5-Ethynyl-2′-Deoxyuridine/Phospho-Histone H3 Dual-Labeling Protocol for Cell Cycle Progression Analysis in Drosophila Neural Stem Cells

In this protocol, we not only provide a detailed explanation of each step of EDU incorporation and immunostaining, but also explain image acquisition, image processing, and provide some tips for analyzing large datasets. This technique allows for the allocation of cells in G1, S, and M phases. This will be particularly useful for cell cycle analysis on studies on non mutations affecting mitotic genes.

We would expect people to struggle for two to three months if they do not have prior knowledge or experience with this technique. To begin, add Schneider’s dissection medium or SM to two consecutive depressions of a gloss spot plate and add PBS to the subsequent depression. Using a pair of forceps, pick wandering third instar larvae and place them on a tissue ratted with PBS to clean off fly food residue.

Place the larva in the depression with PBS followed by the consecutive depression containing SM.Next under a dissection microscope, use fine forceps to remove three quarters of the lower body of the larva. Then gently grab the larval mouth hooks with one pair of forceps and hold the cuticle of the other end with the other pair of forceps. Turn the larval head inside out by pushing the mouth hooks inward and simultaneously peeling off the tissue at the other end.

Observe the larval brain with attached imaginal discs. Remove other tissues attached to the brain and transfer the brain to the subsequent consecutive depression with SM.Add 100 microliters of 100 micromolar EDU SM solution to another depression on the Pyrex plate and incubate approximately 5 to 10 dissected brains in this solution for 2 hours at 25 degrees Celsius. After fixation and immunostaining remove PBST and add the EDU detection cocktail to the dissected brains.

Then incubate them at room temperature in the dark on a neutater. After 30 minutes, wash the samples twice with PBST for 10 minutes per wash in the dark. For DNA staining, prepare 5 micrograms per milliliter Hoechst 33342 solution.

Remove PBST after the second wash and add 500 microliters of the Hoechst solution to the dissected brains, then incubate them in the dark. After 10 minutes, remove the Hoechst and wash the samples with PBST for 10 minutes. Tap the tube onto the lab bench and let the brains settle down.

Remove the PBST from the tube, leaving behind 50 to 100 microliters. Cut the end of a 200 microliter micropipet tip and use it to transfer the brains onto a clean glass slide. Remove access PBST from the slide by blotting it with filter paper strips.

Do not let the filter paper touch the brains. Put a drop of a water soluble, non fluorescing mounting medium onto the brains and orient the brains such that the ventral nerve cord faces the slide and the lobes face upwards. Arrange the brains in a single file so that it is easier to image the brains serially.

Gently place a cover slip on top of the brain and incubate the slide overnight at 2 to 6 degrees Celsius. From the acquisition software select the 63 times objective. Put a drop of immersion oil on the cover slip just above the mounted brains to make it easier to locate the tissue through the eyepiece.

Using the dappy Hoechst 33342 channel, find the brain through the eyepiece and then switch to the acquisition mode in the software. Set up 4 channels to image Hoechst 33342, Alexa floor 4 8 8, Alexa floor 5 6, 8, and Alexa floor 6 4 7. Use the dye assistant tool to automatically set up the selected dyes excitation lasers and emission filters.

Set the field of view, such that it encompasses the entire brain lobe. Image the entire volume of the brain lobe by acquiring this tax spaced 0.8 micron apart. Store all images from an imaging session in a dot L I F library format.

For image analysis, open Fiji, then drag, and drop the LIF files into Fiji. Select data browser from the stack viewing tab and use virtual stack from the memory management tab in the bio formats plugin. Observe the multi-channel image displayed in image J.Change the color of the channels from the menu bar, using image, color, and channels tool.

And monitor the Miranda labeled neuroblasts, which appear as large round cells in the central brain region. Draw a region of interest using the ellipse tool over each neuroblast to avoid counting the neuroblast twice. From the image J menu bar, select analyze, tools, and ROI manager.

Mark all neuroblasts in the current Z section and press T after marking each cell. Once all neuroblasts in the current Z-section are marked, change the channels to pH3 and EDU and manually count the number of EDU positive neuroblasts, pH3 positive neuroblasts, EDU and pH3 positive neuroblasts, and neuroblasts not staining for either marker. Search for neuroblasts and subsequent Z sections, delete old ROIs and add new ROIs to count neuroblasts in different stacks.

Prepare a spreadsheet and calculate percentages of neuroblasts present in all four categories for each lobe. Prepare a bar graph using the spreadsheets software, showing the pools data for percentages of neuroblasts in each category. Central brain neuroblasts from third instar larval brains marked with Miranda, EDU, and pH3 are shown here.

Cells were allocated to G1/G0, S, SG2/M, and M phases. In EDU pH3 analysis, a significantly higher proportion of MMS19, loss of function neuroblasts was found in the M phase compared to wild type neuroblasts. MMS19:eGFP expression in the MMS19 loss of function background rescued the proportion of cells in M phase to wild type levels.

It is important not to use damaged brains for subsequent steps. And ED solution should be prepared freshly before starting dissections. This asset can be used as a quick initial screening to identify defects in specific cell cycle phases.

This could then be followed up by a more detailed analysis of a particular phase.