thesis 4pdf

thesis 4pdf
Ensconsin, a Par-1 regulated
microtubule associated protein,
regulates kinesin dependent
transport
Hsin-Ho Sung
2007
Dissertation
submitted to the
Combined Faculties for the Natural Sciences and for Mathematics
of the Ruperto-Carola University of Heidelberg, Germany
for the degree of
Doctor of Natural Sciences
presented by
Master of Science:
Hsin-Ho Sung
Born in Kaohsiung, Taiwan
Oral-examination: 10.07.2007
Ensconsin, a Par-1 regulated
microtubule associated protein,
regulates kinesin dependent
transport
Referees:
Dr. Anne Ephrussi
Dr. Anne Regnier-Vigouroux
僅此獻給
不辭辛苦的母親和在天之靈的父親
Summary
Maternal proteins and mRNA contribute to both oocyte and embryo development. I
performed a genetic screen to identify genes with maternal function in Drosophila. From
this screen, I isolated PB4170, which affects CG14998 (ensconsin). The human
homologue of this protein, E-MAP-115/Ensconsin, is known to be a microtubule binding
protein that interacts dynamically with microtubules. However, its molecular function is
poorly understood. From ensconsin mutant analysis in Drosophila, I have found that this
gene is specifically required for microtubule-dependent polarity in the oocyte. My results
suggest that Ensconsin does not directly affect the stability or orientation of microtubules,
but acts through Kinesin, a plus end directed motor that moves cargos along microtubules.
I could also demonstrate that Ensconsin is a target of the Par-1 kinase, which has been
shown to be required for the establishment of oocyte polarity in Drosophila. In mammals,
the Par-1 homolog MARK destabilises microtubules through phosphorylation of
microtubule associated proteins (MAPs).In Drosophila, Par-1 directly affects microtubule
stability via unkown MAPs, as the MAPs identified so far do not show any polarity
defects in the oocyte. As ensconsin mutants do show an oocyte polarity phenotype, this
strongly suggests that in Drosophila Par-1 kinase controls oocyte polarity also through the
regulation of Kinesin-based transport by phosphorylation of Ensconsin.
5
Zusammenfassung
Maternale Transkripte und Proteine steuern zur Entwicklung der Eizelle und des Embryos
bei. Um Gene mit einer maternalen Funktion in Drosophila zu identifizieren führte ich
einen genetischen Screen durch. Dabei isolierte ich PB4170, eine Mutante für das Gen
CG14998 (ensconsin). Es ist bekannt, dass das menschliche Homolog dieses Gens,
E-MAP-115/Ensconsin, Mikrotubuli binden und dynamisch mit ihnen interagieren kann.
Die genaue Funktion von Esconsin ist jedoch nicht geklärt. Durch die Analyse der
ensconsin Mutanten fand ich heraus, dass Ensconsin spezifisch für die Polarität der
Oozyte, welche von den Mikrotubuli abhängig ist, benötigt wird. Meine Resultate deuten
darauf hin, dass Ensconsin nicht direkt die Stabilität oder die Orientierung der
Mikrotubuli beeinflusst, sondern spezifisch für die Funktion von Kinesin, einem
Motorprotein welches sich den Mikrotubuli entlang bewegt, gebraucht wird.
Im weiteren konnte ich zeigen, dass Ensconsin durch die Par-1 Kinase, einem Enzym,
welches für die Polarität der Oozyte und der Follikelzellen benötigt wird, phosphoryliert
wird. In Säugetieren destabilisert das Par-1 homolog MARK Mikrotubuli durch
Phosphorylierung von Mikrotubuli-Bindungsproteine (MAPs). Par-1 beeinflusst auch in
Drosophila die Stabilität der Mikrotubuli, doch ist in den Mutanten der bisher bekannten
MAPs die Polarität der Oozyte normal. Da in ensconsin Mutanten die Polarität der
Oozyte beeinträchtigt ist, deuten meine Resultate darauf hin, dass die Par-1 Kinase diese
Polarität durch die Regulation von Kinesin-basiertem Transport über die
Phosphorylierung von Ensconsin kontrolliert.
6
Acknowledgements
When I arrived at EMBL, I was very frustrated and had no idea of future plans for my life.
But now I will finish my Phd soon. I feel that this is a dream. All I want to thank is
Pernille Rørth. She gave me a new life. Thanks for taking care of a stupid and
poor-English-speaking guy, like me. Thanks for teaching me a lot of things in science.
The most important thing that I learned from her is never give up until you get the results,
if you think it is reasonable and worth doing.
Thanks
for
my
thesis
committe
members,
Dr.
Anne
Ephrussi,
Dr.
Anne
Régnier-Vigouroux, Dr. Jurg Muller, and Prof. Dr. Herbert Steinbeisser for good advices
for my experiments. Especially for Dr. Anne Ephrussi, without your support and
suggestion, I cannot finish this work. Thanks for Dr. Anne Ephrussi and Dr. Anne
Régnier-Vigouroux who will read and grade my thesis.
I also want to thanks all previous and present Rørth members: Gaspar, Tudor, Simone,
Luis, Carlos, Anne, Kalman, Andreea, Jan, Lodo, Georgina, Juliette, Celine, Ambra, Oguz,
Katrien, Minna, Adam, Smitha, Nachen, and Issac for discussion in science and for
general support. You are like brothers and sisters to me. I want to thank Gaspar Jekely
particularly. It was great collaborating with him on the cbl project. I want to thank Juliette
Mathieu and Barry Thompson for their works in the genetic screen, especially to Juliette
Mathieu. Without her leading and company, I think I wouldn’t have been able to finish the
screen. Thanks Celine Pugieux for teaching me protein expression. Thanks to Adam
Cliffe, Minna Poukkula, Smitha Vishnu, and Jishy Varghese for reading my thesis. I want
to thank Adam for patiently reading my thesis, and experimental and life support.
I also want to thanks all nice members of the fly community. Thanks to Piyi Papadaki for
kindly sharing her unpublished method, the in vitro kinase assay. Thanks to Anna
Cyrklaff and Natascha Bushati for providing probes and anitbodies. Thanks Lukas
Neidhart for tanslating the thesis summary into german version. Thanks to Juliette, Lodo,
Lukas, and Thelma for hosting me and Yawen during holidays. Thanks to Li-Jung,
Carsten, Felix and Shih-Jung for supporting my life in Heidelberg.
7
謝謝爸爸媽媽的支持和鼓勵,雖然爸等不到我的畢業,但我想他一直都在保佑我。也希望
媽媽可以輕鬆一點,不要太累。謝謝姐姐、姐夫、大哥、二哥和嫂子們的支持,希望你們
身體都健康。最後謝謝雅雯在這五年中的支持和協助,陪我度過許許多多的事,希望未來
的路上我們一路相伴。
8
Table of Contents
Summary
5
Zusammenfassung
6
Acknowledgements
7
Table of Contents
9
List of Figures and Tables
1
13
INTRODUCTION
14
1.1
Cell polarity
15
1.2
Drosophila early oogenesis
16
1.2.1
The germarium development
16
1.2.2
Oocyte specification
17
1.2.3
Microtubules are involved in the oocyte specification
18
1.2.4
Microtubules in early stages of the egg chamber
18
1.3
Mid-oogenesis
1.3.1
Establishment of anterior-posterior axis of the oocyte
20
1.3.2
Patterning of the dorsal-ventral axis
20
1.3.3
Localization of the embryo polarity determinants
22
1.3.3.1
bicoid mRNA localization
22
1.3.3.2
oskar mRNA localization
23
1.3.4
Kinesin patterns the Drosophila oocyte
24
1.4
late oogenesis
26
1.4.1
Ooplasmic streaming
26
1.4.2
The anterior-posterior axis of the embryo
26
1.5
Par-1 function in oogenesis
28
1.5.1
Par-1 affects microtubule-dependent polarity in the oocyte
28
1.5.2
Par-1/MARK regulate microtubules through MAPs
29
1.5.3
E-MAP-115 (Ensconsin), another MAP
29
1.6
2
20
The aim of this thesis
31
MATERIALS AND METHODS
32
9
Fly genetics
2.1
33
2.1.1
Fly husbandry
33
2.1.2
Fly strains
33
2.1.3
Ectopic expression using the GAL4/UAS system
34
2.1.4
Generation of mosaic clones using the FLP/FRT system
35
2.1.5
Generation of Germ line clones using the FLP-DFS system
36
2.2
Mutant analysis in the Screen
38
2.2.1
X-gal staining in the Drosophila egg chamber
38
2.2.2
Cuticle preparation
38
2.2.3
Inverse PCR
38
2.3
ensconsin mutant analysis
40
2.3.1
RNA isolation
41
2.3.2
First-strand cDNA synthesis
42
2.3.3
Microtubule binding assay
42
2.3.4
Western blotting
43
2.3.5
Single Fly PCR
43
2.3.6
Purification of Ensconsin protein
44
2.3.7
In vitro kinase assay
45
2.3.8
The ensconsin Probe preparation
46
2.3.9
In situ hybridization of egg chambers
46
2.3.10
In situ hybridization of embryos
47
2.3.11
Immunofluorescence staining of larvae axons
48
2.3.12
Immunofluorescence staining of embryos
48
2.3.13
Immunofluorescence staining of egg chambers
49
2.3.14
Dhc immunostaining of egg chambers
49
2.3.15
Khc immunostaining of Drosophila egg chambers
50
2.3.16
Live image of tau-GFP in the ovoD1 germ line clone
50
2.3.17
Live image for ooplasmic streaming in the egg chamber
50
2.3.18
Climbing assay
51
2.4
3
Websites
52
RESULTS
3.1
53
Screen for genes affecting border cell migration, oogenesis and embryogenesis
54
3.1.1
Screening method
54
3.1.1.1
Generation of PiggyBac mutants in FRT background
54
3.1.1.2
Mapping
54
3.1.1.3
Germ line clone analysis
55
10
3.1.1.4
Cloning the genes affected by the PiggyBac insertions
55
3.1.1.5
Creation of new ovoD1 lines
55
3.2
Screen result
57
3.2.1
No maternal effect
57
3.2.2
Germ cell lethal
57
3.2.3
Abnormal oogenesis
58
3.2.3.1
Nurse cell membrane defects
58
3.2.3.2
Gurken signaling defects
58
3.2.3.3
Dumpless eggs
59
3.2.3.4
Abnormal nurse cells
60
3.2.3.5
Others
61
3.2.4
Mutations affecting embryo development
61
3.2.4.1
Posterior group
61
3.2.4.2
Segmentation defects
62
3.2.5
Others
65
3.3
Ensconsin
66
3.3.1
PB4170 affects Ensconsin
66
3.3.2
N-terminal region of Ensconsin can bind to microtubules
66
3.3.3
20% of PB4170 egg chambers showed Oskar and Staufen mislocalization
68
3.3.4
Generating new alleles of ensconsin
69
∆C
3.3.5
Oskar, Staufen and Gurken localization are affected but bicoid is not in ens
3.3.6
khc interacts genetically with ensconsin
3.3.7
Ensconsin affects posterior localization of Khc
mutant
70
73
∆C
73
3.3.8
Microtubule polarity is not affected in ens
3.3.9
Ooplasmic streaming is affected in ensconsin mutants
75
3.3.10
Ensconsin is required in neurons
76
3.3.11
Ensconsin is localized to the anterior of the oocyte
78
3.3.12
Ensconsin localization is affected by Par-1
79
3.3.13
Ensconsin is a direct target of Par-1
80
∆C
mutant
74
3.3.14
Follicle cell polarity is not affected in ens
mutant
82
3.3.15
The Par-1 phosphorylation 14-3-3 binding sites in Ensconsin are essential for its localization
83
4
DISCUSSION
4.1
85
The genetic screen using PiggyBac Transposons
86
4.1.1
The PiggyBac screen is effective but laborious
86
4.1.2
The PiggyBac and P element have similar preferential sites
86
4.1.3
Germline clone analysis
87
11
Ensconsin: a microtubule associated protein
4.2
88
4.2.1
Ensconsin affects microtubules in a subtle way
88
4.2.2
Ensconsin affects Khc-dependent transport
88
4.2.3
Ensconsin is regulated by Par-1 phosphorylation
88
4.2.4
Par-1 regulates motor protein transport through MAPs
89
4.2.5
How does Ensconsin affect Khc-dependent transport?
90
5
REFERENCES
6
APPENDIX
93
102
6.1
Screen result
103
6.2
Publications
106
12
List of Figures and Tables
Figure 1. Drosophila ovary.................................................................................................................... 16
Figure 2. The structure of Drosophila germarium................................................................................ 17
Figure 3. Patterning of the oocyte......................................................................................................... 21
Figure 4. Localization of anterior and posterior determinants in the egg............................................ 27
Table 1. Fly strains used for experiments............................................................................................. 33
Figure 5. GAL4-UAS system................................................................................................................. 34
Figure 6. FLP-FRT system.................................................................................................................... 35
Figure 7. FLP-DFS system.................................................................................................................... 36
Figure 8. Ends-out system..................................................................................................................... 41
Figure 9. The scheme of crosses ............................................................................................................ 56
Figure 10. Nurse cell membrane defects............................................................................................... 58
Figure 11. Gurken signaling defects...................................................................................................... 59
Figure 12. Dumpless phenotype ............................................................................................................ 60
Figure 13. Abnormal nurse cells ........................................................................................................... 60
Figure 14. Oskar localization defects.................................................................................................... 61
Figure 15. Posterior group phenotype .................................................................................................. 62
Figure 16. Segmentation defect............................................................................................................. 63
Figure 17. PB2691 mutant phenotype................................................................................................... 65
Figure 18. Others .................................................................................................................................. 65
Figure 19. PB4170 embryo phenotypes and genomic structure ........................................................... 66
Figure 20. N-terminal of Ensconsin can bind microtubules ................................................................. 67
Figure 21. Oskar, Staufen and Gurken are mislocalized in PB4170..................................................... 69
Figure 22. New mutant alleles of ensconsin .......................................................................................... 70
Figure 23. ens∆C mutant phenotype....................................................................................................... 71
Figure 24. Ensconsin affects Dhc localization through Khc mislocalization ........................................ 73
Figure 25. Microtubule formation and microtubule polarity in ens∆C mutant..................................... 74
Figure 26. Ooplasmic streaming is abolished in ens mutants............................................................... 76
Figure 27. Neuronal functions of Ensconsin ......................................................................................... 77
Figure 28. Drosophila Ensconsin is tightly localized during oogenesis ................................................ 78
Figure 29. Ensconsin localization is restricted by Par-1....................................................................... 80
Figure 30. Ensconsin is a direct target of Par-1.................................................................................... 81
Figure 31. Ensconsin does not affect follicle cells polarity.................................................................... 82
Figure 32. Ensconsin is regulated by Par-1 and controls proper posterior marker localization.......... 83
Figure 32. The model for Ensconsin function. ...................................................................................... 91
Table 2. Summary of PiggyBac insertions...........................................................................................103
13
1 INTRODUCTION
14
INTRODUCTION
1.1
Cell polarity
Multicellular organisms are composed of different types of tissues which are derived from
a single fertilized egg. How does a single cell develop into a complex organism?
Embryonic cells and stem cells use their polarity to generate different cell types by
asymmetric cell division. Furthermore, differentiated cells also perform specific functions
by using their polarity. For example, fibroblast cells form an actin-rich leading edge
which aids in their migration. Neuronal cells form distinct axonal and dendritic
compartments which are important for directional signaling. And epithelial cells have
apical-basolateral
domains
necessary
for
maintaining
tissue
impermeability.
(Gonzalez-Reyes, 2003; Siegrist and Doe, 2005; Spradling et al., 2001; Suzuki and Ohno,
2006)
What is cell polarity? Cell polarity has been studied in organisms ranging from
bacteria, yeast, worms, and flies to mammals. Cell polarity is the asymmetric distribution
of proteins that generates an asymmetry of cellular function. One way that asymmetric
protein distribution is accomplished is by asymmetric mRNA localization. The factors
involved in asymmetric localization of mRNA from yeast to Drosophila do not belong to
one common family. Even though diverse molecular mechanisms can control mRNA
localization, the most common method is the transport of mRNAs along cytoskeleton
tracks. In this study, I will focus on the Drosophila oocyte, a relatively large cell with
asymmetric mRNA localization (oskar, gurken and bicoid). It provides a good system for
investigating cell polarity formation (Jan and Jan, 2001; Johnstone and Lasko, 2001;
Nusslein-Volhard and Roth, 1989; Riechmann and Ephrussi, 2001; St Johnston, 2005).
15
INTRODUCTION
1.2
Drosophila early oogenesis
Drosophila females have one pair of ovaries. Each ovary consists of 16-20 ovarioles,
which can be subdivided into the anterior germarium and posterior vitellarium. The egg
chamber is formed in the anterior germarium and matures in the posterior vitellarium
(Figure 1). Thirteen different stages of egg chambers (stages 2-14) can be found in the
vitellarium. The functional unit of oogenesis is the egg chamber which contains 15 nurse
cells and an oocyte. The germ cells are surrounded by a monolayer of follicle cells
(Figure 1). The oocyte and nurse cells are interconnected via cytoplasmic bridges called
ring canals.
Figure 1. Drosophila ovary
The Drosophila ovary (top left) is surrounded by a thin peritoneal sheath (red). Each ovariole
(middle) is surrounded by a layer of muscle (blue). Egg chambers arise in the germarium and
mature in the vitellarium. Each egg chamber contains 15 nurse cells and one oocyte. The
germ cells are surrounded by a monolayer of follicle cells (green). Adapted and edited from
(Frydman et al., 2006)
1.2.1
The germarium development
The germarium is divided into four stages: stage 1, 2a, 2b and 3. The stage 1 area of the
germarium contains two germ cells, associated with a stack of around 10 terminal
filament cells. Terminal filament cells contribute to ovariole development and support
stem cell activity (Figure 2) (King et al., 2001; Lin and Spradling, 1993; Sahut-Barnola et
al., 1995). The germ line stem cell divides asymmetrically, forming a new stem cell and a
cystoblast. The new stem cell remains attached to the neighboring somatic cap cells, at
16
INTRODUCTION
the anterior (Gonzalez-Reyes, 2003; Spradling et al., 2001). The cystoblast undergoes
four rounds of division with incomplete cytokinesis to produce 16 germ cells. During
these divisions the fusome, a vesicle-rich organelle, becomes polarized, with one daughter
cell getting more fusome material than the other during cell division. This asymmetric
division is due to the anchoring of one pole of each spindle to the fusome at every
division. This asymmetric division ensures that the cyst consists of two cells with four
ring canals, two cells with three ring canals, four cells with two canals, and eight cells
with one ring canals. When 16 germ cells are formed, the oogenesis enters region 2a of
germarium (Lin et al., 1994).
Figure 2. The structure of Drosophila germarium
The location of germline stem cells (red cells on left) and somatic stem cells (red cells on top
and bottom). The terminal filament and cap cells (green) retain stem cell activity. The germ
cells and dividing germ cells (yellow) move away from the anterior tip. At the end of
germarium the somatic cells divide and give rise to the follicle cells which then migrate
inwards, surrounding the germ cells. Adapted from (Spradling et al., 2001).
1.2.2
Oocyte specification
At stage 2a, the two cells with four ring canals are selected as pre-oocytes. At stage 2b,
one of them becomes the oocyte, with the other cell becoming a nurse cell. The
determination of the oocyte can be followed with several markers. (1) At stage 2a, the
oocyte specific proteins Bicaudal (BicD), Orb, Barentsz (Btz), Cup, and Egalitarian (Egl)
and mRNAs oskar, BicD, and orb become enriched in the two pre-oocytes. At the end of
stage 2a, these proteins and mRNAs are concentrated in the oocyte (Suter et al., 1989)
(Ephrussi et al., 1991; Keyes and Spradling, 1997; Lantz et al., 1994; van Eeden et al.,
17
INTRODUCTION
2001; Wharton and Struhl, 1989). (2) Initially microtubules are distributed through the 16
germ cells. At the end of stage 2a, their minus ends gradually localize to the selected
oocyte (Bolivar et al., 2001; Grieder et al., 2000). (3) By the end of stage 2a, after the last
mitotic division, the centrosomes appear to become inactived and move into the oocyte
(Cox and Spradling, 2003) (Grieder et al., 2000); (Bolivar et al., 2001). In region 2b, the
germ cells change shape, forming a one cell-thick disc. At the same time, somatic follicle
cells migrate inward and envelop the germ cells. As the germ cells reach stage 3, the
mitochondria, centrosomes, Golgi vesicles, proteins and mRNAs accumulate at the
anterior of the oocyte. The germ cells become rounded and form a sphere, with the oocyte
always staying at the posterior pole. The germ cells leave the germarium and enter the
vitellarium. The oocyte becomes further polarized, as proteins, mRNAs, centrosomes and
a subset of the mitochondria move to the posterior cortex. The oocyte DNA becomes
highly condensed to form a structure called the karyosome, whereas the nurse cells
become polyploid (Dej and Spradling, 1999; Huynh et al., 2001).
1.2.3
Microtubules are involved in the oocyte specification
Microtubules are essential for the specification of the oocyte. Treatment of ovaries with
the microtubule depolymerizing drug, colchicine, results in failure of oocyte specification,
and instead 16 nurse cells are seen (Koch and Spitzer, 1983). Furthermore, the
oocyte-specific proteins and mRNAs are not asymmetrically localized within germ cells
(Theurkauf et al., 1993). Mutations in the microtubule minus-end directed motor, Dynein
heavy chain (Dhc), or in its associated regulator, Lissencephaly (Lis1) affect the
formation of the fusome. Germ cells mutant for these genes show fewer than 16 germ
cells and sometimes lack oocytes (Bolivar et al., 2001; Liu et al., 1999; McGrail and Hays,
1997). This suggests that oocyte-specific proteins and mRNAs are transported using
microtubule network.
1.2.4
Microtubules in early stages of the egg chamber
After the egg chamber leaves the germarium, it enters the vitellarium. The vitellarium is
divided into two stages; the first 6 stages are called the previtellogenic stage and the later
stages vitellogenesis. During the pre-vitellogenic stages, the germ cells grow at almost the
18
INTRODUCTION
same rate. After the pre-vitellogenic stage, vitellogenesis begins and oocyte grows fast,
causing it to increase in size with respect to the nurse cells. In the germarium and the
pre-vitellogenic stages of the vitellarium, the microtubule minus-ends are enriched in the
posterior part of the oocyte and microtubules extend their plus-ends through the ring
canals into the nurse cells (Figure 3A). The oocyte nucleus is largely transcriptionally
inactive throughout oogenesis, as it is arrested in meiotic prophase. At this time, the nurse
cell nuclei start to endo-replicate, becoming highly polyploid (Spradling, 1993). Most of
the RNA and proteins, including grk, oskar and bicoid mRNA, which are required for
oogenesis and embryogenesis are synthesized in the nurse cells and transported into the
oocyte. mRNA and proteins produced in the nurse cells are transported to oocyte on the
microtubule network (Grieder et al., 2000; Pokrywka and Stephenson, 1995; Theurkauf et
al., 1992).
19
INTRODUCTION
1.3
1.3.1
Mid-oogenesis
Establishment of anterior-posterior axis of the oocyte
Establishment of the anterior-posterior (A-P) axis of the oocyte is mediated by gurken,
which encodes a Drosophila TGF-
α homolog, an EGF-receptor ligand. Its expression is
associated with the oocyte nucleus (Nilson and Schupbach, 1999). In previtellogenic
stage egg chambers, the microtubules minus ends are enriched in the posterior pole of the
oocyte, and the oocyte nucleus also localize at posterior pole of the oocyte. mRNA and
proteins which are produced in the nurse cells are transported along microtubules into the
transcriptionally inactive oocyte (Figure 3A) (Spradling, 1993; Theurkauf et al., 1992).
The first wave of Gurken signal is that Gurken produces from perinuclear region at
posterior pole of oocyte and the signal is sent to the terminal follicle cells, specifying
posterior follicle cells fate (stage 6) (Figure 3B) (Gonzalez-Reyes et al., 1995;
Gonzalez-Reyes and St Johnston, 1998; Roth et al., 1995). Then an unknown signal is
sent from the posterior follicle cells to the oocyte resulting in the repolarization of oocyte
microtubule cytoskeleton. In response to this reverse signal, the microtubules minus-ends
at the posterior of the oocyte disassemble and microtubules nucleate from the anterior and
lateral cortex of the oocyte (Figure 3C) (Gonzalez-Reyes et al., 1995; Roth et al., 1995).
This reorganization of the microtubule network is necessary for the oocyte nucleus to
migrate to an anterior position (Figure 3C).
1.3.2
Patterning of the dorsal-ventral axis
The second wave of Grk signaling occurs from the anterior-dorsal corner of the oocyte
and controls dorsal-ventral (D-V) patterning. First, Gurken and Dpp specify the dorsal
follicle cell fate, which will form structures such as the dorsal appendages (Figure 3E, 4)
(Peri and Roth, 2000). Furthermore, Gurken and PVF1 have been identified as guidance
cues for border cells. Border cells, a group of 6-8 follicle cells, delaminate from the
anterior follicle cells and migrate through the nurse cells to the oocyte. When they reach
to the oocyte, they migrate toward to the dorsal part of the oocyte and then form a
micropyle, a hole for sperm entry at the anterior end of the egg. Gurken and PVF1 signal
through their tyrosine kinase receptors (RTKs), EGFR and PVR, respectively.
20
INTRODUCTION
Figure 3. Patterning of the oocyte.
(A) At previtellogenic stage, mRNAs and proteins (red arrows) are produced in the nurse cells
and transported along microtubules into the ooctyte. The oocyte nucleus localizes in the
posterior part of the oocyte, and Gurken is expressed around oocyte nucleus. At stage 6, after
receiving the Grk signal from the oocyte, the terminal follicle cells become posterior follicle cells
fate. (B) After posterior follicle cells formation, posterior follicle cells send an unidentified signal
back to the oocyte at stage 7. (C) The unidentified signal from posterior follicle cells causes the
oocyte microtubules to reorganize. The microtubule minus-ends disassemble and microtubules
21
INTRODUCTION
nucleate at the anterior and lateral cortex. Then the oocyte nucleus moves to the anterior corner
of the oocyte to define the dorsal side of the egg chamber at stage 8. (D) bicoid and oskar
mRNA which are produced in the nurse cells are transported to the oocyte and localized in the
oocyte at stage 9. Microtubules extend from anterior to posterior in the oocyte. bicoid mRNA
accumulates at the anterior cortex of the oocyte. oskar mRNA and Staufen protein are
transported by Kinesin towards the posterior pole. (E) Patterning of the egg by the second Grk
signal at stage 10 egg chamber. Grk protein, localized in the anterior-dorsal corner of the
oocyte, specifies dorsal chorion structures and also guides the migration of border cells, which
are required for the formation of a functional micropyle. Adapted from (Riechmann and
Ephrussi, 2001).
Both RTKs have redundant activity in guiding the anterior-posterior migration of border
cell toward the oocyte, while only the EGFR pathway is essential for dorsal migration
(Figure 3E) (Duchek and Rorth, 2001; Duchek et al., 2001). Formation of the A-P and
D-V axes of the oocyte depends on Gurken signaling. Mutations in gurken or genes
involved in Gurken signaling disrupt oocyte polarity. In these cases, the oocyte nucleus
fails to migrate from the posterior to the anterior-dorsal side and posterior terminal
follicle cells adopt anterior cell fates (Gonzalez-Reyes et al., 1995; Roth et al., 1995).
1.3.3
Localization of the embryo polarity determinants
After the Gurken signaling sets up A-P and D-V axes of the oocyte, A-P and D-V axes
determinants of the embryo start to localize in the oocyte during mid-oogenesis. In
addition to the Gurken signaling specifies the dorsal cell fate of the embryo, the A-P axis
of the embryo also requires for bicoid and oskar mRNA localization in mid-oogenesis
(Figure 3D). bicoid mRNA localizes the anterior pole of the oocyte to determine the
anterior cell fate of embryos, and oskar mRNA localizes to the posterior pole of the
oocyte to define the posterior cell fates of the embryo (Figure 3E, 4) (Ephrussi et al., 1991;
Kim-Ha et al., 1991; Neuman-Silberberg and Schupbach, 1993; Nusslein-Volhard et al.,
1987; Nusslein-Volhard and Roth, 1989). Both mRNAs are produced in the nurse cells
and transferred into the oocyte in mid-oogenesis. After transferring into the oocyte, both
mRNAs are localized and anchored at the poles.
1.3.3.1
bicoid mRNA localization
The early stage of bicoid mRNA localization depends on Exuperantia (Exu), which is also
localized to the anterior of the oocyte. Exu is required in nurse cells to recruit
anterior-targeting factors to the bicoid mRNA. This anterior-targeting complex drives
22
INTRODUCTION
bicoid mRNA to localize at anterior margin of the oocyte (Macdonald et al., 1991; Wang
and Hazelrigg, 1994). bicoid mRNA is seen in a ring at stage 10a and then moves to cover
the whole anterior cortex at stage 10b. From stage 10b onwards, maintaining bicoid
mRNA in the anterior requires Swallow, which is also enriched at the anterior cortex.
Swallow binds Dynein light chain which is a microtubule minus-end directed motor
protein. This suggests that Dynein either anchors or transports bicoid mRNA to the
microtubule minus-end (Berleth et al., 1988; Meng and Stephenson, 2002; Schnorrer et al.,
2000; St Johnston et al., 1989). In addition to Swallow, bicoid mRNA localization also
requires γ-tubulin complex components, γ-tubulin37C and Grip75 that nucleate
microtubules. Thus, the γ-tubulin complex nucleates anterior microtubules, that anchor
bicoid mRNA at right position at stage 10b-11 (Schnorrer et al., 2002). The final stage of
this localization requires Staufen to maintain bicoid mRNA at anterior cortex after stage
11. This process is also microtubule dependent (Ferrandon et al., 1994; St Johnston et al.,
1989).
1.3.3.2
oskar mRNA localization
oskar 3’ untranslated region (3’UTR) is required for the posterior localization of the
mRNA in the oocyte (Kim-Ha et al., 1993). Hrp48 binds to the 5’and 3’UTRs of oskar
mRNA. This represses oskar mRNA translation during its transport and regulates its
localization (Huynh et al., 2004; Yano et al., 2004). Human homologues of Y14 and
Magonashi are core components of the exon-exon junction complex (EJC). They bind to
mRNAs 20-24 nucleotides upstream of exon-exon junctions in a splicing-dependent
manner. oskar mRNA is mislocalized in the Y14/magonashi mutant egg chamber (Hachet
and Ephrussi, 2004). eIF4AIII interacts with Y14 and magonashi and is also suggested to
be involved in oskar mRNA localization (Palacios et al., 2004).
After oskar mRNA is exported from the nurse cell nuclei along with the Hrp48 and EJC
complexes, eIF4AIII recruits Barentsz, a cytoplasmic protein, which also affects oskar
mRNA localization (Palacios et al., 2004; van Eeden et al., 2001). Staufen associates with
oskar mRNA and regulates its translation in the nurse cell cytoplasm (Ephrussi et al.,
1991; Kim-Ha et al., 1991; St Johnston et al., 1991). During oskar mRNA transport,
translational control is also essential for oskar mRNA localization. The translation of
23
INTRODUCTION
oskar mRNA is repressed by Bruno and Hrp48. Bruno recruits Cup protein which binds
to the translation initiation factor eIF4E inhibiting oskar mRNA translation (Chekulaeva
et al., 2006; Kim-Ha et al., 1995; Wilhelm et al., 2003; Yano et al., 2004). This oskar RNP
complex is moved to posterior part of the oocyte by plus-end motor protein on the
microtubules (Cha et al., 2002). Only after oskar mRNA reaches to the posterior pole of
oocyte, it starts to translate Oskar protein at stage 9. As microtubule traffic is important
for the determinants localization, I will focus on one of the microtubule plus end directed
motor, Kinesin.
1.3.4
Kinesin patterns the Drosophila oocyte
Kinesin, a microtubule plus-end directed motor protein, contains 2 Kinesin heavy chains
(Khc) which can form a homodimer and 2 light chains (Klc) (Huang et al., 1994;
Kozielski et al., 1997; Yang et al., 1989). The N-terminal region of Khc contains ATP and
microtubule binding domains (Kull et al., 1996; Yang et al., 1990). Klc binding is
mediated through the C-terminal region of Khc (Cyr et al., 1991; Gauger and Goldstein,
1993; Verhey et al., 1998). The C-terminal portion of Khc has two different functions. (1)
The C-terminal tail (around 60 amino acids) may bind to cargos such as membrane-bound
organelles or vesicles, as well as Klc (Bi et al., 1997; Skoufias et al., 1994). (2) The
C-terminal tail domain also inhibits the ATPase and motor activities of the N-terminus
rendering it inactive. Upon cargo binding, this inhibition is removed and Khc becomes
active (Coy et al., 1999; Hackney et al., 1992). In midoogenesis, oskar mRNA
localization to the posterior pole depends on active transport by motor protein along the
microtubule network. Treatment of egg chambers with colchicine leads to a
mislocalization of oskar mRNA to the oocyte cortex. In khc mutant egg chamber, oskar
mRNA is mislocalized to the oocyte cortex like the phenotypes in the colchicine-treated
egg chamber (Cha et al., 2002). Cha et al. have suggested that during stage 7, before the
reorganizing of the microtubule network, Khc transports oskar mRNA away from cortex
into the middle of the oocyte, then during stage 9-10, oskar mRNA is moved from the
middle of the oocyte to the posterior pole by Khc (Cha et al., 2002). This is confused by
the fact that there is no obvious oskar localization defect in klc mutants. In khc mutants,
dorsal localization of gurken mRNA, proper anterior-dorsal localization of the oocyte
nucleus and Dhc posterior localization are also disrupted, but bicoid mRNA localization
24
INTRODUCTION
is not (Brendza et al., 2000; Duncan and Warrior, 2002). Drosophila khc mutants also
show neuronal defects at the larvae stages (Saxton et al., 1991). Thus, Khc plays a role
not only in oogenesis but also in neurongenesis. In neurons, cargos are transported at two
different speeds along axons. Cytoskeleton elements such as neurofilaments, tubulins and
actins, are transported slowly, whereas membrane bound organelles and synaptic
membrane proteins are transported faster (Cyr et al., 1991; Hirokawa, 1996; Vallee and
Bloom, 1991). khc mutations cause swellings in the axons that are filled with fast
transported cargoes such as Synaptotagmin (syt), an synaptic vesicle membrane protein,
normally concentrated in the terminal boutons (neurite varicosities). Impaired Khc
function causes neuronal defects due to a general disruption of fast axon transport (Hurd
and Saxton, 1996). For large cells, such as neurons or the oocyte, materials that need to be
distributed over long distance are often large and diffuse slowly. Hence, rapid, active
transport via Kinesin, along the microtubule network is necessary.
25
INTRODUCTION
1.4
1.4.1
Late oogenesis
Ooplasmic streaming
After the polarity determinants of the embryo are localized, the nurse cells start to pump
large amounts of their cytoplasm, which contribute the embryo development, into the
oocyte. This occurs from stage 10b. At this stage, microtubules become highly dynamic,
allowing fast and well ordered streaming. This streaming allows the equal distribution of
nurse cell cytoplasm throughout the oocyte (Theurkauf et al., 1992). oskar, bicoid, and
gurken mRNA are anchored to the oocyte cortex, preventing their delocalization during
streaming. In weak khc mutant background, stratified egg chambers are observed, in
which yolk granules accumulate in the posterior of the oocytes, leaving a clear zone in the
anterior part. This stratified egg chamber is caused by a lack of ooplasmic streaming and
remaining pumping force from the nurse cells. The plus-end directed motor (Khc) is
essential for ooplasmic streaming. The fact that a weak allele of khc, in which streaming
was slow or stopped, shows normal oskar mRNA localization, indicating that oskar
localization does not require for ooplasmic streaming (Serbus et al., 2005). Injecting of
Dhc antibodies blocking Dhc function or treatment of egg chambers with cytochalasin,
which disrupts F-actin, leads to premature streaming, suggesting that Khc can drive the
ooplamic streaming before stage 10B, but its function is blocked by Dhc and actin at
earlier stages (Serbus et al., 2005; Theurkauf, 1994).
1.4.2
The anterior-posterior axis of the embryo
After fertilization, Bicoid protein is translated from its anteriorly localized mRNA, and
diffuses towards the posterior of the embryo forming a gradient (Figure 4). The Bicoid
gradient is required to regulate zygotic gap gene expression and pattern the anterior
structures of embryo (Driever, 1993). During late stage oogenesis, Oskar recruits other
posterior pole-plasm components, including Vasa protein and nanos mRNA, which
encode the abdominal determinant of the embryo. Vasa, a DEAD-box RNA helicase, is
essential for posterior patterning and germ line formation in the embryo. Nanos is also
translated after fertilization and forms a gradient from posterior to anterior of the embryo.
26
INTRODUCTION
Nanos also regulates zygotic gap gene expression and abdominal patterning (Bergsten and
Gavis, 1999; Breitwieser et al., 1996; Ephrussi et al., 1991; Gavis and Lehmann, 1994).
Figure 4. Localization of anterior
and posterior determinants in the
egg
A schematic of a late stage egg
showing the dorsal appendages
(green) and micropyle (yellow).
Before fertilization, bicoid mRNA
(blue) is anchored at the anterior pole
and oskar and nanos mRNAs are at
the posterior. Following fertilization,
bicoid and nanos are translated.
Bicoid protein diffuses from the
anterior pole forming a gradient,
concurrently the Nanos gradient
forms from the posterior. The two
gradients regulate zygotic gap gene
expression and regulate early
embryonic patterning. Adapted from
(Riechmann and Ephrussi, 2001).
27
INTRODUCTION
1.5
1.5.1
Par-1 function in oogenesis
Par-1 affects microtubule-dependent polarity in the oocyte
When the germ line cyst reaches region 2b, the oocyte specific proteins and mRNAs
along with the centrosomes are transported into the presumptive oocyte, where they
accumulate at the anterior (Cox and Spradling, 2003). Once the egg chamber reaches
region 3, these proteins and mRNAs shift to the posterior of the oocyte (Huynh et al.,
2001; Pare and Suter, 2000). Par-1 is involved in this transportation. Par-1 is the
Drosophila homolog of Caenorhabditis elegans PAR-1: a serine/threonine kinase
required for the polarization of the C. elegans zygote (Guo and Kemphues, 1995). par-1
mutations disrupt the microtubule network in early oogenesis. In par-1 mutants, the
oocyte is specified and the centrosomes, oocyte specific mRNAs and proteins accumulate
in oocyte in region 2b and 3. But the oocyte specific proteins and mRNAs can not be the
transported to the posterior pole of the oocyte. Later the oocyte de-differentiates and
becomes a nurse cell (Cox et al., 2001; Huynh et al., 2001). Par-1 also plays a role in axis
specification during mid-oogenesis (Shulman et al., 2000; Tomancak et al., 2000).
Mutations in Par-1 disrupt posterior localization of osk RNA and microtubule polarity at
stage 9 (Benton et al., 2002; Shulman et al., 2000; Tomancak et al., 2000).
The proteins aPKC, Bazooka (Par-3) and Par-6 interact with each other and form a
functional complex (aPKC complex). The aPKC complex is localized to the anterior of
the oocyte and apical domains of follicle cells, corresponding to areas where microtubule
minus-ends are enriched. Par-1 localization does not overlap with aPKC complex (Benton
and St Johnston, 2003; Cox et al., 2001; Shulman et al., 2000; Vaccari and Ephrussi,
2002). Instead, Par-1 expresses in the posterior of the oocyte and basolateral regions of
follicle cells (Shulman et al., 2000; Tomancak et al., 2000). The asymmetric localization
of Par-1 and the aPKC complex is essential for cell polarity. PAR-1 can phosphorylate the
conserved 14-3-3 (PAR-5) binding site in Bazooka, allowing the recruitment of 14-3-3.
The phosphorylation of Bazooka destabilizes and delocalizes the aPKC complex,
establishing the mutually exclusive localization of aPKC/Bazooka/Par-6 complex and
Par-1 (Benton and St Johnston, 2003).
28
INTRODUCTION
1.5.2
Par-1/MARK regulate microtubules through MAPs
Par-1 is a homologue of the mammalian MAP/MT affinity regulating kinase (MARK)
family. The MARK family proteins are involved in establishing polarity in many different
cells (Tassan and Le Goff, 2004). MARK proteins are known to phosphorylate
microtubule associated proteins (MAP), such as Tau, MAP2 and MAP4 (Drewes et al.,
1995; Illenberger et al., 1996). These MAPs were originally identified to be copurified
with tubulin in microtubule binding assays. These MAPs contain three to four conserved
microtubule binding domains. They are known to bind and stabilize microtubules. Most
of microtubule associated proteins are found in neuronal cells. MAP2A and MAP2B are
enriched in the dendrites, Tau is abundant in axons, whereas MAP4, originally isolated
from HeLa cells, is abundant in a variety of cell and tissue types (review in (Mandelkow
and Mandelkow, 1995).
Overexpression of Tau in vivo or addition of Tau in excess to microtubules in vitro, leads
to its accumulation on the microtubules and interference with the movement of motor
proteins (Ackmann et al., 2000; Stamer et al., 2002). At normal conditions, Tau and other
MAPs clearly help motor transportation by creating space around microtubules (Chen et
al., 1992). Phosphorylation of MAPs by MARK reduces their affinity for microtubules
and consequently destabilizes microtubules (Drewes et al., 1997; Mandelkow et al., 2004).
This suggests that Par-1 may act as a direct mediator of microtubule organization in the
oocyte. However, mutations in Drosophila Tau or 205KMAP (MAP4 homologue) do not
disrupt oocyte polarity, suggesting that Tau and MAP4 may not be essential Par-1 targets
in the oocyte (Doerflinger et al., 2003; Pereira et al., 1992). Drosophila does not contain
MAP2 homologues (Doerflinger et al., 2003). So to date the essential targets of Par-1 in
Drosophila oocyte remain unknown.
1.5.3
E-MAP-115 (Ensconsin), another MAP
E-MAP-115 was originally isolated from HeLa cells (Bulinski and Borisy, 1979;
Weatherbee et al., 1980). It contains two highly charged regions in the N- and C-terminal
regions. A novel microtubule binding domain is also found in the highly charged region
N-terminal region (Masson and Kreis, 1993; Masson and Kreis, 1995). Several studies
29
INTRODUCTION
have shown that E-MAP-115 stabilizes and organize microtubules. First, cells expressing
high level of E-MAP-115 can stabilize microtubules against nocodazole treatment (which
can depolymerize microtubules). Second, association of E-MAP-115 with microtubules is
reduced when microtubules become dynamic during mitosis. During interphase,
microtubules are involved in the transportation of vesicles, whereas during mitosis,
microtubules are used to ensure accurate chromosomes segregation. The transition
between these two stages requires the rapid rearrangement of the microtubule network
and is accompanied by changes in the dynamic properties of microtubules. The decreased
association of E-MAP-115 with microtubules is correlated with serine/threonine
phosphorylation of E-MAP-115. This suggest that phosphorylation of E-MAP-115
regulate its microtubule binding affinity. However, it is still not known which kinases
regulate E-MAP-115 (Masson and Kreis, 1993; Masson and Kreis, 1995).
Faire et al. have shown that GFP labeled E-MAP115 is present along all microtubules
during mitosis. This suggests that E-MAP-115 may function to modulate the stability or
dynamics of microtubules (Faire et al., 1999). If E-MAP-115 is a microtubule stabilizing
protein, overexpression of it would be expected to alter microtubule stability.
Unfortunately, overexpressing E-MAP-115 4-10 times above endogenous levels does not
result in more bundles of stabilized microtubules.
This suggested that E-MAP-115 may
modulate other microtubule functions or interactions with other cytoskeletal elements
(Faire et al., 1999).
E-MAP-115 knock out mice have been generated and analyzed. Homozygous mice are
viable but male sterile, and show microtubule disruption during spermatogenesis
(Komada et al., 2000). However it is worth noting that mouse genome contains a similar
protein, RPRC1, which may have redundant function with E-MAP-115 in the mouse
genome.
30
INTRODUCTION
1.6
The aim of this thesis
Border cells delaminate from the anterior follicle cells and migrate through nurse cells
toward the oocyte. The original aim of my thesis was to understand the germ line function
in border cell migration. I performed a genetic screen in Drosophila. Unfortunately, I did
not find any good candidate to work on. I also identified genes required in germ line for
oocyte or embryonic development in this screen. In this thesis, I will present the result of
the screen and then focus on the gene, CG14998 (ensconsin) which is disrupted by one
PiggyBac insertion, PB4170.
Oocyte and embryo polarization occur by the specific localization of axis determinants.
Par-1 affects oocyte polarity by regulating microtubules through unkown Microtubule
associated proteins. Microtubules and motor proteins are also important for the
transportation and localization of axis determinants during Drosophila oogenesis. I show
that Drosophila Ensconsin is a microtubule associated protein and its N-terminal domain
binds microtubules, similar to its human homologue (E-MAP-115). In ensconsin mutant
egg chambers, axis determinants are mislocalized in a manner similar (but weaker) to Khc
mutants. Furthermore, ensconsin mutants disrupt the accumulation of Khc in the posterior
part of the oocyte at stage 9. My experiments also show that Ensconsin is a direct target
of Par-1. Par-1 phosphorylates Ensconsin and thereby regulates its localization. This is
crucial for the correct localization of axis determinants. Here, I describe a novel Par-1
target, Ensconsin, which is a microtubule associated protein, which controls
Khc-dependent motor transport.
31
2 MATERIALS AND METHODS
MATERIALS AND METHODS
2.1
Fly genetics
2.1.1
Fly husbandry
Flies were grown on standard corn meal molasses agar. All crosses were carried out at
25ºC. Fly stocks were stored at 18 ºC and flipped once a month.
Fly food recipe
12 g agar, 18 g dry yeast, 10 g soy flour, 22 g turnip syrup, 80 g malt extract, 80 g corn
powder, 6.25 ml propionic acid, and 2.4 g methyl 4-hydroxybenzoate (Nipagin) are mixed
with one liter water.
2.1.2
Fly strains
Table 1. Fly strains used for experiments
Genotype
Source
PB;;FRT40FRT42;;FRT80FRT82
(Mathieu et al., 2007)
If, transposase/CyO
(Horn and Wimmer,
2000)
eyFLP;;FRT40/CyO;;FRT80/TM3ser (Thompson et al., 2005)
eyFLP;;FRT42/CyO;;FRT82/TM3ser (Thompson et al., 2005)
FRT40ovoD1/CyO
Bloomington #2121
FRT42ovoD1/CyO
FRT80ovoD1/TM3ser
FRT82ovoD1/TM3ser
Bloomington #2149
FRTG13Par-19A
(Tomancak et al., 2000)
FRTG13Par-1W3
(Shulman et al., 2000)
kinesinLacZ
(Clark et al., 1994)
tub-tauGFP
(Micklem et al., 1997)
FRTG13ubiGFP
Bloomington #5826
FRT80ubiGFP
Bloomington #5630
PB4170
P{RS3}CB-5457-3
Szeged stock center
Df(3L)ED4341
Szeged stock center
FRT80ens∆N
FRT80ens∆C
UASP-ens
UASP-ens-mutant ∆3’UTR
Tub-ens
Tub-ens∆3’UTR
Tub-ensmutant∆3’UTR
elavGAL4
Pernille Rørth
Pernille Rørth
Pernille Rørth
Pernille Rørth
Pernille Rørth
(Campos et al., 1987)
Description
PB jump starter
Transposase for PB
For mapping
For mapping
For germ line clone analysis
For germ line clone analysis
For germ line clone analysis
For germ line clone analysis
For testing Ens localization
For testing Ens localization
Plus end marker
For marking microtubules
For creating mutant clone
For creating mutant clone
ensconsin mutant allele
ensconsin mutant allele
Deficiency for ensconsin
ensconsin mutant allele from
P{RS3}CB-5457-3
imprecise
excision
ensconsin mutant allele from
end-out experiment
Pan-neuron driver
33
MATERIALS AND METHODS
Continued Table 1.
P{70FLP}11,P{70I-SceI}2B
Sco
noc /CyO
Maternal:: GAL4 VP16
2.1.3
Bloomington #6934
For end-out experiment
(Coutelis and Ephrussi,
2007)
Germ line specific driver
Ectopic expression using the GAL4/UAS system
In Drosophila, genes of interest can be expressed in a temporally and spatially regulated
manner by using the GAL4/UAS system (Brand and Perrimon, 1993). This system uses
the yeast transcription activator, GAL4, and its target sequence, upstream activation
sequence (UAS). When GAL4 binds its UAS, it activates the transcription of the gene
downstream of UAS. GAL4 can be expressed in many different patterns under the control
of various Drosophila promoter sequences and activates expression of target gene or
reporters placed downstream of a UAS. In order to express a target gene in to the pattern
of a specific promoter, transgenic flies carrying the GAL4 driver under the control of a
specific promoter are crossed to transgenic flies carrying UAS followed by the target
gene.
Figure 5. GAL4-UAS system
A fly line expressing GAL4 (a transcriptional activator protein) is
crossed to a line carrying the UAS element upstream of gene X. In the
progeny, GAL4, expressed in the tissue of interest, binds to the UAS
element and activates transcription of the downstream gene X,
specifically in that tissue. Adapted from (Brand and Perrimon, 1993).
34
MATERIALS AND METHODS
2.1.4
Generation of mosaic clones using the FLP/FRT system
Figure 6. FLP-FRT system
FLP-recombinase can induce site-specific exchange at position of the FRT
(FLP-recombinase target sequence) sequence during mitosis. Yellow is the mutant locus,
and green the ubiGFP marker. In the absence of FLP, the genotype of daughter cells is like
that of the mother cell (FRTmutant/FRTubiGFP). When FLP is added, homozygous mutant
ovaries without ubiGFP can be obtained and also homozygous wild type.
In order to obtain a group of clonal cells that are homozygous mutant for a given allele,
the FLP/FRT (flippase recombinase/ flippase recognition target) system was introduced
into fly genetics (Golic, 1991). The mutant allele is recombined onto a suitable FRT
chromosome (Xu and Rubin, 1993) and crossed to flies carrying a heat-shock induced
flippase (hs-FLP) and marker (usually Ubi-GFP) on the matching FRT chromosome.
Ubi-GFP expresses ubiquitous expression of green fluorescent protein under the ubiquitin
promoter. To induce mitotic clones in the germ line or ovarian follicular cells, young
larvae at 48-72 hours after eggs lying (AEL) are heat-shocked at 37°C for 1 hour.
Following heat shock, the FLP induces recombination of homologous chromosomes at
the FRT sites. The recombination event results in two daughter cells: one homozygous for
the GFP transgene carrying chromosome; the other homozygous for the mutant allele.
Subsequently, the two daughter cells will give rise to two clonal groups of cells: a wild
type clone with a higher level of GFP and a mutant clone lacking GFP expression. So,
homozygous mutant cells are marked by the absence of GFP in a background of either
homozygous wild type, GFP-expressing cells (the twin clone) or heterozygous,
35
MATERIALS AND METHODS
GFP-expressing (no recombination event) cells.
2.1.5
Generation of Germ line clones using the FLP-DFS system
This system is similar to FLP-FRT system. However, instead of ubi-GFP, an FRT
chromosome carrying the dominant female sterile (DFS) transgene, ovoD1 is used. In
ovoD1 ovaries, oogenesis stops at early stages, but does not affect adult viability. The
mutant allele is recombined onto a suitable FRT chromosome (Xu and Rubin, 1993) and
crossed to flies carrying heat-shock induced flippase (hs-FLP) and ovoD1 on the same
FRT chromosome. To induce mitotic clones in the germ line cells, the resulting larvae are
heat-shocked at 48-72 hours AEL at 37°C for 1 hour. Following the heat shock, the FLP
induces recombination at the FRT sites. This recombination event may result in two
daughter cells: one homozygous for the ovoD1 transgene chromosome; the other one
homozygous for the mutant allele. Subsequently, the two daughter cells will give rise to
two groups of clonal germ cells: The ovoD1 clone will produce small arrested ovaries;
Figure 7. FLP-DFS system
The FLP-recombinase induces site-specific exchange at the FRT (FLP-recombinase target
sequence) sequence during mitosis. Yellow is PiggyBac insertion, and green is ovoD1.
ovoD1 is a dominant female sterile transgene. Without FLP, the genotype of daughter cells
is like mother cell (FRTPiggyBac/FRTovoD1). Ovaries of this genotype are small. With FLP,
homozygous mutant ovaries can be obtained.
36
MATERIALS AND METHODS
and the mutant clone, ovaries are different sizes depending on the mutant used (Chou and
Perrimon, 1996).
37
MATERIALS AND METHODS
2.2
2.2.1
Mutant analysis in the Screen
X-gal staining in the Drosophila egg chamber
Ovaries were dissected in cold PBS, fixed in 0.5% glutaraldehyde in PBS for 10 min,
washed in PBS-0.1%Triton (PBST), and stained (in 10 mM NaH2PO4/Na2HPO4 (pH7.2),
150 mM NaCl, 1 mM MgCl2, 3.1 mM K4(FeII(CN)6), 3.1 mM K3(FeIII(CN)6), 0.3% Triton
X-100, 0.2% X-Gal) at room temperature in 1 hour. Ovaries were then washed in PBST
and mounted in 50% glycerol.
2.2.2
Cuticle preparation
Embryos were collected and dechorinated in 50% bleach in 1-3 minutes. Embryos were
washed thoroughly to remove the bleach and transferred to a 2 ml eppendorf tube
containing a Methanol/ Heptane (1:1) solution and shaken vigorously for 1 minute to
remove the vitalline membrane. This causes the devitallinzed embryos to sink to the
bottom. All liquid and any embryos at the interface were removed. Embryos were
mounted in the Hoyer's mount and incubated in 65oC overnight.
Hoyer’s mount: 30 g of gum Arabic was added to 50 ml distilled water, and stirred
overnight. While stirring, 200 g chloral hydrate was added in small quantities. Then 20 g
glycerol was added. The solution was centrifuged at least 3 hours at 12000 g to clear.
Lactic acid (1:1) was added in to increase contrast and decrease clearing time.
2.2.3
Inverse PCR
Inverse PCR (IPCR) is a method which allows the rapid amplification of DNA sequences
flanking a region of known sequence (Ochman et al., 1988). The method uses PCR, but
with primers oriented in the opposite direction to usual. The template for the reverse
primers is a restriction fragment that has been self-ligated to form a circle. IPCR is
routinely used to amplify and identify the sequences flanking transposable elements.
Fly genomic DNA was prepared using the DNAeasy kit (Quiagen). Then 15 µl of gDNA
38
MATERIALS AND METHODS
was used for the enzyme digestion in total volume of 20 µl for 2 hours at 37°C. The
enzymes used for inverse PCR are 4-cutters restriction enzymes with recognition sites
present once in the 5’ and 3’ end of the PiggyBac element, such as Sau3AI or HinP1.
After digestion, the mixture was incubated at 65°C for 20 minutes to heat-inactivate the
restriction enzyme, and then transferred to ice immediately. The ligation mixture was set
up in a total volume of 200 µl, (to promote self-ligation), and included T4 DNA ligase,
ligase buffer, and the original 20 µl digest mixture. The reaction was then incubated at
16°C overnight. The next day, DNA was precipitated by adding 20 µl of 3M sodium
acetate and two volumes of absolute ethanol. The mixture was left at -80°C for 30
minutes and centrifuged it at 4°C for 30 minutes. The DNA pellet was washed twice with
70% ethanol and resuspended it in 20 µl of water. The flanking region of the PiggyBac
insertion was amplified with PiggyBac element specific primers, designed to amplify out
from the transposon vector sequence. The primers PLF (5′-CTT GAC CTT GCC ACA
GAG GAC TAT TAG AGG-3′) and PLR (5′-CAG TGA CAC TTA CCG CAT TGA CAA
GCA CGC-3′) were used to amplify the genomic fragment from the 5’ end of PiggyBac
and the primers PRF (5′-CCT CGA TAT ACA GAC CGA TAA AAC ACA TGC-3′) and
PRR (5′-AGT CAG TCA GAA ACA ACT TTG GCA CAT ATC-3′) were used for the 3’
end. The annealing temperature of PCR reaction was set at 60°C. The amplified products
were then sequenced and compared by BLAST search against the Drosophila genome to
identify the insertion site.
39
MATERIALS AND METHODS
2.3
ensconsin mutant analysis
The following ensconsin constructs were used: Full length ens-wt cDNA, ens without the
3’UTR, an ens mutant which removed all putative 14-3-3 binding sites, Ens-X which
lacks amino acids 307-676 and Ens-C which lacks amino acids 1-370 (generated by
Pernille Rørth).
ens∆N was obtained from imprecise excision of P{RS3}CB-5457-3 and checked by single
fly PCR (using the primers CB5’ CAC TAC AGA GCT GGC CAC ACT G and PI3’ GCA
AGA CGA CAA AGG AAC AAC TGC AAG).
ens∆C was generated using the ends-out homologous recombination system. I designed
primers to amplify genomic regions +4146-+7356 (primers A5’ and A3’ were used) and
+9578-+12904 (primers B5’ and B3’ were used) relative to the transcription start site
from genomic DNA. Primer sequences in red are restriction enzyme sites used for further
cloning. The two flanking sequence were cloned into the PW25 vector and used to
generate transgenic (donor) flies. The donor flies were then crossed with flies carrying the
recombination transgenes (P{70FLP}11 P{70I-SceI}2B nocSco/CyO) which expresses the
FLP site-specific recombinase and I-SceI endonuclease. I used a heat-shock to induce
expression of these enzymes. The FLP and I-SceI then linearize the donor molecule,
allowing it to undergo homologous recombination with the target locus. These targeting
events occur in the germ line. Using this method, I replaced the Ensconsin C-terminal
region with the mini-white cassette (Figure 9) (Gong and Golic, 2004).
A5’GCGGCC GCT CGA TCT TCG ATT TGA ATC ACC CGC (sequences in red are
Not1 cutting site).
A3’GGTACC TCA ATC TGC TGC AGG ACG TAG TCG C (sequences in red are Kpn1
cutting site).
B5’ GGCGCGCC ATT ATT GCA GGG GGG AAA CAC AGG G (sequences in red are
Asc1 cutting site).
B3’ CGTACG CCC ACT TCC AAA CAA ACA GCC AAC G (sequences in red are
BsiW cutting site)
40
MATERIALS AND METHODS
Figure 8. Ends-out system
The targeting scheme. (A) Regions
flanking the locus to be deleted, indicated
as L and R, were cloned into the
P-element vector, pW25, with the
indicated features. FLP and I-SceI
generate the extra chromosomal linear
donor, as shown (B). (C) Recombination
of the ensconsin and ensconsin donor
should generate the indicated deletions.
Adapted from (Gong and Golic, 2004).
2.3.1
RNA isolation
5-10 adult ovaries were dissected and put into 1.5 ml eppendorf tube on ice. They where
then dissociated by pipetting up and down with 200 µl of TRIzol (Invitrogen) until no
large pieces could be seen. 800 µl of TRIzol was added and the lysate was kept at room
temperature for 5 minutes. 200 µl of Chloroform was added and samples were vortexed
for 15 seconds. The samples were incubated at room temperature for 15 minutes before a
centrifugation at 14000 rpm at 4°C for 15 minutes. The aqueous phase (top or non-pink
layer) was transferred into a new tube and 500 µl of Isopropanol was added to precipitate
the RNA. Samples were incubated at room temperature for 10 minutes, and then
centrifuged for 15 minutes at 4°C. The pellet was washed with 75 % ethanol (in
RNase-free water) and dried at room temperature for 15 minutes. The pellet was the
resuspended in 50 µl of RNase-free water. Samples were heated to 60°C in order to
dissolve the RNA if necessary. Samples were stored at -80°C.
41
MATERIALS AND METHODS
2.3.2
First-strand cDNA synthesis
First-strand cDNA for PCR reaction was synthesized by reverse transcriptase using the
SuperScriptTM first-strand synthesis system (Cat. 11904-018, Invitrogen). 1 µg of RNA
was mixed with oligo(dT) and dNTPs in a final volume of 10 µl and heated to 65°C to
denature secondary structures of RNA, and then placed them on ice for 1 minute. The
reaction mixture composed of reaction buffer, MgCl2, DTT, RNase inhibitor, and
SuperScriptTM II RT was prepared, and 10 µl of the mixture was added to each
RNA/primer mix. Samples were incubated at 42°C for 50 minutes. The reactions were
then terminated by heating at 70°C for 15 minutes, and transferred to ice for the next
procedure or store them at -20°C. RT5’, RT3' or CB5’, 2RT3’ were used to check
different splicing forms and transcript in PB4170.
RT5’ GCCGACAGTGGTGCTAAGAAGCG
Rt3'GCGATACTGGCGGGCCTCG
CB5’ CACTACAGAGCTGGCCACACTG
2RT3’ TACTCGCGGCGCTCCTTCTCG
2.3.3
Microtubule binding assay
1.5 ml 0-2 hours embryos or 100µl ovaries were collected. Embryos or ovaries were
homogenized with the same volume of BRB-80 buffer (80 mM PIPEs pH6.6, 1 mM
MgCl2, 1 mM EGTA) with protease inhibitor (Roche) on ice, and supplemented with 0.1
% NP40, 1 mM DTT and 1 mM GTP. The solution was centrifuged at 100000g in
TLA100.4 or TLA100 rotors for 20 minutes 4oC. The supernatant was split into two tubes.
The supernatant was heated to 25 oC in water bath. To one sample taxol was added to a
final concentration of 20µM, and an equal volume of DMSO was added to the control. All
reactions were kept at 25 oC 30 minutes to allow microtubules to polymerize. The solution
was then layered onto the 1V cushion of 40 % glycerol in BRB80 buffer. All reactions
were spun 100000g in TLA 100.4 or TLA 100 10 minutes 25 oC. 75 µl of the 1V
supernatant and 25 µl 4X Laemmli Buffer were added. The pellet was dissolved in 1V
BRB80 buffer. After dissolving, 75 µL solution was taken to mix with 25 µl 4X Laemmli
42
MATERIALS AND METHODS
Buffer. Both samples were analyzed by western blot, probed with anti-tubulin and
anti-Ensconsin.
2.3.4
Western blotting
For whole larvae or adults extracts, total protein was obtained by crushing 10 pairs of
ovaries in 200 µl 1x Laemmli Buffer with a plastic pestle. Samples were boiled at 95°C
for 5 minutes and 10 µl from each was loaded on an SDS-polyacrylamide gel. Proteins
were separated by elecrophoresis with a constant current of 20 mA per mini-gel. Proteins
were transferred to nitrocellulose membranes (10401196, Schleicher & Schuell) using a
constant current at 45 mA per mini-gel for 1 hour. The membrane was rinsed with water
and incubated in Ponceau S solution for several minutes. The membrane was then washed
with water several times and checked the total protein levels. The membrane was blocked
with 5 % milk/ PBT (PBS with 0.1% Tween20) for 1 hour. Next, it was incubated with
primary antibody diluted in 5 % milk/ PBT at 4°C overnight. Primary antibodies used in
this study are mouse anti-α-Tubulin DM1A (1:5000) and rat anti-Ensconsin (1:10000).
The membrane was rinsed once and washed for 15 minutes 3 times with PBT. The
membrane was
incubated with HRP-conjugated secondary antibody (Jackson
ImmunoResearch) diluted in 5 % milk/ PBT at room temperature for 1 hour. The
membrane was rinsed once and washed for 3 times 15 minutes with PBT. The membrane
was rinsed with enhanced chemiluminescence reagent by mixing equal volumes of the
Enhanced Luminol Reagent and the Oxidizing Reagent (NEL105, PerkinElmer). The
membrane was exposed to Kodak X-OMAT MR Film for 30 seconds to 15 minutes.
2.3.5
Single Fly PCR
One fly was squashed in 50 µl SB buffer (10 µM Tris-Cl, 1 µM EDTA, and 25 µM NaCl)
with 1 µl ProteinaseK (10 mg/ml). The solution was incubated for 1 hour at 37oC. The
solution was then incubated at 95oC for 2 minutes to inactive the proteinase K. 1 µl
solution was used per PCR reaction.
PCR reaction:
43
MATERIALS AND METHODS
The PCR reaction was performed in a final volume of 20 µl and contained 1 µl of
extracted genomic DNA as amplification template, 1 U Taq DNA polymerase (Roche), 1x
PCR Buffer containing MgCl2 (Roche), 0.2 mM dNTPs (PCR grade, Roche), 1 µM of
forward and reverse primers, and sterile distilled water. The PCR reaction was running on
Thermal Cycler (PTC-200, MJ research) according to the following program: (1) 94°C
for 3 minutes, (2) 94°C for 30 seconds, (3) 60°C for 30 seconds (depend on the Tm values
of primers), (4) 72°C 30 seconds to 1 minute (depend on the length of the product), (5)
repeat step (2)-(4) for 34 times, (6) 72°C 10 min, (7) end at 4°C. The amplified products
were separated by electrophoresis on 1 % agarose gel and visualized by Ethidium
Bromide staining and UV light.
2.3.6
Purification of Ensconsin protein
I designed primers to amplify fragments from LD09646, enswt and ens mutant without
14-3-3 binding sites plasmids which are constructed by Pernille Rørth. Then the
fragments were cloned into PETM11vector (N-his tag labeling) to express the truncated
protein. Plasmids were transformed into BL21 competent cells. The colony was cultured
into in 5 ml LB overnight. 500 µl bacterial culture was mixed with 500 µl 30% glycerol,
flash frozen and kept as a stock. 200 µl bacterial culture was used to inoculate 2 l of LB.
The culture was incubated for about 2 hours at 30oC until the OD reached 0.4-0.6 IPTG
was then added to a final concentration 1 mM to induce protein expression. The culture
was then grown at 30oC for another 4 hours. The bacteria were pelleted and resuspended
in 40 ml Lysis buffer. The lysate flash frozen in liquid nitrogen and kept in -80oC. When
required, the lysate was then thawed on ice. Proteinase inhibitor cocktail (Roche) and
DNAase powder were added to digest DNA. The lysate was passed through a French
press to lyse the cells. The sample was then sonicated 2 times 30sec to further disrupting
the cells. The lysate was spun at 4000 rpm at 4°C. The supernatant was mixed with nickel
beads for 2 hours at 4oC on a shaker. Before mixing, water and Lysis buffer washes were
used twice to remove methanol from 500 µl of beads. The beads were spun at 600 rpm for
5 min and the supernatant was removed. Beads were washed 3 times in wash to remove
non-specific binding proteins. The protein was then eluted in 3 washes of 500 µl elution
buffer.
44
MATERIALS AND METHODS
Lysis buffer (500 mM NaH2PO4, 500 mM NaCl, 5 mM β-Mercaptoethanol, 10% glycerol,
1 mM MgCl2, pH7.4 at room temperature)
Wash buffer (50 mM NaH2PO4, 1 M NaCl, 5 mM β-Mercaptoethanol, 10% glycerol, 1
mM MgCl2, 40 mM Imidazole, pH7.4 at room temperature)
Elution buffer (500 mM NaH2PO4, 500 mM NaCl, 5 mM β-Mercaptoethanol, 10%
glycerol, 1 mM MgCl2, 500 mM Imidazole, pH7.4 at room temperature)
For antigen A, primersA5’ andA3’ were used to do a PCR. For antigen B, primer B5’-B3’
were used. For Ens-N and Ens-N mutant, C5’ and A3’ were used. For C-terminal protein,
D5’and D3’ were used.
A5’ CCATGGctTCCGCACAAACGACACCCAAACG (sequences in red is Nco1
cutting site)
A3’ GAATTCCAGTTCCCGCACCTCTGAGTTC(sequences in red is EcoR1 cutting
site)
B5’ CCATGGctGCAGCTCCTGCCGCAACGG (sequences in red is Nco1 cutting site)
B3’ GAATTCCAGCAGCGATATATCTTTATTTTCGTGACTATC (sequences in red is
EcoR1 cutting site)
C5’ CCATGGctATGGCGAGTCTTGGGGGCCAAC (sequences in red is Nco1 cutting
site)
D5’ CCATGGctTCCACCATCACAAAACCAGCTCCTG (sequences in red is Nco1
cutting site)
D3’ GGATCCTCACAGCAGCGATATATCTTTATTTTCGTGAC (sequences in red is
BamH1 cutting site)
2.3.7
In vitro kinase assay
The in vitro kinase assay was performed as described (Benton et al., 2002). Bacterially
expressed Ensconsin fragments were mixed with the kinase source and 50 µM of γ-32P
label ATP or BnATP (synthesized by Piyi Papadaki) in kinase buffer (20 mM HEPES
pH7.5, 150 mM NaCl, 5 mM MgCl2) for 1 hour at 30oC. The kinase source used was
45
MATERIALS AND METHODS
bacterially expressed MBP-fused Par-1 kinase domain (amino acid 245-521, done by Piyi
Papadaki) or GFP-fused full length GFP-Par-1M329G (done by Piyi Papadaki)
immuno-precipated from ovarian lysate using an anti-GFP monoclonal antibody
(Molecular probes). Samples were run on a 10% protein gel, and then stained in
Coomassiae solution (45% Methanol, 10% Acetic acid and 0.1% Coomassiae Brilliant
Blue R-250) for 30 minutes. The gel was transferred to the destaining solution (50% H2O,
40% Methanol and 10% Acetic acid) for 1 hour to remove the background signal. The gel
was then dried and exposed to Kodak X-OMAT MR Film for 1-3 days.
2.3.8
The ensconsin Probe preparation
The ensconsin cDNA clone LD09626 was linearized using BamH1. 2 µl of the linearized
plasmid was incubated with 2 µl DEPC- water, 2 µl 10X buffer for T7 polymerase, 2 µl
each of ATP, CTP, GTP, 1.5 µl UTP, 4 µl DIG-11-UTP (10nmol/ul), 2 µl T7 polymerase 6
hours at 37oC. Then 1 µl DNase was added in the solution for 15 minutes at 37oC to
digest the template DNA. The probe was purified using a Qiagen RNeasy Kit.
2.3.9
In situ hybridization of egg chambers
Fly ovaries were dissected in the cold PBS and fixed in 4% Paraformaldehyde for 30
minutes. Ovaries were washed for 3 times 10 minutes in PBT (PBS with 0.1% Tween-20).
Ovaries were washed with 1:1 PBST/Hybridization buffer for 5 minutes. Ovaries were
then washed in Hybridization buffer for 5 minutes. Ovaries were prehybridized in the 1ml
Hybridization buffer containing 20 µl Salmon sperm DNA (5mg/ml) for at least 1.5 hours
in a 55oC heat block. Ovaries were then hybridized in 50µl hybridization buffer
containing the probe (1:1000), 10 µl ssDNA at 65oC water bath overnight. Probes used in
this study were oskar, gurken, bicoid (gift from Anna Cyrklaff) and ensconsin. Ovaries
were washed in hybridization buffer 30 minutes at 65oC, then in 1:1 hybridization
buffer/PBST for 30 minutes at 65oC. Ovaries were washed for 4 times 20 minutes in
PBST at 65oC, then in PBST for 20 minutes at room temperature. Ovaries were incubated
with anti-DIG-Hrp (Roche) antibody for 2 hours in PBST. Ovaries were washed for 5x30
min in PBST. Ovaries were incubated in staining solution (0.5 mg/ml DAB and 0.003%
46
MATERIALS AND METHODS
H2O2), washed in the PBST, and then mounted in 70% glycerol.
Hybridization buffer (50% Formamide, 5X SSC, 0.1% Tween-20)
2.3.10
In situ hybridization of embryos
Flies were allowed to lay eggs on apple juice plates. The eggs were then washed with into
a tube sealed at one end with mesh. Embryos were dechorionated in a 50% bleach
solution for 1-2 minutes. Embryos were washed thoroughly to remove bleach and
transferred into a 2 ml eppendorf tube. They were then fixed by shaking in 4 %
Formaldehyde/ PBS/ Heptane for 25 minutes. As the fixed embryos rest at the interface,
the lower, aqueous phase was removed and 500 µl methanol added. The tube was
vigorously shaken for 1 minute to remove the vitalline membrane. The devitallinized
embryos will then sink to the bottom. All liquid and any embryos at interface were then
removed. Embryos were washed twice in methanol and stored at -20°C. Embryos were
then rehydrated in PBT (PBS with 0.1% Tween-20) and fixed again in 4%
paraformadehyde for 30 minutes. They were then washed for 3 times 20 minutes in PBT
(PBS with 0.1% Tween-20). Embryos were washed in a 1:1 mix of PBT/hybridization
buffer for 5 minutes, then in hybridization buffer for 5 minutes. Embryos were
prehybridized in 1 ml hybridization buffer containing 20 µl Salmon sperm DNA (5mg/ml)
for at least 1.5 hours in 55oC heat block. Embryos were hybridized in 50 µl hybridization
buffer with probe (1:1000), 10 µl ssDNA at 65oC water bath O/N. The probes, used in this
study are ftz, and eve (gifts from Natasha Bushati). Embryos were washed in
hybridization buffer for 30 minutes at 65oC. They were then washed in 1:1 hybridization
buffer/PBST for 30 minutes at 65oC. Following this, embryos were washed for 4 times 20
minutes in PBST at 65oC then once in PBST at room temperature. Embryos were
incubated with anti-DIG-HRP antibody (Roche) in PBST for 2 hours. Embryos were
washed for 5 times 30 minutes in PBST. Embryos were stained in 0.5 mg/ml DAB and
0.003% H2O2 for 10-30 minutes in the dark, and then washed in the PBST and mounted
in 70% Glycerol.
Hybridization buffer (50% Formamide, 5X SSC, 0.1% Tween-20)
47
MATERIALS AND METHODS
2.3.11
Immunofluorescence staining of larvae axons
Wandering third instar larvae were dissected in cold PBS as follows: The larva was cut in
half and the head inverted by pushing the mouth hook with forceps. The fat body and
salivary gland were removed from the head to prevent the endo-replicating tissues
depleting the antibodies. Dissected tissues were transferred to a 2 ml eppendorf tube and
fix samples with 4% paraformaldehyde in PBS for 20 minutes on the shocker. The tissues
were washed in PBST (PBS with 0.1% Triton-X100) for 30 minutes. The tissues were
then blocked in PBST with 5% NGS for 1 hour before being incubated with primary
antibody overnight. Primary antibody was used in this study: rabbit anti-Syt (1:100, the
gift from Hugo Bellen). The tissues were washed in several changes of PBST with 0.2%
BSA for 2 hours, blocked in PBST with 5% NGS (1 hour), then incubate with Cy5,
Rodamine and HRP conjugated secondary antibodies in PBST with 5% NGS for 2 hours.
The tissues were washed repeatedly in PBST and mounted in 80% Glycerol with 0.4%
n-Propyl gallate (P313 Sigma) in PBS. Axons were dissected out of the tissue prior to
confocal imaging. Analyses were performed using a Leica TCS SP confocal microscope
and images were editing using Photoshop CS (Adobe, San Jose, CA)
2.3.12
Immunofluorescence staining of embryos
Embryos were collected and fixed as with in situ analysis. Devitallinized, methanol
washed embryos were washed in PBST (PBS with 0.1% Triton-X100) for 30 minutes,
then blocked in PBST with 5% NGS for 1 hour. Embryos were incubated with primary
antibody overnight. Primary antibodies were used in this study: mouse anti-En (1:10
DSHB), rabbit anti-Ftz (1:350), genie pig anti-Eve (1:100), rabbit anti-Knirpe (1:10), and
rat anti-Kruppel (1:100) (a gift from Natasha Bushati). Embryos were washed several
times in PBST with 0.2% BSA for 2 hours. Embryos were blocked in PBST with 5%
NGS 1 hour. Embryos were incubated with FITC, Cy5, Rhodamine conjugated secondary
antibodies (1:100) and Rhodamine conjugated Phallodin (1:500; Molecular probes) in
PBST with 5% NGS for 2 hours. Embryos were finally washed in PBST, and mounted in
80% Glycerol with 0.4% n-Propyl gallate (P313 Sigma) in PBS. Analyses were
performed using a Leica TCS SP confocal microscope and images were editing using
Photoshop CS (Adobe, San Jose, CA).
48
MATERIALS AND METHODS
2.3.13
Immunofluorescence staining of egg chambers
Ovaries were dissected from females in the cold PBS and fixed in 4% paraformaldehyde
for 20 minutes. They were then washed in PBST (PBS with 0.1% Triton) for 10 minutes.
Ovaries were permeablized in PBST (PBS with 1% Triton) for 1 hour then blocked in
PBST with 5% NGS for 1 hour. Ovaries were incubated overnight in primary antibody.
Primary antibodies were used in this study: Rat anti-Staufen (1:2000), rabbit anti-Oskar
(1:2000) (gifts from Anne Ephrussi), rabbit anti-aPKC (1:100, Santa Cruz), and mouse
α-tubulin (1:100, Sigma clone DM1A), mouse anti-Gurken (1:100, DHSB), mouse
anti-α-spectrin (1:100, DHSB). Ovaries were washed several times in the PBST with
anti-
0.2% BSA for 2 hours. Ovaries were then blocked in PBST with 5% NGS for 1 hour
before incubation with FITC, Cy5, or Rhodamine conjugated secondary antibodies (1:100)
and Rhodamine conjugated Phallodin (1:500; Molecular probes) in PBST with 5% NGS
for 2 hours. Ovaries were washed repeatedly in PBST, and mounted in 80 % Glycerol and
0.4 % n-Propyl gallate (P313 Sigma) in PBS. Oskar and Staufen stained samples were
dehydrated in methanol and mounted in clearing solution (1:2 Benzyol alcohol:
Benzylbenzoate). Analyses were performed using a Leica TCS SP confocal microscope
and images were editing using Photoshop CS (Adobe, San Jose, CA).
2.3.14
Dhc immunostaining of egg chambers
Ovaries were dissected in cold PBS and fixed in 4% PFA for 15 minutes. Ovaries were
then washed in PBS with 0.1% Tween-20 (PBT) for 10 minutes, then dehydrated in
methanol and keep in 100% methanol at -20oC O/N. Ovaries were allowed to re-hydrate
in PBT (PBS with 0.1% Tween) for 4-8 hours at 4oC. Ovaries were blocked (PBT with
5% NGS) for 30 minutes. Ovaries were incubated overnight with mouse P1H4 anti-Dhc
(McGrail and Hays, 1997). After staining ovaries were washed several changes of PBT
with 0.2% BSA for 2 hours. Ovaries were blocked again (PBT with 5% NGS) then
incubated in PBT with 5% NGS with secondary antibodies at 4oC overnight. Ovaries
were washing in PBT 3 times then dehydrated in the methanol for 5 minutes and mounted
in cleaning solution (Benzyol alcohol: Benzylbenzoate = 1:2).
49
MATERIALS AND METHODS
2.3.15
Khc immunostaining of Drosophila egg chambers
Ovaries were dissected in the cold PBS, fixed in 4% paraformadehyde in PBS for 30
minutes and washed 3 times 10 min in PBST (PBS with 0.1% Tween-20). Ovaries were
washed in 1:1 PBST/hybridization buffer for 5 minutes, then in hybridization buffer for 5
minutes. Ovaries were prehybridized in 1 ml hybridization buffer for at least 1.5 hours at
55oC. Ovaries were hybridized 50 µl hybridization buffer at 65oC overnight. Ovaries were
washed in hybridization buffer for 30 minutes at 65oC. They were then washed in 1:1
hybridization buffer/PBST 30 minutes at 65oC. Ovaries were washed for 4 times 20
minutes in PBT at 65oC, then in PBT for 20 minutes at room temperature. Ovaries were
incubated with rabbit anti-Khc (1:100, Cytoskeleton) for 2 hours in the PBT. Ovaries
were washed for 3 times 30 minutes in PBT, then incubated in PBT with 5% NGS with
secondary antibodies at 4oC overnight. Ovaries were washed in PBT 3 times. Ovaries
were dehydrated in methanol for 5 minutes and mounted in cleaning solution (Benzyol
alcohol: Benzylbenzoate = 1:2).
Hybridization buffer (50% Formamide, 5X SSC, 0.1% Tween-20)
2.3.16
Live image of tau-GFP in the ovoD1 germ line clone
hsFLP/ +;; Tub-TauGFP/ +; FRT80 ens∆C/FRT80 ovoD1 females were dissected and
mounted on a glass slide in halocarbon oil (Halocarbon products Crop, USA). The
samples were immediate imaged using a confocal microscope (Leica NT, Germany).
2.3.17
Live image for ooplasmic streaming in the egg chamber
Wild type and PB4170 mutant ovaries were dissected in halocarbon oil (Halocarbon
products Crop.USA) and transferred to glass bottomed chamber (MatTek Corp). The
samples were immediately imaged using a by Perkin Elmer UltraView RS spinning disc
confocal. Images were contrast enhanced and maximum projections of 30 sections per
time point were made using ImageJ (NIH).
50
MATERIALS AND METHODS
2.3.18
Climbing assay
A single 3 days old male fly was put into an empty vial. After a 30 minute recovery, the
fly was tapped to the bottom of the vial and thenthe time it took to climb up 5cm was
recording. If flies did not reach 5 cm in 2 minutes, counting was stopped.
51
MATERIALS AND METHODS
2.4
Websites
BDGP
http://www.fruitfly.org/
BLAST http://www.ncbi.nlm.nih.gov/BLAST/
ClustalW http://www.ebi.ac.uk/clustalw/index.html
DHSB
http://www.uiowa.edu/~dshbwww/
Flybase http://flybase.bio.indiana.edu/
NEB cutter
http://tools.neb.com/NEBcutter2/index.php
PubMed http://www.ncbi.nlm.nih.gov/entrez/query.fcgi
SMART http://smart.embl-heidelberg.de/
52
3 RESULTS
RESULTS
3.1
Screen for genes affecting border cell
migration, oogenesis and embryogenesis
Most of the mRNA and proteins required for oogenesis and embryogenesis are
synthesized in the nurse cells and transported into the oocyte. To investigate the functions
of these mRNA and proteins, I performed a screen with Juliette Mathieu, a former
postdoctor fellow in the lab, looking for genes required in border cells migration. I
focused on genes required in the germ line that affect border cell migration as well as
oocyte and embryo development.
3.1.1
3.1.1.1
Screening method
Generation of PiggyBac mutants in FRT background
Mutations in fly can be induced by transposon insertion or EMS (ethyl methanesulfonate).
EMS can induce unbiased mutations but mapping those mutations is very time consuming.
For transposon mutagenesis, the interrupted genes can easily be cloned by inverse PCR.
However, different transposons display different preferential sites biasing the screen at
certain hot spot (Liao et al., 2000). The P element is the most commonly used transposon
in Drosophila screens. In order to perform the FLP-DFS system (see material and method,
Figure 7) to create homozygous mutant germ line cells, we wanted to generate transposon
insertions in FRT background (Chou and Perrimon, 1996). Unfortunately, P-elements can
not be used in FRT background because the FRT sequences are inserted into the genome
using by P elements. Therefore, another transponson, the PiggyBac which is not disrupted
FRT site, was used as a mutagen in our screen. The original PiggyBac line used was on X
chromosome so new PiggyBacs insertions in the autosomes could be selected, and readily
segregated from the original one. Later we performed a mapping cross to localize the
insertions to a chromosome arm.
3.1.1.2
Mapping
We mapped the chromosome arm on which the PiggyBacs were inserted using the
eyFLP-FRT system (Figure 6). eyFLP-FRT drives expression of the FLP-recombinase
54
RESULTS
with the eyeless promoter (in the eye) and homozygous mutant clones (mosaic eyes) were
created only in eye (the other tissues are still normal). Depending on which flies
displayed mosaic eyes, we could distinguish the chromosome PiggyBac was inserted in.
For example, if the PiggyBac was inserted in FRT42 chromosome, the only mosaic eyed
flies were those lacking CyO in the X2 cross in (Figure 9).
3.1.1.3
Germ line clone analysis
After identification of the chromosome the PiggyBac was inserted in, we made stocks for
each mutant line. I then collected mutant virgins to make germ line clones (the FLP-DFS
system, Figure 7) and further analyzed the mutant phenotype. I used a slbo: LacZ marker
which labels the border cell cluster to test for migration delays. To test for embryogenesis
defects, I selected the crosses which produced dead embryos and made cuticle
preparations to check for cuticle defects. If the eggs were unfertilized, I analysed ovaries
stained with phalloidin and DAPI (labelling F-actin and DNA) to check for any obvious
oogenesis defects.
3.1.1.4
Cloning the genes affected by the PiggyBac insertions
For mutants with interesting phenotypes, I used inverse PCR (IPCR) to isolate the
flanking sequence of the insertion. I then blasted the sequences against the Drosophila
genome to identify the insertion site.
3.1.1.5
Creation of new ovoD1 lines
In order to generate germ line clones, FRT40ovoD1, FRT42DovoD1, FRT80ovoD1, and
FRT82ovoD1 were required to create germ line mutant clones (Chou and Perrimon, 1996).
But FRT42ovoD1 and FRT80ovoD1 were not available. It is not possible to create these
lines using recombination in females because ovoD1 is a dominant female sterile
mutation. I chose X-ray to induce homologous recombination of FRT sites with ovoD1.
First, FRTG13(42B)ovoD1 (w+) and FRT2A(79A)ovoD1 (w+) males were crossed with
eyeflp;;FRT42D (w-) virgins, and eyeflp;;FRT80 (w-) virgins separately. When progenies
55
RESULTS
reached the second or third instars larvae, they were irradiated with an X-ray dose of 1000
rad (4 mA, 100 kV, 3'18'', Philips MG102) to induce homologous recombination, including
in the germ line. Using the eyFLP-FRT system, I confirmed the presence of the FRT site
and ovoD1 in the genome. I successfully generated 2 lines of FRT80ovoD1 and 1 line of
FRT42DovoD1.
Figure 9. The scheme of crosses
PB/PB; FRT4042;FRT 8082
Jump cross
X
w/Y;If,Tase/CyO;FRT8082
PB/Y; If,Tase/ FRT4042; FRT8082 X W/W
eyeFlp;FRT40/CyO;FRT80/FRT80
+
X 1 W/Y;FRT4042/+;FRT8082/+ (W )
Mapping
eyeFlp;FRT42*/CyO;FRT82/TM3ser
X2
For example, if there are no mosaic eyes in X1, but in files with ser in X2, it means there is an insertion on
FRT42 chromosome.
hsFLP slbolacZ/Y; FRT42ovoD1/CyO X
hsFLP slbolacZ/Y; FRT40ovoD1/CyO
hsFLP slbolacZ/Y; FRT80ovoD1/TM3
hsFLP slbolacZ/Y; FRT82ovoD1/TM3
eyflp/w; FRT4042/CyO
LHS
Pick up hsflpslboLacZ/W; FRT4042*/FRT42ovoD1 and put them into vials with yeast.
X gal staining for border cells migration
Cuticle preparation for embryo defects
Phallodin and DAPI staining for oogenesis defects
56
RESULTS
3.2
Screen result
We produced over 5000 individual PiggyBac insertions. From them, I checked about
3000 lines for defects in border cell migration, oogenesis or embryogenesis by making
germ line clones. Unfortunately, I only found two mutants which were required in the
germ line for border cell migration. The first mutant is an allele of cornichon, which is
required for early Gurken signaling. Mutant egg chambers show a second, posterior set of
border cells, as the posterior follicle cells are not specified and assume an anterior follicle
cell fate (Roth et al., 1995). The second mutant is an allele of cup, which is a translation
repressor of Gurken, and Oskar (Chekulaeva et al., 2006; Wilhelm et al., 2003). Mutant
cup egg chambers show border cell migration defect and oocyte growth defect. Border
cell migration defect may be a secondary effect, due to abnormal oocyte growth. Below, I
will focus on other classes of mutants that I found in the screen. Mutants displaying
embryonic phenotype or oogenesis defects are listed in Table 2. I divided all mutant lines
into several groups using the well-established criteria: No maternal effect, Germ cell
lethal, abnormal oogenesis and embryonic defects (Perrimon et al., 1989; Perrimon et al.,
1996).
3.2.1
No maternal effect
All PiggyBac mutants in this class can lay eggs which are fertilized and hatched. This
group corresponds to genes that are either not expressed in germ line or not required for
oogenesis and embryogenesis. About 90% PB mutants belong to this group.
3.2.2
Germ cell lethal
In this class, mutant females have ovaries similar to ovoD1, in that oogenesis stops at
very early stage. I identified 300 mutant lines in this category. This group corresponds to
genes that may require for germ cell viability or early oogenesis (Perrimon et al., 1989).
Some of the mtants belonging in this class may be present due to technical reasons, such
as mapping problems. If the mutation was mapped to the wrong chromosome, failure of
recombination will lead to germ cell lethal phenotype (as no clones will be made).
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RESULTS
3.2.3
Abnormal oogenesis
Mutations in this class show ovaries that can develop further than ovoD1 but with some
defects in the oogenesis. I observed several different kinds of oogenesis defects in these
mutants. I divided these mutants into several classes depending on phenotypes.
3.2.3.1
Nurse cell membrane defects
In mutants of this class some of nurse cell membranes are absent based on phalloidin
staining. One such mutant: PB1173, an allele of sds22, mutant eggs can be fertilized and
hatch. Most of PB1173 egg chambers show some nurse cell nuclei that are close each
other and lack any phalloidin-labelled cell membranes between them (Figure 10B). In
yeast, sds22 regulates protein phosphatase 1 (Ceulemans et al., 2002). In Drosophila,
protein phosphatase 1 regulates nonmuscle myosin to control actin cytoskeleton
(Vereshchagina et al., 2004). I proposed that sds22 may affect nurse cell membrane
formation through protein phosphatase 1.
Figure 10. Nurse cell membrane defects
Green is DAPI staining and Red is phalloidin staining in (A) wild type egg
chamber and (B) PB1173 mutant egg chamber.
3.2.3.2
Gurken signaling defects
The dorsal-ventral axis of eggs and dorsal appendages are determined on the second wave
of Gurken signaling. I isolated five mutant alleles of squid, a heterogeneous nuclear
RNA-binding protein, which controls dorsoventral (DV) axis formation during
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RESULTS
Drosophila oogenesis by localizing gurken (grk) mRNA (Kelley, 1993; Norvell et al.,
1999). In late wild type egg chamber, Gurken is normally localized in the anterior-dorsal
part of egg chamber to control formation of two dorsal appendages (Figure 11A, B, B'). In
PB4180 mutan, there are more dorsal appendages due to Gurken localizing not only to the
anterior-dorsal site but also at ventral site of mutant egg chambers (Figure 11C, D, D').
Another mutant, PB4329, affects cornichon, which is required for the first Gurken
signaling which is establishes the anterior-posterior polarization of the oocyte in
mid-oogenesis and later in dorsal-ventral patterning (Roth et al., 1995). In wild type egg
chambers, dorsal appendages and one group of border cells are present at the anterior part
of egg (Figure 11E). In cornichon mutant egg chamber, posterior terminal follicle cells
are transformed to anterior follicle cells, forming another set of border cells at the
posterior part. Dorsal appendages are absent in this mutant (Figure 11F).
Figure 11. Gurken signaling defects
Dorsal appendages in wild type eggs (A) or PB4180 eggs (C). Antibody staining detecting
Gurken in wild type (B) or PB4180 mutant germ line egg chamber (The mutant clone is
marked by lack of GFP, shown in green) (D). Gurken antibody staining (red). (B', D') single
channel of Gurken antibody staining (E, F) arrows indicate border cells, marked by X-gal. One
cluster is present in wild type but two in PB4329 germ line clone egg chamber.
3.2.3.3
Dumpless eggs
The dumpless phenotype is seen when nurse cells fail to transport protein and mRNA into
the oocyte. The oocyte is smaller than in a normal egg chamber, but follicle cells appear
normal. Later, nurse cells may fail to degenerate. One such mutant, PB4057, in the
Bullwinkle locus, shows such a phenotype (Figure 12B). This mutant affects not only
dumping in oogenesis but also anterior-posterior axis formation in embryogenesis
59
RESULTS
(Rittenhouse and Berg, 1995).
I found two mutants of cup, a translation repressor, in which egg chambers have smaller
oocyte (Figure 12 D) and border cells are unable to attach to the oocyte at the right time.
As the Cup protein normally represses the translation of axis determinants such as Oskar,
and Gurken, loss of cup results in reduced oocyte size. Border cell migration defects may
be a secondary effect due to the oocyte growth defects.
Figure 12. Dumpless
phenotype
(A) Late stage wild type
egg chamber with normal
dorsal appendages. (B)
Late stage PB4057
mutant egg chamber.
Arrows indicate smaller
oocyte and
non-degenerated nurse
cells. Stage 10 egg
chamber have a large
oocyte in wild type (C) or
smaller oocyte in PB4975
mutant egg chamber (D).
Green is DAPI staining,
and red is phalloidin
staining.
3.2.3.4
Abnormal nurse cells
An abnormal number of nurse cells may due to egg chamber fusion or cell cycle defects.
Wild type egg chambers are composed of 15 nurse cells and one oocyte (Figure 13A).
PB2955 affects neuralized, which encoded an E3 ubiquitin ligase. PB2955 egg chambers
showed more than 15 nurse cells and one oocyte (Figure 13B).
Figure 13. Abnormal
nurse cells
Stage 10 egg chamber
with 15 nurse cell nuclei
(A) or more nurse nuclei
in PB2955 germ line
clone (B). DAPI staining
is in blue and phalloidin
staining is in red. Mutant
clone is marked by lack
of GFP (green).
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RESULTS
3.2.3.5
Others
This class contains females with germline clones that can lay unfertilized eggs. There are
no obvious oogenesis defects by phalloidin staining. To analyze this class further, I
examined the localization of the axis determinants in some mutant ovaries. In wild type
stage 9-10 egg chamber, Oskar localizes to the posterior pole of the oocyte (Figure 14A).
About 10% of PB3975 egg chambers showed Oskar mislocalization (Figure 14B).
PB3975 affects cropped which is predicted to be a transcription factor. I also observed
Oskar anchoring defects in >10% of PB4949 egg chambers (Figure 14C). PB4949 affects
pipsqueak, a predicted transcription factor.
Figure 14. Oskar localization defects
(A) Wild type egg chamber. (B) PB3975 germ line mutant egg chamber. (C) PB4949 mutant egg
chamber. Green is Oskar antibody staining, and red is phalloidin staining.
3.2.4
Mutations affecting embryo development
RNA and proteins produced by the nurse cells also contribute to early embryonic
development. In this screen, I also identified genes required in the germ line that affect
embryogenesis by checking the cuticle phenotype of embryos. I divided these mutations
into 3 different classes.
3.2.4.1
Posterior group
A general feature in this class is that embryos lack abdominal segments and only contain
the head and tail structure (Figure 15). Posterior group genes are involved in
determination of germ cell fate and posterior cell fate in the embryos. PB456 affects hrp
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RESULTS
48, an RNA binding protein, which is already known to affect oskar RNA localization
(Figure 15D) (Huynh et al., 2004; Yano et al., 2004). Four mutants, PB122, 680, 1094,
3747, affect vasa, which is essential for the assembly of pole plasm, a special type of
cytoplasm found in the posterior portion of the egg and early embryo (Figure 15E)
(Schupbach and Wieschaus, 1986).
Figure 15. Posterior group phenotype
Anterior is left, and posterior is right in all panels. (A) wild type embryo cuticle. (B) PB83. (C)
PB2265. (D) PB456. (E) PB122, PB680, PB1094, PB3747. (F) PB3848. (G) PB4053. (H)
PB5257.
3.2.4.2
Segmentation defects
A general feature in this class is that the number of denticle belts is reduced or denticle
belts are fused or mispatterned (Figure 16). Embryos from PB5090, affecting eIF4E, a
translation initiation factor, show a pair-rule gene mutant like phenotype (Figure 16H).
The eIF4E mutant enhances the phenotype of mutants in ftz, a pair-rule gene. But Kankel
et al. did not see a pair-rule gene mutant like phenotype in their eIF4E mutant (Kankel et
al., 2004). This may be due to the different alleles of eIF4E used.
One mutant in this category, PB2691, affects CG17090 which encodes HIPK 2 kinase.
PB2691 are homozygous viable but embryos from homozygous females show denticle
belt
fusions
(Figure
P{GT1}CG17090
BG00855
17B).
Another
transposon
inserted
in
BG00855
, is homozygous lethal. P{GT1}CG17090
this
region,
and PB2691 fail
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RESULTS
to complement with each other or a deficiency lacking this region (Figure 17C). This
suggested that both PB2691 and P{GT1}CG17090BG00855 affect CG17090.
Figure 16. Segmentation defect
Anterior is left, and posterior is right in all panels. (A) wild type cuticle. (B) PB3296. (C)
PB772. (D) PB1210. (E) PB284. (F) PB4350. (G) PB4399. (H) PB5090. (I) PB4592
In Drosophila embryogenesis, the A/P axis of the fly is structured by both the gradient of
Bicoid and the posterior system that gives rise to Nanos localization (Nusslein-Volhard
and Roth, 1989). Both Bicoid and Nanos gradients determine the location of gap gene
transcription, such as krupple, knirps, and giant. Gap genes control the striped pattern of
pair rule genes like even-skipped (eve), and fushi tarazu (ftz). These striped pattern leads
to the segmental expression of segment polarity genes such as wingless, hedgehog, and
engrailed (en) (Figure 17O). To test whether PB2691 is directly involved in segmentation,
I stained mutant embryos for En. Wild type stage 10-11 embryos express En in 14 stripes
(Figure 17D). Some specific En stripes are missing or discontinuous in the PB2691/Df
mutant embryos (Figure 17E). It suggested that HIPK2 is required in segmentation. As
segment polarity genes are induced by Pair-rule genes, the loss of some En stripes may be
due to a pair-rule gene defect. I examined expression of the pair-rule proteins, Eve and
Ftz in PB2691/Df flies. Wild type embryos show 14 evenly spaced stripes, 7
Even-skipped stripes alternating with 7 Fushi tarazu stripes (Figure 17F). In PB2691/Df
embryos, some specific pair-rule stripes are wider or narrower in mutant background and
the distance between them is not always equal (Figure 17G). It is known that Pair-rule
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RESULTS
gene regulation can be auto-regulatory (Schier and Gehring, 1992). To determine whether
PB2691 affected gap gene or pair-rule autoregulation, I checked the expression of the
Pair-rule genes PB2691/Df embryo by in situ. Both eve and ftz in situ patterns are similar
to the protein expression pattern in PB2691/Df background (Figure 17 I, K). This suggests
that PB2691 affects gap gene or the regulation of pair-rule genes by Gap protein, but not
pair–rule gene autoregulation. To further test this, I check gap gene expression in the
mutant background. Preliminary data suggests, that gap gene expression is not affected
(Figure 17L, M). This suggested that PB2691 does not affected gap gene expression
pattern but may affect the gap gene regulatory network. Zhang et al. found that HIPK2
negatively regulates CtBP (Zhang et al., 2003). And CtBP, a transcriptional co-repressor,
is involved in Gap protein repression in Drosophila (Nibu and Levine, 2001; Nibu et al.,
1998; Strunk et al., 2001). These data suggest that the role of HIPK2 may warrant further
study.
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RESULTS
Figure 17. PB2691 mutant phenotype
(A) wild type cuticle with head structure, tail structure and eight denticle belts. (B,C) PB2691 (B) or
PB2691/Df (C) cuticle still form head and tail structures but show disordered segmentation.
Antibody staining detecting En (red) in wild type (D) or PB2691/Df (E). Arrows indicated these
engrailed stripes that are missing or discontinuous. Antibody staining against Eve (green) and Ftz
(red) in wild type (F) or PB2691/Df (G) . (G) Arrows indicated where the Ftz or Eve stripes are
wider or narrower. Note that more cells are co-stained with Ftz and Eve. In situ hybridization
detecting ftz in wild type (H) or PB2691/Df (I). Arrows indicated that ftz stripes are narrower and the
distance of two stripes is not in order. In situ hybridization detecting eve in wild type (J) or
PB2691/Df (K). Arrows indicated where eve stripes are narrower and the distance of two stripes is
altered. Antibody detecting krupple (green) and knirps (red) in wild type (L) or PB2691/Df (M). (O)
Cartoon of embryo segmentation. Adapted from Dynamic development.
3.2.5
Others
This class contains mutants whose phenotype does not belong to any of the above classes.
PB3292 affected the transcription factor dorsal, which activates and represses zygotic
genes responsible for dorsal-ventral axis patterning in early stages of development. In the
dorsal mutation, ventral cell fates are not formed, and embryos adopt a dorsal cell fate.
PB3292 mutant embryos show the dorsalization phenotype (Figure 18B). Embryos from
PB4170 or PB4496 die at early stages and do not form cuticle (Figure 18E, F).
Figure 18. Others
Anterior is left, and posterior is right in all panels. (A) wild type cuticle. (B) PB3292,
dorsalized cuticle. (C) PB4017 (D) PB28, smaller embryo. (E) PB4170. (F) PB4496. (E,F)
early embryo defects. (G) PB2600, germband retraction defect. (H) PB4211, dorsal opened
embryo. (I) PB4889, opened embryos.
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RESULTS
3.3
3.3.1
Ensconsin
PB4170 affects Ensconsin
From the screen, I retrieved one PiggyBac line, PB4170, which is homozygous viable but
embryos laid by homozygous females arrest at early embryo stages (Figure 19B). By
inverse PCR, I found that PB4170 is inserted in the second intron of CG14998 (ensconsin)
(+1476 relative to transcription start site). I obtained another P element line,
P{RS3}CB-5457-3 which is inserted in the first exon of CG14998 (at +30 relative to
transcription start site) from the Bloomington stock center (Figure 19E). The embryonic
phenotype of P{RS3}CB-5457-3(PCB5457) is similar to that of PB4170. To confirm that
the phenotype is caused by the transposon insertions, I did a complementation test
between PB4170 and PCB-5457 or between PB4170 and a deficiency(Df) removing
ensconsin and other neighboring genes. Embryos from both trans-heterozygous females
gave embryos which died at early stages, like PB4170 (Figures 19C, D). This suggests
that both the PiggyBac and P element alleles affected enconsin, which is required in germ
line for embryogenesis.
Figure 19. PB4170 embryo phenotypes and genomic structure
(A, B, C, D) cuticle preparation. (A) Wild-type embryo. (B) Embryos from PB4170 females.
(C) Embryos from PB4170/Df females. (D) Embryos from PB4170/PCB5457 females. (E)
Schematic of isoform B of ensconsin and two arrows indicated the PB4170 insertion and
PCB5457 insertion. The black bar marks the coding region; the white bar shows
untranslated regions. The two blue bars mark regions conserved with human E-MAP115.
3.3.2
N-terminal region of Ensconsin can bind to microtubules
The ensconsin transcript is predicted to form five different splice forms (A, B, C, D, and
66
RESULTS
E) which differ in the 5’ region and share most of 3’ region. I found by RT-PCR that
isoform B is the most highly expressed in wild type ovaries. In PB4170 mutant ovaries,
two more splicing forms are seen within the PiggyBac sequences, confirming that the
PiggyBac insertion affects ensconsin (ens).
Ensconsin is predicted to be a microtubule associated protein with two conserved, highly
charged regions, EHR1 and EHR2. There are two mouse homologues, E-MAP-115 and
RPRC1 (arginine proline rich coiled-coil protein 1). Vertebrate E-MAP-115 has already
been analyzed. When compared to the mouse homologue, E-MAP-115 shows 31%
identity and 60% similarity in the EHR1, and 43% identity and 62% similarity in the
EHR2 domain (Figure 20A). Masson and Kreis identified human E-MAP115 as a
microtubule binding protein in cell culture (Masson and Kreis, 1993). To test whether
Drosophila Ensconsin is also a microtubule binding protein in Drosophila, I performed a
microtubule binding assay with Taxol (Karpova et al., 2006). Taxol, a yew tree extract,
Figure 20. N-terminal of Ensconsin can bind microtubules
(A) Schematic of Ensconsin protein. EHR1 (Ensconsin homologue region 1) is shown as the
yellow bar, and EHR2 (Ensconsin homologue region 2) is represent by the green bar. White bar
represents Drosophila protein and gray bar is mouse E-MAP-115. Black bars mark the A and B
antigens region. Identity and similarity between Drosophila and mouse E-MAP-115 are shown
below the gray bar. (B, C, D) Microtubule binding assay showing supernatant (S) or pellet (P)
after incubation of cleared exact with either Taxol or DMSO control. (B) 0-2 hours wild type
embryo extracts (C, D) ovaries from tub-Ens-x or tub-Ens-C transgenic females. The blot was
probes with anti-Ensconsin B and anti-tubulin.
67
RESULTS
binds and stabilizes microtubules. When microtubules are polymerized and stabilized
with taxol, they can be spun down as a pellet. If a protein is microtubule associated, most
of the protein will pellet with the microtubules. Such microtubule binding assay is
frequently used to identify microtubule associated protein (MAPs). For microtubule
binding assay, I produced two antigen fragments of E-MAP-115: antigen A (284-499
amino acids) and antigen B (767-851 amino acids) (Figure 20A). I used these to
immunize rats and generate two polyclonal antibodies. When this assay is performed
using taxol treated 0-2 hours wild type embryo extracts, most microtubules are
sedimented, demonstrating that Taxol functions in this experiment. In this experiment I
found that Ensconsin co-sediments with microtubules. This suggested that full length
Ensconsin can bind microtubules in embryonic extract (Figure 20B).
The EHR1 domain of the human protein (77-228) is essential for microtubule binding
(Masson and Kreis, 1993). To examine which regions of Drosophila Ensconsin are
important for microtubule binding, two constructs, lacking either the EHR2 (Ens-X) or
EHR1 domains of Ensconsin (Ens-C) were created by Pernille Rørth. From these, I
generated transgenic flies which express the transgenes under control of tubulin promoter.
I found that Ens-X could co-sediment with taxol treated microtubules (Figure 20C), but
Ens-C could not. (Figure 20D). It has been suggested that the N-terminal region of
Drosophila Ensconsin is required for microtubule binding as human homologue.
3.3.3
20% of PB4170 egg chambers showed Oskar and Staufen
mislocalization
Microtubules are important for the transportation of axis-determinants during oogenesis.
As Ensconsin is a microtubule binding protein, it seemed logical to test whether it played
a role in these events. I examined the localization of the axis determinants Oskar and
Gurken, in PB4170 mutant ovaries. In wild type, from stage 9, Oskar protein is translated
at, and localized to the posterior pole of the oocyte (Figure 21A'). Around 20% of PB4170
egg chambers show Oskar accumulation in the middle of the oocyte (Figure 21 D'). Oskar
protein localization depends on its mRNA localization. I therefore analyzed oskar mRNA
localization by in situ hybridization and Staufen antibody staining. Staufen, a RNA
binding protein, binds oskar RNA in the oocyte. Normally, in stage 9 egg chambers, oskar
68
RESULTS
mRNA is localized to the posterior pole of the oocyte in a pattern similar to Oskar protein
(Figure 21A) and Staufen is also found at the posterior pole (Figure 21B). oskar mRNA
and Staufen also are mislocalized in 20% of PB 4170 egg chambers (Figure 21D,E). I also
checked Gurken antibody staining. Gurken is usually localized to the anterior-dorsal site
of the oocyte and the protein extends along the oocyte cortex to posterior pole (Figure
21C). In PB4170, Gurken is restricted to the anterior-dorsal side and extends less towards
the posterior pole, and some punctate stainings are also found in the middle of the oocyte
(Figure 21F). Interestingly, the localization of bicoid is unaffected (data not shown). This
shows that the localizations of oskar mRNA, Oskar and Gurken proteins are affected, but
localization of bicoid is not affected in PB4170 mutants.
Figure 21. Oskar, Staufen and Gurken are mislocalized in PB4170
(A, B, C) wild type egg chambers. (D, E, F) PB4170 homozygous mutant egg chambers. (A,
D) show oskar mRNA in situs. (A', D') show Oskar antibody staining. (B, E) show Staufen
antibody staining. (C, E) red is Gurken antibody staining, green is DAPI and blue are
phalloidin. (C', E') only Gurken antibody staining is shown.
3.3.4
Generating new alleles of ensconsin
Western blots and immunostaining of PB4170 ovaries show that the mutant is not a
protein null allele. To fully investigate the function of Ensconsin, I created two new
mutant alleles, ens ∆C and ens ∆N. ens ∆N was created by imprecise excision of PCB-5457.
ens ∆N (removing from +31-+1475 of the gene) homozygous flies are viable but embryos
laid by homozygous females die at early stages, much like PB4170 (Figure 22A). ens ∆C
was created by end-out homologous recombination (Gong and Golic, 2004). In ens ∆C,
two third of Ens coding region, including EHR2 are deleted and replaced by the
mini-white cassette (from +7077 to +9632, Figure3-13A). ens ∆C homozygous flies are
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RESULTS
viable but sick and most display wing eversion defects (Figure 22C). Embryos laid from
homozygous females die at early stages. ens ∆C mutant phenotypes can be completely
rescued by tub-ens. It suggests that the phenotypes of ens ∆N and ens ∆C are caused by the
deletion of ens. Western-blot analysis of mutant ovary extracts indicated that the mutant
had decreased levels of Ensconsin protein (wild type>PB4170>ens∆N>ens∆C ) (Figure
22B). ens ∆C protein levels are hard to measure as the antigen region is removed in this
mutant. In ens∆N mutant ovaries, the Ensconsin protein still present may due to alternate
splice forms. I decided to perform most subsequent experiments with ens∆C flies.
Figure 22. New mutant alleles of ensconsin
(A) Schematic of ensconsin gene structure. (B) Western blot of different mutants.
∆N
Ensconsin is about a ~150kD protein. In PB4170, and ens , protein level is reduced. Note
∆C
that the epitope is absent in ens . Tubulin is shown as a loading control. A Star indicates a
∆C
background band. (C) Left is wild type adult, and right is ens adult. Both are 1 day old. In
∆C
ens , the wing can not properly expand.
3.3.5
Oskar, Staufen and Gurken localization are affected but
bicoid is not in ens∆C mutant
My experiments with PB4170 suggest that Ensconsin is required for Oskar and Gurken
localization. I decided to re-examined Oskar, Gurken, and bicoid in ens∆C mutants. ens∆C
mutant stage 9 egg chambers show less accumulation of Staufen and Oskar protein at the
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RESULTS
∆C
Figure 23. ens mutant phenotype
∆C
∆C
(A-F) in situ hybridization of oskar in wild type (A) or in ens (B), bicoid in wild type (C) or in ens
∆C
(D), and gurken in wild type (E) or in ens (F). (G) Quantification of dorsal appendages in w,
∆N
∆C
∆C
PB4170, ens , and ens . (H-K) antibody staining at stage 9 in wild type (H) or in ens (I), at
∆C
stage 10B or 11 egg chamber in wild type (J) or in ens (K).Oskar staining is in green and Staufen
in red. (L, M) nuclei marked by WGA staining and F-actin marked by phalloidin in the wild type (L)
or in ens∆C (M) late stage 9 egg chambers. Arrows indicate the oocyte nucleus and abnormal
accumulation of F-actin. (N, O) antibody staining detecting Gurken (green) in wild type (N) or in
ens∆C (O). The oocyte nucleus can be seen as the round, unstained area, slightly mislocalized in
(O)
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RESULTS
posterior pole. Staufen is also seen around the oocyte cortex (Figure 23 I, I’). In ens∆C
stage 10 egg chambers, both Staufen and Oskar showed a diffuse staining along a large
section of the anterior cortex (Figure 23K, K’). Oskar protein is still translated and
remains attached to the oocyte membrane. This suggests that Ensconsin is not involved in
Oskar translation or anchoring. Normally, oskar mRNA is accumulated at the posterior
pole of the oocyte at stage 10 (Figure 23A). In mutants, oskar mRNA, like Oskar protein
is spread along the anterior cortex (Figure 23B). This confirms that Ensconsin is required
for oskar localization.
gurken mRNA and protein were localized in the anterior-dorsal corner of egg chambers in
wild type egg chamber (Figure 23N and E). In ens ∆C mutant egg chamber, gurken mRNA
levels appear to be reduced but still correctly localized (Figure 23F). Gurken protein was
observed not only to be loosely cortical localization at anterior-dorsal site but also in the
middle or posterior parts of the oocyte (Figure 23O). Gurken signaling is required to
reorganize the microtubule network. This allows the oocyte nucleus to migrate from the
posterior pole to the anterior-dorsal side of the oocyte at mid-oogenesis. Wheat germ
agglutinin (WGA) is a carbohydrate-binding protein that selectively recognizes sialic acid
and N-acetylglucosaminyl sugar residues which are predominantly found on the plasma
membrane. I used WGA-conjugated with FITC to label the oocyte nucleus membrane. In
wild type egg chambers, all oocyte nuclei localize to the anterior-dorsal part of the oocyte
(Figure 23L). Around 10% of ens ∆C mutant egg chamber show a strong mislocalization of
the oocyte nucleus and the others show weaker mislocalization (Figure 23O). Abnormal
F-actin accumulations are also found within the oocyte in ens ∆C mutant egg chambers
(Figure 24M).
Around 50% of ens ∆C eggs showed dorsal appendage fusions which corresponds to the
mis-localization of Gurken but few fusions are seen in ens∆N or PB4170 egg chamber
(Figure 23G). bicioid mRNA remains localized to the anterior of the oocyte in ens∆c
mutant egg chamber (Figure 23D). This confirms that Ensconsin is required for Oskar and
Gurken localization, but not bicoid.
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RESULTS
3.3.6
khc interacts genetically with ensconsin
The Oskar and Gurken mislocalization phenotypes of ens ∆C are similar to kinesin heavy
chain (khc) mutant phenotypes but weaker. Kinesin heavy chain is a microtubule motor
protein which moves its cargo toward the plus ends of microtubules. To test whether
ensconsin interacts with khc, I anaylzed a genetic interaction between ens∆C and khc27
(null allele). I observed a strong interaction between khc and ensconsin, as flies lacking
one copy of khc and homozygous for ens∆C are lethal. 10% of ens∆C egg chambers show
oocyte nucleus mislocalization phenotype, adding one copy of khc27 mutant in ens∆C germ
line clone background increases the oocyte nucleus mislocalization rate to 32%. This
suggests that khc interacts with ensconsin genetically.
3.3.7
Ensconsin affects posterior localization of Khc
Figure 24. Ensconsin affects Dhc localization through Khc mislocalization
∆C
Antibody staining of Khc (green) at stage 9 in wild type (A) or in ens (B). Antibody staining of
∆C
Dhc (red) at stage 10 in wild type (C) or in ens (D). Enlargements of posterior part of the egg
chamber (A', B', C', D').
As I found that khc interacts genetically with ensconsin, I wanted to test if Ensconsin
affects the localization of Khc. To examine this, I did Khc antibody staining in the ens∆C
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RESULTS
mutant background. In wild type stage 9 egg chambers, Khc is enriched the oocyte, and
strongly accumulated in the posterior pole of the oocyte (Figure 24A). However in ens∆C
mutants, Khc is still enriched in the oocyte, but no longer accumulated at the posterior
pole (Figure 24B). Dynein heavy chain (Dhc), a minus-end directed motor protein,
localizes to the posterior pole of the oocyte at stage 9-10 egg chamber. This localization
requires Dhc to be transported in the oocyte by Khc (Figure 24C) (Januschke et al., 2002).
Dhc is also mislocalized in stage 9-10 ens∆C mutant egg chambers (Figure 24D).
Interestingly, Dhc is involved in early oocyte specification, but I did not find similar
defect in the ens∆C mutants (McGrail and Hays, 1997). This suggests that Ensconsin
affects Dhc indirectly, through the localization of Khc to the oocyte.
3.3.8
Microtubule polarity is not affected in ens∆C mutant
∆C
Figure 25. Microtubule formation and microtubule polarity in ens mutant
∆C
Antibody staining of tubulin (green) at stage 9 in wild type (A) or in ens (B). Images of live,
∆C
early stage 9 samples expressing tauGFP in wild type (C) or in ens (D). X-gal staining of
∆C
Kinesin-lacZ at late stage 9 in wild type (E) or in ens (F).
The effect of Ensconsin on Khc and Dhc localization may be caused in several ways: (1)
microtubule formation defects, (2) microtubule polarity defects or (3) Impaired Khc
movement. To test whether Ensconsin affects microtubule formation and microtubule
polarity, I looked at microtubule staining and checked microtubule polarity markers in
ens∆C egg chambers. In wild type egg chambers, microtubules are enriched in the anterior
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RESULTS
part of the oocyte (Figure 25A). In ens∆C mutants, microtubules are also enriched in the
anterior region (Figure 25B). In wild-type oocytes, microtubules are organized in an
anterior to posterior gradient at stage 7–9 that can be visualized by using a Tau: GFP
fusion protein (Figure 25C) (Micklem et al., 1997). Tau: GFP expression pattern is
unaffected in ens∆C mutant oocyte (Figure 25D). Furthermore, microtubules are essential
for early oocyte specification, as no oocyte is seen in egg chambers by treating with
microtubule disrupting drugs (Theurkauf et al., 1993). The fact that ens∆C mutants still
can specify an oocyte implies that Ensconsin is not simply required to assemble or
maintain microtubules. To check microtubule polarity, I used Kinesin-LacZ, LacZ fused
to Khc, lacking the C-terminal regulatory domain (Clark et al., 1994). This fusion protein
is enriched at the posterior end of the oocytes and marks microtubule plus-end in wild
type stage 9 egg chamber (Figure 25E). Kin-LacZ was unaffected in ens∆C mutants
(Figure 25 F). This suggests that Ensconsin does not affect microtubule polarity.
3.3.9
Ooplasmic streaming is affected in ensconsin mutants
In late stage 10 egg chamber, the nurse cells start to pump RNA and proteins into the
oocyte by a process referred to as cytoplasmic dumping. At the same time, the oocyte
begins ooplasmic streaming. This process utilizes microtubules and motor protein to help
distribute the content from the nurse cell within the oocyte (Theurkauf et al., 1992).
Without ooplasmic streaming, nurse cells are still able to pump RNA and proteins into
oocyte, but the lack of mixing causes a stratified ooplasm (Serbus et al., 2005). This
stratification is due to an accumulation of yolk granules near the posterior of the oocyte
and leaving a clear zone at anterior zone. This streaming can be visualised by imaging
and tracking the large, yolk-filled endosomes in live egg chambers (Figure 26A). The
speed and onset of streaming are sensitive to levels of Khc. Several khc point mutants
display reduced or completely halted ooplasmic streaming. khc mutant egg chambers also
show stratified ooplasm. As Ensconsin affects Khc localization, I decided to examine
ooplasmic steaming in ensconsin egg chambers. In the wild type egg chambers, yolk
granules are equally distributed (Figure 26C). All ensconsin mutant egg chambers show
stratified ooplasm (Figure 26D). This suggests that ooplasmic streaming is blocked. To
confirm this, I recorded of ooplasmic streaming. 7/8 wild type egg chambers show normal
streaming (Figure 26A) but 16/16 PB4170 egg chambers showed streaming defects
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RESULTS
(Figure 26B). In khc27/+; ens∆C/+, there are around 10% stratified egg chambers. It
suggested that Ensconsin affects Khc-dependent transport.
Figure 26. Ooplasmic streaming is abolished in ens mutants
(A,B) Yolk granules movement visualized by overlay multiple frames from movies from
stage 11 (A) wild type or (B) PB4170 mutants. The moving particles are seen as
streaming in (A), but not in all ensconsin mutants (B). (C,D,E)Nomarski images of late
stage egg chamber from control (C), PB4170 (D), and transheterozygous
27
khc /+;ens∆C/+ (E) females. The dark material is yolk granules. (F) Quantification of
stratified eggs in different mutant.
3.3.10
Ensconsin is required in neurons
Khc-dependent transportation is not only required to localize axis determinants in the
oocyte, but is also for the delivery of synaptic components to the tips of axons. To
examine whether Ensconsin is also important in the neuron, I checked for potential
neuronal defects in ens∆C mutant. Most wild type flies climb to top of a vial in several
seconds. ens∆C mutant showed wing eversion defects and coordination problems
preventing them from climbing (Figure 27A). ens∆N mutants show normal wing eversion,
but also show climbing defects (Figure 27A). Both the wing eversion defects and
climbing defects can be rescue by expressing UAS-ens with elavGal4, a pan neuron driver.
Unsurprisingly, neuronal expression can not rescue the observed oogenesis defects
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RESULTS
(Figure 27A).
At late larval stages Ensconsin protein is also enriched in the axons connecting the optic
lobe to the eye disc (Figure 27B). This suggests that Ensconsin is required for neuronal
function. Synaptotagmin (Syt) is a major component of synaptic vesicles (DiAntonio et
al., 1993). In Khc mutants, Syt accumulates in the axons (Hurd and Saxton, 1996).
Ensconsin affecting Khc transportation in the oocyte and enrichment of Ensconsin in the
axon suggest that the neuron function of Ensconsin may also be to promote Khc
transportation. To look more directly at kinesin-dependent transport in neurons, I did Syt
antibody staining in ens∆C mutant larvaes. In wild type larvae, syt is efficiently
transported to synapses and no staining is seen in the axons (Figure 27C). In ens∆C, I see
accumulation of Syt along the axon indicating inefficient transport to the tip (Figure 27B).
This also suggests that Ensconsin is required for efficient kinesin-dependent transport in
multiple tissues.
Figure 27. Neuronal
functions of Ensconsin
(A) Climbing assay; A single
male was put into the empty
vial, tapped to the bottom and
their climb to the top was
timed. The graph shows the
cumulative success rate of
different genotypes. (B) HRP
staining, which marks axons in
red and Ensconsin antibody
staining (white in B’) of axons
in the optic stalk (C,D)HRP
(red) and Syt staining (green)
∆C
in control (C) or ens mutant
(D). Syt accumulation in the
axon shows impaired transport
(D).
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RESULTS
3.3.11
Ensconsin is localized to the anterior of the oocyte
Figure 28. Drosophila Ensconsin is tightly localized during oogenesis
Anterior is left in each panel. (A,B) In situ hybridization of endogenous ensconsin in early (A)
and stage 10 (B) egg chambers. (C,D) Antibody staining of endogenous Ensconsin protein
early (C) and at stage 10 (B). (E-G) Staining of Ensconsin when all follicle cells (E), the germ
line (F), or some follicle cells (G) are mutant (ens∆C). Mutant cells are marked by lack of GFP
(green); phalloidin (red). (E'-G') Show only the anti-Ensconsin channel. In situ hybridization
∆C
detecting transgenically expressed tub::ens (H-I) or tub::ens∆3’UTR (K,L) in ens . The
diffuse and delocalized signal in K and L is significantly above background. Staining of
∆C
∆C
transgenically expressed tub::ens (J) or tub::ens∆3’UTR in ens (M). ens alone shows no
signal by in situ or antibody staining.
Correct localization is important in the regulation of motor protein transport. From in situ
data, the ensconsin transcript is enriched in the oocyte from early stages (Figure 28A) and
localizes to the anterior of the oocyte from stage 9 (Figure 28B). Ensconsin protein is also
enriched in the oocyte from early stages (Figure 28C) and begins to form a gradient along
the cortex from anterior to posterior at stage 9 (Figure 28E, D). Ensconsin protein is also
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RESULTS
localized apically in follicle cells (Figure 28D). Apical localization of Ensconsin has been
reported in the mouse intestinal epithelium cells, indicating that this is a conserved feature
of this protein (Fabre-Jonca et al., 1998). This staining pattern is seen with both
antibodies I generated and no signal can be detected in ens∆C mutant cells (which lack the
epitope). (Figure 28E-G) The polarized localization of Ensconsin in the soma and germ
line is similar to the distribution of microtubule minus ends in these tissues.
To determine the mechanism of Ensconsin localization, I first focused on the mRNA. In
the cases of oskar and bicoid, the 3’UTR is an important determinant of RNA localization
(Kim-Ha et al., 1993; Macdonald and Struhl, 1988). I reasoned that as ensconsin has a
1.28Kb 3’UTR, this may play a role its localization. Pernille Rørth created tub:ens
constructs with and without the ensconsin 3’UTR, from these I generated transgenic flies.
When wild type ensconsin is expressed in ens∆C mutants, ensconsin mRNA and protein
are correctly localized (figure 28H-J). Expression of ensconsin without its 3’UTR in
ens∆C mutant results in mislocalization of the transcript (Figure 28K, L), but normal
protein localization (Figure 28M). This suggests that 3’UTR is required for RNA
localization, but that this is not essential for protein localization. It also suggests that there
is another mechanism regulating Ensconsin protein localization. The polar localization of
Ensconsin, in the follicle cells and the oocyte suggests that Par proteins may regulate its
localization.
3.3.12
Ensconsin localization is affected by Par-1
Par-1 is localized to the posterior pole of the oocyte and the basal-lateral domain of
follicle cells, and its localization does not overlap with Ensconsin. To test whether Par-1
regulates Ensconsin localization, I used a Par-1 hypomorphic combination (par-1W3 /
par-19A). This situation allows the need for Par-1 function in oocyte cell fate maintenance
in early oogenesis, allowing me to analyze its potential later role in Ensconsin localization.
In this genotype, most egg chambers are phenotypically normal (Figure 29A), but some
showing mislocalization of posterior markers such as Staufen. In Par-1 mutants,
Ensconsin localization is clearly affected and it is no longer excluded from the posterior
pole of the oocyte (Figure 29B). In par-1W3 mutant follicle cells (which lack any par-1
function), Ensconsin is no longer apically localized, but is evenly distributed throughout
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RESULTS
the cell (Figure 29C, D). This suggests that Par-1 functions, either directly or indirectly, to
restrict Ensconsin localization.
Figure 29. Ensconsin localization is restricted by Par-1
(A,B) Staufen (green) and Ensconsin (red) protein localization in Par-1 hypomorphic
mutant egg chambers (Par-19A/Par-1W3) at late stage 9. Only the posterior part of egg
chamber (including the oocyte) is shown. In some egg chambers, both staufen and
Ensconsin localization is affected at the same time. (A",B") An enlarged posterior image of
W3
the Ensconsin staining. (C,D) Ensconsin protein (red) in Par-1 mutant follicle cells.
Mutant clones are labeled by the lack of GFP (green); phalloidin marks F-actin (blue).
(C',D') Single channel image of Ensconsin staining. In (C), basal is up. (D) A tangential
section through the middle of the cells showing Ensconsin delocalized from the apical
domain in Par-1 mutant follicle cells.
3.3.13
Ensconsin is a direct target of Par-1
Par-1 may regulate Ensconsin localization directly or indirectly, for example, via
microtubule polarity. Human E-MAP-115 (Ensconsin) is a highly phosphoylated protein,
it is unclear which kinases it is a substrate for. Par-1 activity, together with the
phosphorylation induced binding of 14-3-3 (Par-5) to its target proteins, such as Bazooka
(Par-3) is required to restrict proteins to the apical domain of follicle cells. There is also
evidence that a similar mechanism acts downstream of Par-1 in the germ line. We
identified 6 conserved putative 14-3-3 binding sites in Ensconsin. To test whether
Ensconsin is a Par-1 direct target, I performed an in vitro kinase assay with Piyi Papadaki,
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RESULTS
a post-doctoral fellow in Anne Ephrussi’s lab. Several point-mutant constructs of
Ensconsin were generated by Pernille Rørth. I expressed these, and wild type Ensconsin
in bacteria and purified them. Using Par-1 kinase domain purified by Piyi Papadaki, we
first performed an in vitro kinase assay with purified Ensconsin and the kinase domain of
Par-1. We found that an N-terminal truncation of Ensconsin can be strongly
phosphorylated and a C-terminal fragment of Ensconsin was also somewhat
phosphorylated. The N-terminal fragment with mutated 14-3-3 binding sites is more
weakly phosphorylated (Figure 30B).
Figure 30. Ensconsin is a direct target of Par-1
(A) Schematic of Ensconsin constructs used. Red bars mark the predicted 14-3-3
binding sites. (B,C) Star indicated Ensconsin fragments. Top panel is the
autoradiogram and bottom is the Coomassie staining of the same gel. Molecular
weight markers are indicated on the left. Arrow indicates Par-1. (B) In vitro kinase
assay using recombinant Par-1 kinase domain and recombinant Ensconsin fragments.
M329G
fusion protein. With normal ATP,
(C) In vitro kinase assay using GFP-Par
phosphorylation of Ens-N is seen in the absence of the GFP-PAR-1 transgene.
To confirm the in vitro kinase data, we performed the in vitro kinase assay again, using
GFP-Par-1 immunoprecipitated from ovaries (Benton et al., 2002). Unfortunately, the
N-terminal Ensconsin fragment is phosphorylated in the control experiment, which lacks
the GFP transgene. This suggests that an endogenous kinase is non-specifically copurified
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RESULTS
and can phosphorylate Ensconsin. Piyi Papadaki had created a GFP-Par-1M329G mutant
which retains Par-1 kinase activity but only uses the ATP analogue, BenzoATP, as a
substrate.This approach has been used successfully to identify targets of other kinases
(Shah et al., 1997). We hoped that using GFP-Par-1M329G would reduce the background in
our assay. GFP-Par-1M329G was immunoprecipated from ovaries of transgenic flies and
mixed with bacterially purified Ensconsin fragments in the present of radio-labeled,
benzyl-ATP. In the control, lacking the GFP construct, no phosphorylation of the
Ensconsin N-terminal fragment was seen. As with the in vitro kinase assay, the
N-terminal fragment can be phosphorylated, but the C-terminal fragment and mutated
N-terminal fragments are weakly phosphorylated (Figure 30C). This suggests that
Ensconsin is a Par-1 direct target and that its 14-3-3 binding sites mediate much of this
phosphorylation.
3.3.14
Follicle cell polarity is not affected in ens∆C mutant
Par-1 affects follicle cell polarity (Doerflinger et al., 2003; Vaccari and Ephrussi, 2002).
Since Ensconsin is a direct target of par-1, Ensconsin may also affect follicle cell polarity.
To examine this, I made ens∆C mutant clones and checked the distribution of the polarity
markers aPKC (an apical marker) and
α Spectrin (a lateral marker). In large ens
-
∆C
clones, both markers were unchanged (Figure 31A, B). This suggests that Ensconsin does
not affect follicle cell polarity.
Figure 31. Ensconsin does not affect
follicle cells polarity.
(A) Antibody staining of aPKC (blue) in
∆C
ens . (B) Antibody staining of
∆C
-Spectrin (blue) in ens .(A',B') single
channel of aPKC (A') or spectrin(B').
(A,B) Mutant cells are labelled by the
lack of GFP; phalloidin staining is in red.
α
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RESULTS
3.3.15
The
Par-1
phosphorylation/14-3-3
binding
sites
in
Ensconsin are essential for its localization
Figure 32. Ensconsin is regulated by Par-1 and controls proper posterior marker
localization
∆C
(A,B,C) Localization of wild type or mutant tub::Ens, in ens mutants. In (A,B) arrows indicate
the anterior-dorsal accumulation of Ensconsin, which is reduced in Ens-mutant. Inserts show
the areas used quantification, and green is Staufen staining. (C) Follicle cells expressing only
the transgene (marked by lack of GFP, green) the adjacent cells are express both endogenous
and transgenic ensconsin. (D,E,F). Localization of wild type (D) or Ens-mutant, expressed in the
germ line using maternal-Gal4-VP16 driver. (F) The posterior region of the oocyte is shown at
high magnification. (D,E,F) Late stage 9 egg chamber with minimal endogenous Ensconsin
∆C
∆N
(ens / ens ) to visual the transgenic protein. Phalloidin is shown in blue
To investigate the significance of Par-1 phosphorylation in vivo, Pernille Rørth generated
constructs containing the tubulin protmoter driving either full length ensconsin (Ens-wt)
or ensconsin with mutations in the 6 14-3-3 binding sites and without the 3’UTR
(Ens-mutant) to remove any contribution of mRNA localization. I generated transgenic
flies of these constructs. Both constructs rescue the ens∆C mutant phenotypes, including
the wing phenotype and staufen mislocalization (Figure 32A,B). But when
tub::ens-mutant is expressed in the ens∆C background, the protein is less enriched in the
anterior of the oocyte and is less apical in follicle cells (Figure 32C). To quantify the
degree of mislocalization seen in the oocyte, I compared the intensity of staining in the
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RESULTS
same area of the anterior and posterior of the oocyte. Confirming my earlier observations,
I find 3 times more mutant protein at anterior than the posterior, where as for wild type
the difference is about 5-fold (Figure 32A, B). The remaining Ens-mutant signal at the
anterior could be attributed to other, unidentified par-1 phosphorylated sites.
As the tub::ens-mut was only partially delocalized in the oocyte, I tried expressing high
levels of the Ensconsin transgenes (using maternal tubulin-Gal4) in ens∆C / ens∆N flies.
Under this condition, Ensconsin-wt is found not only in the anterior region of the oocyte
but also at the posterior pole and in punctate staining throughout the oocyte (Figure 32D).
Under these conditions, Ens-mut is localized more to the posterior, with some egg
chambers lacking any protein at the anterior. In 50% of stage 9 mutant expressing egg
chambers, Ens-mut is strong mislocalized and Staufen is also mislocalized (Figure 32E).
In other Ens-mut egg chambers, Staufen still can localize to posterior pole of the oocyte
but no longer overlap with Ens-mut (Figure 32F), although it still appears mislocalized.
This suggests localization of Ensconsin to the anterior of the oocyte is controlled by a
saturable process, which may require Par-1 phosphorylation. Mislocalization of
Ensconsin leads to Staufen mislocalization, suggesting that the distribution of posterior
markers is controlled by Par-1, which acts via Ensconsin.
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4 DISCUSSION
DISCUSSION
4.1
The
genetic
screen
using
PiggyBac
Transposons
In this study, I performed a genetic screen for genes required, in the germ line, for
oogenesis and embryogenesis. From the screen, I isolated the mutant PB4170, which
affects ensconsin. I showed that the N-terminal region of Ensconsin can bind to
microtubules, but does not regulate microtubule formation or polarity. The ensconsin
transcript localization in the oocyte depends on its 3’UTR and Ensconsin protein
localization is regulated by Par-1 phosphorylation. In flies lacking Ensconsin, Oskar and
Gurken are mislocalized and this effect is due to the defects in Khc-dependent transport. I
propose that Ensconsin is a Par-1 regulated microtubule associated protein, required for
Kinesin-dependent transport.
4.1.1
The PiggyBac screen is effective but laborious
For this screen, we created around 5000 lines but I only managed to test 3000 with the
GLC system over the course of a year. There are several reasons for this. First, the eye
color of some PiggyBac lines was very weak, making mistakes in the mapping easy.
Second, any lines where the insertion lay between two FRT sites or on the fourth
chromosome could not be used for screening. Third, some mistakes cannot be avoided
during the mapping crosses. Although this screen was laborious; we found that the
phenotypes of some mutations showed identical phenotypes to those of known genes. For
example, I isolated dorsal, vasa, hrp48, and aubergine in the screen. This suggested, in
principle that our screen was working. It should be noted that since I have tested 3000
mutant lines and fly genome contain more than 10000 genes, this screen was not
saturating.
4.1.2
The PiggyBac and P element have similar preferential sites
It has been reported that PiggyBac transposons do not share hot spots or preferential
integration sites with P elements (Hacker et al., 2003; Thibault et al., 2004). In our screen,
I found several PiggyBac “hot spots” , for examples, squid (hit six times), vasa and
86
DISCUSSION
cropped (hit four times), and cup (hit three times). Most of these mutants that we found
interesting also have one or more P element insertions in them. This may reflect that the
near saturating level of P element screens but may also indicate that the insertion sites of
P element and PiggyBac element are perhaps not so different.
4.1.3
Germline clone analysis
From this screen, I found mutants required in germ line for oogenesis, embryogenesis and
border cell migration. The FLP-DFS system works well to analyze germ line function
because only egg chambers with mutant germlines can develop. For border cell migration,
very few genes are known to be involved in the germ lines that affect this process. In this
screen, only two mutants, cup and cornichon, were identified. This may be due to the
limited number of genes that are required in germline for border cell migration or because
relatively few mutants were tested in my screen. It is also possible that many of PiggyBac
mutants were simply not strong enough to cause border cell migration defects.
For oogenesis and embryogenesis, this screen was done by checking for unfertilized eggs
or dead embryos, which does not require dissecting flies, and therefore makes the screen
less labour intensive. Some mutants with high penetrance phenotypes affecting novel
genes like PB2691,PB4350 are worth further analysis. From the ~3000 mutants, I still
managed to find mutants affecting novel genes. This suggests that it may be worth to do a
systemic or saturating screen to study the germ line function in Drosophila.
87
DISCUSSION
4.2
4.2.1
Ensconsin: a microtubule associated protein
Ensconsin affects microtubules in a subtle way
Most of the reported microtubule associated proteins bind microtubules and affect their
stabilization. Tau knock out mice are viable, but show microtubule defects only in some
small calibre axons (Harada et al., 1994) and removal of MAP4 in fibroblast cells does
not cause microtubule defects (Wang et al., 1996). These may be due to a level of
redundancy in MAP functions. In E-MAP-115/Ensconsin knock out mice, microtubule
bundles appear to be thinner and less developed in spermatogenesis, but there are no
obvious defects in microtubule staining in ens∆C mutant egg chamber (Figure 25B,D). It is
still possible that Ensconsin affects microtubules in Drosophila egg chambers, but only in
subtle way and these defects can not be detectable by immunostaining or live imaging.
There may also be other, partially redundant MAPs in the oocyte, which can compensate
for some of the ens∆C mutant defects.
4.2.2
Ensconsin affects Khc-dependent transport
In this study, I found that Ensconsin affects Khc-dependent transport. If there was a strict
dependence of Khc on Ensconsin, then ensconsin mutant phenotype should as strong as
khc. However, ens∆C phenotypes are similar to khc mutant phenotypes, but weaker. It is
possible that the ensconsin mutants that I found are not null alleles, other redundant
proteins exist in the genome or Ensconsin increases the activity or funtion of Khc but is
not completely essential for its funtion.
Neither Ensconsin nor Khc are required for the polarity or morphology of follicle cells,
but affect oskar mRNA localization in the oocyte and vesicle traffic in neurons. It is
possible that because of the smaller size of follicle cells asymmetric protein localization
can be achieved by diffusion rather than requiring active transport.
4.2.3
Ensconsin is regulated by Par-1 phosphorylation
Ensconsin localization in the oocyte is controlled at multiple levels: First, mRNA is
88
DISCUSSION
localized to the anterior part of the oocyte, which is dependent on the 3’UTR. However,
this does not appear to be required for Ensconsin protein localization and function.
Second, Ensconsin is enriched at the oocyte cortex. Third, Ensconsin is kept away from
the posterior pole of the oocyte by Par-1-dependent phosphorylation. The axis
determinants, bicoid, gurken and oskar are transported by microtubule motor proteins
along with microtubule network in the oocyte. Par-1 regulates bicoid mRNA localization
by phosphorylating Exuperantia and also oskar mRNA localization. Furthermore, Par-1
also
regulates
microtubule-dependent
polarity through unknown MAP targets
(Doerflinger et al., 2003; Riechmann and Ephrussi, 2004; Riechmann et al., 2002;
Shulman et al., 2000). Here I have found that Ensconsin is a direct target of Par-1 and
affects oocyte polarity through Kinesin. The phosphorylation of Par-1 sites in Ensconsin
helps to control its proper localization. It is worth noting that mutations in the Par-1
phosphorylation sites do not severly affect the function of Ensconsin, as tub::ens mutant
can completely rescue ens∆C mutant phenotype. It could be that there are still some Par-1
phosphorylation sites in the Ens-mutant, a hypothesis which is supported by the in vitro
kinase assay. These sites may be enough to rescue some Ensconsin localization, or rescue
some its function. I also found that Ensconsin was delocalized from the apical membrane
of follicle cells and mislocalized to posterior pole of the oocyte in Par-1 mutant (Figure
29B, C, D). This suggests that Par-1 prevents Ensconsin localization to the posterior part
of the oocyte.
4.2.4
Par-1 regulates motor protein transport through MAPs
Cell polarity is dependent on microtubule polarity and motor protein function.
MARK/Par-1 regulates microtubule stabilization and polarity through phosphorylation of
MAPs, like Tau, MAP2 and MAP4. However, mutations in Drosophila Tau do not disrupt
oocyte polarity, implying that Tau may not be an essential target of Par-1 in the oocyte
(Doerflinger et al., 2003 ). This suggests that either there are other redundant or unknown
genes, like futsch (MAP1 homologue), or Par-1 can regulate cell polarity through
different mechanisms. I propose that Par-1 can control transportation of the plus-end
directed motor protein Kinesin through phosphorylation of Ensconsin.
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DISCUSSION
4.2.5
How does Ensconsin affect Khc-dependent transport?
How does Ensconsin affect Khc transport? Ensconsin does not appear to directly affect
the microtubule network. The microtubule binding assay shows that the N-terminal region
of Ensconsin can bind to microtubule, but the C-terminal region can not (Figure 20C, D).
Also tub-ens-X, which lacks the EHR2 conserved domain, can not rescue the ens∆C
mutant. This suggests that, although the EHR2 region doesn’t bind microtubules, is still
important for the Ensconsin function. Kinesin-1 transports vesicles or other cargo in a
strictly regulated manner. The C-terminal tail inhibits the motor domain until cargos binds,
causing a change in conformation allowing it to become active (Coy et al., 1999).
Kinesin-LacZ, which contains the motor domain but lacks C-terminal tail (Clark et al.,
1994), still localizes normally in the oocyte of ens∆C mutants, whereas full length Khc
protein does not(Figure 24B,25F). This suggests that the C-terminal domain of Khc may
be regulated by Ensconsin or that Ensconsin somehow regulates cargo loading onto Khc.
It is possible that the C-terminal domain of Ensconsin transiently or indirectly binds to
C-terminal domain of Khc and affects this autoinhibition or Khc binding of microtubules.
However, I have repeatedly searched for any such interaction between Khc and Ensconsin
and have been unable to find one. This does not, however, completely exclude on the
interaction between two.
Another possibility is that Ensconsin plays a role in removing obstacles or helping
Kinesin to bypass obstacles that can block its movement. Tau is involved in
neuron-degenerative disease in human. Aggreated hyperphosphorylated form of Tau
causes neurofibrillary tangles which are the symptom of Alzheimer’s disease and
Parkinson disease. Too much Tau proteins accumulate on microtubule surfaces and
interfere with the movement of motor proteins, like Kinesin, along microtubules,
therefore affecting cargo transport (Ackmann et al., 2000; Stamer et al., 2002; Vershinin
et al., 2007). The phosphorylation of Tau by Par-1/MARK causes Tau to be released from
microtubules (Mandelkow et al., 2004). This suggests that Par-1 could regulate Kinesin
movement via Tau. I have found that Ensconsin, a Par-1 target, also regulate Kinesin
movement. It is worth noting that Ensconsin and Tau display antagonistic function in Khc
transportation. Both Tau and Enconsin are expressed in the axon connecting the eye disc
and optic lobe in Drosophila (Figure 27B') (Heidary and Fortini, 2001). Preliminary data
90
DISCUSSION
shows that expressing TauGFP in ens∆N mutant can enhance the ens∆N phenotype (data not
shown). Overexpressing the wild type or disease mutation of htau causes the same
symptom of the human disorder in the Drosophila, such as Tau accumulation, progressive
neurondegenation, and early death (Ferber, 2001). If Ensconsin and Tau functions are
antagonistic, it may be worth to test if overpressing h-ensconsin or drosophila ensconsin
can suppress the htau overexpression phenotype in Drosophila. By using Drosophila, we
may find new ways to approach the neurodegerenating diseases caused by Tau.
Figure 32. The model for Ensconsin function.
Anterior is left, and posterior is right in the oocyte in the stage 9 or 10 egg chamber. Kinesin
carries cargo (blue) from anterior to the posterior pole of the oocyte along the microtubules
(yellow). After Kinesin reaches to the posterior of oocyte, it may become inactive and diffuse
or be transported by Dynein back to the anterior pole of the oocyte. Ensconsin (Green),
accumulated in the anterior part of oocyte, helps Kinesin loading/ processivity. Par-1 prevents
Ensconsin expression at the posterior pole of the oocyte to ensure the one direction Kinesin
tranportation.
Under normal condition, microtubule minus-ends are abundant in the anterior pole of the
oocyte and plus-ends are enriched in the posterior pole. Kinesin motor proteins transport
cargos from minus- to plus-ends (anterior to posterior) in the oocyte. When Kinesin
reaches to the posterior pole of the oocyte, Kinesin may become inactive. Then Kinesin
may diffuse or be transported by other motor proteins to the anterior pole of the oocyte to
continue its function (Duncan and Warrior, 2002). Ensconsin, a factor that has a positive
91
DISCUSSION
role on motor protein loading/or processivity, is accumulated in the anterior part of oocyte.
Mislocalization of Ensconsin in the posterior pole of the oocyte may stimulate Kinesin to
become active in the posterior pole of the oocyte. This may prevent the normal interaction
and recycling of Kinesin, leading to an accumulation of Kinesin at the posterior or
Kinesin-dependent transport may be stimulated on few microtubules which have their
plus end oriented away from the posterior pole of the oocyte. Kinesin-LacZ may still
localize normally in the ensconsin mutant due to its lack of cargo binding and regulatory
domains. A function of Par-1 may be to ensure that Ensconsin is only present at the
anterior, so allowing Kinesin to cycle normally or transport in one direction. The precise
role of Ensconsin in Kinesin regulation remains to be elucidated and may make a
promising avenue of investigation.
92
5 REFERENCES
REFERENCES
Ackmann, M., Wiech, H., and Mandelkow, E. (2000). Nonsaturable binding indicates
clustering of tau on the microtubule surface in a paired helical filament-like conformation.
J Biol Chem 275, 30335-30343.
Benton, R., Palacios, I. M., and St Johnston, D. (2002). Drosophila 14-3-3/PAR-5 is an
essential mediator of PAR-1 function in axis formation. Dev Cell 3, 659-671.
Benton, R., and St Johnston, D. (2003). Drosophila PAR-1 and 14-3-3 inhibit
Bazooka/PAR-3 to establish complementary cortical domains in polarized cells. Cell 115,
691-704.
Bergsten, S. E., and Gavis, E. R. (1999). Role for mRNA localization in translational
activation but not spatial restriction of nanos RNA. Development 126, 659-669.
Berleth, T., Burri, M., Thoma, G., Bopp, D., Richstein, S., Frigerio, G., Noll, M., and
Nusslein-Volhard, C. (1988). The role of localization of bicoid RNA in organizing the
anterior pattern of the Drosophila embryo. Embo J 7, 1749-1756.
Bi, G. Q., Morris, R. L., Liao, G., Alderton, J. M., Scholey, J. M., and Steinhardt, R. A.
(1997). Kinesin- and myosin-driven steps of vesicle recruitment for Ca2+-regulated
exocytosis. J Cell Biol 138, 999-1008.
Bolivar, J., Huynh, J. R., Lopez-Schier, H., Gonzalez, C., St Johnston, D., and
Gonzalez-Reyes, A. (2001). Centrosome migration into the Drosophila oocyte is
independent of BicD and egl, and of the organisation of the microtubule cytoskeleton.
Development 128, 1889-1897.
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.
Breitwieser, W., Markussen, F. H., Horstmann, H., and Ephrussi, A. (1996). Oskar protein
interaction with Vasa represents an essential step in polar granule assembly. Genes Dev
10, 2179-2188.
Brendza, R. P., Serbus, L. R., Duffy, J. B., and Saxton, W. M. (2000). A function for
kinesin I in the posterior transport of oskar mRNA and Staufen protein. Science 289,
2120-2122.
Bulinski, J. C., and Borisy, G. G. (1979). Self-assembly of microtubules in extracts of
cultured HeLa cells and the identification of HeLa microtubule-associated proteins. Proc
Natl Acad Sci U S A 76, 293-297.
Campos, A. R., Rosen, D. R., Robinow, S. N., and White, K. (1987). Molecular analysis
of the locus elav in Drosophila melanogaster: a gene whose embryonic expression is
neural specific. Embo J 6, 425-431.
Ceulemans, H., Vulsteke, V., De Maeyer, M., Tatchell, K., Stalmans, W., and Bollen, M.
(2002). Binding of the concave surface of the Sds22 superhelix to the alpha 4/alpha
5/alpha 6-triangle of protein phosphatase-1. J Biol Chem 277, 47331-47337.
Cha, B. J., Serbus, L. R., Koppetsch, B. S., and Theurkauf, W. E. (2002). Kinesin
I-dependent cortical exclusion restricts pole plasm to the oocyte posterior. Nat Cell Biol 4,
592-598.
Chekulaeva, M., Hentze, M. W., and Ephrussi, A. (2006). Bruno acts as a dual repressor
of oskar translation, promoting mRNA oligomerization and formation of silencing
particles. Cell 124, 521-533.
Chen, J., Kanai, Y., Cowan, N. J., and Hirokawa, N. (1992). Projection domains of MAP2
and tau determine spacings between microtubules in dendrites and axons. Nature 360,
674-677.
Chou, T. B., and Perrimon, N. (1996). The autosomal FLP-DFS technique for generating
germline mosaics in Drosophila melanogaster. Genetics 144, 1673-1679.
Clark, I., Giniger, E., Ruohola-Baker, H., Jan, L. Y., and Jan, Y. N. (1994). Transient
94
REFERENCES
posterior localization of a kinesin fusion protein reflects anteroposterior polarity of the
Drosophila oocyte. Curr Biol 4, 289-300.
Coutelis, J. B., and Ephrussi, A. (2007). Rab6 mediates membrane organization and
determinant localization during Drosophila oogenesis. Development 134, 1419-1430.
Cox, D. N., Lu, B., Sun, T. Q., Williams, L. T., and Jan, Y. N. (2001). Drosophila par-1 is
required for oocyte differentiation and microtubule organization. Curr Biol 11, 75-87.
Cox, R. T., and Spradling, A. C. (2003). A Balbiani body and the fusome mediate
mitochondrial inheritance during Drosophila oogenesis. Development 130, 1579-1590.
Coy, D. L., Hancock, W. O., Wagenbach, M., and Howard, J. (1999). Kinesin's tail
domain is an inhibitory regulator of the motor domain. Nat Cell Biol 1, 288-292.
Cyr, J. L., Pfister, K. K., Bloom, G. S., Slaughter, C. A., and Brady, S. T. (1991).
Molecular genetics of kinesin light chains: generation of isoforms by alternative splicing.
Proc Natl Acad Sci U S A 88, 10114-10118.
Dej, K. J., and Spradling, A. C. (1999). The endocycle controls nurse cell polytene
chromosome structure during Drosophila oogenesis. Development 126, 293-303.
DiAntonio, A., Burgess, R. W., Chin, A. C., Deitcher, D. L., Scheller, R. H., and Schwarz,
T. L. (1993). Identification and characterization of Drosophila genes for synaptic vesicle
proteins. J Neurosci 13, 4924-4935.
Doerflinger, H., Benton, R., Shulman, J. M., and St Johnston, D. (2003). The role of
PAR-1 in regulating the polarised microtubule cytoskeleton in the Drosophila follicular
epithelium. Development 130, 3965-3975.
Drewes, G., Ebneth, A., Preuss, U., Mandelkow, E. M., and Mandelkow, E. (1997).
MARK, a novel family of protein kinases that phosphorylate microtubule-associated
proteins and trigger microtubule disruption. Cell 89, 297-308.
Drewes, G., Trinczek, B., Illenberger, S., Biernat, J., Schmitt-Ulms, G., Meyer, H. E.,
Mandelkow, E. M., and Mandelkow, E. (1995). Microtubule-associated
protein/microtubule affinity-regulating kinase (p110mark). A novel protein kinase that
regulates tau-microtubule interactions and dynamic instability by phosphorylation at the
Alzheimer-specific site serine 262. J Biol Chem 270, 7679-7688.
Driever, W. (1993). Maternal control of anterior development in the Drosophila embryo
in the development of Drosophila melanogaster 1, 301-324.
Duchek, P., and Rorth, P. (2001). Guidance of cell migration by EGF receptor signaling
during Drosophila oogenesis. Science 291, 131-133.
Duchek, P., Somogyi, K., Jekely, G., Beccari, S., and Rorth, P. (2001). Guidance of cell
migration by the Drosophila PDGF/VEGF receptor. Cell 107, 17-26.
Duncan, J. E., and Warrior, R. (2002). The cytoplasmic dynein and kinesin motors have
interdependent roles in patterning the Drosophila oocyte. Curr Biol 12, 1982-1991.
Ephrussi, A., Dickinson, L. K., and Lehmann, R. (1991). Oskar organizes the germ plasm
and directs localization of the posterior determinant nanos. Cell 66, 37-50.
Fabre-Jonca, N., Allaman, J. M., Radlgruber, G., Meda, P., Kiss, J. Z., French, L. E., and
Masson, D. (1998). The distribution of murine 115-kDa epithelial microtubule-associated
protein (E-MAP-115) during embryogenesis and in adult organs suggests a role in
epithelial polarization and differentiation. Differentiation 63, 169-180.
Faire, K., Waterman-Storer, C. M., Gruber, D., Masson, D., Salmon, E. D., and Bulinski, J.
C. (1999). E-MAP-115 (ensconsin) associates dynamically with microtubules in vivo and
is not a physiological modulator of microtubule dynamics. J Cell Sci 112 ( Pt 23),
4243-4255.
Ferber, D. (2001). Neurodegenerative disease. Using the fruit fly to model tau
malfunction. Science 292, 1983-1984.
Ferrandon, D., Elphick, L., Nusslein-Volhard, C., and St Johnston, D. (1994). Staufen
95
REFERENCES
protein associates with the 3'UTR of bicoid mRNA to form particles that move in a
microtubule-dependent manner. Cell 79, 1221-1232.
Frydman, H. M., Li, J. M., Robson, D. N., and Wieschaus, E. (2006). Somatic stem cell
niche tropism in Wolbachia. Nature 441, 509-512.
Gauger, A. K., and Goldstein, L. S. (1993). The Drosophila kinesin light chain. Primary
structure and interaction with kinesin heavy chain. J Biol Chem 268, 13657-13666.
Gavis, E. R., and Lehmann, R. (1994). Translational regulation of nanos by RNA
localization. Nature 369, 315-318.
Golic, K. G. (1991). Site-specific recombination between homologous chromosomes in
Drosophila. Science 252, 958-961.
Gong, W. J., and Golic, K. G. (2004). Genomic deletions of the Drosophila melanogaster
Hsp70 genes. Genetics 168, 1467-1476.
Gonzalez-Reyes, A. (2003). Stem cells, niches and cadherins: a view from Drosophila. J
Cell Sci 116, 949-954.
Gonzalez-Reyes, A., Elliott, H., and St Johnston, D. (1995). Polarization of both major
body axes in Drosophila by gurken-torpedo signalling. Nature 375, 654-658.
Gonzalez-Reyes, A., and St Johnston, D. (1998). Patterning of the follicle cell epithelium
along the anterior-posterior axis during Drosophila oogenesis. Development 125,
2837-2846.
Grieder, N. C., de Cuevas, M., and Spradling, A. C. (2000). The fusome organizes the
microtubule network during oocyte differentiation in Drosophila. Development 127,
4253-4264.
Guo, S., and Kemphues, K. J. (1995). par-1, a gene required for establishing polarity in C.
elegans embryos, encodes a putative Ser/Thr kinase that is asymmetrically distributed.
Cell 81, 611-620.
Hachet, O., and Ephrussi, A. (2004). Splicing of oskar RNA in the nucleus is coupled to
its cytoplasmic localization. Nature 428, 959-963.
Hacker, U., Nystedt, S., Barmchi, M. P., Horn, C., and Wimmer, E. A. (2003).
piggyBac-based insertional mutagenesis in the presence of stably integrated P elements in
Drosophila. Proc Natl Acad Sci U S A 100, 7720-7725.
Hackney, D. D., Levitt, J. D., and Suhan, J. (1992). Kinesin undergoes a 9 S to 6 S
conformational transition. J Biol Chem 267, 8696-8701.
Harada, A., Oguchi, K., Okabe, S., Kuno, J., Terada, S., Ohshima, T., Sato-Yoshitake, R.,
Takei, Y., Noda, T., and Hirokawa, N. (1994). Altered microtubule organization in
small-calibre axons of mice lacking tau protein. Nature 369, 488-491.
Heidary, G., and Fortini, M. E. (2001). Identification and characterization of the
Drosophila tau homolog. Mech Dev 108, 171-178.
Hirokawa, N. (1996). Organelle transport along microtubules - the role of KIFs. Trends
Cell Biol 6, 135-141.
Horn, C., and Wimmer, E. A. (2000). A versatile vector set for animal transgenesis. Dev
Genes Evol 210, 630-637.
Huang, T. G., Suhan, J., and Hackney, D. D. (1994). Drosophila kinesin motor domain
extending to amino acid position 392 is dimeric when expressed in Escherichia coli. J
Biol Chem 269, 16502-16507.
Hurd, D. D., and Saxton, W. M. (1996). Kinesin mutations cause motor neuron disease
phenotypes by disrupting fast axonal transport in Drosophila. Genetics 144, 1075-1085.
Huynh, J. R., Munro, T. P., Smith-Litiere, K., Lepesant, J. A., and St Johnston, D. (2004).
The Drosophila hnRNPA/B homolog, Hrp48, is specifically required for a distinct step in
osk mRNA localization. Dev Cell 6, 625-635.
Huynh, J. R., Shulman, J. M., Benton, R., and St Johnston, D. (2001). PAR-1 is required
96
REFERENCES
for the maintenance of oocyte fate in Drosophila. Development 128, 1201-1209.
Illenberger, S., Drewes, G., Trinczek, B., Biernat, J., Meyer, H. E., Olmsted, J. B.,
Mandelkow, E. M., and Mandelkow, E. (1996). Phosphorylation of
microtubule-associated proteins MAP2 and MAP4 by the protein kinase p110mark.
Phosphorylation sites and regulation of microtubule dynamics. J Biol Chem 271,
10834-10843.
Jan, Y. N., and Jan, L. Y. (2001). Asymmetric cell division in the Drosophila nervous
system. Nat Rev Neurosci 2, 772-779.
Januschke, J., Gervais, L., Dass, S., Kaltschmidt, J. A., Lopez-Schier, H., St Johnston, D.,
Brand, A. H., Roth, S., and Guichet, A. (2002). Polar transport in the Drosophila oocyte
requires Dynein and Kinesin I cooperation. Curr Biol 12, 1971-1981.
Johnstone, O., and Lasko, P. (2001). Translational regulation and RNA localization in
Drosophila oocytes and embryos. Annu Rev Genet 35, 365-406.
Kankel, M. W., Duncan, D. M., and Duncan, I. (2004). A screen for genes that interact
with the Drosophila pair-rule segmentation gene fushi tarazu. Genetics 168, 161-180.
Karpova, N., Bobinnec, Y., Fouix, S., Huitorel, P., and Debec, A. (2006). Jupiter, a new
Drosophila protein associated with microtubules. Cell Motil Cytoskeleton 63, 301-312.
Kelley, R. L. (1993). Initial organization of the Drosophila dorsoventral axis depends on
an RNA-binding protein encoded by the squid gene. Genes Dev 7, 948-960.
Keyes, L. N., and Spradling, A. C. (1997). The Drosophila gene fs(2)cup interacts with
otu to define a cytoplasmic pathway required for the structure and function of germ-line
chromosomes. Development 124, 1419-1431.
Kim-Ha, J., Kerr, K., and Macdonald, P. M. (1995). Translational regulation of oskar
mRNA by bruno, an ovarian RNA-binding protein, is essential. Cell 81, 403-412.
Kim-Ha, J., Smith, J. L., and Macdonald, P. M. (1991). oskar mRNA is localized to the
posterior pole of the Drosophila oocyte. Cell 66, 23-35.
Kim-Ha, J., Webster, P. J., Smith, J. L., and Macdonald, P. M. (1993). Multiple RNA
regulatory elements mediate distinct steps in localization of oskar mRNA. Development
119, 169-178.
King, F. J., Szakmary, A., Cox, D. N., and Lin, H. (2001). Yb modulates the divisions of
both germline and somatic stem cells through piwi- and hh-mediated mechanisms in the
Drosophila ovary. Mol Cell 7, 497-508.
Koch, E. A., and Spitzer, R. H. (1983). Multiple effects of colchicine on oogenesis in
Drosophila: induced sterility and switch of potential oocyte to nurse-cell developmental
pathway. Cell Tissue Res 228, 21-32.
Komada, M., McLean, D. J., Griswold, M. D., Russell, L. D., and Soriano, P. (2000).
E-MAP-115, encoding a microtubule-associated protein, is a retinoic acid-inducible gene
required for spermatogenesis. Genes Dev 14, 1332-1342.
Kozielski, F., Sack, S., Marx, A., Thormahlen, M., Schonbrunn, E., Biou, V., Thompson,
A., Mandelkow, E. M., and Mandelkow, E. (1997). The crystal structure of dimeric
kinesin and implications for microtubule-dependent motility. Cell 91, 985-994.
Kull, F. J., Sablin, E. P., Lau, R., Fletterick, R. J., and Vale, R. D. (1996). Crystal structure
of the kinesin motor domain reveals a structural similarity to myosin. Nature 380,
550-555.
Lantz, V., Chang, J. S., Horabin, J. I., Bopp, D., and Schedl, P. (1994). The Drosophila
orb RNA-binding protein is required for the formation of the egg chamber and
establishment of polarity. Genes Dev 8, 598-613.
Liao, G. C., Rehm, E. J., and Rubin, G. M. (2000). Insertion site preferences of the P
transposable element in Drosophila melanogaster. Proc Natl Acad Sci U S A 97,
3347-3351.
97
REFERENCES
Lin, H., and Spradling, A. C. (1993). Germline stem cell division and egg chamber
development in transplanted Drosophila germaria. Dev Biol 159, 140-152.
Lin, H., Yue, L., and Spradling, A. C. (1994). The Drosophila fusome, a germline-specific
organelle, contains membrane skeletal proteins and functions in cyst formation.
Development 120, 947-956.
Liu, Z., Xie, T., and Steward, R. (1999). Lis1, the Drosophila homolog of a human
lissencephaly disease gene, is required for germline cell division and oocyte
differentiation. Development 126, 4477-4488.
Macdonald, P. M., Luk, S. K., and Kilpatrick, M. (1991). Protein encoded by the
exuperantia gene is concentrated at sites of bicoid mRNA accumulation in Drosophila
nurse cells but not in oocytes or embryos. Genes Dev 5, 2455-2466.
Macdonald, P. M., and Struhl, G. (1988). cis-acting sequences responsible for anterior
localization of bicoid mRNA in Drosophila embryos. Nature 336, 595-598.
Mandelkow, E., and Mandelkow, E. M. (1995). Microtubules and microtubule-associated
proteins. Curr Opin Cell Biol 7, 72-81.
Mandelkow, E. M., Thies, E., Trinczek, B., Biernat, J., and Mandelkow, E. (2004).
MARK/PAR1 kinase is a regulator of microtubule-dependent transport in axons. J Cell
Biol 167, 99-110.
Masson, D., and Kreis, T. E. (1993). Identification and molecular characterization of
E-MAP-115, a novel microtubule-associated protein predominantly expressed in
epithelial cells. J Cell Biol 123, 357-371.
Masson, D., and Kreis, T. E. (1995). Binding of E-MAP-115 to microtubules is regulated
by cell cycle-dependent phosphorylation. J Cell Biol 131, 1015-1024.
Mathieu, J., Sung, H. H., Pugieux, C., Soetaert, J., and Rorth, P. (2007). A Sensitized
PiggyBac Based Screen for Regulators of Border Cell Migration in Drosophila. Genetics.
McGrail, M., and Hays, T. S. (1997). The microtubule motor cytoplasmic dynein is
required for spindle orientation during germline cell divisions and oocyte differentiation
in Drosophila. Development 124, 2409-2419.
Meng, J., and Stephenson, E. C. (2002). Oocyte and embryonic cytoskeletal defects
caused by mutations in the Drosophila swallow gene. Dev Genes Evol 212, 239-247.
Micklem, D. R., Dasgupta, R., Elliott, H., Gergely, F., Davidson, C., Brand, A.,
Gonzalez-Reyes, A., and St Johnston, D. (1997). The mago nashi gene is required for the
polarisation of the oocyte and the formation of perpendicular axes in Drosophila. Curr
Biol 7, 468-478.
Neuman-Silberberg, F. S., and Schupbach, T. (1993). The Drosophila dorsoventral
patterning gene gurken produces a dorsally localized RNA and encodes a TGF alpha-like
protein. Cell 75, 165-174.
Nibu, Y., and Levine, M. S. (2001). CtBP-dependent activities of the short-range Giant
repressor in the Drosophila embryo. Proc Natl Acad Sci U S A 98, 6204-6208.
Nibu, Y., Zhang, H., and Levine, M. (1998). Interaction of short-range repressors with
Drosophila CtBP in the embryo. Science 280, 101-104.
Nilson, L. A., and Schupbach, T. (1999). EGF receptor signaling in Drosophila oogenesis.
Curr Top Dev Biol 44, 203-243.
Norvell, A., Kelley, R. L., Wehr, K., and Schupbach, T. (1999). Specific isoforms of squid,
a Drosophila hnRNP, perform distinct roles in Gurken localization during oogenesis.
Genes Dev 13, 864-876.
Nusslein-Volhard, C., Frohnhofer, H. G., and Lehmann, R. (1987). Determination of
anteroposterior polarity in Drosophila. Science 238, 1675-1681.
Nusslein-Volhard, C., and Roth, S. (1989). Axis determination in insect embryos. Ciba
Found Symp 144, 37-55; discussion 55-64, 92-38.
98
REFERENCES
Ochman, H., Gerber, A. S., and Hartl, D. L. (1988). Genetic applications of an inverse
polymerase chain reaction. Genetics 120, 621-623.
Palacios, I. M., Gatfield, D., St Johnston, D., and Izaurralde, E. (2004). An
eIF4AIII-containing complex required for mRNA localization and nonsense-mediated
mRNA decay. Nature 427, 753-757.
Pare, C., and Suter, B. (2000). Subcellular localization of Bic-D::GFP is linked to an
asymmetric oocyte nucleus. J Cell Sci 113 ( Pt 12), 2119-2127.
Pereira, A., Doshen, J., Tanaka, E., and Goldstein, L. S. (1992). Genetic analysis of a
Drosophila microtubule-associated protein. J Cell Biol 116, 377-383.
Peri, F., and Roth, S. (2000). Combined activities of Gurken and decapentaplegic specify
dorsal chorion structures of the Drosophila egg. Development 127, 841-850.
Perrimon, N., Engstrom, L., and Mahowald, A. P. (1989). Zygotic lethals with specific
maternal effect phenotypes in Drosophila melanogaster. I. Loci on the X chromosome.
Genetics 121, 333-352.
Perrimon, N., Lanjuin, A., Arnold, C., and Noll, E. (1996). Zygotic lethal mutations with
maternal effect phenotypes in Drosophila melanogaster. II. Loci on the second and third
chromosomes identified by P-element-induced mutations. Genetics 144, 1681-1692.
Pokrywka, N. J., and Stephenson, E. C. (1995). Microtubules are a general component of
mRNA localization systems in Drosophila oocytes. Dev Biol 167, 363-370.
Riechmann, V., and Ephrussi, A. (2001). Axis formation during Drosophila oogenesis.
Curr Opin Genet Dev 11, 374-383.
Riechmann, V., and Ephrussi, A. (2004). Par-1 regulates bicoid mRNA localisation by
phosphorylating Exuperantia. Development 131, 5897-5907.
Riechmann, V., Gutierrez, G. J., Filardo, P., Nebreda, A. R., and Ephrussi, A. (2002). Par-1
regulates stability of the posterior determinant Oskar by phosphorylation. Nat Cell Biol 4,
337-342.
Rittenhouse, K. R., and Berg, C. A. (1995). Mutations in the Drosophila gene bullwinkle
cause the formation of abnormal eggshell structures and bicaudal embryos. Development
121, 3023-3033.
Roth, S., Neuman-Silberberg, F. S., Barcelo, G., and Schupbach, T. (1995). cornichon and
the EGF receptor signaling process are necessary for both anterior-posterior and
dorsal-ventral pattern formation in Drosophila. Cell 81, 967-978.
Sahut-Barnola, I., Godt, D., Laski, F. A., and Couderc, J. L. (1995). Drosophila ovary
morphogenesis: analysis of terminal filament formation and identification of a gene
required for this process. Dev Biol 170, 127-135.
Saxton, W. M., Hicks, J., Goldstein, L. S., and Raff, E. C. (1991). Kinesin heavy chain is
essential for viability and neuromuscular functions in Drosophila, but mutants show no
defects in mitosis. Cell 64, 1093-1102.
Schier, A. F., and Gehring, W. J. (1992). Direct homeodomain-DNA interaction in the
autoregulation of the fushi tarazu gene. Nature 356, 804-807.
Schnorrer, F., Bohmann, K., and Nusslein-Volhard, C. (2000). The molecular motor
dynein is involved in targeting swallow and bicoid RNA to the anterior pole of
Drosophila oocytes. Nat Cell Biol 2, 185-190.
Schnorrer, F., Luschnig, S., Koch, I., and Nusslein-Volhard, C. (2002).
Gamma-tubulin37C and gamma-tubulin ring complex protein 75 are essential for bicoid
RNA localization during drosophila oogenesis. Dev Cell 3, 685-696.
Schupbach, T., and Wieschaus, E. (1986). Germline autonomy of maternal-effect
mutations altering the embryonic body pattern of Drosophila. Dev Biol 113, 443-448.
Serbus, L. R., Cha, B. J., Theurkauf, W. E., and Saxton, W. M. (2005). Dynein and the
actin cytoskeleton control kinesin-driven cytoplasmic streaming in Drosophila oocytes.
99
REFERENCES
Development 132, 3743-3752.
Shah, K., Liu, Y., Deirmengian, C., and Shokat, K. M. (1997). Engineering unnatural
nucleotide specificity for Rous sarcoma virus tyrosine kinase to uniquely label its direct
substrates. Proc Natl Acad Sci U S A 94, 3565-3570.
Shulman, J. M., Benton, R., and St Johnston, D. (2000). The Drosophila homolog of C.
elegans PAR-1 organizes the oocyte cytoskeleton and directs oskar mRNA localization to
the posterior pole. Cell 101, 377-388.
Siegrist, S. E., and Doe, C. Q. (2005). Microtubule-induced Pins/Galphai cortical polarity
in Drosophila neuroblasts. Cell 123, 1323-1335.
Skoufias, D. A., Cole, D. G., Wedaman, K. P., and Scholey, J. M. (1994). The
carboxyl-terminal domain of kinesin heavy chain is important for membrane binding. J
Biol Chem 269, 1477-1485.
Spradling, A. (1993). Developmental genetics of oogenesis. in the development of
Drosophila melanogaster, 1-70.
Spradling, A., Drummond-Barbosa, D., and Kai, T. (2001). Stem cells find their niche.
Nature 414, 98-104.
St Johnston, D. (2005). Moving messages: the intracellular localization of mRNAs. Nat
Rev Mol Cell Biol 6, 363-375.
St Johnston, D., Beuchle, D., and Nusslein-Volhard, C. (1991). Staufen, a gene required to
localize maternal RNAs in the Drosophila egg. Cell 66, 51-63.
St Johnston, D., Driever, W., Berleth, T., Richstein, S., and Nusslein-Volhard, C. (1989).
Multiple steps in the localization of bicoid RNA to the anterior pole of the Drosophila
oocyte. Development 107 Suppl, 13-19.
Stamer, K., Vogel, R., Thies, E., Mandelkow, E., and Mandelkow, E. M. (2002). Tau
blocks traffic of organelles, neurofilaments, and APP vesicles in neurons and enhances
oxidative stress. J Cell Biol 156, 1051-1063.
Strunk, B., Struffi, P., Wright, K., Pabst, B., Thomas, J., Qin, L., and Arnosti, D. N. (2001).
Role of CtBP in transcriptional repression by the Drosophila giant protein. Dev Biol 239,
229-240.
Suter, B., Romberg, L. M., and Steward, R. (1989). Bicaudal-D, a Drosophila gene
involved in developmental asymmetry: localized transcript accumulation in ovaries and
sequence similarity to myosin heavy chain tail domains. Genes Dev 3, 1957-1968.
Suzuki, A., and Ohno, S. (2006). The PAR-aPKC system: lessons in polarity. J Cell Sci
119, 979-987.
Tassan, J. P., and Le Goff, X. (2004). An overview of the KIN1/PAR-1/MARK kinase
family. Biol Cell 96, 193-199.
Theurkauf, W. E. (1994). Premature microtubule-dependent cytoplasmic streaming in
cappuccino and spire mutant oocytes. Science 265, 2093-2096.
Theurkauf, W. E., Alberts, B. M., Jan, Y. N., and Jongens, T. A. (1993). A central role for
microtubules in the differentiation of Drosophila oocytes. Development 118, 1169-1180.
Theurkauf, W. E., Smiley, S., Wong, M. L., and Alberts, B. M. (1992). Reorganization of
the cytoskeleton during Drosophila oogenesis: implications for axis specification and
intercellular transport. Development 115, 923-936.
Thibault, S. T., Singer, M. A., Miyazaki, W. Y., Milash, B., Dompe, N. A., Singh, C. M.,
Buchholz, R., Demsky, M., Fawcett, R., Francis-Lang, H. L., et al. (2004). A
complementary transposon tool kit for Drosophila melanogaster using P and piggyBac.
Nat Genet 36, 283-287.
Thompson, B. J., Mathieu, J., Sung, H. H., Loeser, E., Rorth, P., and Cohen, S. M. (2005).
Tumor suppressor properties of the ESCRT-II complex component Vps25 in Drosophila.
Dev Cell 9, 711-720.
100
REFERENCES
Tomancak, P., Piano, F., Riechmann, V., Gunsalus, K. C., Kemphues, K. J., and Ephrussi,
A. (2000). A Drosophila melanogaster homologue of Caenorhabditis elegans par-1 acts at
an early step in embryonic-axis formation. Nat Cell Biol 2, 458-460.
Vaccari, T., and Ephrussi, A. (2002). The fusome and microtubules enrich Par-1 in the
oocyte, where it effects polarization in conjunction with Par-3, BicD, Egl, and dynein.
Curr Biol 12, 1524-1528.
Vallee, R. B., and Bloom, G. S. (1991). Mechanisms of fast and slow axonal transport.
Annu Rev Neurosci 14, 59-92.
van Eeden, F. J., Palacios, I. M., Petronczki, M., Weston, M. J., and St Johnston, D.
(2001). Barentsz is essential for the posterior localization of oskar mRNA and colocalizes
with it to the posterior pole. J Cell Biol 154, 511-523.
Vereshchagina, N., Bennett, D., Szoor, B., Kirchner, J., Gross, S., Vissi, E., White-Cooper,
H., and Alphey, L. (2004). The essential role of PP1beta in Drosophila is to regulate
nonmuscle myosin. Mol Biol Cell 15, 4395-4405.
Verhey, K. J., Lizotte, D. L., Abramson, T., Barenboim, L., Schnapp, B. J., and Rapoport,
T. A. (1998). Light chain-dependent regulation of Kinesin's interaction with microtubules.
J Cell Biol 143, 1053-1066.
Vershinin, M., Carter, B. C., Razafsky, D. S., King, S. J., and Gross, S. P. (2007).
Multiple-motor based transport and its regulation by Tau. Proc Natl Acad Sci U S A 104,
87-92.
Wang, S., and Hazelrigg, T. (1994). Implications for bcd mRNA localization from spatial
distribution of exu protein in Drosophila oogenesis. Nature 369, 400-403.
Wang, X. M., Peloquin, J. G., Zhai, Y., Bulinski, J. C., and Borisy, G. G. (1996). Removal
of MAP4 from microtubules in vivo produces no observable phenotype at the cellular
level. J Cell Biol 132, 345-357.
Weatherbee, J. A., Luftig, R. B., and Weihing, R. R. (1980). Purification and
reconstitution of HeLa cell microtubules. Biochemistry 19, 4116-4123.
Wharton, R. P., and Struhl, G. (1989). Structure of the Drosophila BicaudalD protein and
its role in localizing the the posterior determinant nanos. Cell 59, 881-892.
Wilhelm, J. E., Hilton, M., Amos, Q., and Henzel, W. J. (2003). Cup is an eIF4E binding
protein required for both the translational repression of oskar and the recruitment of
Barentsz. J Cell Biol 163, 1197-1204.
Xu, T., and Rubin, G. M. (1993). Analysis of genetic mosaics in developing and adult
Drosophila tissues. Development 117, 1223-1237.
Yang, J. T., Laymon, R. A., and Goldstein, L. S. (1989). A three-domain structure of
kinesin heavy chain revealed by DNA sequence and microtubule binding analyses. Cell
56, 879-889.
Yang, J. T., Saxton, W. M., Stewart, R. J., Raff, E. C., and Goldstein, L. S. (1990).
Evidence that the head of kinesin is sufficient for force generation and motility in vitro.
Science 249, 42-47.
Yano, T., Lopez de Quinto, S., Matsui, Y., Shevchenko, A., Shevchenko, A., and Ephrussi,
A. (2004). Hrp48, a Drosophila hnRNPA/B homolog, binds and regulates translation of
oskar mRNA. Dev Cell 6, 637-648.
Zhang, Q., Yoshimatsu, Y., Hildebrand, J., Frisch, S. M., and Goodman, R. H. (2003).
Homeodomain interacting protein kinase 2 promotes apoptosis by downregulating the
transcriptional corepressor CtBP. Cell 115, 177-186.
101
6 APPENDIX
APPENDIX
6.1
Screen result
Table 2. Summary of PiggyBac insertions
FRT
Putatively
chromosome
Molecular nature Phenotype
affected genes
arm
Group 1. No maternal defects
4278
82
rough eye
3297
80
2344
42
CG12765
312
40
rab6
wing can not expand
wing bigger and leg shorter
and bigger
bristle miss
squid
more dorsal appendages
F-box protein
Group 3. Abnormal oogenesis
55
1621
82
squid
more dorsal appendages
2178
82
squid
more dorsal appendages
3508
82
squid
more dorsal appendages
4180
82
squid
more dorsal appendages
5133
82
squid
more dorsal appendages
mod
441
Few
later
stage
egg
chambers.
cuticle defects in adult,
Oskar protein mis-localization
(<10%)
some nurse cell membranes
gone,
dumpless egg
some nurse cell membrane
gone,
10%
border
cell
migration defects
some nurse cell membranes
gone
321
40
lgl
553
42
CG12340
701
82
CG5169
1173
82
sds22
2317
40
pka
2766
80
CG11811
308
40
cropped
transcription factor collapsed eggs
2791
40
cropped
2877
40
cropped
3295
40
cropped
3594
42
3753
40
fray
4044
82
CG31232
4256
40
CG10664
transcription factor collapsed eggs
Expression of Oskar protein is
transcription factor
weaker.
Oskar protein mislocalization,
transcription factor
10% border cells split
Oocytes are smaller,
Oskar protein mislocalization
some nurse cell membranes
are undetectable
Cyclin-like domain all germ cells degenerated
cytochrome c
Border cells migration defects
in somatic clone.
oxidase
Kinase
ppp 1, regulatory
subunit 7
Guanylate kinase
103
APPENDIX
Continued Table 2
FRT
Putatively
chromosome
affected genes
arm
cornichon
4329
82
Molecular nature Phenotype
no dorsal appendage
4371
80
4497
80
4670
82
4741
82
4882
80
CG2034
4949
42
pipsqueak
4744
40
cup
BTB/POZ,
Oskar unanchoring defect
Helix-turn-helix,
Protein kinase-like
Border cell migration defects.
4975
40
cup
Border cell migration defects
5016
40
cup
5493
82
fer1hch
Bab2
Transcription factor
All germ cells die
Border cell migration defects
iron binding protein
in somatic clone
Group 4. Posterior group
83
82
122
40
456
posterior phenotype in embryo
vasa
posterior phenotype in embryo
hrp48
posterior phenotype in embryo
680
40
vasa
posterior phenotype in embryo
1094
40
vasa
posterior phenotype in embryo
3269
82
posterior phenotype in embryo
3747
40
posterior phenotype in embryo
3848
82
posterior phenotype in embryo
4053
40
aubergine
posterior phenotype in embryo
4131
posterior phenotype in embryo
4825
82
posterior phenotype in embryo
5144
82
posterior phenotype in embryo
5175
80
posterior phenotype in embryo
5257
40
posterior phenotype in embryo
Group 5. Segmentation defects
284
80
denticle belts fusion
772
80
CG6854
1210
82
E2F
1280
80
denticle belts fusion
1338
82
denticle belts fusion
2265
82
bess motif
denticle belts fusion
open or denticle belts fusion
denticle belts fusion
2670
80
CG7339
2691
80
CG17090
RNA polymerase
Rpb7
Kinase(hipk2)
denticle belts fusion or missed
104
APPENDIX
Continued Table 2.
FRT
Putatively
chromosome
affected genes
arm
fray
2700
82
Molecular nature Phenotype
denticle belts fusion
2860
82
r1
2970
80
CG7177
Kinase (wnk)
3218
82
CG31048
GTPase
3286
82
4350
82
CG33105
emp24/gp25L/p24, longer denticle and
denticle belts missed
GOLD
4369
82
4399
82
4542
80
4592
82
4824
82
4889
82
CG7023
5059
80
eIF4E
911
40
kismet
Ubiquitin hydrolase open Embryo
pair-rule
like
phenotype
denticle belts fusion
5201
40
kismet
denticle belts fusion
28
40
CG6746
173
82
2600
80
abl
3292
40
dorsal
dorsalized embryo
3311
42
3900
80
tsp66E
embryo defects
4017
82
4170
80
CG14998
4211
80
talin
4496
82
CG31048
4589
80
4735
40
apc or CG31048
sbf
denticle belts fusion
posterior like or denticle belts
fusion
naked embryo or early embryo
defects
some
denticle belts missing
some denticle belts missing
mutant
Group 6. Others
Protein tyrosine
phosphatase-like
protein
smaller embryo
germband retraction defects
Ensconsin
early embryo defects
dorsal open phenotype
Sponge
early embryo defects
germband retraction defects
105
APPENDIX
6.2
Publications
A Sensitized PiggyBac Based Screen for Regulators of Border Cell Migration in
Drosophila. Mathieu, J, Sung, H.H., Pugieux, C, Soetaert, J, Rørth, P. Genetics. 2007
May 4
Tumor suppressor properties of the ESCRT-II complex component Vps25 in
Drosophila. Thompson, B.J., Mathieu, J., Sung, H.H., Loeser, E., Rørth, P. & Cohen, S.M.
Dev Cell 2005 Nov;9(5):711-20.
Regulators of endocytosis maintain localized receptor tyrosine kinase signaling in
guided migration. Jekely, G., Sung, H.H., Luque, C.M. & Rørth, P.
Dev Cell 2005 Aug;9(2):197-207.
106
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