doctoral dissertation ying deng department of cell ... · department of cell physiology ... dengue...

INTERNATIONAL GRADUATE SCHOOL OF BIOSCIENCES

RUHR-UNIVERSITY BOCHUM

MOLECULAR MECHANISM OF OLFACTORY SIGNAL

TRANSDUCTION IN DROSOPHILA MELANOGASTER

Doctoral Dissertation

Ying Deng

Department of Cell Physiology

First thesis advisor: Prof. Dr. Dr. Dr. Hanns Hatt

Second thesis advisor: Prof. Dr. Wolfgang Kirchner

Bochum, October. 2009

INTERNATIONALEN GRADUIERTENSCHULE BIOWISSENSCHAFTEN

RUHR-UNIVERSITÄT BOCHUM

DER MOLEKULAR MECHANISMUS DER OLFAKTORISCHEN

SIGNALTRANSDUKTION IN DROSOPHILA MELANOGASTER

Dissertation

Ying Deng

Lehrstuhl für Zellphysiologie

Referent: Prof. Dr. Dr. Dr. Hanns Hatt

Korreferent: Prof. Dr. Wolfgang Kirchner

Bochum, October. 2009

i

Declaration

I certify herewith that the dissertation at hand was completed and written independently

and without outside assistance. The “Guidelines for Good Scientific Practice” (Leitlinien

guter wissenschaftlicher Praxis und Grundsätze für das Verfahren bei vermutetem

wissenschaftlichen Fehlverhaltens) according to §9, Sec. 3 were adhered to. This work

has never been submitted in this or similar form at this or any other domestic or foreign

institution of higher learning as a dissertation.

Hiermit erkläre ich, dass ich die Arbeit selbständig verfasst und bei keiner anderen

Fakultät eingereicht und dass ich keine anderen als die angegebenen Hilfsmittel

verwandet habe. Es handelt sich bei der heute von mir eingereichten Dissertation um fünf

in Wort und Bild völlig übereinstimmende Exemplare.

Weiterhin erkläre ich, dass digitale Abbildungen nur die originalen Daten enthalten und

in keinen Fall inhaltsverändernde Bildbearbeitung vorgenommem wurde.

Ying Deng

Bochum, 15th Oct. 2009

Error! Reference source not found.

ii

Table of contents 1 Introduction................................................................................................................. 1

1.1 The Drosophila olfactory system........................................................................ 1

1.1.1 Anatomy of the Drosophila olfactory system............................................. 1

1.1.2 The odor coding system in Drosophila....................................................... 7

1.1.3 New insights of Drosophila odorant receptor........................................... 14

1.2 Heterotrimeric G-proteins................................................................................. 16

1.3 Signal transduction pathway in odorant receptor neurons................................ 19

2 Objective ................................................................................................................... 23

3 Materials and Methods.............................................................................................. 24

3.1 Materials ........................................................................................................... 24

3.1.1 Fly food..................................................................................................... 24

3.1.2 Enzymes and Kits and molecularbiological supplies................................ 24

3.1.3 Immunoblotting and Immunohistochemistry............................................ 24

3.1.4 Cell culture................................................................................................ 26

3.1.5 Mammalian cell lines................................................................................ 26

3.1.6 Buffers and Solutions................................................................................ 26

3.1.7 Odorants.................................................................................................... 27

3.1.8 Miscellaneous ........................................................................................... 28

3.2 Methods............................................................................................................. 28

3.2.1 Fly rearing and fly stocks.......................................................................... 28

3.2.2 Molecular cloning and plasmid construction............................................ 29

3.2.3 Cell culture and transfection ..................................................................... 30

3.2.4 Ratiometric calcium imaging in HEK293 cells ........................................ 30

3.2.5 RNA isolation, reverse transcription, RT-PCR and Real-time quantitative

PCR ................................................................................................................... 31

3.2.6 [35S] GTPγS binding assay ...................................................................... 32

3.2.7 Larval chemotaxis assay ........................................................................... 33

3.2.8 Electroantennogram (EAG) ...................................................................... 34


3.2.9 Antennal single sensillum recording......................................................... 35

3.2.10 Labellar single sensillum tip recording..................................................... 36

3.2.11 Odorant application system in EAG and single sensillum recording ....... 37

3.2.12 Immunohistochemistry ............................................................................. 37

3.2.13 Whole mount fluorescent antibody staining of Drosophila larvae........... 38

3.2.14 SDS gel electrophoresis and Western-blotting ......................................... 38

3.2.15 cAMP assay .............................................................................................. 38

4 Results....................................................................................................................... 39

4.1 Different Gα subunits are expressed in Drosophila antenna. ........................... 39

4.1.1 RT-PCR analysis of Gα subunits in Drosophila antenna ......................... 39

4.1.2 Western-blot analysis showed that different classes of Gα are expressed in

the antenna ................................................................................................................ 40

4.1.3 Immunohistochemical analysis of the expression patterns of different Gα

proteins in Drosophila antenna................................................................................. 41

4.1.4 Real-time quantitative PCR analysis of Gαs and Gαq expression in

Drosophila antenna and head.................................................................................... 43

4.2 Functional study of Gα subunit in Drosophila olfactory signal transduction

pathway......................................................................................................................... 44

4.2.1 Overexpression study of different Gα subunits and G-protein specific

protein toxins in Drosophila ORNs .......................................................................... 44

4.2.2 Dose-response curves of EAG responses from CTX and control flies..... 45

4.2.3 CTX blocks the olfactory response of ORNs in Drosophila .................... 47

4.2.4 CTX does not act as a non-specific toxin on cells .................................... 49

4.2.5 Functional study of OR and Gα protein interactivity in the recombinant

HEK293 expression system ...................................................................................... 49

4.2.6 Overexpression of a GTPase deficient Gαs mutant in Drosophila antenna

leads to a change in response dynamics.................................................................... 52

4.2.7 Odorant exposure caused Gαs redistribution in antenna ........................... 53

4.2.8 Generation of a Gαs knockout or knockdown fly ..................................... 54

4.3 Secondary messengers and other components in the ORNs signal transduction

cascade .......................................................................................................................... 56


4.3.1 The cAMP pathway is involved in odorant signaling in fly olfactory

neurons ................................................................................................................... 56

4.3.2 Expression of activated Gαq in ORNs leads to resensitization deficiency. ..

................................................................................................................... 59

4.3.3 Effect of MIA on Drosophila odorant receptors....................................... 61

4.3.4 Functional study of Ih channels in Drosophila odorant receptor neurons 63

4.4 Larva study........................................................................................................ 65

4.4.1 Gαs and Gαq are expressed in larva olfactory receptor neurons ............... 65

4.4.2 CTX larvae have normal chemotaxis behavior......................................... 66

4.4.3 Expression of AcGq3 in larval olfactory receptor neurons causes olfactory

aberration .................................................................................................................. 66

5 Discussion................................................................................................................. 67

5.1 The role of Gαs in the olfactory signal transduction pathway .......................... 67

5.2 Larval olfactory system..................................................................................... 73

5.3 Other signaling components in the olfactory signal transduction pathway ...... 74

6 Conclusion ................................................................................................................ 78

6.1 Summary ........................................................................................................... 78

6.2 Zusammenfassung............................................................................................. 80

References......................................................................................................................... 82

Abbreviations.................................................................................................................. 100

List of Figures and Tables............................................................................................... 102

Curriculum Vitae ............................................................................................................ 104

Acknowledgements......................................................................................................... 107

Introduction

1

1 Introduction

Insects, as the most diverse group of animals, represent more than half of all known

living organisms and potentially represent over 90% of the differing life forms on the

planet. They have an enormous impact on global public health as disease vectors, which

carry pathogens that cause human diseases such as malaria, dengue and yellow fever, and

as agricultural enablers as well as pests (Beaty and Marquardt, 1996). Olfaction acts as an

important sensory input to their behavior, such as processing chemical signals from the

environment, leading to the detection of food, reproductive partners, oviposition sites,

hosts, prey or predators (Rutzler and Zwiebel, 2005). Therefore, it is of great value to

understand the signal transduction pathway in insect olfactory system, in addition to

nurturing a better understanding of insect neurobiology, may ultimately help in devising

novel intervention strategies to reduce crop damage and disease transmission.

The fruit fly, Drosophila melanogaster, has been used as a model orgainism for over 100

years. Today, thousands of scientists use Drosophila to study almost every aspect of

eukaryotic biology from gene organization to developmental biology to behavior and

everything in between. Advantages of choosing Drosophila for these studies include the

completely sequenced genome (Adams et al., 2000), a short live cycle and all kinds of

well-established genetic tools.

1.1 The Drosophila olfactory system

1.1.1 Anatomy of the Drosophila olfactory system

Drosophila has two pairs of olfactory organs, the antennae and the maxillary palps

(Figure 1.1.1 A). They are cuticle-covered appendages, which contain approximately

1200 and 120 olfactory receptor neurons (ORNs) each (Hildebrand and Shepherd,

1997;Riesgo-Escovar et al., 1997;Shanbhag et al., 1999;Shanbhag et al., 2000). ORNs are

compartmentalized into sensory hairs called sensilla, which can be subdivided into three

morphological types: basiconic sensilla (BS) resemble a club, trichoids have sharp tips,

and coeloconics are dome-shaped and have a grooved surface (Figure 1.1.1 B, C, D, E

and F). Each sensillum contains the dendrites of one to four ORNs (Shanbhag et al.,

1999). At the base of sensillum, specialized auxiliary cells surround the cell bodies of the

Introduction

2

ORNs. (Figure 1.1.1 G, H). The sensory dendrites of the ORNs are bathed in the

sensillum lymph that secreted from the auxiliary cells (Shanbhag et al., 2000). Odorant-

binding proteins are also secreted by the auxiliary cells and present in this extracellular

sensillum lymph (Kaissling, 1996;Park et al., 2000).

The antennae contain all three types of olfactory sensilla, whereas the maxillary palps

contain only basiconic sensilla. The respective contributions of the antenna and maxillary

palps to chemosensory-mediated behavior are not yet clear (Shanbhag et al., 1999).

Sensilla of different morphological types respond to different types of chemical stimuli.

Basiconic sensilla are sensitive to many food odorants with a variety of chemical groups

(Park et al., 2002). Trichoid sensilla do not have a strong response to food odors but to fly

odors (van der Goes van Naters and Carlson, 2007). It is also known that the volatile fly

pheromone 11-cis-vaccenyl acetate (cVA) can be detected by the trichoid sensilla (Jin et

al., 2008). One type of coeloconic sensilla (ac3) respond to a remarkably high fraction of

the tested food odors, the rest of the coeloconic sensilla are sensitive to humidity or

ammonia (Benton et al., 2009;Spletter and Luo, 2009;Yao et al., 2005).

Introduction

3

Figure 1.1.1 The Drosophila olfactory organ

(A) Drosophila melanogaster head with the olfactory appendages, the antenna and the maxillary palps. S: Scapus, P: Pedicellus, F: Funiculus. Figure A was adapted from Jürgen Berger, Max Planck Institute for developmental Biology, Tübingen, Germany. (B, C) Low magnification scanning electron micrographs of antenna surface. C: coeloconic sensillum; I: intermediate sensillum; LB, SB, TB large, small, thin basiconic sensillum, respectively; Sp spinule; T s. trichoid sensillum. Scale bar in B and C = 10 μm. (D) High resolution scanning electron micrographs of thin basiconic sensillum (TB), large basiconic sensillum (LB), coeloconic sensillum (C) and trichoid sensillum (T). Different types of sensilly have different surface structure and arrangement of wall pores. Scale bars in TB and LB = 1 μm, Scale bars in T and C = 0.5 μm.(E) Structure of basiconic sensillum, trichoid sensillum and coeloconic sensillum (Stocker, 1994). (F) Cross-sections through various types of sensillum. Scale bars = 0.5 μm. (G, H) Longitudinal section (G) and schematic drawing (H) of a small basiconic sensillum with two ORNs. D1 and D2 are the dendrites of two odorant receptor neurons (R). The three types of auxiliary cells are thecogen Th, trichogen Tr, and tormogen cells To. Epidermal cell E. The large sensillum-lymph cavity (S1) is bordered by the microvilli and microlamellae of trichogen and tormogen cells. Scale bars = 1 μm. B, C, D, F, G were adapted from Shanbhag, S.R. et al. 1999. H was adapted from Park, S.K. et al 2002 (Park et al., 2002).

ORNs send axons to the antennal lobe (AL), which has a functional organization

remarkably similar to that of the olfactory bulb in vertebrates (Hildebrand and Shepherd,

1997). In the AL, ORNs form synapses onto second order neurons called projection

Introduction

4

neurons (PNs) (Stocker, 1994) (Figure 1.1.2 A). The AL can be subdivided into about 43

spherical units called glomeruli (Laissue et al., 1999). Each ORNs send axons to only one

or a few glomeruli (Stocker, 1994), and individual PNs typically innervate only a single

glomerulus (Jefferis et al., 2001;Marin et al., 2002;Wong et al., 2002). The glomeruli also

contain the processes of local interneurons that branch in multiple glomeruli (Stocker et

al., 1990;Stocker, 1994), providing a means for information transfer between glomeruli.

The axons of PNs project to the mushroom body (MB) and lateral horn of the brain

(Figure 1.1.2 A).

Larvae of Drosophila also exhibit a dorsal organ mediated olfactory response (Ayyub et

al., 1990;Cobb et al., 1992;Heimbeck et al., 1999;Monte et al., 1989;Oppliger et al.,

2000). The larva cephalic chemosensory apparatus consists of 3 external sensory organs,

dorsal organ (DO), terminal organ (TO), and ventral organ (VO), plus 3 pharyngeal

organ, as the dorsal, ventral, and posterior pharyngeal sense organs (DPS, VPS, PPS)

(Figure 1.1.2 B) (Gerber and Stocker, 2007;Python and Stocker, 2002). Each of them

includes several sensilla, a sensillum comprising one to several sensory neurons and 3

accessory cells, all housed below a common cuticular structure or terminal pore.

The dorsal organ houses the dendrites of 21 olfactory receptor neurons in the central

dome and six peripheral sensilla. An olfactory function of the dorsal organ is proved by

electrophysiological recordings (Kreher et al., 2005;Oppliger et al., 2000) and by

combined toxin expression and behavioral studies (Fishilevich et al., 2005;Heimbeck et

al., 1999;Larsson et al., 2004). The experiment of genetically ablation the OR83b

receptor and expressing diphtheria toxin in OR83b neurons demonstrate that these 21

sensory neurons, which express OR83b, are the sole larval ORNs.

The DO, TO, and VO all correspond with their internal ganglion. The ganglion of the DO

(DOG) contains 36 to 37 sensory neurons (Python and Stocker, 2002). The 21 ORNs

among them extend their dendrites as seven triplets into the dome. The dendrites of three

of the rest neurons project towards the dorsolateral sensilla of the TO (Heimbeck et al.,

1999;Python and Stocker, 2002), whereas the remaining neruons innervate the 6

peripheral sensilla of the DO. The TO and VO ganglia include 32 and 7 sensory neurons,

respectively (Python and Stocker, 2002). Each of the three pharyngeal sense organs

consists of several sensilla, comprising one to nine sensory neurons. The presence of

Introduction

5

pores and bristles suggest the gustatory and mechanosensory function of these pharyngeal

sense organs. The dorsal and ventral pharyngeal sense organs, both of these sensory

organs are located behind the mouth hooks, contain 17 and 16 neurons, respectively;

while the posterior pharyngeal sense organ contains two sensilla with three neurons each

(Figure 1.1.2 B).

In contrast to adults, all olfactory projections remain ipsilateral in larva. Both olfactory

and gustatory neurons from the DO ganglion connect to the brain by the antennal nerve

(Python and Stocker, 2002;Tissot et al., 1997). The supraesophageal labral nerve receives

information from the dorsal pharyngeal organ and from the posterior pharyngeal organ,

whereas the subesophageal maxillary and labial nerves comprise the afferents from the

TO and VO ganglia and from the ventral pharyngeal organ, respectively (Gendre et al.,

2004;Python and Stocker, 2002) (Figure 1.1.2 B).

Introduction

6

Figure 1.1.2 Organization of the Drosophila olfactory system.

(A) Overview of the Drosophila adult olfactory system organization, with mammalian counterparts in parentheses (Komiyama et al., 2003). Axons of ORNs expressing the same receptor (represented by the same color) project to the same glomeruli in the AL. PNs send dendrites to glomeruli and axons to the mushroom body and the lateral horn, two higher olfactory centers approximately analogous to vertebrate primary olfactory cortex. (B) Overview of larval chemosensory system (Gerber and Stocker, 2007). From the three external chemosensory organs, the DO is a mixed structure composed of the central olfactory dome (grey) and a few putative taste sensilla (small circles). The cell bodies of the sensory neurons are collected in ganglia (DOG, TOG, VOG) below each sense organ (DO, TO, GO). Some of the neurons innervating the dorsolateral sensilla of the terminal organ are situated in the ganglion of the DO. ORNs (blue) send their axons via the antennal nerve (AN) into the larval antennal lobe (LAL). Local interneurons (LN) interconnect the glomeruli of the larval antennal lobe. PNs (PN; green) travel in the inner antenno-cerebral tract (iACT) to link the larval antennal lobe with the mushroom body calyx and the lateral horn (LH). An intrinsic mushroom body Kenyon cell (KC) extending its process via the pedunculus (PD) into the mushroom body lobes (not indicated) is shown in red. Axons from putative taste receptor neurons (brown) extend via the antennal nerve, the labral nerve (LN), the maxillary nerve (MN) and the labial nerve (LBN) to the suboesophageal target region (SOG).

Introduction

7

1.1.2 The odor coding system in Drosophila

1.1.2.1 Identification of Drosophila odorant receptor genes

The discovery of the first ORs was elusive for many years due to the nature of the

receptors themselves. The presence of a large number of ORs, their sequence divergence

and the low expression level made them difficult to detect until, in 1991, the first

mammalian ORs were cloned from the rat olfactory epithelium (Buck and Axel, 1991).

These proteins showed aspects that were consistent with their classification as ORs: they

are expressed specifically in the olfactory epithelium, they are members of the

superfamily of G-protein-coupled receptors (GPCRs) with seven membrane-spanning

domains as hypothesized by previous studies (Jones and Reed, 1989) and their sequences

are related. The protein’s physiological function was confirmed a few years later

(Wellerdieck et al., 1997;Zhao et al., 1998), and genes with similar properties were soon

described in other organisms (Freitag et al., 1999;Nef et al., 1996;Selbie et al.,

1992;Sengupta et al., 1996). Insect ORs were first identified in Drosophila melanogaster

by three independent groups in 1999 using different approaches (Clyne et al., 1999;Gao

and Chess, 1999;Vosshall et al., 1999). One successful approach to their isolation began

with the assumption that odorant receptors in flies, like those in mammals and

Caenorhabditis elegans, were GPCRs. A computer algorithm was then devised that

recognized proteins on the basis of structure and was trained to examine DNA databases

for proteins with structures like those of GPCRs. This algorithm identified members of

the OR gene family from the genome of Drosophila.

The Drosophila OR gene family contains 60 members that are distributed throughout the

genome, often in small clusters (Clyne et al., 1999;Robertson et al., 2003). Two of these

genes are alternatively spliced, resulting 62 odorant receptor proteins in total (Robertson

et al., 2003). 39 ORs are expressed in the adult (no including OR83b) (Vosshall et al.,

2000), and 25 ORs are expressed in the larva (Fishilevich et al., 2005) (Table 1). By

comparison, humans are believed to have about 350 functional OR genes (Malnic et al.,

2004), mice have about 1000 OR genes (Godfrey et al., 2004;Zhang and Firestein, 2002)

and mosquitoes have about 80 OR genes (Hill et al., 2002). In general, Drosophila

odorant receptor proteins are highly diverse, in many cases showing only 20% identity to

Introduction

8

each other and no similarity to mammalian odorant receptors. This diversity among

odorant receptor proteins is apparent throughout the length of the protein, albeit

conserved residues shared by many of the genes have been identified (Clyne et al.,

1999;Vosshall et al., 1999). However, closely linked genes often show a higher degree of

similarity, the two most similar receptors, OR19a and OR19b, differ by only three amino

acids, suggesting that OR gene clusters are likely to have arisen through recent genome

duplication (Robertson et al., 2003). Each ORN expresses only one or a small number of

OR genes, resulting in molecular diversity among ORNs (Clyne et al., 1999;Vosshall et

al., 1999).

Most Drosophila odorant receptor neurons co-express two different types of odorant

receptors: OR83b, a broadly expressed receptor, and one of the 61 ligand specific ORs.

OR83b is highly conserved among insect species whereas the ligand-specific receptors

are highly divergent. Electrophysiological and behavioral experiments in OR83b

knockout flies revealed that OR83b is essential for the normal function of other ORs

(Larsson et al., 2004;Neuhaus et al., 2005). Benton and colleagues later demonstrated that

not only is OR83b served as a chaperone protein that transports the ligand-binding ORs

from the cell body to the dendrite where ORs can detect odorants, but also acted as a

functional part of the receptor-complex (Benton et al., 2006). However, it still remains to

be elucidated whether OR83b is involved at all in the odorant binding. Expression patterns of the complete repertoire of Drosophila odorant receptors Antenna maxillary palp larva not detected Or47a (47F1) Or67c (67D2) Or1a (1A8) Or1a (1A8) Or92a (92E8) Or47b (47F6) Or43b (43F5) Or33c (33B10) Or2a (2E1) Or46b (46E7-8) Or7a (7D14) Or69b (69E8-F1) Or46a (46E7-8) Or7a (7D14) Or98b (98D4) Or85a (85A3) Or69a (69E8-F1) Or59c (59E1) Or13a (13F16-18) Or85f (85D15) Or67a (67B2) Or71a (71B1) Or22c (22C1) Or19a (19B3-19C) Or43a (43A1) Or85e (85B2) Or24a (24E4) Or13a (13F16-18) Or35a (35D1) Or85d (85A11) Or30a (30A3) Or22a (22A5) Or10a (10B15) Or33b (33B10) Or56a (56E1) Or9a (9E1) Or35a (35D1) Or82a (82A3-4) Or88a (88B1) Or42a (42A2) Or2a (2E1) Or49b (49D1) Or42b (42A2) Or23a (23A3) Or98a (98B5) Or45a (45F1) Or65a (65A7-11) Or85b (85A9) Or45b (45F1) Or22b (22A5) Or83c (83D5) Or47a (47F1) Or33b (33B10) Or42b (42A2) Or49a (49A5) Or33a (33B10) Or59b (59E1) Or59a (59E1) Or63a (63B1) Or67b (67B2) Or74a (74A6) Or82a (82A3-4) Or83a (83A6) Or85c (85A9) Or94a (94D9) Or94b (94D9) Table 1 Expression patterns of the complete repertoire of Drosophila odorant receptors.

(Fishilevich et al., 2005;Vosshall et al., 2000)

Introduction

9

Another fact should be noticed is that except for the odorant receptors, different odorant

binding proteins are expressed in the adult and larval olfactory organs. Compared to the

thorough understanding of the odorant receptor map of the odorant receptor neurons,

research on Drosophila odorant binding proteins or pheromone binding proteins are

largely shrinked in the last decade. Except the well described functional requirement of

OBP76a/lush in the trichoide sensillum (Jin et al., 2008;Smith, 2007), general food odor

are thought to activate odorant receptors via direct interactions without the involvement

of OBPs. This idea is supported by the fact that a lot of odorant receptors were

deorphanized in a heterologous system (Nakagawa et al., 2005;Sakurai et al.,

2004;Wetzel et al., 2001), a large number of odorant receptors expressed in the Δhalo

empty neuron system have the same odorant response profile as in the native sensillum

(Hallem et al., 2004;Kreher et al., 2005) and the dissociated olfactory receptor neurons

from the moth Manduca sexta responds to the conponent of the female moth’s sex-

pheromene blend (Stengl et al., 1992b). 35 OBP genes have been identified in the

genome of Drosophila by the use of reporter genes. The expression pattern study

revealed that 9 genes are expressed exclusively in the adult olfactory organ, several genes

that are not found in the adult olfactory organs are expressed in both larval dorsal organ

and adult gustatory organ or peripheral gustatory sensillum, for example OBP19c,

OBP56b and OBP56g (Galindo and Smith, 2001).

1.1.2.2 Characterization of Drosophila odorant receptors

The first Drosophila odorant receptor that has been functionally characterized is OR43a,

which was initially characterized physiologically following antennal overexpression

(Stortkuhl and Kettler, 2001) and heterologous expression in Xenopus oocytes (Wetzel et

al., 2001). Both studies identified that cyclohexanone, cyclohexanol, benzaldehyde and

benzyl alcohol are ligands for OR43a.

Odorants that pass through pores on the sensillum bind to ORs expressed on the dendrite

of ORNs and induce an action potential, which can be monitored using the single

sensillum recording (SSR) technique (Bestmann et al., 1996;Stensmyr et al.,

2003;Wojtasek et al., 1998). By performing the single sensillum recording, a recording

electrode is placed in the desired sensillum and captures voltage changes due to the firing

Introduction

10

of the ORNs (Figure 3.2.11). Because the sensillum contains more than one ORN, the

recording trace represents the summed activity of all the neurons housed within the

sensillum (Figure 3.2.11). In most of the sensilla, it is possible to distinguish the spikes

from different ORNs by their distinct spikes amplitude. Electrophysiological recordings

of antennal basiconic sensilla have revealed that ORNs are classified into distinct

functional classes, each with a unique odorant response spectrum (deBruyne et al., 2001).

A fundamental step forward was achieved when John Carlson’s group established a

mutant fly strain with a deletion in the locus of the receptor OR22a/b, thereby abolishing

odor-evoked responses in the ORN where the receptor is expressed without eliminating

the ORN itself, the so-called ‘empty neuron’ (Hallem et al., 2004). With this system,

based on a combination of the SSR technique and the GAL4-UAS system (Hallem et al.,

2004), it is possible to express virtually any OR, study its properties in vivo and use it as a

medium-throughput tool for Drosophila OR deorphanization, i.e. a simple way to assign

ligands to each OR (Hallem and Carlson, 2004;Kurtovic et al., 2007) (Figure 1.1.3).

Based on this approach, it was shown that the OR is not only responsible for the odorant

response spectrum in ORNs, but also for its spontaneous activity and response dynamics

(Hallem et al., 2004).

Introduction

11

Figure 1.1.3. Analysis of odor response spectra of individual odorant receptors

(Hallem and Carlson, 2004) (A) An in vivo expression system for odorant receptors. A mutant ab3A antennal neuron (Dab3A) lacks odor response due to the deletion of its endogenous receptor genes, OR22a and OR22b. Odorant receptors are introduced specifically into Dab3A using the GAL4–UAS system; an OR22a–GAL4 driver promotes transcription of the OR gene. The odorant response of the neuron (Dab3A:OrX) is then assayed electrophysiologically. (B) Mapping odorant receptors to olfactory neurons. The normal odor response of the ab3A neuron (first panel) is absent from the Dab3A neuron (second panel). Expression of Or7a in the Dab3A neuron (Dab3A:OR7a; third panel) results in an odor response spectrum resembling that of the wild-type ab4A neuron (fourth panel), indicating that E2-hexenal is a ligand for OR7a, and that OR7a is the odorant receptor in ab4A. Graphs depict the response of the neuron in spikes per second (spikes/s) to the diagnostic panel of odorants at the left. (C) Odorant receptors that have been mapped to functional classes of neurons. The eight different functional types of basiconic sensilla are designated ab1–ab8, and the neurons are named according to the sensillum in which they are found (for instance, the ab2 sensillum contains ab2A and ab2B neurons). Odorant receptors that have been mapped to basiconic neurons are indicated below the corresponding neurons. (D) Odor response spectra of antennal odorant receptors. The colored dots depict strong responses.

Electrophysiological studies in vivo have also been complemented by studies in cell

culture: Some insect ORs can be functionally expressed in human embryonic kidney 293

(HEK293) cells, HeLa cells or Xenopus laevis oocytes (Nakagawa et al., 2005;Neuhaus

et al., 2005;Sato et al., 2008;Wetzel et al., 2001;Wicher et al., 2008). The functional

characterization of insect ORs in heterologous expression systems has provided several

new insights into the molecular mechanism of insect ORs, including functional

Introduction

12

interaction between OR subunits (Neuhaus et al., 2005), novel signaling properties of

insect ORs (Sato et al., 2008;Smart et al., 2008;Wicher et al., 2008) and the role of

OR83b (Nakagawa et al., 2005;Neuhaus et al., 2005).

1.1.2.3 Odor representations in the antennal lobe and higher brain center

Several studies showed that in rodents, odorant receptor neurons expressing the same OR

converge to single glomeruli in the olfactory bulb (Mombaerts et al., 1996;Ressler et al.,

1994;Vassar et al., 1994). Studies using OR promoters to drive expression of receptors

have revealed that in Drosophila, axons of ORNs expressing the same OR also converge

onto only one or a few glomeruli (Gao et al., 2000;Vosshall et al., 2000). Recent studies

essentially completed the OR-to-glomerulus map (Couto et al., 2005;Fishilevich and

Vosshall, 2005) (Figure 1.1.4). The ORNs axon projection map is highly rigid in a sense

that the expression of ORs in ORNs does not influence the targeting of glomeruli in the

antenna lobe (Dobritsa et al., 2003;Wang et al., 2003).

Different odorants activate distinct but overlapping subset of glomeruli and the number of

activated glomeruli increases with increasing odorant concentration, as revealed by

optical imaging (Ng et al., 2002;Wang et al., 2003) and metabolic labeling studies

(Rodrigues, 1988). Therefore, odor coding in the antennal lobe appears to involve a

spatial map of odorant receptor activation. Similar electrophysiological analysis revealed

that different odorants activate different populations of projection neurons, and PN

responses were found to differ in breadth of tuning, signaling mode and response

dynamics (Wilson et al., 2004).

Another interesting question raised from these studies is to compare the odorant receptive

field of antenna and the antennal lobe. Based on the optical imaging study, the pre- and

post-synaptic odor-evoked glomerular activity was compared (Ng et al., 2002;Wang et

al., 2003), and it was found that a given odor evokes essentially the same activation

pattern regardless of whether the reporter is driven pre- or post-synaptically, suggesting

that activation of a PN simply reflects activation of its pre-synaptic ORNs.

Another group using electrophysiological recordings obtained different conclusions.

Evidence was provided that PNs are more broadly turned than ORNs, suggesting that

Introduction

13

PNs output is not only modified by the input of ORNs, but also by lateral connections

within the antennal lobe.

Studies have shown that the odor-evoked activity in the mushroom body also has a spatial

pattern (Fiala et al., 2002;Wang et al., 2001). While the response pattern of PNs is

stereotypy, the response spatial pattern in MB appears to be highly inconsistent between

individual flies. Consistent with this observation, a lack of stereotypy among individual

flies was presented in the branching pattern of individual PN axon within the MB (Wong

et al., 2002). Although the functional significance of this variability is unclear, the key

role of the MB in olfactory learning and memory (Heisenberg, 2003;Murthy et al., 2008)

raises the possibility that it might reflect experience dependent plasticity.

A spatial map of odor representation is also likely to exist in the lateral horn, although a

functional analysis of these neurons has not yet been reported. Genetic labeling of

individual PN axons using either the mosaic analysis with a repressible cell marker

(MARCM) (Lee and Luo, 1999) or FLP-out (Basler and Struhl, 1994) techniques has

revealed that PNs that connect to different glomeruli show stereotyped axon branching

patterns within the lateral horn that are distinct but overlapping, thus, allowing for the

integration of olfactory information from multiple AL glomeruli (Marin et al.,

2002;Wong et al., 2002).

Introduction

14

Figure 1.1.4 Molecular neuroanatomy of the adult AL annotated with the molecular and functional

identity of the glomeruli.

Glomeruli receiving projections from the ORNs expressing a given OR or GR are indicated, with antennal basiconic projections indicated in black, antennal trichoid projections in yellow, and antennal coeloconic projections in green. Maxillary palps projections are in cyan, and unmapped glomeruli in black dotted line. Glomeruli innervated by fruitless positive neurons are indicated in pink. This figure was adapted from Vosshall and Stocker, 2007.

1.1.3 New insights of Drosophila odorant receptor

Structural analysis in silico, in vitro and in vivo surprisingly showed that insect ORs have

a flipped topology compared with conventional GPCRs, presenting a cytoplasmic N-

Introduction

15

terminus and an extracellular C-terminus (Benton et al., 2006;Lundin et al., 2007).

Furthermore, a study published last year, pointed out that insect ORs are heteromeric

ligand-gated non-specific cation channels (Sato et al., 2008). The authors showed that

simultaneous measurements of whole-cell currents and Ca2+ influx in HeLa cells

expressing insect ORs have a response onset about 10-fold faster than what is usually

required by GPCRs. Moreover, general pharmacological inhibition of G-proteins did not

impair OR-evoked responses, as would be expected if they were GPCRs. Single-channel

recordings revealed that the response of insect ORs was not dependent on cellular

cytoplasmic components, including second messengers such as cAMP and cGMP.

Finally, different subunit compositions of the OR complex are able to shift the ion

selectivity of the measured current. This is an important finding as ion selectivity is a

unique property of ion channels, making it unlikely that ORs are associated with a

separate ion channel and suggesting that ORs themselves are necessary and sufficient to

produce an odorant-induced response.

However, a contemporary publication came to a hypothesis as regards content between

the provocative ion channel and classical GPCR theories (Wicher et al., 2008). Wicher

and colleagues showed that activation of recombinant expressed Drosophila receptor

OR22a induced opening of a cAMP-dependent CNG channel, suggesting the

involvement of Gαs proteins downstream of OR22a activation. Patch clamp recordings in

whole cell and excised patch configurations from HEK293 cells expressing OR22a and

OR83b revealed a fast odor-dependent response independent of ATP and GTP (a likely

ionotropic response) as well as a slower ATP- and GTP-dependent component. In

contrast to the study by Sato and colleagues (Sato et al., 2008), pharmacological analysis

performed by Wicher and colleagues (Wicher et al., 2008) indicate that the later

component of the odor response is mediated by a G-protein dependent signaling cascade

that includes Gαs, adenylate cyclase and cAMP. Moreover, the co-receptor OR83b alone

can generate currents after an increase of intracellular cAMP/cGMP, similar to the

currents recorded after ligand application. Finally, a mutation in OR83b can directly

modulate the ion permeability of the OR complex, showing that this protein probably

participates in the formation of the channel complex without the involvement of other ion

channels. Together, this study suggest a model in which the unique OR subunit

Introduction

16

contributes ligand selectivity and G-protein coupling, and can mediate fast activation of

the OR83b dependent conductance; OR83b is also stimulated by cAMP, but on a slower

time course.

Moreover, one of the latest studies on insect olfaction has unraveled a new class of

olfactory receptors in Drosophila melanogaster that belong to the ionotropic glutamate

receptor family (iGluRs) (Benton et al., 2009;Spletter and Luo, 2009). Since the

comprehensive analysis of the expression of classical ORs in the Drosophila has been

established (Couto et al., 2005), the lack odorant receptor on coeloconic sensilla (except

for the OR35a/OR83b-expression neuron) has hinted the existence of the other types of

insect chemosensory neurons, it seems that Benton and colleagues have discovered the

missing puzzle pieces of the whole receptor to sensillum map. This study revealed that

iGluR-like receptors (IRs) are expressed in antennal sensory neurons and confer odor

dependent responses to cells. IRs expression patterns are complementary to OR83b-

expressing neurons and might explain the remaining olfactory-mediated responses in

OR83b-null fruit flies. More importantly, this discovery highlights how multiple receptor

families can be recruited to perform similar functions in the same organ but it is yet to be

determined if IRs play a special role in fruit fly olfaction.

1.2 Heterotrimeric G-proteins

Identified by the α subunit, heterotrimeric G-proteins are composed of αβγ subunits. They

transduce, amplify and diversify the signal generated by the occupancy of a receptor by

its hormone or agonist into regulation of one or more effector systems. Receptors activate

G-proteins by increasing the affinity of the G-protein (GDP) complex for magnesium

ions. Bound magnesium causes GDP to dissociate and allows GTP to bind. This is

followed by a subunit dissociation reaction that yields active Gα (GTP) plus a Gβγ

complexed to agonist-occupied receptor. The latter can dissociate further to give free

Gβγ. As signaling molecules both the Gα (GTP) and βγ dimer modulate positively or

negatively a variety of effector functions (McCudden et al., 2005) (Figure 1.2.5).

The Gα subunits remain active until GTP is hydrolyzed to GDP. Gα (GDP) has high

affinity for Gβγ and reconstitutes into trimeric G-protein (GDP), which is ready for

another round of nucleotide exchange, activation by GTP and effector modulation

Introduction

17

through subunit dissociation. Alternatively Gα (GDP) may recombine with the βγ still

associated with the agonist-receptor complex to give a quaternary agonist-receptor-G-

protein (GDP) complex ready to be activated.

Figure 1.2.5 Standard model of the GDP/GTP cycle governing activation of heterotrimeric GPCR

signaling pathway.

(McCudden et al., 2005) Ligand binding induces the Gα GDP dissociation from the trimric Gαβγ complex. Without the ligand, Gβγ acts as a guanine nucleotide dissociation inhibitor (GDI) for Gα·GDP, slowing the spontaneous exchange of GDP for GTP. After Gα GDP dissociation, ligand-bound GPCRs act as guanine nucleotide exchange factors (GEFs) by inducing a conformational change in the Gα subunit, allowing it to exchange GTP for GDP. The cycle returns to the basal state when Gα hydrolyzes the gamma-phosphate moiety of GFP, a reaction that is augmented by GFPase-accelerating proteins (GAP) such as the Regulator of G-protein Signaling (RGS) proteins.

16 genes are identified in mammalian gene families encoding Gα subunits, 5 genes

encoding for Gβ, and 12 genes encoding for Gγ subunits (Downes and Gautam, 1999).

Gα subunit proteins can be divided into four classes based on sequence similarity: Gα(s /

olf), Gα(i / o / t / gust / z), Gα(q / 11), Gα(12/13) (Simon et al., 1991). Sizes of Gα subunits range

from 39 to 45 kilodaltons (kDa) (Nurnberg et al., 1995). Gα subunits contain two

domains: a nucleotide binding domain with high structural homology to the Ras-

superfamily GTPase that is involved in the binding and hydrolysis of GTP and a all-

alpha-helical domain that buries the GTP within the core of the protein (Sprang, 1997).

Gα subunits contain three flexible regions designated switch-I, -II and -III that change

conformation in response to GTP binding and hydrolysis (Lambright et al., 1994) (Figure

1.2.6). The helical domain is the most divergent domain among Gα families and may play

a role in directing specificity of receptor- and effector-G-protein coupling (Cabrera-Vera

et al., 2003).

Introduction

18

G-protein signaling cascade is crucial for three out of five primary senses in vertebrates

(Liman, 2006), namely vision, taste and smell. Gαt1 and Gαt2, which are also called

transducin, are specifically expressed in the outer segments of rod and cone

photoreceptor, couple to rhodopsin and mediate the phototransduction (Goc et al., 2008).

Gαgust, also named gustducin, is expressed in the taste buds, mediate the taste signaling

pathways (McLaughlin et al., 1992). A γ subunit Gγ13, is also involved in the

mammalian response to sweet and bitter compound (Huang et al., 1999). Gαolf and the

olfactory signal transduction pathway will be discussed in detail in the next section.

Figure 1.2.6 Structural features of heterotrimeric G-protein subunits.

(McCudden et al., 2005) (A) The crystal structure of Gα subunit illustrates the Ras-like GTPase domain, the helical domain and the bound nucleotide at the interdomain interface. Switch regions I, II and III are shown in yellow, GDP in green and the phosphate binding loop in blue. (B) Close view of the guanine nucleotide-binding pocket of the Gα subunit.

In the Drosophila genome, 11 genes are predicted to encode for Gα subunits; three for β

subunits and two for γ subunits (Ishimoto et al., 2005). Some of those G-proteins have

been cloned and assigned to different classes (Lee et al., 1990;Ray and Ganguly,

1994;Schmidt et al., 1989;Schulz et al., 1999;Talluri et al., 1995;Wolfgang et al., 1991).

The function of G-proteins, especially the α subunits, has been studied thoroughly in the

field of asymmetric cell division and polarization (Knust, 2001;Matsuzaki, 2005). In

Drosophila sensory system as in the mammals, G-proteins are critical for the vision and

taste. A Gαq and a Gγ protein were reported as the visual specific G-proteins (Lee et al.,

1994;Schulz et al., 1999). Gαs and Gγ1 were thought to be critical in sugar perception

(Ishimoto et al., 2005;Ueno et al., 2006).

Introduction

19

1.3 Signal transduction pathway in odorant receptor neurons

The canonical mammalian olfactory signal transduction pathway (Figure 1.3.7), involves

a G-protein specific to the olfactory system, termed Golf (Jones and Reed, 1989). Odorant

receptors are probably subjected to conformational rearrangements upon odorant binding,

similar to the light-induced structural changes of rhodopsin (Grobner et al., 2000). In

general, this conformational change initiates GDP release from the Gα-subunit that in

turn binds GTP and thereby adopts its active state. This triggers a structural

rearrangement in the heterotrimeric protein (Bunemann et al., 2003) and release of the

βγ-subunit (Hamm, 1998;Janetopoulos et al., 2001). Thereby transmitting the signal from

the extracellular to the intracellular side of the membrane (Paysan and Breer, 2001). Most

odorant activated ORs are linked to the stimulation of adenylate cyclase III (AC3)

(Bakalyar and Reed, 1990) via Golf (Jones and Reed, 1989). Activation of AC3 elicits the

increase of cyclic adenosine-3’, 5’-monophosphate (cAMP) level in the cilia (Jones, Jr. et

al., 1974;Sinnarajah et al., 1998), which then leads to the opening of cyclic-nucleotide

gated cation channels (Nakamura and Gold, 1987;Zufall et al., 1993;Zufall et al., 1994).

The CNG channels are highly permeable to Ca2+ (Dzeja et al., 1999). The influx of

cations through CNG channels depolarizes the cilia membrane, elevates the intracellular

Ca2+ concentration, and then triggers a Ca2+-activated chloride conductance that

significantly amplifies the electrical signal. The ORNs return to the inactive state

following GTP hydrolysis, which is catalyzed by the GTPase domain of the α subunit

(Markby et al., 1993). This is a classic cyclic nucleotide-mediated transduction pathway

in which all of the proteins involved have been identified, cloned, expressed and

characterized. Additionally, many of them have been genetically deleted from strains of

mice, making this one of the most investigated and best understood second-messenger

pathways in the brain (Firestein, 2001).

Introduction

20

Figure 1.3.7 The canonical mammalian olfactory signal transduction pathway.

(Firestein, 2001) Within the compact cilia of the ORNs a cascade of enzymatic activity transduces the binding of an odorant molecule to a receptor into an electrical signal that can be transmitted to the brain. AC, adenylate cyclase; CNG channel, cyclic nucleotide-gated channel; PDE, phosphodiesterase; PKA, protein kinase A; ORK, olfactory receptor kinase; RGS, regulator of G-proteins (but here acts on the AC); CaBP, calmodulin-binding protein. Green arrows indicate stimulatory pathways; red indicates inhibitory (feedback).

Until recently, much of our view of insect olfactory signal transduction pathway was

strongly influenced by this canonical pathway in mammals in addition with some

observations that were made in other vertebrates, crustaceans and nematodes (Hildebrand

and Shepherd, 1997;Krieger and Breer, 1999). Such as in nematodes (Figure 1.3.8 B),

binding of odorants to ORs induces activation of Gi and increases the level of cGMP. The

increase in cGMP levels then triggers the opening of a cyclic nucleotide gated Ca2+

channel (Krieger and Breer, 1999). In lobster olfactory receptor neurons, two second

messengers, namely IP3 and cAMP are produced in response to individual odors

(Boekhoff et al., 1994;Fadool and Ache, 1994;Hatt and Ache, 1994;Michel and Ache,

1992). In insect olfactory system, various second messengers, such as IP3, DAG, cAMP

and calcium, were found to moderate mutiple types of ion channels (Kaissling,

1996;Zufall et al., 1991a;Zufall et al., 1991b;Zufall et al., 1994;Zufall and Hatt, 1991).

Several of the regular molecular suspects have also been identified and in part

functionally characterized in insect olfactory organs. These include odorant binding

proteins (Pelosi and Maida, 1995;Tegoni et al., 2004;Ziegelberger, 1996), heterotrimeric

Introduction

21

G-proteins (Laue et al., 1997), arrestins (Merrill et al., 2002;Merrill et al., 2003;Merrill et

al., 2005) as well as a CNG (Baumann et al., 1994;Dubin et al., 1998b) and a IP3-gated

ion channel (Stengl, 1994).

Before the identification of Drosophila OR genes, there were several clues that suggested

the involvement of GPCR-mediated second messenger pathways based on biochemical

and electrophysiological evidence and the identification of the components of the cAMP

and IP3 signaling pathways in the Drosophila olfactory system. Stimulation with odorants

or pheromones on isolated ORNs increases the production of second messengers like IP3,

and in vivo recordings from antennal neurons showed that action potentials are generated

when IP3 is directly applied to the cells (Stengl et al., 1992a;Stengl, 1993;Talluri et al.,

1995). In addition, reduced expression of the Drosophila Gαq gene, dgq, and other genes

involved in phospholipid signaling induces a decrease of ORNs’ odor-evoked responses

but not their complete abolishment (Kain et al., 2008;Kalidas and Smith, 2002).

Furthermore, Drosophila mutants in phospholipid signaling have reduced olfactory

responses (Kain et al., 2009). These observations led to the assumption that insect

odorant responses are mediated by Gq-coupled GPCRs. However, other groups reported

that altering the expression of the genes rut and dnc, both of which affect the cAMP

transduction cascade, showed abnormal electrophysiological and behavioral responses to

odorants, suggesting that Gαs is also involved in the transduction mechanism (Gomez-

Diaz et al., 2004;Martin et al., 2001).

Introduction

22

Figure 1.3.8 Models of signal transduction mechanisms in odorant receptor neurons (ORNs).

(Pellegrino and Nakagawa, 2009) (A,B) Signal transduction of non-insect olfactory receptors (ORs). (A) Mammalian ORs are G-protein-coupled receptors (GPCRs) coupled to a stimulatory G-protein, Golf. After binding to an odorant, the G-protein activates adenylate cyclase (AC), which increases the intracellular concentration of cAMP. This leads to the opening of a cyclic nucleotide-gated (CNG) channel and the depolarization of the neuron. (B) In nematodes, stimulation of ORs leads to the activation of guanylate cyclase (GC) and an increase in cGMP levels. This leads to the opening of a CNG channel and the depolarization of the ORN. (C,D) Divergent views on signal transduction of insect ORs. (C) Sato et al., provides evidence supporting a model in which the OR83b/ORX complex forms an ion channel that is directly opened by the binding of the odorants and is permeable to cations (Sato et al., 2008). (D) In contrast, in Wicher et al.’s model the ligand-binding subunit (in green) is a GPCR that leads to the increase in cAMP through a stimulatory G-protein (Wicher et al., 2008). This opens the CNG-like channel OR83b (in yellow).

Objective

23

2 Objective

In all animals studied, odorant detection is mediated by seven transmembrane domain

receptors. In vertebrates, the seven transmembrane olfactory receptors are coupled to

Gαolf signaling activating ACIII, which in turn induces an increase in cAMP levels

leading to activation of cyclic nucleotide gated ion channels followed by opening of

chloride conducting channels. Compared to this well-established mechanism, the events

that follow odorant receptor interaction in insects are still controversial. The role of G-

protein-coupled olfactory signal transduction in insect is confounded by the fact that the

highly conserved OR83b receptor heterodimerizes with conventional receptors and both

Drosophila and Bombyx mori OR/OR83b dimmers directly confers ligand stimulated

nonselective channel activity in heterologous systems (Nakagawa et al., 2005;Sato et al.,

2008;Wicher et al., 2008). Cell culture and in vivo studies exhibit that Drosophila OR83b

has an inverse topology compared to the conventional GPCR (Benton et al., 2006). These

findings urged the analysis of the role of G-protein-coupled signaling in invertebrates

olfactory transduction.

The present work aimed (1) to elucidate whether and which heterotrimeric G-proteins are

involved in Drosophila olfactory signal transduction (2) to study the role of second

messengers and (3) to characterize the function of other channels in Drosophila ORNs

and to identify insect olfactory signal inhibitors.

Materials and Methods

24

3 Materials and Methods

3.1 Materials

3.1.1 Fly food

Cornmeal-molasses-yeast medium, Blommington fly food recipe

Semi-defined medium, Blommington fly food recipe

3.1.2 Enzymes and Kits and molecularbiological supplies

Agarose, Biozym

Calf Intestinal Alkaline Phosphatase, Fermentas

dNTPs, Invitrogen

Ethidium bromide, AppliChem

Gene Ruler 1kb DNA ladder, Fermentas

Go-Taq Polymerase, Promega

NuSieve® 3:1 Agarose, Biozym

pCDNA3, Invitrogen

Plasmid Maxi Kit, Qiagen®

pUAST, Brand und Perrimon; P-Element-Transformations vector with Miniwhite-

markergene.

Pure Yield™ Plasmid Maxi-Prep System, Promega

T4 Ligase, Fermentas

Wizard© SV Gel & PCR Clean Up System, Promega

3.1.3 Immunoblotting and Immunohistochemistry

Acrylamide, Sigma

AlexaFluor546, goat anti-rabbit/mouse IgG, Molecular Probes

AlexaFluor488, goat anti-rabbit/mouse IgG, Molecular Probes

Ammonium persulfate, J.T. Baker

anti-Actin, mouse monoclonal, Sigma

anti-DM-Gi, rabbit polyclonal, Juergen Knoblich, (Institute of Molecular Biotechnology,

Austria)


25

anti-DM-Go, rabbit polyclonal, Andrew Tomlinson, (Columbia University, USA)

anti-DM-Gq, rabbit polyclonal, Zvi Selinger (The Hebrew University of Jerusalem)

anti-GFP, rabbit polyclonal, Abcam #ab290-50

anti-m-Gαolf, rabbit polyclonal IgG, Santa Cruz Biotechnology

anti-m-Gαq, rabbit polyclonal IgG, Santa Cruz Biotechnology

Blotting Grade Blocker, Non-fat dry milk, BIO-RAD

Complete® Mini protease inhibitor cocktail tabs, Roche

Dithiothreitol (DTT), BIO-RAD

ECL Advance Western Blotting Detection System, Amersham/GE Healthcare

ECL PLUS, Western Blotting Detection System, Amersham/ GE Healthcare

Fatty acid-free Bovine Serum Albumin, Sigma

Gelatin from cold water fish skin, Sigma-Aldrich

Goat serum, Gibco

Immobilion transfer membrane, PVDF, Millipore

Isopropanol, J.T. Baker

Lysoszyme, Sigma

Methanol, J.T. Baker

Opti-tran BA-s85 reinforced Nitrocellulose, Schleicher & Schuell

Page Ruler Prestained Protein Ladder, Fermentas

Page Ruler Prestained Protein Ladder Plus, Fermentas

Paraformaldehyde, Prolabo

Ponceau S, Fluka Biochemika

ProLong® Antifade mounting medium, Molecular Probes

Roti®-Block, Roth

Sucrose, AppliChem

Top-Block, Fluka Biochemika

TMEMD, Fluka Biochemika

Triton X-100, Amersham Life Sciences

Tween-20, Amersham Life Sciences


26

3.1.4 Cell culture

Bovine Serum Albumin (BSA), Sigma

DMEM, Gibco

FBS, BioChrom

Penicillin/streptomycin, PAA

Trypsin-EDTA, PAA

3.1.5 Mammalian cell lines

HEK293 (Human embryonic kidney cell 293)

3.1.6 Buffers and Solutions

2X HBS: 280 mM NaCl, 50 nM HEPES, 2,8 mM Na2HPO4, pH 7.2

Blocking solution: (immunohistochemistry) 1% gelatin, 0.01% TritonX, 1-10% goat

serum in PBS-/-

Cell lifting buffer: 10 mM HEPES, 0.9% NaCl, 0.2 % EDTA, pH 7.4

Coomassie blue staining: 50% methanol, 10% acetic acid, 39.95% distilled H2O, 0.05%

coomassie blue

Destaining solution: 5% methanol, 7% acetic acid, 88% distilled H2O

DNA loading buffer: 20% ficoll 400, 100 mM EDTA, 0.025% bromophenol blue,

0.025% xylen cyanol

Drosophila ringer’s solution: 3 mM CaCl2, 182 mM KCl, 46 mM NaCl, 10 mM Tris-Cl,

pH 7.2

Electrophoresis buffer: 250 mM Tris, 191.8 mM glycine, 0.1% SDS, pH 8.3

Filter-wash buffer: 10 mM HEPES, 100 mM NaCl, 10 mM MgCl2, pH 7.4

GTP binding assay buffer: 10 mM HEPES, 100 mM NaCl, 10 mM MgCl2, pH 7.4, ±

guanosine 5′-diphosphate (GDP)

Homogenization buffers A: 10 mM HEPES, 10 mM EDTA, pH 7.4

Homogenization buffers B: 10 mM HEPES, 0.1 mM EDTA, pH 7.4

Laemmli buffer: 0.125 M Tris, 4% SDS, 20% glycerin, 0.02% bromophenol blue, pH

6.8

Larvae Whole Mount Fixative: 4% paraformaldehyde, 0.1% Triton X-100 in PBS-/-, pH

7.2


27

LB-Agar medium: 1.5% Agar, 1% bactotrypton, 0.5% yeast extract, 1% NaCl, ± 100

μg/ml Ampicillin, pH 7.2

Luria-Bertani medium: 1% bactotrypton, 0.5% yeast extract, 1% NaCl, ± 100 μg/ml

Ampicillin, pH 7.2

PBS+/+: 136 mM NaCl, 2.68 mM KCl, 0.9 mM CaCl2, 0.48 mM MgCl2, 1.47 mM

KH2PO4, 8.1 m M Na2HPO4, pH 7.4

PBS-/-: 136 mM NaCl, 2.68 mM KCl, 1.47 mM KH2PO4, 8.1 mM Na2HPO4, pH 7.4

Ringer’s solution: 140 mM glucose, 5 mM KCl, 10 mM HEPES, 2 mM CaCl2, 1 mM

MgCl2, 10 mM glucose, pH 7.4

SDS-PAGE buffer A: 0,5 M Tris, 0.4% SDS, pH 6.8

SDS-PAGE buffer B: 1,5 M Tris, 0.4% SDS, pH 8.8

Sensillum lymph ringer’s solution: 150 mM KCl, 20 mM NaCl, 2 mM CaCl2, 5 mM

glucose, 10 mM HEPES, pH 7.2

TBS: 150 mM Tris, 50 mM NaCl, pH 7.4

TBS-T: 150 mM Tris, 50 mM NaCl, 0.1% Tween20, pH 7.4

Transfer buffer: 20 mM Tris, 150 mM glycine, 20% methanol, pH 8.6

Tricholine citrate solution: 0.05 M choline-dihydrogen citrat, adjusted to pH 7.0 by

choline base solution

3.1.7 Odorants

1-octen-3-ol

1-hexanol

2-heptanone Amyl butyrate

2,3-butanedione

Benzaldehyde

Cyclohexanol

E2-hexenal

Ethyl acetate

Ethyl butyrate

Geranyl acetate


28

Isoamyl butyrate

Methyl salicylate

Pentyl acetate

Odorants were from Sigma-Aldrich and were of the highest grade available (97%-99%).

3.1.8 Miscellaneous

[35S]GTPγS (approx 1250 Ci/mmol), NEN Life Science Products

AlphaScreen cAMP Assay Kit, PerkinElmer Life and Analytical Sciences, Inc.

DMSO, J.T. Baker

Forskolin, Sigma

FURA-2, Invitrogen

IBMX, Sigma

3.2 Methods

3.2.1 Fly rearing and fly stocks

All fly stocks were maintained on conventional cornmeal-molasses-yeast medium or semi

defined medium under a 12-hr-light 12-hr-dark cycle at 18 °C or 25 °C. The following fly

stocks were kindly provided: Or83b-Gal4 (John Carlson, Yale University, USA), UAS-

mCD8-GFP, Gr21a-Gal4, Or22a-Gal4 (Bloomingtom Stock Center), UAS-Gαs-wt and

UAS-GαsQ215L (Cahir O’Kane, University of Cambridge, England), UAS-Gαo-wt, UAS-

GαoQ205L, UAS-Gαo-G203T, UAS<[w+]<Ctx and UAS<[w+]<Ptx (Andrew Tomlinson,

Columbia University, USA); UAS-Gαq and UAS-GαqQ203L, also referred as UAS-AcGq3

(Gaiti Hasan, Tata Institute of Fundamental Research, India); UAS-Gαi-wt, UAS-

GαiQ205L and UAS-GαiRNAi (Juergen Knoblich, Institute of Molecular Biotechnology,

Austria); UAS-Pacα (Martin Schwärzel, Saarland University, Germany); Or83b2 (Or83b

knock-out fly) (L. Vosshall, The Rockefeller University, USA); Ih-channel knock-out fly

(Isabelle Canal, Universidad Autonoma de Madrid, Spain). Transgenic constructs were

injected into yw embryo by VANEDIS Drosophila injection service (Oslo, Norway)

using standard procedures.


29

3.2.2 Molecular cloning and plasmid construction

All constructs were cloned according to regular cloning procedures, as outlined in

Methods in Molecular Cloning (Sambrook, 2001). Briefly, the polymerase chain reaction

(PCR) was carried out under various conditions of annealing temperature and elongation

time, using the primers outlined below. The resulting PCR products were restricted using

various restriction enzymes and ligated into the appropriate linearized plasmid vectors.

The ligation product was transformed into E.coli XL1 blue competent cells. Positive

colonies were selected for using ampicillin. Correct orientation of the insert was verified

via restriction analysis. Positive clones were then retransformed and a single colony used

to inoculate 200 ml cultures overnight at 37°C, shaking at 180 rpm. The bacterial cells

were collected by centrifugation and plasmid constructs isolated using the Plasmid Maxi

Kit from Qiagen® or Pure Yield™ Plasmid Maxi-Prep System from Promega. All

constructs were verified by sequencing.

The N-terminus of the human G-protein alpha subunit 16 (Gα16ΔC44) was amplified

from an existing construct (pCISG16 (Amatruda, III et al., 1991)) by PCR using primers

5’-cgg gat cca tgg ccc gct cgc tga cc-3’; 5’-cgg ata tcg ccc tcg ggg ccg tcc ac-3’. The

PCR fragment was cloned into the BamHI and EcoRV sites of the mammalian expression

vector pCDNA3 (Invitrogen). Different cDNA fragments coding for the Drosophila G-

protein alpha subunits’ C-terminus 44 amino acids were generated from Drosophila head

cDNA by polymerase chain reaction and fused into the EcoRV and XbaI sites of the

pCDNA3-Gα16-ΔC44 construct. The primers used for the PCR were: Gαs-fw, 5’-gct agc

gga gac gga aaa c-3’, Gαs-rw, 5’-cgt cta gat agt aac aat tca tat tga cga agg-3’; Gαq-fw, 5’-

cat tag ttt aga tac ata taa-3’, Gαq-rw, 5’-cgt cta gat aga aac aga tta ctt tct ttt agg-3’; Gαo-

fw, 5’-aac aaa tca acc tca aaa g-3’, Gαo-rw, 5’-cgt cta gat agg tac agt cca cag ccg cg-3’;

Gαi-fw, 5’-aac aag cga aaa gac caa aag g-3’, Gαi-rw, 5’-cgt cta gat agg aat aag cca att tgt

ttc ag-3’; Gα73B-fw, 5’-ctg ggt acc tcg gaa agg gag-3’, Gα73B-rw, 5’-cgt cta gat agg aat

agg ccc atg ctg gac ac-3’.

A Gαs-GFP fusion construct was generated from GFP and Drosophila Gαs using

polymerase chain reactions that produced DNA fragments with overlapping ends that

were combined subsequently in a fusion polymerase chain reaction. GFP was inserted

between Gαs residues 71 and 72. A 6-residue linker sequence (SGGGGS) was inserted at


30

both of the junctions between Gαs and GFP. The GFP with linkers was generated first by

polymerase chain reactions, then the linker GFP was used as template in the fusion

polymerase chain reactions. The primers used for the PCR were: GFP-linkerf: 5’-tca ggt

gga gga ggt tct gtg agc aag ggc gag gag c-3’, GFP-linkerr: 5’-gct gcc gcc gcc gcc gct ctt

gta cag ctc ctc cat gc-3’, Gs1f: 5’-ggc cag atc tac cat ggg ttg ctt tgg gtc gcc c -3’, Gs1r:

5’-cac aga acc tcc tcc acc tga tcc gtc gac atg caa tat tcg c-3’, GFPsf: 5’-gaa tat tgc atg tcg

acg gat cag gtg gag gag gtt ctg tga gc-3’, GFPsr: 5’-ctg ttt ctt ttc cga gtc aga aaa gct gcc

gcc gcc gcc gct c-3’, Gs2f: 5’-gcg gcg gcg gcg gca gct ttt ctg act cgg aaa aga aac ag-3’,

Gs2r: 5’-gcg cgg ccg cct ata aca att cat att gac g-3’. The fusion fragment of Gαs-GFP was

inserted in the BglII/NotI site of pUAST. All constructs were verified by sequencing.

3.2.3 Cell culture and transfection

Reagents for cell culture use were purchased from Invitrogen, unless stated otherwise.

Human embryonic kidney (HEK) 293 cells were maintained in Dulbeccos Modified

Eagles Medium (DMEM), containing 10% FBS, 100 units/ml penicillin/streptomycin and

2 mM/l glutamine, to a confluency of ~70%. 5 - 10 μg plasmid DNA was transfected per

35x10mm dish (Falcon). HEK293 cell transfections were performed using a standard

calcium phosphate precipitation technique: The appropriate volume of plasmid DNA

solution and CaCl2 was added to H2O to have a final volume of 100 μl. 100 μl 2XHBS

was then added drop-wise before the solution was gently mixed, incubated at room

temperature for 15 minutes and finally added drop-wise to the cells. 4 - 12 hours post-

transfection, the medium was exchanged with fresh DMEM. Cells were subjected to

experimentation 48 hours post-transfection to allow for an optimum level of olfactory

receptor expression.

3.2.4 Ratiometric calcium imaging in HEK293 cells

HEK293 cells were washed once in PBS+/+ and loaded with a calcium-sensitive

fluorometric dye FURA-2-AM (Molecular Probes) for 45 minutes in the dark. FURA-2-

AM is a ratiometric calcium indicator and exhibits an absorption shift upon Ca2+ binding.


31

This shift can be observed by scanning the excitation spectrum between 300 and 400 nm,

while monitoring the emission at ~510 nm.

Ratiometric calcium imaging was performed as described (Spehr et al., 2003) using an

inverted microscope equipped for ratiometric imaging. Images were acquired from

randomly selected fields of view, and integrated fluorescence ratios (f 340/f 380 ratio)

were measured. WinNT based T.I.L.L Vision software was used to collect and quantify

spatiotemporal Ca2+-dependent fluorescence signals. At time-point 20 seconds, odorant

(500 μM) was applied for 10 seconds using a specialized micro-capillary application

system, followed by washing with Ringer’s solution for 30 seconds. At the end of each

experiment, 20 μM ATP was applied to the cells as a positive control.

3.2.5 RNA isolation, reverse transcription, RT-PCR and Real-time quantitative

PCR

The third antennal segments were collected from 100 flies (3-5 days old). RNA was

isolated with Trizol Reagent (Invitrogen) and cDNA was synthesized using MMLV

reverse transcriptase (Invitrogen).

Real time quantitative PCR was performed in Bio-Rad iQ™5 Real-Time PCR Detection

System using Bio-Rad iQ™ SYBR® Green Supermix kit, according to manufacturer’s

recommendations. The following amplification protocol was used: 40 cycles of 15 s at 95

°C, 20 s at 60 °C and 40 s at 72 °C. Expression of the target gene was normalized to

Drosophila rp49 (ribosomal protein 49) RNA levels. The delta Ct (cycle threshold)

method was used to calculate relative expression levels, as previously described (Livak

and Schmittgen, 2001). Results are reported as fold change in gene expression relative to

control conditions.

The following primer pairs have been used in RT-PCR and real-time quantitative PCR:

CG2812f11: 5’-AAT CGA GGG ACC TGG ATT TG-3’

CG2812r255: 5’-ATG GTG TTT CGT GCG CTT G-3’

CG3004f47: 5’-ACA CCA TTA AGG TGT GGC AGG-3’

CG3004r203: 5’-TTG GAC TCC AGG TCG TAC AGC-3’

CG17760f406: 5’-AAG GAG TGC TAC AAT CGT CG-3’


32

CG17760r612: 5’-TCG CTG ACC AGC GAC GTC CAC C-3’

CG17766f392: 5’-TCG AAC TCG GAT TCG GAA TTG-3’

CG17766r611: 5’-TGG TGC GCT TTA CCC AAT CTG-3’

CG30054f833: 5’-TGT ATT CGC ATT TGG TAG AC-3’

CG30054r1019: 5’-TCC TTA ACT GCA GCG AAC AC-3’

Gsfa440: 5’-TTC TTC AAA CCT ATG AGA GG-3’

Gsra661: 5’-TCC TAC GCT CGT CCC GCT GG-3’

Gsbf428: 5’-AAG ACA AGG GCG TTC TTC AA-3’

Gsbr639: 5’-ACC GAC ATC GAA CAT GTG AA-3’

G49bQ3af471: 5’-TCT CGA TCG TGT GGC TCA ACC-3’

G49bQ3Ar768: 5’-AGG GTA TGT AAT TAT AGT ACG-3’

G49bretinalqaf805: 5’-AAG AAG GAC TTG TTG GAA GAG-3’

G49bretinalqar1028: 5’-ATA ATT GTA TCT TTG ACA GC-3’

G49BQ3bf534: 5’-GCC CAC AAC AGG GAT AAT TG-3’

G49BQ3br786: 5’-TGA CGA ATT TTG AAA CCA AGG-3’

G49bretinalqbf766: 5’-CCT TGG TTT CAA AAT TCG TCA-3’

G49bretinalqbr1019: 5’-TCT TTG ACA GCG CAG AAC AC-3’

Galpha73Bf323: 5’-ATT TCG GCA GCT GTA CCA GCG-3’

Galpha73Br567: 5’-GTG CAG AAT GTC CTC GGT GC-3’

CG7095f1588: 5’-TTC TGG ATA GCA GTC AAT CG-3’

CG7095r1709: 5’-TTG CCA TCA ATA TTA ATC TCG-3’

3.2.6 [35S] GTPγS binding assay

Τransfected cells were homogenized in 10 mM HEPES, 10 mM EDTA (pH 7.4) and

membrane preparation was performed as described previousely (Dowling et al., 2004).

[35S] GTPγS binding was assayed in a final volume of 100 μl (pH 7.4), containing 10

mM HEPES, 100 mM NaCl, 10 mM MgCl2, 100 mM guanosine 5′-diphosphate (GDP)

and 0.1 nM [35S] GTPγS. The incubation was started by the addition of membrane

suspension (about 50 μg of membrane protein per reaction) and was carried out in

triplicate for 30 min at 30 °C. The reaction was terminated by addition of 3 ml of ice-cold


33

filter wash buffer and rapidly vacuum filtered through glass fiber filters (Whatman

GF/C). Filters were washed three times with 3 ml of wash buffer and transferred to

scintillation vials. The scintillation cocktail was added and counted in a Packard Tri-Carb

liquid scintillation counter.

3.2.7 Larval chemotaxis assay

Chemotaxis assay was performed as previously described (Heimbeck et al., 1999). Daily,

flies were transferred into a fresh food vial where they could lay eggs until the following

day when they were transferred to a fresh vial once again. When third-instar larvae

appeared in the vial, usually at 108 h after beginning of the egg laying period in a 25°C

culture condition, a 15% sucrose solution was used to wash out the larvae from the food.

After washing twice with distilled water, 50 larvae were used immediately.

Tests were done on Petri dishes (diameter, 85 mm) covered with a layer of 1.5% agarose.

Plates were air-dried before use to avoid diffusion of the test substance. Odorant and

control diluent (water or paraffin oil) were placed on lids of 1.5 ml eppendorf micro test

tubes to avoid diffusion through the agarose. 50 larvae were handpicked in the center of

the plate before adding the test substances. The dish was immediately covered with the

lid. After 5 min, larvae were counted as shown in Figure 3.2.9 A. Only larvae on

semicircular areas (radius, 30 mm) around the filter disks were included. Thus, the

animals had to move at least one body length toward the source. We then calculated a

response index (RI): (Ns – Nc) /(Ns + Nc). Ns represents the number of animals at 30 mm

from the odorant source (inside area S in Figure 3.2.9); Nc is the number of larvae found

inside an identical surface on the opposite (control) side. Positive RIs indicate attraction;

negative RIs indicate avoidance; and RI = 0 indicates indifferent behavior.


34

Figure 3.2.9 Larval chemotaxis assay

Schematic representation of the test setup. Small filter paper containing a test chemical (S) and control diluent (C) are placed on opposite sides of a Petri dish covered with a layer of agarose. Fifty animals are transferred to the start point and counted after 5 min in indicated semicircular areas. (Heimbeck et al., 1999)

3.2.8 Electroantennogram (EAG)

The EAG responses of wt and mutant flies were recorded as previously described

(Neuhaus et al., 2005;Stortkuhl and Kettler, 2001). Briefly, 2 or 3-day-old flies were

mounted in truncated micropipette tips with the anterior portion of the head protruding

from the end of the tip. The reference electrode was inserted into the haemolymph of the

head capsule. The recording electrode was placed on the frontal surface of the anterior

aspect of the antenna. After obtaining a stable baseline, EAG recordings were initiated by

a short odor pulse (0.3 s, precisely controlled by the odorant application system), applied

into an air stream that was directed toward the antenna (Figure 3.2.10). All odorants were

dissolved in paraffin oil at given concentrations. Odorants used were cyclohexanol,

benzaldehyde, heptanone and ethyl acetate, etc.


35

Figure 3.2.10 Schematic overview of the EAG recording

(A) The reference electrode was inserted into the haemolymph of the head capsule. The recording electrode was placed on the frontal surface of the anterior aspect of the antenna. (B) Illustration of a typical EAG recording setup.

3.2.9 Antennal single sensillum recording

The procedures of extracellular single sensillum recording was essentially similar as

previously reported (Yao et al., 2005). Canton-S flies aged < 1 week were used for wild-

type recordings. hs-flp; Or83b-Gal4, UAS<[w+]<CTX flies were recorded after standard

heat shock (one hour, 38°C) at age 4-5 days. Electrical activity of the neurons was

recorded extracellularly by placing an electrode filled with sensillum lymph ringer

solution in the base of the sensillum. A reference electrode filled with the same ringer

solution was placed in the eye (Figure 3.2.11). Signals were amplified using a patch-

clamp amplifier (L/M-PC, List-Medical Electronic, Darmstadt, Germany) set in voltage-

clamp mode and fed into a computer via a 16-bit analog/digital converter (Digidata

1200A; Molecular Devices, CA, USA). Electrophysiology data were recorded by the

WinEDR software (Strathclyde Electrophysiology Software, University of Strathclyde).

Impulses during the 1 s period before stimulation and the 1 s during stimulation were

counted off-line using the Mini Analysis software (Synaptosoft, Decatur, GA, USA). The

spikes were sorted into different groups according to their amplitude as previously

described (deBruyne et al., 2001). Each sensillum was tested with multiple odorants, and

no more than two sensilla were analyzed per fly.


36

Figure 3.2.11 Schematic overview of single sensillum recording

Electrode positions for the extracellular recording of voltage differences between the sensillum lymph (L) and the haemolymph AC, auxiliary cells; EC, epidermal cells. Figure adapted from de Bruyne et al., 1999

3.2.10 Labellar single sensillum tip recording

The single sensillum tip recordings from fly labellum were preformed on the same setup

as the recording from the antenna. Except for substance application and electrode placing,

most of the experimental procedures and data analysis were similar to the antennal single

sensillum recording. Particularly, the recording electrode was placed on the tip of the

sensillum, and the gustatory substance was applied in the recording electrode. A

tricholine citrate solution was used to dissolve the stimulation substance to inhibit the

water response of gustatory receptor neurons (Ishimoto and Tanimura, 2004) (Figure

3.2.12).

Figure 3.2.12 Schematic overview of single unit tip recording from GRNs

The electrode is used for both stimulating and recording (Ishimoto and Tanimura, 2004).


37

3.2.11 Odorant application system in EAG and single sensillum recording

Flies were under a constant stream of humidified charcoal-filtered air (2.4 l/min, 22-25

°C) to prevent any potential environmental odors from inducing activity during these

studies. Pure odorant substance was diluted in paraffin oil, and 50 μl was applied to a

filter paper and put into a 5 ml syringe, the volume of the syringe was adjusted to 4ml,

and an air stream of 0.4 l/min was applied to go through the syringe for one or three

second. The odorant was then delivered to the fly with the background air. At the same

time, an equal amount of air was cut from the background to make the input air pressure

on the fly constant before and after the stimulation. An electronic device controlled this

application system, and this device is connected to the digital converter, thus the

application process was recorded together with the electrophysiological recording.

Since this odorant application system was controlled by the flowmeter, and the odorant

was contained in a sealed syringe, by weighting the syringe before and after the

application, combined with the whole air volume that pass though the fly, the exact

concentration of the odorant that delivered to the fly could be calculated.

3.2.12 Immunohistochemistry

Fly heads were cut, fixed for 3 hours in 4% paraformaldehyde at 4 °C and subsequently

incubated overnight in 25% sucrose in Drosophila Ringer’s solution. Cryosectioning was

performed to produce 12 µm sections. After blocking with 5% goat serum in Drosophila

Ringer‘s, the primary antibody was applied to the sections overnight at 4 °C or 3 hours at

room temperature. Subsequently slices was washed with PBS containing 0.01% triton X-

100 (30 min, 3 times) and the secondary antibody coupled to A546 or A488 (Invitrogen)

was applied for 1 hr at room temperature. After washing again (30 min, 3 times), slices

were mounted in ProLong® anti-fading mounting medium (Molecular Probes). Pictures

were taken with a Zeiss confocal microscope (LSM510 Meta; Zeiss).


38

3.2.13 Whole mount fluorescent antibody staining of Drosophila larvae

Larvae were removed from the culture vial and transferred into a clean plastic Petri dish

with fixative after food residues were rinsed away. The larvae were dissected in the

fixative solution, the anterior tips, including the brain, imaginal discs, salivary gland and

mouth hooks, were used for further staining. Fixation was carried out on ice for 1 hour.

The larvae pieces were washed with 0.1% Triton X-100 (PBS-Triton), 3 times 10 minutes

at room temperature. The tissue was then blocked for 30 min at RT (1XPBS, 0.1% Triton

X-100, 5% heat inactivated normal goat serum). Afterwards the block solution was

replaced with primary antibody solution, incubated at RT for 2 hours, followed by

washing 3 times 10 minutes with 0.1% Triton PBS. Secondary antibody was then applied

for 1 hour at RT in the dark and samples were washed subsequently three times for 10

minutes with 0.1% Triton PBS. Finally, the tissue was mounted and pictures were taken

with a Zeiss confocal microscope (LSM510 Meta; Zeiss).

3.2.14 SDS gel electrophoresis and Western-blotting

200 or more antennae were homogenized in Laemmli buffer (30% glycerol, 3% SDS, 125

mM Tris/Cl, pH 6.8), resolved by 10% SDS-PAGE and transferred to nitrocellulose

membrane (Protran; Schleicher & Schuell). The membranes were blocked with 5% non-

fat dried milk (Bio-Rad) in TBS-T and incubated with primary antibody (diluted in TBS-

T). After washing and incubation with HRP coupled secondary antibodies, detection was

performed with ECL plus on Hyperfilm (Amersham).

3.2.15 cAMP assay

PerkinElmer alpha screen cAMP assay kit was used for the measurement. Antennae were

cut manually from 2-5 days old adult flies, and immediately homogenized. The

homogenized tissue of approximately 10-15 antennae was finally aliquoted into each

reaction well. The assay was then performed according to the manufacturer’s instruction.

Results

39

4 Results

4.1 Different Gα subunits are expressed in Drosophila antenna.

Drosophila ORs were originally classified as G-protein-coupled receptors, assuming

expression of G-proteins in Drosophila antennae. We therefore set out to determine G-

protein expression and designed RT-PCR, western blot, and immunostaining experiments

to study the expression level and expression pattern of G-protein alpha subunits in the

antenna.

4.1.1 RT-PCR analysis of Gα subunits in Drosophila antenna

By searching the database of Drosophila genome, we collected the sequences of 10

transcript variants translated from 9 genes that were classified to encode for

heterotrimeric G-protein alpha subunits, designed primers against those variants and

performed RT-PCR. Gα q3 and the retina specific Gα q1 are transcribed from Dmel\Gα49B

(CG17759) (Talluri et al., 1995). Dmel\Gsα60A has two transcript variants, which differ

by inclusion or deletion of three amino acids and substitution of a Ser for a Gly (Quan

and Forte, 1990). We could show that both of the transcripts are expressed in the antenna,

however primers used here were designed for the identical sequence of these two

transcript variants. About 200 antennae were cut manually; RNA isolation and cDNA

synthesis were subsequently performed. RT-PCR results revealed expression of all Gα

subunits transcripts in the antenna, with a higher expression level of Gαi, Gαq1-retinal,

Gαq3, Gαs, CG17766, CG17760 and Gαo as compared to that of Gαf, CG3004 and

CG30054 (Figure 4.1.13 A).

Heterotrimeric G-protein α subunits are divided into 4 classes based on their sequence

similarity and downstream effectors. Together with some well-identified Gα subunits

from other species, a phylogenetic tree of Gα subunits was generated by the MegAlign

program using the Clustal V method (Figure 4.1.13 B). As shown, 3 classes of Gα can

also be identified from Drosophila. The ungrouped G-proteins are Gαf, CG3004 and

CG17766, which show a low similarity with those classified Gα. These proteins also have

relatively low expression level in antenna from the RT-PCR result, they were not directly

investigated in my study.

Results

40

Figure 4.1.13 PT-PCR analysis of Gα subunit expression in Drosophila antenna.

(A) RT-PCR analysis showed that most of the Gα subunits found in the Drosophila genome are expressed in the antenna. (B). The phylogenetic tree of Gα subunits from Drosophila. melanogaster, Mus. musculus, Homo sapiens, Anopheles gambiae, Panulirus argus and Rattus norvegicus.

4.1.2 Western-blot analysis showed that different classes of Gα are expressed in

the antenna

Fly antennae were collected manually as described before, followed by a standard

western-blotting procedure. Antibodies against Gαs, Gαq, Gαi and Gαo were used. Each

lane contained protein of approximately 50 antennae. The blot shown in Figure 4.1.14 A

confirms expression of Gαs, Gαq, Gαi and Gαo in the antenna. Gαs was detected by a

mouse Gαolf antibody, since Drosophila Gαs shows strong homology to vertebrate Gαolf,

especially at the C-terminus (Figure 4.1.14 B). The anti-Gq antibody purchased from

Santa Cruz Biotechnology has been raised against the C-terminal peptide

(FAAVKDTILQLNLKE YNLV) of mammalian Gq. This differs from the corresponding

Drosophila Gq3 sequence by a single residue (FAAVKDTILQSNLKEYNLV)

(Ratnaparkhi et al., 2002). Anti-DM-Go was provided by Zvi Selinger (The Hebrew

University of Jerusalem), and anti-DM-Gi was provided by Juergen Knoblich (Institute of

Molecular Biotechnology, Austria).

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41

Figure 4.1.14 Western blot analysis of G-protein α subunits expression in antenna.

(A). Western blot analysis detected the expression of Gαs, Gαq, Gαi and Gαo in antenna. (B). Alignment of Drosophila Gαs, human Gαs, and human Gαolf shows a high degree of homology between these sequences, especially at the C-terminus.

4.1.3 Immunohistochemical analysis of the expression patterns of different Gα

proteins in Drosophila antenna

Although RT-PCR and western-blot analysis strongly supported the idea that G-proteins

are present in fly antenna, we next attempted to study their cellular localization in the

Results

42

antenna. Therefore, we performed immunostaining experiments. As mentioned before,

ORNs are not the only cell type in the antenna; cell bodies of the ORNs are surrounded

by specialized auxiliary cells and epidermal cells. For the purpose of marking the ORNs,

the UAS-GAL4 system was used to generate a fly line that expresses a membrane

targeted GFP (mCD8-GFP) under the control of an OR83b driver. As described

previously, OR83b serves as a general ORN marker. In this fly line, OR83b positive cells

are all labelled with GFP (Figure 4.1.15 B). The UAS-GAL4 system is a biochemical

method used for the ectopic overexpression of transgenes. GAL4 encodes a protein

identified in the yeast Saccharomyces cerevisiae as a transcription activator and UAS

(Upstream Activation Sequence) is a short section of the promoter region to which the

Gal4 protein specifically binds to activate gene transcription. For studies in Drosophila,

the GAL4 gene is placed under the control of a native gene promoter, or driver gene,

while the UAS controls the expression of a target gene. Gal4 is then only expressed in

cells where the driver gene is usually active. In turn, Gal4 should only activate gene

transcription where a UAS has been introduced (Figure 4.1.15 A).

Immunostaining was performed on the antennae cryosection of the OR83b-GAL4, UAS-

mCD8-GFP fly. Gαo and Gαi were homogeneously expressed in the whole antenna (Gαo

data not shown), while Gαq was detected in a subset of ORNs, and Gαs was found to be

expressed in the dendritic part of the ORNs, where the initial olfactory signal

transduction takes place, suggesting that Gαs may play an essential role in the olfactory

signal transduction pathway (Figure 4.1.15 C).

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Figure 4.1.15 Immunohistochemical analysis of G-protein α subunit expression patterns in antenna.

(A) The UAS-GAL4 system. (B) Front view of the OR83b-GAL4; UAS-mCD8-GFP fly. Antennae and maxillary palps express GFP. Scale bar = 100 μm. (C) Immunostaining showing different expression patterns for different Gα proteins. Scale bar = 20 μm.

4.1.4 Real-time quantitative PCR analysis of Gαs and Gαq expression in Drosophila

antenna and head

Since both Gαs and Gαq show an interesting expression pattern in the antenna, a real-time

quantitative PCR was performed to compare the mRNA level of those two proteins in the

antennae and the head. Antennae and fly head cDNA were synthesized and qPCR was

carried out. The delta ct method was used to analyze the relative gene expression level.

The G-protein expression levels were first normalized to the endogenous control gene

rp49, using the expression level in the female head as control. The data shown here are

presented as percentage of controls. The results indicate that Gαs expressed in the

Results

44

antennae was about 40% of the amount expressed in the female head, and the Gαq3 in

antenna was about 80% compared with the female head (Figure 4.1.16). However, a

student t-test showed that the differences between the antenna, male head and female

head are not significant.

Figure 4.1.16 RT-qPCR analysis of Gαs and Gαq expression level in antenna.

(A) The relative Gαs expression level in Drosophila antenna, male head and female head. Gαs expression level in the antenna was about 40% of that expressed in the female head. The Gαs expression level of the male head is significantly higher than that of the female head (p = 0.024). The difference between antenna and male, female head are significant (p < 0.01). (B) The relative Gαq3 expression level in antenna, male head and female head. The expression level of Gαq3 expression in those tissues showed no significant difference. Error bars represent S.E.M.

4.2 Functional study of Gα subunit in Drosophila olfactory signal

transduction pathway

4.2.1 Overexpression study of different Gα subunits and G-protein specific protein

toxins in Drosophila ORNs

Due to the rapid expansion of the use of the UAS-GAL4 system in the last 20 years, it

becomes fairly easy to target gene expression in Drosophila. In our study, we collected

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45

most of the available UAS fly lines of Gα-proteins and toxins (Hampoelz et al.,

2005;Katanaev et al., 2005;Ratnaparkhi et al., 2002), crossing those flies with the

OR83b-GAL4 driver line, to generate different Gα-protein or G-protein specific toxin

overexpression flies. With the use of the OR83b-GAL4 driver, protein overexpression

was targeted specifically to the ORNs. If the G-proteins were essential for the olfactory

signal transduction, we expected olfactory aberration to occur in some of those transgenic

flies. We screened those flies for olfactory deficits by using EAG recordings, and it

turned out that the only fly line showing a severe defect in odorant induced signaling was

a fly line expressing CTX, which is an ADP-ribosyltransferase that typically activates

Gαs proteins (Figure 4.2.17 A). Data are shown for ethyl acetate stimulation. Over-

expression of wt Gαo, GTPase deficient (active) Gαo, constitutively GDP bound

(inactive) Gαo, pertussis toxin (PTX), GTPase deficient (active) Gαq3, wt Gαs, GTPase

deficient (active) Gαs, wt Gαi, GTPase deficient (active) Gαi, and Gαi RNAi did not show

any defect in EAG recordings. Other odors, for example cyclohexanol, benzaldehyde and

heptanone were also tested on those flies and similar results were obtained from EAG

recordings.

4.2.2 Dose-response curves of EAG responses from CTX and control flies.

By measuring the EAG response amplitude of G-protein, mutant G-protein and toxin

overexpression flies, we found that the expression of CTX in ORNs can cause a reduction

in EAG amplitude. CTX is a protein complex secreted by the bacterium Vibrio cholerae.

The cholera toxin is an oligomeric complex made up of six protein subunits: a single

copy of the A subunit, and five copies of the B subunit. Subunit B is responsible for the

toxin binding to the cell surface, and induces endocytosis of subunit A, which acts as a

toxin: once inside the cell, it ribosylates Gαs and leads to a constitutive cAMP production

(Zhang et al., 1995). The cholera toxin subunit A is generally used in heterotrimeric G-

protein studies as a non-cytotoxic and irreversible activator of Gαs (Burton et al., 1991).

Due to its potential toxicity, the protein toxins CTX and PTX were expressed under an

inducible heat-shock promoter based on the yeast Flippase/FLP recognition target

(FLP/FRT) recombination strategy (Strapps and Tomlinson, 2001). The CTX or PTX

gene in the UAS response line was intruded by a marker gene and a stop codon, thus the

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46

toxin expression can start only after the removal of the insertion part. The insertion part

was cut out by the activation of the flippase gene, which was under the control of a heat

shock promoter (Figure 4.2.17 G). To ensure that the observed defects were not caused

by the heat shock process itself, we tested wt flies, OR83b-Gal4; UAS-CTX, hs-flp flies

(CTX), heat shocked OR83b-Gal4; UAS-CTX, hs-flp flies, (CTX hs), and heat shocked

wt flies with four different odorants of different concentrations. We found that only heat

shocked CTX expressing flies showed EAG response deficits (Figure 4.2.17 C-F). CTX

expression did not alter the expression level or cellular distribution of Gαs, and did not

disturb the general cellular morphology in the antenna (Figure 4.2.17 B).

Figure 4.2.17 Screening of G-proteins for participation in the olfactory signal transduction

(A) UAS constructs of different Gα-proteins, mutated Gα-proteins and G-protein affecting toxins were expressed in the sensory neurons of the third antennal segment using an OR83b-Gal4 driver line. EAG amplitudes (mV) in response to application of ethyl acetate (pure) were recorded. Error bars represent SD. (B) Expression pattern of Gαs in the antenna of wt and CTX expression flies. (C-F) EAG amplitudes (mV) upon exposure to different concentrations of 4 odorants in wt flies, OR83b-Gal4; UAS-CTX flies (CTX), heat shocked OR83b-Gal4; UAS-CTX flies (CTX hs), and heat shocked wt flies (wt hs). Error bars represent S.E.M. (G) Schematic representation of the CTX expression strategy. The white gene and a stop codon were inserted upstream of the toxin gene. Once the FLP is activated, it will induce the recombination

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between two FPT sites, the white gene plus stop code will be cut out, and then the protein toxin will be expressed in the target cell.

4.2.3 CTX blocks the olfactory response of ORNs in Drosophila

To further investigate how CTX affects the odorant response on a cellular level, we

performed single sensillum recordings from CTX flies. Single sensillum recording

usually monitors all of the neuronal activities from one sensillum, and the spikes from

different neurons can be resolved into different populations. The sensillum we focused on

here was ab1 (antenna basiconic sensillum 1), which compartments 4 neurons: 3 ORNs

(neuron A, B and D) and one GRN (neuron C). 4 classes of spikes could be distinguished

in the recording traces (Figure 4.2.18).

Figure 4.2.18 Single sensillum recording from wild type ab1 sensillum.

4 populations of spikes can be resolved in the recording trace.

The response of the ab1 sensillum is shown in Figure 4.2.19 A-E. The ab1 sensillum

contains three neurons that express OR83b, and the one expressing the gustatory

receptors Gr21a and Gr63a, which are thought to respond to CO2 (Jones et al.,

2007;Kwon et al., 2007). In our recordings we found that expression of CTX not only

inhibits the response to odorant stimuli, but also blocks the spontaneous activity of ORNs

(Figure 4.2.19 B, G). Only one population of spikes could be observed in the CTX

expressing ab1 sensillum, and odorant responses were abrogated (Figure 4.2.19 B). In

addition to the comprehensive ORNs driver OR83b-GAL4, other OR drivers that only

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direct the expression of one class of ORNs were also used for this study. As expected,

when CTX was expressed in a single class of ORNs, for example OR22a neurons, the

neuronal activity of the target ORNs was disrupted (Figure 4.2.19 F, G). The ab3

sensillum normally houses two ORNs, both of which respond to 2-heptanone (Figure

4.2.19 F). Expression of CTX in ab3A neurons causes loss of spontaneous activity and

only one group of neuronal activity can be observed in the recordings, which is that of the

ab3B neuron still responding to the odorant (Figure 4.2.19 G).

Figure 4.2.19 Single sensillum recordings from CTX and control flies

(A) ab1 sensillum response to ethyl acetate. (B) CTX expressing ab1 sensillum showed only one group of neuronal activity, and does not respond to ethyl acetate. (C) Wt ab1 neuron responses to CO2. (D) The CTX expressing ab1 sensillum showed only one group of neuronal activity and this neuron also responds to CO2. (E) Expression of CTX in Gr21a neurons does not influence the CO2 response. (F) The ab3 sensillum

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responds to 2-heptanone. (G) In the ab3 sensillum, OR22a-GAL4 driven expression of CTX leads to silencing of the OR22a neuron (ab3a neuron). The ab3A neuron activity was gone, however the ab3B neuron still responded to 2-heptanone.

4.2.4 CTX does not act as a non-specific toxin on cells

We already showed that CTX expressing sensilla have a normal morphology and results

from two further experiments also support the idea that CTX does not act as a toxin in

cells.

In the first set of experiments we were able to show that CTX expression in gustatory

sensory neurons does not impair their normal responses to the stimuli tested. The driver

lines we used were Gr21a-GAL4 and Gr5a-GAL4, and no response alterations could be

observed in the recording traces (Figure 4.2.19 C, E and Figure 4.2.20 A, B), which as

well means that the gustatory response of Gr21a and Gr5a neurons is independent of a

Gαs-coupled pathway.

Additionally we performed larval chemotaxis assays on the OR83b-GAL4; UAS-CTX

larvae. Unlike in adult flies, CTX expression in larvae did not influence their olfactory

response. Details of the assay are described in part 5.4.2.

Figure 4.2.20 Gr5a-CTX GRNs response to sucrose

(A, B) Labellar single sensillum tip recordings showed both Gr5a-CTX fly and wt fly respond to sucrose.

4.2.5 Functional study of OR and Gα protein interactivity in the recombinant

HEK293 expression system

4.2.5.1 Chimera Gαq16s protein improves the OR signaling cascade.

To further investigate the functional interactivity between ORs and G-proteins, we

expressed Drosophila OR43a and OR83b in HEK293 cells, and performed single cell

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calcium imaging. In this assay, ligand-mediated OR activation triggers an endogenous

Gαq dependent calcium cascade, and this signal can be monitored and recorded. When the

odorant receptors were transfected without any G-proteins, only a weak response to the

odorant stimulation (1 mM cyclohexanone) could be detected. This might be due to the

weak coupling of Drosophila receptors to the endogenous calcium pathway, or as

demonstrated in the previous experiments be based on OR coupling to Gαs rather than

Gαq. The C-terminus of the G-protein α subunit is a key determinant for the fidelity of

receptor coupling. The Drosophila and human G-protein chimera Gαq16x (human Gαq16,

C-terminus 44 amino acids replaced by Drosophila Gαx C-terminus 44 aa) is supposed to

couple to Drosophila odorant receptor and initiate the HEK293 cell endogenous Gq-

protein cascade. Accordingly, we cotransfected HEK293 cells with constructs encoding

OR43a, OR83b and one of the five different chimaeric G-Proteins (Gαq16q, Gαq16i, Gαq16o,

Gαq16-73b, or Gαq16s) (Figure 4.2.21 A) designed to divert Drosophila Gαq, Gαi, Gαo,

Gα73b, or Gαs dependent signaling to the HEK293 cells calcium pathway (Conklin et al.,

1993;Conklin et al., 1996;Yapici et al., 2008). Calcium imaging experiments were

performed on transfected cells and the number of cells responding to odorant stimulation

was counted. The ratios of the responding cell number from the chimera G-protein

transfected cells to the responding cell number from the cells transfected without the

chimera G-protein were calculated. A higher value of the ratio indicated an increased

coupling efficiency with the transfection of the chimera G-protein. The experiments

revealed that coexpression of Gαq16s leads to the most effectice 7-fold increase in

response to the odorant stimulus (Figure 4.2.21 B). This experiment proves that in a

recombinant system the combination of OR43a/OR83b couples much better to Gαs than

to any other Gα-protein tested.

4.2.5.2 OR stimulation can induce an increase of GTP-binding to the membrane.

Agonist-stimulated [35S] guanylyl-5′-O-(γ-thio)-triphosphate ([35S] GTPγS) binding is

used to measure receptor activation of G-proteins in isolated membranes (Hilf et al.,

1989). The [35S] GTPγS assay is based on the fact that the inactive state of the G-protein

α subunit has a relatively high affinity for GDP over GTP, whereas activation of a

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receptor by its agonist shifts the α subunit into a higher affinity for GTP versus GDP.

Therefore, the [35S] GTPγS assay utilizes excess GDP to shift the G-proteins into the

inactive state and lower basal activity. Addition of agonist decreases the affinity of the α

subunit for GDP and increases its affinity for GTP, so that the receptor-stimulated G-

protein binds GTP. [35S] GTPγS is a hydrolysis-resistant form of GTP allowing to assess

the degree to which an agonist stimulates [35S] GTPγS binding in membranes. We

performed the assay on HEK293 cell membranes that were transfected with ORs and

Drosophila Gαs. Four to five transfections were carried out for each odorant receptor, and

the assay was run in triplicate for each transfection. OR83b was cotransfected with

conventional odorant receptors. Control cells were transfected with empty vector, and the

membranes were stimulated by a combination of the respective odorants. Particularly, 1-

hexanol, cyclohexanol and 1-octen-3-ol were mixed and used for OR43a, ethyl butyrate,

pentyl acetate and 1-octen-3-ol were mixed and used for OR22a (Hallem et al., 2004).

The counting result of the OR transfected sample was normalized to the empty vector

transfected sample. Student’s T-test analysis for the stimulation ratio between transfected

cell membranes and untransfected cell membranes revealed a significant difference,

indicating that activated receptors can stimulate Gαs and induce an increase in GTP

binding to the membrane (Figure 4.2.21 C).

Figure 4.2.21 Functional study of ORs and Gα proteins interactivity in the recombinant HEK293

expression system.

(A). Chimera construct with N-terminus of human Gα16 and the C-terminus of Drosophila Gα-proteins. (B). Ratio of transfected HEK293 cells responding to cyclohexanone in calcium imaging experiments; cells express OR43a, OR83b and the respective G-protein chimera or OR43a, OR83b and the full length human Gα16. An increase in the ratio means that more cells responded upon co-expression of the G-protein chimera. (C). [35S]GTPγS binding assay on transfected HEK293 cell membranes. Error bars represent S.E.M. (** p <0.01, n = 5-7)

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4.2.6 Overexpression of a GTPase deficient Gαs mutant in Drosophila antenna

leads to a change in response dynamics

Although the response amplitude of EAG recordings from the OR83b-GAL4; UAS-Gαs-

GTP flies showed no significant difference to the wt flies, we could clearly observe a

prolonged spike activity in the single sensillum recordings (Figure 4.2.22 A, B). By

counting the spikes in an interval of 200 ms, a significant prolongation of the response

period could be observed (Figure 4.2.22 C, D).

Figure 4.2.22 Overexpression of a GTPase deficient Gαs mutant in Drosophila antenna lead to a

change in response dynamics

(A) Single sensillum recording from wt fly. (B) Single sensillum recording from OR83b-GAL4; UAS-Gαs-GTP fly. (C, D) Spike distribution of the SSRs (n = 5-6). All the recordings are from the ab2 sensillum. Error bars represent S.E.M. (**: p < 0.01, *: 0.01 < p <0.05)

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4.2.7 Odorant exposure caused Gαs redistribution in antenna

The immunostaining of the antenna section shown in Figure 4.2.23 A reveals that Gαs is

expressed at the base of the sensillum but not in the dendrites of ORNs where olfactory

transduction gets initiated. If however Gαs is involved in olfactory signaling, what

explains its location at the base of the sensillum? We looked over our procedure of

immunostaining, and noticed that flies were always fixed immediately after taken out

from the food vials. We decided to repeate the experiment this time keeping flies in a 2%

agar vial for 2 hours before fixation. Interestingly, we observed that Gαs diffused into the

dendrite of ORNs in food odorant deprived flies (Figure 4.2.23 B).

For the purpose of monitoring this translocation process in the living flies, we generated a

GFP-Gαs fusion construct. Because of the importance of both the amino and carboxyl

termini for localization and function of the protein, we adapted a strategy to express a

functional expressed GFP-tagged Gαs (Hughes et al., 2001;Sunahara et al., 1997;Yu and

Rasenick, 2002). Specifically, the GFP sequence was inserted in-between the residues 71

and 72 of Gαs, a 6-residue linker sequence (SGGGGS) was inserted at both of the

junctions between Gαs and GFP (Figure 4.2.23 E, F). The fusion cDNA was first cloned

into the pCDNA3 vector, and the protein expression was tested in transfected HEK293

cells (Figure 4.2.23 G). The fusion cDNA was subsequently cloned into the pUAST

construct and transgenic flies were generated. Afterwards OR83b-GAL4; UAS-Gαs-GFP

flies were produced, and Gαs translocation was studied before and after odorant exposure.

Gαs was spread in the dendrites of the ORNs when the experiment fly was segregated

from food odors (Figure 4.2.23 C), and clustered at the base of the dendrites when the fly

was exposed to the odors (Figure 4.2.23 D).

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Figure 4.2.23 Odorant exposure caused Gαs redistribution in antenna.

(A, B) Immunostaining of Gαs on antenna section. A, + food odor. B, - food odor. (C, D) immunostaining of GFP on the antenna section. C, + food odor. D, - food odor. (E) Schematic representation of the GFP-Gαs fusion construct. GFP sequence was inserted in-between Gαs residues 71 and 72, a 6-residue linker sequence (SGGGGS) was inserted at both of the junctions between Gαs and GFP. (F) Model of Gαs-GFP. GFP is depicted as an insert into the Gαs structure. (G) Expression of GFP-Gαs fusion protein in HEK293 cells. Scale bar in B and G = 10 μm.

4.2.8 Generation of a Gαs knockout or knockdown fly

To further investigate the function of Gαs in odorant receptor neurons, we performed

experiments to generate Gαs knockdown or knockout flies. It is know that Gαs is essential

for the initial stages of larval development, the homozygote Gαs null mutant dgsB19

results in late embryonic-early larval lethality (Wolfgang et al., 2001). We examined the

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olfactory responses of the heterozygote dgsB19 by EAG and SSR and found no significant

phenotype in response amplitude and response kinetics. Therefore, we decided to

generate flies in which the knockdown or knockout was only targeted to the olfactory

sensory organ.

4.2.8.1 Usage of the UAS-RNAi fly line

RNA interference (RNAi) is a system within living cells that helps to control the

activation of genes; it is now a widely used tool for post-transcriptional gene silencing.

To target the RNAi expression in odorant receptor neurons, we used the OR83-GAL4 to

drive the UAS-RNAi expression. UAS-GαsRNAi was obtained from three different

sources: Vienna Drosophila RNAi center, NIG fly center and fly lines generated by our

own lab.

Performing EAG and SSR, we observed no obvious olfaction deficiency in all those flies.

We therefore tested RNAi efficiency by expressing them under a different driver, the elav

driver. Unfortunately, all RNAi lines have generated robust offspring. It seems that all

Gαs RNAi lines tested do not work the way they were designed, or at least, do not down

regulate the post Gαs transcription to a level that would affect either neuronal

development at embryonal stages or influence signal transduct in ORNs.

4.2.8.2 Generation of mosaic Gαs knockout flies

To avoid lethality in the early stage of embryonic development, we used a strategy based

on the FLP-induced mitotic recombination (Xu and Rubin, 1993) to generate genetic

mosaic flies that only have the homozygote dgsB13 cells in a part of their sensory organ.

FLP-induced somatic clones appear to result from a simple reciprocal recombination

event between FRT sites. A mitotic recombination event in a cell heterozygous for a

marker gene would produce one daughter cell with two copies of the marker and a sibling

cell with no copies (Figure 4.2.24). Each of these daughter cells will divide to give a

clone of cells in the adult (twin-spot clones). If neither cell is defective in proliferation or

differentiation, the twin-spot clones will be of similar size. In our experiment, a ey-FLP

construct (Jefferis et al., 2004;Sweeney et al., 2007) was used to induce the chromosome

recombination between the P[neoFRT]42D dgsB13 and the P[neoFRT]42D Ubi-

GPF(S56T). If the homozygote dgsB13 cells in the eye-antennal disc could still develop

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similar to wild type cells, the adult fly could have some Gαs empty clusters in the

antennae.

The flies were generated and the cryosection of the antenna was examined. Cells without

GFP expression were not observed, arguing that either recombination did not occur or

that Gαs is essential for antenna cell development. Since this ey-Flp construct was well

described in several studies (Hummel et al., 2003;Jefferis et al., 2004;Sweeney et al.,

2007), it is very likely that the Gαs plays a crucial role in the antennal neuron

development.

Figure 4.2.24 Schematic overview of genetic mosaic fly generation.

Homozygous mutant cells are identified as unstained cells if the marker gene is placed distal to the FRT site on the homologous chromosome arm in trans to the mutant gene.

4.3 Secondary messengers and other components in the ORNs signal

transduction cascade

4.3.1 The cAMP pathway is involved in odorant signaling in fly olfactory neurons

4.3.1.1 A cAMP level increase in ORNs causes a rise in neuronal activity

A photoactivated adenylate cyclase (PAC) was expressed in ORNs to manipulate the

cAMP level. This photoactivated adenylate cyclase was identified in the unicellular

flagellate Euglena gracilis (Iseki et al., 2002). It is composed of two PACα and two

PACβ subunits, which exhibit adenylate cyclase activity that is enhanced by blue light.

Each subunit harbors two BLUE-type photoreceptor domains, binding flavin adenine

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dinucleotide (Anderson et al., 2005;Gendre et al., 2004), and two catalytic domains that

are homologous to class III adenylate cyclase. Studies showed that in a heterologous

expression system, the PACα subunit exhibits a strong cAMP synthesis efficiency

(Schroder-Lang et al., 2007). Similar results were obtained from Xenopus oocytes, HEK

cells and transgenic Drosophila (Schroder-Lang et al., 2007). The same transgenic fly

was used in our experiment. By crossing the UAS-PACα responder line with the OR83b

driver line, flies expressing the photoactivated adenylate cyclase in odorant receptor

neurons were generated. During the whole fly culture, the animals were kept in the dark,

or shortly exposed to red light for the requirement of handling. During single sensillum

recording flies were alternately exposed to light stimulation and dark rest. The neuronal

activity increased upon light stimulation and turned back to normal in the dark (Figure

4.3.25 A C). Light stimulation was carried out for 10 seconds (Figure 4.3.25 A), 30

seconds (Figure 4.3.25 C), or even several minutes (similar data not shown). After

stimulation, neuronal activity always returned to basal level. The longest stimulation time

tested was 5 minutes. Spike numbers were counted for ab1, ab2 and ab3 sensilla

recordings. Except for the ab1C neurons, which could not be covered by the OR83b

driver, all other neurons analyzed showed a statistically significant increase in spiking

frequency (Figure 4.3.25 B).

Figure 4.3.25 cAMP level moderate the activity of ORNs

(A) Representative recording of an OR83b-PACα ab2 sensillum. The whole trace represents a 30 seconds recording. (B) Comparison of spike number between light and dark stimulations. Error bars represent S.D., n = 5-7. (C) Repetitive model of PACα-ORNs in response to light stimulation. Data were obtained from the ab2A neuron. Error bars represent S.D., n = 5-7.

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4.3.1.2 Activation of PAC recovers the spontaneous neuronal activity in OR83b

knockout ORNs.

To test whether an increase in cAMP levels would affect odorant receptor neurons of

OR83b knock-out flies (OR83b-/-), we generated a mutant fly expressing PACα in the

odorant receptor neurons on a OR83b knock-out background (Figure 4.3.26 A). Single

sensillum recordings from the ab1 sensillum of OR83b-/- flies have only one group of

spikes (Figure 4.3.27 A), representing the Gr21a neuron activity, instead of four from the

wt ab1 sensillum (Figure 4.3.26 C). When the PACα was expressed by the OR83b-GAL4

driver, and the fly antenna was exposed to blue light, additional neuronal activity could

be recorded (Figure 4.3.26 B). We sorted all spikes in a given time and analyzed the

spike amplitude distribution (Figure 4.3.26 D, E). 453 spikes were analyzed from 8

seconds of recording of an OR83b PACα (OR83b-/-) fly (Figure 4.3.26 E), 229 spikes

from wt fly were as well analyzed (Figure 4.3.26 D). We observed a similar spike

distribution pattern, indicating that PACα expression in olfactory receptor neurons does

efficiently recover spontaneous neuronal activity.

Figure 4.3.26 An increase in cAMP levels can recover the spontaneous activity of ORNs.

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(A) Crossing scheme of the OR83b-GAL4 / UAS-PACα; OR83b2 / OR83b2 fly. (B) Single sensillum recording from the OR83b-GAL4 / UAS-PACα; OR83b2 / OR83b2 fly with blue light illumination. The whole trace is 3 seconds in duration. (C) SSR recording from a wt fly exposed to blue light. The whole trace is 3 seconds in duration. (D) Spike amplitude distribution of the wt ab1 sensillum. (E) Spike amplitude distribution of the blue light illuminated OR83b-GAL4 / UAS-PACα; OR83b2 / OR83b2 ab1 sensillum.

Figure 4.3.27 SSR of the OR83b-GAL4 / UAS-PACα; OR83b2 / OR83b2 and control flies

This figure shows orginal single sensillum recordings of flies: (A) OR83b2 / OR83b2 (B) OR83b2 / OR83b2 with blue light (C) OR83b-GAL4 / UAS-PACα; OR83b2 / OR83b2 (D) OR83b-GAL4 / UAS-PACα; OR83b2 / OR83b2 with CO2 stimulation. (E) OR83b-GAL4 / UAS-PACα; OR83b2 / OR83b2 with odorant stimulation. (F) OR83b-GAL4 / UAS-PACα; OR83b2 / OR83b2 with blue light stimulation. (G) OR83b-GAL4 / UAS-PACα; OR83b2 / OR83b2 with blue light stimulation, then plus odorant stimulation. All of the traces are 3 seconds in duration.

4.3.2 Expression of activated Gαq in ORNs leads to resensitization deficiency.

In general, the continuously activated Gq3 functions as a dominant gain-of function allele

in a tissue and cell specific manner (Ratnaparkhi et al., 2002). The UAS-AcGq3; OR83b-

GAL4 fly was used in an EAG screen for olfactory deficient flies. As described in section

4.2.1, this fly showed no olfactory defect in EAG amplitude. However, while studying

desensitization, we observed a deficiency in resensitization of EAG responses.

Two sets of assays were performed to evalute desensitization. At first, short pulse odorant

stimuli (pure odorant) were applied every minuteand EAG response were recorded. In

wild type flies, EAG amplitudes of consecutive responses were unaltered, while EAG

responses of UAS-AcGq3;OR83b-GAL4 flies declined upon repetitive stimulation

eventually reaching a steady level (Figure 4.3.28 A). In a second previously decribed

assay (Stortkuhl et al., 1999), flies were pre-treated with a given odorant (pure odorant)

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for one minute followed by a short pulse odorant stimulation every minute (1:100

dilution). Normalizing EAG amplitudes to the none pre-treated condition, UAS-

AcGq3;OR83b-GAL4 flies appeared to exhibit a resensitization deficiency (Figure 4.3.28

B, C).

Figure 4.3.28 Expression of an AcGαq in ORNs leads to a resensitization deficiency

(A) The UAS-AcGq3;OR83b-GAL4 fly showed an abnormal desensitization to a continuous odorant stimulation. EAG amplitudes are normalized to the response of the first stimulation. (B, C) Ethyl acetate and benzaldehyde were used in B and C. The UAS-AcGq3; OR83b-GAL4 fly showed an abnormal resensitization after constant odorant stimulation. Error bars represent S.E.M. n = 5-7.

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4.3.3 Effect of MIA on Drosophila odorant receptors

MIA, an amiloride derivative, was reported as a lobster TRP channel blocker (Bobkov

and Ache, 2007) (Figure 4.3.29 D). As demonstrated by Dr. Günter Gisselmann, MIA

can block odorant-induced currents in oocyte expressing OR47a and OR83b (Figure

4.3.29 A-C). To further investigate whether this substance works on the fly as well, we

applied the substance on the surface of the antenna and measured olfactory responses.

Different approaches of substance application were tested, and finally we chose to mount

the antenna in a micropipet tip filled with 1 mM MIA solution. 20 minutes after MIA

mounting, a filter paper fiber was used to wipe up the antenna surface. The fly was

subsequently exposed to a constant airflow to allow the antenna surface to dry. After 20

minutes, the fly was subjected to single sensillum recording. Since the antenna surface is

covered by cuticle, the efficiency of substance delivery by using this approach is unclear.

From all single sensillum recordings, only a few sensilla show a phenotype in olfactory

response. About one tenth of the recorded sensilla showed a lack of spontaneous activity

and a reduction of odorant responses (Figure 4.3.30). On one hand, we observed spike

traces from the sensilla missing only one group of neuronal activity, on the other hand, a

sensillum which losing the neuronal spontaneous activity of all neurons is hard to access

by the recording electrode in the SSR. As a control, the solvent of MIA (DMSO) was also

applied directly on the antenna, but no significant olfactory response alteration could be

observed.

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Figure 4.3.29 MIA blocks odorant-induced currents in oocytes

(A) Pentyl acetate induced currents could be observed in oocytes expressing OR47a and OR83b. (B) MIA blocked the pentyl acetate induced current in oocytes. (C) Dose-inhibition curve showing the blocking efficiency of MIA. (D) The chemical structure of MIA (5-(N-methyl-Nisobutyl) amiloride). Error bars represent S.E.M. These data were produced by Dr. Günter Gisselmann (shown with permission).

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Figure 4.3.30 Direct application of MIA on antenna blocks the spontaneous neuronal activity

(A, C) MIA treated antenna, ab2 sensillum, the spontaneous activity of the ab2A neuron was gone after MIA treatment. Although the ab2A neuron still respond to odorant stimulation (1:100 diluted ethyl acetate and 1:10000 diluted ethyl butyrate), the response is largely decreased. (B, D) 1% DMSO treated antenna, DMSO was used as the solvent for MIA.

4.3.4 Functional study of Ih channels in Drosophila odorant receptor neurons

Ih (HCN, Hyperpolarization-activated, cyclic-nucleotide-gated ion channel) channels are

widely expressed in the mammalian heart and brain (Brown et al., 1979;Halliwell and

Adams, 1982). They are responsible for a variety of physiological functions ascribed to

the depolarizing cation current (Ih) in the cellular context. In physiological context Ih

channels are involved in control of pacemaker activity in both heart and brain

(DiFrancesco, 1993;Pape and McCormick, 1989), determination of the resting membrane

potential (Williams and Stuart, 2000), control of membrane resistance and synaptic

integration in dendrites (Magee, 1999) and primary sensory transduction (Stevens et al.,

2001). Ih channels have also been cloned form arthropod, for example Drosophila (Marx

et al., 1999), Apis mellifera (Gisselmann et al., 2003;Gisselmann et al., 2004), Heliothis

(Krieger et al., 1999), and Panulirus (Gisselmann et al., 2005b). The sequence of these

channels exhibits homology to both CNG channels (Finn et al., 1996;Santoro et al., 1997)

and voltage-gated potassium channels (Catterall, 1995). In addition, mRNA of Ih-channel

was detected in the antennae of different insects and lobster (Gisselmann et al.,

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2005a;Gisselmann et al., 2005b;Krieger et al., 1999;Marx et al., 1999;Roeper et al.,

1998), suggesting a possible role of this channel in olfactory signal transduction.

Two mutant fly lines have been used in our study: UAS-DoIh was generated by Martina

Köper as previously described (Köper M, 2008), and Ih-/- mutant was obtained from

Inmaculada Canal (Universidad Autonoma de Madrid, Spain). The Ih-/- fly was measured

directly. The dominant negative Ih construct encoding a truncated protein was expressed

by OR83b-GAL4. Single sensillum recordings from both mutants has been performed,

and the recording results were analyzed by counting the spikes in an interval of 200 ms.

Both of the mutants showed a response dynamic change compared to control flies. The

odorant-induced firing rate exhibited a faster decline in Ih-/- flies as compared to control

flies (Figure 4.3.31 A). The UAS-DoIh; OR83b-GAL4 ORNs showed a spontaneous

neuronal activity increase, and a prolonged activity after odorant stimulation (Figure

4.3.31 B).

Figure 4.3.31 Single sensillum recording analysis from Ih channel mutants

(A) Single sensillum recordings on Ih channel knockout flies. Odorant: ethyl butyrate, 1:10000 diluted. Odorant application lasted for 1 second. The sensillum measured here is ab3. The response of neuron A is shown here. (B) Single sensillum recordings on UAS-DoIh, OR83b-GAL4 ab2 sensillum. Odorant: ethyl acetate, 1:100 diluted. Odorant application lasted for 1 second. The response of neuron A was analyzed. Error bars represent S.E.M. n = 5-7. **: p < 0.01, *: 0.01 < p < 0.05.

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4.4 Larva study

4.4.1 Gαs and Gαq are expressed in larva olfactory receptor neurons

To investigate whether G-proteins are also involved in the larval olfactory system, we

carried out similar experiments in larvae as we did in adult flies. First, we checked the

expression of Gαs and Gαq in larval olfactory receptor neurons. OR83b-GAL4; UAS-

mCD8-GFP larvae were used for the whole mount and cryosection staining. As described

earlier in section 1.1.1, the OR83b expression cells are the only ORNs in larvae.

Therefore, OR83b-GAL4 driver can be used to mark the ORNs in larvae as well as in the

adult olfactory organs. Immunohistochemical stainings revealed that both Gαs and Gαq

are expressed in the cell body of larval odorant receptor neurons. The cilia of the larval

odorant receptor neurons are housed in the dorsal organ, as shown in Figure 4.4.32 A.

The dorsal organ exhibits strong autofluorescence, which makes the detection of the Gαs

and Gαq almost impossible.

Figure 4.4.32 Immunostaining of larval ORNs

(A) Whole mount staining of the OR83b-GAL4; UAS-mCD8-GFP larval olfactory sensory neurons. Gαs is expressed in the cell bodies of the ORNs. Since dorsal organs (DO) have a strong autofluorescence, Gαs is expression in the dendritic area cannot be assessed. (B, C) Cryosection of the larva (cross section of the head tip). Both Gαs and Gαq are expressed in the cell body of the ORNs. Scale bar = 20 μm.

Results

66

4.4.2 CTX larvae have normal chemotaxis behavior

Larval chemotaxis assays were performed with OR83b-CTX larvae, which have the same

genotype as the CTX adult described before (Chapter 4.2.2). It turned out that OR83b-

CTX larvae behave completely normal like wt larvae in the chemotaxis assay (Figure

4.4.33). It has already been shown that OR83b neurons are essential for the larval

chemosensation, and OR83b knock out larvae are anosmic to a wide range of fly odors

(Larsson et al., 2004). Our result showed that larval ORNs expressing CTX function

normally, which substantiate that the signal transduction pathways in larvae olfactory

receptor neurons is Gαs independent.

4.4.3 Expression of AcGq3 in larval olfactory receptor neurons causes olfactory

aberration

For the expression of AcGq3, females of the genotype UAS-AcGq3/FM7-GFP were

crossed to homozygous males of OR83b-GAL4. The non-GFP expressing larvae were

picked up manually under the fluorescence microscope and used for the larval

chemotaxis assay. Ethyl acetate, cyclohexanol and benzaldehyde were applied as odorous

stimuli.

As indicated in Figure 4.4.33, AcGq larvae have a reduced response to ethyl acetate as

well as a high concentration of cyclohexanol. To benzaldehyde and low concentration of

cyclohexanol behavior responses remained unaltered.

Figure 4.4.33 Chemotaxis assays on AcGq and CTX larvae

(A, B, C) Ethyl acetate (A), cyclohexanol (B) and benzaldehyde (C) are used for the test. OR83b-GAL4; UAS -CTX larvae have a normal chemotaxis behavior. UAS-AcGq3; OR83b-GAL4 larvae showed a response decrease to ethyl acetate and high concentration of cyclohexanol. Error bars represent S.E.M.

Discussion

67

5 Discussion

5.1 The role of Gαs in the olfactory signal transduction pathway

In recent years, it has been realized that Drosophila ORs are not classical GPCRs since

these proteins neither show a homology to conventional GPCRs nor have the

extracellular N- terminus and intracellular C- terminus membrane topology (Benton,

2006;Clyne et al., 1999;Gao and Chess, 1999;Lundin et al., 2007;Vosshall et al., 1999).

Moreover, two studies published last year provided evidence that heterologously

expressed Drosophila ORs have an ion channel capacity (Sato et al., 2008;Wicher et al.,

2008). These studies raise the question of whether heterotrimeric G-proteins are involved

in the signal transduction pathway of odorant receptor neurons. In the present study, we

investigated whether Drosophila ORs couple to G-proteins to trigger downstream signal

transduction events.

Previous studies provide plenty of evidence indicating that G-protein signaling is

involved in invertebrate olfactory signal transduction. Two G-protein signaling

transduction pathways depending on the second messengers 1,4,5-inositol triphosphate

and cAMP, respectively, appeared to be involved in olfactory perception (Breer,

1994;Ronnett and Moon, 2002;Stengl et al., 1992a). The existence of both signaling

cascades has been proven by using different approaches: at the molecular level, by

expression of genes encoding for intermediate products of both signal transduction

pathways in ORNs (Baumann et al., 1994;Dubin et al., 1998a;Hasan and Rosbash,

1992;Martin et al., 2001;Marx et al., 1999;Riesgo-Escovar et al., 1997;Yoshikawa et al.,

1992) and at the cellular level by electrophysiological measurements of vertebrate and

invertebrate ORNs (Fadool and Ache, 1994;Hatt and Ache, 1994;Martin et al.,

2001;McClintock et al., 1997). In our study, convincing evidence is provided to

substantiate the involvement of Gαs-mediated cAMP signaling in Drosophila olfactory

signal transduction.

Discussion

68

First of all, the expression levels and expression patterns of different G-proteins in the

antennae of Drosophila were investigated. Two proteins of interest were specified due to

their expression pattern in ORNs. The most suspicious goes to Gαs, which is expressed in

the dendrites of odorant receptor neurons, where the olfactory signal transduction cascade

is initiated. Further investigation on Gαs localization revealed that food odor exposure

causes Gαs protein translocation from the end of the dendrite to the base of the dendrite.

This redistribution is similar to the light induced transducin, the eye specific Gαt,

translocation in rod photoreceptors (Philp et al., 1987;Whelan and McGinnis, 1988),

suggesting that Gαs might serve a similar function in ORNs as transducin in rod

photoreceptors. Gαq was found to be expressed in some of the ORNs. Quantitative PCR

also showed that Gαq was highly expressed in the antennae compared to brain tissue, but

the expression pattern of Gαq in antennae showed that Gαq protein is localized in the cell

body of ORNs but not in the dendrite. Although a recent publication revealed that a

mutation in the Gαq gene can cause a reduction in amplitude and spiking frequency,

respectively, in EAG and single sensillum recordings (Kain et al., 2008), it is not likely

that Gαq directly couples to ORs as the protein is not located in the cellular location

where the receptor and the downstream factor should interact. Also, it is important to

notice that although homozygote Gαq null mutant neurons were generated, no anosmic

phenotype has been found. Precisely how is involved in OR signal transduction interacts

requires further investigation.

Secondly, a functional screening of distinct G-proteins as to be involved in olfactory

transduction was performed in both the fly antennae as well as heterologous expression

system. In Drosophila, different wt G-proteins, mutated G-proteins, and G-protein

affecting toxins were expressed in the ORNs. Only CTX flies showed a strong reduction

of odorant-induced signals in EAG recordings. Further investigation by single sensillum

recordings on CTX flies showed that odorant-induced spikes in CTX expressing neurons

are completely gone. Since CTX is an ADP-ribosyltransferase that typically activates Gαs

proteins (Moss and Vaughan, 1988), the strong olfactory deficiency of CTX expressing

flies indicated that Gαs, the only stimulative type of G-proteins in Drosophila, is crucially

important for odorant induced signal transduction in olfactory sensory neurons. Although

Discussion

69

other G-protein over-expressing flies exhibited no change in EAG amplitude, olfactory

response aberration could be demonstrated by other experimental procedures. For

example, single sensillum recordings of Gαs-GTP flies, which express a mutated Gαs-

protein with decreased GTPase activity and therefore can be regarded as constitutively

active, revealed a significant prolongation of spike activity after odorant stimulation as

compared to control flies. The absence of phenotype in EAG recordings might be

reflected by the fact that the initial increase in neuronal firing was not significantly

different in these flies. Changes were observed in the duration of the period of increased

firing in the mutant flies, which may be caused by the deficiency in GTPase activity of

the activated G-protein subunit.

For the screening of odorant receptor coupled G-proteins in the heterologous expression

system, we used the G-protein chimera strategy to switch the Drosophila odorant

receptor signals to the endogenous calcium signal in HEK293 cells (Conklin et al.,

1993;Conklin et al., 1996;Mody et al., 2000). Using calcium imaging, HEK293 cells

cotransfected with OR and Gαq16s showed stronger odorant-induced signals as compared

to other chimeric G-proteins (Gαq16q, Gαq16i, Gαq16o and Gαq16-73b). The increased

coupling efficiency emphasizes that Gαs serves as downstream G-protein of ORs.

Conventional odorant receptors were cotransfected with OR83b to increase the

expression efficiency (Neuhaus et al., 2005), albeit the conventional odorant receptor

alone expressed in HEK cells or Xenopus oocytes responds to odorant stimulation

(Neuhaus et al., 2005;Wetzel et al., 2001).

To further examine our hypothesis that Drosophila odorant receptors are coupling to Gαs,

the classical [35S]GTPγS binding assay was carried out to test whether activation of ORs

could increase GTP binding to the G-protein. We coexpressed Gαs and odorant receptors

in HEK293 cells. As expected, the binding assay showed a significant increase of

[35S]GTPγS binding in OR transfected cell membranes, indicating the odorant-treated

Drosophila ORs can interact with Gαs. Although further speculations have to await

investigations on the mode of receptor G-protein coupling in the olfactory system, it is

tempting to speculate that Drosophila ORs couple to heterotrimeric G-proteins although

Discussion

70

bearing an inverse membrane topology compared to other GPCRs. In fact, there are other

reversed GPCR like receptors that show a possibility of coupling to G-proteins, for

example the human adiponectin receptors (AdipoRs). Human adiponectin receptors and

membrane progestin receptors (mPRs) belong to the PAQR (Progestin, AdipoQ-

Receptor) family of proteins, which are seven transmembrane receptors of a novel type,

that similar to Drosophila OR83b share little sequence homology with other GPCRs

(Yamauchi et al., 2003;Zhu et al., 2003). As Drosophila OR83b (Benton et al., 2006),

AdipoRs have been shown to have intracellular N- and extracellular C-termini (Deckert

et al., 2006;Yamauchi et al., 2003), but location of the termini of mPR has yet to be

confirmed. Nevertheless, the fish mPR has been shown to be a plasma membrane protein

whose activation leads to inhibition of adenylate cyclase in a pertussis toxin-sensitive

manner, consistent with mPR being a novel type of GPCR (Zhu et al., 2003).

Gαs typically activates adenylate cyclases to produce cAMP. To test whether odorant

stimulation could induce a cAMP increase in Drosophila antennae, a cAMP assay was

performed. Fly antennae were cut manually and the PerkinElmer alpha screen kit was

used for the assay. Most of the time no significant cAMP increase could be observed.

This may either be due to the fact that a too small amount of tissue was used for each

assay, or be due to the nature of the odorant receptor neurons in Drosophila, in a way that

certain odorants can cause excitation in some ORNs and inhibition in other ORNs

(deBruyne et al., 2001;Hallem et al., 2004). If an odorant can induce both excitation and

inhibition at the same time in the antenna, this may lead to an unchanged cAMP level

before and after odorant stimulation. Even though no cAMP increase could be observed

after odorant application, a cAMP increase in odorant receptor neurons can certainly

cause neuronal excitement.

Expression of a light-activated adenylate cyclase (PACα) (Schroder-Lang et al., 2007) in

olfactory neurons enabled us to show that a cAMP increase in these cells results in

increased firing rates, providing hints for the existence of a cAMP dependent excitatory

signaling pathway. Also, overexpression of a cAMP-phosphodiesterase in olfactory

neurons was indicative for an excitatory role of cAMP in the Drosophila olfactory system

Discussion

71

(Gomez-Diaz et al., 2004). Moreover, cyclic nucleotide–gated channels and Ih channels

as potential cAMP targets are known to be expressed in the antennae of Drosophila

(Baumann et al., 1994;Gisselmann et al., 2005a;Marx et al., 1999) and mutants of a cyclic

nucleotide-modulated potassium channel show olfactory deficits (Dubin et al., 1998a). To

further understand the function of cAMP in the olfactory signal transduction pathway, the

light-activated adenylate cyclase was expressed in the OR83b-/- odorant receptor neurons.

By lightening up the antenna, neuronal activity can be recorded by single sensillum

recording. After grouping the spikes by their amplitude, similar spontaneous activity

patterns can be observed. From previous publications (Dobritsa et al., 2003;Larsson et al.,

2004;Neuhaus et al., 2005) and our own recording data, it is clear that the ORNs of

OR83b-/- flies show a complete lack of spontaneous activity, but the Δhalo fly, which

have no functional OR expressed in the ab3a neuron, still maintains a spontaneous

neuronal activity of the ab3a neuron. Our data suggest that the spontaneous activity is

largely dependent on the cellular level of cAMP.

As the gain of function of Gαs has already been investigated, studying the loss of function

of Gαs became critical. It is known that the complete knockout of Gαs in the entire animal

causes early embryo lethality (Wolfgang et al., 2001). Two strategies were used for

generating the olfactory organ targeted Gαs knockdown or knockout flies. One is based

on the GAL4-UAS RNAi method; the other one is based on the genetic mosaic fly

generation. Unfortunately, none of these efforts achieved the predicted effect. For the

RNAi method, three sources of UAS-Gαs-RNAi were used, but none of them could induce

an efficient post-transcriptional gene silencing. The recordings from those flies also did

not show any obvious olfactory deficiency. For the mosaic fly, the prerequisite of

successfully generating such a fly is that the homozygote mutant cells are healthy enough

to generate offspring cells. In our case, it seems that the homozygote dgsB13 cells in the

embryo stage cannot survive through the development. No homozygote dgsB13 cells could

be observed in cryosections of the antenna. Since the driver line used (ey-Flp) was well

described and examined in several studies (Hummel et al., 2003;Jefferis et al.,

2004;Sweeney et al., 2007), it is very likely that the Gαs plays a crucial role in antennal

neuron development.

Discussion

72

Putting aside the Gαs coupling, the odorant receptor ion channel hypothesis raises a series

of new questions. First, there is no clear consensus on the position where the pore of the

channel is located and to what extent different subunits in the OR complex contribute to

the pore itself. Only a weak similarity was found between the suspected odorant receptor

pore motif and the well-characterized potassium channel pores (Wicher et al., 2008).

Second, there is little data on the exact stoichiometry of the OR complex. Although it was

showed by Benton et al. that at least two subunits each of OR83b and the conventional

odorant detecting OR are included in the receptor complex (Benton et al., 2006), the

composition of the functional complex is still unknown and it might even vary for

different OR83b/OR combinations. Third, if Drosophila odorant receptors form a

heteromeric ligand-gated ion channel, and raise action potentials immediately after

odorant binding, the ablation of any one of the channel subunits should lead to a similar

phenotype. The fact is that the OR83b knockout fly results in a lack of spontaneous

action potential firing of the ORNs (Larsson et al., 2004), while conventional OR knock

out ORNs still generate spontaneous action potentials (Dobritsa et al., 2003). Of course

none of these ORNs respond to odorants any more. In addition, it is still not well

understood whether classical GPCRs form dimers or higher ordered oligomers, and the

general functional significance of this polymerization or dimerization is not clear.

Vertebrate class C GPCRs, such as metabotropic glutamate receptors and γ-aminobutyric

acid type B receptors clearly form homo- and heterodimeric structures, essential for both

trafficking of receptors to the cell surface and G-protein coupling (Pin et al., 2005). The

relevance of the monomeric or dimeric state for G-protein activation for other GPCRs is

currently under debate. For example, NTS1, a dimerizing class A receptor, was recently

shown to alter the mode of the receptor G-protein interaction (White et al., 2007). The

fact that two distinct families of seven transmembrane domain receptors, namely

vertebrate ORs as classical GPCRs and Drosophila ORs seem to make use of similar

intracellular signaling cascades could demonstrate an interesting case of convergent

evolution (Benton, 2006). It moreover demonstrates that olfactory signaling pathways

seem to be principally conserved between species.

Discussion

73

5.2 Larval olfactory system

We have demonstrated the possible role of Gαs in the adult olfactory system. Since the

larval olfactory neurons also coexpress conventional odorant receptors and OR83b, it is

of great interest to understand if their olfactory signal transduction pathways are similar.

For this purpose, we examined all those different wt G-proteins, mutated G-proteins, and

G-protein affecting toxins expressing lines using the larval chemotaxis assay.

Surprisingly, the CTX expressing larvae show normal chemotaxis behavior. It has

already been proven that OR83b neurons in the Drosophila larvae dorsal organ ganglia

are the only essential odorant receptor neurons that are responsible for larval olfaction

(Fishilevich et al., 2005;Heimbeck et al., 1999;Larsson et al., 2004), suggesting that Gαs

is not required for the larval olfactory signal transduction pathway. The OR83b-GAL4;

UAS-AcGq larvae showed a reduced sensitivity to some of the odorants that were used,

indicating that Gαq may be important for the larval olfactory signal transduction.

Immunostaining of larval olfactory organ was performed with Gαs and Gαq antibodies on

OR83b-GAL4; UAS-mCD8-GFP larvae. Adults of the same fly line were used for the

immunostaining of the antennae. The immunohistochemistry showed that both Gαs and

Gαq are expressed in the cell body of larvae ORNs. The dendrites of the ORNs in larvae

are housed in the dorsal organ, which exhibits a strong autofluorescence, making it very

difficult by immunostaining to detect whether the Gαs and Gαq are expressed in the

dendrites.

It is known that olfactory organs of adults and larvae have different developmental

origins. During metamorphosis, the larval dorsal organ is histolysed (Stocker, 1994), and

the antennae and palps develop de novo from the eye-antenna imaginal disc (Postlethwait

and Schneiderman, 1971). Larvae and adults inhabit in different phases. The demands of

olfaction are also different: for the larvae, the major purpose is foraging; while for the

adult, foraging, courtship and oviposition become equally important. It seems likely that

Drosophila larvae and adult depend on different signal transduction pathways.

Discussion

74

5.3 Other signaling components in the olfactory signal transduction

pathway

Despite the recent discovery of the ion channel capacity of Drosophila odorant receptors,

it has long been assumed that the canonical model of olfactory signal transduction would

also hold true in insects. Indeed, most of the molecules involved in mammalian OR or

GPCR signal transduction cascade have been identified in insect, including OBPs (Pelosi

and Maida, 1995;Tegoni et al., 2004), arrestins (Merrill et al., 2002;Merrill et al.,

2003;Merrill et al., 2005), AC (Gomez-Diaz et al., 2004;Martin et al., 2001), CNG

(Baumann et al., 1994;Dubin et al., 1998b;Marx et al., 1999), and IP3 gated ion channel

(Stengl, 1994). In the first part of this chapter, we have already discussed the

participation of cAMP in the transduction. Therefore, we will only discuss the function of

Gαq, Ih channels, and the amiloride derivative MIA as OR83b blocker in this section.

Although Gαq3 is not expressed in the cilia of odorant receptor neurons, and over-

expression of the continuously activated Gαq3 in the antenna does not induce any

olfactory response aberrations in the single pulse EAG measurement, the UAS-

AcGq3;OR83b-GAL4 fly did show a phenotype in the desensitization test. Two sets of

tests were performed for the desensitization evaluation by the EAG recordings. The

difference between these two assays is whether the fly had a pretreatment of

desensitization. With or without pre-exposure, the short pulse odorant stimulations were

applied every minute, and the amplitudes of the olfactory response were recorded. The

test without odorant pre-exposure showed that at the frequency of one stimulation per

minute, ethyl acetate, benzaldehyde and cyclohexanol all evoked a response

desensitization in the UAS-AcGq3; OR83b-GAL4 fly. While in the wt fly, the same pattern

of repetitive stimulation did not induce such phenotype. In the odorant pre-exposure test,

similar results could be observed, compared to the wt fly, the AcGq3 fly showed a

deficiency in resensitization. In the Drosophila visual system, it was described that a

prolonged Gαq activity could trigger the rhodopsin endocytosis and degradation, thus

resulting in a reduced photoreceptor sensitivity (Han et al., 2007). Albeit the Drosophila

odorant receptor are not likely coupled to the Gαq as the light receptor Rh1, similar

receptor endocytosis and degradation could be induced by the continuously activated Gαq

Discussion

75

and lead to this resensitization defect. The same UAS-AcGq3 construct used by us was

expressed under the hs-GAL4 driver by Kain and colleagues, since the activated Gαq can

cause an intercellular calcium level increase and probably lead to up or down regulation

of other genes. This strategy of conditional expression has the advantage of avoiding this

complication in the development of odorant receptor neurons. The EAG recording from

these heat-shocked mutant flies showed a mildly reduced or similar amplitude compared

to the control flies (Kain et al., 2008). This observation further supports the idea that Gαq

is not the direct downstream effector of the odorant receptor. A lack of odorant induced

EAG amplitude aberration and the phenotype in the desensitization assays indicate that

Gαq is not essential for the depolarization of odorant receptor neurons, but somehow

crosstalks with the Gαs signaling cascade to regulate the olfactory transduction pathway.

The presence of both transduction system depending on Gαq and Gαs in the same

olfactory receptor neurons has been suggested using pharmacological approaches in

vertebrates (Noe and Breer, 1998), the crosstalk between both systems had also been

reported (Vogl et al., 2000). All these data reinforced previous hypotheses of a functional

meaning of this coexistence and of the role of the olfactory receptor cell as primary

complex integrating unit in the olfactory system (Ache, 1994).

It is clear that cyclic nucleotide-gated ion channels serve as downstream targets of

signaling pathways in vertebrate olfactory sensory neurons. Since the first identification

of CNG channes in Drosophila, it has been proposed that CNG channels are involved in

signal transduction cascade of invertebrate olfactory neurons (Baumann et al., 1994). The

CNG channels we discussed here belong to a heterogeneous gene superfamily of ion

channels that share a common transmembrane topology and pore structure and that

harbor in their C-terminal region a binding domain for nucleoside 3’,5’-cyclic

monophosphates, Ih channels are also members of this superfamily. CNG channels form

heterotetrameric complexes consisting of two or three different types of subunits. Six

genes encording CNG channels have been found in human genome (four A subunits, A1

to A4, and 2 B subunits, B1 and B3), while the Drosophila genome contains 4 CNG

channel genes (dmA, dmB, dm3, dm4) (Kaupp and Seifert, 2002). Although CNG

Discussion

76

channels are widely expressed in the Drosophila olfactory tissue, their function is still

largely unknown.

Like CNG channels, Ih channels (HCN, hyperpolarization-activated, cyclic-nucleotide-

gated ion channel) have also been detected in the antennae of different insect and lobster

(Gisselmann et al., 2005a;Gisselmann et al., 2005b;Krieger et al., 1999;Marx et al.,

1999;Roeper et al., 1998). Signal sensillum recording from the Ih channel knockout fly

showed a response dynamic change compared to the control fly. The firing rate of the

odorant stimulated ORNs showed a relatively faster decline in the Ih channel knockout fly

than in the control fly. In addition to the knockout fly, a truncated Ih channel mutant

construct has also been generated, and this truncated Ih channel protein was supposed to

act as a dominant negative mutant when expressed in the flies. The UAS-DoIh, OR83b-

GAL4 ORNs showed a spontaneous neuron activity increase. Both of these mutant fly

lines had no obvious EAG amplitude variation in response to the odorant stimulation. The

response dynamic change in the Ih channel knockout fly is in accordance with the

phenotype of the continuously activated Gαs overexpression fly, which showed a

prolonged activation in the same type of measurement, indicating that Ih channel is one of

the downstream effectors of the cAMP in Drosophila ORNs.

Since the molecular mechanism of insect repellent DEET has been published (Ditzen et

al., 2008), our lab started a screening of a series of chemicals as potential novel insect

odorant repellent on the oocyte recombinant system. To achieve a high-throughput

screening, the insect odorant receptor and OR83b or OR83b homologue have been

expressed in the oocyte. Odorant was applied to maintain the current; the chemicals were

then applied to check if they could block the odorant-induced current. By this procedure,

an amiloride derivative, MIA, was identified as a novel odorant receptor blocker. This

substance was published before as a lobster TRP channel blocker (Bobkov and Ache,

2007). To understand if MIA worked on the natural system as well, it was applied

directly on the antenna, and followed by the single sensillum recording. In the single

sensillum recordings, only a part of the sensilla showed a missing spontaneous activity

and a reduction in odorant evoked spike frequency. This relative low number of affected

Discussion

77

sensilla observed in the living animal indicated the low delivery efficiency of the MIA

substance through the antennae cuticle, or, there are molecules other than conventional

OR and OR83b essential for the ORNs to generate a depolarization.

Conclusion

78

6 Conclusion

6.1 Summary

Drosophila melanogaster has 60 odorant receptor (OR) genes, which are expressed in the

third antenna segments as well as in the maxillary palps. The receptor proteins have well

understood odor response profiles. A given odorant receptor gene is expressed only in a

small fraction of olfactory neurons and each neuron expresses a very small number of

odorant receptor genes. This arrangement is similar to the mammalian olfactory system.

On a cellular level, olfactory signal transduction starts with the activation of olfactory

receptors, which are known to recognize a wide range of structurally highly variable

substances. While vertebrate olfactory receptors are 7-transmembrane proteins, which

activate heterotrimeric G-proteins after ligand binding, olfactory receptor proteins in

Drosophila were reported to have an inverse membrane topology compared to classical

G-protein-coupled receptors, presenting an intracellular N- and an extracellular C-

terminus. Moreover, it was shown recently that the Drosophila odorant receptors could

function as ligand gated ion channels. Controversial findings were reported concerning

the additional involvement of heterotrimeric G-proteins in olfactory receptor signaling.

One arising question is therefore, whether these 7-transmembrane receptors also couple

to heterotrimeric G-proteins, in addition to the reported novel direct activation. Our data

involving in vivo pharmacological studies, electrophysiological recordings and protein

redistribution analysis, as well as investigations using recombinantly expressed olfactory

receptors, now demonstrate that odorant receptor signaling in Drosophila indeed involves

G-proteins for signal transduction. Moreover, our results provide compelling evidence

that the stimulatory Gαs protein is involved in the olfactory signaling cascade of odorant

receptor neurons. In conformity with Gαs signaling we could show that increased cAMP

levels lead to excitation of olfactory sensory neurons. Furthermore, the manipulation of

cellular cAMP level can lead to a spontaneous neuronal activity restoration in the OR83b

knockout fly, which shows no spontaneous activity of ORNs, indicating that OR83b

might play a role in cAMP-dependent modulation. Results on the Ih channel knockout fly

let us suggest that Ih channels serve as one of the downstream effectors of cAMP.

Conclusion

79

A previous study on Gαq, PLC and DAG mutant flies also revealed a possible

involvement of the IP3 pathway in insect olfactory transduction. In the present study,

Gαq3 was found highly expressed in Drosophila antennae, but not in the cilia of olfactory

cells. Expression of a continuously activated Gαq3 in the olfactory neurons led to a

resensitization deficiency. These results suggest that Gαq3 is not the downstream target of

the odorant receptor, but might play a role in the receptor endocytosis and degradation or

for a crosstalk with the Gαs signaling cascade to regulate the olfactory signal

transduction.

A preliminary study showed that in larvae olfaction a different cellular signaling pathway

might be engaged with a participation of Gαq but not Gαs.

Conclusion

80

6.2 Zusammenfassung

Drosophila melanogaster hat 60 Geruchsrezeptorgene, die in den dritten antennalen

Segementen sowie den Maxillarpalpen exprimiert werden. Die Rezeptorproteine weisen

ein gut charakterisiertes Antwortprofil für Geruchsstoffe auf. Ein bestimmtes

Geruchsrezeptorgen wird nur in einer kleinen Population von olfaktorischen Neuronen

exprimiert. Jedes Neuron exprimiert nur eine sehr kleine Anzahl an

Geruchsrezeptorgenen vergleichbar mit dem olfaktorischen System von Säugetieren.

Auf zellulärer Ebene beginnt die olfaktorische Signaltransduktion mit der Aktivierung

von olfaktorischen Rezeptoren, die in der Lage sind, ein breites Spektrum von strukturell

unterschiedlichen Substanzen zu erkennen. Während die olfaktorischen Rezeptoren der

Vertebraten 7-Transmembranproteine sind, die nach der Bindung von Liganden

heterotrimere G-Proteine aktivieren, wurde für die olfaktorischen Rezeptoren von

Drosophila eine inverse Membrantopologie im Vergleich zu den klassischen G-Protein

gekoppelten Rezeptoren festgestellt, mit einem intrazellulären N- und einem

extrazellulären C-Terminus. Darüber hinaus wurde kürzlich gezeigt, dass die

olfaktorischen Rezeptoren von Drosophila als ligandengesteuerte Ionenkanäle

funktionieren können. Ergebnisse anderer Arbeitsgruppen lassen jedoch auf eine

zusätzliche Beteiligung von heterotrimeren G-Proteinen in der Geruchsrezeptor-

abhängigen Signalweiterleitung schließen. Es stellt sich daher die Frage, ob diese 7-

Transmembranrezeptoren ungeachtet der Tatsache, dass diese direkt aktiviert werden

können, auch an heterotrimere G-Proteine koppeln. Die von uns erzielten Ergebnisse aus

pharmakologischen in vivo Studien, elektrophysiologischen Messungen, Protein-

Umverteilungsanalysen sowie Untersuchungen an rekombinant exprimierten

olfaktorischen Rezeptoren zeigen, dass für die Geruchsrezeptor-abhängige

Signaltransduktion in Drosophila tatsächlich G-Proteine benötigt werden. Unsere Daten

erbringen klare Beweise für eine Beteiligung des stimulatorischen Gαs-Proteins in der

Signalkaskade von olfaktorischen Rezeptorneuronen. Passend zu einer Gαs-abhängigen

Signalkaskade konnte gezeigt werden, dass ein erhöhtes Niveau von cAMP zu einer

Erregung von olfaktorischen Neuronen führt. Weiterhin bewirkte die Manipulation des

zellulären cAMP Niveaus eine Wiederherstellung der spontanen neuronalen Aktivität in

Conclusion

81

OR83b-knockout-Fliegen, die prinzipiell keine spontane Aktivität von olfaktorischen

Rezeptorneuronen zeigen, was möglicherweise auf eine Rolle von OR83b in der cAMP-

abhängigen Modulation schließen lässt. Die Messungen an Ih Kanal knockout-Fliegen

lassen vermuten, dass Ih-Kanäle als ein Typ von Effektoren stromabwärts der cAMP-

Bildung dienen könnten.

Vorangegangene Studien mit Gαq, PLC und DAG Mutanten deuten auf eine mögliche

Beteiligung des IP3-Signalweges in der olfaktorischen Signaltransduktion von Insekten

hin. In der vorliegenden Studie wurde ein hoher Expressionslevel von Gαq3 in den

Antennen von Drosophila gefunden, jedoch nicht in den Zilien der olfaktorischen Zellen.

Die Expression eines kontinuierlich aktivierten Gαq3 in den olfaktorischen Neuronen

führte zu einer Verminderung in der Resensitisierung. Diese Ergebnisse weisen darauf

hin, dass Gαq3 nicht an der primären Signaltransduktionskaskade beteiligt ist, aber

möglicherweise eine Rolle in der Endozytose und Degradation der Rezeptoren spielt oder

die olfaktorische Signaltransduktion in Abstimmung mit dem Gαs Signalweg reguliert.

Erste Untersuchungen zur Geruchswahrnehmung von Larven zeigten, dass hier ein

unterschiedlicher zellulärer Signalweg vorliegen könnte, an dem Gαq und nicht Gαs

beteiligt ist.

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82

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Abbreviations

100

Abbreviations

AC Adenylate Cyclase

AL Antennal Lobe

ATP Adenosine 5’-triphosphate

BSA Bovine Serum Albumin

cAMP Adenosine 3’,5’-cyclic monophosphate

CNG Cyclic Nucleotide-Gated Channel

CTX Cholera Toxin

DMEM Dulbecco’s Modified Eagles Medium

DMSO Dimethylsulfoxide

DO Dorsal organ

EAG Electroantennogram

EDTA Ethylene-diamine-tetra-acetic acid

FBS Foetal Bovine Serum

GFP Green Fluorescent Protein

GPCR G-protein-coupled receptor

GRN Gustatory Receptor Neuron

GTP Guanosine-5’-triphosphate

GTPγS guanosine 5’-O-[gamma-thio]triphosphate

HEK Human Embryonic Kidney Cells

HEPES 4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid

IBMX 3-isobutyl-1-Methylxanthine

IP3 Inositol (1,4,5)-trisphosphate

LH lateral horn

MB mushroom body

MIA 5-(N-methyl-Nisobutyl) amiloride

OBP Odorant-Bingding Protein

OR Odorant Receptor

ORN Odorant Receptor Neuron

Abbreviations

101

OSN Olfactory Sensory Neuron

PBP Pheromone-Binding Protein

PCR Polymerase chain reaction

PLC Phospholipase C

PN Projection Neuron

SDS sodium dodecyl sulfate

SEM Standard Error of the Mean

SSR Single sensillum recording

TO Terminal Organ

UAS Upstream Activation Squence

VO Ventral Organ

wt wild-type

List of Figures and Tables

102

List of Figures and Tables

Figure 1.1.1 The Drosophila olfactory organ .................................................................... 3

Figure 1.1.2 Organization of the Drosophila olfactory system. .........................................6

Figure 1.1.3. Analysis of odor response spectra of individual odorant receptors..............11

Figure 1.1.4 Molecular neuroanatomy of the adult AL annotated with the molecular and

functional identity of the glomeruli. .................................................................................14

Figure 1.2.1 Standard model of the GDP/GTP cycle governing activation of

heterotrimeric GPCR signaling pathway. ..........................................................................17

Figure 1.2.2 Structural features of heterotrimeric G-protein subunits...............................18

Figure 1.3.1 The canonical mammalian olfactory signal transduction pathway. ..............20

Figure 1.3.2 Models of signal transduction mechanisms in odorant receptor neurons

(ORNs)...............................................................................................................................22

Figure 3.2.1 Larval chemotaxis assay ...............................................................................34

Figure 3.2.2 Schematic overview of the EAG recording...................................................35

Figure 3.2.3 Schematic overview of single sensillum recording .......................................36

Figure 3.2.4 Schematic overview of single unit tip recording from GRNs .......................36

Figure 4.1.1 PT-PCR analysis of Gα subunit expression in Drosophila antenna..............40

Figure 4.1.2 Western blot analysis of G-protein α subunits expression in antenna...........41

Figure 4.1.3 Immunohistochemical analysis of G-protein α subunit expression patterns in

antenna. ..............................................................................................................................43

Figure 4.1.4 RT-qPCR analysis of Gαs and Gαq expression level in antenna. .................44

Figure 4.2.1 Screening of G-proteins for participation in the olfactory signal transduction46

Figure 4.2.2 Single sensillum recording from wild type ab1 sensillum. ...........................47

Figure 4.2.3 Single sensillum recordings from CTX and control flies..............................48

Figure 4.2.4 Gr5a-CTX GRNs response to sucrose...........................................................49

Figure 4.2.5 Functional study of ORs and Gα proteins interactivity in the recombinant

HEK293 expression system. ..............................................................................................51

Figure 4.2.6 Overexpression of a GTPase deficient Gαs mutant in Drosophila antenna

lead to a change in response dynamics ..............................................................................52

Figure 4.2.7 Odorant exposure caused Gαs redistribution in antenna................................54

Figure 4.2.8 Schematic overview of genetic mosaic fly generation. ................................56

Figure 4.3.1 cAMP level moderate the activity of ORNs..................................................57

Figure 4.3.2 An increase in cAMP levels can recover the spontaneous activity of ORNs.58

Figure 4.3.3 SSR of the OR83b-GAL4 / UAS-PACα; OR83b2 / OR83b2 and control flies59

Figure 4.3.4 Expression of an AcGαq in ORNs leads to a resensitization deficiency ......60

Figure 4.3.5 MIA blocks odorant-induced currents in oocytes ........................................62

Figure 4.3.6 Direct application of MIA on antenna blocks the spontaneous neuronal

activity................................................................................................................................63

Figure 4.3.7 Single sensillum recording analysis from Ih channel mutants.......................64

Figure 4.4.1 Immunostaining of larval ORNs ...................................................................65

Figure 4.4.2 Chemotaxis assays on AcGq and CTX larvae ..............................................66

Table 1 Expression patterns of the complete repertoire of Drosophila odorant receptors. .8

Curriculum Vitae

104

Curriculum Vitae

Personal:

name: Ying Deng data of birth, place of birth: 09.09.1980, Wuhan nationality: China

EDUCATION

PhD Department of Cellphysiology, 2005-present

Ruhr-Universität Bochum, Bochum, Germany Pre-PhD Department of Cellphysiology, 2004-2005

Ruhr-Universität Bochum, Bochum, Germany Master Department of Biology, 2002-2004

Xiamen University, Xiamen, China Bachelor Department of Biology, 1998-2002

Xiamen University, Xiamen, China

RESEARCH EXPERIENCE

International Max-Planck Chemical Biology Program 2005-present Ruhr-Universität Bochum, Germany Doctoral thesis research conducted with Prof. Dr. Dr. Dr. Hanns Hatt Thesis: Molecular mechanisms of olfactory signal transduction in Drosophila melanogaster International Graduate School of Neuroscience 2004-2005 Ruhr-Universität Bochum, Germany Pre-PhD training with Prof. Dr. Dr. Dr. Hanns Hatt Thesis: Interaction partners of odorant receptors from Drosophila melanogaster Regulatory Biology, Department of Biology, 2002-2004 Xiamen University, Xiamen, China Master study conducted with Prof. Dr. Shengcai Lin Thesis: The structural basis of Axin function in the JNK MAPK pathway. Regulatory Biology, Department of Biology, 2001-2002 Xiamen University, Xiamen, China Bachelor degree thesis study conducted with Prof. Dr. Shengcai Lin Thesis: Identification of genes required for TNF-induced cell death.

Curriculum Vitae

105

LIST of PUBLICATIONS

Wong CK, Luo W, Deng Y, Zou H, Ye Z, Lin SC. J Biol Chem. 2004 Sep

17;279(38):39366-73.

The DIX domain protein coiled-coil-DIX1 inhibits c-Jun N-terminal kinase activation by

Axin and dishevelled through distinct mechanisms.

Neuhaus EM, Zhang W, Gelis L, Deng Y, Noldus J, Hatt H. J Biol Chem. 2009 Jun

12;284(24):16218-25.

Activation of an olfactory receptor inhibits proliferation of prostate cancer cells.

Deng Y, Zhang W, Farhat K, Hatt H, Gisselmann G, Neuhaus EM.

The stimulatory heterotrimeric G-protein Gαs mediates olfactory signal transduction in

Drosophila. (In preparation)

Gisselmann G, Deng Y, Neuhaus EM, Werner M, Hatt H,

Pharmacological characterization of MIA as an insect odorant receptor blocker. (In

preparation)

Köper M, Deng Y, Schreiner B, Störtkuhl K, Hatt H, Gisselmann G

Molecular and functional characterization of an Ih-channel in Drosophila olfactory

receptor neurons. (In preparation)

POSTERS

Deng Y, Gisselmann G, Zhang W, Hatt H, Neuhaus EM.

Role of heterotrimeric G-proteins in the olfactory signal transduction cascade in

Drosophila. CSHL Meeting on Neurobiology of Drosophila. Oct. 3 –7, 2007.Cold spring

harbor, New York (poster)

Deng Y, Gisselmann G, Zhang W, Hatt H, Neuhaus EM.

The stimulatory heterotrimeric G-protein Gαs is involved in olfactory signal transduction

in Drosophila. 12th European Drosophila Neurobiology Conference. Sep. 6-10, 2008.

Würzburg, Germany (poster)

Deng Y, Zhang W, Gisselmann G, Hatt H, Neuhaus EM.

Curriculum Vitae

106

The stimulatory heterotrimeric G-protein Gαs is involved in olfactory signal transduction

in Drosophila. 8th Goettingen metting of the German neuroscience society. Mar. 25-29,

Göttingen, Germany (poster)

REFERENCES

Prof. Dr. Dr. Hanns Hatt Ruhr-Universität Bochum, 44780 Institute of Cellphysiology +49 234 32 24586 [email protected] PD. Dr. Eva Neuhaus Ruhr-Universität Bochum, 44780 Institute of Cellphysiology +49 234 32 24315 [email protected] Prof. Dr. Martin Engelhard Max Planck Institute of Molecular Physiology Dept. of Physical Biochemistry Otto-Hahn-Strasse 11 44227 Dortmund Phone: +49 231 133 2302 Email: [email protected]

Acknowledgements

107

Acknowledgements

My sincerest gratitude goes to my supervisor Prof. Dr. Dr. Dr. Hanns Hatt for giving me

the opportunity to become a member of his successful and dynamic team at the

Department of Cell Physiology.

I would like to thank my direct supervisors PD. Dr. Eva Neuhaus and Dr. Günter

Gisselmann for their endless energy and enthusiasm for my work. Their constant

guidance in the right direction, rapid development of new ideas in order to unravel

important questions, clear and concise experimental design and unceasing determination

were invaluable to me.

For their technical support I would like to thank Mr. Harry Bartel, Ms. Farideh Salami,

Ms. Jasmin Gerkrath, Mr. Thomas Lichtleitner, Ms. Andrea Stoeck, Ms. Ute Müller, and

Mr. Grabowski.

A special thanks goes to Julia Dörner, Weiyi Zhang, Ruth Dooley, Nico Bredendiek,

Sabrina Baumgart, Nicole Schöbel, Lain Gelis, Stefan Kurtenbach and Sebastian Rasche

for their friendship and support, interesting discussions, unconditional help during my

thesis writing and ideas for problem-solving.

To my parents I am forever indebted, for their unwavering support throughout my years

of study.

This work was financially supported by The International Max-Planck Research School

in Chemical Biology and The Deutsche Forschungsgemeinschaft.

doctoral dissertation ying deng department of cell ... · department of cell physiology ... dengue...

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