow.

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The presence of typical and atypical footprint sequences in the genome was confirmed by the determination of DNA sequences (Fig. 4B). In this analysis, the frequency of animals having footprint sequences was approximately 25.6% of the GFP-negative progeny (n = 39), which is slightly higher than the PCR-based detection of the footprint. This may be because some atypical footprint sequences could not be amplified by PCR. From these results, we concluded that Minos transposase expressed in the eggs can cause the excision of a single transposon insertion in the Ciona genome. Our analyses did not find an animal with a recognizable deletion of a flanking genomic sequence during excision of the Minos copy (0/39).

Transposition of a Single Transposon Copy Creates New Insertions

The presence of progeny with GFP expression patterns different from that of the transposon donor line suggests that some of the excised Minos copies were integrated again in the genome. To examine whether the progeny from eggs of double transgenic animals contained new insertion sites, we performed two experiments. First, genomes of animals with new enhancer detection events were subjected to Southern blot analysis. As shown in Figure 5, the progeny with a new enhancer detection event had a single insertion that was different from the original insertion site (n = 10), suggesting that remobilization of a single transposon copy yielded a new, single transposon insertion.

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Figure 5. Southern blot analysis of enhancer detection lines using digoxigenin-labeled gfp probe. WT, wild-type genome that showed no band; Ori, the genome of E[MiTSAdTPOG]18 that showed a single band. ET, genomes of six enhancer detection lines derived from eggs of a double transgenic animal between E[MiTSAdTPOG]18 and Ju[CiNutMiTP]1. Genomes of the enhancer detection lines showed a single band whose size is different from the original insertion.

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Next, the positions of the new insertion sites in the genome were determined by TAIL-PCR (Liu et al., 1995). Both animals showing constitutive GFP expression patterns of pMiTSAdTPOG and those with new enhancer detection patterns were analyzed. We identified 32 and 7 new insertion sites from progeny showing constitutive GFP expression patterns and new enhancer detection patterns, respectively (Table 5), suggesting that remobilization of Minos occurred in the eggs of double transgenic animals. The positions of the insertion sites with respect to the nearby genes were investigated; 58.9% of the insertions were inside genes, and 17.9% were in exons.

Table 5. Sequence of Minos Insertion Sites
GFP expression patternaSequencebScaffold number in the JGI version 1 genome browserNearest gene modelcPosition in the geneChromosome
Original insertionTACTCTTTATTTTGCAGGCATTACTGCGAACGAATGTACTCTTATT61KH.C6.19∼2.2 kbp upstream6
TPOTACATGTAATCCGTTACCATCGAAAAAAGTTGTTTCTTCAAGGCGCGTAT162KH.C1.25Exon1
TPOTATACTGGTTGTTTTTTGCATGTAGAGTAACTTAAAAAATATAACAAC281KH.C1.346Intron1
ETAGAGGGCCAGTTTCTGCCTTATACAGACGGCACGTATTACATCCTGTGAAT148KH.L171.24∼220bp downstream1
TPOTATACTGCTGATATAGTTAATTATTTCTTATAAATATAGTTGAAATTTGAGT80KH.C2.618∼20bp upstream2
ETATAGGGTGGGGTAGGATGGGACACCTTTAGCACAAAACGTCCAAATATCCT28KH.C2.133Exon2
TPOTAGTGTAAATATAGTTTGTCTGTATAATGTACATGGTTAAGCACACAGAATA46KH.C2.656Intron2
ETATAACTGATAAACAATTGCTTTAAACGGATTTATCGTTCGCGCGTTTTTCA233KH.L108.4∼2kbp upstream2
ETACATCATTAGTGTCTTCCGGTTTTCGTTAAAATTTATATTAAGAAATTGAG57KH.C3.314∼4.5kbp downstream3
TPOTACCTGGCTGTAGAACCTAATGTATGGCTGATAATTTGGACAACCCATCAGT57KH.C3.488Intron3
ETATGTAAACATGTAATTCTTTTATTTAGCACTGGTGGAAAGAAGAAGCAGTT20KH.C3.656Intron3
ETAGTGTAGACCACAAATCATCGCGAACACACGTGGAGCGAAACGGCTAAGTC871KH.C3.102∼3.4kbp upstream3
TPOTATACTATTATAAGCACAAATTTTGAAAGCCCAAAATAGAAAAGATTAGTAC25KH.C3.642Exon3
TPOTATGCTGCGAGGTTTCTCGATAAAACAGTAACAAGAGAGCACTGCTTTAATG2KH.C3.736Exon3
TPOTATATACATGTATATATAACACAGCCTATAGCAGTGGTTTCCAAAGTTATTT44KH.C4.748Intron4
TPOTAGGATTGAAAGAATACACCCAGTTCGTACTGCGCAGACGACGCGAACAGTG6KH.C4.214∼1.1kbp downstream4
TPOTAGGTGCTCTCATCATGATTGTGCTTGCTGTTATCGGAAAGTTCGGAGCTCT6KH.C4.330Exon4
TPOTACACTATGGTATAGGATAGAGCTGCACACACACACATAACTACACATGCAC99KH.C4.215Intron4
TPOTAGCTCCGGTAACACAATGCGTTAATCTGTGCAGGGGTAAGTTGGGACACAG324KH.C5.346∼900bp upstream5
TPOTAGACGAAAACTCCACCAGTCACCTCACAACTTGGTGAAATGTGTAACAG284KH.L3.16Intron5
TPOTATGTGTTTATCTATTTTGTAAAAAATTAAATAAAACATCATTGAAAAAAAC341KH.L4.5Intron5
TPOTATATTAGATTATTTTTTTGTTAAATCGAATTTTCCTGAAGTTCGTTTGCGT8KH.C6.112∼1.4kbp upstream6
TPOTAGTAACCGAAAGCTGTGCAAGCCACCCGCATGGATCAACATGCTACGCGCA8KH.C6.43Intron6
TPOTAGTAGGATGGGGGAAGACGGGACACCTTCGGCGCATAATATCCGAATATTC61KH.C6.78Exon6
TPOTATATACATTTTTTTGTAGCTTTACGCACGTTTACTAACGATTCAAAAATGC61KH.C6.292∼7.3kbp downstream6
TPOTAACTAATGAATATAACATACAATACAGGACACTGGGACAGAGGACATAAAT71KH.C7.225Intron7
ETATGGGGAAAATGGGACAACTTTAGCACATTCTGAATATCCTAATCGCGTTT83KH.C7.695Intron7
TPOTACTGTGAGTATATAGTGGTCGAGAGAGATGAATAAAGGCGTTTGACTCGGT242KH.C7.61Intron7
TPOTAGGTATATTACCCAAACACCCAGGGTGCTACCATCTAGACCCCACTGAGAC127KH.C7.254Intron7
TPOTATACGGTGGCAAGAGGTATGACTTCTACCTGAGAAGGCTAAACTTCGCTGG13KH.C8.433Exon8
TPOTATTCTATAAGCATTAGAGTATCTATCGTCTAAGTCTGAATTTTATTGGTTT4KH.C9.354∼130bp downstream9
TPOTACAATTTAATGTTATTCATACTTTCTCGTACATTTTGGTAGTAAACAAAGA12KH.C11.337∼2.3kbp upstream11
TPOTATGATTACCTTTCGCAAGTAAAAAAACAAACGTAAAATCACAAGGCTACGG256KH.S256.5∼1.7kbp upstream11
TPOTATACATGAAGTTAAAGTTATCGCATGGTAAATGACGCTCAAATGGGTCTGA63KH.C12.100∼140bp downstream12
TPOTACCTTGTGTAGCCATTTTATAACGACTGTCGTTGGCCGCCACGCGAAGCTA37KH.C12.634Intron12
TPOTAGTTGTAGGGTGGGGTAAGACGGGACGCCTTTAGCACATAATATCCAAATA215KH.S215.18Intron13
ETACAATGTAAGGTATATAAGTGGATTGAGGCAGGCTAAGGGAACAGGGATTT55KH.C14.433Intron14
TPOTATACCCATGTGAAAAAGCTATCTTTAGTTACGTTGCTGATAAAATTACCTA143KH.C14.42∼400bp downstream14
TPOTATATGTGCTATAAGTGTCCTATTAGGCCACAGTACTATACATACA2334no gene model  
TPOTACTCATATAGTACTCCTATGTGAATGAATGTAACTTACTTTATCCTCGCGTRepetitive sequence  

The new insertion sites were mapped onto the chromosomes of Ciona (Shoguchi et al., 2005, 2006, 2008) to determine the genomic distribution. We found that 10.2% of the insertions were on the same chromosome (chromosome 6) as the original E[MiTSAdTPOG]18 insertion (Fig. 6A). Among the intrachromosomal transpositions, two events occurred close to the original insertion site (Fig. 6B), suggesting that Minos has low but consistent activity to cause local hopping. The remaining 89.7% of the insertion was on different chromosomes, suggesting that the majority of the transposition occurred interchromosomally. Most of the insertions seem to have been distributed evenly among the chromosomes.

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Figure 6. Transposition of a single Minos copy in the Ciona genome. A: The new insertion sites generated by the remobilization of E[MiTSAdTPOG]18 insertion were mapped onto the Ciona intestinalis genome. The number of chromosomes according to Shoguchi et al. (2008) appears at the bottom. The original insertion site of E[MiTSAdTPOG]18 is shown by a red arrow. New insertions identified from progeny having TPO-pattern green fluorescent protein (GFP) expression and enhancer detection pattern GFP expression are shown by black and blue arrowheads, respectively. B: Local hopping of Minos. The genetic organization around the insertion site of E[MiTSAdTPOG]18 is portrayed in this diagram. The location of the original insertion of E[MiTSAdTPOG]18 and two new insertions generated by the transposition of the original insertion are shown by a black and red two-headed arrows, respectively. Exons of genes predicted in the KH gene models (Satou et al., 2008) are shown by boxes. Ci-meta2 is fragmented in the gene model (KH.C6.19, KH.C6.197, KH.C6.50, and KH.C6.78). A gap of approximately 43 kbp is omitted in this figure.

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DISCUSSION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

In the present study, we created a transgenic line expressing Minos transposase in eggs. This egg transposase line can excise and catalyze the transposition of a single transposon insertion in the Ciona genome, and new insertions can be created. This experimental system provides us with a new method to establish transgenic lines. Because Minos causes frequent interchromosomal transposition, genome-wide enhancer detection and insertional mutagenesis can be conducted with this method. Of the insertions created by this method, 17.9% hit exons of genes. These insertions probably disrupt genes, which suggests that transgenic lines created by this method will be a valuable resource for insertional mutagenesis. In the following sections the usefulness of this system and the characteristics of Minos transposition in the Ciona genome are discussed.

Comparison of the Methods for Creating Minos Insertions in Ciona intestinalis: Advantages of the Egg Transposase Line system

There are five established methods to create Minos-transgenic lines in Ciona: (1) microinjection-mediated transformation by microinjecting the transposon and transposase (Sasakura et al., 2003b); (2) electroporation-mediated transformation by electroporating the transposon and transposase (Matsuoka et al., 2005); (3) enhancer detection by microinjecting transposase mRNA into eggs of the transposon donor line (Awazu et al., 2007); (4) a jump-starter system using a sperm transposase line (Sasakura et al., 2008) and (5) a jump-starter system using an egg transposase line as described in this study. The first two methods are simple ways to introduce the Minos vector and transposase mRNA into eggs to create transgenic lines, while the latter three methods use remobilization of Minos copies present in the genome. The advantage of the remobilization methods over the simple introduction of Minos vectors is their high efficiency in the creation of transgenic lines without laborious experiments, as was noted in the introduction. In addition, the simple microinjection or electroporation methods sometimes create tandem arrayed transposon insertions by unknown mechanisms (Sasakura, 2007), which makes the identification of insertion sites difficult. This disadvantage can be overcome by the egg transposase line-mediated method, because this method usually creates single insertions.

The major advantage of an egg transposase line over a sperm transposase line is the ability to remobilize single transposon insertions. Sperm transposase lines are less efficient at remobilizing Minos insertions, which may be a major reason why the remobilization of a single transposon donor has not been detected with sperm transposase lines. In a previous study, the efficiency of generating enhancer detection lines by sperm transposase lines was approximately 1.0–4.2% per progeny (Sasakura et al., 2008). The transposon donor line used in that study had a concatemer of Minos and a vector backbone that contained 255 transposon copies (Awazu et al., 2007). The frequency of enhancer detection in the system used by Sasakura et al. (2008) was approximately 0.004–0.016% per Minos copy, which is 38–152 times lower than the frequency of enhancer detection in the present study (0.61%).

The other advantage of the transposition of a single transposon insertion is local hopping. In our study, we identified two new insertions close to the original insertion site, which represents 5.1% of the total insertion number (Fig. 6B). These insertions were within approximately 70 kbp of the original insertion sites, which represents 0.04% of the total Ciona genome. If the goal is to create such an insertion by random insertion, too many founders have to be screened, and this is not feasible. The local hopping with the egg transposase line will be a powerful screening strategy to isolate insertions in a specific region of a genome.

A disadvantage of the egg transposase line is that the maturation of eggs is required to isolate new transgenic lines. During the culturing of Ciona, which is hermaphroditic, animals always come to sperm maturation (1–2 months in good culturing condition) more quickly than egg maturation (2–3 months or later). Therefore, longer culturing of double transgenic animals is necessary before the screening of transgenic lines is carried out. In addition, the number of eggs produced by an adult is fewer than the number of sperm. Generally, an animal produces several thousands of eggs under our culturing conditions, while it can produce a much higher number of sperm. To overcome this disadvantage, we have to culture the double transgenic animals in the best possible conditions to produce as many eggs as possible and to shorten the culturing period. If the double transgenic animals are cultured in good conditions, they rear several thousands of eggs, which will yield ∼100 new transgenic lines.

Minos Causes Frequent Interchromosomal Transposition in the Ciona Genome

The Minos transposon caused interchromosomal transposition more frequently than intrachromosomal transposition in Ciona. Approximately 90% of the transposition was interchromosomal. This characteristic of Minos provides a strong advantage for saturation mutagenesis and enhancer detection, because the whole genomic region can be the target of Minos.

The global transposition of Minos has also been observed in mice (Drabek et al., 2003), which suggests that the frequent interchromosomal transposition is not specific to Ciona. However, the mosquito Anopheles stephensi has provided a contrasting result in which the Minos transposon caused more frequent intrachromosomal transposition than interchromosomal transposition (Scali et al., 2007). These different manners of transposition suggest that the mobilization mechanisms of Minos can be changed in a host-dependent manner. Differences in the physiological conditions, host factors and genomic context such as frequent/infrequent CpG methylation or histone modifications may be the cause.

The tendency toward frequent interchromosomal transposition of Minos in Ciona and mice is not the conserved nature of Tc1/mariner transposons, because the SB transposon causes frequent local hopping in the mouse genome (Fischer et al., 2001). This difference may be due to the different natures of Minos and SB. Identification of the factors affecting the transposition of these two transposons is necessary to elucidate the transposition mechanisms of Tc1/mariner transposons.

EXPERIMENTAL PROCEDURES

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

Animals

Wild-type Ciona intestinalis was collected from or cultivated in the Onagawa Bay (Miyagi), Maiduru Bay (Kyoto), and Usa Bay (Kochi). Sperm and eggs were collected by cutting sperm ducts and egg ducts, respectively. Transgenic animals were cultured by an inland system described elsewhere (Joly et al., 2007).

Constructs

pSPCiNutMiTP: An approximately 1-kb segment of the 5′ upstream region from Ci-Nut (Kitaura et al., 2007) was amplified by PCR with primers 5′-ccggatccggggttatgtgagtgaaatagc-3′ and 5′-ccggatccgttgttgatatagtgtaca-3′. The PCR product was digested with BamHI and was inserted into the BamHI site of pSP-MiTP to create pSPCiNutMiTP.

pSBFr3dTPORCiNutMiTP: The Ci-Nut-MiTP fusion cassette of pSPCiNutMiTP was subcloned into pSBFr3dTPORDestF (Sasakura et al., 2008) with a gateway system (Invitrogen) to create pSBFr3dTPORCiNutMiTP.

pSP-Kaede: An open reading frame of Kaede (Amalgaam) was amplified with primers 5′-nngcggccgccatggtgagtctgattaaaccag-3′ and 5′-nngaattcttacttgacgttgtccggc-3′. The PCR product was digested with NotI and EcoRI, and was inserted into the NotI and EcoRI site of pSP-eGFP to create pSP-Kaede.

pSPCiAKRK: An approximately 1-kb segment of the 5′ upstream region from Ci-AKR (Tokuoka et al., 2004) was amplified by PCR. The PCR product was digested with BamHI and was inserted into the BamHI site of pSP-Kaede to create pSPCiAKRK.

pMiCiAKRK: The Ci-AKR-Kaede fusion cassette of pSPCiAKRK was subcloned into pMiDestF (Sasakura et al., 2008) with a gateway system (Invitrogen) to create pMiCiAKRK.

Transgenic Lines

Tg[MiCiTnIG]2 and E[MiTSAdTPOG]18 were described previously (Joly et al., 2007; Awazu et al., 2008). Ju[SBFr3dTPORCiNutMiTP]1 was created by transgenesis with electroporation (Matsuoka et al., 2005; Sasakura, 2007). In this process, 60 μg of pSBFr3dTPORCiNutMiTP was electroporated into wild-type eggs. To increase the efficiency of transgenesis, electroporated animals with bright RFP signals were selected and cultured. A total of 21 electroporated animals were screened, and one founder was obtained.

Tg[MiCiAKRK]1 was created by electroporation of 60 μg of pMiCiAKRK into eggs of Ju[SBFr3dTPORCiNutMiTP]1. In this case, selection of the electroporated animals was not performed according to the method in the previous report (Matsuoka et al., 2005).

Remobilization of Minos in the Genome

Ju[SBFr3dTPORCiNutMiTP]1 and E[MiTSAdTPOG]18 were crossed, and both GFP- and RFP-positive juveniles were selected and cultured until they reached the reproductive stage. Eggs and sperm of the double transgenic animals were collected and crossed with wild-type counterparts. Expression of GFP and RFP in the progeny was observed at the larval and juvenile stages.

To extract genomes from juveniles, we subjected juveniles to digestion in 50 μl of 1 × TE solution containing 0.2 μg/μl Proteinase K for 3 hr at 50°C, followed by 15 min at 95°C to inactivate the Proteinase K. This extract was used for the following PCR analyses.

Detection of Minos Excision From the Genome

We subjected 1 μl of the extract of juveniles to PCR with the primers 5′-cagcaacttgcaactgactatac-3′ and 5′-gcctgcaaaataaagagtacgag-3′ to detect the footprint sequence shown in Figure 4A. To isolate genomic DNA with the footprint sequence, we performed the following procedures. We subjected 1 μl of the extract of juveniles to PCR with the primers 5′-cagcaacttgcaactgactatac-3′ and 5′-gaactgtacagactcttcgatg-3′. These primers flank the insertion site. The PCR products were subcloned into pGEMT (Promega). Transformed E. coli was subjected to PCR with the primers 5′-cgtagtatataaatactcttt-3′ and 5′-gaactgtacagactcttcgatg-3′. The primers perfectly matched the wild-type locus in Figure 4B, and the DNA fragments containing footprint sequences could not be amplified with the PCR. Plasmid DNAs were recovered from the band-negative E. coli clones, and their sequence was determined.

Southern Blotting

Genomic DNA was recovered from sperm or somatic cells of transgenic animals with Wizard Genomic DNA Purification Kit (Promega). The amount of DNA was estimated by electrophoresis. DNA was digested with EcoT14I overnight at 37°C, electrophoresed and blotted on Hybond N+ nylon membranes (GE Healthcare). Digoxigenin (DIG) -labeled probes were synthesized by DIG PCR labeling kit (Roche), and membranes were hybridized with the probes and washed under high-stringency conditions. Signals were detected by CDP-Star (GE Healthcare).

Thermal Asymmetric Interlaced (TAIL)-PCR

For TAIL-PCR (Liu et al., 1995), 40 μl of the extract of juveniles was twice extracted with phenol-chloroform and ethanol precipitated. The purified DNA, or the DNA isolated from sperm, were digested with KpnI and SacI, twice extracted with phenol-chloroform, ethanol precipitated, and used as templates. The PCR was performed as previously reported (Sasakura et al., 2003b).

RT-PCR

Total RNA was isolated from eggs of wild-types or Ju[SBFr3dTPORCiNutMiTP]1 by AGPC methods (Chomczynski and Sacchi, 1987). Residual DNA was digested with DNaseI (Takara). Reverse transcription was performed with Superscript III reverse transcriptase (Invitrogen). PCR was performed with ExTaq DNA polymerase (Takara). PCR primers were described in a previous report (Sasakura et al., 2008).

Acknowledgements

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

We thank Dr. Kazuo Inaba, Hiromi Takahashi, Yasutaka Tsuchiya, Toshihiko Sato, Hideo Shinagawa, and members of the Shimoda Marine Research Center at the University of Tsukuba for their kind cooperation with our study. We thank Dr. Yutaka Satou, Kazuko Hirayama, Yasuyo Kasuga, Yoko Kitagawa, Dr. Koji Akasaka, Dr. Manabu Yoshida, and all members of the Maizuru Fishery Research Station of Kyoto University, the Education and Research Center of Marine Bioresources at Tohoku University, and Dr. Shigeki Fujiwara for the collection of Ciona adults. Dr. Stephen Ekker and Dr. Charalambos Savakis are acknowledged for their kind provision of SB and Minos. We thank Dr. Koichi Kawakami and Dr. Akihiro Urasaki for meaningful discussions. Y.S. was funded by Grants-in-Aid for Scientific Research from JSPS and MEXT and was funded by the NIG Cooperative Research Program.

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  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES
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