Department of Zoology, University of Wisconsin, 250 North Mills Street, Madison, WI 53706, USA e-mail: ssblair{at}wisc.edu
SUMMARY
Genetic screens for recessive mutations continue to provide the basis for much of the modern work on Drosophila developmental genetics. However, many of the mutations isolated in these screens cause embryonic or early larval lethality. Studying the effects of such mutations on later developmental events is still possible, however, using genetic mosaic techniques, which limit losses or gains of genetic function to specific tissues and cells, and to selected stages of development. A variety of genetic mosaic techniques have been developed, and these have led to key insights into developmental processes in the fly. Variations on these techniques can also be used to screen for novel genes that are involved in non-embryonic patterning and growth.
Introduction
Genetic mosaic techniques are those that induce genetic changes in a subset of cells or tissues in an individual organism. Such techniques have been crucial for our current understanding of a number of developmental processes in Drosophila. There are several reasons for this. Most importantly, the techniques provide a way of examining genetic changes that would be lethal if applied to the entire organism. For example, many of the mutations that completely remove the function of a given gene are homozygous lethal in embryonic or early larval stages of development. This is not a problem if one is studying early development; indeed, the isolation and characterization of embryonic lethal mutations was a landmark in Drosophila developmental genetics. However, if one is interested in studying the later stages of development, the only ways of circumventing early lethality are to use viable alleles or temperature-sensitive alleles (provided that they exist or can be generated), or to use genetic mosaic techniques. If the genetic change can be limited to small numbers of cells, or can be induced at later stages of development, early lethality can often be avoided. Our present understanding of the development of late-developing adult tissues in the fly, such as the appendages, compound eye, internal organs and oocyte, owes much to the use of genetic mosaic techniques.
Genetic mosaics also provide a powerful tool for teasing apart complex
developmental interactions, no matter what developmental stage is being
examined. If removing a gene prevents the development of a specific structure,
is it because the precursors of that structure require the gene? Or is the
requirement in some different group of cells? If the gene is expressed in both
sets of cells, genetic mosaics provide the only means of answering these
questions. Genetic mosaics also juxtapose wild-type and genetically altered
cells, and this can be used to test the `cell-autonomy' of a mutant phenotype.
That is, are the mutant cells affected by the presence of neighboring
wild-type cells? Can the mutant phenotype be rescued by wild-type cells and,
if so, over what range? Conversely, can the mutant cells affect the
development of their wild-type neighbors (sometimes referred to as
`domineering' nonautonomy)? Testing the cell autonomy of a mutation again
helps to determine how directly a gene is involved in a given developmental
decision. And for a novel mutation that affects signaling, one can determine
whether it is required in the sending or receiving cells. For example, cell
autonomy was used to determine that Notch was a receptor, and Delta its
ligand, before any details were known about the Notch signaling pathway
(Heitzler and Simpson,
1991)
Mosaic techniques have also provided much of what we know about normal cell
lineages in Drosophila tissues. Although dye-tracing techniques can
be used to follow cell lineages in the embryo (e.g.
Vincent and O'Farrell, 1992;
Bossing and Technau, 1994
), the
extensive cell proliferation that occurs between embryonic and adult stages
dilutes lineage dyes to undetectable levels. Thus, researchers interested in
later stages of development have relied upon genetic mosaics to mark wild-type
cells and their progeny; the `compartmental' lineage restrictions of adult
appendages were discovered using these techniques
(Garcia-Bellido et al., 1973
;
Garcia-Bellido et al.,
1976
).
In this article I briefly summarize the most popular techniques for generating genetic mosaics in Drosophila. I include not only those mosaic techniques used for examining mutations in endogenous genes, but also the techniques used to limit the expression of gene constructs to particular regions or stages of development.
Making mosaics the old-fashioned way
Surgical approaches
Before the advent of engineered constructs and the like, there were
essentially two approaches to creating genetic mosaics in flies. The first
involved transplantation between different fly strains, either of cells or
nuclei (Geyer-Duszynska, 1967;
Illmensee, 1968
;
Illmensee, 1973
;
Technau, 1986
) [for techniques
see Santamaria (Santamaria,
1986
)] or whole tissues (such as imaginal discs, which form most
of the adult epidermis) (Ephrussi and
Beadle, 1936
) [for techniques, see Ashburner
(Ashburner, 1989
)]. Although
often useful, these surgical techniques have been underused of late,
presumably because of the technical difficulties involved. The small size of
the fly embryo can make cell and nuclear transplantation difficult.
Transplanting mature imaginal discs is easier; they will grow after being
injected into an adult or larva, and will differentiate in pupae. However, it
is difficult to get them to incorporate into the adult epithelium, as they
usually differentiate as a crumpled mass inside the developing pupa, leaving
some structures difficult to analyze. Immature imaginal discs from embryos or
early larval stages are also small and difficult to isolate, although discs
can develop from whole embryos after they have been transplanted into a host
(Simcox, 1997
). And, in many
cases, one would like to study changes to only part of a disc.
Given the power of Drosophila genetics, it was therefore natural for researchers to use purely genetic techniques for creating mosaics. Initially, researchers relied on two techniques to study recessive mutations in flies: chromosome loss and the recombination of homologous chromosomes during mitosis.
Chromosome loss
Several mutations can induce the random loss of the three autosomes, the X
chromosome or the Y chromosome during early cleavages of the fly embryo
(reviewed by Ashburner, 1989).
However, although individual flies with a single X or fourth chromosome are
mostly viable, loss of a second or third chromosome usually causes death.
Certain morphologically abnormal chromosomes are also unstable, such as the
muchused ring-X chromosome. Loss of a ring-X occurs randomly during the first
few divisions of the zygotic nuclei, and creates a gynandromorph, which has a
mixture of female (XX) and male (XO) tissues. This chromosome has been used to
study normal cell lineages and to study mutations on the X. However, nearly
half of the resultant early embryo is XO, so if the X carries a lethal
mutation, the embryo is unlikely to survive to later stages of
development.
Mitotic recombination
A more generally applicable method is the induction of mitotic
recombination between the arms of homologous chromosomes
(Stern, 1936). As shown in
Fig. 1, this can generate, from
a heterozygotic parent cell, two daughter cells that are homozygous for
everything distal to the site of recombination. A low level of mitotic
recombination occurs spontaneously in flies, and this rate can be increased to
a useful, although still low, level by exposing flies to X-rays or
-rays (Patterson,
1929
).
|
Marking cells
One problem that chromosome loss and irradiation-induced mitotic
recombination techniques share is that the location of homo- or hemizygous
cells within the tissue is essentially random. These cells therefore need to
be marked (reviewed by Ashburner,
1989; Lawrence et al.,
1986
). Identifying mutant cells in the adult epidermis is usually
done by linking the mutation being studied to a benign marker mutation on the
same chromosome arm; the marker mutation changes the color (yellow)
or morphology (forked, multiple wing hair, etc.) of the cuticle that
is secreted by a given cell. In the eye, pigmentation mutants or constructs,
such as white, are used in a similar manner.
A variety of techniques have been used to mark the cells of internal
organs, or cells at stages of development before cuticle or pigment are made.
Initially, researchers used mutations in ubiquitously expressed enzymes, such
as a temperature-sensitive mutation in succinate dehydrogenase
(Lawrence, 1981). Homozygotic
cells were identified by their failure to stain after appropriate
histochemical reactions. More recently, constructs encoding several
non-endogenous, histologically identifiable tags have been inserted into the
genome; available tags include ß-galactosidase (ß-gal) (e.g.
Blair, 1992
), a Myc epitope
(Xu and Rubin, 1993
) and green
fluorescent protein (GFP). In mitotic recombination-based approaches, the
histological tags are usually located on the wild-type chromosome, so that the
homozygous mutant cells are identified by the absence of the marker
(Fig. 1). In some cases, the
marker/marker sister of the mutant/mutant
cell can also be identified; this `twin-spot' is useful both as a control for
the effects of the sister mutant/mutant cells, and as an
indication of the location of the recombination event within the tissue
(Fig. 1). However, it should be
pointed out that mutant/mutant cells are occasionally
difficult to identify because of the absence of the marker, especially if the
cells are few in number or the tissue is complex. For one solution to this
problem, see the Gal80 section below.
Clones
Even following irradiation, mitotic recombination is still a relatively
rare event. However, this rarity has also led to the discovery of a useful
feature of Drosophila development, that cell intermixing and
migration are quite limited in Drosophila epithelia. The homozygous
daughter cell produced by a single mitotic recombination event almost always
forms a single, spatially coherent `clone' of descendents
(Fig. 1B). This is unlike the
salt-and-pepper patterns that are commonly seen in genetic mosaics in
vertebrates, and can simplify the analysis of phenotypes. The size of the
clone observed in an adult depends on the developmental stage at which the
larva was irradiated. As the percent of cells undergoing recombination is
constant, the number of clones observed at later stages depends on the number
of target cells that were present during irradiation. For example, clones
induced earlier in imaginal disc development are large but infrequent, whereas
those induced later are smaller but more frequent.
The Minute technique
In some cases, it is helpful to have mitotic recombinant clones that are as
large as possible, for example, when looking for restrictions in cell
migration, or when removing a broadly expressed signaling molecule. This can
be accomplished using the Minute technique (reviewed by
Ashburner, 1989;
Lawrence et al., 1986
)
(Fig. 2). Dominant mutations at
several different Minute loci slow cell division rates. Those
Minute mutations that have been characterized disrupt ribosomal
proteins. When a wild-type (+/+) clone forms in Minute/+ tissue, the
clone has a growth advantage over its slow-dividing neighbors. Interestingly,
the abnormally large +/+ clones do not usually alter developmental patterning.
The Minute technique can thus be used to increase the size of
mutant clones. This technique can also increase the chances that a
mutant clone survives. For example, mutant clones in imaginal discs
are often lost because of a phenomenon called clone competition, where
abnormal slow-growing cells are eliminated in some way by the surrounding
wild-type cells [for recent work on this phenomenon, see Moreno et al. (Moreno
et al., (2002
)]. Giving the
clone a growth advantage using the Minute technique often rescues
it.
|
Additions and improvements to mosaic techniques
The stable insertion of DNA constructs into the fly genome via engineered
transposable elements (most commonly the P element)
(Rubin and Spradling, 1982;
Spradling and Rubin, 1982
) has
made several modifications of and additions to the early mosaic techniques
possible (see Duffy, 2002
).
The initial insertion of a new P element construct into the genome is
still somewhat laborious, requiring the injection of many embryos to generate
one transformant. However, constructs that have already been incorporated into
the genome can be remobilized, and thus `hopped' from one position to another
in the genome, by mating the transformed fly strain to another that carries a
constitutively expressed transposase
(Robertson et al., 1988
).
The mosaic techniques discussed in the following sections use variations on
two different systems, both derived from yeast. The first uses targeted DNA
recombination at FLPase recombination targets (FRTs), which can be driven in
flies by the FLP recombinase (FLPase)
(Golic and Lindquist, 1989).
The second uses the Gal4 transcription factor to drive the expression of
constructs that are coupled to the UAS enhancer sequence
(Brand and Perrimon, 1993
).
Flies carrying most of the constructs discussed below are available as
community-wide resources through the Bloomington Drosophila Stock
Center
(http://flystocks.bio.indiana.edu/).
FRT-mediated mitotic recombination
FRTs have been inserted into proximal locations on each of the chromosome
arms, and several stocks have been generated that express FLPase under the
control of the hsp70 heat-shock promoter (hs-FLPase)
(Chou and Perrimon, 1992;
Golic, 1991
;
Xu and Rubin, 1993
). If a fly
has two FRTs in identical positions on homologous chromosomes,
heat-shock-induced expression of FLPase can cause recombination between the
FRT sites (Fig. 3). This
technique has several advantages over irradiation-induced recombination.
FRT-mediated mitotic recombination rates are much higher than those caused by
irradiation, although they are still low enough to ensure that only a small
percentage of cells will be homozygous. The site of the recombination is also
controlled, so that one no longer has to worry about recombination occurring
in the chromosomal region distal to the mutation, or between the mutation and
the marker. Heat shock also induces less cell death than irradiation. However,
there are also disadvantages to FRT-mediated recombination. Mutations and
markers must first be meiotically recombined onto the appropriate FRT-bearing
chromosome before the technique can be used. Not only does this take time, it
also prevents the technique from being used for extant mutations on the fourth
chromosome, where meiotic recombination does not occur. Moreover, the
technique cannot be used for those genes that are proximal to any available
FRT insertion.
|
|
|
Many UAS constructs have been generated using cloned genes. Another way of
driving the expression of UAS-gene constructs that is more random, but is
proving to be increasingly useful, uses EP constructs
(Rorth, 1996;
Rorth et al., 1998
)
(Fig. 5D). EP constructs
contain multiple UAS sequences that are coupled to a weak promoter; as with
enhancer trap constructs, and have been hopped around the genome. If an EP
construct lands in a favorable position it can, in the presence of Gal4, drive
the expression of the neighboring gene (see
Fig. 5E). Many of these EP
insertions have been precisely mapped in the genome, and thus provide a
ready-made resource for the misexpression of wild-type genes (the insertion
sites are mapped at
http://flybase.bio.indiana.edu,
or available by BLAST at
http://www.fruitfly.org/blast/).
The Gal4-UAS system can also be used to generate loss-of-function mosaics.
The genes in UAS constructs can be engineered to ones liking; and thus
dominant-negative constructs can be used. Another, increasingly popular way of
generating loss-of-function mosaics uses UAS-hairpin constructs to drive
expression of double-stranded (ds) RNA
(Kennerdell and Carthew, 2000;
Fortier and Belote, 2000
;
Piccin et al., 2001
)
(Fig. 5F). dsRNA has not been
effectively delivered into Drosophila except by injecting embryos,
and the effects of such injections do not last until late stages of
development (Kennerdell and Carthew,
1998
). UAS-hairpin constructs, when coupled with appropriate Gal4
drivers, provide another way of generating dsRNA. The hairpin constructs
contain forward and reverse coding regions, which are coupled end to end by a
short spacer sequence. The RNA transcribed from these constructs folds back
upon itself to generate dsRNA, which knocks down the expression of the
corresponding endogenous gene. Such knockdowns are often incomplete
(Kennerdell and Carthew,
2000
). Nonetheless, this approach provides a powerful way of
analyzing genes for which no mutants are available.
Combinations and variations
Given the utility of the FRT and the Gal4 UAS systems, it was almost inevitable that they would end up being united. This has been achieved in several ways.
FLPout-Gal4
Previously, researchers wanting to drive the mosaic misexpression of a gene
had to choose between building a FLPout or a UAS construct. This is no longer
the case, as several FLPout lines have now been developed that express high
levels of Gal4 (de Celis and Bray,
1997; Ito et al.,
1997
; Pignoni and Zipursky,
1997
). Thus, a researcher can simply build a UAS-gene
construct, and then drive the expression of that gene in FLPout clones
expressing Gal4, or in the regions defined by any other Gal4 driver. Many of
the FLPout-Gal4 chromosomes also carry UAS-GFP or a lacZ marker,
providing a positive marker for the clones.
UAS-FLPase
FRT-based mitotic recombination or FLPouts can be induced in spatially
restricted domains in the fly by crossing flies that express Gal4 in a
restricted manner to those that carry a UASFLPase construct
(Duffy et al., 1998).
Interestingly, this provides a way of tracing the history of a gene's
expression patterns during development
(Weigmann and Cohen, 1999
)
(Fig. 6). In this approach, a
given Gal4 line is used to drive the expression of FLPase under the control of
UAS. These flies also contain a FLPout-lacZ construct, and the
FLPase-induced recombination of this construct causes the irreversible
expression of lacZ. As a result, lacZ is expressed in the
descendents of all of the cells that ever expressed Gal4, even if they are no
longer expressing Gal4. The only difficulty here is telling whether some
unexpected pattern of lacZ expression is due to previously undetected
gene expression, or due to the leakiness or inaccuracy of the Gal4 driver. For
example, the patched Gal4 enhancer trap (ptcGal4)
is thought to mirror accurately the anterior compartment-specific expression
of the endogenous ptc gene (Hinz
et al., 1994
; Speicher et al.,
1994
) (Fig. 5D).
However, when ptcGal4 is used to drive UAS-FLPase and a
FLPout-construct, FLPout clones are often found in the `wrong' (posterior)
compartment (Fig. 6B).
|
Gal80
The Gal4 inhibitor Gal80 provides yet another way of controlling the Gal4
UAS system (Lee and Luo,
1999). When Gal80 expression is driven with a tubulin promoter
(tub-Gal80), it can inhibit the activity of a tubulin promoter-Gal4 construct.
If Gal80 is removed (see below), Gal4 is disinhibited and drives the
expression of a UAS construct.
In the MARCM (mosaic analysis with a repressible cell marker) technique,
the tub-Gal80 is removed using FRT-mediated mitotic recombination
(Lee and Luo, 1999)
(Fig. 7). The advantage of this
technique over FLPouts is that it simultaneously generates a mitotic
recombinant clone. This can be used, for example, to generate a clone of
homozygous mutant neurons that also express a membrane-associated GFP, thus
marking the mutant axons; in fact, this technique can be used in all cases
where one needs to positively mark homozygous mutant clones. Moreover, this
technique can also be used to generate clones that are not only homozygous for
a given mutation, but also simultaneously express any chosen UAS
construct.
|
Therefore, several laboratories have developed ways of regulating the Gal4
UAS system by building hormone or drug sensitivity into the Gal4 or UAS, or
even adding a FLPout cassette after the UAS sequence (reviewed by
Duffy, 2002). However, all of
these techniques require using novel Gal4 or UAS constructs, and thus cannot
be used with the lines that were previously generated. A new technique that
solves this problem uses a temperature-sensitive form of Gal80 (Gal80ts) (R.
Davis, personal communication). Fly lines carrying Gal80ts under the control
of a tubulin promoter have been generated and crossed to lines containing Gal4
and UAS constructs. At low permissive temperatures, the Gal80ts blocks the
effectiveness of Gal4, while at higher restrictive temperatures, it fails to
inhibit Gal4.
Genetic screens
Mitotic recombination can be used to make not only known mutations
homozygous, but also mutagenized chromosomes. An advantage of this technique
is that the screening can be done in the first (F1) generation, reducing the
labor involved. In such a screen, mitotic recombination is used to generate
homozygous cells in developing F1 flies that are each heterozygous for a
different mutagenized chromosome. F1 adults that have clones with interesting
phenotypes are recovered and a stock established. This technique was used even
before the advent of FRT-based recombination, but the high rate of
recombination that can be achieved using FRTs makes the technique much more
efficient (Xu and Rubin,
1993). In the developing wing disc, for example, the appropriate
induction of hs-FLPase can generate several clones in every wing blade of
every F1 adult. Using UAS-FLPase and an appropriate Gal4 driver can further
increase the rate of recombination and direct it to a particular tissue or
subregion. Variations on this technique have been used to isolate several
mutations that affect cell differentiation and growth patterns in the adult,
such as slimb (which encodes a ubiquitin ligase that regulates
Hedgehog and Wingless/Wnt signaling)
(Jiang and Struhl, 1998
;
Theodosiou et al., 1998
) and
warts (which encodes a Lats family tumor suppressor)
(Justice et al., 1995
;
Xu et al., 1995
).
The use of FRT-based mitotic recombination in mosaic screens does have a
few problems, however. First, the FRT and FLPase constructs are all inserted
via P elements. This means that P-element-based insertional
mutagenesis cannot be used in these flies without remobilizing the constructs.
Therefore, most screens to date have used chemical mutagens or irradiation to
generate mutations, which greatly slows down the subsequent identification and
molecular analysis of the mutations. In the future, it should be possible to
solve this problem by using other types of transposable elements for
mutagenesis, especially those, like the piggyBac element, whose rate
of mobilization in Drosophila can be as high as that of P
elements (Horn et al., 2002;
Häcker et al., 2003
).
Second, if chemical mutagens are used, they cause germ line mosaicism in the
F1 flies; even though a particular mutagenized chromosome is present in the
somatic cells, it might be lost in the germ line. Finally, the F1 adults must
be kept alive, and this limits the types of phenotypes that can be screened.
Thus, some laboratories have used mitotic recombination in F2 screens on
individual mutant stocks.
A very different use of mosaics is to screen for novel genes whose
misexpression causes interesting phenotypes. This is done using any
appropriate Gal4 driver, including FLPout-Gal4 for mixexpression in clones,
and mating these to different EP lines
(Rorth et al., 1998). As noted
above, EP constructs contain multiple UAS sequences, and these constructs have
been inserted randomly around the genome. In the presence of Gal4, an EP
insertion may drive the expression of a neighboring gene. After screening,
those EP insertions that generate interesting phenotypes are mapped, and
neighboring genes are tested for their developmental functions.
The future of mosaic techniques
New variations to and combinations of these techniques are constantly being developed and being made available to the Drosophila community. Given the ever-increasing power of these methods, anyone wanting more could with some justification be accused of ingratitude.
Nonetheless, development is an extremely complex and, in some cases, an
annoyingly redundant process, and it is as yet difficult to manipulate more
than a few genes at once using mosaic techniques in flies. A traditional way
around the problem of redundancy and complexity in Drosophila is to
screen for mutations that act as enhancers and suppressors of mutant
phenotypes, and these techniques can be applied to mosaics (e.g.
Parker et al., 2002). Still,
in the age of genomics, one cannot also help being envious of the ease with
which mixtures of mRNAs or dsRNAs can be used to disrupt embryogenesis in
Drosophila and other organisms. It is not yet clear how complex
mixtures of constructs could be delivered to late-developing tissues in
Drosophila. Solving that problem could open up entirely new frontiers
for mosaic analyses.
ACKNOWLEDGMENTS
The author's research is supported by grants from NIH and NSF.
REFERENCES
Ashburner, M. (1989). Drosophila, A Laboratory Handbook. Plainview: Cold Spring Harbor Laboratory Press.
Blair, S. S. (1992). engrailed expression in the anterior lineage compartment of the developing wing blade of Drosophila. Development 115, 21-34.[Abstract]
Bossing, T. and Technau, G. M. (1994). The fate
of the CNS midline progenitors in Drosophila as revealed by a new
method for single cell labelling. Development
120,1895
-1906.
Brand, A. H. and Perrimon, N. (1993). Targeted
gene expression as a means of altering cell fates and generating dominant
phenotypes. Development
118,401
-415.
Calleja, M., Moreno, E., Pelaz, S., and Morata, G.
(1996). Visualization of gene expression in living adult
Drosophila. Science
274,252
-255.
Chou, T. B. and Perrimon, N. (1992). Use of a
yeast site-specific recombinase to produce female germline chimeras in
Drosophila. Genetics
131,643
-653.
Chou, T. B. and Perrimon, N. (1996). The
autosomal FLP-DFS technique for generating germline mosaics in Drosophila
melanogaster. Genetics
144,1673
-1679.
Chou, T. B., Noll, E. and Perrimon, N. (1993).
Autosomal P[ovoD1] dominant female-sterile insertions in
Drosophila and their use in generating germ-line chimeras.
Development 119,1359
-1369.
Conley, C. A., Silburn, R., Singer, M. A., Ralston, A.,
Rohwer-Nutter, D., Olson, D. J., Gelbart, W. and Blair, S. S.
(2000). Crossveinless 2 contains cysteine-rich domains and is
required for high levels of BMP-like activity during the formation of the
cross veins in Drosophila. Development
127,3947
-3959.
de Celis, J. F. and Bray, S. (1997). Feed-back
mechanisms affecting Notch activation at the dorsoventral boundary in the
Drosophila wing. Development
124,3241
-3251.
Duffy, J. B. (2002). GAL4 system in Drosophila: a fly geneticist's Swiss army knife. Genesis 34,1 -15.[CrossRef][Medline]
Duffy, J. B., Harrison, D. A. and Perrimon, N.
(1998). Identifying loci required for follicular patterning using
directed mosaics. Development
125,2263
-2271.
Ephrussi, B. and Beadle, G. W. (1936). A technique for transplantation in Drosophila. Am. Nat. 70,218 -225.[CrossRef]
Fortier, E. and Belote, J. M. (2000). Temperature-dependent gene silencing by an expressed inverted repeat in Drosophila. Genesis 26,240 -244.[CrossRef][Medline]
Garcia-Bellido, A., Morata, G. and Ripoll, P. (1973). Developmental compartmentalization of the wing disk of Drosophila. Nature New Biol. 245,251 -253.[Medline]
Garcia-Bellido, A., Ripoll, P. and Morata, G. (1976). Developmental compartmentalization in the dorsal mesothoracic disc of Drosophila. Dev. Biol. 48,132 -147.[Medline]
Geyer-Duszynska, I. (1967). Experiments on nuclear transplantation in Drosophila melanogaster. Preliminary report. Rev. Suisse Zool. 74,614 -615.[Medline]
Golic, K. G. (1991). Site-specific recombination between homologous chromosomes in Drosophila. Science 252,958 -961.[Medline]
Golic, K. G. and Lindquist, S. (1989). The FLP recombinase of yeast catalyzes site-specific recombination in the Drosophila genome. Cell 59,499 -509.[Medline]
Häcker, U., Nystedt, S., Padash Barmchi, M., Horn, S. and
Wimmer, E. A. (2003). piggyBac-based insertional mutagenesis
in the presence of stably integrated P elements in
Drosophila. Proc. Natl Acad. Sci. USA
100,7720
-7725.
Hassan, B. A., Bermingham, N. A., He, Y., Sun, Y., Jan, Y. N., Zoghbi, H. Y. and Bellen, H. J. (2000). atonal regulates neurite arborization but does not act as a proneural gene in the Drosophila brain. Neuron 25,549 -561.[Medline]
Heitzler, P. and Simpson, P. (1991). The choice of cell fate in the epidermis of Drosophila. Cell 64,1083 -1092.[Medline]
Hepker, J., Wang, Q. T., Motzny, C. K., Holmgren, R. and Orenic,
T. V. (1997). Drosophila cubitus interruptus forms a
negative feedback loop with patched and regulates expression of
Hedgehog target genes. Development
124,549
-558.
Hinz, U., Giebel, B. and Campos-Ortega, J. A. (1994). The basic-helix-loop-helix domain of Drosophila lethal of scute protein is sufficient for proneural function and activates neurogenic genes. Cell 76, 77-87.[Medline]
Horn, C., Offen, N., Nystedt, S., Hacker, U. and Wimmer, E. A. (2002). piggyBac-based insertional mutagenesis and enhancer detection as a tool for functional insect genomics. Genetics 163,647 -661.
Illmensee, K. (1968). Transplantation of embryonic nuclei into unfertilized eggs of Drosophila melanogaster. Nature 219,1268 -1269.[Medline]
Illmensee, K. (1973). The potentialities of transplanted early gastrula nuclei of Drosophila melanogaster. Production of their imago descendants by germline transplantation. Roux Arch. EntwMech. Organ. 171,331 -343.
Ito, K., Awano, W., Suzuki, K., Hiromi, Y. and Yamamoto, D.
(1997). The Drosophila mushroom body is a quadruple
structure of clonal units each of which contains a virtually identical set of
neurones and glial cells. Development
124,761
-771.
Jiang, J. and Struhl, G. (1998). Regulation of the Hedgehog and Wingless signalling pathways by the F-box/WD40-repeat protein Slimb. Nature 391,493 -496.[CrossRef][Medline]
Justice, R. W., Zilian, O., Woods, D. F., Noll, M. and Bryant, P. J. (1995). The Drosophila tumor suppressor gene warts encodes a homolog of human myotonic dystrophy kinase and is required for the control of cell shape and proliferation. Genes Dev. 9,534 -546.[Abstract]
Kennerdell, J. R. and Carthew, R. W. (1998). Use of dsRNA-mediated genetic interference to demonstrate that frizzled and frizzled 2 act in the Wingless pathway. Cell 95,1017 -1026.[Medline]
Kennerdell, J. R. and Carthew, R. W. (2000). Heritable gene silencing in Drosophila using double-stranded RNA. Nat. Biotechnol. 18,896 -898.[CrossRef][Medline]
Lawrence, P. A. (1981). A general cell marker for clonal analysis of Drosophila development. J. Embryol. Exp. Morphol. 64,321 -332.[Medline]
Lawrence, P. A., Johnston, P. and Morata, G. (1986). Methods of marking cells. In Drosophila, A Practical Approach (ed. D.B. Roberts), pp.229 -242. Oxford: IRL Press.
Lee, T. and Luo, L. (1999). Mosaic analysis with a repressible neurotechnique cell marker for studies of gene function in neuronal morphogenesis. Neuron 22,451 -461.[Medline]
Moreno, E., Basler, K. and Morata, G. (2002). Cells compete for decapentaplegic survival factor to prevent apoptosis in Drosophila wing development. Nature 416,755 -759.[CrossRef][Medline]
O'Kane, C. and Gehring, W. J. (1987). Detection in situ of genomic regulatory elements in Drosophila. Proc. Natl. Acad. Sci. USA 84,9123 -9127.[Abstract]
Parker, D. S., Jemison, J. and Cadigan, K. M.
(2002). Pygopus, a nuclear PHD-finger protein required for
Wingless signaling in Drosophila. Development
129,2565
-2576.
Patterson, J. T. (1929). The production of mutations in somatic cells of Drosophila melanogaster by means of X-rays. J. Exp. Zool. 53,327 -372.
Piccin, A., Salameh, A., Benna, C., Sandrelli, F., Mazzotta, G., Zordan, M., Rosato, E., Kyriacou, C. P. and Costa, R. (2001). Efficient and heritable functional knock-out of an adult phenotype in Drosophila using a GAL4-driven hairpin RNA incorporating a heterologous spacer. Nucleic Acids Res. 29, E55.[Medline]
Pignoni, F. and Zipursky, S. L. (1997).
Induction of Drosophila eye development by decapentaplegic.
Development 124,271
-278.
Robertson, H. M., Preston, C. R., Phillis, R. W.,
Johnson-Schlitz, D. M., Benz, W. K. and Engels, W. R. (1988).
A stable genomic source of P-element transposase in Drosophila
melanogaster. Genetics
118,461
-470.
Rorth, P. (1996). A modular misexpression
screen in Drosophila detecting tissue-specific phenotypes.
Proc. Natl. Acad. Sci. USA
93,12418
-12422.
Rorth, P., Szabo, K., Bailey, A., Laverty, T., Rehm, J., Rubin,
G., Weigmann, K., Milan, M., Benes, V., Ansorge, W. and Cohen, S.
(1998). Systematic gain-of-function genetics in
Drosophila. Development
125,1049
-1057.
Rubin, G. M. and Spradling, A. C. (1982). Genetic transformation of Drosophila with transposable element vectors. Science 218,348 -353.[Medline]
Santamaria, P. (1986). Injecting eggs. inDrosophila, a practical approach (ed. D.B. Roberts), pp. 159-173. IRL Press Ltd., Oxford.
Simcox, A. (1997). Differential requirement for EGF-like ligands in Drosophila wing development. Mech. Dev. 62,41 -50.[CrossRef][Medline]
Speicher, S. A., Thomas, U., Hinz, U. and Knust, E.
(1994). The Serrate locus of Drosophila and its
role in morphogenesis of the wing imaginal discs: control of cell
proliferation. Development
120,535
-544.
Spradling, A. C. and Rubin, G. M. (1982). Transposition of cloned P-elements into Drosophila germ line chromosomes. Science 218,341 -347.[Medline]
Staehling-Hampton, K., Jackson, P. D., Clark, M. J., Brand, A. H. and Hoffmann, F. M. (1994). Specificity of bone morphogenetic protein-related factors: Cell fate and gene expression changes in Drosophila embryos induced by decapentaplegic but not 60A. Cell Growth Differ. 5, 585-593.[Abstract]
Stern, C. (1936). Somatic crossing over and
segregation in Drosophila melanogaster.
Genetics 21,625
-730.
Struhl, G. and Basler, K. (1993). Organizing activity of wingless protein in Drosophila. Cell 72,527 -540.[Medline]
Technau, G. M. (1986). Lineage analysis of transplanted individual cells in embryos of Drosophila melanogaster. Roux Arch. Dev. Biol. 195,389 -398.
Theodosiou, N. A., Zhang, S., Wang, W. Y. and Xu, T.
(1998). slimb coordinates wg and dpp
expression in the dorsal-ventral and anterior-posterior axes during limb
development. Development
125,3411
-3416.
Vincent, J. P. and O'Farrell, P. H. (1992). The state of engrailed expression is not clonally transmitted during early Drosophila development. Cell 68,923 -931.[Medline]
Weigmann, K. and Cohen, S. M. (1999).
Lineage-tracing cells born in different domains along the PD axis of the
developing Drosophila leg. Development
126,3823
-3830.
Xu, T. and Rubin, G. M. (1993). Analysis of
genetic mosaics in developing and adult Drosophila tissues.
Development 117,1223
-1237.
Xu, T., Wang, W., Zhang, S., Stewart, R. A. and Yu, W.
(1995). Identifying tumor suppressors in genetic mosaics: the
Drosophila lats gene encodes a putative protein kinase.
Development 121,1053
-1063.