Relating proton pumps with gap junctions: colocalization of ductin, the channel-forming subunit c of V-ATPase, with subunit a and with innexins 2 and 3 during Drosophila oogenesis
© The Author(s). 2016
Received: 6 February 2016
Accepted: 29 June 2016
Published: 13 July 2016
Ion-transport mechanisms and gap junctions are known to cooperate in creating bioelectric phenomena, like pH gradients, voltage gradients and ion fluxes within single cells, tissues, organs, and whole organisms. Such phenomena have been shown to play regulatory roles in a variety of developmental and regenerative processes. Using Drosophila oogenesis as a model system, we aim at characterizing in detail the mechanisms underlying bioelectric phenomena in order to reveal their regulatory functions. We, therefore, investigated the stage-specific distribution patterns of V-ATPase components in relation to gap-junction proteins.
We analysed the localization of the V-ATPase components ductin (subunit c) and subunit a, and the gap-junction components innexins 2 and 3, especially in polar cells, border cells, stalk cells and centripetally migrating cells. These types of follicle cells had previously been shown to exhibit characteristic patterns of membrane channels as well as membrane potential and intracellular pH. Stage-specifically, ductin and subunit a were found either colocalized or separately enriched in different regions of soma and germ-line cells. While ductin was often more prominent in plasma membranes, subunit a was more prominent in cytoplasmic and nuclear vesicles. Particularly, ductin was enriched in polar cells, stalk cells, and nurse-cell membranes, whereas subunit a was enriched in the cytoplasm of border cells, columnar follicle cells and germ-line cells. Comparably, ductin and both innexins 2 and 3 were either colocalized or separately enriched in different cellular regions. While ductin often showed a continuous membrane distribution, the distribution of both innexins was mostly punctate. Particularly, ductin was enriched in polar cells and stalk cells, whereas innexin 2 was enriched in the oolemma, and innexin 3 in centripetally migrating follicle cells. In lateral follicle-cell membranes, the three proteins were found colocalized as well as separately concentrated in presumed gap-junction plaques.
Our results support the notion of a large variety of gap junctions existing in the Drosophila ovary. Moreover, since ductin is the channel-forming part of a proton pump and, like the innexins, is able to form junctional as well as non-junctional membrane channels, a plethora of cellular functions could be realized by using these proteins. The distribution and activity patterns of such membrane channels are expected to contribute to developmentally important bioelectric signals.
The concerted action of ion-transport mechanisms and gap junctions gives rise to bioelectric phenomena, like pH gradients, voltage gradients and ion fluxes within cells, tissues, organs, and even organisms. Such bioelectric phenomena are known to play regulatory roles in various developmental and regenerative processes, e.g. proliferation, differentiation, polarisation and migration [1–5].
Using a variety of approaches, we have gathered considerable information on the electrical properties as well as on the distribution and activity patterns of ion-transport mechanisms and gap junctions in the ovary of Drosophila melanogaster [6–19]. Following this strategy in a model system, we are trying to further clarify the roles that bioelectric phenomena play during development, e.g. for pH-regulation, osmoregulation, cell-cell communication, cell migration, cell proliferation, cell death, vitellogenesis and growth.
During the course of Drosophila oogenesis, characteristic extracellular current patterns [6, 20] as well as membrane-potential changes in germ-line and soma cells have been observed that partly depend on the exchange of protons, potassium ions and sodium ions [7, 21, 22]. Moreover, striking stage-specific patterns of membrane potentials and intracellular pH that characterize distinct cell populations have been described recently . These bioelectric patterns were found to be related to the distribution patterns of various membrane channels, namely V-ATPases, L-type Ca2+-channels, amiloride-sensitive Na+-channels and Na+,H+-exchangers, as well as innexin-3-containing gap junctions. Therefore, these membrane proteins are likely to be involved in the regulation of membrane potentials and/or intracellular pH .
An ovarian follicle of Drosophila consists of 16 germ-line cells surrounded by a layer of somatic follicle cells [23, 24]. The oocyte and its 15 nurse cells form a cytoplasmic continuum via intercellular bridges as well as via gap junctions, and the same holds true for the follicle cells. On the other hand, the germ-line cells are connected to the soma cells only via gap junctions [25, 26]. By way of the intercellular exchange of microinjected fluorescent tracers, stage-specific communication between germ-line cells and soma cells has been detected, and a variety of treatments was found to either inhibit or stimulate this communication .
Innexins are considered the main gap-junction proteins of invertebrates [27–30], but there is some evidence that additional proteins, like e.g. ductin, can be part of gap junctions [11, 15, 31]. Therefore, it is tempting to analyse whether or not ductin and members of the innexin family are found in common gap-junction plaques. Previously, the localization of ductin [11, 16] as well as the localization of innexins 1 to 4  have been separately analysed in Drosophila ovarian follicles in detail.
In the present study, we try to clarify the relationship of ductin, known as a channel-forming subunit of V-ATPases, with innexins 2 and 3, two well known components of gap junctions, using double-immunolabeling. We apply combinations of four antisera directed against (1) ductin (proteolipid or subunit c), (2) another V-ATPase component in the membrane-spanning V0 domain (subunit a), (3) innexin 2, and (4) innexin 3, respectively. These antisera have previously been shown to specifically recognize cellular antigens in the Drosophila ovary [18, 19].
Preparation of follicles
Drosophila melanogaster wild-type Oregon R flies were reared at about 20 °C on standard food with additional fresh yeast. Individual 2–3 days old females were killed by crushing the thorax with tweezers without previous anesthetization. The ovaries were dissected with tweezers in Drosophila PBS , and single follicles of different stages were isolated by pulling at the anterior tip of an ovariole.
To localize V-ATPase and gap-junction proteins, we used (1) a rabbit antiserum (Anti-ductin ) raised and affinity-purified against a highly conserved region (amino acids 115 to 126 in putative loop 3) of the 16 kDa-protein ductin, which forms subunit c of V-ATPases and is also supposed to be a part of gap junctions (AB5496, Chemicon International, USA), (2) a guinea-pig antiserum (Anti-V-ATPase a ) raised against an N-terminal region of subunit a of the Manduca sexta V-ATPase, kindly provided by B. Walz and O. Baumann (Potsdam, Germany), (3) a guinea-pig antiserum (Anti-Inx2 ) raised against the cytoplasmic loop of innexin 2, and (4) a guinea-pig antiserum (Anti-Inx3 ) raised against the C-terminus of innexin 3 from Drosophila, kindly provided by R. Bauer and M. Hoch (Bonn, Germany). A rabbit non-immune serum (NIS [11, 15–18]) was used as control.
Immunoblots were performed as described previously . In short, homogenates of ovaries were sonicated and briefly boiled. The proteins were separated using 12 % SDS-PAGE and transferred to nitrocellulose membranes. Nonspecific binding sites were blocked with 5 % skimmed milk powder/PBS and the blots were incubated in Anti-ductin diluted 1:100 up to 1:500 with 1 % BSA/PBS. NIS diluted 1:200 was used as a control . Subsequently, biotinylated goat-anti-rabbit IgG (Jackson, PA, USA; diluted 1:1000), streptavidin-peroxidase (Dianova, Germany; diluted 1:1000) and H2O2/4-chloro-1-naphthol (Sigma, Germany) were applied, and photographs were taken using a digital camera. Experiments were performed three times.
Indirect immunofluorescence preparations
Follicles were fixed for 30 min at 4 °C in 4 % formaldehyde dissolved in PBS, washed in PBS and blocked for 1 h at 20 °C with 2 % BSA/0,1 % Triton X-100 in PBS. Thereafter, the follicles were incubated overnight at 4 °C in PBS containing 0,5 % BSA/0,1 % Triton X-100 and two of the following antisera for double-labeling: Anti-ductin diluted 1:100, Anti-V-ATPase a diluted 1:1000, Anti-Inx2 diluted 1:20, Anti-Inx3 diluted 1:20. Controls were performed without antiserum and using NIS 1:200 [11, 18], respectively.
After washing six times for 10 min, the follicles were treated with goat-anti-rabbit-Cy3 (Jackson, USA; diluted 1:2000) and donkey-anti-guinea-pig-FP488 (FluoProbes/Interchim, France; diluted 1:100) for 1 h in PBS containing 0,5 % BSA/0,1 % Triton X-100. Washing was repeated six times and the nuclei were stained with 0,2 μg/ml DAPI (Sigma, Germany) in PBS for 3 min. Thereafter, the follicles were mounted in Fluoromount G (Interchim) and viewed, by using x20/0.5, x40/0.75 or x63/1.25oil objectives and the appropriate filter sets, either on a Zeiss Axiovert 200 wide-field fluorescence microscope (WFM), equipped with a Hamamatsu Orca ER camera, or they were mounted in glycerine/PBS 1:1 and viewed on a Leica DMRE laser-scanning microscope (LSM). Each experiment was repeated at least three times.
Immunoblot analysis of Anti-ductin
Localization of ductin in indirect immunofluorescence preparations
Using Anti-ductin and laser-scanning microscopy (LSM), we investigated the distribution of ductin during the course of oogenesis (Fig. 1b-g). As shown previously using the two other anti-ductin sera on sections prepared for immuno-fluorescence and immuno-electron microscopy [11, 16], the continuous as well as punctate distribution of ductin was found to change stage-specifically. In the following, we concentrate on ductin in specialized types of follicle cells (polar cells, border cells, stalk cells and centripetally migrating cells) and on some main features of its distribution relevant to the present comparative analysis.
During previtellogenesis (up to stage 7), ductin is located in cytoplasmic particles or vesicles as well as in plasma membranes and is enriched in polar cells (Fig. 1b). Beginning with vitellogenesis, ductin increasingly becomes located in plasma membranes of nurse cells and follicle cells (Fig. 1c). During vitellogenesis (from stage 8 onward), ductin is located (1) in cytoplasmic vesicles, (2) in plasma membranes of centripetally migrating follicle cells (Fig. 1d), (3) in lateral and apical plasma membranes of columnar follicle cells, (4) in plasma membranes of squamous follicle cells and nurse cells as well as around nurse-cell nuclei, (5) in the oolemma, and (6), most prominently, in polar cells and stalk cells (Fig. 1e and f). Predominantly in lateral and apical follicle-cell membranes, ductin is localized in a punctate pattern resembling gap-junction plaques (Fig. 1g; (cf. )). No specific staining was found in control preparations without primary antiserum or using NIS (cf. [11, 18]).
Colocalization of ductin with V-ATPase subunit a
During early previtellogenesis, ductin and subunit a co-occur (1) in polar cells, (2) in lateral and apical follicle-cell membranes and (3) in the cytoplasm of follicle cells, nurse cells and the oocyte. However, while ductin was found to be more prominent in stalk cells, the distinct particles or vesicles in germ-line cells were found to contain predominantly subunit a (Fig. 2a). During late previtellogenesis, both ductin and subunit a are enriched in apical follicle-cell membranes. On the other hand, ductin was found to be more prominent in posterior polar cells and in stalk cells, while subunit a was more prominent in the cytoplasm of follicle cells, nurse cells and the oocyte (Fig. 2b).
During vitellogenesis, ductin and subunit a co-occur especially in columnar and squamous follicle cells and in border cells. In columnar follicle cells, the amount is usually higher on one side of the follicle (Fig. 2c), namely the ventral side (cf. ). Here, ductin and subunit a are colocalized in the plasma membranes and in the cytoplasm while, in squamous follicle cells, they are are colocalized in vesicles (Fig. 2d). On the other hand, ductin is enriched in nurse-cell membranes, while subunit a is enriched in border cells (Fig. 2c) as well as in nuclear particles or vesicles in follicle cells (Fig. 2e). During late vitellogenesis, ductin and subunit a are colocalized in cytoplasmic vesicles in most of the follicle cells, whereas, in the cytoplasm and nuclei of anterodorsal follicle cells, predominantly subunit a was found (Fig. 2f). Taken together, ductin and subunit a are stage-specifically either colocalized or separately enriched in different cellular regions of germ-line and soma cells. Frequently, ductin is more prominent in plasma membranes, while subunit a is more prominent in cytoplasmic and nuclear particles or vesicles (for summary, see Fig. 5a and a’).
Colocalization of ductin with innexins
Since ductin is known to form intercellular communication channels [11, 15, 31], it might localize, in part, together with members of the channel-forming innexin family in the same cellular regions [27–30]. In Drosophila ovarian follicles, the localization of innexins 1 to 4 has been described in detail previously using LSM and WFM . For the present comparative analysis, we concentrate on some main features concerning Inx2 and Inx3, that both showed distribution patterns similar to ductin.
Ductin and Inx2
Also during vitellogenesis (Fig. 3c), ductin and Inx2 co-occur in lateral and apical follicle-cell membranes. On the other hand, ductin is still enriched in posterior polar cells, while Inx2 is now enriched in the oolemma. In lateral follicle-cell membranes, both proteins are often found concentrated in different plaques that are presumed to represent different gap junctions (arrowheads in Fig. 3c and c’).
Ductin and Inx3
In mid to late vitellogenic stages (Fig. 4b, c), ductin and Inx3 co-occur in lateral follicle-cell membranes and in nurse-cell membranes. While ductin is enriched in apical follicle-cell membranes and in the oolemma, Inx3 forms prominent plaques in lateral follicle-cell membranes. Here, ductin and Inx3 are often found concentrated in different plaques that are presumed to represent different gap junctions (arrowheads in Fig. 4c and c’ and arrows in c”).
The channel-forming protein ductin has been found in both V-ATPases and gap-junctional structures . In V-ATPases, it forms the proteolipid component or subunit c, whereof the C- and N-termini as well as loop 2 are located either on the vacuolar or on the extracellular side of the membrane. In gap-junctional structures, where ductin is supposed to form membrane channels, these regions are located on the cytoplasmic side of the membrane [33, 34]. Using light and electron microscopy, it has been shown that antibodies against ductin bind to antigens located in the cytoplasm and in plasma membranes of various Drosophila tissues, especially of ovarian follicles and embryos [11, 15–17]. In part, cytoplasmic labeling might represent storage or transport of ductin. However, since V-ATPases are required for the acidification of organelles (e.g. ), cytoplasmic labeling is likely to also represent secretory vesicles, lysosomes or endosomes in which ligand-receptor complexes become dissociated (e.g. [16, 36]).
On the other hand, V-ATPases coupled to secondary active antiport mechanisms are known to energize transport processes across plasma membranes (e.g. ). These pumps are expected to exert direct or indirect influence on intracellular pH and membrane potentials. In the follicle-cell epithelium of Drosophila, antibodies against both subunits a and c revealed an asymmetric distribution of V-ATPases which is in accordance with the gradual dorsoventral distribution of epithelial membrane potentials . This observation points to a possible involvement of V-ATPases in the regulation or maintainance of the spatial coordinates during oogenesis, a phenomenon that has been observed in other systems (e.g. [38–40]). Moreover, V-ATPases are also known to be involved in the regulation of cell division, migration and differentiation (e.g. [41, 42]).
Labeling of plasma membranes with antibodies against ductin was either continuous or punctate (cf. [11, 16, 19]). Continuous labeling is presumed to originate from ductin as part of V-ATPases, while punctate labeling might originate from ductin as part of gap-junctional structures. By microinjection experiments, it has been shown that antibodies directed against presumed cytoplasmic regions of ductin specifically block intercellular communication and exert adverse influence on oogenesis as well as on embryogenesis [11, 15].
Interestingly, ductin labeling was prominent in both stalk cells and polar cells that derive from a common cell population in the germarium. Polar cells are known to control stalk formation as well as axis formation, they recruit the border cells and they express various cell-specific markers [43, 44].
Evidence amounts that ductin, or subunit c, is a multi-functional channel protein. It has been observed to form Ca2+-inducible membrane pores that are permeable to hydrophilic molecules . It has also been demonstrated that hexamers of subunit c form stacked rings containing an intercellular channel [46, 47]. And, in addition, ductin seems to play a role in the process of vesicle-membrane fusion . The present study lends further support to the notion that ductin might be engaged in several of these functions also during Drosophila oogenesis.
Although various proteins have been described to be part of Drosophila gap junctions (cf. [15, 27]), members of the innexin family are assumed to be the main invertebrate gap-junction proteins [29, 30, 49]. In the Drosophila ovary, the mRNAs of innexins 1, 2, 3, 4 and (to a minor extent) 7 have been detected . Using various antibodies against innexins (Inx), Inx1 was found to be predominantly located in the baso-lateral domain of follicle cells, whereas Inx2 is positioned apico-laterally as well as apically between follicle cells and germ-line cells. Inx3 was observed laterally in follicle cells and also in nurse cells, and Inx4 was detected in the oolemma up to stage 8 and in nurse-cell membranes up to stage 12. While Inx2 and Inx3 are colocalized between somatic cells, Inx2 and Inx4 are colocalized between somatic cells and germ-line cells .
Microinjected antibodies directed against presumed cytoplasmic regions of Inx2 specifically blocked intercellular communication and lead to inhibitory effects on oogenesis that were similar to those obtained with antibodies directed against presumed cytoplasmic regions of ductin . Even during spermatogenesis, Inx2 was found to be involved in soma-to-germ line intercellular communication . Moreover, it has been demonstrated that, within the germarium, Inx2 supports germ-cell survival, cyst formation and early follicle development .
During embryogenesis, Inx2 was shown to participate in organizing epithelia by interacting with core proteins of adherens and septate junctions, and it has been presumed that Inx2 exerts its structural functions indirectly through the formation of channels for intercellular communication [53–56]. Recently, Inx2 has also been identified as a regulator of eye development .
Contrary to Inx2, Inx3 shows a striking non-uniform distribution in the follicle-cell epithelium : it is enriched in the lateral membranes of centripetally migrating follicle cells. This indicates distinct coupling conditions between these cells concerning e.g. regulatory signals, membrane potentials or intracellular pH. Moreover, Inx3 has been described to be involved in maintaining tissue integrity in response to tension , a feature that is particularly important for migrating cells. The spatial restriction of specialized intercellular channels and the independent regulation of their permeability is suspected to provide mechanisms to generate or maintain boundaries between distinct follicle-cell populations . By the use of various proteins in either homomeric, heteromeric or heterotypic combinations, a wealth of different gap-junction channels might be formed (cf. ).
Compared to neighboring cells, the centripetally migrating follicle cells are specialized in several respect: (1) they are more acidic and more depolarized , (2) they possess higher amounts of active voltage-dependent L-type Ca2+-channels  (becoming activated under depolarized conditions ), (3) they contain higher Ca2+-concentrations , (4) they possess higher amounts of Na+,K+-ATPase [13, 16] (becoming stimulated under alkaline conditions ), and (5) they remain coupled via gap junctions to the oocyte when neighboring cells are already uncoupled .
The permeability of innexin channels in general has been reported to be sensitive to membrane potential, intracellular pH, K+- and Ca2+-concentrations (e.g. [61, 62]). Gap-junctional communication between oocyte and follicle cells was found to be inhibited by low extracellular pH, by low extracellular K+-concentrations (leading to hyperpolarization [21, 22]), and by high Ca2+-concentrations in the oocyte . Taken together, these findings support the notion that intercellular communication in Drosophila ovarian follicles is precisely regulated.
Similar to ductin and to the pannexins in vertebrates, innexins have also been found to form non-junctional channels (innexons) allowing the secretion of small molecules, including ATP, into the extracellular space [47, 62–64]. Innexons were observed to open in response to mechanical stress, depolarization, high intracellular Ca2+- and high extracellular K+-concentrations, and to close due to low intracellular pH [62, 63].
In recent years, the origin as well as the physiological and cellular functions of bioelectric phenomena have attracted growing attention and, thus, their meaning becomes more and more elucidated (e.g. [4, 65–68]). Accordingly, the distribution and activity patterns of membrane channels, like ductin and innexins, during the course of Drosophila oogenesis are supposed to contribute to developmentally important bioelectric signals .
Since ductin shows overlapping distribution patterns with both Inx2 and Inx3 (for summary, see Fig. 5), these proteins might be, in part, located within the same gap-junctional structures, especially between neighboring follicle cells. However, the analysed proteins are often found concentrated in different membrane plaques which supports the notion of different gap junctions with a large variety of protein compositions.
Particularly, ductin is enriched in polar cells and stalk cells, while Inx2 is enriched in the oolemma, and Inx3 is enriched in centripetally migrating follicle cells, but the other proteins are not absent from these regions. Since ductin is part of a proton pump and, like the innexins, can form junctional as well as non-junctional membrane channels, a plethora of cellular functions could be realized by using only the three proteins analysed in the present study.
a, anterior; ad, anterodorsal; BC, border cells; BSA, bovine serum albumine; c, centripetally migrating; DAPI, 4’,6-diamidino-2-phenylindole; DMSO, dimethyl sulfoxide; FC, follicle cells; Inx2, innexin 2; Inx3, innexin 3; LSM, laser-scanning microscopy; M, membrane; N, nucleus; NC, nurse cells; NIS, non-immune serum; Ooc, oocyte; Ool, oolemma; p, posterior; PBS, phosphate buffered saline; PC, polar cells; SC, stalk cells; SDS-PAGE, sodium dodecyl sulphate polyacrylamide gel-electrophoresis; WFM, wide-field microscopy
We are indebted to Bernd Walz and Otto Baumann (Potsdam, Germany) for providing the V-ATPase antiserum, to Reinhard Bauer and Michael Hoch (Bonn, Germany) for providing the innexin antisera, and to Isabel Weiß for designing Fig. 5. Financial support by RWTH Aachen University is acknowledged.
Availability of data and materials
The dataset supporting the conclusions of this article is included within the article.
JL carried out the experiments under the supervision of JB. JB conceived the study, reviewed the data and wrote the manuscript. Both authors read and approved the final version.
The authors declare that they have no competing interests.
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