RESEARCH ARTICLE

The developing Drosophila eye – a new model to study centriolereductionMaria G. Riparbelli, Veronica Persico, Marco Gottardo and Giuliano Callaini*

ABSTRACTIn the developing Drosophila eye, the centrioles of the differentiatingretinal cells are not surrounded by the microtubule-nucleatingγ-tubulin, suggesting that they are unable to organize functionalmicrotubule-organizing centers. Consistent with this idea, Cnn andSpd-2, which are involved in γ-tubulin recruitment, and the scaffoldprotein Plp, which plays a role in the organization of the pericentriolarmaterial, are lost in the third-instar larval stage. However, thecentrioles maintain their structural integrity, and both the parentcentrioles accumulate Asl and Ana1. Although the loading of Aslpoints to the acquisition of the motherhood condition, the daughtercentrioles fail to recruit Plk4 and do not duplicate. However, it issurprising that the mother centrioles that accumulate Plk4 also neverduplicate. This suggests that the loading of Plk4 is not sufficient, inthis system, to allow centriole duplication. By halfway through pupallife, the centriole number decreases and structural defects, rangingfrom being incomplete or lacking B-tubules, are detected. Asl, Ana1and Sas-4 are still present, suggesting that the centriole integrity doesnot depend on these proteins.

KEY WORDS: Centrosome, Centriole elimination, Centriolestructure, Drosophila

INTRODUCTIONThe centrosome is a structured protein complex that recruitsmicrotubule-nucleating proteins and tubulin (Woodruff et al.,2017), acting as the main microtubule-organizing center (MTOC)of the animal cells (Sanchez and Feldman, 2017). Through its abilityto nucleate the cytoplasmic microtubule network, the centrosomeplays essential roles in various cellular activities, includingcytoplasmic transport, cell movement, chromosome segregationand primary cilia formation (Bettencourt-Dias et al., 2011; Arquintet al., 2014; Conduit et al., 2015). The centrosome is also involved insome aspects of the cell division because it organizes the mitoticspindle and dictates its orientation throughout the cell cycle (Roubinetand Cabernard, 2014; Meraldi, 2016). Therefore, the accurate controlof centrosome dynamics is instrumental to avoiding a plethora ofcellular defects including ciliopathies, microcephalies, aneuploidyand cancer (Zyss and Gergely, 2009; Crasta et al., 2012; Vitre andCleveland, 2012; Chavali et al., 2014; Godinho and Pellman, 2014).Centrosome elimination is a common feature during gametogenesis

of most organisms (Manandhar et al., 2005; Mikeladze-Dvali et al.,2012; Pimenta-Marques et al., 2016). Female germ cells, indeed, loseor inactivate their centrosomes to avoid multipolar spindles at

fertilization. Thus, the assembly of the zygotic centrosome is drivenby the sperm-provided centrioles (Schatten, 1994). Centrosomeelimination has been also reported in endoreduplicating intestinalcells of Caenorhabditis elegans (Lu and Roy, 2014) and follicle cellsof Drosophila melanogaster (Mahowald et al., 1979). The alterationof the centrosome integrity has been also observed in human post-mitotic cells (Zebrowski et al., 2015) and in Drosophila somatic cystcells (Riparbelli et al., 2009).

Since the organization and integrity of the centrosome dependson a pair of centrioles at its heart (Sluder and Rieder, 1985),understanding centriole dynamics is crucial to decipheringcentrosome behavior. We have now a quite detailed knowledge ofcentriole composition and architecture, and the process of theirduplication (Lattao et al., 2017), but there is a poor understanding ofhow the centrioles are eliminated or inactivated in differentiated cells.

It has been recently reported that the starfish oocytes lose mothercentrioles before the daughters (Borrego-Pinto et al., 2016). In thissystem, the mother centriole retains its microtubule-organizingactivity and moves through a dynein-dependent process towards theplasma membrane to be eliminated with the extrusion of the polarbody. Thus, the elimination of the parent centrioles is associatedwith their different activity. The basal bodies of Caenorhabditissensory neurons also degenerate early in neuronal differentiation,after the formation of the ciliary structures (Serwas et al., 2017).

In an attempt to study the mechanisms underlying centrioleelimination in post-mitotic cells, we examined centriole behaviorduring the developments of the Drosophila eye. The adultDrosophila eye is formed from ∼750 ommatidial units that arederived by a complex differentiation process that requires dramaticcell transformations (Wolff and Ready, 1993; Carthew, 2007). Theeye-antennal imaginal disc of the third-instar Drosophila larvae iscrossed by the morphogenetic furrow that represents the boundarybetween the anterior region in which undifferentiated epithelial cellsproliferate and the posterior region that holds the differentiatingrhabdomeric cells (Cagan, 2009; Treisman, 2013). The epithelialcells ahead of the morphogenetic furrow proliferate randomly, butarrest in G1 as they enter the morphogenetic furrow (Thomas et al.,1994; Treisman and Heberlein, 1998). The cells posterior to themorphogenetic furrow are organized into regularly spaced groups,the ommatidial preclusters, within which the differentiation of thephotoreceptors occurs (Tomlinson and Ready, 1987; Wolff andReady, 1991). The remaining uncommitted cells re-enter the cellcycle and undergo a single round of cell division within the secondmitotic wave, before they arrest terminally (Wolff and Ready, 1991;de Nooij and Hariharan, 1995; Firth and Baker, 2005; Escudero andFreeman, 2007). As the distance from the furrow increases, theommatidial units became complete with the recruitment of the lastthree rhabdomeric cells and the differentiation of pigment- and lens-secreting cone cells (Ready et al., 1976; Wolff and Ready, 1991).Since the post-mitotic cell differentiation occurs progressively,successive developmental stages are present in the anterior to theReceived 25 September 2017; Accepted 15 January 2018

Department of Life Sciences, Via A. Moro 2, University of Siena, 53100 Siena, Italy.

*Author for correspondence (callaini@unisi.it)

G.C., 0000-0003-2252-0309

1

© 2018. Published by The Company of Biologists Ltd | Journal of Cell Science (2018) 131, jcs211441. doi:10.1242/jcs.211441

Journal

ofCe

llScience

posterior region of the imaginal disc. Thus, the neuroepithelial cellsof the developing Drosophila eye represent a suitable model tostudy the changes in centriole dynamics and organization thataccompany terminal cell fate specification.We show here that the centrosomes of theDrosophila retinal cells

lose their activity early during the larval stage, whereas thecentrioles start to disappear later by halfway through the pupallife. Centriole elimination begins with the gradual disassembly ofthe microtubule wall, followed by the disappearance of thecartwheel and the loss of the ninefold symmetry. Moreover, thecentrioles of post-mitotic cells fail to duplicate even if theyaccumulate Asl and Plk4.

RESULTSNeuroepithelial cells lose functional centrosomesThe eye-antennal disc of theDrosophila third-instar larva is crossedby a distinct groove, the morphogenetic furrow, that establishes theanterior and posterior regions of the disc (Fig. 1A). The apical

cytoplasm of the retinal cells in the posterior region showed focalaccumulations of tubulin continuous with longitudinal bundles ofmicrotubules (Fig. 1B). These microtubules radiated from electron-dense material lining the plasma membrane of short microvillus-likeprojections (Fig. 1C). Distinct centrioles were found in the apicalcell cytoplasm, but they did not contact the microtubules nor theperipheral electron-dense material (Fig. 1D). These observationspoint to non-conventional microtubule-organizing centers like thosedescribed in the cone cells of theDrosophila ommatidia (Mogensenet al., 1993) and suggest that centrioles of the epithelial cells withinthe posterior region of the imaginal discs were unable to properlyrecruit centrosomal material. To verify whether these centrioles losetheir ability to recruit the main centrosomal proteins during thedifferentiation process of the ommatidia, we first analyzed thelocalization of γ-tubulin, the master protein for microtubulenucleation. γ-tubulin was found in the anterior region of the third-instar larval imaginal disc as small spots associated with thecentrioles of the interphase cells and as large aggregates at the polesof the mitotic cells (Fig. 1E). Behind the morphogenetic furrow,only the cells that underwent a new cell division within the secondmitotic wave and a few scattered mitotic cells in the more posteriorregion of the disc displayed distinct accumulations of γ-tubulin attheir spindle poles (Fig. 1E). Although γ-tubulin did not accumulateat the interphase centrosomes of the differentiating rhabdomericcells, a weak labeling was found at their apical surface (Fig. 1F).This staining did not overlap the centrioles but was presumablyassociated with the nucleation sites for the longitudinal microtubulebundles seen at the plasma membrane.

γ-Tubulin recruitment at the centrosome mainly depends on thepericentriolar protein centrosomin (Cnn) (Megraw et al., 1999).Thus, we asked whether the accumulation of this protein may bereduced during the differentiation of the rhabdomeric cells. We finda weak Cnn staining on centrioles of the interphase epithelial cellslocated both in the anterior and posterior regions of the imaginaldisc and within the morphogenetic furrow (Fig. 2A). By contrast, astrong Cnn accumulation was seen at the poles of the dividing cellsin the anterior proliferating region and at the poles of the spindleswithin the second mitotic wave (Fig. 2A). The centrioles of theinterommatidial cells were weakly stained, whereas the majority ofthe centrioles of the rhabdomeric cells did not accumulate Cnn(Fig. 2A). During the early pupal stages only a few spots of lowintensity were observed (data not shown).

The incorporation of Cnn into the pericentriolar material (PCM)is facilitated in somatic Drosophila cells by Spd-2 (Conduit et al.,2014a). Thus, we expected a temporal and spatial colocalization ofthese proteins during the early phases of the eye development. Theanti-Spd-2 antibody, indeed, mainly recognized the centrioles of theinterommatidial cells, whereas most of the centrioles associatedwith the apical region of the rhabdomeric cells were devoid of theSpd-2 protein (Fig. 2B).

Centrioles disappeared during eye developmentSince the behavior of the centrosome depends on the centrioles at itsheart we analyzed their dynamics during eye development. Wetraced centrioles through the localization of the conserved centriole-specific core protein Sas-4, which provides a link between thecartwheel and the microtubule wall (Hsu et al., 2008; Tang et al.,2011) and may represent a bona fide marker of centrioles in thedeveloping eye.

Distinct centriole pairs were found behind the morphogeneticfurrow on the narrow apical regions of the cells that formedthe ommatidial preclusters and within the undifferentiated

Fig. 1. Loss of centrosome function in the larval imaginal disc eye.(A) Whole-mount of the eye-antennal disc of the third-instar larva: themorphogenetic furrow (mf) represents the boundary between the anterior (a)and the posterior (p) regions of the disc. Anterior is to the left. (B) Behind themorphogenetic furrow (mf), longitudinal bundles of microtubules (arrowheads)cross the cytoplasm of the retinal cells and end in the apical region wherestrong accumulations of tubulin are found (arrows). (C) The cell membranebulges out to form short microvillus-like projections containing electron-densematerial in which the cytoplasmic microtubules end (arrowheads). (D) Detail ofthe apical region of a retinal cell showing two separated centrioles that lackPCM and are not in contact with cytoplasmic microtubules. (E–E″) γ-tubulin isassociated with the centrioles of the proliferating cells in the anterior region ofthe eye disc (a) and inside the morphogenetic furrow (mf); behind themorphogenetic furrow, a distinct accumulation of γ-tubulin is found at thespindle poles of the cells within the second mitotic wave (arrowheads),whereas γ-tubulin does not associate with the centriole clusters of thedifferentiating rhabdomeric cells (asterisks). (F–F″) Weak γ-tubulin spots(arrowheads) that do not overlap with the Plp signal (arrows) are present in theapical region of the rhabdomeric cells. Scale bars: 100 µm (A); 2.5 µm (B);250 nm (C,D); 15 µm (E,E′), 5 µm (inset E′); 8 µm (F–F″), 5 µm (inset F″).

2

RESEARCH ARTICLE Journal of Cell Science (2018) 131, jcs211441. doi:10.1242/jcs.211441

Journal

ofCe

llScience

interommatidial cells (Fig. 3A). Each rhabdomeric cell alsodisplayed two apical centrioles. Since the ommatidia consist ofeight rhabdomeric cells, we found evenly spaced groups of eightcouples of centrioles within the posterior region of the disc(Fig. 3A).Surprisingly the Sas-4 signal had a different intensity within the

centrioles of the same pair (Fig. 3B). To decipher whether thissignal was a property of only one parent centriole, we counterstainedthe eye imaginal discs with an antibody against centrobin (Cnb),which specifically recognizes the daughter centrioles (Gottardoet al., 2016). The Cnb stain overlapped with the centrioles thathad lower Sas-4 intensity (Fig. 3B) that can be, therefore, taken asdaughters. The lower Sas-4 accumulation at the daughter centrioleswas also observed when the parent moved away (Fig. 3B).The eye imaginal disc dramatically changed shape during the

transition to the pupal stage becoming a thin disc-like epitheliumin which the ommatidia are separated by a matrix of unpatternedinterommatidial cells arranged in double or triple rows. At ∼25 hafter puparium formation (APF), the interommatidial cells sort intosingle rows disposed in a precise hexagonal pattern (Fig. 3C).At this developmental stage, the centrioles of the rhabdomericcells were barely detectable because the apical surface of thesecells had retracted below the cone cells. By contrast, the centriolesof the interommatidial, cone and primary pigment cells could beeasily identified because they were present at approximately thesame focal plane at the surface of the ommatidial units. Each cellhad one distinct centriole pair at this stage of development.Therefore, apical views of the retina show groups of 12 centrioles,eight from cone cells and four from primary pigment cells,surrounded by the centriole pairs of several interommatidial cells(Fig. 3C). At ∼45 h APF the interommatidial cells flattened anddistinct mechano-sensory bristles were visible at the anteriorvertex of each ommatidium (Fig. 3D). The majority of theinterommatidial, cone and pigment cells showed single Sas-4 spots(Fig. 3D). By 65 h APF, only two or three spots of Sas-4 werefound in the apical region of the ommatidial units (Fig. 3E). Aquantification of the centrioles as defined by Sas-4 stainingconfirmed the progressive reduction of their number from 25 h to65 h APF (Fig. 3F,G).

Centrioles of the pupal eye lost the scaffold protein Plp, butmaintained the core proteins Asl and Ana1It has recently been suggested that centriole elimination inDrosophila somatic cells and female germ cell line could be aconsequence of the loss of different components of the PCM(Pimenta-Marques et al., 2016). This prompted us to verify whetherthe centriole disappearance during later stages of eye developmentwas also associated with the loss of distinct centriolar constituentsthat play the main roles in centriole and centrosome biogenesis.Since we found that γ-tubulin was the first PCM component to belost, and then Cnn and Spd-2, we asked whether the dynamics of themain core centriole proteins involved in PCM recruitment andorganization might be also affected during eye development.

A distinct role in PCM organization is played by the coiled-coilPericentrin-like protein (Plp), which is radially arranged around thecentriole wall and organizes a distinct scaffold before the PCM isrecruited (Mennella et al., 2012). Moreover, the direct interactionbetween Plp and Cnn is required for normal centrosomeorganization and activity during interphase and mitosis of theDrosophila syncytial embryo (Lerit et al., 2015; Richens et al.,2015). We then sought to analyze the distribution of this peripheralcentriolar component during the eye development. Plp wasassociated with all the centrioles of the rhabdomeric cells in thelarval eye imaginal disc (Figs 1E and 4A), but found that they start todisappear early in the pupal stage. By 25 h APF, the anti-Plpantibody recognized seven or eight small spots within the apicalsurface of each ommatidium out of 12 recognized at this stage by theSas-4 antibody (Fig. 4A). At 45 h APF there were usually six Sas-4spots in the apical region of each ommatidium but only two or threeof them maintained a detectable Plp signal (Fig. 4A).

The assembly of the PCM around the centrioles requires theproducts of the genes asterless (asl) and anastral spindle 1 (ana1)(Lattao et al., 2017). The recruitment of Spd-2 seems, indeed, to beinitially supported by Asl (Conduit et al., 2014a), although additionalobservations have suggested that Drosophila centrioles lacking Aslmay efficiently recruit PCM (Galletta et al., 2014). Asl, in turn, isrecruited and maintained at the centriole by Ana1 (Fu et al., 2016;Saurya et al., 2016). These proteins extend from the inner centriole tothe outermost part of it and are usually recruited at the daughter

Fig. 2. PCM dynamics in the third-instar larval eye. Cnn(A,A′) accumulation is less evident than Spd-2 (B,B′). Thecentrioles of the rhabdomeric cells (arrows) lack both Cnn andSpd-2 (arrowheads). Scale bars: 15 µm (main images); 5 µm(insets).

3

RESEARCH ARTICLE Journal of Cell Science (2018) 131, jcs211441. doi:10.1242/jcs.211441

Journal

ofCe

llScience

centrioles when they acquire motherhood (Fu and Glover, 2012).Both Ana1 (Fig. 4B) and Asl (Fig. 4C) were found within thecentrioles of the rhabdomeric cells during the larval stage. When theapical surface of the photoreceptor cells turned by 90° at thebeginning of pupal life, the staining for Asl and Ana1 was barelydetectable. By contrast, the centrioles of the cone and primarypigment cells displayed strong Ana1 and Asl signals (Fig. 4B,C).Thus, the apical surface of each ommatidium at 25 h APF displayed acluster of 12 centrioles surrounded by the centriole pairs of the

interommatidial cells (Fig. 4B,C). By 45 h APF, we find six or sevenspots of Ana1 or Asl within each ommatidial unit (Fig. 4B,C). At65 h APF the number of spots reduced to two or three (Fig. 4B,C).

Daughter centrioles failed to recruit Plk4To understand how long the centrioles maintain their duplicationproperties, we looked at the localization of Plk4, a protein kinase atthe head of the centriole duplication process (Bettencourt-Diaset al., 2005; Habedanck et al., 2005). Plk4 has a distinct cell

Fig. 3. Timeline of centriole reduction during Drosophila eye development. (A–A′′′′) Distinct ommatidial preclusters (p) are visible just behind themorphogenetic furrow (mf) and complete ommatidial units (asterisks) form in the posterior region of the larval eye disc; the rhabdomeric cells have distinctcentriole pairs (arrowheads) that are clustered in evenly spaced groups (arrows). (B–B″) The daughter centrioles, identified by Cnb staining, display a lowerSas-4 accumulation (arrowheads). (C–E″) Top panels are surface views of the Drosophila retina at different developmental times; middle and lower panels aredetails of single ommatidia and surrounding interommatidial cells with their centrioles defined by Sas-4 localization. Centrioles of the photoreceptor cells, whenpresent, are out of focus and are not visible at the apical surface. (C–C″) Before the activation of the apoptotic machinery, the ommatidial and interommatidialcells display distinct centriole pairs. (D–D″) As the cell death reduces the interommatidial cells, the centriole number within the retina cells also decreases and onlyone Sas-4 spot for each cell is usually found. (E–E″) At later developmental stages only two or three Sas-4 spots are observed. Arrows and arrowheadspoint to centrioles of ommatidial and interommatidial cells, respectively. b, mechano-sensory bristles. (F) Schematic illustration of the apical profiles of singleommatidia at different developmental times showing two primary pigment cells (yellow) and four cone cells (gray). For simplicity, we did not include theinterommatidial cells. The eight photoreceptor cells lie below the cone cells and are not visible. The red spots represent parent centrioles. (G) Quantificationanalysis of the centriole numbers within the ommatidia at different times of development as defined by Sas-4 localization. n, total number of the ommatidia scoredfor each developmental stage. Scale bars: 1 µm (A); 0.25 µm (A′–A′′′′); 1 µm (B,B′); 4 µm (B″); 1 µm (C–E); 0.25 µm (C′–E″).

4

RESEARCH ARTICLE Journal of Cell Science (2018) 131, jcs211441. doi:10.1242/jcs.211441

Journal

ofCe

llScience

cycle-dependent localization on the centrioles of the anterior regionof the larval imaginal disc where the undifferentiated cellsproliferated randomly. At the beginning of interphase, Plk4 wasassociated with only one centriole of each pair (Fig. 5A). Thiscentriole expressed more Sas-4 and was identified as the mother onthe basis of the lack of Cnb (Fig. 3B). As interphase progressed thedaughter centrioles gradually accumulated Plk4 and reached thesame fluorescence intensity as the mothers. Both the parentcentrioles soon displayed a small daughter lacking Plk4. The tightcentriole pairs then migrated to the opposite poles of the cells duringprometaphase/metaphase (Fig. 5A) and disengaged at anaphase(Fig. 5A). At the end of telophase, the parent centrioles were widelyseparated and each sister cell inherited a pair of centrioles with onlythe mother expressing Plk4 (Fig. 5A). Electron microscopy (EM)

analysis of the anterior region of the larval eye imaginal discconfirmed the presence of interphase cells with duplicatedcentrioles (Fig. 5B). The short procentrioles formed by the nineA-tubules and some growing B-tubules were orthogonal to theproximal end of the mother centrioles that were in turn formed bythe nine doublet microtubules (Fig. 5B).

No sign of centriole duplication was seen behind themorphogenetic furrow, except within the second mitotic wavearea. The retinal cells of the posterior region of the imaginal discdisplayed a centrioles pair in which only the mother showed adistinct Plk4 accumulation (Fig. 5C). However, the mothercentrioles did not support procentriole assembly despite the factthat they showed accumulation of Plk4. At 25 h APF each retinalcell expressed two Sas-4 spots, but only one Plk4 spot was present

Fig. 4. Dynamics of the centriole core proteinsduring eye development. Distributions of Plp(A–A″), Ana1 (B) and Asl (C) at different stages ofeye development. All the centrioles accumulateAna1 and Asl, but some of them lack Plp during thepupal stages. Scale bars: 0.25 µm.

5

RESEARCH ARTICLE Journal of Cell Science (2018) 131, jcs211441. doi:10.1242/jcs.211441

Journal

ofCe

llScience

(Fig. 5D). As the eye development proceeded the number ofcentrioles expressing Plk4 further reduced. The Plk4 labeling wasbarely detectable by 45 APF when each cone and pigment celldisplayed a single Sas-4 spot (data not shown).EM analysis revealed that distinct procentrioles closely

associated with the mothers only in the region just posterior themorphogenetic furrow where the cells start to divide to enter thesecondmitotic wave (data not shown). Behind this tight proliferativearea each cell had only two centrioles and these had lost theirorthogonal configuration and became un-orientated (Fig. 5E). Onecentriole of the pair was shorter and looked like a procentriole(Fig. 5E). This centriole was usually composed of a distinctcartwheel and single A-tubules; only occasionally were one or twoB-tubules observed. Therefore, in differentiating rhabdomeric cells,the procentrioles did not complete their doublet content and failed toelongate properly. Despite examining serial sections from severalcells in the posterior region of the larval imaginal disc (n=47), wenever observed parent centrioles in association with a newly formed

daughter. The different length of the parent centrioles duringinterphase is unusual, since sister cells inherit, at the end of mitosis,two disengaged and equally sized centrioles. Serial sections of latetelophase cells revealed that there was a pair of different sizedcentrioles at the cell poles (Fig. 5F) pointing to the failure ofcentriole elongation during the previous interphase.

Centrioles of post-mitotic cells lose their structural integrityThe mother centrioles within the anterior and the posterior regionsof the larval imaginal disc were built by nine doublets ofmicrotubules and a central cartwheel (Fig. 6A). This organizationpersisted through the early pupal life. By 45 h APF all the centriolesscored (n=27) still maintained a central cartwheel, but often theydisplayed distinct defects of the microtubule wall. These defectswere more evident in cross sections and ranged from incomplete tolacking B-tubules (Fig. 6B,C). We excluded the possibility thatthese centrioles were remnant procentrioles that failed to grow. Theassembly of the B-tubule occurs, indeed, during centriole

Fig. 5. Centriole duplication failure inpost-mitotic retinal cells. (A) Cellcycle-dependent Plk4 localization inproliferating neuroepithelial cells relativeto the centriole marker Sas-4: note thatPlk4 docking is restricted to the centrioleexpressing more Sas-4; I, interphase,M, metaphase, A/T, anaphase/telophase. (B) EM analysis ofproliferating late interphase cellsshowing two pairs of engagedcentrioles; the daughter centriole isformed by A-tubules and some growingB-tubules and it is orthogonal to themother that is built by a set of ninecomplete doublets. At the end of thelarval stage (C) and during the earlypupal life (D–D″) Plk4 is restricted to thecentrioles expressing more Sas-4.(E–E″) The apical region of the cellsbehind the morphogenetic furrow showspairs of disoriented centrioles: one ofthem is shorter with an incomplete walland looks like a procentriole.(F,F′) Differently sized centrioles arefound at the poles of late telophasespindles. Scale bars: 0.8 µm (A); 500 nm(B,E); 1 µm (B′,B″,E′,E″,F′); 1 µm (C);0.25 µm (D–D″); 2 µm (F).

6

RESEARCH ARTICLE Journal of Cell Science (2018) 131, jcs211441. doi:10.1242/jcs.211441

Journal

ofCe

llScience

elongation from arc-like projections starting from the external sideof the A-tubule, whereas the disassembly of the B-tubule occurred atthe opposite side (Fig. 6B) or at the middle region of the same tubule(Fig. 6C). Themajority of the centrioles scored during later stages ofeye development (75%, n=34) lost their cartwheel and the centriolewall was often built by eight singlet-doublet tubules (Fig. 6D).Thus, the diameter of these centrioles (148±2.3 nm; n=9) wasreduced compared to the diameter of the centrioles that maintainedthe ninefold symmetry (165±1.9 nm: n=21).

DISCUSSIONRecent findings suggest that the centrioles may be modified duringdevelopment, yet still maintain their function. In the Caenorhabditisembryo, indeed, the centrioles remodel to nucleate the axoneme ofsensory neurons (Nechipurenko et al., 2017), and during Drosophilaspermiogenesis the sperm basal body is modified both in proteincomposition and in ultrastructure (Khire et al., 2016). However, thecentrioles do not persist at the base of the mature sensory cilia inCaenorhabditis suggesting that they are dispensable for ciliamaturation and maintenance (Serwas et al., 2017). Similarly, thecentrioles of the differentiatingDrosophila ommatidia gradually losetheir structural organization and then disappear. It is unclear why itwould be advantageous to remove the centrioles from the developingDrosophila eye. Perhaps, centriole elimination could prevent theassembly of too many centrosomes and unnecessary mitotic spindlesthat may impair cell differentiation by imposing abnormal divisions.It has been, indeed, proposed that centrosome inactivation indifferentiated cells may function as a barrier restricting cell cyclere-entry (Wong et al., 2015).

PCM is lost before pupationγ-Tubulin and Cnn were the first PCM components that disappearedfrom the centrioles during the development of the Drosophila eye(Fig. 7). This agrees with similar observations showing the loss ofγ-tubulin within the posterior region of the larval eye imaginal disc(Fernandes et al., 2014). These findings might be explained by thespatial localization of Cnn and γ-tubulin within the PCM. 3D-structured illumination microscopy has indeed revealed that the PCMcomponents are organized in two discrete layers around the centrioleof Drosophila somatic cells (Fu and Glover, 2012; Mennella et al.,2012): an inner layer formed by the fibrous proteins Asl, pericentrinand Plp that span outwards from the centriole wall, and a peripheralmatrix composed of Spd-2, Cnn and γ-tubulin (Mennella et al.,2014). Thus, the more-external proteins could be the first to beremoved in differentiating retinal cells. Since Cnn organizes ascaffold to ensure the proper PCM architecture during interphase andmitosis (Conduit et al., 2014b; Lerit et al., 2015; Feng et al., 2017), itis possible that the reduction in the amount of Cnn at the centriolesleads to the instability or defective recruitment of γ-tubulin.It has been demonstrated that Spd-2 helps the incorporation of

Cnn into the centrosome in Drosophila cells (Dix and Raff, 2007;Giansanti et al., 2008; Conduit and Raff, 2010). Moreover, Cnn

seems to be required to maintain Spd-2 within the PCM (Conduitet al., 2014a). We, thus, expected a similar behavior of Spd-2 andCnn in the Drosophila eye imaginal discs. However, the Spd-2signal was present within the centrioles of the whole imaginal disc,whereas a distinct Cnn labeling was restricted to only the anteriorregion of the disc, the morphogenetic furrow and the dividing cellsof the secondmitotic wave. It has been reported that Spd-2 is presentat the centrosome of the Drosophila somatic cells with two distinctpopulations (Fu and Glover, 2012): one inner population close to thecentriole wall and another that localized at the PCM duringcentrosome maturation and presumably interacts with Cnn.Therefore, the most extensive Spd-2 labeling with respect to theCnn staining may be due to the inner Spd-2 population. However,we were surprised to find that Spd-2 was stably associated with thecentrioles of the interommatidial cells in the posterior region of theeye imaginal disc, whereas it is lost or is barely detectable at thecentrioles of the rhabdomeric cells. Since all these cells are post-mitotic, the diverse timeline of the PCM loss might reflect adifferent differentiation degree for the cells scored. Rhabdomericcells are, indeed, fully differentiated, whereas most of theinterommatidial cells have still to be finally committed.

Although γ-tubulin is not associated with the centrioles of theretinal cells in the posterior region of the larval imaginal disc, adistinct population of this protein is found within the apicalprotrusions of the rhabdomeric cells. Remarkably, Spd-2 and Cnnappear to be redundant regarding ensuring the proper localization ofγ-tubulin in these cytoplasmic domains, which are the sites forthe nucleation of longitudinal bundles of microtubules and mayrepresent a non-conventional microtubule-organizing center.Accordingly, differentiated animal cell types often display non-centrosomal microtubule-organizing centers that function apicallyin many epithelia to generate longitudinal microtubule bundles(Bartolini and Gundersen, 2006; Sanchez and Feldman, 2017).

The ‘young configuration’ of the daughter centriole does notprevent Asl accumulationThe recruitment of PCM requires Asl (Conduit et al., 2014a), whichaccumulates into the daughter centrioles at mitosis (Novak et al.,2014; Fu et al., 2016). Thus, in flies, Asl plays a crucial role inallowing the daughter centriole to mature in a mother able to recruitPCM and to duplicate (Novak et al., 2014; Fu and Glover, 2016). Theappropriate loading of Asl is ensured byAna1, which is recruited laterin interphase (Fu et al., 2016). At the end of mitosis, sister cells,therefore, inherit a pair of centrioles both expressing Asl and Ana1.This is also the case for the post-mitotic retinal cells in whichdaughter centrioles display Ana1 and Asl, suggesting that they haveacquired a motherhood condition. However, the parent centrioles ofretinal cells are unable to recruit the main PCM proteins and lose thescaffold protein Plp. Moreover, daughter centrioles maintain ‘young’characteristics and look like procentrioles, suggesting that they fail toundergo full maturation. Daughter centrioles are, indeed, shorter thanthe mothers and do not complete their microtubule wall. It has been

Fig. 6. Centriole disassembly during late pupal stages.Cross sections of mother centrioles from retinal cells duringlarval (A) and pupal stages (B–D). The mother centriolesduring the larval stage and early pupal life always displaycomplete sets of nine doublets with a distinct cartwheel (A).The disassembly of the B-tubule occurs at the sideopposite to the region of its assembly (B, arrows) or at themiddle of the tubule (C, arrowheads). (D) Abnormalcentriole symmetry in older retinal cells. Scale bars:250 nm (A–D).

7

RESEARCH ARTICLE Journal of Cell Science (2018) 131, jcs211441. doi:10.1242/jcs.211441

Journal

ofCe

llScience

shown that Asl is incorporated into daughter centrioles of thesyncytial embryos not as they acquire its complete structure at theinterphase exit but later in mitosis, but rather when the parentcentrioles have disengaged (Novak et al., 2016). Presumably, theaccumulation of Asl to the daughter centrioles of the retinal cellscould require the travel through mitosis rather that their full growth. Itis, therefore, possible that the acquisition of the motherhoodcondition needs additional players in these cells.Sas-4 has an important role in fly cells to promote the recruitment

of Asl to daughter centrioles during mitosis (Conduit et al., 2014a;Novak et al., 2016). However, in contrast to the constant level of Aslon both parent centrioles of the retinal cells, we observed a lowintensity of the Sas-4 signal at the daughter centrioles, consistentwith a reduced antigen amount due to their incomplete structure.This finding suggests that the accumulation of Asl is not directlycorrelated to the amount of Sas-4, but that a low threshold level ofSas-4 might be enough to recruit high levels of Asl.

Plk4 loading is not sufficient to trigger centriole duplicationIt has been suggested that daughter centrioles have to be modifiedbefore they can duplicate (Wang et al., 2011) and this modification inflies might be the disengagement and the accumulation of Asl (Novaket al., 2014). Daughter centrioles that lack Asl are unable to duplicateeven if they disengage from their mothers at the end of mitosis(Novak et al., 2014). However, the parent centrioles in post-mitoticretinal cells of the Drosophila eye are disengaged and accumulateAsl, but they are unable to duplicate. The need for Asl in centrioleduplication is explained by its crucial role in the loading of Plk4, themaster protein that marks the site where the new centriole is going tobe built (Bettencourt-Dias et al., 2005; Ohta et al., 2014) both in

Drosophila (Dzhindzhev et al., 2010) and humans (Sonnen et al.,2012). The inability of the daughter centrioles to duplicate is expectedbecause they lack Plk4, but it is surprising that the mother centriolesnever duplicate even if they accumulate Plk4. This suggests that theloading of Plk4 is not sufficient, in this system, to allow centrioleduplication. The usual pathway of centriole duplication might be thusinactivated in post-mitotic retinal cells despite the presence of theproteins directly involved in centriole assembly.

It has been recently demonstrated that Plk4 influences the properradial positioning of Asl on mature somatic centrioles (Galletta et al.,2016). However, we do not find a direct correlation between Aslaccumulation at the centrioles and Plk4 recruitment. Plk4 is lost earlyduring eye development, whereas the Asl signal is maintained at thecentrioles until later stages. Moreover, although all the centrioles ofthe retinal cells accumulate Asl, only the mothers displays a distinctPlk4 signal. Since the only significant difference between the parentcentrioles is the incomplete structure of the daughters, the asymmetriclocalization of Plk4 may be explained by additional components thatdelay the accumulation of Plk4 until the daughter centrioles acquiretheir complete organization. The low accumulation of Sas-4 on thedaughter centrioles might play some roles in this process. Therefore,loading Plk4 to the daughter centriole does not only require theaccumulation of Asl during the previous cell cycle, but Plk4recruitment might also depend on the correct assembly of the ninedoublet microtubules that form the centriole wall.

Maintaining the proper centriole structure in post-mitoticretinal cells does not depend on the PCMThe external PCM proteins γ-tubulin, Cnn, Spd-2 and Plp are lostduring larval or early pupal stages, but the centriole architecture is not

Fig. 7. Schematic representation of centriole and PCMdynamics during ommatidia development in theDrosophila eye.Parent centrioles are fully evidentuntil early pupal stages (red, mothers; green, daughters), but become barely distinguishable by 45 h APF when the centriole number reduces (blue,unspecified parent centrioles). The centriole-associated proteins assayed at different developmental stages are listed in the table: +, present; −, absent; and +/−,partially present.

8

RESEARCH ARTICLE Journal of Cell Science (2018) 131, jcs211441. doi:10.1242/jcs.211441

Journal

ofCe

llScience

affected by their depletion. This apparently contrasts with the findingthat the down-regulation of the PCM leads to centriole elimination inthe Drosophila female germline (Pimenta-Marques et al., 2016).Thus, centriole deconstruction may experience different pathways inmeiotic and post-mitotic cells. The first structural defects of the retinalcell centrioles are observed during later pupal stages when Sas-4,Ana1 and Asl are still detected. Therefore, maintaining the propercentriole organization does not directly depend on these proteins.Centriole deconstruction in post-mitotic retinal cells starts with theprogressive disassembly of the B-tubules pointing to the low stabilityof the external components of the centriole wall. It has been reportedthat antibodies against polyglutamylated tubulin lead to theprogressive destabilization of the microtubule wall in vertebratecultured cells, suggesting the involvement of this tubulin isoform inmaintaining the stability of the centriole microtubules (Bobinnecet al., 1998). We do not know whether polyglutamylated tubulin alsoinfluences the dynamics of the centrioles in the Drosophila retinalcells. It is, however, remarkable that the abnormal centrioles we foundin post-mitotic retinal cells look like the incomplete centriolesobserved in vertebrate cells following the incubation with antibodiesagainst polyglutamylated tubulin.

MATERIALS AND METHODSDrosophila strainsWe used fly stocks containing Ana1–GFP (Blachon et al., 2009) and Asl–GFP (Dzhindzhev et al., 2010) transgenes. Oregon-R stock was also used ascontrol. Flies were raised on a standard Drosophila medium at 24°C.

AntibodiesWe used the following antibodies: mouse anti-γ-tubulin-GTU88 (1:100;Sigma-Aldrich); rabbit anti-Cnb (1:200; Sigma-Aldrich); mouse anti-acetylated tubulin (1:100; Sigma-Aldrich); mouse anti-Sas4 (1:200;Gopalakrishnan et al., 2011); rabbit anti-Spd-2 (1:500; Rodrigues-Martinset al., 2007); chicken anti-Plp (1:1500; Rodrigues-Martins et al., 2007);rabbit anti-Cnn (1:400; Vaizel-Ohayon and Schejter, 1999); rabbit anti Plk4(1:50; Bettencourt-Dias et al., 2005). The secondary antibodies used (1:800)were Alexa Fluor 488 and 555-conjugated anti-mouse-IgG, anti-rabbit-IgGand anti-chicken-IgG, and were obtained from Invitrogen.

Immunofluorescence preparationsEye imaginal discs from larvae and pupae at different times after pupariumformation were dissected in phosphate-buffered saline (PBS) and fixed incold methanol for 10 min at −20°C. For antigen localization, the sampleswere washed for 20 min in PBS and incubated for 1 h in PBS containing0.1% bovine serum albumin (PBS-BSA, from Sigma-Aldrich) to block non-specific staining. The samples were incubated overnight at 4°C with thespecific antisera in a humid chamber. After washing in PBS-BSA, thesamples were incubated for 1 h at room temperature with the appropriatesecondary antibodies. In all cases, DNA was visualized after an incubationof 3–4 min in Hoechst 33258 (1 µg/ml, Sigma-Aldrich). Imaginal discswere mounted in small drops of 90% glycerol in PBS. Images were taken byan Axio Imager Z1 microscope (Carl Zeiss), using an 100× objective,equipped with an AxioCam HR cooled charge-coupled camera (Carl Zeiss).Gray-scale digital images were collected separately and then pseudocoloredand merged using Adobe Photoshop 5.5 software (Adobe Systems).

Transmission electron microscopyThe eye imaginal discs were isolated from third-instar larvae and pupae atdifferent stages of development and transferred in 2.5% glutaraldehydebuffered in PBS overnight at 4°C. Samples were subsequently rinsed in PBSand post-fixed in 1% osmium tetroxide in PBS for 2 h at 4°C. The materialwas washed in PBS, dehydrated in a graded series of ethanol, embedded in amixture of Epon-araldite resin, and then polymerized at 60°C for 48 h. Thinsections (50–60 nm thick) were obtained with a Reichert Ultracut Eultramicrotome equipped with a diamond knife, mounted upon copper

grids, and stained with samarium triacetate and lead citrate. Samples wereobserved with a Tecnai Spirit Transmission Electron Microscope (FEI)operating at 100 kV and equipped with a Morada CCD camera (Olympus).

AcknowledgementsWe would like to thank Jay Gopalakrishnan (Center For Molecular Medicine,University of Cologne, Germany), Monica Bettencourt-Dias (Gulbenkian Institute ofScience, Oeiras, Portugal), Ana Rodrigues-Martins (Gulbenkian Institute ofScience, Oeiras, Portugal) and Eyal Schejter (Department of Molecular Genetics,Weizmann Institute of Science, Rehovot, Israel) for generously providing theantibodies used in this study. We also thank Tomer Avidor-Reiss (Department ofBiological Sciences, University of Toledo, Toledo, USA) and Nikola Dzhindzhev(Department of Genetics, University of Cambridge, Cambridge, UK) for the fliescarrying the GFP transgene.

Competing interestsThe authors declare no competing or financial interests.

Author contributionsConceptualization: G.C., M.G.R.; Methodology: M.G.R., M.G.; Formal analysis:M.G.; Investigation: M.G.R., V.P.; Data curation: V.P., M.G.; Writing - original draft:G.C.; Supervision: G.C.

FundingThis research received no specific grant from any funding agency in the public,commercial or not-for-profit sectors.

ReferencesArquint, C., Gabryjonczyk, A.-M. and Nigg, E. A. (2014). Centrosomes as

signalling centres. Phil. Trans R. Soc. 369, 20130464.Bartolini, F. and Gundersen, G. G. (2006). Generation of noncentrosomal

microtubule arrays. J. Cell Sci. 119, 4155-4163.Bettencourt-Dias, M., Rodrigues-Martins, A., Carpenter, L., Riparbelli, M.,

Lehmann, L., Gatt, M. K., Carmo, N., Balloux, F., Callaini, G. and Glover, D. M.(2005). SAK/PLK4 is required for centriole duplication and flagella development.Curr. Biol. 15, 2199-2207.

Bettencourt-Dias, M., Hildebrandt, F., Pellman, D., Woods, G. andGodinho, S. A.(2011). Centrosomes and cilia in human disease. Trends Genet. 27, 307-315.

Blachon, S., Cai, X., Roberts, K. A., Yang, K., Polyanovsky, A., Church, A. andAvidor-Reiss, T. (2009). A proximal centriole-like structure is present inDrosophila spermatids and can serve as a model to study centriole duplication.Genetics 182, 133-144.

Bobinnec, Y., Khodjakov, A., Mir, L. M., Rieder, C. L., Edde, B. and Bornens, M.(1998). Centriole disassembly in vivo and its effect on centrosome structure andfunction in vertebrate cells. J. Cell Biol. 143, 1575-1589.

Borrego-Pinto, J., Somogyi, K., Karreman, M. A., Konig, J., Muller-Reichert, T.,Bettencourt-Dias, M., Gonczy, P., Schwab, Y. and Lenart, P. (2016). Distinctmechanisms eliminate mother and daughter centrioles in meiosis of starfishoocytes. J. Cell Biol. 212, 815-827.

Cagan, R. (2009). Principles of Drosophila eye differentiation. Curr. Top. Dev. Biol.89, 115-135.

Carthew, R. W. (2007). Pattern formation in the Drosophila eye. Curr. Opin. Genet.Dev. 17, 309-313.

Chavali, P. L., Putz, M. and Gergely, F. (2014). Small organelle, big responsibility:the role of centrosomes in development and disease. Phil. Trans. R. Soc. B. 369,20130468.

Conduit, P. T. and Raff, J. W. (2010). Cnn dynamics drive centrosome sizeasymmetry to ensure daughter centriole retention inDrosophila neuroblasts.Curr.Biol. 20, 2187-2192.

Conduit, P. T., Richens, J. H., Wainman, A., Holder, J., Vicente, C. C., Pratt, M. B.,Dix, C. I., Novak, Z. A., Dobbie, I. M., Schermelleh, L. et al. (2014a). A molecularmechanism of mitotic centrosome assembly in Drosophila. Elife 3, e03399.

Conduit, P. T., Feng, Z., Richens, J. H., Baumbach, J., Wainman, A., Bakshi,S. D., Dobbelaere, J., Johnson, S., Lea, S. M. and Raff, J. W. (2014b). Thecentrosome-specific phosphorylation of Cnn by Polo/Plk1 drives Cnn scaffoldassembly and centrosome maturation. Dev. Cell. 28, 659-669.

Conduit, P. T., Wainman, A. and Raff, J. W. (2015). Centrosome function andassembly in animal cells. Nat. Rev. Mol. Cell Biol. 16, 611-624.

Crasta, K., Ganem, N. J., Dagher, R., Lantermann, A. B., Ivanova, E. V., Pan, Y.,Nezi, L., Protopopov, A., Chowdhury, D. and Pellman, D. (2012). DNA breaksand chromosome pulverization from errors in mitosis. Nature 482, 53-58.

de Nooij, J. C. and Hariharan, I. K. (1995). Uncoupling cell fate determination frompatterned cell division in the Drosophila eye. Science 270, 983-985.

Dix, C. I. and Raff, J. W. (2007). Drosophila Spd-2 recruits PCM to the spermcentriole, but is dispensable for centriole duplication. Curr. Biol. 17, 1759-1764.

Dzhindzhev, N. S., Yu, Q. D., Weiskopf, K., Tzolovsky, G., Cunha-Ferreira, I.,Riparbelli, M., Rodrigues-Martins, A., Bettencourt-Dias, M., Callaini, G. and

9

RESEARCH ARTICLE Journal of Cell Science (2018) 131, jcs211441. doi:10.1242/jcs.211441

Journal

ofCe

llScience

Glover, D. M. (2010). Asterless is a scaffold for the onset of centriole assembly.Nature 467, 714-718.

Escudero, L. M. and Freeman, M. (2007). Mechanism of G1 arrest in theDrosophila eye imaginal disc. BMC Dev. Biol. 7, 13.

Feng, Z., Caballe, A., Wainman, A., Johnson, S., Haensele, A. F. M., Cottee,M. A., Conduit, P. T., Lea, S. M. and Raff, J. W. (2017). Structural basis for mitoticcentrosome assembly in flies. Cell. 169, 1078-1089.

Fernandes, V. M., McCormack, K., Lewellyn, L. and Verheyen, E. M. (2014).Integrins regulate apical constriction viamicrotubule stabilization in theDrosophilaeye disc epithelium. Cell Rep. 9, 1-13.

Firth, L. C. and Baker, N. E. (2005). Extracellular signals responsible for spatiallyregulated proliferation in the differentiating Drosophila eye. Dev. Cell 8, 541-551.

Fu, J. and Glover, D. M. (2012). Structured illumination of the interface betweencentriole and peri-centriolar material. Open Biol. 2, 120104.

Fu, J. and Glover, D. M. (2016). How the newborn centriole becomes amother. CellCycle 15, 1521-1522.

Fu, J., Lipinszki, Z., Rangone, H., Min, M., Mykura, C., Chao-Chu, J., Schneider,S., Dzhindzhev, N. S., Gottardo, M., Riparbelli, M. G. et al. (2016). Conservedmolecular interactions in centriole-to-centrosome conversion. Nat. Cell Biol. 18,87-99.

Galletta, B. J., Guillen, R. X., Fagerstrom, C. J., Brownlee, C. W., Lerit, D. A.,Megraw, T. L., Rogers, G. C. and Rusan, N. M. (2014). Drosophila pericentrinrequires interaction with calmodulin for its function at centrosomes and neuronalbasal bodies but not at sperm basal bodies. Mol. Biol. Cell. 25, 2682-2694.

Galletta, B. J., Jacobs, K. C., Fagerstrom, C. J. and Rusan, N. M. (2016).Asterless is required for centriole length control and sperm development. J. CellBiol. 213, 435-450.

Giansanti, M. G., Bucciarelli, E., Bonaccorsi, S. and Gatti, M. (2008). DrosophilaSPD-2 is an essential centriole component required for PCM recruitment andastral-microtubule nucleation. Curr. Biol. 18, 303-309.

Godinho, S. A. and Pellman, D. (2014). Causes and consequences of centrosomeabnormalities in cancer. Philos. Trans. R. Soc. Lond. B Biol. Sci. 369, 20130467.

Gopalakrishnan, J., Mennella, V., Blachon, S., Zhai, B., Smith, A. H., Megraw,T. L., Nicastro, D., Gygi, S. P., Agard, D. A. and Avidor-Reiss, T. (2011). Sas-4provides a scaffold for cytoplasmic complexes and tethers them in a centrosome.Nat. Commun. 2, 359.

Gottardo, M., Callaini, G. and Riparbelli, M. G. (2016). Does Unc–GFP uncoverciliary structures in the rhabdomeric eye of Drosophila? J. Cell Sci. 129,2726-2731.

Habedanck, R., Stierhof, Y.-D., Wilkinson, C. J. and Nigg, E. A. (2005). The Polokinase Plk4 functions in centriole duplication. Nat. Cell Biol. 7, 1140-1146.

Hsu, W.-B., Hung, L.-Y., Tang, C.-J., Su, C.-L., Chang, Y. and Tang, T. K. (2008).Functional characterization of the microtubule-binding and -destabilizing domainsof CPAP and d-SAS-4. Exp. Cell Res. 314, 2591-2602.

Khire, A., Jo, K. H., Kong, D., Akhshi, T., Blachon, S., Cekic, A. R., Hynek, S., Ha,A., Loncarek, J., Mennella, V. et al. (2016). Centriole Remodeling duringSpermiogenesis in Drosophila. Curr. Biol. 26, 3183-3189.

Lattao, R., Kovacs, L. and Glover, D. M. (2017). The centrioles, centrosomes,basal bodies, and cilia of Drosophila melanogaster. Genetics 206, 33-53.

Lerit, D. A., Jordan, H. A., Poulton, J. S., Fagerstrom, C. J., Galletta, B. J., Peifer,M. and Rusan, N. M. (2015). Interphase centrosome organization by the PLP-Cnn scaffold is required for centrosome function. J. Cell Biol. 210, 79-97.

Lu, Y. and Roy, R. (2014). Centrosome/cell cycle uncoupling and elimination in theendoreduplicating intestinal cells of C. elegans. PLoS One 9, e110958.

Mahowald, A. P., Caulton, J. H., Edwards, M. K. and Floyd, A. D. (1979). Loss ofcentrioles and polyploidization in follicle cells of Drosophila melanogaster. Exp.Cell Res. 118, 404-410.

Manandhar, G., Schatten, H. and Sutovsky, P. (2005). Centrosome reductionduring gametogenesis and its significance. Biol. Reprod. 72, 2-13.

Megraw, T. L., Li, K., Kao, L. R. and Kaufman, T. C. (1999). The centrosominprotein is required for centrosome assembly and function during cleavage inDrosophila. Development. 126, 2829-2839.

Mennella, V., Keszthelyi, B., McDonald, K. L., Chhun, B., Kan, F., Rogers, G. C.,Huang, B. and Agard, D. A. (2012). Subdiffraction-resolution fluorescencemicroscopy reveals a domain of the centrosome critical for pericentriolar materialorganization. Nat. Cell Biol. 14, 1159-1168.

Mennella, V., Agard, D. A., Huang, B. and Pelletier, L. (2014). Amorphous nomore: subdiffraction view of the pericentriolar material architecture. Trends CellBiol. 24, 188-197.

Meraldi, P. (2016). Centrosomes in spindle organization and chromosomesegregation: a mechanistic view. Chromosom. Res. 24, 19-34.

Mikeladze-Dvali, T., von Tobel, L., Strnad, P., Knott, G., Leonhardt, H.,Schermelleh, L. and Gonczy, P. (2012). Analysis of centriole eliminationduring C. elegans oogenesis. Development. 139, 1670-1679.

Mogensen, M. M., Tucker, J. B. and Baggaley, T. B. (1993). Multiple plasmamembrane-associated MTOC systems in the acentrosomal cone cells ofDrosophila ommatidia. Eur. J. Cell Biol. 60, 67-75.

Nechipurenko, I. V., Berciu, C., Sengupta, P. and Nicastro, D. (2017). Centriolarremodeling underlies basal body maturation during ciliogenesis inCaenorhabditiselegans. Elife 6, e25686.

Novak, Z. A., Conduit, P. T., Wainman, A. and Raff, J. W. (2014). Asterlesslicenses daughter centrioles to duplicate for the first time in Drosophila embryos.Curr. Biol. 24, 1276-1282.

Novak, Z. A., Wainman, A., Gartenmann, L. and Raff, J. W. (2016). Cdk1Phosphorylates Drosophila Sas-4 to recruit Polo to daughter centrioles andconvert them to centrosomes. Dev. Cell 37, 545-557.

Ohta, M., Ashikawa, T., Nozaki, Y., Kozuka-Hata, H., Goto, H., Inagaki, M.,Oyama, M. and Kitagawa, D. (2014). Direct interaction of Plk4 with STIL ensuresformation of a single procentriole per parental centriole. Nat. Commun. 5, 5267.

Pimenta-Marques, A., Bento, I., Lopes, C. A. M., Duarte, P., Jana, S. C. andBettencourt-Dias, M. (2016). A mechanism for the elimination of the femalegamete centrosome in Drosophila melanogaster. Science 353, aaf4866.

Ready, D. F., Hanson, T. E. and Benzer, S. (1976). Development of the Drosophilaretina, a neurocrystalline lattice. Dev. Biol. 53, 217-240.

Richens, J. H., Barros, T. P., Lucas, E. P., Peel, N., Pinto, D. M. S., Wainman, A.and Raff, J. W. (2015). The Drosophila Pericentrin-like-protein (PLP) cooperateswith Cnn to maintain the integrity of the outer PCM. Biol. Open 4, 1052-1061.

Riparbelli, M. G., Colozza, G. and Callaini, G. (2009). Procentriole elongation andrecruitment of pericentriolar material are downregulated in cyst cells as they enterquiescence. J. Cell Sci. 122, 3613-3618.

Rodrigues-Martins, A., Riparbelli, M., Callaini, G., Glover, D. M. andBettencourt-Dias, M. (2007). Revisiting the role of the mother centriole incentriole biogenesis. Science 316, 1046-1050.

Roubinet, C. and Cabernard, C. (2014). Control of asymmetric cell division. Curr.Opin. Cell Biol. 31, 84-91.

Sanchez, A. D. and Feldman, J. L. (2017). Microtubule organizing centers: from thecentrosome to noncentrosomal sites. Curr. Opin. Cell Biol. 44, 93-101.

Saurya, S., Roque, H., Novak, Z. A., Wainman, A., Aydogan, M. G., Volanakis,A., Sieber, B., Pinto, D. M. andRaff, J.W. (2016).DrosophilaAna1 is required forcentrosome assembly and centriole elongation. J. Cell Sci. 129, 2514-2525.

Schatten, G. (1994). The centrosome and its mode of inheritance: the reduction ofthe centrosome during gametogenesis and its restoration during fertilization. Dev.Biol. 165, 299-335.

Serwas, D., Su, T. Y., Roessler, M., Wang, S. and Dammermann, A. (2017).Centrioles initiate cilia assembly but are dispensable for maturation andmaintenance in C. elegans. J. Cell Biol. 216, 1659-1671.

Sluder, G. and Rieder, C. L. (1985). Centriole number and the reproductivecapacity of spindle poles. J. Cell Biol. 100, 887-896.

Sonnen, K. F., Schermelleh, L., Leonhardt, H. and Nigg, E. A. (2012). 3D-structured illumination microscopy provides novel insight into architecture ofhuman centrosomes. Biol. Open. 1, 965-976.

Tang, C.-J., Lin, S.-Y., Hsu, W.-B., Lin, Y.-N., Wu, C.-T., Lin, Y.-C., Chang, C.-W.,Wu, K.-S. and Tang, T. K. (2011). The humanmicrocephaly protein STIL interactswith CPAP and is required for procentriole formation. EMBO J. 30, 4790-4804.

Thomas, B. J., Gunning, D. A., Cho, J. and Zipursky, L. (1994). Cell cycleprogression in the developing Drosophila eye: roughex encodes a novel proteinrequired for the establishment of G1. Cell 77, 1003-1014.

Tomlinson, A. and Ready, D. F. (1987). Neuronal differentiation in the Drosophilaommatidium. Dev. Biol. 120, 366-376.

Treisman, J. E. (2013). Retinal differentiation in Drosophila. Int. Rev. Dev. Biol. 2,545-557.

Treisman, J. E. and Heberlein, U. (1998). Eye development in Drosophila:formation of the eye field and control of differentiation. Curr. Top. Dev. Biol. 39,119-158.

Vaizel-Ohayon, D. and Schejter, E. D. (1999). Mutations in centrosomin revealrequirements for centrosomal function during early Drosophila embryogenesis.Curr. Biol. 9, 889-898.

Vitre, B. D. and Cleveland, D. W. (2012). Centrosomes, chromosome instability(CIN) and aneuploidy. Curr. Opin. Cell Biol. 24, 809-815.

Wang, W.-J., Soni, R. K. R., Uryu, K. and Tsou, M. F. B. (2011). The conversion ofcentrioles to centrosomes: essential coupling of duplication with segregation.J. Cell Biol. 193, 727-739.

Wolff, T. and Ready, D. F. (1991). The beginning of pattern formation in theDrosophila compound eye: the morphogenetic furrow and the second mitoticwave. Development 113, 841-850.

Wolff, T. and Ready, D. F. (1993). Pattern formation in the Drosophila retina. In TheDevelopment of Drosophila melanogaster, Vol. 2 (ed. M. Bate and A. MartinezArias), pp. 1277-1325. Cold Spring Harbor Laboratory Press.

Wong, Y. L., Anzola, J. V., Davis, R. L., Yoon, M., Motamedi, A., Kroll, A., Seo,C. P., Hsia, J. E., Kim, S. K., Mitchell, J. W. et al. (2015). Reversible centrioledepletion with an inhibitor of Polo-like kinase 4. Science 348, 1155-1160.

Woodruff, J. B., Ferreira Gomes, B., Widlund, P. O., Mahamid, J., Honigmann,A. and Hyman, A. A. (2017). The centrosome is a selective condensate thatnucleates microtubules by concentrating tubulin. Cell 169, 1066-1077.

Zebrowski, D. C., Vergarajauregui, S., Wu, C.-C., Piatkowski, T., Becker, R.,Leone, M., Hirth, S., Ricciardi, F., Falk, N., Giessl, A. et al. (2015).Developmental alterations in centrosome integrity contribute to the post-mitoticstate of mammalian cardiomyocytes. Elife 4, e05563.

Zyss, D. and Gergely, F. (2009). Centrosome function in cancer: guilty or innocent?Trends Cell Biol. 19, 334-346.

10

RESEARCH ARTICLE Journal of Cell Science (2018) 131, jcs211441. doi:10.1242/jcs.211441

Journal

ofCe

llScience