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Methods and Protocols for Studying

Cell Death in Drosophila

Donna Denton, Kathryn Mills, and Sharad Kumar

Contents

1. Introduction 18

1.1. Drosophila as a model system to study apoptosis 18

1.2. Cell death pathway 20

1.3. Protocols described 21

2. Cell Death Analysis During Development 22

2.1. Fixation of samples 22

2.2. Detection of apoptotic cells with TUNEL 24

2.3. Acridine orange staining 25

2.4. Immunostaining: Caspase-3 and cytochrome c 26

2.5. g-Irradiation–induced apoptosis 27

3. Biochemical Analysis 27

3.1. Cell culture 27

3.2. Measurement of caspase activity by use of synthetic

peptide substrates 29

3.3. Immunoblotting cleaved substrates 31

4. Genetic Analysis 31

4.1. Genetic dissection of cell death pathway 31

References 35

Abstract

Drosophila melanogaster is a highly amenable model system for examining

programmed cell death during animal development, offering sophisticated

genetic techniques and in vivo cell biological analyses. The reproducible pat-

tern of apoptosis, as well as the apoptotic response to genotoxic stress, has

been well characterized during Drosophila development. The main cellular

components required for cell death are highly conserved throughout evolution.

Central to the regulation of apoptosis is the caspase family of cysteine pro-

teases, and studies in Drosophila have revealed insights into their regulation

and function. This chapter describes protocols for detecting apoptotic cells

in Enzymology, Volume 446 # 2008 Elsevier Inc.

076-6879, DOI: 10.1016/S0076-6879(08)01602-9 All rights reserved.

Institute, Institute of Medical and Veterinary Science, Adelaide, Australia

17

18 Donna Denton et al.

during Drosophila development, as well as the use of Drosophila cell lines.

Commonly used methods for detecting apoptosis are described, including

TUNEL, acridine orange, and immunostaining with specific components of the

apoptotic pathway such as active caspases. A crucial step in the induction of

apoptosis is caspase activation and cleavage, which can be measured by use of

fluorogenic peptide substrates or detection of cleaved protein products by

immunoblotting, respectively. In addition, one of the advantages of the use of

Drosophila as model is the ability to examine genetic interactions with various

components of the cell death pathway.

1. Introduction

Programmed cell death (PCD) or apoptosis is an essential process fornormal animal development and can be distinguished by morphologicallydistinct characteristics, including cellular shrinkage, membrane blebbing,and nuclear DNA fragmentation (reviewed in Baehrecke, 2002). This formof cell death is distinct from necrosis which, as a result of cellular injury,culminates in cell swelling and lysis. During development, the balancebetween cell proliferation, differentiation, and death is critical. Apoptosisis vital to remove unnecessary or excess cells during tissue pattern formationand also in maintaining adult homeostasis. In addition, apoptosis eliminatesdamaged or abnormal cells such as those subjected to DNA damage or thoseinfected with pathogens. Misregulation of apoptosis can have severe effectson the organism and can lead to various developmental abnormalities ordiseases including cancer.

1.1. Drosophila as a model system to study apoptosis

As the main components of the cell death machinery are highly evolution-arily conserved, Drosophila is an exceptional model system for examiningapoptosis during animal development. In addition to its well-characterizeddevelopmental program and complete genome sequence, Drosophila offerssophisticated genetic techniques and in vivo cell biological analyses that arenot possible in other systems (Adams et al., 2000; Richardson and Kumar,2000). The life cycle of Drosophila consists of a series of developmentalstages: embryo, larvae (three larval instar stages), pupae, and adult flies. Thesteroid hormone ecdysone plays a significant role in controlling and coor-dinating progression of the animal through the major developmental stages.Increases in ecdysone titer initiate the onset of the larval molts, late larvalwandering behavior, pupariation, pupation, and adult development(Fig. 2.1). Modulation of ecdysone can lead to severe developmental con-sequences arising from perturbations to apoptosis (reviewed in Baehrecke,2000).

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Figure 2.1 The steroid hormone ecdysone regulates morphogenic events. Increases inecdysone titers initiate the onset of the larval molts, late larval wandering behavior,pupariation, pupation, and adult development. The late larval and prepupal ecdysonepulses correspond to histolysis of obsolete larval tissues. Midgut (MG) death canbe detected at 120 h AED (0 h RPF), and 15 h later salivary gland (SG) death can bedetected.Modified fromRiddiford,1993.

Studying Cell Death in Drosophila 19

The reproducible pattern of apoptosis during Drosophila developmenthas been well characterized (Abrams et al., 1993). As early as 6 h after eggdeposition (AED), apoptosis can be detected in a small number of cells.After this, apoptotic cells become more widespread, and at later stages ofembryogenesis an increase in apoptotic cells can be identified in tissues suchas the central nervous system. During metamorphosis, a dramatic increase inPCD is observed when obsolete larval tissues are removed in response to thedevelopmental pulses of ecdysone (reviewed in Baehrecke, 2000; Kumarand Cakouros, 2004) (Fig. 2.1). The larval midgut is removed by apoptosisin response to the large ecdysone pulse during the late larval stage, and theadult gut begins to develop in its place. Approximately 12 h later, anotherecdysone pulse signals the larval salivary glands to undergo histolysis. Cor-rect patterning of the adult eye during the pupal stage also requires devel-opmentally regulated PCD. The interommatidial cells of the pupal eye discare rearranged to form a single layer around the photoreceptor clusters, withthe excess interommatidial cells eliminated by apoptosis (reviewed inBrachmann and Cagan, 2003). During normal oocyte development, nursecells undergo PCD after they deposit their cytoplasmic contents into the

20 Donna Denton et al.

developing oocyte. Another role for apoptosis during oogenesis is toremove defective egg chambers unable to develop into fertile eggs(reviewed in Buszczak and Cooley, 2000). Unlike PCD during embryo-genesis or metamorphosis, upstream components of the apoptotic pathwayare not required for nurse cell death. Thus in Drosophila, the regulation ofPCD can occur differently in specific tissues during development.

In addition to developmental PCD, apoptosis can also be induced inresponse to DNA damage induced by genotoxic stress such as ionizingradiation. This can be observed in embryos and in the proliferating larvalimaginal disc tissue that undergoes cell death after irradiation. In wingimaginal discs an increase in cell death is observed 4 to 6 h after irradiationto remove damaged cells (Wichmann et al., 2006).

1.2. Cell death pathway

Central to apoptosis is the highly conserved caspase family of cysteineproteases. These enzymes are produced as inactive zymogens and, uponactivation after death-inducing signals, cleave multiple cellular proteinsresulting in apoptosis (reviewed in Hay and Guo, 2006; Kumar, 2007).Caspases can be grouped into two classes: ‘‘initiator’’ caspases contain longprodomains and act as signal transducers that are required to cleave andactivate the downstream ‘‘effector’’ caspases that subsequently orchestratecleavage of cellular substrates and dismantling of the cell. The Drosophilagenome encodes seven caspases, three initiator (Dronc, Dredd, and Strica),and four effector caspases (Drice, Dcp-1, Decay, and Damm) (Chen et al.,1998; Dorstyn et al., 1999a,b; Doumanis et al., 2001; Fraser and Evan, 1997;Harvey et al., 2001; Song et al., 1997). Dronc is the only caspase recruitmentdomain (CARD) containing caspase in Drosophila and is essential for devel-opment, unlike the other initiator caspases. dronc null mutants are pupal lethaland display various cell death defects in embryos, larvae, and prepupae, aswell as defects in response to stress-induced apoptosis (Chew et al., 2004;Daish et al., 2004). Although strica mutants are viable, strica seems to play aredundant role with dronc in PCD during oogenesis (Baum et al., 2007).The other initiator caspase, Dredd, is not essential for developmental PCDbut is required for innate immune response (Leulier et al., 2000). The effectorcaspase drice is required for developmental and stress-induced apoptosis andmutants are pupal lethal with approximately 20% escapers (Kondo et al., 2006;Muro et al., 2006; Xu et al., 2006).Dcp-1 plays a relatively minor role in PCD,required during mid-oogenesis, and overlaps with drice function (Kondo et al.,2006; Muro et al., 2006; Xu et al., 2006). decay mutants are viable andfertile with normal developmental PCD (Kondo et al., 2006), whereas a rolefor Damm in PCD has not been established, because no specific mutantsare available. Thus, Dronc is believed to be the predominant initiator caspase

Ark

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Drice

Apoptosis

DIAP1RHG

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Figure 2.2 The main caspase activation pathway in Drosophila.On death stimulus theRHG family of proapoptotic proteins bind to DIAP1, which is then degraded by anubiquitination-dependent mechanism.This allows released Dronc to become activatedafter bindingwithArk and activate downstream effector caspaseDrice.

Studying Cell Death in Drosophila 21

andDrice the main effector caspase required for both developmental PCD andstress-induced apoptosis (Fig. 2.2).

Activation of Dronc requires association with the adaptor protein Ark(Apaf-1 related killer), and similar to dronc, ark is an essential gene withmutants showing severe defects in developmental PCD and stress-inducedapoptosis (Mills et al., 2006; Srivastava et al., 2006). Until the appropriatedeath signal is received, Dronc is maintained in an inactive form by associa-tion with DIAP1, the Drosophila member of the inhibitor of apoptosisprotein (IAP) family (reviewed in Kumar, 2007; Mills et al., 2005)(Fig. 2.2). A loss-of-function mutation in the gene encoding DIAP1, thread,is embryonic lethal as a result of an increase in cell death during embryo-genesis (Hay et al., 1995). In Drosophila, initiation of cell death requires theproducts of the reaper (rpr), grim and head involution defect (hid/Wrinkled)genes, as deletion of all three genes shows an absence of cell death duringembryogenesis (White et al., 1994). In addition to Rpr, Grim, and Hid,other cell death regulators include Sickle and Jafrac2 and are collectivelyreferred to as the RHG proteins (Fig. 2.2). This group of proteins acts toinitiate cell death by binding to and sequestering DIAP1 from interactingwith caspases (Hay and Gao, 2006; Kumar, 2007).

1.3. Protocols described

This chapter describes methods used to examine apoptosis in Drosophila. Thefirst section describes protocols to examine cell death during development andincludes methods for detecting apoptosis in situ by DNA fragmentation withTUNEL and by use of the vital dye acridine orange (AO) in live tissue.

22 Donna Denton et al.

Another in situ approach includes immunostaining with specific markers ofapoptosis such as antibodies to active caspase-3. The second section detailsbiochemical approaches. A crucial step in the induction of apoptosis is caspaseactivation and cleavage, which can bemeasured by use of synthetic peptides ordetection of cleaved substrates by immunoblotting. In addition to detectingapoptosis in whole animals, a variety of Drosophila cell lines can also be used.The third section illustrates one of the great advantages of the use ofDrosophilaas model to examine apoptosis by the ability to test genetic interactions byexpression of components of the cell death pathway in the animal.

2. Cell Death Analysis During Development

The cellular basis of Drosophila development, including the reproduc-ible patterns of PCD, is well described (Abrams et al., 1993). In addition,differential regulation of PCD has been characterized in the embryo, larvae,and adult (ovaries). The protocols described in this section include TUNEL,acridine orange staining, and immunostaining requiring either fixed orlive samples. Different tissues have different requirements for fixation, asdescribed in the following.

2.1. Fixation of samples

2.1.1. Collection and fixation of embryosFlies can be set to lay in ‘‘lay tubes’’ on top of grape juice agar plates (seebelow) with a small amount of yeast paste. We use plastic tubing sealed atone end with fine wire mesh as lay tubes. The embryos are then harvestedfrom the agar plate by washing with water and brushing with a small paintbrush into a collection basket (made by placing fine wire mesh over a cutoff1.5-ml Microfuge tube or by use of cell strainers available from BD Falcon).The chorion is then removed by use of a pipette to continually rinseembryos with a 50% bleach solution (domestic brand) for 3 min. Thedechorionated embryos are washed thoroughly with water and transferredto a tube containing a two-phase mix of equal parts 4% formaldehyde in 1�HEN buffer (10 � HEN buffer: 1 M HEPES, 0.5 M EGTA, 0.1% NP-40,pH to 6.9, and filter sterilize) and heptane. The sample is then shaken for20 min on an orbital platform such that the interface between the liquidphases is disrupted and the embryos are in an emulsion. After fixation, theembryos will settle on the interface and the bottom (aqueous) phase can beremoved and replaced with an equal volume of methanol. Shake vigorouslyfor 30 s to remove the vitelline membrane and let stand for 1 min. Devi-tellinized embryos will sink from the interface and can be collected from thebottom by use of a cut-off pipette tip (discard embryos with vitelline

Studying Cell Death in Drosophila 23

membrane as they will not stain). Embryos are then rinsed in methanol atwhich point they can be stored at �20 �C in ethanol. Before staining,embryos are rehydrated by serial incubation in 75, 50, and 25% (v/v)methanol in PBT for 5 min each and then washed with PBT.

Note: As the strength of bleach can vary, dechorionation can bemonitored by use of a dissection microscope until 80% of the dorsalappendages have dissolved. It is important not to overfix embryos, becausethis results in decreased efficiency of devitellinization. If cloudiness orprecipitation is present in the 10 � HEN solution, the NP-40 can beomitted. The addition of heptane in the fixative is essential to create holesin the vitelline membrane. Overnight storage in methanol often improvesimage quality, however, extended periods of storage (more than a month)can result in loss of image quality.

2.1.1.1. Grape juice agar plates0.3% agar, 25% grape juice, 0.3% sucrose, 0.03% tegosept (10% para-hydroxybenzoate in ethanol). Combine agar, grape juice, sucrose, andwater together and boil by heating in a microwave. Mix and to cool to60 �C before adding tegosept and pouring into petri dishes (appropriatelysized to fit the lay tube). Once the plates have set, they can be stored at 4 �C.

2.1.2. Staging and fixation of larval tissuesTo stage larvae for analysis of midgut (MG) and salivary gland (SG) histoly-sis, larvae can be propagated on food containing 0.05% bromophenol blue.As larvae commence the wandering behavior, they cease feeding and beginemptying their gut contents. Therefore, their age relative to pupariation,when the cuticle hardens and forms a pupal case, can be estimated on thebasis of gut clearance (Andres and Thummel, 1994; Maroni and Stamey,1983). Wandering larvae are collected from the side of the vial with a wetpaint brush and transferred to a petri dish lined with wet Whatmann paper.Animals can be monitored regularly (every 15, 30, or 60 min) for gutclearance or pupariation. Larvae with dark blue guts represent �24 to�12 h relative to pupariation (RPF), light blue guts �12 to �5 h RPF,and clear guts �6 to �1 h RPF. Immediately before pupariation, the larvaeshorten, evert spiracles, and stop moving. During the first 1 h after puparia-tion, the cuticle appears white, after which the cuticle tans. This coloringcan be used to stage animals at 0 toþ1 h RPF and is also referred to as whiteprepupae (WPP) (Andres and Thummel, 1994; Maroni and Stamey, 1983)(see Fig. 2.1). At the desired stage, animals are transferred to another petridish lined with wet Whatmann paper and aged until the desired time forexperimental analysis. For analysis of apoptosis in the MG, we use clear gutlarvae (�6 to �1 h RPF) and WPP. For SG analysis we age animals to 14 hRPF that is equivalent to 2 h after head eversion (AHE). Head eversionmarks the transition between prepupal and pupal stages. During this stage,

24 Donna Denton et al.

abdominal muscles contract, pushing an air bubble from the posterior to theanterior end, resulting in a space for the head to evert. This process occursrapidly in approximately 10 min. Dissect tissue in 1� PBS and fix in 4%formaldehyde in PBTW (0.1% Tween-20 in PBS) for 20 min at roomtemperature. Wash twice with PBTX (0.1% TritonX-100 in PBS).

2.1.3. Fixation of ovariesTo ensure ovaries are well developed and all stages represented, females areaged at 25 �C for 3 to 7 days in uncrowded conditions on food supplemen-ted with fresh yeast in the presence of an equal number of males to stimulateegg production. Dissect females in PBS in a depression plate by use of fineforceps to hold the submerged fly between the thorax and abdomen and asecond pair of forceps to hold the external genitalia and pull to releaseovaries into PBS. Remove any debris from the ovary and tease apartovarioles by use of a fine needle. Transfer ovarioles into a 1.5-ml tubecontaining a two-phase fix solution of equal parts heptane (400 ml) and 4%formaldehyde in PBS (400 ml) and shake for 25 min at room temperature.Remove heptane/fix and wash twice with PBT, ensuring removal of allheptane.

Note: SylgardÒ 184 Silicone Elastomer Kit (Dow Corning) can be usedaccording to manufacturers instructions to make dissection dishes with a softsilicone base, which are useful to prevent dissecting forceps and needlesfrom damage.

2.2. Detection of apoptotic cells with TUNEL

A frequently used method for detecting apoptosis by DNA fragmentation isterminal deoxynucleotidyl transferase (TdT)–mediated dUTP nick-endlabeling (TUNEL) staining and has been used routinely on whole-mounttissue samples (White et al., 1994). This technique specifically detectsapoptotic cells by preferentially labeling DNA strand breaks. Several com-mercial kits are available and should be used according to manufacturersprotocols. Both fluorescent and color detection kits are available (extralabeling steps are required for color detection) and offer reliable results.

After fixation (appropriate for your tissue of interest), wash twice withPBTX (0.1% TritonX-100) then twice with PBTX5 (0.5% TritonX-100)and permeabilize in 100 mM sodium citrate in PBTX at 65 �C for 30 min.Wash three times with PBTX5 before addition of TUNEL mix according tothe manufactures instructions and incubate at 37 �C for 3 h on a rotatingplatform. Wash three times with PBTX and store in 80% glycerol in PBS.To mount samples place a strip of double-sided tape across each end of theslide, placing samples between the tape, dissect the midgut or tissue ofinterest away from remaining tissue. Gently place a coverslip restingon the tape over samples. Examine samples by epifluorescence or confocalmicroscopy.

Studying Cell Death in Drosophila 25

Whilst TUNEL staining produces highly reproducible results, it is advis-able to set up both positive and negative controls. For TUNEL staining, aDNase-treated sample will serve as a positive control, and negative controlcan be performed by the addition of water instead of the active enzyme TdT.

Note: Ensure the sodium citrate solution is made up fresh. If difficulties inlabeling are encountered, an alternative permeabilization to increase pene-trance can be achieved by proteinase K treatment (10 mg/ml in PBS for 3 to5 min at room temperature) (Arama and Stellar, 2006; McCall and Peterson,2004). Antibody staining can be performed sequentially in combinationwith TUNEL staining. Samples can also be co-stained to detect nuclei, byuse of Hoechst 33258 at 4 mg/ml in PBS for 1 min before storage in 80%glycerol in PBS. When assaying for cell death after irradiation, it is prefera-ble to use acridine orange instead of TUNEL, because TUNEL can (inprinciple) detect the DNA damage induced by the treatment.

2.3. Acridine orange staining

The vital dye acridine orange (AO) can be used to observe apoptosis in livetissue (Arama and Steller, 2006). The advantage of AO compared withTUNEL staining is the speed of the staining. However, because it is carriedout on live tissue, multiple labeling cannot be performed, and the tissuemust be examined immediately. A stock solution of AO can be made up inethanol (1 mM stock: 1.85 mg AO in 5 ml ethanol) and stored in the dark atroom temperature for several months. The AO stock solution is thendiluted to the final concentration in PBS immediately before use.

2.3.1. Embryo stainingAfter egg collection and dechorionation, transfer embryos to a tube con-taining equal volume of heptane and 5 mg/ml AO in PBS. Shake vigorouslyfor 5 min at room temperature to generate a fine emulsion. Allow the liquidphases to separate and transfer the embryos from the interface onto a glassmicroscope slide in a drop of PBS.

2.3.2. Staining larval tissueFrom appropriately aged larvae dissect tissue of interest away from othertissue. Incubate the dissected tissue in a drop (20 to 30 ml) of 5 mg/ml AOsolution for 5 to 15 min.Wash briefly by transferring tissue to a fresh drop ofPBS. Mount in a drop of PBS on a glass microscope slide.

2.3.3. Ovary stainingTransfer dissected ovarioles into a 1.5-ml tube containing equal volumes ofheptane and 10 mg/ml AO in PBS. Mix gently and rotate for 5 min. Allowovarioles to sink to bottom and replace AO solution with PBS to rinsebriefly. Transfer ovarioles to a slide and gently spread out.

26 Donna Denton et al.

To prevent damage and squashing of samples, double-sided tape can bepositioned across each end of the microscope slide to support the coverslip.Examine samples immediately by epifluorescence/confocal microscopy.The best sensitivity can be observed by use of the green channel(522 nm); however, the red channel (568 nm) can be used to provide lessbackground (Arama and Steller, 2006).

In addition to the test samples, positive and negative control samplesshould be examined. Comparisons can be made between tissues from wild-type animals at different stages of development, depending on the level ofapoptosis at that specific stage.

Note: Ensure no detergent is present in dissection and AO stainingsolutions, because this will abolish staining. Additional protocols for AOand TUNEL staining in embryos and testis can be found in Arama andSteller (2006). Halocarbon oil can be used as an alternative mountingsolution if PBS evaporates too quickly.

2.4. Immunostaining: Caspase-3 and cytochrome c

Antibodies that specifically recognize active caspases can be used to detectapoptotic cells in vivo. Various tissue and developmental stages can beanalyzed and require fixation (as described previously), before incubationwith a primary antibody. Of particular use is the antibody against the activeform of caspase-3, which has been shown to cross-react with Drosophilaeffector caspases (anti-active caspase-3, Cell Signaling Technology). In addi-tion, the anti-cytochrome c antibody that recognizes an altered configurationof the protein can be used to specifically label dying cells (Varkey et al., 1999).However, it should be noted that, unlike inmammalian cells, cytochrome c isnot generally released during apoptotic signaling inDrosophila cells and is notrequired for caspase activation (Dorstyn et al., 2002; 2004).

After fixation, block sample in 5 to 10% normal goat sera for 1 h at roomtemperature in PBT then incubate in primary antibody diluted in blockingsolution overnight at 4 �C with gentle shaking. Remove the primaryantibody and wash three times with PBT over an hour. Longer washesand more rinses will result in less background. Add fluorescently labeledsecondary antibody, also diluted in blocking solution, and incubate at roomtemperature for 1 h. Once the fluorescent secondary has been added,samples should be kept in the dark. Remove the secondary antibody andwash in PBT as described previously. Store samples in 80% glycerol in PBSbefore mounting for microscopy.

Note: Alternative blocking solutions such as 1% BSA in PBT can be used,depending on the primary antibody. Samples can also be co-stained to detectnuclei (as described). Alexa Fluor–conjugated secondary antibodies are avail-able in a wide range of fluorescence emission wavelengths (we commonly useAlexa Fluor 488 and 568) and offer increased intensity and photostability

Studying Cell Death in Drosophila 27

compared with others products. If photobleaching occurs, commerciallyavailable anti-fade products can be used as the mounting medium.

2.5. g-Irradiation–induced apoptosis

Induction of apoptosis can be observed after genotoxic stress in proliferatinglarval imaginal tissue. Wandering third-instar larvae are collected andexposed to varying doses of g-irradiation, usually ranging from 20 up to40 Gy, by use of a 137Cs source (Kondo et al., 2006). After irradiation, larvaeare allowed to recover at 25 �C for an appropriate length of time beforedissection and staining of tissue. A dose response should be determined and anonirradiated control included.

Note: Alternative sources such as ultraviolet irradiation and X-irradiationmay also be used, however, suitable doses and recovery times should beoptimized to suit the equipment used.

3. Biochemical Analysis

The protocols described in this section include the use of cell lines toanalyze the apoptotic response to death stimuli, measurement of caspaseactivity by cleavage of synthetic peptide substrates, and detection ofcaspase processing by immunoblotting with specific antibodies.

3.1. Cell culture

Drosophila cell culture can be useful to complement animal studies and,consistent with Drosophila development, central death activators induce celldeath in cultured cells. Various Drosophila cell lines are available from theDrosophila Genomics Resource Center (DGRC), and general protocols canbe found on their web site (https://dgrc.cgb.indiana.edu/). Commonlymaintained lines include SL2, mbn2, BG2 and Kc cells.

3.1.1. Maintaining cell linesDrosophila cell lines are grown in Schneider’s cell medium supplementedwith penicillin and streptomycin, 10% fetal bovine serum, and 1% gluta-mine. Some cell lines require additional growth factors, for example BG2cells are supplemented with insulin.

The advantage of SL2 and BG2 cells is that they are adherent, fastgrowing, relatively robust, and good for localization studies. Whereas othercells such as mbn2 are responsive to ecdysone and are, therefore, useful foranalyzing ecdysone responsive cell death pathways. Several different vectorsare available for inducible or constitutive expression and can be obtained

28 Donna Denton et al.

from the DGRC or other Biotech companies. The inducible vector,pRmHa-3, contains a metallothionein promoter that is induced by theaddition of CuSO4 to the cell culture medium. Vectors for constitutiveexpression are also available. We routinely use pIE (Novagen), however,others that also contain epitope tags are available.

3.1.2. Transient transfectionsTransient transfections can be performed with CellfectinÒ (Invitrogen)transfection reagent according to manufacturers guidelines with a resultanttransfection efficiency of approximately 20%. Plate exponentially growingcells at 6 � 105 cells/ml in six-well plates (or 35-mm plates) in Schneider’smedium and leave for 1 h to overnight for cells to adhere. Prepare thetransfection solutions, by diluting 1 to 2 mg DNA into 100 ml of mediawithout serum and antibiotics and dilute 1.5 to 9 ml of CellfectinÒ into100 ml of media without serum and antibiotics. Combine the two solutions,mix gently, and incubate at room temperature for 20 to 30 min. Removethe media from the plated cells and wash once with media without serumand antibiotics. To the transfection mix add 0.8 ml serum and antibiotic-free media, mix gently, and add to the cells after the removal of the washmedia. Incubate cells at 27 �C for 5 h. Add 2 ml of complete media contain-ing serum and antibiotics. Incubate cells at 27 �C for the desired experi-mental time, usually 24 to 48 h. If an inducible expression system is used,after 24 to 48 h incubation, add 0.7 mM CuSO4 to induce expression andincubate for the desired experimental time. Be sure to include a sample thathas not been treated with CuSO4 as noninduced control. Cells are harvestedeither by scraping by use of a commercial sterile cell scraper or by pipetting.

3.1.3. Viability assaysCell viability can be assessed by use of the Trypan blue exclusion assay.Trypan blue is taken up by dying cells, and the percentage viable cells can beestimated by counting the number of Trypan blue–negative (unstained)cells compared with Trypan blue–positive (blue) cells. Alternately, cells canbe stained with DAPI (2 mg/ml in methanol) and scored on the basis ofnuclear morphology. The nuclei of apoptotic cells appear condensed, and atlater stages apoptotic nuclear membrane blebbing is visible. To score trans-fected cells for cell death, cells can be co-transfected with a b-gal or GFPreporter vector and morphology of b-gal or GFP-positive transfected cellsscored by microscopy.

Note:Drosophila cells can be susceptible to low cell density and variationsin batches of media. The same transfection protocol can be used to generatestable cell lines by use of vectors with appropriate selection.

Studying Cell Death in Drosophila 29

3.2. Measurement of caspase activity by use of syntheticpeptide substrates

A frequently used method to detect caspase activity after the induction ofapoptosis is measuring the cleavage of synthetic substrates that have beenincubated with cell lysates. The technique was originally described byPennington and Thornberry (1994) and has been further developed witha variety of synthetic substrates becoming commercially available. Thesynthetic peptides are conjugated to a fluorochrome, such as 7-amino-4-methylcoumarin (AMC) (other fluorometric or colorimetric substratesare available). The basis of the assay is that the peptide substrate is cleaved bycaspases that recognize the substrate cleavage site, releasing the fluoro-chome. The intensity of the fluorescent signal measured by a spectropho-tometer is proportional to the amount of cleaved substrate, which dependson caspase activity from the apoptotic cells present in the cell population.

3.2.1. Preparation of protein lysatesProtein lysates can be prepared from appropriately staged whole animals,dissected tissue samples, or tissue culture cell lysates. The sample is homo-genized in lysis buffer (below) by use of a pestle in a 1.5-ml Microfuge tubeand subjected to three rounds of freezing in liquid nitrogen and thawing.Debris is removed by spinning the extracts at 13,000 rpm for 20 min at 4 �Cthen transferring the supernatant to a fresh tube. After determining proteinconcentration, the lysates can be used in caspase activity assays as describedbelow or can be stored at �70 �C in aliquots for several months.

Note: Samples can be snap frozen and stored at �70 �C before prepara-tion of lysates. The volume of lysis buffer used can vary, depending on theprotein yield of the sample. As a guide, we use 10 ml per whole animal or5 ml per dissected tissue.

3.2.1.1. Lysis buffer 50 mM HEPES, pH 7.5; 100 mM NaCl; 1 mMEDTA; 0.1% CHAPS; 10% sucrose; 5 mM DTT; 0.5% TritonX-100; 4%glycerol; 1� protease inhibitor cocktail (Complete Roche).

3.2.2. Measurement of caspase activityCleavage of the caspase peptide substrates VDVAD-AMCandDEVD-AMCcan be used to determine enzyme activity, because they show the highestlevel of activity for Drosophila caspases. Caspase activity assays should be setup in triplicate with 20 to 50 mg of protein lysate incubated with 100 mM ofVDVAD-AMC or DEVD-AMC in a final volume of 100 ml made up withlysis buffer. The assay should also include positive and negative controlsamples, as well as a fluorescence calibration standard. Fluorescence is quan-tified by use of a spectrophotometer (excitation, 385 nm; emission, 460 nm)over a time course of up to 5 h at 37 �C. The fluorescence standard curve is

30 Donna Denton et al.

obtained by plotting the relative fluorescence of the calibration standardover the concentration, and the slope of the line is used as the conversionfactor (1/slope) when performing the caspase assay. The relative fluorescenceunits for each sample can be calculated by plotting the linear region of thechange in fluorescence over time, whereby the slope of the line canbe determined. This is then used with the conversion factor to calculate theactivity expressed as pmol/min (Fig. 2.3).

Note: This method of detecting caspase activity is widely used, because itis quantitative, relatively quick, and sensitive. Even though different peptidesubstrates are available to distinguish caspase activities, there is redundancyin the sequence specificity of caspases for substrates. That is, several caspaseswill cleave the same substrates, although with different efficiency, so thepeptide sequences should be viewed as preferred substrate motifs and ratherthan absolute (Berger et al., 2006).

05000

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Figure 2.3 Determining the specific activity for cleavage of the synthetic fluorogenicsubstrate. (A)Example of rawdataof the substrate cleavage activityexpressed as fluores-cence units and the change over time. (B) Plot of the linear region of the change in fluo-rescence over time and use an appropriate linear regression program to obtain the slopeof the line (m). (C) Calculations used to determine the activityof the samples expressedas pmol substrate cleaved permin. (D) Plot of the substrate cleavage activity is expressedin pmol/min.

Studying Cell Death in Drosophila 31

3.3. Immunoblotting cleaved substrates

The activation of caspases coincides with the cleavage of a caspase precursor,therefore, the appearance of large and small subunits can be used as a markerfor processing and activation of individual caspases. Cell extracts can beanalyzed by immunoblotting with antibodies to detect cleaved caspase sub-units. The standard protocols for immunoblot analysis can be used(Sambrook and Russell, 2001). The protein lysates (as made previously forcaspase assay) can be prepared for electrophoresis by mixing with an equalvolume of 2� protein loading buffer. The proteins are separated by sodiumdodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and trans-ferred to a membrane (nitrocellulose or polyvinylidene difluoride, PVDF).The membrane is blocked with 5% dry milk powder dissolved in phosphate-buffered saline (PBS) and probed with a primary antibody against thecaspase of interest. Useful antibodies include anti-Dronc (Dorstyn et al.,2002; Quinn et al., 2000) and anti-Drice (Dorstyn et al., 2002). The primaryantibody can be detected by use of horseradish peroxidase (HRP) or alkalinephosphatase (AP)-conjugated secondary antibodies and visualized byenhanced chemiluminescence (ECF) substrate or enhanced chemifluores-cence ECF substrates.

4. Genetic Analysis

4.1. Genetic dissection of cell death pathway

Genetic interaction screens have been commonly used to identify genes thatmodify a phenotype caused by misexpression of a gene of interest in theDrosophila eye. This type of screen is also particularly useful in determiningthe order of new components within a biological pathway. The followingsection is designed to give an introduction into the principles of examininggenetic interactions in the Drosophila adult eye.

The Drosophila adult compound eye is an excellent tissue to examinegenetic interactions, because it is not essential for viability and is easy toexamine (reviewed in Thomas and Wassarman, 1999). The compound eyeconsists of approximately 800 units called ommatidia in a precise array and isderived from a single layer epithelium, the eye imaginal disc, that is set asideduring embryogenesis and proliferates during larval development increasingseveral fold in size (Wolff and Ready, 1991a). Differentiation of the eye discinitiates during the third-instar larval stage and requires cell death as part ofits normal development, and perturbations to this process are visible in theadult eye (Wolff and Ready, 1991b). Several components of the cell deathpathway, when expressed in the eye imaginal disc, perturb eye developmentand result in a disorganized or rough adult eye (see below).

32 Donna Denton et al.

The GAL4/UAS system is a powerful tool in Drosophila for drivingexpression of a gene of interest by specific promoters (Brand and Perrimon,1993; reviewed in Duffy, 2002). It is a two-component systemwhereGAL4expression is driven by tissue-specific enhancers, and the presence of GAL4results in transcriptional activation from the GAL4 binding sites, UAS,driving a gene of interest (Fig. 2.4). The GMR (glass multimer reporter)promoter drives expression of GAL4 in the posterior region of the eyeimaginal disc (Hay et al., 1994) (GMRGAL4 is available from BloomingtonDrosophila Stock Center). The expression of several apoptotic regulators inthe eye has been characterized such as the baculovirus caspase inhibitor p35,inwhichGMRp35 results in adults with a rough eye because of the survival ofcells (supernumerary retinal cells) that would normally be lost (Hay et al.,1994). Conversely, expression of the apoptotic activator hid in the eye,

UAS Target gene

UAS Target gene

UAS Interacting gene

GMR GAL4

GMR GAL4

GMR GAL4

Control

Rough eye

Suppressor or enhancer

Figure 2.4 The GAL4/UAS system used to examine genetic interactions in the adulteye. The GMR enhancer is used to drive eye-specific expression of GAL4. The targetline containing theUAS sites is crossed to flies containingGMRGAL4, enabling controlof target gene expression, producing a rough eye phenotype.Testing for genetic interac-tion is achieved by co-expression from the UAS sites present in both the target and thetest interactor gene.

Studying Cell Death in Drosophila 33

GMRhid, results in rough eyes because of an increase in cell death (Hay et al.,1994). A number of fly strains have been generated with cell death genesunder the control of UAS sites that can be used for ectopic expression in theeye and some of these are shown in Table 2.1.

To uncover a genetic interaction between your candidate gene andknown components of the cell death pathway several approaches can beundertaken. These include (1) gene activation screen, where the gene isoverexpressed; (2) dominant modifier screen by use of single gene disruptionalleles; and (3) target gene inactivation screen undertaken by the knockdownof gene expression. These are all based on the ability of the gene expression,either increased or decreased, to modify a sensitized rough eye phenotype.

To achieve overexpression of a gene in the eye, the GAL4/UAS systemis used such that the gene of interest is inserted downstream of UAS. Thiscan be achieved by direct cloning into the P-element vector, pUAST, thatcontains UAS sites (available from Drosophila Genomics Resource Centerhttps://dgrc.cgb.indiana.edu/) and then generating transgenic flies byP-element mediated transformation (by standard procedures; Ashburneret al., 2005). Alternately, a collection of misexpression Drosophila lines(P{EP}) are available that contain a P-element with UAS sites, that wheninserted near the 50 end of gene in the correct orientation can be activatedby GAL4 to drive expression of the downstream gene (R�rth, 1996; R�rthet al., 1998). Approximately 2300 P{EP} lines are available from the various

Table 2.1 Cell death transgenes available for expression in the eye

TransgeneEffect on cell deathin the eye Source

GMRhid* Increase Bloomington

GMRgrim* Increase Bloomington

UASreaper Increase Bloomington

UASprodronc Increase Meier et al., 2000; Quinn et al., 2000

GMRdrice Increase Song et al., 2000

GMRDcp-1 Increase Song et al., 2000

UASp35 Decrease Bloomington

UASDIAP1 Decrease Bloomington

UASdroncDN Decrease Meier et al., 2000; Quinn et al., 2000

UASRNAi Increase/decrease Available from VDRC

* If hid or grim overexpression is used, direct GMRhid and GMRgrim lines will need to be used asGMRGAL4, UAShid (or grim) is lethal. GMRGAL4 will still be required to co-express an interactinggene with UAS sites.

A full description of the transgene can be found on FlyBase: http://flybase.bio.indiana.edu/BloomingtonDrosophila Stock Center at Indiana University: http://flystocks.bio.indiana.edu/The Vienna DrosophilaRNAi Center (VDRC): www.vdrc.at

34 Donna Denton et al.

stock centers, and a list of Drosophila stock centers is available at FlyBase(http://flybase.bio.indiana.edu/) (reviewed in Adams, 2002).

Traditionally, genetic screens have required the generation of mutationsin genes resulting in decreased function, and many thousands of mappedgene disruption lines are available (Bellen et al., 2004). Another approach isto knockdown a gene of interest by use of RNA-mediated gene interfer-ence (RNAi). RNAi can be triggered in Drosophila by the presence of longdouble-stranded hairpin RNA that can be expressed from a transgene.UAS-RNAi transgenic lines are available from Vienna Drosophila RNAiCenter (VDRC, www.vdrc.at; Dietzl et al., 2007) and the DrosophilaGenetic Resource Center (www.dgrc.kit.ac.jp/en/) in Japan or can begenerated by construction of inverted repeats by use of a vectors such aspRISE and pWIZ, available from theDrosophilaGenomics Resource Center(https://dgrc.cgb.indiana.edu/).

The availability of such overexpression and knockdown fly lines makes itfeasible to examine genetic interactions with components of the cell deathpathway. This can be done by screening for modification, that is, eithersuppression (more mild) or enhancement (more disrupted) of the eye-specific phenotype from expression of a cell death regulator (e.g., GMRhidor GMRp35). To achieve this, the gene of interest (e.g., P{EP} or RNAi)line can be crossed to the sensitized cell death eye phenotype line (Fig. 2.4).

The use of loss-of-function mutations can also be examined in thismanner, where the effect of a reduction in gene dose on eye phenotypecan be determined. A prerequisite for this type of analysis is that a reductionin gene dose by half must be sufficient to modify the phenotype, thereforeenabling the dissection of the contribution of your gene of interest to thecell death pathway.

A rough eye phenotype associated with either increased or decreased celldeath (Table 2.1) can be used in crosses to examine the effect of yourtransgene of interest. Crosses with GMRGAL4 are routinely set up at25 �C, because the activity of GAL4 is temperature dependent (Duffy,2002). If a severe effect is observed, reducing the temperature to 18 �Cmay enable more subtle differences to become evident. It is also necessary tokeep in mind that GMRGAL4 alone induces cell death in the eye (Kramerand Staveley, 2003) and so must always be included as a control by crossingto a wild-type line (does not contain UAS transgene).

The phenotype of the eye can be examined by use of a dissectingmicroscope comparing the gene of interest to an appropriate control.Photographic images can be taken under the dissecting microscope, orsamples can be dehydrated in serial dilutions of acetone (25, 50, 75 and100% acetone) for scanning electron microscopy. After dehydration, fliescan be air-dried, mounted onto EM studs, and viewed without coating.Alternately, they can be dried by critical point drying and coated beforeelectron microscopy.

Studying Cell Death in Drosophila 35

Note: The exact requirements for setting up crosses will vary betweenexperiments, therefore, detailed instructions cannot be given within thescope of this section. Useful resources include Ashburner et al. (2005),Greenspan (2004), and Sullivan et al. (2000). The UAS constructs can beused with alternative tissue-specific or ubiquitous GAL4 drivers to express atdifferent stages of development to induce or inhibit apoptosis. A P-elementis a type of modified transposable element commonly used in generatingtransgenic flies as well as insertional mutagenesis.

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