Materials and Reagents

1.7 mL microtubes (Axygen, catalog number: MCT-175-C)
48-well Nunc cell plate (Thermo Scientific, catalog number: 150687)
Micropipette 200 µL tips (Axygen, catalog number: T-200-Y)
Double-sided tape
Cotton (Genesee Scientific, catalog number: 51-101)
25 × 95 mm polystyrene Drosophila culture vials (Genesee Scientific, catalog number: 32-109RL)
Microscopy slides (Westlab, catalog number: 663-249)
Cover slips (Trajan, catalog number: 471112440M)
90 mm Petri dishes
MitoTimer reporter Drosophila strain (Bloomington Drosophila Stock Center, # line: 57323)
Neuronal Gal4 driver Drosophila strain (Bloomington Drosophila Stock Center, # line: 458)
Phosphate-buffered saline (PBS) (Sigma-Aldrich, catalog number: P5493)
16% Formaldehyde solution (Thermo Fisher Scientific, catalog number: 28908)
Spinosad (Sigma-Aldrich, catalog number: 33706)
Dimethyl Sulfoxide (DMSO) (Sigma-Aldrich, catalog number: 276855-100ML)
Vectashield 10 mL Mounting medium (Vector, catalog number: VEH1000)
Analytical standard sucrose (Merk, product number: 47289)
Fly media (1 L) (see Recipes)
Apple juice-agar plates (1 L) (see Recipes)
Tegosept (250 mL) (see Recipes)
Acid mix (250 mL) (see Recipes)
20% Sucrose solution (200 mL) (see Recipes)
5% Sucrose solution (200 mL) (see Recipes)
1,000 ppm spinosad solution in DMSO (see Recipes)
4% Formaldehyde solution (see Recipes)

Equipment

Dissecting forceps (Fine Science Tools, Dumont #5 Forceps, catalog number:1195-10)
Brush (Artist First Choice Taklon round size 3)
Stereo microscope (Zeiss, model: Stemi 2000-C)
Confocal microscope (Leica, model: SP8 Confocal microscope)
Delicate task wipers (Kimtech Science Kimwipes, manufacturer code: 34120A)
Micropipette P200 (Gibson, catalog number: FA10005M)
Incubator 25 °C (Thermoline Scientific)
500 mL laboratory bottles (Sigma-Aldrich, catalog number: Z305197-10EA)
Agitator (Benchmark Scientific Inc, Super Nutation Mixer, catalog number: B3D 1320)
Stove

Software

ImageJ/FIJI (NIH, https://fiji.sc/)
Excel (Microsoft)
R (The R Foundation, https://www.r-project.org/)
Inkscape (https://inkscape.org/)

Procedure

Drosophila rearing
1. Set a vial containing fly media with twenty Neuronal Gal4 driver Drosophila adult virgin females and five MitoTimer reporter Drosophila adult males.
2. Keep vials in an incubator at 25 °C, and transfer adult flies to a new vial containing fly media every 24 h.
3. Set 2 to 4 vials with adults laying eggs per day, to collect enough larvae of the desired stage (early third instar larvae are used here).
Larvae collection
1. To collect early third instar larvae, wait 4 to 5 days from egg laying, depending on larvae population density per vial.
2. Recover larvae from the food using the sucrose extraction method (Nichols et al., 2012). Select a vial containing larvae, and cover half of it with 20% sucrose solution (Figure 1A).
3. Using a brush, gently disrupt the top food layer and wait for a couple of minutes. The larvae will float, and the food will sink back to the bottom.
4. Using a brush, collect the larvae and transfer them to an apple juice-agar plate (Figure 1B).
Insecticide exposure
1. Using the micropipette, load a 48-well Nunc plate with 200 µL of 5% sucrose solution per well.
2. Under the stereo microscope, select early third instar larvae (Figure 1C) and, using a pair of forceps, gently transfer 25 larvae per well (Figure 1D).
3. For the insecticide exposure, create a 5x insecticide solution by diluting a 1,000 ppm insecticide stock solution in 5% sucrose solution (or equivalent dose of DMSO for controls) (Denecke et al., 2015).
4. Add 50 µL of the 5× stock solution to each well containing larvae to be exposed, and give the plate a gentle swirl (Figure 1D).
5. Keep the plate with larvae under insecticide exposure at 25 °C and protected from light for 2 h (or desirable number of hours), until exposure is over.
6. Once exposure time is over, using a pair of forceps, gently transfer larvae to a new well containing 250 µL of 1× PBS (Figure 1E).
Tissue dissection and fixation
1. Transfer larva from the PBS containing well to a microscopy slide containing a drop of 1× PBS.
2. Under the stereo microscope, begin the dissection. Using two pairs of forceps, hold the larval body at the midpoint, and pull the anterior and posterior regions apart (Figure 1F).
3. Using a pair of forceps, gently transfer the anterior body section to a microtube containing 500 µL of 4% formaldehyde solution (Figure 1G).
4. Keep the microtubes at slow agitation in a bench agitator for 20 min, to fix the sample.
5. With a micropipette, slowly and completely remove the fixative solution, whilst avoiding touching the sample. Dispose of the toxic formaldehyde waste appropriately.
6. Gently add 500 µL of 1× PBS back to the microtube, to avoid damaging the sample, and place the microtube back on the agitator for another 5 min.
7. Repeat the previous step two more times.
8. Under the stereo microscope, add a drop of 1× PBS to a new microscopy slide. Using a pair of forceps, gently transfer the fixed sample from the microtube onto the slide (Figure 1H).
9. Conclude the dissection by holding the anterior body section sideways with two pairs of forceps, and gently pull them apart from each other to tear the cuticle. Once the brain is located, use the forceps to clear the surrounding tissues. If PBS starts to dry out, add more of it accordingly.
Mounting slides for fluorescence microscopy
1. Clear the remaining tissue from the slide, leaving only the brain.
2. Using a delicate task wiper’s tip, carefully drain the excess of PBS. Moistening the wiper’s tip in distilled water before touching the PBS, as far away from the fixed sample as possible, will reduce its dragging capacity, and prevent the sample from being sucked into the wiper.
3. Add a small drop of Vectashield on the top of the fixed sample, only enough to cover it. Using a pair of forceps, if necessary, adjust sample orientation.
4. Cut two small pieces of double-sided tape, and place them flanking the sample in Vectashield.
5. Place the coverslip on the top, aligning its border with the double-sided tape (Figure 1I).
Figure 1. Experimental procedure. A. Use the sucrose extraction method to recover larvae from the vial. B. Using a brush, transfer larvae to an apple-juice agar plate. C. Selected early third instar larvae. D. Transfer 25 larvae to a well pre-loaded with 5% sucrose solution, and expose them to insecticide. E. Once exposure is over, transfer larvae to a new well containing 1× PBS solution. F. Under the microscope, begin dissection by separating anterior and posterior body parts. G. Transfer the anterior body part to a microtube containing fixative solution. After fixing the sample, perform three washing steps by replacing the solution in the microtube with 1× PBS. H. Under the microscope, conclude dissection, and isolate the brain from the other tissues. I. Mount the slide for microscopy using double-sided tape, Vectashield, and a cover slip.
Imaging
1. Proceed to image acquisition with a fluorescent microscope immediately.
2. To ensure consistency and accuracy in the measurements, set optimal values of laser power and gain with a control sample, and maintain the same settings across all samples.
3. Once the desired area is localized, acquire both green (excitation/emission 488/518 nm) and red (excitation/emission 543/572 nm) signals at 200× magnification.
4. Acquire at least 20 images across the z-stack for every sample.

Data analysis

Analyzing microscopy images on ImageJ
1. Open the image files on ImageJ and use the freehand selection tool to create a selection area around the area of interest (e.g., whole brain, ventral nerve cord, or optic lobes—used here).
2. On the top menu, under Analyze, select Measure.
3. Copy the mean values of fluorescence intensity generated to an Excel spreadsheet.
4. For every sample, acquire three independent measurements along the z-stack that are representative of the sample signal.
5. Use the same selection area and same z-stack images for both green and red MitoTimer signal measurements per sample.
6. Obtain measurements from the same regions in the different samples.
7. Use at least 10 individual samples per treatment condition for a consistent evaluation of changes on mitochondrial turnover.
8. Identify microscopy images that are representative of control and treated conditions and, using a suitable program (e.g., Inkscape), organize these images for publication (Figure 2A).
Statistical analyses
1. On an Excel spreadsheet, organize the data in columns to show treatment condition, nature of the signal, sample replicate number, and signal values (Figure 2B).
2. Save the file as mitotimer.csv
3. Open the software R and download the packages lme4 and car
4. To perform statistical analysis, use the following script:
  library(lme4)
  library(car)
  mitotimer=read.csv(file.choose("mitotimer"),header=TRUE)
  model<-lmer(value~signal*treatment+(1|replicate),REML=FALSE,na.action= na.omit,data=mitotimer)
  summary(model)
  car::Anova(model)
5. The script generates a generalized linear mixed model that compares treatment and MitoTimer signal values. It also uses the three independent measurement replicates per sample as a random effect. The analysis is followed by an ANOVA test that provides test statistics and the p-values.
Figure 2. Data analysis and representation of mitochondrial turnover results. A. Example confocal microscopy image of green and red MitoTimer signal in the optic lobes of third instar larval brains. Control and spinosad exposed individuals are represented. B. Example of how to organize a .csv table containing the MitoTimer fluorescence signals measured using ImageJ. This table is used to perform the statistical analysis in R. C. Normalized mean fluorescence intensity of MitoTimer signals in optic lobes (n = 20 larvae/treatment; three image sections/larva). Generalized linear mixed model followed by ANOVA. (***) p-value <0.001.
Data normalization and plotting
1. To facilitate data visualization and plotting, represent the data in terms of normalized mean fluorescence intensity.
2. Independently for green and red MitoTimer signals, calculate the mean fluoresce intensity of control samples. For a given control group, divide each individual sample value by the group mean and then multiple it by 100. Using that same group mean value, perform the same operation for the respective treated samples. At the end, control groups should have mean green and red signal values of 100, and the treated samples should have values normalized to their respective controls.
3. Using a suitable program (e.g., R) graphically represent the data in the form of bar plots or box plots, ideally including dot plots (Figure 2C).

Notes

Spinosad was used here to test its capacity to generate oxidative stress. However, this protocol can be applied to the same end for testing the effect of other drugs, rearing conditions, and/or mutations on mitochondrial turnover.
Given the exposure method used here, early third instar larvae are preferable, as late wandering third instars may crawl out of the wells or start pupation during exposure, and first or second instars would offer extra challenges for dissection, given their smaller size. However, other life stages may be selected depending on the research focus, including pupal and adult stages.
Image acquisition under the confocal microscope should be performed immediately after microscopy slides are ready, to guarantee sample integrity and the reliability of the signal acquired.
If performing insecticide/drug exposures for a fixed time with large sample groups, do not start exposure of all groups at the same time. Instead, start the exposure of each group at a different time, creating regular intervals of 10 or 15 min between them (or as required). That will allow time for sample dissection and microscopy of smaller sample batches, while ensuring that all individuals are exposed to roughly the same time.
Handle formaldehyde inside the fume hood, as it is toxic, and dispose of its waste as required.
More than one sample can be transferred to the same microtube with 4% formaldehyde solution. However, if doing so, transfer all these samples from the dissection slide into the microtube at once, to ensure that they are all fixed for the same amount of time.
Here, an elav-Gal4 driver was used to drive MitoTimer expression in neurons. Other Gal4 drivers can be used to different ends. Drosophila Gal4 driver strains can be found on the website of Bloomington Drosophila Stock Center (https://bdsc.indiana.edu/).

Recipes

Fly media (1 L)
Water 800 mL
Maize meal 60 g
Dried active yeast 15 g
Agar 6 g
Acid mix 7.5 mL
Tegosept 5 mL
Set the stove heat to high and in a pot add water and agar. Simmer and bring it to boil. After boiling the mixture for a couple of minutes, reduce the heat to medium, and add maize meal and dried active yeast. Simmer the mixture until homogenous, and turn off the heat. Wait a couple of minutes before adding acid mix and Tegosept while simmering. Dispense around 10 mL of fly media per vial, and wait for 12 h for it to set before closing the vials with cotton. Vials with fly media can be kept in a 4 °C fridge for a month but must be brought to room temperature before usage.
Apple juice-agar plates (1 L)
Water 720 mL
Agar 20 g
Apple juice 200 mL
Brewer’s yeast 7 g
Glucose 52 g
Sucrose 26 g
Tegosept 6 mL
Set the stove heat to high and, in a pot, add water and agar. Simmer and bring it to boil. After boiling the mixture for a couple of minutes, reduce the heat to medium and add apple juice, brewer’s yeast, glucose, and sucrose. Simmer the mixture until homogeneous, and turn off the heat. Wait a couple of minutes before adding Tegosept, and do it while simmering. Dispense enough media to fully cover the petri dish. Wait for it to solidify, and keep it upside down in the fridge at 4 °C; it must be brought to room temperature before usage.
Tegosept (250 mL)
Distilled water 137 mL
Propionic acid 103 mL
Orthophosphoric acid 10 mL
Add distilled water, propionic acid, and orthophosphoric acid into a 500 mL glass bottle. The solution can be kept at room temperature.
Acid mix (250 mL)
Ethanol 95% 250 mL
p-Hydroxybenzoic acid methyl ester 25 g
Add ethanol, distilled water, and p-Hydroxybenzoic acid methyl ester into a 500 mL glass bottle. The solution can be kept at room temperature.
20% Sucrose solution (200 mL)
Distilled water 160 mL
Analytical standard sucrose 40 g
Add distilled water and sucrose into a 500 mL glass bottle and give it a swirl until completely dissolved. It can be kept in the 4 °C fridge for a month, but must be brought to room temperature before usage.
5% Sucrose solution (200 mL)
Distilled water 190 mL
Analytical standard sucrose 10 g
Add distilled water and sucrose into a 500 mL glass bottle and give it a swirl until completely dissolved. It can be kept in the 4 °C fridge for a month but must be brought to room temperature before usage.
1,000 ppm spinosad solution in DMSO
DMSO 1 mL
Spinosad 1 mg
Add DMSO and Spinosad into a 1.7 mL microtube and vortex vigorously for a few minutes until solubilized. Store the insecticide dilution in a freezer (-20 °C) protected from light to prevent degradation. Create small volume aliquots (approximately 100 µL each) to avoid multiple freeze-thaw cycles.
4% Formaldehyde solution
Distilled water 375 µL
16% Formaldehyde solution 125 µL
Add distilled water and 16% formaldehyde to a 1.7 mL microtube.

Acknowledgments

The author was supported by a Victorian Latin America Doctoral Scholarship (Victorian Government and the University of Melbourne), an Alfred Nicholas Fellowship (University of Melbourne), and a University of Melbourne Faculty of Science Travelling Scholarship.

Original research paper: Martelli, F., Hernandes, N. H., Zuo, Z., Wang, J., Wong, C. O., Karagas, N. E., Roessner, U., Rupasinghe, T., Robin, C., Venkatachalam, K., et al. (2022). Low doses of the organic insecticide spinosad trigger lysosomal defects, elevated ROS, lipid dysregulation, and neurodegeneration in flies.Elife 11: e73812.

Competing interests

The author declares no conflicts of interest.

Ethics

Genetic modified Drosophila strain were maintained in a physical containment insectary facility following the biosafety guidelines by the Office of Research Ethics & Integrity at the University of Melbourne.

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