Here, we provide a dissection protocol required to live-image the late embryonic Drosophila male gonad. This protocol will permit observation of dynamic cellular processes under normal conditions or after transgenic or pharmacological manipulation.
The Drosophila melanogaster male embryonic gonad is an advantageous model to study various aspects of developmental biology including, but not limited to, germ cell development, piRNA biology, and niche formation. Here, we present a dissection technique to live-image the gonad ex vivo during a period when in vivo live-imaging is highly ineffective. This protocol outlines how to transfer embryos to an imaging dish, choose appropriately-staged male embryos, and dissect the gonad from its surrounding tissue while still maintaining its structural integrity. Following dissection, gonads can be imaged using a confocal microscope to visualize dynamic cellular processes. The dissection procedure requires precise timing and dexterity, but we provide insight on how to prevent common mistakes and how to overcome these challenges. To our knowledge this is the first dissection protocol for the Drosophila embryonic gonad, and will permit live-imaging during an otherwise inaccessible window of time. This technique can be combined with pharmacological or cell-type specific transgenic manipulations to study any dynamic processes occurring within or between the cells in their natural gonadal environment.
The Drosophila melanogaster testis has served as a paradigm for our understanding of many dynamic cellular processes. Studies of this model have shed light on stem cell division regulation1,2,3, germ cell development4,5, piRNA biology6,7,8, and niche-stem cell signaling events9,10,11,12,13. This model is advantageous because it is genetically tractable14,15 and is one of the few where we can live-image stem cells in their natural environment3,16,17,18. However, live-imaging of this model has been limited to adult tissue and early embryonic stages, leaving a gap in our knowledge of gonadal dynamics in the late embryo, the precise stage when the niche is first forming and beginning to function.
The late stage embryonic gonad is a sphere, consisting of somatic niche cells at the anterior, and germ cells encysted by somatic gonadal cells throughout more posterior regions19. This organ can be imaged live in vivo up until early embryonic Stage 1717,20,21. Further imaging is prevented due to initiation of large-scale muscle contractions. These contractions are so severe that they push the gonad out of the imaging frame, and such movement cannot be corrected with imaging software. Our lab is interested in unveiling the mechanisms of niche formation, which occurs during this elusive period for live-imaging. Therefore, we generated an ex vivo approach to live image the gonad starting at embryonic Stage 16, facilitating the study of the cell dynamics during this crucial period of gonad development. Previous work from our lab shows that this ex vivo imaging faithfully recapitulates in vivo gonad development17. This technique is the first and only of its kind for the Drosophila embryonic gonad.
Here, we present the dissection protocol required for ex vivo live-imaging of the gonad during late embryonic stages. This protocol can be combined with pharmacological treatments, or transgenic manipulation of specific cell lineages within the gonad. Using this technique, we have successfully imaged the steps of stem cell niche formation17. This imaging approach is thus instrumental for the field of stem cell biology, as it will enable visualization of the initial stages of niche formation in real time within its natural environment15,17. While this method is beneficial for the field of stem cell biology, it is additionally applicable for visualizing any dynamic processes occurring in the gonad during this developmental timepoint, including cellular rearrangements22, cell adhesion2,12,23, and cell migration23. This dissection protocol will thus enhance our understanding of many fundamental cell biological processes.
1. Day-before-dissection preparation
2. Embryo collection—15–17 h before dissection
3. Day-of-dissection preparation
4. Dissection
NOTE: These steps must be carried out under a stereo-fluorescent microscope.
5. Imaging
We illustrate preparation of the imaging dish in Figure 1, as described in “Day-of-dissection preparation.” These methods should ultimately result in well-hydrated embryos adhered to a cover slip strip, which is temporarily fixed to the bottom of the dish and submerged in Ringer’s solution (Figure 1F). A diamond-tipped knife allows one to cleanly slice a 22 x 22 mm cover slip into three to four smaller strips (Figure 1A). While handling these strips with forceps, we use a pipette to transfer enough heptane-glue to coat these strips, which gives them an adhesive, textured surface (Figure 1B). The coated strips are easily stored in an empty cover slip box to keep the adhesive surfaces clean for up to 2 h (Figure 1C). Once these coated strips are made, the goal is to adhere embryos to the strip in an aggregate, near the long edge of the strip (Figure 1E). It is necessary to first transfer a few embryos from the watch glass onto a clean glass slide and dry them (Figure 1D). Then, quickly use forceps to gently touch the coated strip to the embryos so that they adhere (Figure 1E). We then immediately press the strip to the bottom of the dissection dish with embryos facing up, and cover the embryos with Ringer’s solution (Figure 1F). If this goal is met, then viewing embryos under a traditional brightfield stereomicroscope will reveal fully hydrated, healthy embryos within their vitelline membranes (Figure 1G). If, instead, the process of transferring these embryos onto the strip and covering them with solution takes more than approximately 30 s, the embryos will dehydrate and become flaccid (Figure 1G’). Flaccid embryos are not healthy and are incredibly challenging to dissect, so working efficiently during this process is vital.
Figure 1: Mounting and hydrating embryos prior to dissection. (A–F) Steps for adhering embryos to a strip of glue-coated coverslip, and securing the strip to an imaging dish. (A) Coverslip scored once by diamond-tipped knife (knife indicated by arrowhead) to create a strip. (B) Application of heptane-glue to severed coverslip strip, held with a pair of forceps. (C) Four glue-coated strips drying in a coverslip box. (D–E) Arrows point to embryos. (D) Dechorionated embryos that have been collected in heptane and expelled onto the edge of a microscope slide. Excess heptane was removed with a kimwipe. (E) Glue-coated strip with embryos attached. (F) The final setup of the dish immediately prior to dissection. Note the shallow layer of Ringer’s solution (arrowhead) and placement of the strip (arrow) above the inner dissection circle. (G–G’) Embryos adhered to the strip in the dish. (G) Properly-hydrated, turgid embryos. (G’) Embryos that have become dehydrated due to prolonged air exposure, evident by collapsed vitelline membranes. Asterisks indicate flaccid embryos. Scale bars are 0.5 mm. Please click here to view a larger version of this figure.
Viewing an embryo under a stereo-fluorescent microscope allows clear visualization of the gut, which auto-fluoresces in the GFP channel. Gut morphology serves as a proxy for embryonic age when choosing embryos to dissect. Because embryos adhered to the strip of cover slip will vary slightly in age, they will present a diverse array of gut morphologies (Figure 2A). To live-image gonad niche morphogenesis, we dissect early Stage 16 embryos. These embryos present four regionalized gut sections that are stacked in an even row (Figure 2B’, dotted lines). Younger embryos present a sac-like, non-regionalized gut (Figure 2C), and do not yet have sufficient extracellular matrix (ECM) around their gonads to allow for efficient culturing of the intact organ. Older embryos that have already begun niche compaction present four gut regions that are not evenly stacked and are instead shifted relative to one another (Figure 2D’, dotted lines). These embryos have thicker cuticle, which makes the dissection process more challenging.
Figure 2: Selecting appropriately-aged embryos for gonad dissection. Embryos adhered to a glue-coated coverslip prior to dissection. Embryos express six4-eGFP::moesin (green), which marks the gonad and fat body cells. Note that the gut auto-fluoresces in green. Arrows indicate male gonads (discerned by the presence of brightly fluorescing msSGPs) that are visible in each panel. Scale bars indicate 0.25 mm. (A) Embryos of various stages. (B–D) Embryos of distinct stages in the center of the image, oriented with anterior to the left and dorsal up. (B) An early Stage 16 embryo that is aged appropriately for dissection. (B’) The four stacked gut regions are indicated with dotted white lines. (C) A Stage 15 embryo that is too young to dissect (lower embryo). Note that the fluorescent gut sac just anterior to the gonad does not yet have discrete regions. (D) A late Stage 16 embryo that is difficult to dissect due to developing cuticle. Note that the four gut regions have begun to rotate relative to one another in preparation for gut looping. (D’) The four gut regions are outlined. Please click here to view a larger version of this figure.
It is important to dissect embryos that express an indelible fluorescent marker in the gonad, and this marker must be visible under a stereo-fluorescent microscope. Here, we have chosen to use six4-eGFP::moesin20 to mark the gonad (Figure 2 and Figure 3, arrows) for visualization during dissection.
We illustrate the gonad dissection process in Figure 3. The dissection of appropriately-staged embryos on the poly-lysine-coated dish allows for clean isolation of the gonads from these embryos (Figure 3D). Embryos that are devitellinized will adhere to the dish, and can be arranged in a convenient row prior to dissection (Figure 3A). Embryo devitellinization and transfer to the poly-lysine is an imprecise process such that tissue may become extruded from the confines of the embryo body (see piece of tissue between Embryo 1 and Embryo 2, and mangled Embryo 4; Figure 3A). This imprecision is of no importance, as long as the gonads remain unscathed. A standard stereo-fluorescent microscope enables identification of the gonad within the devitellinized embryo body (Figure 3B, arrow). The first few manipulations with a sharp dissection needle should separate the gonads from the embryo carcass, though some auto-fluorescent tissue will remain adhered (Figure 3C). Additional manipulations result in isolated gonads that adhere directly to the coated dish (Figure 3D).
Figure 3: The dissection process. (A–D) Embryos that express six4-eGFP::moesin (green) at sequential stages in the dissection protocol. (A) Four devitellinized embryos that have been transferred to the poly-lysine-coated dissection region of the dish. Note how the embryos are aligned in a neat row to facilitate successive downward dissections in the dish. The small piece of tissue between Embryos 1 and 2 is part of the gut from Embryo 2, extruded during hand devitellinization. (B) Higher magnification view of embryo from (A), indicated by the asterisk. (C) The remaining carcass of an embryo that has been filleted down the midline. The arrowhead indicates an occluded gonad, still heavily embedded in embryonic tissues. (D) A completely dissected gonad. Note only minimal extraneous tissue (arrowhead) remains near the gonad. Arrows point to male gonads. Scale bars are 0.25 mm. Please click here to view a larger version of this figure.
Once the gonads are dissected, one can replace the Ringer’s solution with imaging media, and image cultured gonads directly in the same dish used for dissection. We use a spinning disk confocal microscope. Brightfield visualization of an isolated gonad on a confocal microscope reveals a dark shadow surrounding the gonad periphery (Figure 4A–B, arrows), which is the ECM that maintains integrity of the gonad during imaging. We present an example of a healthy, well-cultured gonad that expresses GFP-labeled F-actin in somatic gonadal cells20 and an RFP-labeled histone marker to visualize all nuclei28 (Figure 4C–C’). Because all cells in this embryo express the RFP histone marker, both gonadal cells, and cells of other adherent tissue are observed in viewing red emission. We discern the boundaries of the gonad using the gonad-specific GFP marker (Figure 4C’, outline). It is clear that this cultured gonad is healthy because the gonad boundary is smooth and round (Figure 4C’, outline), and because gonadal cells have even levels of RFP histone fluorescence throughout nuclei (Figure 4C’, arrow). If, instead, gonads are not sufficiently hydrated during imaging, nuclei and six4-eGFP::moesin fluorescence can become punctate (Figure 4D, arrow) as the gonad tissue shrivels. Further, if the gonad ECM is damaged excessively during dissection, the gonad boundary is compromised, which is evident by the presence of gonad-specific cells (Figure 4E, asterisks) outside of the confines of the gonad (Figure 4E, arrow).
Figure 4: Locating healthy gonads during live-imaging. (A) Low and (B) high magnification brightfield views of two dissected gonads adhered to the coverslip. (C–C’) One movie frame from ex vivo imaging of niche compaction in a healthy gonad. Somatic gonadal cells express six4-eGFP::moesin (green), and all cells express His2Av-mRFP (red). (C’) Gonad boundary marked with a white dotted line. His2Av-mRFP visible outside of the gonad boundary is likely fat body that is still attached to the dissected gonad. Arrow points to a germ cell nucleus with uniform His2Av-mRFP signal, indicating the gonad is healthy. (D–E) Representative negative outcomes of the ex vivo dissection protocol. (D) A frame from an imaging session in which the gonad has dehydrated because of media evaporation. Note pyknotic nuclei as His2Av-mRFP condenses (arrow), and discontinuous spots of six4-eGFP::moesin along the gonad boundary. (E) Ex vivo imaging frame in which extracellular matrix has been compromised during dissection. Note that some germ cells (asterisks) are exiting the gonad boundary (arrow). Germ cells are labeled with nos-lifeact::tdtomato (magenta), and somatic gonadal cells with six4-eGFP::moesin (green). Scale bars show 20 μm. Please click here to view a larger version of this figure.
Gonads can be cultured for about 5 h using this ex vivo imaging method, enabling image acquisition of the dynamic morphogenesis events that occur in late embryonic gonad development. In our lab, we have successfully used this protocol to image the compaction of the forming stem cell niche17 (Figure 5). Prior to compaction, the niche is a loose aggregate of somatic cells at the gonad anterior (Figure 5A, green), surrounded by germline cells labeled with nos-lifeact::tdtomato29 (see Table of Materials), the first tier of which will be germline stem cells (Figure 5A, magenta). This niche aggregate initially presents with an irregular boundary (Figure 5A’, dotted line). Throughout the course of imaging, we observe individual niche cells rearranging their positions while the niche aggregate acquires smoother borders. These cellular rearrangements also result in a decrease in niche area (Figure 5C). Our observations of cell rearrangements, and neighboring germ cell divisions that occur concurrently with this phase of development informed our understanding of mechanisms underlying niche compaction17. This protocol thus affords the ability to visualize cell rearrangements, shape changes, divisions, and other cellular events with the resolution required to analyze morphogenetic events in late stage gonads.
Figure 5: An ex vivo cultured gonad undergoing niche compaction. Stills from an imaging series acquired promptly following dissection of gonads from early Stage 16 embryos. (A–C) Germ cells (magenta) are labeled by nos-lifeact::tdtomato and somatic gonadal cells (green) are labeled by six4-eGFP::moesin. The niche (dotted white line) is outlined in A’–C’. Scale bars show 10 μm. Anterior is to the left, and posterior is to the right. (A) At the first timepoint (t = 0 min) in the imaging series, niche cells have finished assembling at the gonad anterior and are at an early stage of compaction. (B) Midway through the imaging series and the compaction process (t = 1 h 48 min), the niche has begun to circularize, but its edge remains irregular. (C) Near the end of the imaging series (t = 4 h 21 min), the niche has a highly smoothened, circularized boundary. Please click here to view a larger version of this figure.
During gonadogenesis, the embryonic gonad, and particularly the stem cell niche within the male gonad15, undergoes rapid morphological changes. Developmental mechanisms that underlie these dynamic changes are best understood through live-imaging techniques. However, at embryonic Stage 17, in vivo imaging of the gonad is rendered impossible by the onset of large-scale muscle contractions17. With this protocol, we provide a successful alternative: dissection of the gonads directly onto an imaging dish for ex vivo live-imaging. This protocol presents the only method available to accomplish live-imaging of the late stage embryonic gonad.
The critical steps of the protocol should be executed with an acute focus on dexterity and timing. Prior to dissection, rapid mounting of embryos is paramount to ensure that the embryos do not dehydrate and remain healthy and turgid, which eases devitellinization and dissection. During dissection, it is vital to avoid disruption of the gonadal extracellular matrix, which is accomplished using only delicate and precise manipulations. Use of such dexterity will also ensure that the gonad is in direct contact with the imaging dish, thereby enabling clear imaging directly through the coverslip. After dissection, Ringer’s solution must be switched for imaging media using only gentle suction and expulsion with a pipette to prevent gonad dislodgement. Finally, it is important to limit the overall dissection time to 25 min. This ensures that the amount of time spent in Ringer’s solution during dissection, and location of the tissue at the confocal, is limited to a non-toxic exposure. With increased practice, dissection speed will improve, and personalization of the dissection sequence will occur. In our experience, sufficient dissection may be achieved after about a week of regular practice, with full mastery of the dissection attainable in about 1 month. To ease learning, we recommend practicing individual steps before executing the entire protocol.
There are a number of best practices that ameliorate the challenges associated with this protocol, starting with the genotype of the embryos used for dissection. Incorporation of multiple copies of the transgene used to mark the gonad will make the gonad brighter, and therefore easier to visualize and dissect. The dissection itself works best with a sharp needle, though it should not be overly sharpened, as it will bend against the bottom of the imaging dish and become burdened with extraneous tissue. Schneider’s imaging media auto-fluoresces in the green emission spectrum, which makes it challenging to locate gonads marked with GFP once the media is added. Therefore, it is critical to use the confocal imaging software to mark the location of gonads while they are still in the Ringer’s solution. If locating gonads at the confocal remains difficult, after the next dissection we suggest sketching or taking an image of the relative positioning of gonads in the dish while still at the dissection microscope. Then, mark the outside of the dish to indicate its orientation during dissection, and match this orientation when transitioning to the confocal microscope. Once located at the confocal, if a gonad appears disrupted or mangled, in the next dissection try leaving more adherent tissue surrounding the gonad. On the contrary, if a gonad is present but appears blurry throughout all planes, there is likely tissue between the coverslip and the gonad, and future dissections should strive for better isolation of the gonad from the adherent tissue. In our experience, adopting these practices has facilitated learning and execution of this protocol.
During the formation of this protocol, we identified several alterations that could be applied. For niche compaction studies, we aim to dissect embryos at Stage 16. At this stage, the embryo has not yet developed significant amounts of cuticle, and easily sticks to the poly-lysine-coated dish. However, this technique may be modified to dissect older embryos with cuticle by sticking them to the inner wall of the poly-lysine-coated region for the first incision. Once internal tissues are exposed, the embryo carcass will stick to the poly-lysine, and dissection can proceed as normal. In addition to adjustments for embryo age, we have successfully adjusted this technique to dissect multiple genotypes in the same dish. For example, homozygous mutants can be separated from sibling heterozygotes by the use of an easily discernable marker such as deformed-YFP on the balancer chromosome. After devitellinization, embryos should be transferred to either side of the coverslip based on the presence or absence of the marker. Dissection should proceed downwards in the dish as described in the protocol, with extra care taken to maintain segregation of different genotypes. Also, this protocol could potentially be modified to use culture medium other than Schneider’s, though a superficial investigation of Shields and Sang M3 Insect Media yielded unviable gonads (data not shown). Additionally, this protocol pairs well with pharmacological manipulations by simply combining the drug with the imaging media. Over a 5 h period of imaging, re-addition of drug might be necessary to maintain an effective concentration. Finally, although this protocol was developed with the intention of visualizing niche compaction, a phenomenon specific to male gonads, in theory this protocol could also be implemented to live-image the developing ovary by simply dissecting and imaging gonads that lack a niche and msSGPs.
Limitations associated with this technique relate to the length of culturing time, and the ex vivo nature of the culture. As is the case with mid-stage Drosophila egg chambers30, cultured gonads begin to die after about 5 h of ex vivo live-imaging, evidenced by loss of tissue integrity (nuclei become pyknotic and cell membranes shrivel). Thus, if one wished to explore later-stage events in the male gonad, one would need to dissect gonads from 1st instar or older larvae by making modifications to the protocol presented here, and then image those under conditions similar to those presented here. However, we do not rule out the possibility of ex vivo live-imaging past five hours if improved culture conditions are developed, though there may be a limit if mechanical cues from within the embryo are necessary for gonad development. Perhaps the most severe drawback of the technique is due to its inherent ex vivo nature. When cultured ex vivo, cells can have unnatural potential that is unrepresentative of in vivo biology31. Additionally, subtleties of culturing environments, including media content and matrix stiffness, can have drastic influence over cell behavior32. With this sensitivity in mind, it is vital to ensure that culturing conditions accurately reflect in vivo biology. We have taken measures to attest that in vivo niche development and signaling is recapitulated ex vivo17. Briefly, we ascertained that niche cell fate is maintained and number of niche cells is unchanged using the markers Fasciclin-III and E-cadherin. Also, STAT signaling is present and germline stem cells divide orthogonally, suggesting that niche functionality is maintained. However, we have not verified that the relevant in vivo biology remains intact in other, non-niche tissues within the gonad. Future ex vivo investigations of these other tissues should include a comprehensive analysis to ensure that this is indeed the case.
A future direction one might take with this protocol would include incorporation of a barrier within the imaging dish, such that one dissection session may contain both a control group as well as a drug-treatment group. This enhancement would allow for minimization of error between technical replicates, thereby improving scientific rigor.
There are no other true alternatives to this protocol, as it is the only method available to examine the live events of this organ during late embryonic development. As such, previously unanswerable questions about gonad morphogenesis are now accessible with this advancement. These questions include the intricacies of cell divisions, cytoskeletal changes, and cell intercalation events that govern stem cell niche formation and gonadogenesis. None of these events are detectable in the still images of these processes acquired using a fix-and-stain technique. Overall, this method unlocks the possibility of investigating the dynamics in the late embryonic Drosophila gonad, and such a breakthrough has strong implications for the advancement of a myriad of biological fields, including stem cell-niche biology, piRNA biology, and organogenesis.
The authors have nothing to disclose.
We would like to thank Lindsey W. Plasschaert and Justin Sui for their substantial contributions to the early development of this protocol. The authors are grateful to the fly community for their generosity with reagents, and particularly to Ruth Lehmann and Benjamin Lin for their gift of the nos5’-Lifeact-tdtomato p2a tdkatushka2 Caax nos3' line prior to its publication. Stocks obtained from the Bloomington Drosophila Stock Center (NIH P40OD018537) were used in this study. This work was supported by NIH RO1 GM060804, R33AG04791503 and R35GM136270 (S.D) as well as training grants T32GM007229 (B.W.) and F32GM125123 (L.A.).
Alfa Aesar Tungsten wire | Fisher Scientific | AA10408G6 | 0.25mm (0.01 in.) dia., 99.95% (metals basis) |
D. melanogaster: His2Av::mRFP1 | Bloomington Drosophila Stock Center (BDSC) | FBtp0056035 | Schuh, Lehner, & Heidmann, Current Biology, 2007 |
D. melanogaster: nos-lifeact::tdtomato | Gift from Ruth Lehmann Lab | Lin, Luo, & Lehmann, Nature Communications, 2020: nos5'- Lifeact-tdtomato p2a tdkatushka2 Caax nos3' | |
D. melanogaster: P-Dsix4-eGFP::Moesin | FBtp0083398 | Sano et al., PLoS One, 2012 | |
Diamond-tipped knife | |||
Double-sided tape | Scotch | 665 | |
Fetal Bovine Serum | GIBCO | 10082 | |
Imaging dish | MatTek | P35GC-1.5-14-C | |
Imaging software | Molecular Devices | MetaMorph Microscopy Automation and Image Analysis Software v7.8.4.0 | |
Insulin, bovine | Sigma | l0516 | Store aliquots at 4 °C |
Needle holder | Fisher Scientific | 08-955 | |
Nytex basket | |||
Penicillin/streptomycin | Corning | 30-002-Cl | |
Ringer's solution | 2 mM MgCl2, 2 mM CaCl2, 130 mM NaCl, 5mM KCl, 36 mM Sucrose, 5mM Hepe’s Buffer; adjusted with NaOH until pH of 7.3 is achieved | ||
Schneider's Insect Media | GIBCO | 21720-024 |