CHAPTER

Imaging cilia in Drosophila melanogaster

15

Jennifer Vieillard, Jean-Luc Duteyrat, Elisabeth Cortier, Be´ne´dicte Durand1 Centre de Ge´ne´tique et de Physiologie Mole´culaire et Cellulaire, Universite´ Claude Bernard Lyon-1, Lyon, Villeurbanne, France 1

Corresponding author: E-mail: [email protected]

CHAPTER OUTLINE Introduction ............................................................................................................ 280 1. Immunodetection of Cilia .................................................................................... 283 1.1 Embryonic Cilia................................................................................... 285 1.1.1 Collection of embryos ....................................................................... 285 1.1.2 Dechorionation ................................................................................. 285 1.1.3 Staining of whole mount embryos...................................................... 287 1.1.4 Staining of dissected embryos........................................................... 288 1.2 Pupal Antennae Cilia ........................................................................... 290 1.2.1 Preparation of antennae ................................................................... 290 1.2.2 Immunostaining ............................................................................... 291 1.3 Materials ............................................................................................ 291 2. Observation of Cilia Ultrastructure by Transmission Electron Microscopy............... 293 2.1 Methods ............................................................................................. 293 2.1.1 Dissection ........................................................................................ 293 2.1.2 Fixation ............................................................................................ 293 2.1.3 Postfixation....................................................................................... 294 2.1.4 Embedding....................................................................................... 294 2.1.5 Sectioning ........................................................................................ 295 2.2 Materials ............................................................................................ 296 2.3 Data Analysis ...................................................................................... 298 Acknowledgments ................................................................................................... 298 References ............................................................................................................. 298

Methods in Cell Biology, Volume 127, ISSN 0091-679X, http://dx.doi.org/10.1016/bs.mcb.2014.12.009 © 2015 Elsevier Inc. All rights reserved.

279

280

CHAPTER 15 Imaging cilia in Drosophila melanogaster

Abstract Drosophila melanogaster is a powerful genetic model organism to understand the function of proteins in specific cellular processes. Cilia have been extensively studied in Drosophila playing various sensory functions that are essential for fly survival. Indeed, flies defective in cilia formation cannot walk, fly, or feed properly. Drosophila harbors different types of cilia that can be motile or immotile or that can show compartimentalized (intraflagellar transport (IFT)-dependent) or cytoplasmic (IFT-independent) mode of assembly. Therefore, Drosophila represents an advantageous model organism to study the function of novel ciliary candidates and to address specific questions such as their requirement for IFT-dependent processes versus other aspects of cilia-associated functions. This chapter describes protocols to visualize cilia by direct or indirect fluorescent labeling and protocols to analyze ciliary ultrastructure by electron microscopy.

INTRODUCTION The fruit fly Drosophila melanogaster is a powerful model system to study ciliogenesis as there are only a few types of ciliated cells, which resume the different types of cilia found in many other organisms. In addition, Drosophila ciliary functions can be assessed by a wide range of sensory and behavioral assays. In Drosophila, cilia are responsible for transducing all senses except vision. Sensory perception is mediated by cilia found on type I sensory neurons (monodendritic neurons) of the peripheral nervous system (PNS), which are found in two types of sensory organs (Figure 1(A)) (Bate & Martinez arias, 1993): the neurons of the external sense (ES) organs, which play mechano, chemo, or olfactory-sensory functions and the neurons of the chordotonal (CH) organs that are involved in proprioception and hearing (Gogendeau & Basto, 2009; Jarman, 2002; Keil, 2012; Kernan, 2007). These two categories of sensory organs are described for all Drosophila stages from late embryos to adults. In Drosophila embryos and larvae, both ES organs and CH organs are located on the body wall of each thoracic and abdominal segments according to a stereotyped pattern (Bate & Martinez arias, 1993; Orgogozo & Grueber, 2005). In adult flies, ES organs include the bristles covering the body, the different sensilla of the wings, legs, and proboscis and olfactory sensilla of the third segment of the antenna. CH organs are present in the limb joints, the abdomen, and a specialized array of CH organs are found in the antennae that are involved in hearing (Eberl & BoekhoffFalk, 2007; Eberl, Hardy, & Kernan, 2000). ES organs are composed of one neuron, one or several glial cells, and two support cells (socket and hair cells) that form the specialized structures on the cuticle. CH organs are composed of one (limb joints) to two or three neurons (antennae) and several associated cells: the scolopidia which ensheath the neuron and produce specialized structures around the dendrite and cap and ligament cells at both ends to connect the neuron to the cuticle (Figure 1(A)). CH and ES neurons harbor a single primary cilium at the distal end of their dendrite. Cilia of sensory neurons require intraflagellar transport (IFT) for their assembly (Han, Kwok, & Kernan, 2003; Sarpal et al., 2003) and

Introduction

FIGURE 1 The different types of ciliated cells in Drosophila. (A) Cilia are found at the tip of the monodendritic neurons of type 1 sensory organs of Drosophila melanogaster. These can be divided in two groups: external sensory organs and chordotonal organs. Cilia have a 9 þ 0 architecture. The proximal segment of the chordotonal cilia also shows dynein arms that ensure cilia motility required for hearing. Cilia assembly in the PNS relies on IFT. (B) Flagella are found on spermatozoa and have a 9 þ 2 architecture with a central pair, radial spokes, nexin links, and dynein arms to ensure motility. Sperm flagella assembly does not rely on IFT and is carried inside the cytoplasm. Primary like cilia are assembled in late spermatocytes and engulfed inside the cell during meiosis. These have been well described and protocols to detect these cilia-like structures have been recently published (Gottardo et al., 2013; Martins et al., 2010; Riparbelli, Callaini, & Megraw, 2012). IFT, intraflagellar transport.

show a characteristic 9 þ 0 ultrastructure but different subtypes can be distinguished for the different kinds of sensory neurons. The cilium of the CH neurons is separated in two parts by the ciliary dilation (Moulins & Mill, 1976; Uga & Kuwabara, 1965) (Figures 1 and 2). The axoneme of the proximal part harbors dynein arms that serve motility and force generation to actively amplify sound-induced antennal vibrations (Effertz, Wiek, & Go¨pfert, 2011; Go¨pfert, Humphris, Albert, Robert, & Hendrich, 2005; Go¨pfert & Robert, 2003; Kavlie, Kernan, & Eberl, 2010). The distal part of the axoneme is devoid of dynein arms. It concentrates several transient receptor potential ion channels such as NompC also required for hearing (Lee & Ashrafi, 2008). The cilium of the ES organs does not include axonemal dynein arms and shows specialized distal outer segments (for review, see Keil, 1997, 2012). The second type of ciliated cell is found in the male germ cells in Drosophila. In spermatocytes, each of the four centrioles first dock to the plasma membrane and

281

282

CHAPTER 15 Imaging cilia in Drosophila melanogaster

FIGURE 2 Organization of the chordotonal neurons and position of some useful markers of the cilia (see Table 2 for more details).

protrude at the cell surface forming a cilium-like extension (Fabian & Brill, 2012; Gottardo, Callaini, & Riparbelli, 2013; Riparbelli, Megraw, Callaini, & Megraw, 2012). The motile flagellum of spermatozoa is further extended from the centriole in spermatids and shows a typical 9 þ 2 axoneme (Figure 1(B)). Whereas the sperm axoneme shares many features with human motile axoneme, one specificity of Drosophila is that flagellar axoneme formation does not require IFT for its assembly and is thus described as a cytoplasmic mode of cilia assembly (Han et al., 2003; Sarpal et al., 2003). This makes Drosophila particularly suitable to distinguish IFT-related functions versus other ciliary associated functions. Several studies in the past years have shown that functional analyses in Drosophila were powerful to understand the role of many proteins involved in cilia assembly. For example, functional analysis associated with genome-wide studies helped to pinpoint on several novel genes required for cilia assembly (for example, Avidor-Reiss et al., 2004; Laurenc¸on et al., 2007; Newton et al., 2012). As well, analysis of RNAs specifically enriched in CH neurons helped to identify novel genes required for motility and was informative for human diseases linked with motility defects (for example, Cachero et al., 2011; Moore et al., 2013; Senthilan et al., 2012). In this chapter, we describe three protocols to visualize cilia by immunofluorescence in either embryos

1. Immunodetection of cilia

(whole mount or dissected) or in pupal antennae. We also detail one protocol to perform electron microscopic observations of cilia ultrastructure.

1. IMMUNODETECTION OF CILIA Cilia in the PNS can be visualized at all different stages: embryos, larvae, pupae, or adults. In adults, the presence of the cuticle renders antibody stainings more difficult but have been described successfully to visualize cilia either in wings and more often in antennae (for example, Cheng, Song, Looger, Jan, & Jan, 2010; Shaham, 2009). More sensitive labeling can be performed on sections of the antennae to visualize cilia in adult stages. Detailed description of adult antennae cryosection and whole mount immunostaining can be found in Saina and Benton (2013). Use of pupal antennae for immunostaining will be described in this chapter. At this stage, antibody penetration is improved. In embryos, cilia can be visualized by whole mount staining using standard protocols for immune detection. However, in our hands, we obtained images with better resolution and sensitivity after opening and flattening embryos. In some cases, like for epitopes that do not resist to methanol treatment, this is the only option. Both protocols will be presented here. To facilitate cilia observations, the PNS neurons can be labeled using either the 22C10 antibody that stains all neuronal membranes but not the ciliary membrane (Hummel, Krukkert, Roos, Davis, & Kla¨mbt, 2000) (Figures 2 and 3(E), (H), and (J)), or the anti-Horseradish peroxidase (HRP) antibody that also labels the ciliary ending (Jan & Jan, 1982) and some associated scolopale structures (Figures 2 and 3(F)). Cilia of the CH neurons (Figure 3(G)) are longer and hence easier to observe. In this chapter, we focus mainly on CH cilia but the same protocols can be used to observe ES cilia. Several useful markers of different cells or ciliary compartments can be used (Table 1, Figure 2). The ciliary axoneme can be specifically labeled using GT335 antibody that recognizes polyglutamylated tubulin (Wolff et al., 1992) (Figure 3(I)). In antennal segments, specific structures that are associated with the scolopale cell of CH neurons can be labeled using phalloidin and are useful to point to the ciliated region of the CH neuron (see, for example, Senthilan et al., 2012). Several GFP reporter strains have also been published and can be used to label specific substructures of the cilia: Nomp-A::GFP (Chung, Zhu, Han, & Kernan, 2001) allows the labeling of the ciliary dilation, CBY::GFP and Unc::GFP label the ciliary transition zone, Nomp-C (antibody or NompC-GFP) labels the distal segment beyond the ciliary axoneme (Cheng et al., 2010; Lee, Moon, Cha, & Chung, 2010; Walker, Willingham, & Zuker, 2000) and centriole markers (for example, Sas4, PLP, or Asterless) have been used to label the basal body (Enjolras et al., 2012; Ma & Jarman, 2011). A specific review of the centriole-associated markers and of protocols to detect centrioles in Drosophila is available in a previous chapter (Martins, Machado, Callaini, & Bettencourt-Dias, 2010).

283

284

(B)

(D)

(E)

(C)

(G)

(F)

(H)

(I)

(J)

(K)

FIGURE 3 Imaging cilia in embryos. (A) Egg-collecting chamber and agar plate used to collect embryos. (B) Sieve used to collect embryos for dechorionation. (C) Slide used to dissect embryos. (D) Observation of a stage 16 embryo on the binocular. Note the three major gut lobes clearly distinguishable at this stage. (E) Whole mount embryo labeled with 22C10 antibody and Diaminobenzidine (DAB) staining showing the regular arrangement of type I neurons in

CHAPTER 15 Imaging cilia in Drosophila melanogaster

(A)

1. Immunodetection of cilia

1.1 EMBRYONIC CILIA 1.1.1 Collection of embryos In Drosophila embryos, PNS neurons and their support cells begin to differentiate at stage 13 and are easily observable until stage 17. Embryonic stages can be easily recognized based on the shape of the gut. To collect embryos between stage 13 and 17: 1. Place young adults Drosophila (3e4-days old, both males and females) on egglaying plates covered with fresh yeast paste in egg-collecting cages and maintain at 25  C (Figure 3(A)). 2. On the morning of the second day, change egg-laying plates every 2 h in order to habituate flies to tremor and to avoid egg retention by females. Throw away plates. Change the egg-laying plates at 2 pm. Replace collecting plates at 6 pm. Incubate harvested egg-laying plates at 18  C. Let the females lay eggs overnight at 25  C. 3. At 9 am the third day, collect the embryos from the night egg-laying plates and the laying plates kept at 18  C. The majority of the collected embryos will be between stage 13 and 17.

1.1.2 Dechorionation During embryonic development, embryos are protected by the chorion that must be removed before further treatments. This can be done using bleach (alternatively this can also be done by hand by rolling embryos on double-sided tape, which avoids contact with methanol). 1. Pour PBS1X/0.1%Triton X-100 (PBT) on the laying plate and suspend the embryos using a paintbrush. 2. Transfer the embryos on a sieve (Figure 3(B)). Wash the embryos with PBT. 3. Dechorionate the embryos by incubating 3 min the sieve in a petri dish filled with bleach (9.6% sodium hypochlorite solution) diluted 1/5 in water. Do not over

=

abdominal segments A1eA7. (F) Chordotonal neurons labeled for HRP and DAB staining showing the cilia inside the scolopale-associated structure. HRP also stains the scolopale cap (arrow). (G) Scheme of the distribution of type I sensory neurons in one Drosophila abdominal segments. (Adapted from Orgogozo and Grueber (2005).) Empty circles represent ES organs, the straight lines show the direction of the dendrite. Filled black triangles represent chordotonal neurons. The sharp tip of the triangle points toward the tip of the dendrite. (H) Whole mount embryos showing chordotonal cilia labeled with 22C10 and anti-GFP antibodies (Arl13b::GFP transgene). (I) Dissected embryos showing chordotonal neurons labeled with anti-HRP and GT335 (polyglutamylated tubulin) antibodies. (J) Chordotonal neurons of dissected embryos labeled with 22C10 and anti-GFP antibodies (Arl13b::GFP transgene). (K) Higher magnification of chordotonal neurons of dissected embryos labeled with 22C10 and anti-GFP antibodies (Arl13b::GFP transgene). Note the cilia (arrowhead) and the ciliary dilation (arrow). Scale bars: 10 mm. ES, external sense.

285

286

CHAPTER 15 Imaging cilia in Drosophila melanogaster

Table 1 Useful Makers of Ciliated Neurons and of Specific Cilia Compartments Published in Drosophila Tools

Ciliary Compartment

References

Tagged proteins (endogenous or ubiquitous promoter) IAV::GFP Nan::GFP NompB::GFP

Proximal ciliary segment Proximal ciliary segment Outer dendritic segment

Unc::GFP

Basal body and transition zone Transition zone Ciliary dilation Ciliary cap Proximal ciliary segment Dendrite, ciliary rootlet Basal body (ubiquitously expressed) Transition zone Inner and outer dendritic segment Inner and outer dendritic segment Inner and outer dendritic segment

CBY::GFP RempA::YFP NompA::GFP Arl13b::GFP (CG11356) Rootletin::GFP Ubq::Sas-4-GFP B9D1::MYC (CG14870) Ccdc151::GFP Hmw::GFP Zmynd10::Venus

Gong et al. (2004) Newton et al. (2012) Han et al. (2003) and Moore et al. (2013) Baker, Adhikarakunnathu, and Kernan (2004) Enjolras et al. (2012) Lee et al. (2008) Chung et al. (2001) Enjolras et al. (2012) Laurenc¸on et al. (2007) Peel, Stevens, Basto, and Raff (2007) Enjolras et al. (2012) Jerber et al. (2013) Soulavie et al. (2014) Moore et al. (2013)

Tagged proteins (UAS control) Doublecortin-GFP

DILA–Flag Klp64D-GFP Klp68D-YFP IAV-GFP GFP-Oseg1 Oseg2-GFP Oseg3-GFP Ose4-GFP Oseg5-GFP NompC-GFP TilB-GFP

Ciliary dilation in CH neurons/distal segment (tubular body) in ES neurons Transition zone Inner and outer dendritic segment Inner and outer dendritic segment Proximal ciliary segment Outer dendritic segment* Outer dendritic segment* Outer dendritic segment* Outer dendritic segment* Outer dendritic segment* Distal end of the cilia (after ciliary dilation) Inner and outer dendritic segment

Bechstedt et al. (2010)

Ma and Jarman, (2011) Jana, Girotra, and Ray (2011) Jana et al. (2011) Kim et al. (2003) Avidor-Reiss et al. (2004) Avidor-Reiss et al. (2004) Avidor-Reiss et al. (2004) Avidor-Reiss et al. (2004) Avidor-Reiss et al. (2004) Cheng et al. (2010) Kavlie et al. (2010)

1. Immunodetection of cilia

Table 1 Useful Makers of Ciliated Neurons and of Specific Cilia Compartments Published in Drosophiladcont’d Tools

Ciliary Compartment

References

Tagged proteins (endogenous or ubiquitous promoter) GFP-Bug22

Proximal ciliary segment

Mendes Maia, Gogendeau, Pennetier, Janke, and Basto (2014)

Other useful transgenic reporter lines UAS-DsRED UAS-CD8-GFP UAS-myr-mRFP

Cytoplasm Membrane-tagged GFP Membrane-tagged RFP

Bechstedt et al. (2010) Han et al. (2003) Sun et al. (2009)

Cell body, axon and dendrite, excluded from the cilia Neuronal membrane- and scolopale-associated structures Luminal space that surrounds the cilia (base of the outer segment and proximal part of the cilia) Cilia proximal segment Cilia distal segment

Fujita, Zipursky, Benzer, Ferru´s, and Shotwell (1982) and Hummel et al. (2000) Dubruille et al. (2002), Husain et al. (2006), and Jan and Jan (1982) Cook, Hardy, McConnaughey, and Zuker (2008), Fujita et al. (1982), and Husain et al. (2006) Park et al. (2013) Lee et al. (2010) and Liang, Madrid, Saleh, and Howard (2011)

Actin-rich scolopale rods

See, for example, Senthilan et al. (2012)

Antibodies Futsch/22C10

HRP

Eyes shut/21A6 antibody/Spam

Tulp NompC antibody

Other Phalloidin

Both transgenic strains for tagged proteins and available antibodies are reported. For specific images, see associated reference. ES, external sense. * Oseg protein distribution is only partially described.

treat the embryos with bleach, the vitelline membranes should remain intact after treatment. 4. Wash extensively the embryos under tab water to completely remove bleach.

1.1.3 Staining of whole mount embryos 1.1.3.1 Removal of vitelline membrane 1. Collect embryos with a paintbrush and resuspend the embryos in an Eppendorf tube containing 500 mL of heptane and allow embryos to sink to the bottom of the tube.

287

288

CHAPTER 15 Imaging cilia in Drosophila melanogaster

2. Add 500 mL of fixative solution A (PBS1X/3.7% paraformaldehyde (PFA)/ 0.25 M EGTA) and incubate 20 min at room temperature on a rotating wheel. Embryos should be at the interface between the heptane and fixative solution. If they are too many embryos, increase the volumes of heptane and fixative solution and use bigger tubes to expand the surface of the interphase. 3. Eliminate the inferior phase containing the fixative solution without removing embryos. 4. Add 1 mL of cold methanol (20  C) and shake vigorously for 20 s. Note: The embryos should sink to the bottom of the tube. Remove the heptane/methanol mix. 5. Wash the embryos once with methanol. 6. Remove the methanol and rehydrate the embryos in PBS1X/ethanol 50%, two times 5 min and then in PBT. 7. Rinse two times with PBT. Note: You can keep the embryos in methanol at 20  C. However, long exposure to methanol alters epitopes for immunostaining. It is best to rehydrate and use immediately embryos for immunostaining.

1.1.3.2 Staining All incubations are performed with gentle rocking. 1. Incubate embryos 30 min at room temperature in 500 mL of blocking solution (PBT/3% BSA/5% normal goat serum (NGS)). 2. Remove the blocking solution and add the primary antibodies diluted in the blocking solution. Incubate overnight at 4  C (see Table 2 for antibody concentrations). 3. Wash three times 30 min in PBT/0.1% BSA. 4. Incubate between 3 and 4 h at room temperature with secondary antibodies diluted in PBT/0.1% BSA. 5. Wash three times 10 min in PBS1X/0.1% Tween 20 (PBTW). 6. Remove almost all PBTW. Using a cut pipet tip transfer embryos to a glass slide. 7. Remove all PBTW using an absorbing paper. 8. Add a drop of Dako mounting medium and spread the embryos on the slide. 9. Put a coverslip over the embryos and seal with nail polish. Note: Triton X-100 can be increased to 0.3% in all steps.

1.1.4 Staining of dissected embryos This protocol is particularly suited to look at specific subcompartments of cilia or for weak signals. It is also efficient for epitopes that are sensitive to methanol treatment as vitelline membrane retrieval is hand performed. It allows collection of highresolution images of cilia from the 5 lateral CH neurons (Figure 3). Cilia from ES neurons at this stage are very short. Embryos are collected and dechorionated as described above.

1. Immunodetection of cilia

Table 2 Antibodies Concentration Used in Immunostaining Protocols Primary Antibodies

Mouse anti-Futsch Rabbit anti-GFP Rabbit anti-HRP antibody GT335 antibody Anti-myc antibody

Source

Catalog n

Dilution

DSHB Invitrogen Jackson Immuno Research Enzo life sciences Euromedex

22c10 A11122 325-005-021

1/250 1/1000 1/500

ALX-804-885-C100

1/500

Myc-1A1

1/1000 to 1/2500

Invitrogen

A11008

1/1000

Invitrogen

A21135

1/1000

Invitrogen

A21141

1/1000

Invitrogen

A21428

1/1000

Secondary antibodies Goat antirabbit ALEXA FLUORÒ488 Goat antimouse ALEXA FLUORÒ594 Goat antimouse ALEXA FLUORÒ488 Goat antirabbit ALEXA FLUORÒ555

1.1.4.1 Preparation of dissecting slides The day before, coat slides with polylysine to allow embryos to strongly stick for subsequent treatments. 1. 2. 3. 4.

Wash the slide with SDS 5%. Rinse two times in distilled water. Wash the slide with ethanol 96% and let dry. Pour 500 mL of polylysine 0.1 mg/mL on the slide and incubate 20 min at room temperature. 5. Remove the polylysine and let the slide dry for 15 min. 6. Delimit a pool with silicone around the polylysine surface (Figure 3(C)). 7. Keep at 4  C over night.

1.1.4.2 Dissection, fixation, and immunostaining This is the critical step because embryos must be correctly oriented and are, at this point, very fragile and easily breakable. 1. On slides prepared as above, place a piece of double-sided tape on one side of the silicone pool (Figure 3(C)). 2. Collect and dechorionate embryos as described above. 3. Using a paintbrush, transfer the embryos from the sieve (after dechorionation) in a watch glass containing PBS1X.

289

290

CHAPTER 15 Imaging cilia in Drosophila melanogaster

4. Under the binocular magnifier, select embryos of correct stage based on gut shape. With a paintbrush or a straight needle probe transfer selected embryos on an agar juice plate and remove all the PBS1X using an absorbing paper. Note: The easiest embryos to dissect (because they stick easily to slides) are at stage 16, when the midgut forms four chambers (Figure 3(D)) (Ashburner, 1989). The three major ones are clearly distinguished under the binocular and make three parallel stripes (Figure 3(D)). Moreover, the ciliary ending is best developed at stage 16. 5. Transfer embryos on the tape of the dissecting slide using a straight needle probe. Be careful to remove all the PBS1X before depositing embryos on the tape. Orient embryos ventral side toward the polylysine surface with the micropyle on the dorsal face pointing on top. Press gently the embryos to allow them to stick strongly on the tape. 6. When several embryos are placed, fill the silicone pool with 400 mL of PBS1X. 7. Using a tungsten needle, pierce the vitelline membrane in the posterior side, take out the embryos, and stick them on the polylysine surface. Cut the dorsal face of the embryos and move apart body walls to stick them on the polylysine (you can see the ventral cord of the CNS). Finally, remove the gut. 8. When all the embryos are dissected, remove the PBS1X and add 400 mL of the fixative solution B (PBS1X/4% PFA). Incubate 20 min at room temperature. 9. Remove the fixative solution and rinse with PBS1X. 10. Incubate 2 h in the blocking solution (PBT/3%BSA/5% NGS). 11. Incubate overnight at 4  C with primary antibodies diluted in blocking solution (see table below for antibody concentrations). 12. Wash three times 15 min with PBT/0.1% BSA. 13. Incubate 2 h at room temperature with secondary antibodies diluted in PBT/ 0.1% BSA. 14. Wash three times 15 min with PBTW. 15. Remove the silicone using a razor blade. 16. Mount between slide and coverslip in Dako mounting medium. 17. Seal the coverslip with nail polish. See Figure 3(E), (F) and (H)e(K) for some examples of embryonic cilia staining. Note: A critical point is to be careful not to allow samples to dry at any stage. Always use moist chambers for incubations and work quickly after removing solutions during washing steps.

1.2 PUPAL ANTENNAE CILIA 1.2.1 Preparation of antennae 1. Pick young pupae (pupal stage P0), let them mature for 36e40 h at 25  C in a fly raising tube to reach pupal stage P5eP7 (Ashburner, 1989). 2. Cut the pupae between the abdomen and thorax with a microscalpel (Figure 4(A)). Extract the head from the cuticule and remove thorax tissues.

1. Immunodetection of cilia

3. Fix the head 30 min at room temperature with fixative solution B (PBS1X/4% PFA) and wash three times 5 min with PBS1X. 4. Transfer the head on a glass slide to dissect more precisely the antennae and keep the superior part of the cuticle of the head. Put the antennae in a 0.65 mL lowbinding tube containing PBS1X. 5. Antennae can be used immediately for immunostaining or can be stored at 4  C for a few days.

1.2.2 Immunostaining All incubations are performed with gentle rocking. 1. Permeabilize tissues 1 h at room temperature with PBS1X/0.3% Triton X-100. 2. Replace PBS1X/0.3% Triton X-100 with blocking buffer (PBS1X/0.3% Triton X-100/5% NGS) and incubate 1e2 h at room temperature. 3. Remove the blocking buffer and incubate with primary antibodies diluted in blocking buffer during 48 h at 4  C (see Table 2 below for antibody concentrations). 4. Wash four times 15 min with PBS1X. 5. Replace PBS1X with secondary antibodies diluted in blocking buffer and incubate 48 h at 4  C. For this step we use secondary antibodies diluted to 1/500. 6. Wash four times 15 min with PBS1X. 7. Place the antenna on a glass slide, isolate the antenna, and add a drop of Vectashield. Before covering with the coverslip, add on both sides two drops of silicone to avoid complete crushing of the antennae (Figure 4(B)). Note: Some typical images of cilia from Johnston’s organ are presented in Figure 4(C).

1.3 MATERIALS Egg-laying plates grape fruit and agar plates (composition: 15 g Agar, 14.5 g saccharose, 29 g glucose, 100 mL grape fruit juice, 400 mL distilled water, and 1.25 mL NaOH 10 N. After sterilization add 2.3 mL propionic acid and 0.2 mL phosphoric acid. Mix and distribute the medium in small petri dishes 60 mm diameter) Mesh for sieves (Sefar nitex, 03-150/50 102 cm) Microscalpel (Sharpoint microsurgical knife, f 72-2201) Fine forceps (Dumont #5 ForcepsdInox Medical Biologie Fine Tips) Dissecting needle probe (Moria nickel plated pin holder and 0.2 mm minutien pins) Tungsten needle (Tungsten needlesd0.125 mm diameter) 0.65 mL low-binding tube (Dutscher, 027200) PBS1X (Eurobio, CS0PBS01-08) NGS (Euromedex, GTX73206) PBT (PBS1X/01% Triton X100)

291

292

CHAPTER 15 Imaging cilia in Drosophila melanogaster

FIGURE 4 Imaging cilia on whole mount antennae. (A) Different steps to dissect pupal heads and antennae. (B) Schemes of a Drosophila head showing the antennae with dotted lines

2. Observation of cilia ultrastructure by transmission electron microscopy

PBTW (PBS1X/0.1% Tween 20) Polylysine (Poly-L-lysine hydrobromide SIGMA, P1524) Slides (Dutscher, 100,000) Dako mounting medium (Dako fluorescence mounting medium, S3023) Vectashield mounting medium (Vector laboratories, H-1000) Yeast paste (Dutscher, 789,093) PFA (Sigma Aldrich, P6148) Formaldehyde solution (Sigma Aldrich, F8775) Arl13b::GFP tagged transgenic flies have been previously described (Enjolras et al., 2012)

2. OBSERVATION OF CILIA ULTRASTRUCTURE BY TRANSMISSION ELECTRON MICROSCOPY Cilia ultrastructure is usually observed in adult stages because ciliated neurons are easier to locate. The antenna is particularly suited to look for either CH cilia or olfactory cilia, which are both present in high number. CH neurons of the leg joints are also quite easy to locate, but because only a few of these neurons are present in each limb it is more convenient to work on antennae.

2.1 METHODS 2.1.1 Dissection Whole adult fly heads are cut from the body and antennae are dissected by cutting on both eye sides (Figure 5(B)) and immersed as fast as possible in the fixative solution. Make sure the antennae do not float on top of the solution otherwise fixation will not be efficient. Remove any tiny air bubbles attached to the antennae by pipetting the fixative solution to allow efficient fixation of the antenna.

2.1.2 Fixation 1. Fix by immersion for 48 h at 4  C in 0.1 M sodium cacodylate buffer, pH 7.4/2% glutaraldehyde/0.5% PFA. If possible check the osmolarity of your sodium cacodylate buffer by measuring the freezing point with an osmometer (at least the first time you use the stock cacodylate solution). Osmolarity and pH must absolutely be controlled to preserve ultrastructure. Osmolarity should be 200 mOsM for the cacodylate buffer. The fixative should not contribute to more than

=

indicating the region that is dissected and of the mounted antennae with the silicone drops between slides and coverslips. (C) Immunostaining of cilia in the Johnston’s organ labeled with 22C10 and anti-GFP antibodies (Arl13-b::GFP transgene; C1) or with anti-HRP and GT335 (polyglutamylated tubulin) antibodies (C2eC3). Arrowhead points to the dendrite. Arrows point to the ciliary ending. (C3) The two arrows point to the proximal and distal end of the cilium. Scale bars: 10 mm. (See color plate)

293

294

CHAPTER 15 Imaging cilia in Drosophila melanogaster

FIGURE 5 Electron microscopy analysis of chordotonal cilia from antennae. (A) Scheme of the organization of the antennae and of the orientation of the antennae in silicone molds. Note that only one antenna is presented in each bloc but up to six antennae can be embedded together in one bloc. On the right is a representative scheme of a cross (C) section or a longitudinal (L) section of the second segment of the antennae (Johnston’s organ). (B1e2) Representative longitudinal sections of the antennae showing cross sections of the chordotonal neurons present in the Johnston’s organ.

400 mOsM (2% glutaraldehyde contributes to 200 mOsM and 0.5% PFA to 100 mOsM). PFA penetrates faster into tissues, hence it is better for fixing quickly samples whereas glutaraldehyde allows a better fixation but shows a slower penetrance. In addition, glutaraldehyde fixation is irreversible whereas PFA fixation is reversible. Antennae can be kept several days or months in the fixation solution before subsequent treatment. 2. Wash four times 15 min at room temperature with 0.15 M sodium cacodylate pH 7.4.

2.1.3 Postfixation 1. Postfix in 0.1 M sodium cacodylate pH 7.4/1% osmium tetroxide (OsO4) for 4 h at room temperature in the dark. Osmium tetroxide essentially reacts with double lipid unsaturated connections of lipid membranes and enhances contrast of these structures. 2. Rinse in distilled water for 5 min.

2.1.4 Embedding 1. Dehydrate through several ethanol series (30%, 50%, 70%, 80%, 95%) for 30 min each, followed by three incubations of 15 min in ethanol 100% and two

2. Observation of cilia ultrastructure by transmission electron microscopy

baths of propylene oxide for 15 min each. To conserve unsaturated lipids, the first four baths of ethanol (until 80%) have to be performed at 4  C. 2. Immerse antennae in successive baths of Epon resin mix (volume v)/propylene oxide (volume v) at room temperature: 1v/3v for 1 h, 1v/1v for 1 h, and finally 1v/3v for 1 h. 3. Immerse in Epon resin mix for 1 h then replace with fresh Epon resin mix and incubate 12 h at room temperature. 4. Immerse in a freshly prepared solution of Epon resin mix for 2 h at room temperature. Embed antennae in flat silicone molds. Under the binocular, orientate antennae either parallel or perpendicular to the surface of the bloc (Figure 4(A)). Place molds in an oven for 48 h at 56  C to harden the Epon resin. Epoxy monomers upon heating form a polymer branched network. The length of the chain of polymers depends on the temperature. Be aware that overheating can lead to excessively hard resin that may be difficult to section.

2.1.5 Sectioning 1. Semithin sections (500 nm to 1 mm) are cut on an ultramicrotome, collected on glass slides, and stained with methylene blue/toluidine blue (1:1 ratio) to check sections and structures. When appropriate, serial 75 nm ultrathin sections are cut and collected on a formvar-coated copper grid. Semithin and ultrathin sections are both obtained with a diamond knife. 2. Stain ultrathin sections of antennae with aqueous uranyl acetate in an Ultrostainer (1 h at 56  C followed by 4 min at 20  C with lead citrate (Leica Ultrostain II (lead citrate solution). The staining program ends with a 4 min wash with 1 ml 65% nitric acid in 1L distilled water and several washes with pure distilled water. An optional wash with water can be performed: fill a crystallizer with distilled water and dive grids at least 20 times into the water. Air-dry grids. Protect from light during staining with uranyl acetate to avoid precipitates to appear. During lead citrate treatment, avoid exposure to air because of steam/water interactions with lead citrate that will form black aggregates. In the absence of an Ultrostainer, contrast can be performed manually as follow: incubate grids 15 min (at room temperature in the dark) upside down on drops of uranyl acetate 7% in methanol (previously filtered). Rinse two times in methanol (dive grids at least 20 times in methanol). Rinse grids (as above, dive grids at least 20 times in solution) once with 50% methanol/50% distilled water, and once in distilled water. Let dry 1 h at room temperature. Incubate 10 min at room temperature in a closed chamber (containing NaOH pastilles to absorb humidity) with lead citrate (prepared as described by Reynolds (1963): 1.33 g of Pb(NO3)2 and 2.13 g of Na3(C6H507),5H2O in 30 mL distilled water). Rinse with NaOH 0.02 N and three times with distilled water. Air-dry grids. Uranyl acetate stains nucleoproteins whereas lead citrate contrasts all membrane systems. This overall staining procedure is critical for proper contrasts. Grids can be stored in grid boxes for years.

295

296

CHAPTER 15 Imaging cilia in Drosophila melanogaster

Note: Both cross sections and longitudinal sections can be successively analyzed from a single bloc because CH neurons have a spherical arrangement inside the antennae. However, it is better to use longitudinal sections of antennae to look for cross sections of cilia (Figure 5(A) and (L)) and cross sections of the antennae to look for longitudinal sections (Figure 5(A) and (C)). We also put a group of up to six antennae in one block to have the highest probability to find useful sections of cilia. Incubation steps for embedding should not be shortened to avoid incomplete impregnation of the antennae that could lead to its tearing from the resin while sectioning. Always use fresh fixative solutions (diluted fixative solution should not be kept more than a week and concentrated solutions should not be kept more than a month after opening). Small conditioning of electron microscopy (EM) grade fixative should be preferred. The same procedure can be used to visualize primary cilium like extensions in spermatocytes and spermatid flagella.

2.2 MATERIALS Glutaraldehyde EM Grade 50% aqueous solution (10 mL vial, Electron Microscopy Sciences) Formaldehyde aqueous solution EM Grade 16% (10 mL vial, Electron Microscopy Sciences) Sodium cacodylate trihydrate (12310, Electron Microscopy Sciences) Osmium tetroxide 4% (1 g vial, Laurylab) BDMA (11,400, Electron Microscopy Sciences) Epon resin mix: Epon A 1 volume þ Epon B 1 volume þ 1.7% BDMA Epon A: 62 mL glycidyl ether 100 (Fluka) þ 100 ml dodecenyl succinic anhydride (Fluka) Epon B: 100 mL glycidyl ether 100 (Fluka) þ 88 ml methylnadic anhydride (Fluka) Lead citrate (Leica Ultrostain 2, Laurylab) Uranyl acetate 0.5% solution (Ultrostain, Laurylab) Absolute Alcohol (VWR) Propylene oxide (VWR) Toluidine blue (89,640dFluka) Methylene blue (Chimie-Plus Laboratories, 81,044) Silicone molds: Flat Embedding Molds 70,900 (Electron Microscopy Sciences) Osmometer: Fiske QF osmometer model 330D Uxbridge, Massachusetts Ultramicrotome Leica UC7 Ultrostainer Leica Diamond knife Diatome ultra 45 MF 1185 Diamond knife Diatome histo 45 Hi 8669

2. Observation of cilia ultrastructure by transmission electron microscopy

FIGURE 6 Observations of Drosophila Chordotonal cilia by electron microscopy (EM). (A) Scheme of a longitudinal section of an antennal chordotonal organ composed of two or three neurons ensheathed by a scolopale cell (only the dense structures produced by the scolopale are drawn) and linked to the cap cell at the tip of the ciliary endings. (B1eB3) Corresponding EM images of longitudinal sections. (B1eB2) The ciliary dilation is visible on these sections (arrow). (B1) The transition zone is clearly visible at the base of the cilia (arrowhead). (B3) The cilia are anchored on a distal basal body (DBB) aligned above the proximal basal body (PBB) at the end of the dendrite. The rootlet anchors the basal body apparatus into the dendrite and the cell body. (C1eC6) Cross sections of a scolopidium cut at different levels of the organ as reported on the scheme (A). (C1eC4) For each panel, a higher representative image of the structure is present on right panels. (C1) Sections of the distal end of the cilium after the ciliary dilation. Only microtubule singlets are observed at this level. (C2) Sections at the level of the ciliary axoneme composed of nine peripheral microtubule doublets. The dynein arms are visible (arrows). Two or three axonemes (neurons) are present in each scolopidia. The differences between scolopidia with two or three neurons are still unclear but could reflect specialized function of subsets of chordotonal neurons (Kamikouchi et al., 2009; Sun et al., 2009; Todi, Sharma, & Eberl, 2004). (C3) Section at the distal transition zone. (C4) Section at the proximal level of the transition zone. A dense ring structure of ninefold symmetry is observed. (C5) Section at the level of the proximal or between both basal bodies. Tight junctions connect the two dendrites. (C6) Section at the base of the basal body showing the connections between the basal body and electron dense structures inside the dendrite. Tight junctions can be observed between the two dendrites.

297

298

CHAPTER 15 Imaging cilia in Drosophila melanogaster

2.3 DATA ANALYSIS Examination of ultrathin sections were performed using a Philips CM120 transmission electron microscope at an accelerating voltage of 80e120 Kv. Images were acquired with a 2k  2k digital camera (ORIUS 200; Gatan) and digital micrograph software. See Figure 6 and legend for representative images and analysis of EM sections of the antennae.

ACKNOWLEDGMENTS We are grateful to Charline Maire, who contributed very efficiently to set up the antennal staining procedure. We thank Je´roˆme Schmitt for Drosophila medium and stock maintenance. Electron microscopy was performed at the Centre Technologique des Microstructures of the University of Lyon (CTmu). This work was supported by grants to B. Durand from the Fondation pour la Recherche Me´dicale (e´quipe FRM, DEQ2000329168), the ANR (Ciliopath-X). J.V. is supported by a doctoral fellowship from the University of Lyon-1. We thank Joe¨lle Thomas for critical reading of the manuscript.

REFERENCES Ashburner, M. (1989). Drosophila Laboratory Manual. Avidor-Reiss, T., Maer, A., Koundakjian, E., Polyanovsky, A., Keil, T., Subramaniam, S., et al. (2004). Decoding cilia function: defining specialized genes required for compartmentalized cilia biogenesis. Cell, 117(4), 527e539. Baker, J. D., Adhikarakunnathu, S., & Kernan, M. J. (2004). Mechanosensory-defective, male-sterile unc mutants identify a novel basal body protein required for ciliogenesis in Drosophila. Development (Cambridge, England), 131(14), 3411e3422. http:// dx.doi.org/10.1242/dev.01229. Bate, M., & Martinez arias, A. (1993). The development of Drosophila melanogaster (Vols I, II). Bechstedt, S., Albert, J. T., Kreil, D. P., Muller-Reichert, T., Go¨pfert, M. C., & Howard, J. (2010). A doublecortin containing microtubule-associated protein is implicated in mechanotransduction in Drosophila sensory cilia. Nature Communications, 1, 11. http:// dx.doi.org/10.1038/ncomms1007. Cachero, S., Simpson, T. I., Lage, Zur, P. I., Ma, L., Newton, F. G., Holohan, E. E., et al. (2011). The gene regulatory cascade linking proneural specification with differentiation in Drosophila sensory neurons. PLoS Biology, 9(1), e1000568. http://dx.doi.org/ 10.1371/journal.pbio.1000568. Cheng, L. E., Song, W., Looger, L. L., Jan, L. Y., & Jan, Y.-N. (2010). The role of the TRP channel NompC in Drosophila larval and adult locomotion. Neuron, 67(3), 373e380. http://dx.doi.org/10.1016/j.neuron.2010.07.004. Elsevier Ltd. Chung, Y. D., Zhu, J., Han, Y., & Kernan, M. J. (2001). NompA encodes a PNS-specific, ZP domain protein required to connect mechanosensory dendrites to sensory structures. Neuron, 29(2), 415e428. Cook, B., Hardy, R. W., McConnaughey, W. B., & Zuker, C. S. (2008). Preserving cell shape under environmental stress. Nature, 452(7185), 361e364. http://dx.doi.org/10.1038/ nature06603.

References

Dubruille, R., Laurenc¸on, A., Vandaele, C., Shishido, E., Coulon-Bublex, M., Swoboda, P., et al. (2002). Drosophila regulatory factor X is necessary for ciliated sensory neuron differentiation. Development (Cambridge, England), 129(23), 5487e5498. Eberl, D. F., & Boekhoff-Falk, G. (2007). Development of Johnston’s organ in Drosophila. The International Journal of Developmental Biology, 51(6e7), 679e687. http:// dx.doi.org/10.1387/ijdb.072364de. Eberl, D., Hardy, R., & Kernan, M. (2000). Genetically similar transduction mechanisms for touch and hearing in Drosophila. Journal of Neuroscience, 20(16), 5981e5988. Effertz, T., Wiek, R., & Go¨pfert, M. C. (2011). NompC TRP channel is essential for Drosophila sound receptor function. Current Biology, 21(7), 592e597. http:// dx.doi.org/10.1016/j.cub.2011.02.048. Elsevier Ltd. Enjolras, C., Thomas, J., Chhin, B., Cortier, E., Duteyrat, J. L., Soulavie, F., et al. (2012). Drosophila chibby is required for basal body formation and ciliogenesis but not for Wg signaling. The Journal of Cell Biology, 197(2), 313e325. http://dx.doi.org/10.1083/ jcb.201109148. Fabian, L., & Brill, J. A. (2012). Drosophila spermiogenesis: big things come from little packages. Spermatogenesis, 2(3), 197e212. http://dx.doi.org/10.4161/spmg.21798. Fujita, S. C., Zipursky, S. L., Benzer, S., Ferru´s, A., & Shotwell, S. L. (1982). Monoclonal antibodies against the Drosophila nervous system. Proceedings of the National Academy of Sciences of the United States of America, 79(24), 7929e7933. Gogendeau, D., & Basto, R. (2009). Centrioles in flies: the exception to the rule? Seminars in Cell and Developmental Biology, 21(2), 163e173. http://dx.doi.org/10.1016/j.semcdb. 2009.07.001. Gong, Z., Son, W., Chung, Y., Kim, J., Shin, D., McClung, C., et al. (2004). Two interdependent TRPV channel subunits, inactive and Nanchung, mediate hearing in Drosophila. Journal of Neuroscience, 24(41), 9059e9066. Go¨pfert, M. C., Humphris, A. D. L., Albert, J. T., Robert, D., & Hendrich, O. (2005). Power gain exhibited by motile mechanosensory neurons in Drosophila ears. Proceedings of the National Academy of Sciences of the United States of America, 102(2), 325e330. http:// dx.doi.org/10.1073/pnas.0405741102. Go¨pfert, M. C., & Robert, D. (2003). Motion generation by Drosophila mechanosensory neurons. Proceedings of the National Academy of Sciences of the United States of America, 100(9), 5514e5519. http://dx.doi.org/10.1073/pnas.0737564100. Gottardo, M., Callaini, G., & Riparbelli, M. G. (2013). The cilium-like region of the Drosophila spermatocyte: an emerging flagellum? Journal of Cell Science, 126(Pt 23), 5441e5452. http://dx.doi.org/10.1242/jcs.136523. Han, Y., Kwok, B., & Kernan, M. (2003). Intraflagellar transport is required in Drosophila to differentiate sensory cilia but not sperm. Current Biology, 13(19), 1679e1686. Hummel, T., Krukkert, K., Roos, J., Davis, G., & Kla¨mbt, C. (2000). Drosophila Futsch/ 22C10 is a MAP1B-like protein required for dendritic and axonal development. Neuron, 26(2), 357e370. Husain, N., Pellikka, M., Hong, H., Klimentova, T., Choe, K., Clandinin, T., et al. (2006). The agrin/perlecan-related protein eyes shut is essential for epithelial lumen formation in the Drosophila retina. Developmental Cell, 11(4), 483e493. Jana, S. C., Girotra, M., & Ray, K. (2011). Heterotrimeric kinesin-II is necessary and sufficient to promote different stepwise assembly of morphologically distinct bipartite cilia in Drosophila antenna. Molecular Biology of the Cell, 22(6), 769e781. http:// dx.doi.org/10.1091/mbc.E10-08-0712.

299

300

CHAPTER 15 Imaging cilia in Drosophila melanogaster

Jan, L., & Jan, Y. (1982). Antibodies to horseradish peroxidase as specific neuronal markers in Drosophila and in grasshopper embryos. Proceedings of the National Academy of Sciences of the United States of America, 79(8), 2700e2704. Jarman, A. P. (2002). Studies of mechanosensation using the fly. Human Molecular Genetics, 11(10), 1215e1218. Jerber, J., Baas, D., Soulavie, F., Chhin, B., Cortier, E., Vesque, C., et al. (2013). The coiledcoil domain containing protein CCDC151 is required for the function of IFT-dependent motile cilia in animals. Human Molecular Genetics, 23(3), 563e577. http://dx.doi.org/ 10.1093/hmg/ddt445. Kamikouchi, A., Inagaki, H. K., Effertz, T., Hendrich, O., Fiala, A., Go¨pfert, M. C., et al. (2009). The neural basis of Drosophila gravity-sensing and hearing. Nature, 458(7235), 165e171. http://dx.doi.org/10.1038/nature07810. Kavlie, R. G., Kernan, M. J., & Eberl, D. F. (2010). Hearing in Drosophila requires TilB, a conserved protein associated with ciliary motility. Genetics, 185(1), 177e188. http:// dx.doi.org/10.1534/genetics.110.114009. Keil, T. A. (1997). Functional morphology of insect mechanoreceptors. Microscopy Research and Technique, 39(6), 506e531. http://dx.doi.org/10.1002/(SICI)1097-0029(19971215) 39:63.0.CO;2-B. Keil, T. A. (2012). Sensory cilia in arthropods. Arthropod Structure and Development, 41(6), 515e534. http://dx.doi.org/10.1016/j.asd.2012.07.001. Elsevier Ltd. Kernan, M. J. (2007). Mechanotransduction and auditory transduction in Drosophila. Pflu¨gers Archiv, 454(5), 703e720. http://dx.doi.org/10.1007/s00424-007-0263-x. Kim, J., Chung, Y., Park, D., Choi, S., Shin, D., Soh, H., et al. (2003). A TRPV family ion channel required for hearing in Drosophila. Nature, 424(6944), 81e84. Laurenc¸on, A., Dubruille, R., Efimenko, E., Grenier, G., Bissett, R., Cortier, E., et al. (2007). Identification of novel regulatory factor X (RFX) target genes by comparative genomics in Drosophila species. Genome Biology, 8(9), R195. http://dx.doi.org/10.1186/gb-2007-8-9-r195. Lee, J., Moon, S., Cha, Y., & Chung, Y. D. (2010). Drosophila TRPN(¼NOMPC) channel localizes to the distal end of mechanosensory cilia. PLoS One, 5(6), e11012. http:// dx.doi.org/10.1371/journal.pone.0011012. Lee, B., & Ashrafi, K. (2008). A TRPV channel modulates C. elegans neurosecretion, larval starvation survival, and adult lifespan. PLoS Genetics, 4(10), e1000213. Lee, E., Sivan-Loukianova, E., Eberl, D. F., & Kernan, M. J. (2008). An IFT-A protein is required to delimit functionally distinct zones in mechanosensory cilia. Current Biology, 18(24), 1899e1906. http://dx.doi.org/10.1016/j.cub.2008.11.020. Liang, X., Madrid, J., Saleh, H. S., & Howard, J. (2011). NOMPC, a member of the TRP channel family, localizes to the tubular body and distal cilium of Drosophila campaniform and chordotonal receptor cells. Cytoskeleton (Hoboken, N.J.), 68(1), 1e7. http://dx.doi.org/ 10.1002/cm.20493. Ma, L., & Jarman, A. P. (2011). Dilatory is a Drosophila protein related to AZI1 (CEP131) that is located at the ciliary base and required for cilium formation. Journal of Cell Science, 124(15), 2622e2630. Martins, A., Machado, P., Callaini, G., & Bettencourt-Dias, M. (2010). Microscopy methods for the study of centriole biogenesis and function in Drosophila. Methods in Cell Biology, 97, 223e242. Mendes Maia, T., Gogendeau, D., Pennetier, C., Janke, C., & Basto, R. (2014). Bug22 influences cilium morphology and the post-translational modification of ciliary microtubules. Biology Open, 3(2), 138e151. http://dx.doi.org/10.1242/bio.20146577.

References

Moore, D. J., Onoufriadis, A., Shoemark, A., Simpson, M. A., Lage, Zur, P. I., de Castro, S. C., et al. (2013). Mutations in ZMYND10, a gene essential for proper axonemal assembly of inner and outer dynein arms in humans and flies, cause primary ciliary dyskinesia. American Journal of Human Genetics, 93(2), 346e356. http://dx.doi.org/10.1016/ j.ajhg.2013.07.009. Moulins, M., & Mill, P. (1976). Ultrastructure of chordotonal organs. Structure and Function of Proprioreceptors in the Invertabrates, 387e426. Newton, F. G., Lage, Z. P. I., Karak, S., Moore, D. J., Go¨pfert, M. C., et al. (2012). Forkhead transcription factor Fd3F cooperates with Rfx to regulate a gene expression program for mechanosensory cilia specialization. Developmental Cell, 22(6), 1221e1233. http:// dx.doi.org/10.1016/j.devcel.2012.05.010. Elsevier Inc. Orgogozo, V., & Grueber, W. B. (2005). FlyPNS, a database of the Drosophila embryonic and larval peripheral nervous system. BMC Developmental Biology, 5, 4. http://dx.doi.org/ 10.1186/1471-213X-5-4. Park, J., Lee, J., Shim, J., Han, W., Lee, J., Bae, Y. C., et al. (2013). dTULP, the Drosophila melanogaster homolog of tubby, regulates transient receptor potential channel localization in cilia. PLoS Genetics, 9(9), e1003814. http://dx.doi.org/10.1371/ journal.pgen.1003814. Peel, N., Stevens, N. R., Basto, R., & Raff, J. W. (2007). Overexpressing centriole-replication proteins in vivo induces centriole overduplication and de novo formation. Current Biology, 17(10), 834e843. http://dx.doi.org/10.1016/j.cub.2007.04.036. Reynolds, E. S. (1963). The use of lead citrate at high pH as an electron-opaque stain in electron microscopy. The Journal of Cell Biology, 17, 208e212. Riparbelli, M. G., Callaini, G., & Megraw, T. L. (2012). Assembly and persistence of primary cilia in dividing Drosophila spermatocytes. Development Cell, 23(2), 425e432. http:// dx.doi.org/10.1016/j.devcel.2012.05.024. Elsevier Inc. Saina, M., & Benton, R. (2013). Visualizing olfactory receptor expression and localization in Drosophila. Methods in Molecular Biology (Clifton, NJ), 1003, 211e228. http:// dx.doi.org/10.1007/978-1-62703-377-0_16. Sarpal, R., Todi, S., Sivan-Loukianova, E., Shirolikar, S., Subramanian, N., Raff, E., et al. (2003). Drosophila KAP interacts with the kinesin II motor subunit KLP64D to assemble chordotonal sensory cilia, but not sperm tails. Current Biology, 13(19), 1687e1696. Senthilan, P. R., Piepenbrock, D., Ovezmyradov, G., Nadrowski, B., Bechstedt, S., Pauls, S., et al. (2012). Drosophila auditory organ genes and genetic hearing defects. Cell, 150(5), 1042e1054. http://dx.doi.org/10.1016/j.cell.2012.06.043. Elsevier Inc. Shaham, S. (2009). Chemosensory organs as models of neuronal synapses. Nature Reviews Neuroscience, 11(3), 212e217. http://dx.doi.org/10.1038/nrn2740. Soulavie, F., Piepenbrock, D., Thomas, J., Vieillard, J., Duteyrat, J.-L., Cortier, E., et al. (2014). hemingway is required for sperm flagella assembly and ciliary motility in Drosophila. Molecular Biology of the Cell, 25(8), 1276e1286. http://dx.doi.org/ 10.1091/mbc.E13-10-0616. Sun, Y., Liu, L., Ben-Shahar, Y., Jacobs, J. S., Eberl, D. F., & Welsh, M. J. (2009). TRPA channels distinguish gravity sensing from hearing in Johnston’s organ. Proceedings of the National Academy of Sciences of the United States of America, 106(32), 13606e13611. http://dx.doi.org/10.1073/pnas.0906377106. Todi, S. V., Sharma, Y., & Eberl, D. F. (2004). Anatomical and molecular design of the Drosophila antenna as a flagellar auditory organ. Microscopy Research and Technique, 63(6), 388e399. http://dx.doi.org/10.1002/jemt.20053.

301

302

CHAPTER 15 Imaging cilia in Drosophila melanogaster

Uga, S., & Kuwabara, M. (1965). On the fine structure of the chordotonal sensillum in antenna of Drosophila melanogaster. Journal of Electron Microscopy (Tokyo), 14, 173e181. Walker, R. G., Willingham, A. T., & Zuker, C. S. (2000). A Drosophila mechanosensory transduction channel. Science (New York, NY), 287(5461), 2229e2234. Wolff, A., de Ne´chaud, B., Chillet, D., Mazarguil, H., Desbruyeres, E., Audebert, S., et al. (1992). Distribution of glutamylated alpha and beta-tubulin in mouse tissues using a specific monoclonal antibody, GT335. European Journal of Cell Biology, 59(2), 425e432.