Insertional mutagenesis in Arabidopsis can be conveniently accomplished either by Agrobacterium mediated T-DNA transformation, or by transposable element mobilization. Recent advances have made T-DNA transformation one of the most efficient methods to generate large numbers of insertions in the Arabidopsis genome (1-3). Typically, several hundred independent T-DNA insertions can be generated per transformed plant. This is many more insertions than in most transposon schemes, but despite the labor involved, transposons have some important advantages:

• Firstly, transposons typically integrate into the genome as simple, single-copy insertions. T-DNA, on the other hand, tends to be integrated in multiple arrays.
• Secondly, transposed elements can be remobilized resulting in reversion and short range transposition, while T-DNA insertions are more-or-less permanently integrated into the genome.

These features make transposable elements uniquely suitable vehicles for a powerful form of insertional mutagen, known as the gene trap, or enhancer trap.

Enhancer traps and gene traps are reporter gene constructs that can respond to cis-acting transcriptional signals when integrated into chromosomal DNA (reviewed in 4, 5). Insertional mutagenesis using these "traps" involves generating a large number of individuals that have the reporter gene integrated at different sites throughout the genome. Their progeny are collected and examined for (1) expression of the reporter gene; and (2) mutant phenotypes caused by insertion. In lines in which the reporter gene is inserted within or near a chromosomal gene, reporter gene expression mimics that of the chromosomal gene.

In the last few years enhancer traps and gene traps have been extensively exploited in Drosophila and in mouse developmental genetics (6-14), and several modifications have been made to the basic systems. In plants, gene traps have taken a number of different forms, depending on the reporter gene construct and the vector used for insertional mutagenesis. T-DNA and transposons have both been used to introduce promoter and enhancer traps into Arabidopsis (15-17). In this review, we will describe the enhancer trap and gene trap transposon system that we and our colleagues have developed in Arabidopsis (18, 19). First, though, we will review additional transposon systems in Arabidopsis in order to illustrate some of the genetic strategies employed.

Heterologous transposon mutagenesis in Arabidopsis

Transposon mutagenesis in Arabidopsis has become increasingly sophisticated in recent years. Although some endogenous, mobile Arabidopsis transposons have been found (20), heterologous transposons from maize have so far proven most useful (21-23). This is largely because the genetic properties of these transposons are relatively well understood, so that the timing, range and frequency of transposition can be carefully controlled.

Two transposable element systems from maize have been widely used in Arabidopsis. They are:

• Suppressor-mutator(also called Enhancer) -abbreviated Spm (or En) (24)
• Activator/ Dissociation - abbreviated Ac/Ds (25-28)

In each case, transposase genes, encoded by the autonomous elements Ac and Spm, are required to mobilize non-autonomous elements, Ds and dSpm, respectively. Non-autonomous elements are deletion derivatives of autonomous elements that retain terminal sequences but lack transposase genes. The properties of Spm and Ac are summarized below:

For transposon tagging, transposase genes and non-autonomous elements are separately introduced into Arabidopsis by T-DNA transformation. Transformants are selected that have the transposon-bearing T-DNA integrated at a single locus. Mutagenesis is typically initiated by crossing homozygous plants that have non-autonomous elements with homozygous plants that have transposase genes. In the F1 progeny of these crosses, non-autonomous elements will transpose to new locations during development.

In order to recover stable germinal insertions, F1 progeny are self-pollinated, and F2 plants are selected that have inherited a transposed element, but have lost the transposase gene by segregation. Loss of the transposase is important because unstable transposed elements will revert or transpose germinally causing confusion in progeny analysis. They will also revert or transpose somatically, which could obscure some mutant phenotypes caused by insertion. These problems can be avoided by selecting against the T-DNA that carries the transposase gene, typically by using a negative selectable marker on the T-DNA. There are several ways in which the F2 progeny of a mutagenesis cross can be enriched for those progeny that carry stable transposed elements. Two of these are described below.

Mutagenesis is initiated with the Ds or dSpm element located in the untranslated leader of an antibiotic resistance gene (Figure 1A). Chloroplast biogenesis in plants is sensitive to growth on medium supplemented with some antibiotics, resulting in bleached inviable seedlings. Excision of the transposon from the resistance gene results in restoration of marker gene expression, so that green seedlings can be selected for antibiotic resistance (36). By using a different antibiotic resistance gene within the transposon, seedlings that have the transposon integrated at a new site can be selected by including both antibiotics in the selection medium.

This type of selection scheme has been widely used in plants and has been generally successful. However, it is relatively inefficient for the following reasons:

• excision events that occur late in the development of F1 plants will be inherited by the male, but not the female gametes (or vice versa) (37). In this case, self-pollination results in F2 plants that are resistant to both antibiotics, because they are heterozygous for the post-excision and pre-excision T-DNA alleles. These plants do not necessarily carry a transposed element.
• most transpositions are short range (38, 39), so that two linked insertions must be generated for every unlinked insertion.
• post- and sometimes pre-excision T-DNA loci are deliberately retained in selected progeny. Marker genes on the T-DNA can interfere with marker and reporter gene activity on transposed elements.

As an alternative to selection by excision, transposed elements can be selected directly (Figure 1B). For this purpose, a negative marker is placed on the T-DNA that carries the transposon (19). A positive marker is included within the transposon itself. F2 progeny that carry a transposed element can be identified by selecting for the transposon and against the T-DNA. Doubly resistant plants carry the transposon but not the T-DNA, typically because the transposon has moved away. Because loss of the T-DNA depends on segregation from the transposed element following meiosis, unlinked elements are preferentially selected. Furthermore, loss of the T-DNA ensures that it cannot influence transposon expression and inheritance. Positive markers include antibiotic resistance genes, as before. Negative markers include enzymes that activate herbicide analogs that are otherwise non-toxic (40-43).

A minor problem with this scheme is that very short-range transpositions in F1 plants can result in disruption of the negative marker gene on the T-DNA. This will result in F2 progeny that inherit the T-DNA but do not have a long-range transposition. Although only a handful of F1 plants will normally undergo transposition into the T-DNA, pooling F2 progeny from different F1 plants should be avoided because of this problem.

Figure 1. Selection schemes for transposed elements. A. Selection for excision. Donor Ds element carries a hygromycin resistance gene and interrupts a streptomycin resistance gene. In the presence of Ac, the Ds excises, resulting in streptomycin resistance. F2 plants that are resistant to streptomycin and hygromycin have a Ds element that has excised from its original location. Both linked and unlinked transpositions are recovered. B. Selection for unlinked transposition. Donor Ds element carries a kanamycin resistance gene and is adjacent to the IAAH gene which acts as a negative-selectable marker. In the presence of Ac, the Ds excises. F2 plants that are resistant to both kanamycin and NAM will carry a Ds element that has transposed and segregated away from its original location.

Enhancer and gene trap transposons

Enhancer trap reporter genes have a minimal promoter that is only expressed when inserted near cis-acting chromosomal enhancers. Gene trap reporter genes (promoter traps and exon traps) have no promoter, so that reporter gene expression can occur only when the reporter gene inserts within a transcribed chromosomal gene, creating a transcriptional fusion. Our system uses both enhancer trap and gene trap reporters, and is based on the maize Ac and Ds transposable elements (Figure 2). The Ds elements carry the b -glucuronidase (GUS) gene as a reporter (44) and the Neomycin phosphotransferase (NPTII) gene (conferring resistance to kanamycin) as a selectable marker. There are two classes of Ds elements: (1) an enhancer trap element which has a minimal promoter fused to the GUS gene, and (2) a gene trap element, which has a multiple splice acceptor fused to the GUS gene. Random insertions of the Ds element throughout the genome allow us to detect chromosomal gene expression through the activation of the GUS gene. We can detect this expression in the absence of a mutant phenotype, so genes performing redundant functions can be identified.

Figure 2. T-DNA constructs used to introduce Ac transposase and gene trap and enhancer trap Ds elements. IAAH=indole acetic acid hydrolase, NPTII=neomycin phosphotransferase, GUS=b -glucuronidase.

Most importantly, it is possible to assess the role of essential genes during development by using gene traps. This is because the GUS expression pattern can be examined in viable heterozygotes even when homozygotes (or haploid gametes) are inviable. Plant development is highly reiterative, so that many genes have both early and late functions. Other types of insertional mutagenesis would be unable to uncover late functions when the early function is essential, resulting instead in a large number of poorly distinguished lethal mutations.

Mutagenesis is initiated by crossing plants homozygous for one of the Ds elements to plants containing the Ac transposase gene (Figure 3). The resulting F1 seed are planted, and the plants are allowed to self-pollinate. The F2 seed are harvested from each individual F1 plant, and plated on media containing kanamycin (selects for the Ds element) and NAM (selects against both T-DNAs). The double resistant F2 seedlings (called transposants) contain a transposed Ds element. The NamR KanR seedlings are transplanted to soil, and allowed to self-fertilize. F3 seed are collected, and grown, and the resulting plants are stained for GUS activity and examined for mutant phenotypes.

Figure 3. Crossing scheme used to generate transposant insertions.

Protocol 1. Mutagenesis

• Fine pointed forceps
• Dissecting microscope with fiber optic light source
• Plant growth cabinets and soil

1. Sow parental seed homozygous for the transposase gene in individual 2" pots and grow under optimal conditions: 16 hour days are recommended. Plant homozygous Ds seed one week later.
2. Emasculate the first ten flowers from each Ds parent as they appear (before pollen is shed) in groups of 3 flowers at a time. Leave each flower overnight before pollination with homozygous transposase pollen. Remove any unfertilized buds, and mark the branch.
3. Collect the seed approximately 2 weeks later, just before the siliques open. Collect and store in coin envelopes sealed with tape.
4. Plant the F1 seed individually with a damp spatula on moistened compost in the greenhouse. On bolting, provide each plant with a small plastic or wire stake, and a 12" polythene collection tube.
5. Grow F1 plants until they emerge from the tube and stop producing flowers. Stop watering them, and harvest 5-10,000 F2 seed from each plant 1 week later. It is convenient to work in batches of 2-4,000 F1 plants at a time.

• Germination media: 4.3 g/ l MS salts (GIBCO/BRL), 10.0 g/ l sucrose (tissue culture grade, Sigma), pH to 5.7 with 1 M KOH.
• Plates: Add 7 g agar per litre GM and autoclave. Allow to cool to 60°C, then add 325 m l per litre NAM (alpha-napthalene acetamide, Sigma) from a 2 mg/ml stock in 95% EtOH. Add 1 ml kanamycin from a 50 mg/ml filter sterilized stock. Final concentrations are 0.65 m g/ml NAM, 50 µg/ml kanamycin. Pour plates at least 8 mm deep; this is important for the selection.
• Top agar: Add 7 g agar per liter GM and autoclave. Cool to 60°C before use. This can be stored at room temperature, and melted in the microwave before using.

• 20% household bleach (1% Na hypochlorite) with 0.1% Tween 20.

1. Aliquot F2 seed into approximately 500 seed (10 mg) batches in 15 ml plastic tubes.

2. Sterilize the seed by following the protocol below. Work in the hood, and use sterile technique.

b. Add 1 ml of bleach solution to the seed, leave 5 min.

3. Add top agar to 4 ml. Replace the cap, invert to mix, and pour quickly onto 100 mm square plate. Swirl gently to evenly distribute seeds.
4. Place at 4°C for 4 days to encourage even germination, and then move plates to a growth chamber for 3-5 days. Grow at 21°C under reduced light: exact conditions will vary from chamber to chamber, but reduced light encourages hypocotyl elongation, which enhances NAM selection. Initial growth for 2-3 days in the dark can improve selection.
5. Seedlings resistant to NAM can be recognized by their long, unbranched roots, long hypocotyl and upright stature. Most of these will be pale green indicating kanamycin sensitivity. Select those NAM resistant seedlings that are dark green (kanamycin resistant) and transfer onto fresh NAM/kan GM plates. Use sterile forceps, and work in the hood. After 5-7 days, the phenotypes will be more extreme on fresh rather than on the crowded plates, and easily recognized. Always transfer some sensitive seedlings as a control.
6. Transfer seedlings to soil when the phenotypes are unambiguous. The number of double resistant seedlings varies from 0 to 2.5% in each family. Transplant from 1-5 seedlings to soil, but thin all but one plant on flowering. Collect F3 seed.

Screening transposant lines

Transposant lines can be screened for mutant phenotypes caused by insertion, and for patterns of reporter gene expression. Many different screens for the phenotypic consequences of Ds insertion can, of course, be devised. In some cases, for example, it may be beneficial to stain transposants first, and then use the staining pattern to guide phenotypic examination. This is very useful in cases where subtle phenotypes are expected. However, it should be remembered that only a fraction of insertions into genes result in reporter gene expression, so that lines without reporter gene expression can still be very useful. Simple screens and tests for bona fide insertion phenotypes are discussed below by way of illustration. We have found these procedures to be applicable to most stages and tissue types examined, although specific modifications will be required for some stages.

Individual F2 plants can be either homozygous or heterozygous for the transposed Ds element, so any phenotype associated with the insertion could either be present in every F3 plant (in the case of a F2 homozygote) or could segregate 3:1 normal to mutant (if the F2 plant was heterozygous and the mutation is recessive). Segregation ratios substantially more than 3:1 may indicate that the mutation arose after transposition and is not associated with the insertion.

Lethal insertions can be scored most conveniently in F2 transposants, by opening developing siliques and scoring unfertilized ovules and colorless embryos (e.g. (18)). Lethal mutations in our lines arise at a frequency of about 4%, but should be carefully tested for heritability and association with the Ds insertion. This can be accomplished by plating F3 seed on kanamycin. The Ds insertion carries a kanamycin resistance gene. Thus, if the insertion is responsible for the lethal phenotype, all resistant F3 plants (which are heterozygous for the insertion) should give rise to plants with semisterile siliques or defective embryos. Lethality in the female gametophyte or in the embryo should result in poor transmission of the kanamycin resistant trait, with ratios of between 1:1 and 2:1 resistant to sensitive F3 seed.

Screening for reporter gene expression patterns should be done wherever possible in heterozygous plants or in segregating families that include phenotypically normal individuals. This is because homozygosity for the insertion can affect reporter gene expression patterns in many cases. 5 progeny from each line should be stained to ensure that at least one has an insertion. Alternatively, plants can be pre-selected on kanamycin before staining, although this may affect seedling reporter gene expression patterns. Reporter gene expression patterns in flowers can be tested directly in F2 plants without killing them. However, it is wise to examine several individual plants wherever possible to avoid artefactual staining patterns, so that staining in the F3 is recommended.

Potassium ferricyanide is included in the staining reaction in order to catalyze dimerization of the indigo monomer, which is the product of the glucuronidase reaction. The monomer is soluble, while the dimer is not, resulting in a precipitate at the site of enzyme activity. In the absence of at least 1.5 mM ferricyanide, diffusion of the indigo monomer results in artefactual staining patterns. Unfortunately, ferricyanide ions also inhibit the GUS enzyme at these concentrations. This inhibition can be partially overcome by using long incubation times for weaker staining patterns. Clearing of the tissue following staining is most simply and gently accomplished using 70% ethanol, which we find adequate for all tissues we have examined. However, other more drastic clearing methods can be employed, but need to be carefully controlled with respect to re-dissolving the stain and damaging the tissue. A good discussion of clearing procedures can be found in Sylvester (45). Cleared tissue is mounted in glycerol for optimal Nomarski optics.

• GUS Stain: 100 mM NaPhosphate pH 7, 10 mM EDTA, 0.1% Triton X-100, 1 mg/ml X-Gluc., 100 µg/ml Chloramphenicol, 2 mM Potassium Ferricyanide and 2 mM Potassium Ferrocyanide
• Forceps
• Vacuum dessicator
• 24-well microtitre plates

1. Sterilize 20 - 30 seeds from each F3 in an eppendorf tube according to Protocol 2, step 2, but using smaller volumes. Solutions can be easily removed with an aspirator.
2. Resuspend seed in 0.7 ml 0.1% agar. Spread on 60 mm GM plates without selection, distributing evenly.
3. Place plates at 4°C for 4 days, then transfer plates to growth chamber for 4-8 days.

1. With fine forceps, carefully transfer seedlings from each plate into a microtitre well containing 500 µl of GUS stain. Some staining protocols call for a mild fixation before or after staining (46). This may be useful for some tissue types.
2. After seedlings are transferred, place the microtitre dish in a dessicator, and draw vacuum for 10 minutes.
3. Release vacuum slowly, wrap the dishes in aluminum foil (X-Gluc is light sensitive) and incubate them at 37°C overnight to 3 days. If incubation is longer than overnight, the plates should be sealed to prevent evaporation.
4. Remove the staining solution and replace with 70% EtOH. Clearing is more rapid at 37°C, and several changes may be necessary. The stained seedlings are stable for several weeks in ethanol at 4°C.
5. Examine stained seedlings in 70% ethanol under a dissecting scope. Mount selected seedlings in a drop of 50% glycerol under a coverslip, and examine at high power using Nomarski optics. The stain will begin to recrystallize in glycerol after a few hours, resulting in large needles. These can be avoided by rapid examination, and by the use of shorter incubation times and higher levels of ferricyanide in the stain. For long staining times (2-3 days) it is advisable to use 2-4 mM ferri/ferrocyanide to limit stain diffusion.

Molecular analysis of transposants

Each transposant line that has a staining pattern or phenotype of interest can be analyzed molecularly to determine the location of the transposed element in the Arabidopsis genome. Flanking genomic DNA or cDNA corresponding to the gene nearest the insertion site can be rapidly obtained by PCR amplification. The resulting products can be sequenced directly, or used as hybridization probes for further analysis.

Although most insertions are randomly distributed around the genome, about 5 percent of our transposants have Ds insertions into the IAAH gene on the T-DNA. These insertions typically have strong ubiquitous reporter gene expression in seedling tissues and can often be discarded on the basis of this expression pattern or on the basis of further molecular analysis (see below). However, if this is not possible, then each GUS positive line of interest can be tested for the presence of the IAAH gene directly. This is done by tissue PCR using primers from two different regions of the IAAH gene (primers Nam1-4). These allow amplification of the gene even if one region has been disrupted by Ds insertion. Primers from the prolifera gene (148AR and 148D) provide an internal control for each reaction. The PCR reactions are performed using plant tissue according to Klimyuk et al. (47).

Protocol 4: Screening for linked insertions into T-DNA marker genes

• 0.25N NaOH
• 0.25N HCl
• 0.5 M Tris, pH 8.0; 0.25% Nonidet P-40
• Boiling water bath
• Thermocycler
• Primers:

NAM1: 5′-GTTTATTTCGGCGTGTAGGACATG-3′

NAM2: 5′-GTAGTCTTTCCGTTTCAGGTG-3′

NAM3: 5′-CATTCCCCACCTTGACGAACTG-3′

NAM4: 5′-GGTCTGAATCCGCTAATCCA-3′

148AR: 5′-AACTGCAGATGCCCTGAACTGGATCTCCA-3′

148D: 5′-AGCTGCAGATTCCTCCAAACACTC-3′

1. Pick a small amount of tissue from one stained seedling from each line (1/2 to 1 cotyledon works well, and too much inhibits the reaction).
2. Collect tissue into a 0.2 ml reaction tube containing 40 µl of 0.25N NaOH. Boil for 30 seconds in a boiling water bath or PCR machine.
3. Add 40 µl 0.25 N HCl and 20 µl 0.5 M Tris pH 8; 0.25% Nonidet P-40. Boil for 2 further minutes.
4. Transfer the tissue to a fresh 0.2 ml tube, and perform PCR reactions according to Klimyuk et al. (47). Run a 2.0% agarose gel; control primers give a 175 bp band; the Nam1 - Nam2 primer pair gives a 368 bp band and the Nam3 - Nam4 primer pair gives a 494 bp band. The presence of either of the Nam bands indicates that the T-DNA is still present. Positive controls (Ac and Ds parents) should be included.

Chromosomal sequences flanking both gene trap and enhancer trap insertions can be amplified using I (inverse) PCR or TAIL (Thermal Asymmetric InterLaced) PCR using standard protocols and primers from the Ds element. These PCR products can be sequenced directly after purification on spin columns or gels by cycle sequencing using dye terminators. Each of these methods depends on the fortuitous location of a primer sequence or restriction site close to the insertion, and consequently is only successful about 50% of the time for a given primer/enzyme combination. It is wise, therefore, to use several primer combinations or several approaches when attempting to amplify a given insertion site.

Amplification of cDNA from gene trap fusions

Gene trap insertions result in transcriptional fusions between the reporter gene and the chromosomal gene into which it is inserted. Consequently, flanking sequences can be amplified by 5′RACE PCR using RNA isolated from the gene trap line. This is useful when the transposon is inserted into a large intron, or when multiple introns make chromosomal sequence hard to interpret. RACE PCR products can also be sequenced directly, except that alternate splicing will result in mixed sequence reads in many cases. In these cases, subcloning will be required.

In either case, it is wise to confirm the assignment of a given sequence to a given line by hybridization of the products to genomic Southern blots, or by PCR using specific primers derived from the sequence. The genomic DNA purification procedure described below allows restriction digestion and Southern blotting as well as PCR.

Protocol 5. Preparation of genomic DNA

Flowers and inflorescences have much smaller cells than leaves or roots, and consequently, they have a lot more DNA. 5 µl DNA (0.5-1.0 µg) in a 20 µl digest is sufficient for a Southern blot with single copy probes. Spermidine (pH 7, 2.5 mM final concentration) greatly aids digestion.