The gap junction protein Innexin3 is required for eye disc growth in Drosophila

Author’s Accepted Manuscript The gap junction protein Innexin3 is required for eye disc growth in Drosophila Mélisande Richard, Reinhard Tavosanis, Michael Hoch

Bauer,

Gaia

www.elsevier.com/locate/developmentalbiology

PII: DOI: Reference:

S0012-1606(16)30281-0 http://dx.doi.org/10.1016/j.ydbio.2017.04.001 YDBIO7410

To appear in: Developmental Biology Received date: 11 May 2016 Revised date: 30 March 2017 Accepted date: 3 April 2017 Cite this article as: Mélisande Richard, Reinhard Bauer, Gaia Tavosanis and Michael Hoch, The gap junction protein Innexin3 is required for eye disc growth i n Drosophila, Developmental Biology, http://dx.doi.org/10.1016/j.ydbio.2017.04.001 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting galley proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

The gap junction protein Innexin3 is required for eye disc growth in Drosophila

Mélisande Richard1,3, Reinhard Bauer1, Gaia Tavosanis2 and Michael Hoch1*

1

Life & Medical Sciences Institute (LIMES)

Development, Genetics & Molecular Physiology Unit University of Bonn Carl-Troll-Straße 31 53115 Bonn, Germany 2

German Center for Neurodegenerative Diseases (DZNE)

Dendrite Differentiation Unit Sigmund-Freud-Str. 27 53127 Bonn, Germany 3

present address: German Center for Neurodegenerative Diseases (DZNE)

Dendrite Differentiation Unit Sigmund-Freud-Str. 27 53127 Bonn, Germany *

Corresponding author: [email protected]

Keywords: Drosophila, eye, Innexin, growth, disc, peripodial epithelium

ABSTRACT The Drosophila compound eye develops from a bilayered epithelial sac composed of an upper peripodial epithelium layer and a lower disc proper, the latter giving rise to the eye itself. During larval stages, complex signalling events between the layers contribute to the control of cell proliferation and differentiation in the disc. Previous work in our lab established the gap junction protein Innexin2 (Inx2) as crucial for early larval eye disc growth. By analysing the contribution of other Innexins to eye size control, we have identified Innexin3 (Inx3) as an important growth regulator. Depleting inx3 during larval eye development reduces eye size, while elevating inx3 levels increases eye size, thus phenocopying the inx2 loss- and gain-of-function situation. As demonstrated previously for inx2, inx3 regulates disc cell proliferation and interacts genetically with the Dpp pathway, being required for the proper activation of the Dpp pathway transducer Mad at the furrow and the expression of Dpp receptor Punt in the eye disc. At the developmental timepoint corresponding to eye disc growth, Inx3 colocalises with Inx2 in disc proper and peripodial epithelium cell membranes. In addition, we show that Inx3 protein levels critically depend on inx2 throughout eye development and that inx3 modulates Inx2 protein levels in the larval eye disc. Rescue experiments demonstrate that Inx3 and Inx2 cooperate functionally to enable eye disc growth in Drosophila. Finally, we demonstrate that expression of Inx3 and Inx2 is not only needed in the disc proper but also in the peripodial epithelium to regulate growth of the eye disc. Our data provide a functional demonstration that putative Inx2/Inx3 heteromeric channels regulate organ size.

INTRODUCTION The adult Drosophila eye presents a regular hexagonal array of approximately 750 facets, each consisting of 8 photoreceptors and 11 accessory cells (cone cells, pigment cells and bristle cells) (Wolff and Ready, 1991). The compound eye develops from the eye imaginal disc, a flattened epithelial sac which arises from a 20-cell invagination of the embryonic blastoderm (Ready et al., 1976). The eye disc is composed of two epithelial layers: an upper peripodial epithelium of flattened cells (PE) and lower disc proper of columnar cells (DP) (Auerbach, 1936; Cohen, 1993). While the DP will give rise to the eye itself, the PE will form external cuticular structures of the adult head (Chouinard and Kaufman, 1991; Fristrom and Fristrom, 1993; Pilot and Lecuit, 2005; Stultz et al., 2006; Lee et al., 2007). Cells of the eye disc proliferate extensively in both layers throughout larval development, to reach a final number of about 2000 cells at third instar (Ready et al., 1976). Interestingly, growth of the DP depends on signals from the PE (Cho et al., 2000; Gibson and Schubiger, 2000; Atkins and Mardon, 2009). The PE also contributes cells to the DP during larval development (Pallavi and Shashidhara, 2003; Lim and Choi, 2004; McClure and Schubiger, 2005). During the middle of the third instar phase, an indentation within the DP, called the morphogenetic furrow, is formed and advances from posterior to anterior (Wolff and Ready, 1991). As the furrow traverses the DP, the field of anterior undifferentiated cells is transformed into an array of periodically spaced photoreceptor cell clusters, separated by accessory cells (Ready et al., 1976; Wolff and Ready, 1991). During the course of pupal stages, accessory cells reorganise and morphogenetic changes take place to finally produce the neurocrystalline lattice observed in the adult eye (Cagan and Ready, 1989). We recently described a role for the gap junction protein Innexin2 in the control of Drosophila eye disc growth (Richard and Hoch, 2015). Loss of innexin2 in the larval eye disc led to a reduced adult eye size, a consequence of the fact that inx2 function is a prerequisite for cell proliferation and morphogenetic furrow movement in the disc proper (Richard and Hoch, 2015). Gap junctions fulfill the fundamental requirement of intercellular communication in developing multicellular animals, since they allow for transfer of small molecules between neighboring cells (Goodenough et al., 1996; Wei et al., 2004; Bauer et al., 2005; Phelan, 2005). Gap junctions consist of an array of intercellular channels formed by gap junction proteins which are encoded by three different gene families: the connexins and pannexins in deuterostomes (respectively 21 genes and 3 genes in the human) and the innexins in protostomes (8 genes in Drosophila, termed inx1 to inx8) (Panchin et al., 2000; Phelan and Starich, 2001; Söhl and Willecke, 2003; Phelan, 2005; Bauer et al., 2005; Barbe et al., 2006; Dahl and Locovei, 2006). All gap junction proteins, although distinct in their primary structure, fold into four-pass transmembrane proteins and oligomerize into hexa- or

octameric hemichannel subunits (Yeager et al., 1998; Oshima et al., 2016). Each communicating cell produces a hemichannel at the plasma membrane that dock head-to-head with its neighbor in the extracellular space to create a functional intercellular channel (Martin and Evans, 2004; Segretain and Falk, 2004). In addition, a growing amount of evidence shows that hemichannels also function as non-junctional channels, connecting the cytoplasm with the extracellular space (Scemes, 2012; Sáez and Leybaert, 2014). Out of the eight described Drosophila Innexins, five were shown to be transcribed in pupal eye imaginal discs (inx1, inx2, inx3, inx6 and inx7) (Stebbings et al., 2002). Since we know that inx2 controls eye size (Richard and Hoch, 2015), we decided to analyse whether inx1, inx3, inx6 and inx7 sustain eye disc growth. Several reports indicate that Inx1, Inx2 and Inx3 often associate and function together. In paired Xenopus oocytes, expression of Inx2 with Inx3 or Inx1 leads to the formation of voltage-sensitive channels (Stebbings et al., 2000; Holcroft et al., 2013). In larval glial cells, Inx1 and Inx2 colocalise, are mutually dependent for expression and are required for proper neuroblast proliferation (Holcroft et al., 2013; Spéder and Brand, 2014). Similarly, Inx2 and Inx3 colocalise in embryonic epithelia, depend on each other for localisation and are required to maintain epithelial integrity (Lehmann et al., 2006; Giuliani et al., 2013). Importantly, Inx2 and 3 were shown to interact via their C-terminal cytoplasmic domains to form heteromers in Drosophila embryos (Lehmann et al., 2006). In the amnioserosa, Inx3 was also shown to mediate the stability of Inx1, Inx2 and DE-Cadherin by forming a complex (Giuliani et al., 2013). A single report indicates that Inx6 can form channels with Inx7 in specific neurons of the mushroom body and that these are important for memory formation (Wu et al., 2011). Here we show that the Drosophila gap junction proteins Inx2 and Inx3 are cooperatively required in DP and PE to sustain eye disc growth. We propose that Inx2 and Inx3 heteromerise in the eye disc to fulfill their function.

RESULTS

Inx2 colocalises with Inx1 and Inx3 during eye development Our published observations have demonstrated that Inx2 is expressed throughout eye development (Richard and Hoch, 2015). Inx2 firstly localised apico-laterally in larval disc proper cells, was then restricted to the accessory cells posterior to the morphogenetic furrow and was finally expressed in pigment cells at pupal stages (Richard and Hoch, 2015). In addition, Inx2 was also expressed cortically in peripodial epithelium cells (Richard and Hoch, 2015). In several organs, Inx2 was shown to colocalise (at least partially) with Inx1 and Inx3 (Lehmann et al., 2006; Bohrmann and Zimmermann, 2008; Giuliani et al., 2013; Holcroft et al., 2013; Spéder and Brand, 2014). In addition, in situ hybridisation of pupal eye discs showed that inx1, inx2 and inx3 mRNAs are co-expressed in the pigment cells at mid-pupal stage (Stebbings et al., 2002). We thus analysed protein expression of Inx1 and Inx3 throughout eye development and compared it with that of Inx2. During the third instar larval stage, Inx3 strongly colocalised with Inx2 at the plasma membrane and was enriched apico-laterally in all disc proper cells anterior to the morphogenetic furrow (Fig. 1A’’’, supplementary Fig. 1A-A’’’ and 1C). Inx1 was also colocalised with Inx2 in those cells (Fig. 1C’’’), though not as perfectly as Inx3 (also see Suppl. Fig. 1D). The Inx1 staining observed in the nuclei was unspecific (asterisk in Fig. 1C’’’, Suppl. Fig.1D, E). Posterior to the furrow, expression of Inx1, Inx2 and Inx3 was restricted to the uncommitted progenitors located around the recruited photoreceptor cell clusters (Fig. 1B’, 1D’). In peripodial epithelium cells, Inx2 and Inx3 colocalised predominantly cortically in all cells and were enriched at the contact points between cells (Fig. 1E’’’, Suppl. Fig. 1B-B’’’ and 1C). Inx1 also colocalised with Inx2 in PE cells, though not as strongly as Inx3 (Fig. 1F’’’). At mid-pupal development (45% p.d. - pupal development), Inx3 was enriched at the membrane between secondary and tertiary pigment cells and, in a reduced manner, at the border between secondary pigment cells and bristle cells (Fig. 2A’’). Expression was apico-lateral (Fig. 2B’) but also present in the pigment cell feet contacting the photoreceptor cell feet at the floor plate (Fig. 2B’). Inx2 and Inx3 perfectly colocalised at this stage (Fig. 2D’’). As for Inx2 and Inx3, Inx1 expression was restricted to the pigment cells at this stage (Fig. 2C’) and Inx1 perfectly colocalised with Inx2 (Fig. 2E’’). During late pupal development (75-90% p.d.) Inx1 and Inx2 were still expressed in the pigment cells (Fig. 2H’), seen as dots on a longitudinal section (Fig. 2I’, J). Interestingly, Inx3 was strongly expressed in the forming rhabdomeres of photoreceptor cells, though moderate expression remained

in the pigment cells (Fig. 2F’, G’, J’), especially close to the cornea at the level of the cone cells (a colocalisation between Inx2 and Inx3 is shown at this level on Fig. 2K’’) and in the pigment cell feet. (Fig. 2G’). These results demonstrate that Inx1, Inx2 and Inx3 colocalise throughout eye development. Since inx6 and inx7 transcripts were also detected in the pupal eye disc, though not colocalising with inx1, inx2 and inx3 transcripts (Stebbings et al., 2002), we used anti-Inx6 and anti-Inx7 antibodies to detect protein expression in third instar eye discs. Inx6 was mostly detected as dots in all disc cells while Inx7 was present in nuclei (Suppl. Fig. 2A-A’’ and B-B’’), thus showing that Inx1, Inx2 and Inx3 do not obviously colocalise with Inx6 and Inx7 during eye disc larval development. Taken together, our results show that Inx1, Inx2 and Inx3 colocalise throughout eye development. Our analysis of the protein expression patterns of Inx1, Inx3, Inx6 and Inx7 shows that Inx3 is the gap junction protein that colocalises the best with Inx2 in the larval eye discs, at the timepoint at which disc cells proliferate and photoreceptor cells differentiate.

Adult eye size is controlled by inx3 We reported previously that eye discs lacking inx2 expression during larval stages do not grow properly. As a consequence, inx2 mutant adult eyes were reduced in size (Richard and Hoch, 2015). This phenotype could be phenocopied by downregulating inx2 expression with an inx2 RNAi construct under the control of eyGal4 or eyaGal4, both driving expression early in the whole eye field (Hazelett et al., 1998; Bui et al., 2000; Richard and Hoch, 2015; also see Suppl. Fig. 2C). Since we observed Inx1, Inx3, Inx6 and Inx7 protein expression in the larval eye discs, we asked whether expression of those Innexins contributes to growth regulation during larval stages. Using RNA interference, we depleted inx1, inx3, inx6 or inx7 in the larval eye disc with eyaGal4 and measured the resulting adult eye size (Figure 3). The RNAi lines used were verified to be functional in the eye disc (Suppl. Fig. 3A). We observed a statistically significant reduction in adult eye size in flies with an eye disc depletion in inx3 but no change upon depletion of inx1, inx6 or inx7 (Figure 3; one-way ANOVA: F(10.125)=11.46, p<0.01; post-hoc comparisons using the Tukey HSD revealed statistically significant differences (p<0.01) only between controls and inx3 RNAi conditions). Since an inx1 null mutant line was available (inx1 KO, (Giuliani et al., 2013)), we also measured adult eye size of those flies and they were similar to the size of the corresponding w controls (Figure 3), supporting the view that inx1 is not involved in eye size control. Since only the inx3 RNAi lines showed a substantial reduction in protein levels (Suppl. Fig. 3A), the jury is still out on whether

inx6 or 7 have a role in eye development. Overexpression of a full-length Inx2 construct under the control of eyGal4 led to a slight increase in eye size (Richard and Hoch, 2015). Interestingly, expression of an Inx3 transgenic construct with eyaGal4 also slightly increased eye size (103.9±2.2% of eyaGal4/+ control eye size, see Suppl. Fig. 3B), a similar increment as observed after Inx2 overexpression in the eye disc (Richard and Hoch, 2015). These results show that inx3, as inx2, controls eye size both in loss- and gain-of-function experiments.

inx3 controls proliferation in the eye disc and interacts with the Dpp pathway to control eye size We showed previously that loss-of-function of inx2 during larval eye disc development was accompanied by a decreased proliferation rate in the anterior eye disc compartment (Richard and Hoch, 2015). Since it is known that proper eye development necessitates a precisely regulated balance between growth on the one hand and apoptosis on the other hand (Neufeld and Hariharan, 2002), we asked which of these processes might be affected in inx3-depleted eyes⁠. We first examined the phenotype of adult eyes by preparing semi-thin sections of eyGal4/+ (control, Fig. 4A) or eyGal4 UAS RNAi inx3 fly eyes (Fig. 4B). We observed no difference in the size of photoreceptor cells or ommatidia (Fig. 4A, B) and the overall morphology of the inx3 knock-down eyes appeared wild-type (Fig. 4B). In addition, apoptosis rate was not increased in inx3-depleted eye discs, since they did not show an increased staining for activated caspase 3 in comparison to controls (activated caspase 3-positive cells counts in the anterior eye disc compartment: eyaGal4 UAS RNAi inx3: 81.2 ± 12.2% in comparison to the control 100.0 ± 26.9%, meand±s.d. of n=5, Fig. 4E’, F’). Since the amount of cells located anteriorly to the furrow sets the upper limit of ommatidia that can be generated (Kumar, 2011), we examined cell proliferation in the eye disc by counting the number of cells positive for the metaphase marker phospho-H3 histone in the anterior disc compartment. For this, we used the DEGal4 driver to express the inx3 RNAi only in the dorsal half of the eye disc (Morrison and Halder, 2010)⁠. We observed a 30% decrease in cell proliferation in the dorsal anterior compartment of knockdown eyes (reduced inx3 expression) when comparing pH3 histone counts with those obtained in the ventral anterior compartment (wild-type levels of inx3 expression) (Fig. 4C, D). These results demonstrate that inx3, as inx2 (Richard and Hoch, 2015), controls proliferation in the anterior compartment of third instar eye discs.

Dpp is known to promote eye disc growth throughout larval stages (Masucci et al., 1990; Burke and Basler, 1996; Firth et al., 2010) and we demonstrated previously that inx2 genetically interacts with the Dpp pathway, being required for proper activation of the Dpp pathway transducer Mad and expression of Dpp receptors Thickveins and Punt in L3 eye discs (Richard and Hoch, 2015). We thus examined whether activation of the Dpp pathway could also be controlled by inx3. The amount of phosphorylated Mad protein (Raftery et al., 1995; Sekelsky et al., 1995)⁠, a transcription factor phosphorylated upon activation of the Dpp pathway at the MF (Vrailas and Moses, 2006)⁠, was reduced considerably when inx3 was downregulated with DEGal4 (Fig. 5B’). pMad is known to accumulate in a broad band of cells anterior to and within the MF, and in a second stripe of more intensely-labelled cells around columns 3 and 4 at the posterior edge of the furrow (Firth et al., 2010)⁠. Within the dorsal inx3 knockdown area, both stripes were strongly reduced in intensity (Fig. 5B’’’). A close-up on the dorso-ventral boundary is presented in Fig. 5CC’’’. Similarly, levels of Punt, the type-II Dpp receptor (Childs et al., 1993; Wrana et al., 1994; Letsou et al., 1995; Ruberte et al., 1995)⁠, were clearly diminished in the whole dorsal part of the eye disc upon inx3 depletion (Fig. 5A’’’). Since we observed a decreased activation of the Dpp pathway in inx3-depleted eye discs, we asked whether changes in eye size observed upon inx3 knock-down were depending on the Dpp pathway. For this, we performed genetic interaction experiments in adult fly eyes, taking advantage of a medea RNAi line that has been shown to be functional in the eye (Richard and Hoch, 2015). Medea codes for a Drosophila SMAD protein (Raftery et al., 1995; Hudson et al., 1998; Raftery and Wisotzkey, 2008) and eye-specific knock-down of medea leads to flies with small eyes (Marinho et al., 2013; Richard and Hoch, 2015). Genetic interactions between medea and inx3 were studied by comparing the measured eye phenotype of flies expressing two RNAis with the expected phenotype of a combination of both RNAis based on individual single RNAis (Costanzo et al., 2011; Marinho et al., 2013)⁠. We thus used eyaGal4 to drive expression of the medea RNAi alone or in combination with our inx3 RNAi construct in the eye. We firstly observed that depletion of medea gave rise to a small percentage of flies with only one eye or no eyes at all (respectively 6% and 8%, see Fig. 5D). Co-depletion of inx3 in this background aggravated the phenotype (respectively 42% and 14% of flies having no or only one eye, Fig. 5D). Eye size of flies expressing the medea RNAi under the control of eyaGal4 alone or in combination with the inx3 RNAi was then measured and we compared the observed eye size to the expected one. To obtain the expected eye size (eye size without genetic interaction), we used a model that assumes that the expected double mutant phenotype can be the result of the multiplicative combination of the single mutant phenotypes (Costanzo et al., 2011; Marinho et al., 2013)⁠. The expected eye size value is depicted as a black

horizontal bar in Fig. 5E (72.5% of control eye size). The majority of medea inx3 RNAi flies (Fig. 5E green triangles) have smaller eyes than expected, pointing to a negative genetic interaction between inx3 and the Dpp pathway. These results show that inx3, as inx2 (Richard and Hoch, 2015), is required for activation of the Dpp pathway at the MF and for expression of Dpp receptor Punt. In addition, inx3 interacts genetically with the Dpp signaling pathway to control eye size.

Levels of Inx1 and Inx3 proteins depend on inx2 and Inx2 levels depend on inx3 in the eye disc Since Inx1, Inx2 and Inx3 colocalised throughout eye development (Fig. 1, Fig. 2), we produced small eye clones for a null mutation of inx2 (see Materials and Methods) in order to monitor Inx1 and Inx3 protein levels in the absence of inx2. In third instar larval eye discs, we observed that inx2clones, expressing no RFP (Fig. 6A, B, C), showed a strong reduction in protein expression of Inx1 and Inx3, both in the disc proper and overlaying peripodial epithelium (Fig. 6A’, B’, C’). Inx1 and Inx3 expression also depended on inx2 during pupal eye development, as seen in MARCM clones for inx2 (expressing GFP in Fig. 6D, E, F and G) stained for Inx1 (Fig. 6F’, G’) or Inx3 (Fig. 6D’’, E’). Taken together, these results show that Inx1 and Inx3 levels depend on inx2 throughout eye development. We then depleted inx1 and inx3 in larval eye discs to monitor protein levels of the other Innexins. In order to compare levels within the same larval eye disc, we used the DEGal4 driver to deplete inx1 or inx3 specifically in the dorsal part of the disc and compare protein levels in the dorsal vs. ventral area of the disc (Fig. 7). The top panels in Figure 7 show the region that we focused on. Depletion of inx1 in the dorsal half of the disc (Fig. 7A, B, arrowhead) led to minor changes in Inx2 levels (86.2±5.3% of control levels, n=5, see Mat. and Meth., Fig. 7A’) but without leading to a delocalisation of the protein from the plasma membrane, and did not change Inx3 levels (99.6±2.2% of control levels, n=5, Fig. 7B’). In contrast, expressing a inx3 RNAi line in the dorsal part of the eye disc (Fig. 7C, D, arrowhead) led to a slight decrease in Inx2 levels (70.3±5,2% of control levels, n=5, Fig. 7C’) accompanied by a strong delocalisation of the protein in the cytoplasm (arrowhead in Fig. 7C’), while Inx1 protein levels were not modified (99.6±2,1% of control levels, n=5, Fig. 7D’). Taken together, these results show that Inx1 and Inx3 levels depend on inx2 throughout eye development. inx3 influences Inx2 protein levels in the larval eye disc. In contrast, inx1 does nearly not influence Inx2 or Inx3 levels in the eye disc.

Overexpression of Inx2 and Inx3 rescues the small eye phenotype observed upon inx2 or inx3 depletion Since Inx2 and Inx3 colocalise and control each other’s levels in the eye disc (Fig.1, Figs. 6 and 7) and the loss of either leads to the same phenotype (Fig. 3, Fig. 4, Fig.5 and (Richard and Hoch, 2015)), we asked if they cooperate functionally to sustain eye disc growth during larval stages. We thus tested whether transgenic expression of inx1, inx2, inx3 or a combination thereof could rescue the small eye phenotype observed upon depletion of inx2 with eyaGal4 (Fig. 8). Expression obtained by one or two copies of a full-length UAS inx2 transgene in the inx2 knockdown background with eyaGal4 rescued the small-eye phenotype only partially but significantly, as did expression obtained by one UAS inx2 copy with one copy of UAS inx1 (78.0%±2.5% of control eye size for inx2 depleted eyes vs. 87.2%±8.3%, 87.6%±1.3% or 85.4%±4.0% for rescues, Fig. 8; oneway ANOVA: F(7.81)=22.23, p<0.01; post-hoc comparisons using the Tukey HSD revealed statistically significant differences (p<0.05) between all rescues containing UAS Inx2 and the inx2 RNAi condition). An explanation for the partial rescue could be that the the transgenic overexpression of inx2 did not reach sufficient levels of expression. Expression of UAS inx3 or UAS inx1 alone or in combination did not rescue the small eye phenotype (Fig. 8). Interestingly, overexpression of both full-length UAS inx2 and UAS inx3 nearly completely rescued the small eye phenotype (97.5%±3.6% of control eye size for the rescue, Fig. 8; Tukey HSD p<0.01), thus showing that expression of both Inx2 and Inx3 proteins is necessary to enable eye disc growth. We also tried to rescue the small-eye phenotype observed in inx3 knock-down animals (Suppl. Fig. 3C). Upon inx3 depletion in the eye disc, eye size is not affected to the extent observed in inx2 depleted animals (90.1% of control eye size upon inx3 depletion vs. 78.0% of control eye size for inx2 depletion, both with eyaGal4). This observation probably arises because some inx2 and inx3 transcription still takes place in the discs (Suppl. Fig. 5A) and as a matter of fact, Inx2 protein is not completely downregulated upon inx3 depletion (Fig. 7C’). In these mildly affected eyes, driving expression of either UAS inx3 or UAS inx2 with eyaGal4 at larval stages nicely rescued the small eye phenotype, as did a combination of both transgenes (Suppl. Fig. 3C; one-way ANOVA: F(3.41)=25.65, p<0.01; post-hoc comparisons using the Tukey HSD revealed statistically significant differences (p<0.01) between all rescues and the inx3 RNAi condition), confirming that both proteins are important to enable eye disc growth. Furthermore, a depletion of both inx2 and inx3 with eyaGal4 gave rise to flies with eyes that are markedly smaller as those observed in the single knock-downs (Suppl. Fig. 3D), highlighting the synergistic function of both Innexins.

Taken together, these results highlight the functional cooperation between Inx2 and Inx3 in sustaining larval eye disc growth.

Inx2 and Inx3 function in the peripodial epithelium and in the disc proper Since the peripodial epithelium (PE) has been shown to be required for disc proper (DP) cell proliferation and to regulate signalling pathways in the DP (Cho et al., 2000; Gibson and Schubiger, 2000; McClure and Schubiger, 2005), we sought to analyse whether inx2 and/or inx3 function in the PE. Inx2 and Inx3 colocalise in the plasma membrane of PE cells at larval stages (Fig. 1E’’’, Suppl. Fig. 1B-B’’’) and Inx3 protein levels depends on inx2 in the PE (Fig. 6B’). Similarly, Inx2 protein levels depend on inx3 in the PE (Suppl. Fig. 4A-A’’’). Since eyGal4 and eyaGal4 drive expression both in DP and PE (Suppl. Fig. 2C and (Gibson and Schubiger, 2000)), our inx depletion experiments (for inx2: (Richard and Hoch, 2015), for inx3: Fig. 3 of this paper) did not allow us to focus on potential layer-specific functions for Inx2 or Inx3. We thus used a PE-specific Gal4 driver, c311-Gal4 (Gibson and Schubiger, 2000), and a DP-specific driver, optix-Gal4 (a gift of M. Atkins and G. Mardon), to deplete inx2 or inx3 specifically in one cell layer during larval disc development. Those drivers were crossed to UAS-GFP and verified to express constructs only in the appropriate cell layer at larval stages (Suppl. Fig. 2D and E). As expected, a depletion of inx2 or inx3 in the larval eye disc proper produced small-eyed flies (Fig. 9A; one-way ANOVA: F(2.30)=49.0, p<0.01; post-hoc comparisons using the Tukey HSD revealed statistically significant differences (p<0.01) between control and knock-downs). Interestingly, flies with a depletion of inx2 or inx3 in the larval PE also developed small eyes (Fig. 9A; one-way ANOVA: F(2.34)=97.7, p<0.01; post-hoc comparisons using the Tukey HSD revealed statistically significant differences (p<0.01) between control and knock-downs). Upon inx2 knock-down in the DP with optixGal4, Inx2 levels in the anterior compartment were decreased by about 20% in comparison to controls (80.2±5.2% of control levels, n=5) and Inx3 levels by about 60% (40.8±3.2% of control levels, n=5). This decrease was accompanied by a delocalization of both proteins from the plasma membrane (Suppl. Fig. 4B-B’’’ and C-C’’’). In the peripodial epithelium, Inx2 was strongly delocalised from the plasma membrane in c311Gal4 UAS RNAi inx2 eye discs (Suppl. Fig. 4E) though the overall Inx2 levels in the PE were only slightly modified (93.8±3.1% of control levels, n=5). Despite the strong delocalization of Inx3 from the plasma membrane upon inx2 depletion (Suppl. Fig. 4E’), Inx3 levels were, as observed for Inx2 levels, only slightly diminished (95.2±4.3% of control levels, n=5). These results suggest that a delocalization of Inx2 and Inx3 from the plasma membrane in the PE is sufficient to produce a small eye phenotype.

Since we have shown that the inx2 and inx3 small eye phenotypes arise from defects in cell proliferation in the DP (Fig. 4C, D of this paper; Richard and Hoch, 2015), this experiment suggests that a proper inx2/inx3 expression in the PE is important to regulate cell proliferation in the larval eye disc proper. To analyse this question, we stained L3 eye discs depleted for inx2 in the PE (c311Gal4 UAS RNAi inx2) with the anti-pH3 histone antibody and compared histone counts in the anterior disc compartment of the DP with those of control eye discs (c311Gal4/+). Those counts are presented in Fig. 9D: a depletion of inx2 in the PE decreases proliferation in the underlying DP by about 30%. To analyse whether Innexins also sustain proliferation cell autonomously in the PE, we compared size of small inx2 clones with that of their twin clones in the PE (Fig. 9B). For this, we stained the discs with RFP, to delineate clone boundaries (part of an inx2 clone is marked with a yellow line in Fig. 9B), Homothorax (Hth) as a nuclear marker (Hth is a transcription factor strongly expressed in peripodial cell nuclei, see Bessa et al., 2002) (Fig. 9B’) and DE-Cadherin as a membrane marker (Fig. 9B’’). All inx2 clones observed were smaller as their respective twin clones (n=14, Fig. 9C), showing that inx2 also controls proliferation cell autonomously in the peripodial epithelium during eye disc development. The lateral PE of the eye-antennal disc has also been shown to contribute significantly to external cuticular structures in the adult fly head, including cuticle in the midline of the head, the vibrissae and ventral head structures such as the maxillary palps (Chouinard and Kaufman, 1991; Fristrom and Fristrom, 1993; Stultz et al., 2006; Lee et al., 2007; Atkins and Mardon, 2009). Upon inx2 downregulation in the whole eye disc (e.g. with eyGal4), we observed with a moderate penetrance (ca. 50% of the adult flies) and often only on one side, a lack of vibrissae in the ventral head (Fig. 9F). Taken together, our data show that Innexin2 and Innexin3 are required in the peripodial epithelium to sustain cell proliferation in the disc proper. Inx2 is also required cell autonomously for PE growth and its absence leads to cuticular defects. Our data also highlight the tight functional cooperation existing between Inx2 and Inx3 in the PE.

DISCUSSION Previous work has shown that the mRNA of inx1, inx2, inx3, inx6 and inx7 is found in pigment cells during pupal eye development (Stebbings et al., 2002). Our present analysis extends this observation at the protein level and throughout eye development for Inx1, Inx2 and Inx3 (Figs. 1, 2). We observe a strong colocalisation of Inx2 with Inx3 in eye discs throughout larval and pupal development (Figs. 1, 2). A similar colocalisation of Inx2 and Inx3 has already been observed in embryonic epithelia (Lehmann et al., 2006; Giuliani et al., 2013) and between soma cells of the ovary (Bohrmann and Zimmermann, 2008). Since Inx2 and Inx3 were demonstrated to directly interact via their C-termini and form heteromers in Drosophila embryos (Lehmann et al., 2006), those heteromers are likely to be present in the eye disc, too. An argument for the presence of heteromers comes from the fact that overexpression of an Inx2Myc transgene in the larval eye disc is able to recruit Inx3 to specific membrane domains (Suppl. Fig. 5B). In addition, Inx3 expression at the plasma membrane critically depends on inx2 throughout eye development (Fig. 6). In some cases, instead of the prominent membrane localisation of Inx3, we observed a faint cytoplasmic staining (Fig. 6A’ and B’), suggesting that the turnover of Inx3 might be affected in the absence of inx2. Connexin protein levels are known to be modified through differential gene transcription or altered through trafficking and degradation mechanisms (Su and Lau, 2014). In inx2 mutant eye discs, inx3 transcripts levels were not affected, excluding the possibility of a transcriptional control (Suppl. Fig. 5A) and suggesting that Inx2 might regulate Inx3 levels post-transcriptionally. Inx2 protein levels also depended on inx3 in larval eye discs (Fig. 7C’). A similar observation has been made in the amnioserosa, where a deficiency uncovering inx3 led to a 3-fold reduction in Inx2 levels at the plasma membrane (Giuliani et al., 2013). Since inx2 transcription was neither obviously affected in inx3 depleted eye discs (Suppl. Fig. 5A), it seems likely that heteromerisation of Inx2 and Inx3 is required for complex stability at the plasma membrane of the eye disc, as suggested previously for embryonic epithelia (Lehmann et al., 2006). Heteromerisation of Inx2 and Inx3 is known to be crucial for epithelial organisation and polarity of the embryonic epidermis (Lehmann et al., 2006). Our new observations strengthen the functional cooperation between Inx2 and Inx3 during development. We demonstrate here that a reduced expression of inx3 in the larval eye disc leads to small-sized eyes (Fig. 3) thus phenocopying the inx2 loss-of-function situation (Richard and Hoch, 2015). Conversely, inx3 overexpression increases eye size, phenocopying the inx2 gain-of-function situation (Richard and Hoch, 2015). Moreover, we show that inx3 (i) controls proliferation in the anterior compartment of L3 eye discs, (ii) is required for activation of the Dpp pathway at the MF and for expression of Dpp receptor Punt

in the disc as well as (iii) genetically interacts with the Dpp signaling pathway to control eye size (Figs. 4 and 5). All those observations were also made in inx2 mutant eye discs (Richard and Hoch, 2015). We also demonstrate that Inx2 and Inx3 expression rescues the small-eye phenotype of inx2 or inx3-depleted eyes (Fig. 8 and Suppl. Fig. 3C), suggesting that the formation of Inx2/Inx3 heteromers is required to sustain larval eye disc growth. Furthermore, our results point to a new role for Inx2/Inx3 channels during epithelial development. In contrast to the observations made in the embryo, where Inx2/Inx3 heteromers were shown to be required for epithelial maintenance (Lehmann et al., 2006), our results now demonstrate that Inx2 and Inx3 are required for epithelial proliferation. Further biochemical work will be required to unravel the molecular connection between gap junction and Dpp-dependent communication. Our studies thus identify Innexin3 as the second important gap junction protein besides Inx2 regulating Drosophila eye disc growth. In contrast, Inx1 is not a critical component in the control of eye size (Fig. 3) and though it colocalises with Inx2 at larval stages, its presence influences Inx2 levels only slightly (Fig. 7A’). In addition, overexpression of Inx1 does not rescue the small eye phenotype observed upon inx2 depletion during larval stages (Fig. 8). Interestingly, our work also points to a new crucial role for Innexins in the connection between peripodial epithelium and disc proper cells, inx2 and inx3 being two new genes among very few described to be involved in communication between both layers (Atkins and Mardon, 2009). We have shown previously that inx2 influences Dpp signaling and controls proliferation in a cellautonomous manner in the disc proper (Richard and Hoch, 2015). Here we show that inx2 also controls proliferation in the PE cell autonomously (Fig. 9B-B’’’, C). Furthermore, we show that a depletion of inx2 or inx3 in the PE impairs proper growth of the DP (Fig. 9D), leading to a smallsized eye in the adult (Fig. 9A). Moreover, inx2 depletion in the eye disc leads to malformed head structures (Fig. 9F), phenocopying head cuticle defects observed in flies with a disrupted dpp expression in the PE (Stultz et al., 2006). These last two observations suggest that Innexins might influence signal transduction pathways in the PE too. Our preliminary results show that inx2 is responsible for activation of the hedgehog pathway in the PE (Suppl. Fig. 4F’). Since Hedgehog signaling from the PE is required for DP growth (Cho et al., 2000), it might be that Innexins influence PE-DP communication indirectly. In this scenario, Innexins would regulate expression of ligands in the PE and those would then traffic between layers, in a not fully understood manner, to modify growth in the DP (Atkins and Mardon, 2009). What about the direct transmission of metabolites through gap junctions between both layers? We show here that Inx2 and Inx3 colocalise to the plasma membrane of eye peripodial epithelium cells at larval stages (Fig. 1E-E’’’, Suppl. Fig. 1B-B’’’ and 1C) and that DP cells express Inx2/Inx3 apico-laterally (Suppl. Fig. 1A-A’’’ and 1C).

Considering the fact that in many areas of the eye disc, a lumen separates the apical sides of both PE and DP (Pallavi and Shashidhara, 2003), it seems that there is no possibility for a direct gap junctional contact between both cell layers. However, gap junction proteins can also form hemichannels, allowing for release of metabolites, such as ATP, from the cells (Lohman and Isakson, 2014). Signalling from the vertebrate retinal pigment epithelium (RPE) is known to support growth, survival and patterning of the retina, and in zebrafish the RPE contributes cells to the retina (Raymond and Jackson, 1995; Li et al., 2000) mimicking the situation observed in Drosophila between PE and DP (Atkins and Mardon, 2009). It was shown that Connexin 43 hemichannels are required to release ATP from the RPE cells to speed up cell division and proliferation in the neural retina. In this system, opening of hemichannels is evoked by spontaneous elevations of Ca2+ in the RPE (Pearson et al., 2005). It could thus be that a similar mechanism exists in Drosophila: Inx2/Inx3 hemichannels would allow for ATP release from the peripodial epithelium to modulate proliferation in the disc proper. More experiments will be needed to address this question.

ACKNOWLEDGEMENTS

For providing fly stocks we thank M. Atkins, N. Bulgakova, G. Halder, E. Knust, G. Mardon, M. O’Connor and the Bloomington Stock Center. For providing antibodies we thank A. Brandt, P. ten Dijke, H. Sun and A-S. Chiang. For providing plasmids we thank R.W. Carthew. We thank F. Eckardt for technical support and all the Hoch lab members for discussion. We thank Katinka Ostrowski for the cloning of UAS-Inx1 and UAS-RNAi inx1 constructs. This work was supported by grants from the DFG to M.H. (SFB 645, TRR83, Excellence Cluster ImmunoSensation) and the Helmholtz cross program topic “Metabolic Dysfunction” as well as by a grant from the DFG to R.B. (SFB 645) and to G.T. (SPP 1464).

AUTHOR CONTRIBUTION STATEMENTS

MH and MR conceived the study. MR performed all experiments. RB designed the UAS-Innexin1 and UAS-RNAi Inx1 1 and 2 fly lines. MH and MR designed and analysed the experiments. MH, MR, GT and RB discussed the data. MH and MR wrote the manuscript.

REFERENCES Atkins, M., Mardon, G., 2009. Signaling in the third dimension: the peripodial epithelium in eye disc development. Dev. Dyn. 238, 2139–48. Auerbach, C., 1936. The development of the legs, wings and halteres in wild type and some mutant strains of Drosophila melanogaster. Trans. R. Soc. Edin. LVIII, Par. Barbe, M.T., Monyer, H., Bruzzone, R., 2006. Cell-cell communication beyond connexins: the pannexin channels. Physiology (Bethesda). 21, 103–14. Bauer, R., Lehmann, C., Martini, J., Eckardt, F., Hoch, M., 2004. Gap junction channel protein innexin 2 is essential for epithelial morphogenesis in the Drosophila embryo. Mol. Biol. Cell 15, 2992–3004. Bauer, R., Löer, B., Ostrowski, K., Martini, J., Weimbs, A., Lechner, H., Hoch, M., 2005. Intercellular communication: the Drosophila innexin multiprotein family of gap junction proteins. Chem. Biol. 12, 515–26. Bauer, R., Weimbs, A., Lechner, H., Hoch, M., 2006. DE-Cadherin, a Core Component of the Adherens Junction Complex Modifies Subcellular Localization of the Drosophila Gap Junction Protein Innexin2. Cell Commun. Adhes. 13, 103–114. Bessa, J., Gebelein, B., Pichaud, F., Casares, F., Mann, R.S., 2002. Combinatorial control of Drosophila eye development by eyeless, homothorax, and teashirt. Genes Dev. 16, 2415– 27. Bohrmann, J., Zimmermann, J., 2008. Gap junctions in the ovary of Drosophila melanogaster: localization of innexins 1, 2, 3 and 4 and evidence for intercellular communication via innexin-2 containing channels. BMC Dev. Biol. 8, 111. Brand, A. H., Perrimon, N. 1993. Targeted gene expression as a means of altering cell fates and generating dominant phenotypes. Development 118, 401-15. Bui, Q.T., Zimmerman, J.E., Liu, H., Gray-Board, G.L., Bonini, N.M., 2000. Functional analysis of an eye enhancer of the Drosophila eyes absent gene: differential regulation by eye specification genes. Dev. Biol. 221, 355–64. Bulgakova, N.A., Rentsch, M., Knust, E., 2010. Antagonistic functions of two stardust isoforms in Drosophila photoreceptor cells. Mol. Biol. Cell 21, 3915–3925. Burke, R., Basler, K., 1996. Hedgehog-dependent patterning in the Drosophila eye can occur in the absence of Dpp signaling. Dev. Biol. 179, 360–8. Cagan, R.L., Ready, D.F., 1989. The emergence of order in the Drosophila pupal retina. Dev. Biol. 136, 346–62. Childs, S.R., Wrana, J.L., Arora, K., Attisano, L., O’Connor, M.B., Massagué, J., 1993. Identification of a Drosophila activin receptor. Proc. Natl. Acad. Sci. U. S. A. 90, 9475–9. Cho, K.O., Chern, J., Izaddoost, S., Choi, K.W., 2000. Novel signaling from the peripodial

membrane is essential for eye disc patterning in Drosophila. Cell 103, 331–42. Chouinard, S., Kaufman, T., 1991. Control of expression of the homeotic labial (lab) locus of Drosophila melanogaster: evidence for both positive and negative autogenous regulation. Development 113, 1267–1280. Cohen, S.M., 1993. Imaginal disc development., in: Bate, M., Martinez Arias, A. (Eds.), The Development of Drosophila Melanogaster. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, pp. 747–841. Costanzo, M., Baryshnikova, A., Myers, C.L., Andrews, B., Boone, C., 2011. Charting the genetic interaction map of a cell. Curr. Opin. Biotechnol. 22, 66–74. Dahl, G., Locovei, S., 2006. Pannexin: to gap or not to gap, is that a question? IUBMB Life 58, 409–19. Firth, L.C., Bhattacharya, A., Baker, N.E., 2010. Cell cycle arrest by a gradient of Dpp signaling during Drosophila eye development. BMC Dev. Biol. 10, 28. Fristrom, D., Fristrom, J., 1993. The Metamorphic Development of the Adult Epidermis., in: Bate, M., Martinez-Arias, A. (Eds.), The Development of Drosophila Melanogaster. Cold Spring Harbor Laboratory Press., Cold Spring Harbor, NY, pp. 843–897. Gibson, M.C., Schubiger, G., 2000. Peripodial cells regulate proliferation and patterning of Drosophila imaginal discs. Cell 103, 343–50. Giuliani, F., Giuliani, G., Bauer, R., Rabouille, C., 2013. Innexin 3, a new gene required for dorsal closure in Drosophila embryo. PLoS One 8, e69212. Goodenough, D.A., Goliger, J.A., Paul, D.L., 1996. Connexins, connexons, and intercellular communication. Annu. Rev. Biochem. 65, 475–502. Hazelett, D.J., Bourouis, M., Walldorf, U., Treisman, J.E., 1998. decapentaplegic and wingless are regulated by eyes absent and eyegone and interact to direct the pattern of retinal differentiation in the eye disc. Development 125, 3741–51. Holcroft, C.E., Jackson, W.D., Lin, W.-H., Bassiri, K., Baines, R.A., Phelan, P., 2013. Innexins Ogre and Inx2 are required in glial cells for normal postembryonic development of the Drosophila central nervous system. J. Cell Sci. 126, 3823–34. Hudson, J., Podos, S., Keith, K., Simpson, S., Ferguson, E., 1998. The Drosophila Medea gene is required downstream of dpp and encodes a functional homolog of human Smad4. Development 125, 1407–1420. Kumar, J.P., 2011. My what big eyes you have: how the Drosophila retina grows. Dev. Neurobiol. 71, 1133–52. Lechner, H., Josten, F., Fuss, B., Bauer, R., Hoch, M., 2007. Cross regulation of intercellular gap junction communication and paracrine signaling pathways during organogenesis in Drosophila. Dev. Biol. 310, 23–34. Lee, H., Stultz, B.G., Hursh, D.A., 2007. The Zic family member, odd-paired, regulates the

Drosophila BMP, decapentaplegic, during adult head development. Development 134, 1301–10. Lee, T., Luo, L., 1999. Mosaic Analysis with a Repressible Cell Marker for Studies of Gene Function in Neuronal Morphogenesis. Neuron 22, 451–461. Lee, Y. S., Carthew, R. W., 2003. Making a better RNAi vector for Drosophila: use of intron spacers. Methods 30, 322-9. Lehmann, C., Lechner, H., Löer, B., Knieps, M., Herrmann, S., Famulok, M., Bauer, R., Hoch, M., 2006. Heteromerization of innexin gap junction proteins regulates epithelial tissue organization in Drosophila. Mol. Biol. Cell 17, 1676–85. Letsou, A., Arora, K., Wrana, J.L., Simin, K., Twombly, V., Jamal, J., Staehling-Hampton, K., Hoffmann, F.M., Gelbart, W.M., Massagué, J., O’Connor, M.B., 1995. Drosophila Dpp signaling is mediated by the punt gene product: A dual ligand-binding type II receptor of the TGFβ receptor family. Cell 80, 899–908. Li, Z., Joseph, N.M., Easter, S.S., 2000. The morphogenesis of the zebrafish eye, including a fate map of the optic vesicle. Dev. Dyn. 218, 175–88. Lim, J., Choi, K.-W., 2004. Drosophila eye disc margin is a center for organizing long-range planar polarity. Genesis 39, 26–37. Livak, K.J., Schmittgen, T.D., 2001. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) Method. Methods 25, 402–8. Lohman, A.W., Isakson, B.E., 2014. Differentiating connexin hemichannels and pannexin channels in cellular ATP release. FEBS Lett. 588,1379-88. Marinho, J., Martins, T., Neto, M., Casares, F., Pereira, P.S., 2013. The nucleolar protein Viriato/Nol12 is required for the growth and differentiation progression activities of the Dpp pathway during Drosophila eye development. Dev. Biol. 377, 154–65. Martin, P.E.M., Evans, W.H., 2004. Incorporation of connexins into plasma membranes and gap junctions. Cardiovasc. Res. 62, 378–87. Masucci, J.D., Miltenberger, R.J., Hoffmann, F.M., 1990. Pattern-specific expression of the Drosophila decapentaplegic gene in imaginal disks is regulated by 3’ cis-regulatory elements. Genes Dev. 4, 2011–23. McClure, K.D., Schubiger, G., 2005. Developmental analysis and squamous morphogenesis of the peripodial epithelium in Drosophila imaginal discs. Development 132, 5033–42. Morrison, C.M., Halder, G., 2010. Characterization of a dorsal-eye Gal4 Line in Drosophila. Genesis 48, 3–7. Neufeld, T.P., Hariharan, I.K., 2002. Regulation of Growth and Cell Proliferation During Eye Development, in: Moses, K. (Ed.), Drosophila Eye Development, Results and Problems in Cell Differentiation. Springer Berlin Heidelberg, Berlin, Heidelberg, pp. 107–133. Oshima, A., Matsuzawa, T., Murata, K., Tani, K., Fujiyoshi, Y., 2016. Hexadecameric structure of

an invertebrate gap junction channel. J. Mol. Biol. 428, 1227–36. Ostrowski, K., Bauer, R., Hoch, M., 2008. The Drosophila innexin 7 gap junction protein is required for development of the embryonic nervous system. Cell Commun. Adhes. 15, 155–67. Pallavi, S.K., Shashidhara, L.S., 2003. Egfr/Ras pathway mediates interactions between peripodial and disc proper cells in Drosophila wing discs. Development 130, 4931–41. Panchin, Y., Kelmanson, I., Matz, M., Lukyanov, K., Usman, N., Lukyanov, S., 2000. A ubiquitous family of putative gap junction molecules. Curr. Biol. 10, R473–4. Pearson, R.A., Dale, N., Llaudet, E., Mobbs, P., 2005. ATP released via gap junction hemichannels from the pigment epithelium regulates neural retinal progenitor proliferation. Neuron 46, 731–44. Phelan, P., 2005. Innexins: members of an evolutionarily conserved family of gap-junction proteins. Biochim. Biophys. Acta 1711, 225–45. Phelan, P., Starich, T.A., 2001. Innexins get into the gap. Bioessays 23, 388–96. Pilot, F., Lecuit, T., 2005. Compartmentalized morphogenesis in epithelia: from cell to tissue shape. Dev. Dyn. 232, 685–94. Raftery, L.A., Twombly, V., Wharton, K., Gelbart, W.M., 1995. Genetic screens to identify elements of the decapentaplegic signaling pathway in Drosophila. Genetics 139, 241–54. Raftery, L.A., Wisotzkey, R.G., 2008. Characterization of Medea, A Gene Required for Maximal Function of the Drosophila BMP Homolog Decapentaplegic. Ann. N. Y. Acad. Sci. 785, 318– 320. Raymond, S.M., Jackson, I.J., 1995. The retinal pigmented epithelium is required for development and maintenance of the mouse neural retina. Curr. Biol. 5, 1286–1295. Ready, D.F., Hanson, T.E., Benzer, S., 1976. Development of the Drosophila retina, a neurocrystalline lattice. Dev. Biol. 53, 217–40. Richard, M., Grawe, F., Knust, E., 2006. DPATJ plays a role in retinal morphogenesis and protects against light-dependent degeneration of photoreceptor cells in the Drosophila eye. Dev. Dyn. 235, 895–907. Richard, M., Hoch, M., 2015. Drosophila eye size is determined by Innexin 2-dependent Decapentaplegic signalling. Dev. Biol. 408, 26–40. Ruberte, E., Marty, T., Nellen, D., Affolter, M., Basler, K., 1995. An absolute requirement for both the type II and type I receptors, punt and thick veins, for dpp signaling in vivo. Cell 80, 889–97. Rubin, G.M., Spradling, A.C, 1982. Genetic transformation of Drosophila with transposable element vectors. Science 218, 348-353. Sáez, J.C., Leybaert, L., 2014. Hunting for connexin hemichannels. FEBS Lett. 588, 1205–11.

Scemes, E., 2012. Nature of plasmalemmal functional “hemichannels”. Biochim. Biophys. Acta 1818, 1880–3. Segretain, D., Falk, M.M., 2004. Regulation of connexin biosynthesis, assembly, gap junction formation, and removal. Biochim. Biophys. Acta 1662, 3–21. Sekelsky, J.J., Newfeld, S.J., Raftery, L.A., Chartoff, E.H., Gelbart, W.M., 1995. Genetic characterization and cloning of mothers against dpp, a gene required for decapentaplegic function in Drosophila melanogaster. Genetics 139, 1347–58. Söhl, G., Willecke, K., 2003. An update on connexin genes and their nomenclature in mouse and man. Cell Commun. Adhes. 10, 173–80. Spéder, P., Brand, A.H., 2014. Gap junction proteins in the blood-brain barrier control nutrientdependent reactivation of Drosophila neural stem cells. Dev. Cell 30, 309–21. Stebbings, L.A., Todman, M.G., Phelan, P., Bacon, J.P., Davies, J.A., 2000. Two Drosophila Innexins Are Expressed in Overlapping Domains and Cooperate to Form Gap-Junction Channels. Mol. Biol. Cell 11, 2459–2470. Stebbings, L.A., Todman, M.G., Phillips, R., Greer, C.E., Tam, J., Phelan, P., Jacobs, K., Bacon, J.P., Davies, J.A., 2002. Gap junctions in Drosophila: developmental expression of the entire innexin gene family. Mech. Dev. 113, 197–205. Stowers, R.S., Schwarz, T.L., 1999. A genetic method for generating Drosophila eyes composed exclusively of mitotic clones of a single genotype. Genetics 152, 1631–1639. Stultz, B.G., Lee, H., Ramon, K., Hursh, D.A., 2006. Decapentaplegic head capsule mutations disrupt novel peripodial expression controlling the morphogenesis of the Drosophila ventral head. Dev. Biol. 296, 329–39. Su, V., Lau, A.F., 2014. Connexins: mechanisms regulating protein levels and intercellular communication. FEBS Lett. 588, 1212–20. Tepass, U., Hartenstein, V., 1994. The development of cellular junctions in the Drosophila embryo. Dev. Biol. 161, 563–96. Vrailas, A.D., Moses, K., 2006. Smoothened, thickveins and the genetic control of cell cycle and cell fate in the developing Drosophila eye. Mech. Dev. 123, 151–65. Wei, C.-J., Xu, X., Lo, C.W., 2004. Connexins and cell signaling in development and disease. Annu. Rev. Cell Dev. Biol. 20, 811–38. Wolff, T., Ready, D.F., 1991. The beginning of pattern formation in the Drosophila compound eye: the morphogenetic furrow and the second mitotic wave. Development 113, 841–50. Wrana, J.L., Tran, H., Attisano, L., Arora, K., Childs, S.R., Massagué, J., O’Connor, M.B., 1994. Two distinct transmembrane serine/threonine kinases from Drosophila melanogaster form an activin receptor complex. Mol. Cell. Biol. 14, 944–50. Wu, C.-L., Shih, M.-F.M., Lai, J.S.-Y., Yang, H.-T., Turner, G.C., Chen, L., Chiang, A.-S., 2011. Heterotypic gap junctions between two neurons in the drosophila brain are critical for

memory. Curr. Biol. 21, 848–54. Yeager, M., Unger, V.M., Falk, M.M., 1998. Synthesis, assembly and structure of gap junction intercellular channels. Curr. Opin. Struct. Biol. 8, 517–524.

MATERIALS AND METHODS Cloning of pUAST-Inx1-Myc and pWiz-Inx1 pUAST-Innexin1-Myc was generated by cloning the open reading frame of inx1-RA (CG3039), cterminally tagged with a Myc-tag in the pUAST vector (Brand and Perrimon, 1993). The insert was amplified by PCR using the following primers: GGA ATT C ATG TAT AAG TTG CTG GGT AGC CTG as a forward primer containing an EcoRI restriction site and GCT CTA GAG CTA CAG ATC CTC TTC AGA GAT GAG TTT CTG CTC CTT GGC ACG GTC GTG as a reverse primer containing an XbaI restriction site, stop codon and Myc-tag. Selected constructs were injected into w1118 fly embryos (Rubin and Spradling, 1982; Brand and Perrimon, 1993). Cloning of recombinant plasmids for the generation of stable RNAi-inducible fly strains was performed as described by Lee and Carthew (2003). A 300-bp fragment of inx1-RA (bases 1199 to 1499) was amplified by polymerase chain reaction (PCR), with PCR primers containing at their 5’ ends a XbaI restriction site, which is compatible with the AvrII and NheI sites (forward primer: GCT CTA GAT GTA CGC CAT TCG GTA CTG, reverse primer: GCT CTA GAT CCG ATG AGC AGG ACG A). The described insert was cloned in two different orientations (“tail to tail”) into the AvrII and NheI site of pWiz (a gift from R.W. Carthew), which was dephosphorylated with alkaline shrimp phosphatase prior to ligation. For transformation, competent SURE cells were used (Stratagene). Recombinants in “tail to tail” orientation were screened, selected and injected into w1118 fly embryos.

Fly strains Flies were kept on standard medium at 25°C. Following fly stocks were used: w1118, OregonR, a inx2 null mutant line w ΔP16 FRT19A/FM7i (Richard and Hoch, 2015), a inx1 null mutant line (inx1 KO, (Giuliani et al., 2013)). To produce small inx2 mutant clones for stainings in larval eye discs or pupal eyes, females of the genotype Ubi-mRFP.nls w hsFLP FRT19A (Bloomington #31418) were crossed to w ΔP16 FRT19A/ Dp(1;Y) ct+ y+ males (Richard and Hoch, 2015) and heat-shocked twice for 2 hours at 37,8°C on day 2 and 3 after egg laying. For measuring the size of inx2 clones and their twin clones, a single heat-shock of 1 hour was performed on day 3 after egg laying. For MARCM experiments (Lee and Luo, 1999), hsFLP tub-Gal80 FRT19A; act-Gal4 UASCD8-GFP/CyO females (Bulgakova et al., 2010) were crossed to w ΔP16 FRT19A/ Dp(1;Y) ct+ y+ males and heat-shocked twice for 2 hours at 37,8°C on day 2 and 3 after egg laying. For producing nearly complete mutant larval eye discs for inx2, the EGUF/hid technique was applied (Stowers and Schwarz, 1999)⁠. yw GMR-hid FRT19A, l(1)CL[1]/FM7 KrGal4 UAS-GFP;

eyFLP females (adapted from (Stowers and Schwarz, 1999)⁠, a gift of N. Bulgakova, Cambridge, UK) were crossed with males of the genotype w ΔP16 FRT19A/ Dp(1;Y) ct+ y+ (this duplication is available e.g. in Bloomington #5280) for producing inx2 mutant eye discs or w FRT19A/Y for control eye discs and selected female third instar larvae without GFP expression under a Zeiss Discovery.V8 binocular. The UAS-innexin1-Myc (UAS inx1) transgenic flies were generated by injection of the pUASTInx1-Myc construct into w1118 fly embryos and selected by eye color (Rubin and Spradling, 1982; Brand and Perrimon, 1993). Expression of the Myc-tagged construct was verified in the eye disc (Suppl. Fig. 5C-C’’). UAS RNAi inx1 1 and 2 transgenic fly lines were generated by injecting pWizInx1 construct into w1118 fly embryos (Rubin and Spradling, 1982; Brand and Perrimon, 1993). We also used following UAS lines: UAS-CD8-GFP (Bloomington #5136), two insertions of UAS inx2-Myc (Bauer et al., 2006), UAS RNAi inx2 (Lechner et al., 2007), UAS inx3 (Stebbings et al., 2000), UAS RNAi inx3 (A9 and BA14 are two different inx3 RNAi lines produced in our lab, (Lehmann et al., 2006)), UAS RNAi inx1 (103816 and 7136 are VDRC IDs, Vienna), UAS RNAi inx6 (8638 and 17690 are VDRC IDs), UAS RNAi inx7 1 (Ostrowski et al., 2008) and UAS RNAi inx7 103256 (VDRC, Vienna), medea RNAi (VDRC # 19688). Following Gal4 drivers were used in this study: eyGal4 (Bloomington #8227), eyaGal4 (Bui et al., 2000), DEGal4 (Morrison and Halder, 2010) (a gift from G. Halder, Houston/TX, USA), c311Gal4 (Bloomington #5937), optixGal4 (gift from M. Atkins and G.Mardon, from Kyoto #NP2631).

Antibodies, immunofluorescence analyses and imaging Dissection of pupal eyes from staged pupae and staining was performed as previously described (Richard et al., 2006). Larval eye discs were dissected in PBS and fixed for 1 hour in 4% PFA. After an overnight incubation with primary antibodies in PBS Tween 0,1% BSA 0,1% discs were washed in PBS Tween 0,1% and incubation was pursued in secondary antibodies in PBST BSA 0,1% for 2 hours before mounting in Fluoromount-G (Southern Biotech, Birmingham, AL). Primary antibodies were as follows: anti-Innexin 1 (rabbit, 1:40 used for Fig.2 and Fig.6 or guinea pig, 1:100 used in Figs. 1, 7 and Suppl. Fig. 1) (Bauer et al., 2004; Spéder and Brand, 2014), antiInnexin 2 (rabbit or guinea pig, 1:50) (Bauer et al., 2004; Bohrmann and Zimmermann, 2008), antiInnexinx3 (rabbit or guinea pig, 1:50) (Bohrmann and Zimmermann, 2008; Lehmann et al., 2006), anti-Armadillo (mouse, 1:10; Hybridoma Bank), anti-DE-cadherin (goat, 1:10; Santa Cruz Biotechnology), anti-GFP (mouse,1:100; Santa Cruz Biotechnology), anti-RFP (mouse or rabbit, 1:100 and 1:500 respectively, Abcam), anti-activated Caspase3 (rabbit, 1:20, Cell Signalling), anti-

phospho-Histone H3 (mouse, 1:500, Cell Signalling), anti-Myc (mouse or rabbit, 1:50, Santa Cruz Biotechnology), anti-Innexin6 (rabbit, 1:2500, (Wu et al., 2011)), anti-Innexin7 (rabbit, 1:100, (Ostrowski et al., 2008)), anti-full-length Ci (2A1, rat, 1:3, DSHB), anti-phospho Mad (rabbit, 1:1000, gift from P. ten Dijke, Leiden, Netherlands), anti-Punt (rabbit, 1:200, Abcam), anti-Hth (goat, 1:50, dG20, gift from Henry Sun, Taiwan, also available at Santa Cruz Biotechnology). Secondary antibodies recognizing different species were bought coupled to Alexa Fluor 488 (1:200; MoBiTec, Goettingen, Germany), Cy3 (1:200; Dianova, Hamburg, Germany), and Alexa Fluor 633 (1:200, MoBiTec, Goettingen, Germany). For detecting F-actin, Phalloidin Alexa Fluor 488 or 633 (1:100, Molecular Probes, Invitrogen) was added with the secondary antibodies. Fluorescent images were recorded using a confocal microscope (Zeiss LSM 710), and images of multilabeled samples were acquired sequentially on separate channels. All images were processed with the Adobe Photoshop software. For producing the intensity plots of pMad and Punt levels, we used the “Interactive 3D surface plot” Plugin from the ImageJ freeware (http://imagej.nih.gov/ij). Pictures of adult eyes were taken with an Olympus AX70 binocular. Image processing and eye measurements were performed with the ImageJ freeware (http://rsbweb.nih.gov/ij) and Adobe Photoshop. To measure adult eye size, the dorso-ventral length on its longer segment (from the top to the bottom of the eye) was measured. Control eyes measured values were considered as 100%. For analysing mitotic figures in the eye disc, the number of phospho-Histone H3 positive cells in the anterior disc compartment (anterior to the furrow) of c311Gal4 UAS RNAi inx2 or control flies (Fig. 9D) was divided by the surface of the anterior compartment (all pictures with the same magnification). Alternatively, for downregulation of inx3 (Fig. 4C), the number of phospho-Histone H3 positive cells counts in the dorsal or ventral area of the anterior compartment were divided by the surface of the respective compartment. For quantifying apoptosis in the eye disc, the number of activated caspase 3-positive cells in the anterior disc compartment (anterior to the furrow) of eyaGal4 UAS RNAi inx3 or control flies (Fig. 4E, F) was divided by the surface of the anterior compartment (all pictures with the same magnification). For quantifying Innexin levels in the dorsal vs. ventral area of the eye disc upon dorsal knock-down using DEGal4 (Fig. 7, Suppl. Fig. 3A), we defined an area (of a size of approximately one fourth of the anterior part of the disc) and measured pixel intensity with ImageJ either on the ventral anterior side (control) or on the dorsal anterior side (knock-down area), paying attention to choose areas of similar disc flattening. Values correspond to pixel intensity of the depleted area divided by the pixel intensity of the control area (as a percentage of control). Five discs were analysed for each

condition and mean±sd is presented in Fig. 7 or Suppl. Fig. 3A. For quantifying Innexin levels in the anterior compartment of optixGal4 UAS RNAi inx2 eye discs or their controls (Suppl. Fig. 4B-B’’’ and C-C’’’), we used the exact same confocal settings for taking all pictures and defined an area corresponding to ca. two third of the anterior compartment for measuring pixel intensity with ImageJ. For c311Gal4 UAS RNAi inx2 discs and their controls (Suppl. Fig. 4D-D’’’ and E-E’’’), we used the same confocal settings as for optixGal4 discs and pixel intensity was measured in an area corresponding to that of the picture seen in Fig. 4D. Values mentioned in the text correspond to pixel intensity of the inx2-depleted discs divided by the pixel intensity of the control discs (as a percentage of control). Five discs were analysed and values are mean±sd. For measuring the size of inx2 clones and their corresponding twin clones (Fig.9 C) we used the area measuring tool of ImageJ. We checked that the area of the clone corresponded to its number of cells, counting the cells of each clone/twin clone using the Hth staining (nuclei) or the DE-Cadherin staining (cell membranes). Statistical

analyses

were

done

using

Microsoft

Excel

and

the

GraphPad

freeware

(www.graphpad.com) for t Tests, as well as http://statpages.info/anova1sm.html for one-way ANOVA and their post-hoc tests.

Semi-thin sections Sections were prepared according to (Tepass and Hartenstein, 1994)⁠ with modifications. In brief, 0.1 M phosphate buffer (pH 7.4) was used to fix bisected heads in 25% glutaraldehyde, followed by simultaneous fixation in 1% osmium tetroxide/2% glutaraldehyde, followed by 2% OsO4. After dehydration, eyes were embedded in Araldite, and semi-thin (2.5 μm) sections were cut on a Leica RM2255 microtome and stained with toluidine blue. Imaging was done in Adobe Photoshop.

Quantitative real-time RT-PCR For RNA isolation, we collected 30-60 3rd instar eye discs (without antennal discs) in lysis buffer. Total RNA was isolated using the NucleoSpin RNA II kit (Macherey & Nagel). RNA concentration was measured with a NanoDrop spectrophotometer (Thermo Scientific). For first strand cDNA synthesis, ±100 ng of total RNA was transcribed using the QuantiTect Reverse Transcription Kit (Qiagen) including DNaseI treatment and following supplier’s protocol. Real-time PCR was performed with iQ SYBR Green Supermix (Biorad) and analyzed on a CFX Connect Real-time

PCR Detection System (Biorad). cDNA samples were run in triplicates. Average CTs were used to analyze the expression levels with the 2(-ΔΔCT) method (Livak and Schmittgen, 2001)⁠. Experiments were repeated with at least 2 different RNA samples. Expression analysis was performed using CFX Manager software (Biorad) and Microsoft Excel. Actin 5C and ribosomal protein L32 were used as reference genes. Primer selection was done using Primer 3 plus (http://www.bioinformatics.nl/cgibin/primer3plus/primer3plus.cgi/) and primers were selected to analyze all possible transcripts and to lie outside of the region targeted by the RNAi. Following

oligonucleotides

were

used:

ribosomal

protein

L32

forward

CTAAGCTGTCGCACAAATG, ribosomal protein L32 reverse GTTCGATCCGTAACCGATGT, actin5C

forward

GTGCACCGCAAGTGCTTCTAA,

actin

5C

reverse

TGCTGCACTCCAAACTTCCAC, inx2 forward CCTACTCCGAGCCCGTTCC, inx2 reverse TGCCCAGCTGATAGAGCAGG, inx3 forward GATCGGTCCAGAAACACGACA, inx3 reverse GAGATGGTGGCCAAGATGAT.

FIGURE LEGEND

Figure 1: Expression and colocalisation of Inx1, Inx2 and Inx3 during larval eye disc development. Single optical sections of wild-type larval eye discs at the same apical level in the disc proper (AA’’’ to D-D’’’) or in the peripodial epithelium (E-E’’’, F-F’’’). In all images, posterior is to the left and dorsal to the top. The morphogenetic furrow is indicated (mf). PE=peripodial epithelium, DP= disc proper. The image in the fourth column is a merge picture of stainings in the first two columns. All scale bars = 10 m. A-A’’’: cross-section through a L3 eye disc at the level of the DP. The region anterior to the mf (mf highlighted by DE-Cadherin in blue, A’’) was imaged. Inx2 (green, A) and Inx3 (magenta, A’) colocalise cortically in all DP cells (A’’’). B-B’’’: same disc as in A-A’’’ but the region posterior to the mf (DE-Cadherin in blue, B’’) was imaged. Inx2 (green, B) and Inx3 (magenta, B’) colocalise in the uncommitted progenitors between the recruited photoreceptor cell clusters (*, B’’’). C-C’’’: cross-section in a L3 eye disc at the level of the DP. The region anterior to the mf (mf highlighted by DE-Cadherin in blue, C’’) was imaged. Inx2 (green, C) and Inx1 (magenta, C’) colocalise in DP cells (C’’’). The asterisk marks an unspecifically stained nucleus. D-D’’’: same disc as in C-C’’’ but the region posterior to the mf was imaged. Inx2 (green, D) and Inx1 (magenta, D’) colocalise to the uncommitted progenitors between the recruited photoreceptor cell clusters (*, D’’’). E-E’’’: cross-section through a L3 eye disc at the level of the PE. Inx2 (green, E) and Inx3 (magenta, E’) colocalise cortically in all PE cells (E’’’). F-F’’’: cross-section through a L3 eye disc at the level of the PE. Inx2 (green, F) and Inx1 (magenta, F’) colocalise in PE cells (F’’’).

Figure 2: Expression and colocalisation of Inx1, Inx2 and Inx3 in the pupal eye. The image in the third column is a merge picture of stainings in the first two columns. A-A’’, B-B’’: optical sections through a pupal eye at about 45-50% pupal development stained for F-actin (green) and Inx3 (magenta). A-A’’: cross section. Secondary (sp) and tertiary (tp) pigment cells as well as bristle cells (b) are located between each photoreceptor cell cluster (PRCs

rhabdomeres are highlighted with F-actin, arrowhead) (A). Inx3 (A’) accumulates at the contact point between secondary (sp) and tertiary (tp) pigment cells and to a minor extent between secondary pigment cells and bristle cells (b). B-B’’: longitudinal section of the pupal eye displaying photoreceptor cell clusters (rhabdomeres showed by arrowheads in B) ensheathed in pigment cells (p) in the upper part of the picture and cross section at the retinal floor plate in the lower part of the picture showing the regular arrangement of the pigment cell feet (pf). Inx3 is expressed in the pigment cells. C-C’’: cross section through a pupal eye at about 45-50% pupal development stained for F-actin (green) and Inx1 (magenta). Rhabdomeres of a PRCs cluster are marked with an arrowhead. Inx1 (C’) accumulates at the contact point between secondary and tertiary (tp) pigment cells and less between secondary pigment cells and bristle cells (b). D-D’’: optical cross section of a pupal eye at 45% p.d. Inx2 (magenta, D) colocalises (D’’) with Inx3 (green, D’) in the pigment cells (tp, tertiary pigment cells). E-E’’: optical cross section of a pupal eye at 45% p.d. Inx2 (magenta, E) colocalises (E’’) with Inx1 (green, E’) in the pigment cells (tp, tertiary pigment cells). F-F’’ and G-G’’: optical sections of late pupal eyes (70% p.d. for F-F’’ and 90% p.d. for G-G’’) stained for Armadillo (white), highlighting the adherens junctions, and Inx3 (magenta). F-F’’: cross section showing that the forming rhabdomeres of the photoreceptor cells (rh in F), located apically to adherens junctions between photoreceptor cells (Arm dots in F), express Inx3 (F’). At this stage, Inx3 is still expressed in the pigment cells (tp in F’). G-G’’: longitudinal section through the retina, cornea is to the left (c in G’’). Inx3 localises both in the pigment cells (p) and in the rhabdomeres of the photoreceptor cells (rh). H-H’’: cross-section through the retina of a late pupal eye (ca. 85% p.d.) stained for F-Actin (green, H) and Inx1 (magenta, H’). Inx1 localises in the pigment cells (tp are indicated in H’) between the photoreceptor cell clusters (rh in H). I-I’’: longitudinal section through the retina of a late pupal eye (ca. 90% p.d.) stained for F-Actin (green, I) and Inx1 (magenta, I’). Inx1 localises to the pigment cells (p in I’) between the photoreceptor cell clusters (rh in I). Cornea is to the left (c in I). J-J’’: longitudinal section through the retina of a late pupal eye (ca. 90% p.d.) stained for Inx2 (green, J) and Inx3 (magenta, J’), cornea is to the left (c). Inx3 partially colocalises with Inx2 in the pigment cells (p in J’) but is strongly expressed in the rhabdomeres of the PRCs (rh in J’) where Inx2 is not expressed.

K-K’’: optical cross section through a late pupal retina just below the cornea. Inx2 (green, K) is expressed in the membrane of primary pigment cells (pp) and in secondary, tertiary pigment and bristle cells (sp, tp and b). Inx3 (magenta, K’) colocalises with Inx2 in pigment and bristle cells (K’’). All scale bars = 10 m.

Figure 3: inx3 depletion in the eye disc decreases adult eye size. Pictures of eyes from flies expressing different inx1, inx3, inx6 and inx7 RNAi lines under the control of eyaGal4 or mutant for inx1. The control for the knock-down eyes is eyaGal4/+ and is presented three times to easily appreciate size differences in the knock-downs. The control for the last panel (inx1 KO) is w (penultimate panel). Scale bar for all eye pictures = 250 m. The bottom graph presents the mean adult eye size of eyaGal4/+ flies (control, first column) or flies expressing different RNAi constructs against inx1, inx3, inx6 and inx7 under the control of eyaGal4. Mean eye size of a null mutant for inx1 is presented on the right-hand side (the control is w and presented in the penultimate column). The dorso-ventral length on its longer segment (from the top to the bottom of the eye) was measured. Values represent mean ± s.d. and the controls (first and penultimate columns) were independently considered as 100%. n values from the left to the right column: 17, 13, 14, 10, 12, 11, 13, 12, 12, 11, 9, 10, 11. ***: extremely statistically significant in Unpaired t test (P<0,001).

Figure 4: inx3 controls proliferation in the eye disc. A, B: Semi-thin sections of eyGal4/+ (A) and eyGal4/UAS RNAi inx3-BA14 (B) adult eyes. Morphology and ommatidium size is not affected in inx3-depleted eyes. Scale bars=10 μm. C and D-D’’’: Proliferation is decreased in inx3-depleted third instar eye discs. D-D’’’: Optical section of a DEGal4 UAS RNAi inx3-BA14 third instar larval eye disc with a depletion for inx3 (green in D’) in the dorsal part of the disc (arrowhead in D’) and stained with DE-Cadherin (D’’) and phospho-histone H3 (red in D) monitoring proliferating cells. The anterior dorsal area of the disc (arrowhead in D) shows less proliferating cells as the ventral control area. Scale bar = 50 μm. C: Proliferation counts anterior to the morphogenetic furrow in third instar DEGal4 UAS RNAi inx3-BA14 eye discs. Left column represents mean± s.e.m. of pH3 histone counts in the ventral control area of the discs while right column is mean±s.e.m. of counts in the dorsal inx3-depleted area. Values in the columns indicate mean±s.e.m. and number of discs (n).*: statistically significant

in Unpaired t test (P<0.05). E-E’ and F-F’: Optical sections of third instar eyaGal4/+ control (E,E’) or eyaGal4 UAS RNAi inx3-BA14 (F,F’) larval eye discs stained with DE-Cadherin (E,F) and activated caspase 3 (E’, F’) monitoring apoptotic cells. Scale bar=50 μm.

Figure 5: Dpp signalling is affected in inx3-depleted eye discs and inx3 interacts with the Dpp pathway to control eye size. A-A’’’ to C-C’’’: Optical sections through 3rd instar eye discs of DEGal4 UAS RNAi inx3-BA14 flies. Posterior is to the left, ventral to the bottom. Scale bars = 50 μm. A-A’’’: Punt expression (A’) is decreased in the whole dorsal compartment when inx3 expression (green, A) is reduced. A pixel intensity plot is shown in A’’’ in which colors follow a thermal scale: blue is the lowest intensity, red the highest. B-B’’’: At the furrow, both stripes of pMad staining (red, B’) are decreased upon inx3 downregulation (dorsal area in B). B’’ shows DE-Cadherin staining and an area encompassing the dorso-ventral boundary, which is magnified below (C-C’’’) in order to better appreciate the staining difference. B’’’ and C’’’: Pixel intensity plots of the pMad staining showing the decreased pMad activation in the dorsal eye disc area at the furrow. D: Percentage of adult flies possessing no, 1 or 2 eyes (n = number of flies examined). Downregulation of inx3 increases number of flies lacking eyes. E: Percentage of eye size in comparison to controls. Left eye size of single flies was measured. Each dot in the graph represents the percentage of a measured single fly eye size in comparison to the measured mean control eye size value (eyaGal4, 100%). Expected eye size for genetic interaction was obtained by multiplying the means of observed eye values and is depicted as a black horizontal bar. inx3 depletion aggravates the small eye phenotype observed after Dpp pathway downregulation. Description of the genotypes for D and E: eyaGal4 (w;eyaGal4/+), eyaGal4 UAS RNAi inx3 (w;eyaGal4/UAS-RNAi inx3-BA14), eyaGal4 UAS RNAi medea (w;eyaGal4/+;UAS-RNAi medea/+), eyaGal4 UAS RNAi medea UAS RNAi inx3 (w;eyaGal4/UAS RNAi inx3-BA14; UASRNAi medea/+).

Figure 6: Inx1 and Inx3 expression depend on inx2. A-A’’’ until C-C’’’: Optical sections in 3rd instar larval eye discs containing inx2 null clones. Anterior to the furrow in DP (A-A’’’) or in PE cells (B-B’’’), Inx3 expression (green, A’, B’) is

diminished and delocalized from the plasma membrane (arrowheads in A’’’, B’’’) in inx2 clones (lacking RFP in red, A, A’’’, B, B’’’). Similarly, Inx1 expression (green, C’) in the progenitor cells (arrowheads in C’’’, future pigment cells) is diminished in the inx2 clones (lacking RFP, red in C, C’’’). D-D’’’ until G-G’’: Optical sections of pupal eyes containing inx2 null MARCM clones marked with GFP (green in D, E, F, G). D-D’’’: Late pupal eye (at the level of the cone cell, close to the cornea) showing that Inx2 (magenta, D’) is not expressed in the MARCM clone. Inx3 expression is diminished in the clone, but only in the pigment cells (D’’). The remaining Inx3 staining is localized in the rhabdomeres. E-E’’ and F-F’’: Early pupal eyes (45% p.d.) showing that Inx3 (E’) and Inx1 (F´) expression in the pigment cells (p) is diminished in inx2 clones expressing GFP (E, F). G-G’’: Section in a late pupal eye at the level of the cone cells (cc) showing reduced Inx1 (G’) expression in the absence of inx2 (GFP, green. G). pp, primary pigment cell, p, secondary pigment cell. All scale bars = 10 m.

Figure 7: Inx2 expression depends on inx3 but neither Inx2 nor Inx3 expression depend on inx1. Optical sections through 3rd instar eye discs of DEGal4 UAS RNAi inx1-7136 or DEGal4 UAS RNAi inx3-BA14 flies at the level of the DP. Posterior is to the left, ventral to the bottom. In all discs, the knock-down area is dorsal (to the top) and control area is ventral (to the bottom). The two panels at the top depict the magnified anterior region of the discs shown in A-A’’’ to D-D’’’, with the morphogenetic furrow on the left (mf). DE-Cadherin staining (blue, A’’ to D’’) is used to visualise details of the disc. The last column contains a merge image of the first two single-channel images on the left. Scale bars = 10 μm. Upon inx1 (red, A, B) depletion in the dorsal area (A, B, arrowhead), Inx2 (green, A’) and Inx3 (green, B’) expression levels and localization remain nearly unmodified (A’’’, B’’’). In contrast, inx3 (red, C, D) depletion in the dorsal part of the disc (arrowhead in C, D) leads to a strong delocalization of Inx2 from the membrane and decreased Inx2 levels (arrowhead in C’), while Inx1 levels are not changed (D’).

Figure 8: Rescue of the inx2 small-eye phenotype with different UAS-Innexin transgenes. Pictures of eyes from flies depleted for inx2 with eyaGal4 and expressing different combinations of UAS inx1, UAS inx2 and UAS inx3. The inx2-depleted eyes are presented twice to better appreciate

size differences upon rescue. Scale bar for all eye pictures = 250 m. The bottom graph represents the mean adult eye size of eyaGal4/+ flies (first column, control) or flies expressing the UAS inx2 RNAi construct under the control of eyaGal4 (second column). Rescue with UAS inx1, UAS inx2, UAS inx3 and combinations thereof were performed in the eyaGal4 UAS RNAi inx2 background. Values represent mean ± s.d. and the control (first column) was considered as 100%. n values from the left to the right column: 10, 11, 12, 11, 14, 10, 13, 7, 11. Statistical results for mean comparison of rescued flies with eyaGal4 UAS RNAi inx2 flies are indicated above the columns. **: very statistically significant in Unpaired t test (0.001
Figure 9: Functions of inx2 and inx3 in the peripodial epithelium. A: Cell layer-specific downregulation of inx2 or inx3. Adult eye size measurements of control eyes (optix Gal4/+, c311Gal4/+) or respective eye knock-downs for inx2 or inx3 (BA14 line) in the disc proper (optixGal4) or in the peripodial epithelium (c311Gal). Values in the columns: mean ± s.d. n=number of eyes. ***: extremely statistically significant in Unpaired t test (P<0,001). B-B’’’ and C: inx2 controls proliferation cell autonomously in the PE. B-B’’’: Optical section through a L3 eye disc containing inx2- clones at the level of the peripodial epithelium. The disc was stained with DE-Cadherin to visualize cell membranes and Hth to visualize nuclei. RFP was used to visualise the clones. An inx2- clone is marked by the absence of RFP (yellow line in B, B’ and B’’’). Twin clones can be recognized by the strong RFP staining (red lines in B, B’ and B’’’). The twin clone in the middle has no partner inx2- clone. Scale bar = 10μm. C: Quantification of the size of an inx2- clone as a percentage of the twin clone size. Each dot represents a clone. The surface of inx2clones is reduced in comparison to the size of their corresponding twin clone in the PE. D: inx2 depletion in the PE decreases proliferation in the DP. Proliferation counts anterior to the morphogenetic furrow in the DP of third instar control (c311Gal4/+) or peripodial inx2-depleted (c311Gal4/UAS RNAi inx2) eye discs. Columns represent mean± s.e.m. of pH3 histone counts in the anterior DP area of the discs (n= number of discs). *: statistically significant in Unpaired t test (P<0.05). E, F: Ventral head structures such as the vibrissae (arrowhead) are sometimes missing upon inx2 depletion in the eye (genotypes: in F knock-down eyGal4/UAS RNAi inx2; in E control ey Gal4/+). Scale bars = 250 μm.

Supplementary figures legend

Supplementary Figure 1 Expression of Inx2 and Inx3 in the peripodial epithelium and in the disc proper. A-A’’’: Optical cross section through a fold in the dorsal part of a 3rd instar OregonR eye disc to visualize the peripodial epithelium (PE) in close contact to the disc proper (DP) (DE-Cadherin, blue in A’’). Inx2 (green, A) and Inx3 (magenta, A’) colocalise (merge of A and A’ in A’’’) and are enriched in the lateral membranes of PE cells and apico-laterally in DP cells. Scale bar = 10 m. B-B’’’: Optical cross section through a 3rd instar OregonR eye disc to visualize the peripodial epithelium (PE) in close contact to the disc proper (DP) (DE-Cadherin, blue in B’’). In the PE, Inx2 (green, B) and Inx3 (magenta, B’) localize predominantly at the contact points between PE cells (arrows in the merge picture in B’’’) but are also detected cortically around the cell (arrowhead in B’, the cortical staining for Inx2 is more dotty as that of Inx3). B’’’ is a merge of B and B’. Scale bar = 10 m. C: Schematic representation of the Inx2 and Inx3 stainings observed in the 3rd instar eye disc. The flattened peripodial epithelium cells (PE) are located above the disc proper cells (DP). A lumen is separating the layers in some areas of the disc. The cell apical sides of both layers face each other in the lumen. In the PE, Inx2 and Inx3 are enriched at the contact points between the cells (big red dots). However, smaller amounts of proteins are also observed cortically around the cells (on the basal and apical sides, here represented as small red dots). In the DP, Inx2 and Inx3 are enriched at the apico-lateral plasma membrane (big red dots) but, as for the PE, can be found in smaller amounts all over the cells (small red dots). The guinea pig anti-Inx1 antibody unspecifically stains nuclei in L3 eye discs. Optical sections through control (D-D’’’) or inx1KO (E-E’’’) L3 eye discs stained with the guinea pig anti-Inx1 antibody (see Mat. and Methods) (red in D, E) and for Inx2 (D’, E’ in green). DECadherin staining was used to visualize cells and morphogenetic furrow (blue in D’’, E’’). The guinea pig anti-Inx1 antibody stains nuclei in the inx1 null mutant (E). The staining at the plasma membrane of eye discs cells anterior to the furrow as well as the staining in the progenitors surrounding the PRCs clusters posterior to the furrow are specific, since they were not observed in the null mutant. Scale bar = 10 m.

Supplementary Figure 2

A-A’’, B-B’’: Expression of Inx6 and Inx7 in the eye disc. Optical sections of OregonR 3rd instar eye discs stained for Inx6 (red, A) or Inx7 (red, B) and DE-Cadherin (A’, B’) to visualize the disc. Posterior is to the left, ventral to the bottom. A’’ and B’’ are merge images. Scale bars = 50 m. C, D, E: eyaGal4, c311Gal4 and optixGal4 expression domains in L3 eye discs. Each image is a representative XY optical section of a series of sections obtained through a L3 eye disc (with the PE at the top and the DP at the bottom, except for panel E where the PE is at the bottom and DP at the top) of the following genotypes: eyaGal4 UAS CD8-GFP (C), c311Gal4 UAS CD8-GFP (D), optixGal4 UAS CD8-GFP (E). Scale bars = 50m. The small panel located above each XY-image is a XZ-section through the disc corresponding to the green line on the XY-section, the small panel located to the right of each image is a YZ-section of the disc corresponding to the red line on the XY-section. In addition, blue lines in the XZ and YZ panels indicate the location of the cut through the Z-stack (XY section). eyaGal4 drives expression in both DP and PE (shown with the arrowheads in the XY section in C). c311Gal4 drives expression of GFP only in the PE (the XY section is at the level of the PE in D). optixGal4 does not drive in the PE (arrowhead in E) but only in the DP.

Supplementary Figure 3 A: Efficiency of the RNAi lines used in this study. Efficiency of each RNAi line was measured in situ in the L3 eye disc by measuring protein expression levels of the corresponding Innexin. For this, each RNAi line was crossed with the DEGal4 driver and intensity of the Inx staining was measured with ImageJ upon depletion. We used a defined area (of a size of approximately one fourth of the anterior part of the disc) to measure pixel intensity in the anterior part of the disc, either on the ventral side (control) or on the dorsal side (knock-down area), paying attention to measure intensity only in areas of similar disc flattening. Values represented in the graph correspond to pixel intensity of the depleted area divided by the pixel intensity of the control area (as a percentage of control). Five discs were analysed for each condition and columns represent mean±sd. B: Inx3 overexpression increases eye size. Mean adult eye size of eyaGal4/+ flies (control, first column) or flies expressing a full-length Inx3 transgene under the control of eyaGal4 (second

column). The dorso-ventral length on its longer segment (from the top to the bottom of the eye) was measured. Values represent mean ± s.d. and the control was considered as 100%. n values are depicted on the columns. ***: extremely statistically significant in Unpaired t test (P<0,001). Eye pictures representative of each genotype are presented under the graph. Scale bar = 250 m. C: Rescue of the inx3-depleted eyes with Inx transgenes. Mean adult eye size of eyaGal4/+ flies (first column, control) or flies expressing the UAS inx3 RNAi BA14 construct under the control of eyaGal4 (second column). Rescue with UAS inx3 or UAS inx2, or a combination of both transgenes was performed in the eyaGal4 UAS RNAi inx3 background (respectively third, fourth and fifth column). Values represent mean ± s.d. and the control (first column) was considered as 100%. n values from the left to the right column: 10, 11, 10, 11, 13. Statistical results for mean comparison of rescued flies with eyaGal4 UAS RNAi inx3 flies are indicated above the columns. ***: extremely statistically significant in Unpaired t test (P<0.001). D: Synergistic effect on eye size upon depletion of inx2 and inx3. Mean adult eye size of eyaGal4/+ flies (first column, control) or flies expressing the UAS RNAi inx3-BA14 construct (second column), the UAS RNAi inx2 construct (third column) or both (fourth column) under the control of eyaGal4. Values represent mean ± s.d. and the control (first column) was considered as 100%. n values are indicated on the columns. Mean comparison of the double knock-down flies (last column) with either eyaGal4 UAS RNAi inx3 or eyaGal4 UAS RNAi inx2 flies are extremely statistically significant in an Unpaired t test (P<0.001). Representative pictures of each genotype is presented on the right (scale bar = 250 μm).

Supplementary Figure 4 Last column pictures are merge images of the first three images. A-A’’’: Inx2 levels are decreased upon inx3 depletion in the peripodial epithelium. Optical section through the PE of a 3rd instar eye disc with a depletion for inx3 in the dorsal part of the disc (DEGal4 UAS RNAi inx3-BA14). Posterior is to the left, ventral to the bottom. The knock-down area is dorsal (to the top) and control area is ventral (to the bottom). When Inx3 (red, A) is depleted, Inx2 levels (green, A’, arrow) are also diminished in the PE. DE-Cadherin staining (blue, A’’) is used to visualize details of the disc. A’’’ is a merge image of the single-channel images on the left. Scale bar = 50 μm. B-B’’’ until E-E’’’: Changes in Inx3 protein levels in DP vs. PE upon inx2 knock-down. Optical sections through control L3 eye discs (B-B’’’ and D-D’’’) or L3 eye discs depleted of inx2 in the DP (with optixGal4, imaged in the DP in B-B’’’ and C-C’’’) or depleted of inx2 in the PE (with

c311Gal4, imaged in the PE in D-D’’’ and E-E’’’). Posterior is to the left, ventral to the bottom. The furrow (mf) is shown in panels B’’ and C’’. Scale bars = 10 m. F-F’’’: Ci expression is decreased in the PE upon inx2 depletion. Optical cross-section through a DEGal4 UAS RNAi inx2 L3 eye disc at the level of the PE. Ci (green, F’, arrow) levels are decreased upon inx2 knock-down (red, F) in the dorsal part of the disc. Scale bar = 50 m.

Supplementary Figure 5 A: Quantitative real-time PCR analysis of inx2 and inx3 transcript levels in inx2 mutant eye discs (EGUF inx2) and their controls (EGUF control) as well as in eye discs depleted for inx3 (eyaGal4 UAS RNAi inx3-BA14) and their controls (eyaGal4-/+). The remaining inx2 transcript expression in EGUF inx2 eye discs probably comes from cells that did not die upon GMR-hid expression (see Mat. and Methods). Values in the columns indicate mean ± s.d. B: Overexpression of Inx2-Myc recruits Inx3 to specific membrane domains. Optical cross sections through the dorsal overexpressing area of a DEGal4 UAS Inx2 Myc L3 eye disc at the level of the DP (lower panels) or at the level of the PE (upper panels) and stained for Myc (magenta), Inx3 (green) and DE-Cadherin (blue). A merge image of Myc and Inx3 is presented on the right side. Inx2-Myc accumulates at specific subdomains in the plasma membrane of eye disc cells, both in the anterior DP and PE, and recruits Inx3 to these domains. Scale bars= 10 m. C: UAS-Inx1-Myc expression is detected in the eye disc. Optical cross section through a L3 eye disc of genotype DEGal4 UAS inx1 Myc. The construct is expressed in the dorsal part of the eye disc (Myc in red; DE Cadherin in blue). Scale bar = 50m.

Richard, M., Bauer, R., Tavosanis, G. and Hoch, M. The gap junction protein Innexin3 is required for eye disc growth in Drosophila HIGHLIGHTS  The gap junction protein Innexin3 controls eye size by modulating the Dpp signaling pathway.  Innexin3 cooperates functionally with Innexin2 to allow for eye disc growth.  Growth of the eye disc requires expression of both Innexins in the peripodial epithelium.