Transformation of C. elegans is traditionally carried out by microinjection of DNA into the cytoplasm of the syncytial part of the gonad 36. This predominantly results in formation of long, extrachromosomal arrays (a special characteristic of C. elegans) that can pass through the germline. Infrequently, chromosomal integrations can also occur, depending on the injection conditions. Because this method is efficient, and the extrachromosomal arrays can be maintained in the transgenic strains for a long period of time, the transposon systems have not typically been utilized for the purpose of simple transgenesis.

Reverse genetics in C. elegans has been hampered by the practical inability to target specific mutations to a selected gene via homologous recombination based gene targeting. Developing this technology in C. elegans has been unsuccessful for a long time, because of the inefficient homologous recombination and chromosomal transformation in this model organism. Recently, these difficulties were addressed by the introduction of DNA into the worms using novel methodology, which involved more efficient harvesting of chromosomal integrations and use of the new selection marker unc-119 37. This method is not yet in routine use, however.

TE based mutagenesis is suitable for the purpose of target selected gene modifications in C. elegans. This approach relies on screening mutant libraries created by transposon insertions. In C. elegans it is relatively easy to isolate a specific transposon insertion in any genomic region of interest, using a polymerase chain reaction (PCR) based sib-selection method 38. One difficulty in generating loss of function phenotypes with transposon insertions is that by using endogenous, unmodified Tc1-like elements, the chance for insertions creating null alleles is very low. For example, it was shown that animals homozygous for transposon insertions in the myosin light chain 2 (mlc-2) gene are phenotypically wild type. Interestingly, not even insertions in exons caused loss of function effects, because different atypical splicing events removed the Tc1 part out of the mlc-2 pre-mRNA, leaving only small, in-frame deletions and insertions 39. Analysis of five more Tc1 insertions in two other genes revealed that each mutant expressed substantial quantities of mature mRNA from which most or all of the Tc1 sequences have been removed.

To circumvent the above problem and to obtain knockouts via TE insertions, a strategy of searching for chromosomal deletions following TE excision can be used. The following mechanism is suggested to explain transposition associated deletions. TEs generally create target site duplications upon insertion and leave footprints upon excision. These footprints are generated when the double strand break (DSB), created by the transposase at the excision site, is repaired by the nonhomologous end joining DNA repair pathway (Figure 1b). Alternatively, the DSB can undergo homology dependent DNA repair (HDR). This may not result in any change to the original sequence, if the wild-type homologous chromosome is available as template, and it is used for the repair. HDR can also fill back transposon sequences to different extents, in case a homologous template containing the transposon is available for repair (Figure 1b). However, one of the free DNA ends can scan the DNA at the other end for a short homology; when this is encountered, DNA repair reactions ligate the two ends, thereby deleting the intervening DNA. In the study conducted by Zwaal and coworkers 40, gene specific PCR primer pairs were used to detect Tc1 insertions in five genes, and subsequently to detect the DNA repair mediated partial deletions of these genes following transposon excision. Such deletion derivatives were detected at a surprisingly high frequency (>10^-3). This approach is definitely useful in a model organism such as C. elegans that has short generation time and easy culturing properties.

However, the transposon insertion-deletion method described above has limitations as compared with homologous recombination based approaches. Namely, it is limited to isolating only those deletions that are of a size that is detectable by PCR and that occur between the two predetermined gene specific primers; therefore, it is not useful for creating gene mutations other than null alleles. To circumvent these limitations, a method based on transgene directed, transposon mediated gene conversion was recently introduced 41. This approach capitalizes on the observation that an episomal transgene construct can also serve as a template for the repair after the excision of the TE. In this protocol, a plasmid carrying an engineered gene modification and some regions homologous to the targeted locus is introduced into a mutator C. elegans strain harboring the gene specific Tc1 insertion. This strain is then expanded into independent populations, in some of which the DSBs generated by transposon excision are repaired from this extrachromosomal template. These populations are subsequently tested for the presence of the desired mutation. The efficiency of this method seems to be about tenfold higher than that of the standard insertion-deletion approach mentioned above. Recently, the protocol was adapted to the use of Mos1 transposition in worms 42. The advantage of using a heterologous TE is that background transposition events can practically be ruled out.

Mutagenic insertions for forward genetics involve molecular tags that are used to clone the mutated gene rapidly. Therefore, the above strategy utilizing the Tc1 transposon is rather cumbersome for use in forward genetics research in C. elegans, because the Tc1 insertions do not provide unique tags, and so it is difficult to identify the relevant insertion responsible for a given phenotypic change. Moreover, the Tc1 elements are active in all C. elegans strains, at least in their soma. Therefore, a phenotype driven insertional mutagenesis screen is more promising by using the heterologous transposon Mos1, which expedites the process of identifying the mutated genes 3543. It has been shown that mobilization of Mos1 can also be carried out via the generally used binary system of a non-autonomous transposon and a separate transposase source (Figure 1a). These two components are maintained as extrachromosomal arrays in separate strains, and expression of the transposase protein is controlled by a heat shock promoter 34. The mutagenicity of the Mos1 system is in the range of that of Tc elements in mutator strains 43. Therefore, the newly developed Mos1 system is a promising tool for use in insertional mutagenesis in C. elegans.

Genetic transformation in Drosophila is traditionally done by the injection of preblastoderm embryos 47. Enhancement of this transformation procedure via TE transposition into the genome is a widespread technique. Practically, all TEs listed above are successfully used for this purpose in Drosophila. The injection of in vitro synthesized mRNA as a transposase source can further enhance the efficiency of this technique because of the more rapid availability of the transposase, resulting in reduced transgene mosaicism in the embryo and therefore elevated germline transmission rates. For example, in the case of Minos based transformations, such methods have been shown to yield germline transgenesis rates about tenfold higher as compared with straight plasmid injections 48. Moreover, the application of insulator sequences in combination with TEs can facilitate reliable transgene expression, and may protect endogenous genes from the effect of enhancer elements carried by the transposon construct. Indeed, a recent study demonstrated the usefulness of scs/scs', gypsy, and β-globin HS4 insulators to minimize position effects influencing the expression of transgenes delivered by the piggyBac transposon in Drosophila 49.

Homologous recombination is fairly inefficient in flies. Thus, classical gene targeting, as applied in mouse embryonic stem (ES) cells, has not been successfully established in Drosophila. However, induced DSBs can enhance recombination to some extent, and can trigger HDR at the breakpoints; therefore, the excision of TEs can efficiently facilitate gene conversion events. Utilizing this phenomenon, targeted gene replacement in Drosophila via P element induced gap repair has already been reported (some 15 years ago) 50. P elements have also been used for the isolation of rare deletion derivatives after element excision from a gene of interest, to obtain null alleles 51, long before such a strategy was applied in C. elegans.

Insertional mutagenesis using engineered TEs has proven to be one of the most productive and versatile approaches to disrupting and manipulating Drosophila genes on a genome-wide scale, and by far it leads the field among metazoan model systems. The experimental strategies generally utilize the 'jump starter/mutator' experimental setup, as described above. The usefulness of these screens is strongly promoted by the highly developed classical genetic tools and methods in Drosophila.

The P element has been the most widely used vehicle for these purposes. The mutagenicity of P element insertions is higher than that of Tc1/mariner elements (Table 1). Moreover, P elements appear to transpose efficiently with large cargo sequences inserted within the transposon (Table 1). The early mutagenesis screens carried out in Drosophila utilized vectors that harbor marker genes that are easy to screen such as white, and functional bacterial components (antibiotic resistance genes, origins of replication) that aid molecular analysis of the transposon insertion sites. Vectors of later generations were equipped with gene trapping features, representing an improvement to the basic design. The basic strategies employed to enhance the mutagenicity as well as reporting capabilities of insertional vectors by trapping transcription units are shown in Figure 2. Moreover, elements of binary systems for controlled gene expression such as the GAL4 DNA binding transcription factor (GAL4)/GAL4-upstream activator sequence (UAS) system, or for site-directed recombination such as the flip recombinase (FLP)/FLP recombinase target (FRT) system have also been incorporated into advanced vectors. Thus, a range of versatile experimental designs using P elements for insertional mutagenesis has been developed.

Enhancer trap screens are designed for the identification of enhancers by activating a reporter gene within the P element (Figure 2b). In early enhancer trap versions, the transposase promoter itself was utilized as a weak promoter to capture the enhancers in the genome 52. Gene trapping is based on the activation of a promoterless reporter gene that is dependent on splicing between the exons of the trapped gene and a splice acceptor site carried by the transposon (Figure 2c). Thus, gene trap vectors both report the insertion of the transposon into an expressed gene, and have a mutagenic effect by truncating the transcript through imposed splicing.

Both enhancer and gene trap vectors have been combined with the yeast GAL4/UAS transcription activation system (Figure 2). For example, enhancer detection vectors that direct the expression of GAL4 in a genomic integration site dependent manner have been developed 53. Therefore, such vectors report the expression pattern of trapped enhancer elements, and can also activate a cloned, UAS driven gene of interest in a tissue specific manner (specifically, in those cells in which GAL4 is expressed). The system proved to be highly useful for identifying genes that are involved in a variety of biological processes, and many GAL4 driver lines have been created and made available to the research community 54.

More sophisticated vectors that contain a polyA trap cassette that reports insertion into a Pol II transcription unit have also been developed (Figure 2d). Because polyA trap cassettes have their own promoters, they can report insertion into genes irrespective of their expression status in a given cell type. Importantly, polyA trapping is not expected to be mutagenic, because the vector is not designed to express the downstream exons of the targeted gene at the protein level. Therefore, polyA trap insertions are unlikely to cause dominant effects. Dual tagging systems that combine both gene trap and polyA trap elements (Figure 2e) have been used both in mouse 55 and in Drosophila 56.

Protein trapping (Figure 2f), similar to the trapping systems described above, is also based on hybrid splicing events between an endogenous gene and the transgene cassette carried by the transposon; however, the vector is designed to ensure that the inserted reporter manifests at the protein level. A protein trap strategy to detect GFP tagged proteins expressed from their endogenous loci has been developed by Morin and coworkers 57. The P element based PTT vector was constructed to tag proteins randomly with an enhanced GFP, without disrupting their subcellular localization. PTT carries an artificial exon that encodes GFP and is deprived of initiation and stop codons but flanked by splice acceptor and donor sequences. To enhance protein tagging efficiency, a series of constructs were created to allow reporter gene activation in each of the three reading frames (Figure 2). GFP chimeras typically retain the localization properties of the trapped proteins, except when GFP disrupts a domain necessary for subcellular targeting. Thus, transgenic lines exhibiting tissue specific GFP expression and targeting the GFP signal to virtually any compartment of the cell can be recovered. A disadvantage of the system is that it relies on the relatively rare intronic insertions of the P element. Other transposons such as piggyBac may produce more intronic insertions, which may be better suited to the requirements of such a screen (Table 1; also see below).

Another way to manipulate a trapped transcription unit that has already been proven to be useful is the targeted over-expression and/or mis-expression system (Figure 2g). Using this method, one can bring about over-expression of the full length or truncated protein product (depending on the position of transposon insertion) of the targeted gene, thereby producing dominant phenotypes by overdosing the affected gene product. An improved version of this method is the modular mis-expression system developed by Rorth 58. It allows directed mis-expression of P element targeted genes in any temporal or spatial pattern. It is practically the reversal of the GAL4/UAS system. This modular mis-expression system benefits from the insertional preference of P element; specifically, it tends to insert upstream of the ATG codon of transcription units 58. Therefore, the integrated transposon carrying UAS enhancer and promoter elements can frequently over-express the native full length protein. Such insertion screens can identify genes that, when over-expressed or mis-expressed in a pattern of interest, give a specific phenotype or modulate an existing mutant phenotype. For example, in the first screen conducted by Rorth 58, when activated in the developing eye, 4% of insertions gave a dominant phenotype. The next study by Rorth and coworkers 59 demonstrated the usefulness of the system in genetic interaction screens. They identified many known and new genes involved in the migration of border cells in the ovary, by suppressing the cell migration defect on a mutant background.

An important consideration for the mutagenicity of any transposon is its insertional preferences. The insertion pattern of P elements (and that of all other TEs) is non-random (Table 1). Moreover, P element insertions have numerous 'hotspots' and 'cold regions' on a genome wide scale. Investigators at the Berkeley Drosophila Genome Project have created and analyzed large collections of insertions utilizing some of the vectors mentioned above 60. They found that, interestingly, the observed hotspots can be divided to 'common hotspots' and 'screen specific hotspots'. The first class is commonly hit by all P element vectors, whereas the second is preferentially hit only in a particular mutator strain 60. Hotspots seen in local hopping are excluded from the second class. The existence of screen specific hotspots suggests that the specific parameters of the screen, such as the structure and location of the mutator transposon, can affect the spectrum of hotspots and the diversity of the targeted genes. This phenomenon is poorly characterized, and the underlying cellular mechanisms remain unclear.

Common hotspots represent the main obstacle to full genome coverage with P element vectors. It is estimated that about 150,000 P element insertions might need to be screened to obtain 87% saturation of the estimated 13,500 Drosophila genes 61. To complement the use of P elements, alternative transposon systems have successfully been contributing to large scale insertional mutagenesis of the Drosophila genome. For example, investigators at the Berkeley Drosophila Genome Project have already included piggyBac and Minos element based screens in the collection. A large scale comparison of the insertion patterns of piggyBac and P element vectors has already been reported 62. It has been found that piggyBac exhibits an insertional preference that is distinct from that of P elements. It does not share chromosomal hotspots that are associated with P elements and, although piggyBac favors genes as targets, it lacks the bias for 5' regulatory sequences 62. The target preference of Minos also differs from both P elements and piggyBac, and appears to be similar to that of SB (Table 1). Therefore, these TEs are used as complementary tools for Drosophila mutagenesis.

The combined use of different TEs is common in Drosophila. Practically all of the insertional mutagenesis screens have been established by the creation of 'jump starter' and 'mutator' P element lines, whose generation is facilitated by use of other TEs, generally the Hobo or Hermes elements. For some more sophisticated screening strategies, it is also important to have alternative TEs for effective insertional mutagenesis on a background already carrying P elements and possibly also Hobo or Hermes. Hacker and coworkers 63 used piggyBac for insertional mutagenesis on a chromosome harboring P element inserted FRT sites for the generation of mitotic recombinants. The FRT sites remained stably integrated, demonstrating that the two systems are compatible.

Chromosomal deletions in Drosophila are indispensable, classic genetic tools for mapping mutations, characterizing alleles, and identifying interacting loci. The P element based methods for creating deletions are extremely useful because the end-points of the deletions are molecularly marked. Furthermore, their positions can be designed, which is of particular importance in avoiding haplolethal or haplosterile loci. Two P element based methods are in predominant use to create chromosomal deletions 64. One is based on the observation that P transposase often induces chromosomal aberrations that involve the sites of two distinct P element insertions. The process behind this is called 'hybrid element transposition', in which the TE ends come from separate elements rather than a single element (the 5' end of one element pairs with the 3' end of another one at a different location) 65. This results in different chromosomal rearrangements from which the deletions can be selected. An improved version of the transposon based method for creating deletions makes use of the FLP/FRT site specific recombination system (Figure 3a). Transposon vectors, usually the P element, are used to facilitate single copy chromosomal integration of FRT sites. FLP mediated recombination between two of these P elements can result in chromosomal rearrangements. With a sufficiently high density of starting insertions, even single gene deletions can be created. Progress is currently being made toward covering the genome of Drosophila with characterized deletions 64.

This technique can also create predesigned chromosomal translocations between the chromatids in flies with transgenic FRT sites at identical positions on the homologous chromosomes 66. Thus, FLP mediated recombination can be used to generate mitotic clones (Figure 3b), in which only clones of the cells of interest are homozygous for the studied mutation, whereas the rest of the organism is heterozygous. Spatially or temporally controlled expression of FLP recombinase can create homozygous clones in any tissue or developmental stage of interest.

Chromosomal inversions are also powerful tools in classical Drosophila genetics. They have been applied in the majority of genetic screens in Drosophila as balancer chromosomes, because they can block recombination within the region of the inversion. For practical reasons, the balancers carry visible markers, and are also designed to be recessive lethal. Most of them were isolated as spontaneous or induced chromosomal rearrangements having break points in vital genes. TEs have not been typically used to engineer balancers. The balancers greatly help to map and maintain mutations in the Drosophila genome.

Classic ways to induce expression of foreign genes in vertebrates rely on microinjection of nucleic acids into oocytes or fertilized eggs. Two main drawbacks of these approaches are the low rates of genomic integration and that the injected DNA generally integrates as a concatemer. Both drawbacks can be circumvented by utilizing transposition mediated gene delivery, because it can increase the efficiency of chromosomal integration and facilitates single copy insertion events. Single units of expression cassettes are presumably less prone to transgene silencing than are the concatemeric insertions created using classical methods. Retroviral vectors are also useful tools for the same purpose, but their integration pattern is potentially more mutagenic because of their preference for the 5' end of transcription units (for review 14). In case of transgenesis, a single copy insertion away from endogenous genes is clearly desired. The insertional spectrum of Tc1/mariner elements satisfies this need the best (Table 1), because these elements integrate randomly at the genome level, and do not exhibit pronounced bias for integration into genes. Another particular problem concerning transgenesis is that founders that develop from the injected oocytes or eggs are predominantly mosaic for the transgene, because integration generally occurs relatively late during embryonic development. Therefore, in order to promote successful transmission of the transgene through the germline to the next generation, it is necessary to shift the window of integration events to as early as possible. This can be facilitated by co-injection of engineered transposons with transposase mRNA. This method has been employed to generate transgenic zebrafish with Tc3 73, Mos1 74, Tol2 72 and SB 75; transgenic Xenopus with SB 76 and Tol2 77; and transgenic mice with SB 787980.

To carry out phenotype-driven forward genetic screens in vertebrates, one must achieve efficient germline mutagenesis. To date, the most efficient mutagen in the mouse germline is the chemical N-ethyl-N-nitrosourea (ENU). Mutagenesis rates using ENU are two to three orders of magnitude higher than those with insertional mutagens 81. However, ENU is an ethylating agent that generally causes single base pair mutations. Therefore, the average phenotypic effect of an insertional vector can be more dramatic, and the total number of insertions required to reach the same mutagenic effect is expected to be lower. Moreover, identification of the point mutations causing a phenotype requires positional cloning, which is time consuming and laborious work. Therefore, insertional mutagenesis is an attractive alternative to ENU screens.

Both TEs and retroviral vectors can successfully be applied to insertional mutagenesis. Cultured cells can be efficiently infected with viruses 82. A pseudotyped retrovirus has also been applied to generate insertions in zebrafish via microinjection of virus particles into blastula stage embryos 83. In this approach, however, the transgenic founder fish aremosaic and must be out-crossed to establish F₁ fish with retrovirus insertions. Therefore, one drawback of the application of retroviruses is that the convenient 'jump starter' and 'mutator' method is not applicable with these vectors. Moreover, handling of such viruses may bring up safety issues.

Using TEs as tools for insertional mutagenesis is simpler, and the 'jump starter' and 'mutator' scheme can be applied. In Drosophila, transposon mediated insertional mutagenesis is predominantly carried out using such a scheme. This is also the preferred method in vertebrates, and it has successfully been adopted for germline mutagenesis in the mouse, but not in zebrafish, in which the screens are currently performed by co-injection of the transposon DNA and the transposase mRNA.

In zebrafish, SB and Tol2 were shown to be useful for insertional mutagenesis in co-injection experiments 8485868788. In the mouse, it has been demonstrated that SB, Minos, and piggyBac transposases can function in transgenic animals 1718687889909192. Recently, SB based insertional mutagenesis was also established in the rat 9394. In these experiments, chromosomally resident transposon vectors were mobilized in transgenic animals that either ubiquitously expressed the transposase or expressed the transposase in the male germline using the protamine 1 (Prm1) promoter. SB based insertional mutagenesis has successfully been applied in mice to recover a range of mutant phenotypes in a crossing scheme applying a balancer chromosome 95. This and other studies 96 demonstrated that local saturation mutagenesis of a genomic region is a realistic goal using the SB transposon system with a chromosomally resident transposon donor site. The Minos transposase has also been shown to mobilize non-autonomous Minos elements in mice by transposase expression in the oocytes using ZP3 69 and in the lymphocytes using CD2 promoters 91. PiggyBac has also been used in co-injection experiments in mice 68. The activity of Tol2 element has already been demonstrated in mouse ES cells 97 and in vivo in the mouse liver 98.

All of the vectors used in vertebrate insertional mutagenesis to date are versions of gene trapping insertional mutagenic constructs (Figure 2), equipped with elevated mutagenicity and other useful properties. The mutagenicity of gene trap vectors is higher than that of simple insertional vectors, and they enable easy identification of the mutagenized gene by reverse transcription PCR of composite transcripts composed of sequences of the insertional vector and the endogenous gene. In cell culture, drug resistance markers are generally in use, whereas in animal systems other reporters such as GFP are sufficient for this purpose. Similar to the GAL4/UAS system in Drosophila, a conditional, tetracycline regulated system has been shown to be applicable to TE mediated insertional mutagenesis in mice 99.

As an alternative to the loss of function approaches, targeted over-expression and/or mis-expression has been shown to be efficient in somatic tissues of mice using SB. Viral enhancer promoter elements incorporated into SB vectors were shown to be useful in inducing cancer in experimental animals 100101. These screens can also capitalize on TEs with an intronic preference for insertion, such as members of the Tc1 family. In order to devise customized screens for cancer development, a current approach is focusing on establishing mouse lines that conditionally express the transposase 102. One approach is to express the transposase from tissue specific promoters. The second is to generate a Cre recombinase inducible transposase allele, and to take advantage of the many existing Cre strains to induce mutagenesis in specific tissues in mice 102.