Here, we discuss a workflow to prepare, dissect, mount, and image live explant brains from Drosophila melanogaster third instar larvae to observe the cellular and subcellular dynamics under physiological conditions.
Drosophila neural stem cells (neuroblasts, NBs hereafter) undergo asymmetric divisions, regenerating the self-renewing neuroblast, while also forming a differentiating ganglion mother cell (GMC), which will undergo one additional division to give rise to two neurons or glia. Studies in NBs have uncovered the molecular mechanisms underlying cell polarity, spindle orientation, neural stem cell self-renewal, and differentiation. These asymmetric cell divisions are readily observable via live-cell imaging, making larval NBs ideally suited for investigating the spatiotemporal dynamics of asymmetric cell division in living tissue. When properly dissected and imaged in nutrient-supplemented medium, NBs in explant brains robustly divide for 12-20 h. Previously described methods are technically difficult and may be challenging to those new to the field. Here, a protocol is described for the preparation, dissection, mounting, and imaging of live third-instar larval brain explants using fat body supplements. Potential problems are also discussed, and examples are provided for how this technique can be used.
Asymmetric cell division (ACD) is the process by which subcellular components such as RNA, proteins, and organelles are partitioned unequally between daughter cells1,2. This process is commonly seen in stem cells, which undergo ACD to give rise to daughter cells with different developmental fates. Drosophila NBs divide asymmetrically to produce one NB, which retains its stemness, and one ganglion mother cell (GMC). The GMC undergoes further divisions to produce differentiating neurons or glia3. Asymmetrically dividing NBs are abundant in the developing brains of third-instar larvae, which are readily observed via microscopy. At the third instar larval stage, there are roughly 100 NBs present in each central brain lobe3,4,5,6.
Asymmetric cell division is a highly dynamic process. Live-cell imaging protocols have been used to measure and quantify the dynamics of cell polarity7,8,9,10, spindle orientation11,12,13, the dynamics of the actomyosin cortex14,15,16,17,18, microtubule and centrosome biology19,20,21,22,23,24,25,26,27, and membrane10,28 and chromatin dynamics29. Qualitative and quantitative descriptions of ACD rely on robust methods and protocols to image dividing NBs in intact living brains. The following protocol outlines methods to prepare, dissect, and image third instar larval brains for live-cell imaging in vivo using two different mounting approaches. These methods are best suited for researchers interested in the spatiotemporal dynamics of stem cell divisions, as well as divisions in other brain cells, as they allow for short- and long-term observations of cellular events. Additionally, these techniques are readily accessible to newcomers to the field. We demonstrate the effectiveness and adaptability of this approach with larval brains expressing fluorescently tagged microtubule and cortical fusion proteins. We additionally discuss methods of analysis and considerations for application in other studies.
NOTE: Figure 1 shows the materials required to perform this study.
1. Considerations and preparations for the experiment
2. Larvae staging and collection (Figure 2)
3. Larval fat body dissection (Figure 3)
NOTE: This protocol describes dissections using a 3-well dissection dish.
4. Larval brain dissection (Figure 3)
5. Mounting and imaging (Figure 4)
6. Data processing and management best practices
7. Example quantification of cell cycle length (Figure 5)
NOTE: in this example, larvae expressing the polarity marker Pins (Pins::EGFP16) and the microtubule-binding protein Jupiter25 (cherry::Jupiter13) were imaged. The subsequent analysis was performed using Imaris software.
8. Example quantification of cell spindle alignment (Figure 5)
NOTE: In this example, the analysis is performed using Imaris software.
Dissection and imaging of central brain lobe NBs expressing Pins::EGFP and Cherry::Jupiter
To showcase this protocol, larvae expressing UAS-driven Cherry::Jupiter13 and endogenously tagged Pins::EGFP16 (w; worGal4, UAS-cherry::jupiter/CyO; Pins::EGFP/TM6B, Tb) were imaged for 4 h using the described protocol using multi-well imaging slides (Figure 5C,D). Additional data were taken from larvae expressing UAS-driven Cherry::Jupiter13 and endogenously tagged Miranda::EGFP (w; worGal4, UAS-cherry::Zeus/CyO; UAS-Miranda::GFP/TM6B), which were imaged for 10 h using a membrane-bound slide (Figure 5E,F). Larvae were reared in cages as described in section 1 of this protocol. Upon reaching 72-96 h old, the larvae were dissected (Figure 3), mounted (Figure 4), and imaged. For the experiments performed here, a 561 nm laser was used at 10% laser power with 100 ms of exposure time, and a 488 nm laser was used at 15% laser power with 100 ms of exposure time. Z-stacks (41 µm) were acquired with a 1 µm step size. Images were acquired every 5 min at RT on an Intelligent Imaging Innovations (3i) spinning disc confocal system, consisting of a Yokogawa CSU-W1 spinning disc unit and two Prime 95B Scientific CMOS cameras. A 60x/1.4NA oil immersion objective mounted on a Nikon Eclipse Ti microscope was used for imaging. The live imaging voxels were 0.22 µm x 0.22 µm x 0.75 µm (60x/1.4NA spinning disc).
Consistent with previous reports16, the Pins formed a pronounced apical crescent in dividing NBs during mitosis, and the mitotic spindles consistently aligned to this apical crescent (Figure 5C). The cell cycle length was determined by measuring the time between successive metaphases of the individual NBs (Figure 5D,F).
In samples that were imaged on a multi-well imaging slide for 4 h without fat body supplementation, the cell cycle length increased with increasing imaging time (Figure 5C,D). Samples that were imaged in fat body-supplemented medium on a membrane-bound slide did not show an increase in cell cycle length (Figure 5E, F). Furthermore, NBs with four divisions were observed on the 10 h membrane-bound slide (Figure 5D vs. Figure5F).
Lastly, the angle between the division axis and the mitotic spindle was determined using GFP-tagged Pins as a reference (schematic shown in Figure 5G). The division axis was determined by identifying the midpoint of the apical crescent formed by the Pins in mitosis and bisecting the cell in half (Figure 5G, red dashed line). As previously described, the wild-type NBs displayed mitotic spindles that were oriented no more than 30° from the division axis (Loyer and Januschke36 and Figure 5H).
Figure 1: Materials. (A) Dissection microscope. (B) Collection cage containing flies of the desired genotype and a meal cap with growing larvae. (C) Dissection and imaging medium, 5 mL. (D) Meal caps with larvae from three different collections. (E,E') Microdissection tools, from left to right: microdissection scissors, forceps. (F,F') Dissection dish. (G) An 8-welled µ-slide for imaging the samples. Please click here to view a larger version of this figure.
Figure 2: Example meal caps. (A) Two vials with male and female flies to be crossed, an empty embryo collection cage, and a fresh meal cap. (B) The bottom of the embryo collection cage with the new meal cap (left) and the top of the cage with flies (right). (C) The fully assembled fly cage. (D) An example of a well-staged meal cap with larvae for dissection. Note that the food is disturbed by the larvae, but not over-disturbed. (E,F) Two examples of 4 day old, overcrowded meal caps. Note that that food now has a soup-like consistency to it. Please click here to view a larger version of this figure.
Figure 3: Dissection. (A) Dorsal view of a third instar larva. The red line on the posterior end of the larva denotes where the first cut should be made. (B) Dorsal view of the larva after removing the posterior end. (C) Diagram showing how to invert the larva. Using one set of forceps, hold the larva by the cuticle near where the first cut was made. Using the other forceps, press into the anterior end of the larva to invert. The black arrows denote the direction of the forceps, with one "pushing into" the larvae from the anterior side and the other moving the cut posterior end towards the anterior end. The smaller red arrow denotes a cartoon of fat bodies. (D) View of an inverted larva with the fat bodies and digestive tract still attached. (E) View of an inverted larva with the non-CNS tissue removed. The red dashed line outlines the still-attached brain. (F) Schematic showing how to remove the brain from the cuticle. The red dashed line indicates the path to cut with microdissection scissors to release the brain from the inverted cuticle, and the black arrows denote the removal of the brain from the cuticle. (G) A view of the inverted brain that is still attached to the cuticle by a small number of axonal connections under the ventral nerve cord (VNC). (H) An isolated larval brain. The brain lobes are outlined with dashed orange lines. (I) Isolated fat bodies. Please click here to view a larger version of this figure.
Figure 4: Mounting and imaging. (A) View of the components for assembling a membrane-bound metal slide. (B) Schematic of the components of the membrane-bound slide and their assembly. (C) Side view of the membrane inserted into the metal slide, held in place by the split metal ring. (D) Top view of the assembled imaging slide without a coverslip. The dissection and imaging medium containing fat bodies and dissected brains has been placed onto the membrane. (E) Zoomed-in view of the fat bodies and brains in the drop of medium. (F) Top view of the metal slide after adding the glass coverslip. (G) Top view of the assembled slide with the glass coverslip fixed to the slide with melted petroleum jelly. Covering the edges of the coverslip with petroleum jelly also prevents evaporation of the medium. (H) Schematic of the assembled membrane slide with brains oriented for observation on an inverted microscope. The blue circles denote NBs in the central brain lobes and VNC. (I) View of an empty multi-well slide. (J) Zoomed-in view of one well of the multi-well imaging slide. (K) Schematic showing the brain orientation for imaging the central brain lobes on an inverted microscope with a multi-well slide. The blue circles denote NBs in the central brain lobes and VNC. Please click here to view a larger version of this figure.
Figure 5: Quantification of the cell cycle length. (A) Diagram of a third instar larval brain, highlighting the brain lobes, the optic lobe (OL, dark gray), the ventral nerve cord (VNC), NBs (dark blue), GMCs (light blue), and neurons (purple) within the central brain lobes and VNC. (B) Schematic of NB and GMC divisions. (C) Image series of a wild-type NB with microtubules labeled in white (MTs, UAS-Cherry::Jupiter) and apical Pins (Pins::EGFP in green) imaged in a multi-well slide without fat body supplementation for 4 h. Merged images and the corresponding lineage tree with cell cycle timing are shown below. Scale bar = 10 µm. (D) Quantification of the cell cycle length (metaphase – metaphase) for the first, second, and third divisions in samples imaged on a multi-well slide without fat bodies.(E) Image series of a wild-type NB with microtubules labeled in white (MTs, UAS-Cherry::Jupiter) imaged with a membrane-bound slide with fat body supplementation for 10 h with the corresponding lineage tree and cell cycle timing shown below. Scale bar = 10 µm. (F) Quantification of cell cycle length (metaphase – metaphase) for the first, second, third, and fourth divisions in samples imaged on a membrane-bound slide with fat bodies. (G) Schematic of how the angle between the spindle axis (orange dashed line) and division axis (θ, red dashed line) was determined. (H) Quantification of θ from 10 cells imaged using a multi-well slide without fat bodies. Please click here to view a larger version of this figure.
This protocol outlines one approach for the imaging of live explant brains from Drosophila melanogaster larvae. The protocol described here allows for explant brains to be observed for 12-20 h under the right experimental conditions. Special consideration must be given to the preparation of samples and the design of the desired experiments. As mentioned above, one of the most critical factors that determines the quality of the dissected tissue is the health of the larvae. To achieve the highest quality possible, one must ensure that larvae are well-fed before collection. Unhealthy larvae most commonly originate from overcrowding. To address this, one must ensure that overcrowding is minimized, either by increasing the frequency of harvest or by splitting dishes with freshly laid eggs with an empty dish.
Another critical element of this protocol is the method of dissection to isolate the brain tissue. The brain and the NBs within them are extremely sensitive to outside factors, such as the temperature of the dissection medium and the invasiveness of the dissection itself. Medium that is too cold will tend to depolymerize the microtubules. Similarly, a dissection that stretches or ruptures the brain will have detrimental effects on the quality of the NBs. To prevent this, one should avoid pulling on the brain tissue directly and instead anchor the dissection tools on the cuticle or other tissue. Microdissection scissors greatly assist in this regard, as they minimally pull on the brain tissue. If scissors are not available, tweezers can be used to carefully remove the connecting tissue between the ventral nerve cord and the cuticle.
This protocol presents two methods to mount larval brains for confocal microscopy. From a technical standpoint, mounting samples in a multi-well slide is simpler than mounting on a membrane-bound slide. However, each method is best suited for different types of experiments. The data shown here demonstrate that for shorter movies (i.e., less than 4 h) or movies with high temporal resolution (i.e., acquiring a z-stack every 10 s), imaging in a multi-well slide is sufficient to observe multiple divisions in the larval central brain. For longer acquisition windows, mounting on a membrane-bound slide is ideal, as samples prepared this way divide more frequently throughout the movie. Normally, wild-type larval NBs divide once every 40-90 min13. Although multi-well slides can be used for long (> 4 h) movies, an increase in the cell cycle length and a decrease in proliferating NBs has been observed in this condition (Figure 5D). Therefore, the method used to mount samples and the type of data to be measured must be considered when designing experiments.
This protocol recommends imaging multiple brains in one well or slide, as this increases the efficiency of data collection for any given experiment. However, the orientation and placement of the brains within the well will affect how much shifting occurs throughout the movie due to the stage moving between brain positions. Clustering the brains together in one centralized spot in a well minimizes this issue. However, some shifting may occur over the course of longer movies in experiments using multi-well slides. In many cases, the shifting observed in these longer movies is minimal and can be corrected during analysis. Brain movement is also preventable by using a metal slide setup because the capillary forces prevent the dissected brains from shifting.
With a multi-well slide, it is technically possible to perform multi-well imaging experiments. However, this requires adjustments in the stack size and temporal resolution to account for the extra time spent by the microscope moving between positions. This may be beneficial for large-scale genetic screening experiments, where one may image multiple genotypes in different wells.
There are instances where brains will display little or no dividing NBs over the course of an experiment. This is likely due to several factors that depend on the nature of the sample being imaged, the quality of the food used to rear the larvae, the quality of the dissection, and the acquisition settings used to generate the data. Although it is advisable to remove as much tissue as possible when preparing larval brains for imaging, it is ill-advised to over-prune the brain to minimize mechanical stress, as this may negatively impact the quality of the data. Additionally, frequent exposure to powerful imaging lasers will negatively impact the neuroblast health. Thus, one should consider adjusting the laser power, exposure time, and imaging frequency when collecting the imaging data.
For mounting samples, alternative methods may be used. For instance, explant brains can be mounted in a solid matrix37. The availability of different mounting protocols provides an opportunity to image larval brain explants on a wide variety of microscopes.
The authors have nothing to disclose.
This research is supported by R35GM148160 (C. C.) and a National Institutes of Health (NIH) Training Grant T32 GM007270 (R. C. S)
0.22 µm polyethersulfone (PES) Membrane | Genesee | 25-231 | Vacuum-driven filters |
Agar | Genesee | 20-248 | granulated agar |
Analytical Computer | Dell | NA | Intel Xeon Gold 5222 CPU with two 3.80 GHz processors running Windows 10 on a 64-bit operating system |
Bovine Growth Serum | HyClone | SH30541.02 | |
Chambered Imaging Slides | Ibidi | 80826 | |
Confocal Microscope | Nikon | NA | |
Custom-machined metal slide | NA | NA | See Cabernard and Doe 2013 (Ref. 34) for specifications |
Dissection Dishes | Fisher Scientific | 5024343 | 3-well porcelain micro spot plate |
Dissection Forceps | World Precision Instruments | Dumont #5 | |
Dissection Microscope | Leica | NA | |
Dissection Scissors | Fine Science Tools (FST) | 15003-08 | |
Embryo collection cage | Genesee | 59-100 | |
Flypad with access to CO2 to anesthetize adult flies | Genesee | 59-172 | |
Gas-permeable membrane | YSI | 98095 | Gas-permeable membrane |
Glass Cover Slides | Electron Microscopy Sciences | 72204-03 | # 1.5; 22 mm x 40 mm glass coverslips |
Imaris | Oxford Instruments | NA | Alternatives: Fiji, Volocity, Aivia |
Imaris File Converter | Oxford Instruments | NA | |
Instant Yeast | Saf-Instant | NA | |
Molasses | Genesee | 62-117 | |
Petri dish | Greiner Bio-One | 628161 | 60 mm x 15 mm Petri dish |
Petroleum Jelly | Vaseline | NA | |
Schneider's Insect Medium with L-glutamine and sodium bicarbonate liquid | Millipore Sigma | S0146 | |
SlideBook acquisition software | 3i | NA | |
Vacuum-Driven Filtration Unit with a 0.22 µµm PES membrane filter | Genesee Scientific, GenClone | 25-231 |