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Vol. 15, Issue 10, 4512-4521, October 2004
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Cell Biology and Metabolism Branch, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD 20892
Submitted June 11, 2004;
Revised July 22, 2004;
Accepted July 23, 2004
Monitoring Editor: Allan Spradling
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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; Büning, 1994; de Cuevas et al., 1997). In Drosophila a single oocyte develops within the environment of a 16-cell cyst. The development of this cyst occurs in a series of morphologically defined regions (see Figure 1A). Cyst production begins when a germline stem cell divides asymmetrically to produce a cystoblast. The cystoblast undergoes four synchronous mitotic divisions (regions 1 and 2a), with incomplete cytokinesis, to produce a cluster of 16 cells that are connected via arrested mitotic cleavage furrows (reviewed in de Cuevas et al., 1997). Later in oogenesis, the arrested cleavage furrows mature into stable ring canals and are reinforced with an actin cytoskeleton (reviewed in Mahajan-Miklos and Cooley, 1994). Initially, all 16 cystocytes develop in synchrony. However, as development proceeds, differentiation within the cyst becomes asynchronous. Ultimately, only one cell commits to meiosis and differentiates as the oocyte, whereas the other 15 cystocytes enter the endocycle and develop as trophic nurse cells.
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A unique membranous organelle, the fusome, grows and branches along the spindle equators after each mitotic division to physically connect all cells within a cyst. The fusome is found in the germline cysts of Drosophila as well as other insects (reviewed in Telfer, 1975; McKearin, 1997). The Drosophila fusome is comprised of cytoplasmic endomembranes, membrane skeletal proteins and polarized microtubules (reviewed in de Cuevas et al., 1997; McKearin, 1997).
Molecular genetic studies over the last ten years indicate the fusome mediates several essential steps in germline cyst development (reviewed in McKearin, 1997). The differentiation of the oocyte depends on the production of a polarized network of microtubules within the cyst, that facilitates the directional transport of specific mRNAs and proteins from the pronurse cells to the single cell that will become the oocyte (Koch and Spitzer, 1983; Theurkauf et al., 1992, 1993). The fusome organizes the network microtubules within developing ovarian cysts and is required for the microtubule-dependent restriction of oocyte markers (Grieder et al., 2000). In addition, the fusome directs the pattern of cystocyte interconnections by anchoring one pole of each mitotic spindle, thus orienting the plane of cell division (Telfer, 1975; Storto and King, 1989; Lin and Spradling, 1995).
Mutations in genes that encode the cytoskeletal component of the fusome including 
-spectrin and 
-spectrin, and the adducin-like protein hts, result in a reduced number of cyst divisions and a block in oocyte differentiation (Lin et al., 1994; de Cuevas et al., 1996; de Cuevas and Spradling, 1998). In the absence of the fusome, cysts develop with random patterns of interconnections and no signs of polarity. However, it is unclear if the fusome plays an instructive role in the differentiation of the oocyte through participation in specific signaling pathways or if its sole function in oocyte differentiation is to facilitate the organization of microtubules into a polarized network. Thus, the fusome provides a useful model to examine how developmentally directed modifications of an intracellular organelle might function to facilitate differentiation.
Although progress has been made toward revealing the molecular composition of the proteinaceous components of the fusome, the origin and nature of the membranous component remains poorly understood. Based on electron microscopy studies, the ribosome deficient endomembranes of the fusome have been described as vesicles or an interconnected network of tubules that resemble modified endoplasmic reticulum (ER) or Golgi (Koch and King, 1966; Mahowald, 1971; Lin et al., 1994; McKearin and Ohlstein, 1995). Consistent with this observation, León and McKearin (1999) reported the enrichment on Drosophila fusome structures of the cytoplasmic protein TER94, which associates with both the Golgi complex and the ER in vertebrates (Uchiyama et al., 2002). In this study, we have used live imaging and photobleaching of green fluorescent protein (GFP)-tagged proteins to gain insight into the membranous component of the Drosophila fusome.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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) was a gift from Allan Spradling. Dhc643–2 and Dhc646–12 (McGrail and Hays, 1997) were from Margaret de Cuevas. The P[w+ Pvas-gfp] flies were a gift from Akira Nakamura (Nakamura et al., 2001). The Sec61-GFP line (ZCL0488 from the Flytrap collection) was generously provided by Lynn Cooley. Sec61 is a 10-membrane–spanning resident ER protein that associates in a heterotrimer with Sec61 and 
to form the core of the ER protein translocation channel, the translocon (Johnson and van Waes, 1999).
Transgenic lines were generated in a w1118 background using standard techniques (Spradling, 1986). Two independent insertions of P{w+, UASp::Lys-GFP-KDEL} and two independent insertions of P{w+, UASp: galactosyltransferase-GFP} into the second and third chromosomes were obtained. The nanos-Gal4:VP16 driver (Van Doren et al., 1998) was used to express UASp-transgenes in the germline. Transgenic flies of the following genotypes were used in this article: w1118; P{w+, UASp::Lys-GFP-KDEL}; P{w+, nanos-Gal4:VP16}; w1118; P{w+, UASp::galactosyltransferase-GFP}; P{w+, nanos-Gal4:VP16}; w1118; hts1/hts1; P{w+, UASp::Lys-GFP-KDEL}/P{w+, nanos-Gal4: VP16}; w1118; P{w+, UASp::Lys-GFP-KDEL}/+; Dhc643–2/Dhc646–12, P{w+, nanos-Gal4:VP16}.
Dhc646–12, P{w+, nanos-Gal4:VP16}, w1118; P{w+ Pvas-GFP}; hts1/hts1 chromosomes were generated by meiotic recombination. All additional stocks were obtained from the Bloomington Stock Center.
Plasmid Constructions
To visualize the ER, a marker protein, lysozyme-GFP-KDEL (Lys-GFP-KDEL), was expressed in the germline using the Drosophila Gal4-UAS system. Lysozyme-GFP-KDEL was constructed by fusing the complete coding sequence of hen's egg lysozyme with a COOH-terminal enhanced GFP (BD Biosciences, Clontech, Palo Alto, CA) followed by a KDEL ER retention signal (Munro and Pelham, 1987). The Golgi apparatus marker protein, galactosyl-transferase (GalTase-GFP) has been previously described (Cole et al., 1996). Separate DNA fragments encoding Lys-GFP-KDEL or GalTase-GFP were cloned into pUASp (Rorth, 1998) to generate a p[w+, UASp-lysozyme-GFP-KDEL] or a p[w+, UASp-galactosyltransferase-GFP] transgene. Lys-GFP-KDEL and GalTase-GFP expressing flies produced viable adults.
Immunofluorescence and Imaging of Live and Fixed Ovaries
Ovaries were dissected, immunostained, and mounted as described previously (Lin et al., 1994). Antibodies were used at the following concentrations: rabbit anti--spectrin (Byers et al., 1987; Dubreuil et al., 1987) at 1:100, mouse anti--spectrin 3A9 (Dubreuil et al., 1987) at 1:5 (Developmental Studies Hybridoma Bank, University of Iowa), mouse anti-Hts (Zaccai and Lipshitz, 1996) at 1:5, rabbit anti-GFP (Molecular Probes, Eugene, OR) at 1:800 and mouse anti-GFP (Molecular Probes; JL-8) at 1:1000. Secondary antibodies conjugated to Alexa 594 or Alexa 488 (Molecular Probes) were used at 1:800 dilution.
Live ovaries were placed flat on poly-lysine–coated Lab-Tek chambered coverglasses from Nalge Nunc (Naperville, IL). Chambers were then filled with Drosophila Ringer's solution (Tübingen and Düsseldorf). Confocal microscope images of live and fixed ovaries were captured on a LSM 510 confocal microscope (Zeiss, Thornwood, NY) using a 488-nm line of a 40-mW Ar/Kr laser with a 505–530 emission filter for GFP and a 543 nm HeNe laser line with a 560–615 emission filter. Images were captured with a 1.2 NA 63x water objective and a 1.3 NA plan-neo 40x oil objective. Image analysis was performed using NIH Image 1.62 and Zeiss LSM image examiner software. Composite figures were prepared using Photoshop 7.0 and Illustrator 9.0 software (Adobe, San Jose, CA).
Photobleaching and Analysis
Fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP) were performed by photobleaching a small region of interest (ROI) and monitoring fluorescence recovery or loss over time, as described previously (Siggia et al., 2000; Snapp et al., 2003). Fluorescence intensity plots and Deff measurements were calculated as described previously (Siggia et al., 2000; Snapp et al., 2003). To create the fluorescence recovery curves, the fluorescence intensities were transformed into a 0–100% scale and were plotted using Kaleidagraph 3.5 (Synergy Software, Reading, PA). For Figure 3, the postbleach asymptote intensity of the ROI was designated as 100%. For Figure 4, 5, 6, the prebleach intensity of the ROI was designated as 100%. p values were calculated using a Student's two-tailed t test in Excel (Microsoft, Redmond, WA).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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; Uchiyama et al., 2002). To characterize the structure and origin of the membranous component of the fusome, we expressed the lumenal ER-marker protein, Lys-GFP-KDEL, the resident ER membrane protein, Sec61-GFP, or the Golgi marker, GalTase-GFP (Cole et al., 1996), in ovarian cysts. In confocal micrographs of live germaria, Lys-GFP-KDEL and Sec61-GFP both labeled amorphous tubular elements that extend from the nuclear envelope throughout the cytoplasm (Figure 1, B and C), consistent with an ER distribution. Interestingly, the fluorescence of both proteins was more intense in thick branching structures between adjacent cystocytes. To determine whether these accumulations of ER membranes correspond to fusomes, we performed immunofluorescence with antibodies against the fusomal components -Spectrin and Hts. We observed extensive colocalization of Lys-GFP-KDEL and Sec61-GFP with the fusomal markers (Figure 2, A–D). In contrast, GalTase-GFP localized to discrete punctate structures throughout the cytoplasm of all cystocytes. The puncta did not colocalize with the fusome marker Hts (Figure 2E). This pattern is similar to the distribution of the Golgi marker anti--mannosidase 2 previously described by Cox and Spradling (2003). Additional membranes of other origins may be present within the fusome. Nonetheless, our data support the conclusion that fusomes are significantly enriched in ER membranes and that these probably represent the primary source of fusome membranes.
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In stem cells, spectrin accumulates in a structure termed the spectrosome (Lin et al., 1994). It has been proposed that the spectrosome is the precursor of the fusome and may serve as a source of cytoskeletal proteins and membranes (Lin et al., 1994). In Figure 2, A and C, the ER colocalizes with spectrosomes. However, the ER is less enriched than in the fusome structures for either Lys-GFP-KDEL or Sec61-GFP, similar to previous observations (Lin et al., 1994).
Ovarian Cysts Share a Common ER
Whether the fusome membranes consist of discrete ER-derived vesicles or form a network of interconnected tubules has remained unresolved with evidence for both types of fusome membrane structures observed by electron microscopy (Telfer, 1975; Büning, 1994; Lin et al., 1994; McKearin, 1997). It is worth noting that the ER of animal cells is an ideal example of an interconnected membranous compartment and typically exists as a continuous network of branching tubules connected to the nuclear envelope. However, the fusomal membranes may not be continuous with the cytoplasmic ER of the cystocytes.
We exploited photobleaching methods to directly assess the membrane continuity of fusomal membranes (Ellenberg et al., 1997; Lippincott-Schwartz et al., 2001). First, we confirmed the suitability of Lys-GFP-KDEL and Sec61-GFP (unpublished data) as highly mobile diffusing molecular markers by performing Fluorescence Recovery after Photobleaching (FRAP) of Lys-GFP-KDEL expressed in cystocytes from region 1 of the germarium (unpublished data) and stem cells (Figure 3A). For example, a small ROI in a stem cell expressing Lys-GFP-KDEL was photobleached with a high intensity laser beam, destroying the GFP fluorescence. Recovery of fluorescence into the photobleached ROI indicated that unbleached Lys-GFP-KDEL molecules outside of the photobleached ROI had diffused into the photobleached ROI, whereas photobleached molecules had diffused out of the ROI. The fluorescence recovery was rapid and did not involve gross structural changes of the ER membranes, consistent with the diffusion of a highly mobile protein. The diffusion coefficient (Deff) within the cystocytes (Deff = 0.84 ± 0.4 µm2/s, Mobile fraction 
100%, n = 5) represents a value consistent with diffusion of a protein within the crowded lumen of the ER (Dayel et al., 1999). Sec61-GFP also diffuses readily. However, as expected, the membrane protein diffuses more slowly through the more viscous environment of the membrane (Deff = 0.2 ± 0.02 µm2/s, Mobile fraction 100%, n = 2). Thus, both proteins freely diffuse within the ER and therefore represent suitable markers to examine membrane continuity.
To determine whether fusome membranes were vesicular or formed a continuous network, we compared the Deff of Lys-GFP-KDEL and Sec61-GFP within the fusome ER-derived membranes. The organization of the membranes should affect the apparent mobility of the lumenal and membrane proteins. If the fusome were composed of discrete vesicles, then movement of both proteins would depend on the movement of molecular motors or diffusion of the whole vesicles. Both proteins would be predicted to move with identical dynamics. In contrast, if the proteins reside within continuous tubules, then the distinct environments of the ER lumen and the more viscous ER membrane would dominate protein mobility. The two proteins would be predicted to diffuse at different rates.
FRAP of the fusome membranes revealed that Lys-GFP-KDEL fluorescence recovers rapidly within the fusome (Deff = 1.5 ± 0.4) µm2/s, n = 3; Figure 3, B and E), whereas Sec61-GFP recovers significantly more slowly (p = 0.0002; Deff = 0.15 ± 0.03 µm2/s, n = 5). The Deff values for each protein are not statistically significantly different for each protein in cystocytes (p = 0.062 for Lys-GFP-KDEL and 0.094 for Sec61-GFP). In both cases, recovery occurred on both sides of the photobleached ROI, consistent with diffusion, a nondirectional process. Thus, the FRAP results are most consistent with the organization of the fusome as a continuous network of membranes, similar to cytoplasmic ER.
The FRAP results raised an important question. Are the ER-derived fusome membranes continuous with the cytoplasmic ER of cells within the cyst? While FRAP can be used to infer compartment continuity, a second photobleaching method, Fluorescence Loss in Photobleaching (FLIP; Ellenberg et al., 1997), can be used to directly probe membrane continuity of an entire structure by repeatedly photobleaching a discrete ROI (Lippincott-Schwartz et al., 2001). If all of the fluorescent molecules are mobile and can diffuse through the ROI in a short period, all of the fluorescence within a compartment will be depleted. We performed FLIP on cysts with 2, 4, or 8 cells in region 1. Data for a representative 8 cell cyst are shown in Figure 4, A and B. Surprisingly, when we bleached a small ROI of the cytoplasmic ER from an individual cystocyte, within a cyst, we observed a rapid depletion of fluorescence from the entire fusome, as well as the cytoplasmic ER in all cystocytes throughout the cyst (Figure 4A). Similar results were obtained if the photobleach ROI was placed within the fusome (unpublished data). In all FLIP experiments, fluorescence in other cysts within the same germarium was not significantly depleted, confirming that fluorescence loss in cells within the cyst of interest was not caused by photobleaching due to the imaging conditions. These dramatic results reveal that not only are the ER membranes within the fusome continuous, but more remarkably, the ER of all cystocytes are interconnected to form a common ER.
Cyst ER Connectivity Is Developmentally Regulated
Next, we investigated the maintenance of the common ER during oogenesis. The structure of the cytoskeletal component of the fusome changes after the completion of the final mitotic cyst division (region 2a, see Figure 1a). Staining with antibodies against Hts and -Spectrin reveals a decrease in the thickness of the fusome beginning in region 2a cysts. This change in fusome structure correlates with the time that cell cycle synchrony is lost as cystocytes asynchronously progress through premeiotic S phase (Lin et al., 1994). By region 2b, the cytoskeletal component of the fusome begins to deteriorate and by region 3, characteristic fusome structures are no longer visible (Lin et al., 1994; de Cuevas et al., 1996).
We asked whether the ER of cells within an ovarian cyst remain interconnected after the completion of the mitotic cyst divisions, as the nurse cells and oocyte begin to pursue their unique developmental and cell cycle programs. To address this question, we used FLIP to compare the continuity of the ER within cysts before (region 1) and after the final mitotic division (region 2a). Figure 4, B and C, graphically reveal an abrupt change in ER connectivity within cysts between region 1 and region 2a. In contrast to the rapid fluorescence loss within the 8 cystocyte region 1 cyst in Figure 4B, the rate of fluorescence loss in cystocytes within region 2a cyst decreases dramatically (Figure 4C). The fluorescence intensities of the directly photobleached cystocytes (red circles) decrease rapidly and exponentially. However, while the cystocyte closest to the cystocyte being photobleached does lose fluorescence, the loss is at a much slower linear rate. In addition, the fluorescence loss from other cystocytes within the same cyst now resembles the minimal loss of fluorescence of cystocytes in adjacent cysts (black triangles). Similar results were observed for FLIP experiments of regions 2b and 3 cysts (Figure 4, D and E). The abrupt change from a steep exponential drop in fluorescence in adjacent cystocytes, to a shallow linear decrease in fluorescence between cysts in regions 1 and 2a, argues for a substantial loss of cyst ER connectivity after the completion of the mitotic cyst divisions. This result is surprising, in that Lys-GFP-KDEL does not noticeably redistribute and continues to colocalize with the cytoskeletal component of the fusome in region 2a (unpublished data). The change in ER connectivity at the mitotic/meiotic boundary represents a newly defined step in cyst development.
ER Connectivity Correlates with Mitotic Synchrony
The temporal coincidence of the loss of ER connectivity and the onset of cell cycle asynchrony suggest a potential role for the shared ER in region 1 cysts; the ER may be important for synchronizing the mitotic cyst divisions. To explore this possibility, we examined ER connectivity in female sterile mutants of hu-li-tai-shao (hts) and the Dynein subunit, Dhc64c. In hts1 cysts, no fusome structures can be detected by immunofluorescence or by electron microscopy (Lin et al., 1994). Conversely, cysts from Dhc64c3–2/Dhc64c6–12 transheterozygous females retain fusome structures, but these structures frequently are fragmented and do not extend into all cells of the cyst (McGrail and Hays, 1997). Both hts1 and Dhc64c3–2/Dhc64c6–12 dramatically reduce the number of cyst divisions leading to the production of egg chambers that contain fewer than 16 cells. However, the mutants have very different effects on mitotic synchrony. hts1 cysts undergo asynchronous divisions as evidenced by the presence of egg chambers that contain cyst cell numbers that do not conform to multiples of 2n (i.e., 2, 4, 8, 16; Yue and Spradling, 1992). In contrast, 95% of Dhc64c3–2/Dhc64c6–12 egg chambers contain a 2n number of germline cells, indicating mitotic synchrony is maintained (McGrail and Hays, 1997). We reexamined both the hts1 and Dhc64c3–2/Dhc64c6–12 mutant phenotypes and determined that mitotic divisions involving four or more cells (4–8 and 8–16) require a synchronizing regulatory mechanism that is disrupted in hts1 but not in Dhc64c3–2/Dhc64c6–12 mutants (unpublished data).
Given the differences in the ability of hts and Dhc64c mutants to support mitotic synchrony during cyst divisions, we then asked whether the two mutants differ in ER connectivity. We probed the degree of ER continuity between cystocytes in the two mutants by FLIP (Figure 5). Repetitive photobleaching of individual cystocytes of hts1 mutants failed to deplete fluorescence in adjacent cystocytes in the same cyst (Figure 5, A and B). In contrast, FLIP of Dhc64c3–2/Dhc64c6–12 mutant cystocytes rapidly depleted fluorescence of all cystocytes within the same cyst (Figure 5, C and D). Thus, the mitotically synchronous Dhc64c3–2/Dhc64c6–12 cysts maintain an interconnected ER, whereas hts1 cysts do not. These data are consistent with EM studies that indicate that the membrane component of the fusome is absent in hts1 mutant germaria (Lin et al., 1994). As a control, we determined that cytoplasmic connectivity is unaffected in hts1 mutants (Figure 6). Specifically, we found that a cytoplasmic GFP marker can freely diffuse between the cytoplasms of interconnected cystocytes in both wild-type and mutant cysts (Figure 6). These data confirm that cytoplasmic connectivity is insufficient to coordinate mitotic synchrony between cystocytes and support the model that the fusome is required to promote cell cycle synchrony. Our results raise the intriguing possibility that the membranous ER component of the fusome coordinates cell cycle synchrony during the mitotic cyst divisions.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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How might a common ER contribute to cyst development during early oogenesis? One potential functional role for the fusomal membranes was first suggested by the observation that mutations in bam and bgcn, two genes that control the developmental switch from stem cell to cystoblast, alter the membranous composition of the spectrosome (McKearin and Ohlstein, 1995), the likely precursor of the fusome (Lin et al., 1994). Another possibility is that the common ER may promote mitotic synchrony in cystocytes that are not directly connected. Mitotic synchrony is essential for the production of the invariant pattern of interconnections found in Drosophila ovarian cysts. Mitotic effectors such as cyclin A (CycA) have been shown to transiently associate with the fusome during G2 and prophase (Lilly et al., 2000). At least one signal must exist upstream of CycA to coordinate the activity of CycA and other mitotic effectors. Modulation of calcium levels is an attractive potential signaling candidate. Calcium signaling plays several roles in mitotic pathways (Berridge, 1995) and the ER is the primary site of calcium storage and release in eukaryotic cells (Baumann and Walz, 2001). Another possible mechanism by which the fusome ER may regulate the cell cycle is suggested by the findings that the SCF component Cul-1, as well as polyubiquitinated cyclin E, localize to the fusome (Ohlmeyer and Schupbach, 2003). Cul-1 has been implicated in the regulated destruction of a wide array of proteins involved in both cell cycle regulation and signal transduction (reviewed in Deshaies, 1999). Interestingly, the membrane-associated protein TER94/VSC/p97, which localizes to the fusome (Leon and McKearin, 1999), has also been implicated in ubiquitin-dependent degradation in mammals (Dai and Li, 2001). Thus, the ER component of the fusome may provide a surface for regulated protein degradation. An additional possibility is that the common ER may serve as a transport system that regulates trafficking of signal containing transmembrane and membrane-associated proteins between cystocytes in a manner analogous to the ER of plasmodesmata in plants (Zambryski, 2004). It will be important in future studies to distinguish between the roles of the cytoskeletal proteins and the ER in fusome function.
Our results have implications for communication in other syncitial systems as well, such as spermatozoa in mammals (Fawcett et al., 1959; Rasmussen, 1973; Clermont and Rambourg, 1978) and the ectoderm of hydra cnidoblasts. In both of these examples, groups of synchronously developing cells contain evidence of intercellular ER connections between dynamic ring canals (Fawcett et al., 1959). Thus, a shared ER may be general mechanism for coordinated cyst development.
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ACKNOWLEDGMENTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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Footnotes |
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Abbreviations used: ER, endoplasmic reticulum; FRAP, fluorescence recovery after photobleaching; FLIP, fluorescence loss in photobleaching; Deff, effective diffusion coefficient; Mf, mobile fraction; ROI, region of interest.
* Corresponding author. E-mail address: mlilly{at}helix.nih.gov.
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) loses fluorescence exponentially, whereas adjacent cystocytes (2 and 3; 
and +'s) lose minimal fluorescence. Compare with 
