Materials and Reagents

1. Mini Petri dishes, 30 mm (Sigma-Aldrich, catalog number: SLW1480/12D)

2. Microsurgical scalpel (Stab Knife, 22.5 Degree, Straight; Sharpoint, catalog number: 72-2201)

3. Glass microcapillaries (50-μl calibrated pipets; Drummond, catalog number: 2-000-050)

4. Aspirator tube assemblies for calibrated microcapillary mouth pipette (Sigma-Aldrich, catalog number: A5177-5EA)

5. Syringe filter (0.2 μm; Pall Life Sciences, catalog number: PN 4612)

6. Epredia^TM PTFE Diagnostic Slides, 3-well (Thermo Fisher Scientific, catalog number: 10632391)

7. Epredia^TM PTFE Diagnostic Slides, 8-well (Thermo Fisher Scientific, catalog number: 10727951)

8. Animals: C57BL/6J female mice (3-5 weeks old)

9. Gonadotropin from pregnant mare serum; PMSG (Sigma-Aldrich, catalog number: G4877)

10. Chorionic gonadotropin human; hCG (Sigma-Aldrich, catalog number: CG10)

11. Fibronectin [Sigma-Aldrich, catalog number: F1141-2MG (solution; from bovine plasma) or 11051407001 (powder; from human plasma)]

12. M2 medium (Sigma-Aldrich, catalog number: M7167)

13. Mineral oil (Sigma-Aldrich, catalog number: M8410)

14. PBS (Sigma-Aldrich, catalog number: P4417)

Equipment

1. An animal room with a 12-hour light/dark cycle, red-light option, and a light timer that allows the researcher to shift the light schedule to be more compatible with normal working hours

2. Stereo microscope (Nikon, model: SMZ1270)

Procedure

1. Timed mating of female mice in the estrous stage

   1. For timed mating, hyper-stimulated (super-ovulated) or non-stimulated female mice can be used. The hyper-stimulation procedure increases the chance of successful timed mating.

      1. Hyper-stimulation: Hyper stimulate female mouse (22-24 days old optimally but up to 35 days old) with an intraperitoneal (i.p.) 5-IU injection of PMSG, followed by a 5-IU i.p. injection of hCG 48 h after PMSG. Calculate the time of both injections to start timed mating 12-14 h after the hCG injection. The protocol can be modified according to the animal stock provider’s suggestions (e.g., The Jackson Laboratory; https://www.jax.org/).

         Alternatively, the estrous cycle can be determined by examining washes of cell smears of the vagina to increase the success rate of the timed mating.

      2. Determining the stage of the estrous cycle in female mice is described in detail in Champlin et al. (1973) and Caligioni (2009).

   2. Introduce a male to a female mouse housed in the single cage during the last dark hour of the 12 h long dark cycle (30 min to 1 h). Work under red light when handling mice for mating to not disrupt the light/dark cycle.

   3. Check the presence of vaginal plugs as soon as the light cycle begins (set this time as 1 h post-coitus). Females with plugs can be used for the experiment at a desired time point (i.e., 3 h or 7 h post-coitus). Save females without plugs as they can be used for another run of timed mating.

   4. Hormone injection and mating time should be customized according to the availability of the animal room with flexible light/dark cycle adjustment and the experimenter’s schedule. The following example is based on the lack of such facility and flexible schedule of an experimenter: In a facility with a 12-h light/dark cycle with the light on at 7 am and off at 7 pm, PMSG can be injected on Day 1 at 6 pm, followed by 6 pm hCG injection on Day 3. At 6 am (12 h after hCG) on Day 4, start timed mating under red light without disrupting the light/dark cycle by introducing a male to females. When the light is on at 7 am, the females are allowed to mate for 2 h. In the next 30 min or so, check the plug and sacrifice the plugged females at 9:30-10:30 am for collecting in vivo capacitated sperm from 3-4 h post-coitus or 4:30-5:30 pm for collection in vivo capacitated sperm at 7-8 h post-coitus.




2. Microdissection and isolation of in vivo capacitated sperm

   1. After the sperm remain inside the female reproductive tract for a sufficient duration to allow in vivo capacitation, sacrifice the female mice with the confirmed vaginal plugs either by CO₂ asphyxiation or cervical dislocation.

   2. Under the well-lit stereo microscope, grip the fat pad around the ovary and stretch gently to clearly identify the membranous boundary between the ovary and the oviduct (line 1) and the oviduct and the uterus (line 2) (Figure 1).

   3. Carefully cut through the membrane around the proximal oviduct to slightly de-coil the oviduct first (line 2), followed by the membrane around the bursa and just outside infundibulum (line 1), and lastly, cut through the uterus just outside UTJ (line 3) (Figure 1).

      
      

      
      Figure 1. Scheme of mouse female reproductive tract dissection. Drawing slightly modified from Nagy (2003) in Manipulating the Mouse Embryo.

      
      

   4. The trimmed female tract (right) and the number and distribution of in vivo capacitated sperm (left) will look like Figure 2A.

   5. (Optional) If Acr-EGFP or Su9-DsRed/Acr-EGF mice are used, the distribution and number of sperm can be validated under the fluorescent microscope (Figure 2B).

      
      

      
      Figure 2. Distribution of sperm along the female reproductive tract. (A-B) Number of sperm recovered from each region of the female tract, aligned with an actual example image of a dissected female tract as described. Scale bar, 50 μm. (C) Validating location and number of Acr-EGF sperm in the isthmus with fluorescence microscopy.

   
   

   6. Place individual trimmed tract into a 0.3 ml-droplet of M2 media under mineral oil (Figure 3A) and further dissect them carefully with a microsurgical scalpel into ampullar, middle isthmus (2-3 mm), and UTJ parts (Figure 3B).

   7. Immediately place each part into an individual a 100-μl droplet of M2 media in a mini-Petri dish under mineral oil and gently press to release sperm (Figure 3B).

      
      

      
      Figure 3. Microdissection and aspiration of sperm from different regions of the female reproductive tract. (A) Oviduct dissected by microsurgical scalpel placed into a 0.3-ml M2 droplet. (B) Placing individual oviductal parts into a 0.1-ml M2 droplet. (C) Observation under stereo microscope and picking sperm by glass microcapillary. (D) A setup of mouth aspiration pipette with a glass capillary tube and a filter. (E) A schematic cartoon representation of visible components of the droplet under the microscope before and after additional transfer/washing.

   
   

   8. Observe the droplets (optimally by three persons) under (stereo) microscopes (e.g., for 10 min) and pick individual sperm with an aspirator connected to a filter in the middle and 50-ul glass microcapillary by mouth pipetting (Figure 3C and 3D).

   9. Depending on the speed of dissection and handling of the tissue, there will be other cell types, such as discharged epithelial cells and red blood cells (RBC), and debris together with sperm cells (Figure 3E). As sperm cells are smaller than these somatic cells (except fine debris), transferring the liquid to the 2^nd droplet by avoiding these contaminants can help to obtain a preparation more enriched with sperm cells.




3. Slide preparation

   1. Coat the surface of the slides with fibronectin. (A) Method A uses the fibronectin solution from bovine plasma: take 50-100 µl directly from the vial and spot onto the surface multiples times, making a domed solution with the desirable size. After 2-5 min when the edge of the dome is about to dry, rinse the coating solution with water. The slide/coverslip is ready to use for the next step. Method A does not require additional incubation time. (B) Method B prepares fresh fibronectin solution from fibronectin powder from human plasma. Reconstitute in sterile water by incubating for 30 min at 37°C to yield 1 mg/ml stock solution. Make small aliquots and keep frozen at -20°C until use. Prepare a fresh working solution (50 µg/ml in water) from an aliquot and apply 50 µl per well (3-well slide) or 10 µl per well (8-well slide). Incubate the slide/coverslip in a humid chamber for ~1 h. Rinse the coating solution with water and keep the slides in the chamber prior to sperm deposition. The time between the coating and sperm deposition should be as short as possible. Ideally, the whole procedure should be tested to optimize the cell attachment and reduce the background for the subsequent immunofluorescent microscopy due to the lot-to-lot variation of fibronectin.

   2. Place the picked-up sperm to 10 µl PBS droplet on the fibronectin-coated 8-well Teflon slides (Figure 4A: At a minimum, an average of 10 sperm per sample should be obtained). If too much tissue material is transferred by micropipette, a PBS washing step can be added (Figure 3E).

      
      

      
      Figure 4. Slide preparation for downstream experiments and example images. (A) Sperm transfer by glass micropipette into 8-well Teflon microdroplet slides. (B) An example of setting up a 3-well Teflon smeared slide.

   
   

   3. Subject the rest of the material to finer dissection, remove macroscopic pieces manually, mix the rest of the material with PBS (material/PBS 1:10), and smear it on the fibronectin-coated 3-well Teflon slides (Figure 4B: ~30-50 µl per well).

Data analysis

Obtained microscopic slides can be subjected to classical procedures for immunofluorescent staining, followed by confocal or super-resolution imaging. Shown here is an example of raw data (Figure 4C) used for P-Tyr quantification shown in Ded et al. (2020) to illustrate a downstream experimental outcome.

Notes

Mice hyperstimulation and vaginal smear analysis procedures are complex general protocols, and their detailed description can be found in provided references/links.

Acknowledgments

This protocol details an experimental method reported in the original research paper published in eLife (Ded et al., 2020; DOI: 10.7554/eLife.62043). This work was supported by start-up funds from Yale University School of Medicine, a Yale Goodman-Gilman Scholar Award-2015, and NIH (R01HD096745) to J-JC; by the Czech Science Foundation (GACR No. GJ20-17403Y) to LD; and by project BIOCEV (CZ.1.05/1.1.00/02/0109) from the ERDF and the Institute of BiotechnologyRVO:86652036.

Competing interests

Authors declare no competing interests.

Ethics

This study was performed in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. All the mice were treated in accordance with guidelines approved by Yale (protocol number: 20079, valid 2015-current) Animal Care and Use Committees (IACUC).

References

1. Austin, C. R. (1951). Observations on the penetration of the sperm in the mammalian egg. Aust J Sci Res B 4(4): 581-596.
2. Caligioni, C. S. (2009). Assessing reproductive status/stages in mice. Curr Protoc Neurosci 48: A.4I.1-A.4I.8.
3. Chang, M. C. (1951). Fertilizing capacity of spermatozoa deposited into the fallopian tubes. Nature 168(4277): 697-698.
4. Chung, J. J., Miki, K., Kim, D., Shim, S. H., Shi, H. F., Hwang, J. Y., Cai, X., Iseri, Y., Zhuang, X. and Clapham, D. E. (2017). CatSperzeta regulates the structural continuity of sperm Ca²⁺ signaling domains and is required for normal fertility. Elife 6: e23082.
5. Champlin, A. K., Dorr, D. L. and Gates, A. H. (1973). Determining the stage of the estrous cycle in the mouse by the appearance of the vagina. Biol Reprod 8(4): 491-494.
6. Ded, L., Hwang, J. Y., Miki, K., Shi, H. F. and Chung, J. J. (2020). 3D in situ imaging of the female reproductive tract reveals molecular signatures of fertilizing spermatozoa in mice. Elife 9: e62043.
7. Li, H., Hung, P. H. and Suarez, S. S. (2015). Ejaculated mouse sperm enter cumulus-oocyte complexes more efficiently in vitro than epididymal sperm. PLoS One 10(5): e0127753.
8. Nagy, A., Gertsenstein, M., Vintersten, K. and Behringer, R. (2003). Manipulating the Mouse Embryo: A Laboratory Manual. (3rd edition). Cold Spring Harbor Laboratory Press (2003 re-edition; 1986 org. ed.) 764 pp.
9. Steptoe, P. C. and Edwards, R. G. (1976). Reimplantation of a human embryo with subsequent tubal pregnancy. Lancet 1(7965): 880-882.