Materials and Reagents

1. Worm pick (flattened platinum wire mounted on a glass handle). Platinum wire with a diameter of 0.2 mm (Agar Scientific, catalog number: AGE404-2 )
2. Parafilm
3. Falcon tubes (50 ml, Greiner Bio-One, Germany)
4. Microscope slides (76 x 26 mm x 1 mm, Engelbrecht, catalog number: 11102 )
5. Empty plate or small container
6. Filter paper Whatman Grade 589/2 (diameter: 90 mm, Merck, catalog number: WHA10300109 ). The filter paper was cut radially into 12 pieces
7. Polystyrene beads (Polybead^® Microspheres 0.10 μm; Polysciences, catalog number: 00876-15 )
8. High-precision cover slips (18 mm x 18 mm x 0.17 µm, Carl Roth, catalog number: LH22.1 )
9. Candle wax
10. Glass Pasteur pipettes
11. C. elegans strain N2 (reference strain; Brenner, 1974)
12. C. elegans strain TMR17 (genotype: unc-119(ed3) III; ddIs6[tbg-1::GFP + unc-119(+)]; ltIs37[pAA64; pie-1::mCherry::HIS-58 + unc-119(+)] IV; strain maintained in the Müller-Reichert lab)
13. E. coli strain OP50 (Brenner, 1974)
14. Agarose (Carl Roth, catalog number: 3810.2 )
15. Peptone (BD Biosciences, catalog number: 4197928 )
16. NaCl (Carl Roth, catalog number: 9265.1 )
17. Cholesterol (Carl Roth, catalog number: 8866.1 )
18. CaCl₂ (Carl Roth, catalog number: CN93.1 )
19. MgCl₂ (Carl Roth, catalog number: 2189.1 )
20. NaOCl (Carl Roth, catalog number: 9062.3 )
21. KH₂PO₄ (Carl Roth, catalog number: P749.2 )
22. Na₂HPO₄ (Carl Roth, catalog number: P030.1 )
23. MgSO₄ (Carl Roth, catalog number: P027.1 )
24. NGM-agar plates (see Recipes)
25. KPO₄ buffer (see Recipes)
26. Bleaching solution (see Recipes)
27. M9-buffer (see Recipes)

Equipment

1. Fine tweezers
2. Incubator set to 20 °C (Thermo Fisher, model: Heraeus BK 6160 )
3. Incubator set to 30 °C to perform a heat-shock to animals (IKA, model: KS 4000 i control )
4. Stereo microscope with transmitted light (Motic, model: SMZ-168 BL )
5. Microwave
6. Spoon/spatula
7. Spinning disc confocal microscope (Olympus, model: IX83 )
8. EMCCD camera (Andor, model: iXon Ultra 897 )
9. Spinning disk unit (Yokogawa, model: CSU-X1 )
10. 60x water immersion objective (Olympus, model: UPlanSApo 60x W , N.A. 1.2)
    

Software

1. Fiji software package (www.fiji.sc, Schindelin et al., 2012)
2. Arivis Vision4D (arivis AG, Munich)
3. Microsoft Excel
4. Python (https://www.python.org/)
   

Procedure

1. Culture of C. elegans males
   1. Obtain male worms by applying the heat-shock method (Sulston and Hodgkin, 1988).
   2. Place 10 young L4 hermaphrodites on a fresh NGM-agar plate containing E. coli OP50.
   3. Seal the plate with Parafilm.
   4. Expose the plate to 30 °C for 6 h.
   5. Remove the parafilm and place the plate in an incubator at 20 °C for 2-3 days.
   6. Check then regularly for the presence of males on the plate. There should be 5-30 male worms on the plate.
   7. Transfer as many males as possible to a fresh NGM-agar plate and place 5 L4 hermaphrodites on this plate. Aim to have for 4-5 times more male individuals than hermaphrodites per plate. The progeny of the hermaphrodites should produce ~50% males.
   8. Maintain a culture of C. elegans males by transferring ~30 males and 5 L4 hermaphrodites to a fresh plate every three days.
      
      
      
2. Synchronization of male worms
   1. Pipette 50 µl of bleaching solution on a fresh NGM-agar plate containing E. coli OP50.
   2. Add 10-20 adult hermaphrodites from a male culture plate into the bleaching solution.
   3. Incubate the plate for three days at 20 °C.
      
      
      
3. Preparation of agarose pads
   1. Prepare about 30 clean glass slides and place them next to each other on the bench (Figure 2A).
   2. Prepare 10% (w/v) agarose powder in M9 buffer in a 50 ml Falcon tube (e.g., 0.5 g in 5 ml).
   3. Melt the agarose in a microwave with short heating intervals. Open the lid shortly in between the intervals.
      Caution: Melted agarose is hot; protective gloves should be worn.
   4. Use a small spoon or spatula to place a drop of the melted 10% agarose on a glass slide (Figure 2B).
   5. Immediately add a second glass slide on top and gently apply pressure for 5 s (Figure 2C).
   6. Repeat this procedure until all glass slides have agarose in between.
   7. Remove the top slide.
   8. Use a round object (e.g., cap of a permanent marker) to cut out agarose pads of a defined diameter of 10 mm. For this, select areas that do not contain any or only a few trapped air bubbles. Such bubbles might impair worm immobilization or the image quality later. Multiple agarose pads can be cut out of a single agar preparation (Figure 2D).
   9. Grab each agarose pad with fine tweezers and put it to a small container (e.g., empty plate) filled with M9 buffer (Figure 2E).
      
      
      Figure 2. Immobilization of worms for live-imaging using agarose pads and Polystyrene beads. A. Arrangement of glass slides on the laboratory bench. This way, agarose pads can be produced quickly before the agarose starts to solidify. B. Placing of melted agarose on one glass slide using a small spoon. C. Placing of a second glass slide on top of the agarose droplet and simultaneously application of pressure until the agarose is solid (~5 s). D. Excision of an agarose pad (diameter of about 10 mm) using a cap of a permanent marker. E. Transfer of the agarose pad with a fine tweezer to M9 buffer until further use. F. Placing of the agarose pad on a glass slide to start a live-cell experiment and removal of excess of liquid from the side with a piece of filter paper. G. Application of a Polystyrene-bead solution (bead diameter 0.1 µm) to the top of the agarose pad. H. After addition of worms, a cover slip was placed on top of the agarose pad. Afterwards, a few microliters of M9 buffer were added to avoid dehydration of the animals. I. Sealing of the cover slip with candle wax and imaging of the slide using fluorescence microscopy.
      
      
   10. Store agarose pads for several days in M9 buffer at 4 °C until further use. However, best results for worm immobilization are achieved with freshly prepared agarose pads.
       
       
       
4. Live-imaging of meiotic spindles in male worms using fluorescent markers
   1. Select an agarose pad, place it with tweezers in the middle of a new glass slide. Be careful not to damage the surface of the agarose pad as this will reduce the immobilization effect on individual worms.
   2. Aspirate excess of liquid from the side of the agarose pad with a piece of filter paper (Figure 2F). Be careful not to touch the surface of the agarose pad as also cellulose fibers on the surface of the pad will reduce the immobilization of worms.
   3. Add 1 µl of a polystyrene bead solution on the agarose pad (Figure 2G).
   4. Place 5-10 three-day old male worms within the bead solution.
   5. Gently add a cover slip (Figure 2H). It is important to lower the covers slip gently as a dropping of the glass might damage the worms or activate spermatids within the males. A sliding of the mounted cover glass should also be avoided as this might impair the worms as well. If worms are still moving after the assembly procedure, discard the sample and start again with point no. 1.
   6. Add a few microliters of M9 buffer from the side under the cover slip. Be careful not to add too much as excess liquid will make the cover slip float and decrease the amount of immobilization.
   7. Seal the edges of the cover slip with melted candle wax (Figure 2I).
   8. Check again under a dissection microscope if the worms are immobile. If worms are perfectly immobilized at this point, they will not become mobile again, thus long-term imaging will be possible by using the settings described below.
   9. Place the prepared sample on a fluorescence microscope.
   10. Select a fully immobilized worm for imaging. Start image acquisition. Samples can be imaged for about 90 min without any detectable effects on cell division. Typically, z-stacks are recorded on a confocal spinning disk microscope in two fluorescence channels with an image stack every 30 s containing about 60 planes (0.33 µm z-spacing). For the analysis of spindle dynamics, use a worm strain expressing γ-tubulin::GFP and histone H2B::mCherry in the germ line. With a 60x water immersion objective (N.A.: 1.2), we generated images with a voxel size of 0.222 x 0.222 x 0.33 µm. As the specifications among microscopes and lasers can vary, it is not possible here to give absolute values for the laser power to be used. It is advisable to assess the degree of photobleaching after one hour by comparing the signal intensities in the recorded images at the beginning and at the end of data acquisition. If the signal is bleached very fast or cells stop dividing, reduce the laser power or the exposure time per frame. Then check, if the signal/noise ratio in a new recording allows to detect specific signals. This process needs to be repeated iteratively until the optimal imaging settings for the specific microscope have been determined.
       
       
       
5. Image pre-processing
   1. Import the Z-stacks to ImageJ/Fiji. A user manual for ImageJ/Fiji can be found here.
   2. Convert them to a hyperstack.
   3. Correct for bleaching using the method “Exponential fit”.
   4. Save images.
      
      
      
6. Image processing
   1. Import images into arivis Vision4D (Figure 3A).
   2. Crop out and export individual meiotic cells from the meiotic region of the male gonad (Figure 3B).
   3. Make a note in which frame anaphase starts.
   4. Use a 3D median filter with a kernel size of 1 pixel (Figure 3C) to reduce the noise.
   5. Segment individual centrosomes in each meiotic cell with the tool “Blob finder” (Figure 3D). The settings have to be chosen in accordance to the used imaging setup. As the image quality varied among the live recordings the segmentation settings had to be adjusted within a narrow range. Typically, we used the following settings for the “Blob Finder” tool: diameter: 1 µm, threshold: 10-20, split sensitivity: 85-95.
   6. Track the segments over time using the tool “Track segments”. Typically, we used the following settings: type: “Brownian motion”, max. distance: 2-4 µm.
   7. Edit and combine tracks in order to have one track connecting the individual centrosomes over time using the tool “Track editor”.
   8. Export a Microsoft Excel file containing the x, y and z coordinates of the geometric center of mass of each of the segmented centrosomes.
   9. Create a semicolon-delimited CSV-file for the first centrosome track, containing in each line the x, y, and z coordinate of the center of mass each time point.
   10. Create analogously a second CSV-file for the second centrosome track.
   11. Use a custom-made Python script to reorient (Watt, 1999) and resample the image data in order to create a kymograph along the spindle axis with a defined radius for each fluorescence channel (Figure 3F, https://cdn.elifesciences.org/articles/50988/elife-50988-code1-v4.py).
       1. Give the path to both CSV-files containing the coordinates of the center of the centrosomes (PATH).
       2. Indicate which fluorescence channel should be processed (CHANNEL).
       3. Indicate the frame of the data set that should be used as a start (START_FRAME; typically, this was ten time points before the frame of anaphase onset).
       4. Indicate the pixel size in the resampled output data set (LINE_PIXEL_SIZE; typically 0.1 µm was used).
       5. Indicate the radius around the spindle axis that is used to calculate the fluorescence pixel values within the kymograph (LINE_RADIUS; typically a ‘9’ was used, which means a radius of 0.9 µm around the spindle axis, given that the pixel size was set to 0.1 µm).
       6. Indicate the length of extrapolation along the spindle axis after the start and end coordinate (LINE_EXTRAPOLATE, typically 1.0 µm before and after the centrosome centers was added to each kymograph).
       7. Define an output filename for the kymograph CSV-file (OUTPUT_FILENAME).
   12. Calculate the Gaussian weighted sum of fluorescence (Burger and Burge, 2013) in each plane in 0.1 µm steps along the spindle axis with a radius of 0.9 µm (Figure 3G). This was repeated for all time points (Figure 3H) and the kymograph was saved as an image (Figure 4A) and a csv-file.
       
       
       Figure 3. Workflow for image processing using arivis Vision4D. A. Data after pre-processing showing a meiosis I spindle in dual color fluorescence within one image plane. A selected area (indicated by a yellow box) is exported to a new file. Scale bar, 5 µm. B. Exported spindle as indicated in (A). C. Exported data after application of a 3D median filter (kernel size: 1 pixel). This algorithm removes the noise in the image while preserving the edges of structures. D. Segmentation of spindle poles (GFP-channel) using the “Blob finder” tool. The segmented volume is shown in yellow. Scale bar for B-D, 2 µm. E. 3D visualization of the exported image volume. The individual segmented spindle poles were traced over time as represented by lines on each pole. For 16 time points of the recording, white lines show the 3D position of the poles in the ‘past’ and gray lines the ‘future’ positions. F. Image showing a rotated and resampled spindle. Yellow dots indicate the center of mass of the segmented spindle poles. G. The volume that is used to assemble a kymograph is indicated by a box (green). Numbers in the coordinate system in (E-G) show distances in µm. H. Projections of the exported volumes at different time points that were used for the calculation of kymographs. Scale bar, 1 µm.
       
       
   13. Analyze the kymograph for each fluorescence channel to determine the position of the peak intensity from both sides of the spindle over time (Figures 4B-4C).
   14. Calculate the distance of the peak intensities for each channel over time. This way, the autosome-to-autosome distance (Figure 4D) and the pole-to-pole distance (Figure 4E) can be calculated. In addition, the pole-to-autosome distance (Figure 4F) can also be determined by calculating the distance of the peak intensities from both fluorescence channels on each side of the spindle.
       
       
       Figure 4. Kymographic analysis of spindle dynamics. A. Dual-color kymograph for the spindle as presented and processed in Figure 3. Autosomes are shown in magenta and spindle poles in cyan. Scale bar (white), 1 µm, time bar (orange), 2 min. B. Analysis of autosome dynamics. The kymograph of the histone::mCherry channel with the individual time points indicated by blue dashed lines is shown on the left, the fluorescence intensity profiles with highlighted the two peak intensities (red) arrows on the right. C. Analysis of spindle pole movements. The kymograph of the γ-tubulin::GFP channel (left) and the fluorescence intensity profiles are shown (right). D. Analysis of the autosome-to-autosome distance over time for 31 data sets aligned at anaphase onset (t₀). Circles represent the mean, shaded area the standard deviation. All mean values are connected by a solid line. E. Analysis of the pole-to-pole distance over time for 31 data sets. F. Analysis of the pole-to-autosome distance over time for 31 data sets.

Data analysis

For each spindle, the pole-to-pole, the autosome-to-autosome and the pole-to-autosome distance was calculated for each time point from the kymograph data. In addition, the time of anaphase onset was determined as time zero (t₀) by the first frame showing a clear separation of the autosomes. Accordingly, spindle dynamics in all data sets were then aligned to t₀. By calculating the average distance (± standard deviation) for each time point the dynamic properties of both meiotic spindles were calculated, i.e., the initial and final spindle length, the initial rate of elongation and the duration of spindle elongation or segregation.

Recipes

1. NGM-agar plates (Stiernagle, 2006)
   0.25% (w/v) peptone
   51 mM NaCl
   25 mM KPO₄ buffer
   0.005 g/L cholesterol
   1 mM CaCl₂
   1 mM MgCl₂
   2% (w/v) agar
   The solution was autoclaved for 15 min at 120 °C, afterwards plates were poured
2. KPO₄ buffer (1 M)
   0.8 M KH₂PO₄ and 0.8 M K₂HPO₄ in double-distilled water
   Adjust the pH to 6.0
   Filter sterilize the solution by using a vacuum filtration system (0.22 µm filter mesh size)
3. Bleaching solution (Stiernagle, 2006)
   500 mM NaOH
   2.6% (v/v) NaOCl in double-distilled water
   Protect solution from light and renew solution every 3-6 month as NaOCl gets less reactive over time
4. M9-buffer (Brenner, 1974)
   22 mM KH₂PO₄
   42.3 mM Na₂HPO₄
   85.6 mM NaCl
   1 mM MgSO₄
   The solution was filtered afterwards (filter pore size 0.2 μm)
   

Acknowledgments

The authors would like to thank the members of the Core Facility Cellular Imaging of the Faculty of Medicine “Carl Gustav Carus” (TU Dresden, Germany), especially Silke Tulok and Dr. Anja Nobst, and the light microscopy facility at the MPI-CBG (Dresden, Germany) for technical assistance. We also would like to thank Drs. Leocadia Paliulis and Robert Haase for the critical reading of this manuscript. Some strains were provided by the CGC, which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440). Research in the Müller-Reichert laboratory is supported by the Deutsche Forschungsgemeinschaft (MU 1423/10–1 to TMR).
  This protocol was part of and derived from Fabig et al. (2020).

Competing interests

FL and CG are employed by arivis AG. This company develops commercial image processing software (Vision4D). This software was utilized to process fluorescence microscopy data used in this protocol.

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