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Isolation and Manipulation of Mouse Gametes and Embryos
C H A P T E R
F I V E
Isolation and Manipulation of Mouse Gametes and Embryos Eveline S. Litscher and Paul M. Wassarman Contents 1. Introduction 2. Materials, Media, and Solutions 3. Collection and Storage Conditions 3.1. Isolation of growing and fully-grown oocytes 3.2. Isolation of ovulated eggs 3.3. Isolation of two-cell embryos 3.4. Isolation and capacitation of sperm 3.5. Storage of oocytes, ovulated eggs, and two-cell embryos 4. Sperm-Egg Interactions 4.1. Incubation of sperm with fully-grown oocytes, ovulated eggs, and two-cell embryos 4.2. Analysis of sperm binding to fully-grown oocytes and ovulated eggs 5. Analysis of the Status of the Sperm’s Acrosome 5.1. Coomassie-staining of sperm 5.2. Lectin–FITC staining of sperm 6. Visualization of Gold-Labeled Antibody on Sperm Acknowledgments References
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Abstract Many experimental approaches to answer questions about mammalian development begin with the isolation of mouse gametes and/or embryos. Here, methods used to isolate and store growing and fully-grown oocytes, ovulated (unfertilized) eggs, cleavage-stage embryos, and sperm from mice are described. Procedures used to carry out capacitation of sperm, binding of sperm to oocytes and ovulated eggs, and induction and assessment of the acrosome reaction in vitro are also described. Department of Developmental and Regenerative Biology, Mount Sinai School of Medicine, New York, New York, USA Methods in Enzymology, Volume 476 ISSN 0076-6879, DOI: 10.1016/S0076-6879(10)76005-5
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2010 Elsevier Inc. All rights reserved.
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1. Introduction In mammals, oogenesis and spermatogenesis begin during early fetal development with formation of primordial germ cells from a small number of stem cells at an extragonadal site (Austin and Short, 1982). Primordial germ cells, in turn, become either oogonia in females or spermatogonia in males; cells that proliferate mitotically. Oogonia and spermatogonia then enter meiosis and become oocytes and spermatocytes, respectively. During meiosis crossing-over and recombination occur, and at the end of meiosis haploid gametes, eggs and sperm are produced. Here, methods currently used to obtain and store growing and fullygrown oocytes, ovulated (unfertilized) eggs, cleavage-stage embryos, and sperm from mice are described.1 This methodology should be useful to investigators interested in studying various aspects of oogenesis, spermatogenesis, fertilization, and embryogenesis in mice.
2. Materials, Media, and Solutions The following alphabetical list includes materials, media, and solutions used to obtain, store, and manipulate mouse gametes and cleavage-stage embryos: A23187 (Ca2þ ionophore): 40 mM stock solution in dimethylsulfoxide (DMSO), diluted 1:40 with PBS, pH 7.2, and mixed with M199-M to a concentration of 20 mM. Capacitated sperm are pipetted into this solution at a 1:1 dilution (10 mM final concentration). Used to induce sperm to undergo the acrosome reaction. Ammonium acetate: 0.1 M, pH 9.0. Used to assess the status of the sperm’s acrosome. Antibody–gold conjugate: Antibody of choice conjugated to 5 nm gold particles, colloidal suspension (Sigma). Used to label the surface of sperm. Blocking buffer: PBS/PVP with 0.05% Tween 20 and 5% goat serum. Used in gold-labeling of sperm. CD-1 mice: Randomly bred Swiss mice (Charles River). EGTA: 0.5 M ethyleneglycoltetraacetic acid (EGTA), pH 7.4 (stock solution), diluted 125 to 4 mM EGTA in M199-M. Used to prevent sperm from undergoing the acrosome reaction.
1
Three particularly relevant sources for the methodology described here are Rafferty (1970), Nagy et al. (2003), and Wassarman and DePamphilis (2003).
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Coomassie stain: 0.04% Coomassie brilliant-blue G-250 (BioRad), dissolved in 3.5% perchloric acid. Used to stain the sperm’s acrosome. Gelatin-coated glass slides: 0.3% gelatin (autoclaved), 0.01% chromium potassium sulfate, sterile filtered. Slides are dipped for several seconds and airdried. Used to assess the status of the sperm’s acrosome. Human chorionic gonadotropin (hCG; Sigma): 5 International Units (IU)/ 100 ml/female mouse (see manufacturer’s instructions for reconstitution). Used to obtain ovulated eggs. Hyaluronidase: 1 mg/ml, dissolved in M199-M and prewarmed at 37 C (from bovine testes, type VI-S; Sigma). Used to remove cumulus cells from ovulated eggs. Light mineral oil: 225 ml light mineral oil is shaken vigorously with 25 ml M199, autoclaved, allowed to separate into aqueous and oil phases at room temperature (RT), and stored in the incubator at 37 C at least 2 days prior to use. Used to culture mouse gametes and embryos. Medium 199 (M199): Containing Earle’s salts, L-glutamine, 2.2 g/l sodium bicarbonate, 25 mM HEPES buffer. Medium 199-M (M199-M): M199 supplemented with 4 mg/ml bovine serum albumin (BSA; Sigma, fraction V, A-4503) and 30 mg/ml sodium pyruvate. Used to culture mouse gametes and embryos. Mounting medium: PBS–30% glycerol (for Coomassie stain), SlowFade Antifade Kit (for PNA–FITC; Molecular Probes). Used to examine the status of the sperm’s acrosome. Pasteur pipettes: Borosilicate glass pipettes, 9 in. length, drawn to an internal diameter of 150 mm (medium-bore), 150–200 mm (wide-bore), or not drawn (large-bore). Phosphate buffered saline (PBS): 30 mM sodium phosphate, pH 7.2, 150 mM NaCl. PBS–formaldehyde: PBS with 5% paraformaldehyde. Used as a fixative for sperm. PBS/PVP: PBS containing polyvinylpyrrolidone-40 (PVP-40), 4 mg/ml, 0.02% sodium azide. Used for storage of oocytes, eggs, and embryos. PBS/PVP–formaldehyde: PBS/PVP with 1% (a fixative for oocytes, eggs, and embryos) or 2% (a fixative for sperm bound to oocytes, eggs, and embryos) paraformaldehyde. PBS/PVP–glutaraldehyde: PBS/PVP with 2.5% glutaraldehyde. A fixative used for gold-labeling of sperm. Peanut agglutinin (PNA)–FITC (Arachis hypogaea) (Sigma): 15 mg/ml in PBS. Used to stain the sperm’s acrosome. Pregnant mare’s serum (PMS; Sigma): 5 IU/100 ml/female mouse (see manufacturer’s instructions for reconstitution). Used to obtain ovulated eggs. Silver Enhancer Kit: Enlarges colloidal gold particles for signals visible by light microscopy. Contains silver enhancement solutions A, B, and sodium thiosulfate pentahydrate (SE-100, Sigma). Used to visualize gold-labeled antibodies.
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Sterile steel needles: PrecisionGlide Needles 18G 1½ in. and 30G ½ in. (Becton Dickinson) connected to 1 ml syringes. Sterile syringe filter: 0.2 mm, connected by flexible tubing and a mouthpiece. Storage buffer: M199 containing 50 mM Tris–HCl, pH 7.5, 4 mg/ml PVP40, 0.02% sodium azide. Used to store oocytes, eggs, and embryos.
3. Collection and Storage Conditions Collection of mouse gametes and their culture in vitro are carried out in sterile-filtered medium M199-M at 37 C in a humidified atmosphere of 5% CO2 in air, unless otherwise indicated. M199-M, stored at 4 C, should be used within 2 weeks of preparation. Light mineral oil saturated with M199 is used to cover drops of M199-M in sterile tissue culture dishes (60 mm diameter) and the dishes are incubated at least 30 min at 37 C prior to addition of gametes and embryos. For fixation and storage of oocytes, eggs, and embryos, drops of PBS/PVP (with or without formaldehyde) and storage buffer are covered with light mineral oil at RT. Oocytes, eggs, and embryos are transferred from one droplet to another under a dissecting microscope using a mouth-operated, medium-bore Pasteur pipette (drawn to an internal diameter of 150 mm). The pipette is connected by flexible tubing via a sterile syringe filter (0.2 mm) to a mouthpiece. It is best to prefill a medium-bore Pasteur pipette with a small amount (a few microliters) of oil and an equal small amount of M199-M (as a visible oil-M199-M border line) before taking up small batches of cells (e.g., 5–10), and carefully monitoring the uptake as well as the downloading of cells. Capacitated sperm are transferred into drops of medium by Pipetman and microtips (e.g., 10 ml tips). Samples are always transferred through the oil layer covering the drops of M199-M.
3.1. Isolation of growing and fully-grown oocytes To obtain growing oocytes, ovaries are excised from juvenile (< 3 weeks old) female mice (Qi et al., 2002; Sorensen and Wassarman, 1976). Juvenile mice possess an unusually large number of growing oocytes arrested in the Dictyate stage of first meiotic prophase (Wassarman, 1996). Fully-grown oocytes are excised from adult (>3 weeks old) females. The ovaries are placed in a tissue culture dish containing 5 ml of prewarmed (37 C) M199-M and any fat and connective tissue are removed manually by using scissors and tweezers. Ovaries are poked extensively with fine steel needles (30G ½ in.) under a dissecting microscope. Released, healthy oocytes are visible as shiny and round (sometimes oval) cells amid cellular debris and torn ovaries. Oocytes are washed by carefully transferring them through
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three 100 ml drops of M199-M. On average, 10–15 growing or fully-grown oocytes can be recovered from a single ovary (CD-1 females). Typically, ovaries excised from 13- to 16-day-old juvenile female mice harbor a relatively large number of growing oocytes that are 60 10 mm in diameter. For fully-grown oocytes (diameter 80 mm), it is best to use ovaries from young adult females (3–4 weeks old) as the yield is similar to that of growing oocytes from juvenile mice and better than that from adult mice 5 weeks of age and older (Table 5.1; Fig. 5.1A). Table 5.1 Relationship between age of juvenile female mice and diameter of their oocytes Age of female (days)
Diameter of oocytes (mm)
9 12 15 18 21
50 55 60 70 80
A
B
C
D
Figure 5.1 Light micrograph of (A) a mouse oocyte (bar 20 mm), (B) an ovulated egg, (C) mouse sperm bound by their heads to the ZP of an ovulated mouse egg (bar 20 mm) and (D) a two-cell embryo. Samples are fixed in 1% formaldehyde.
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3.2. Isolation of ovulated eggs Fully-grown oocytes undergo meiotic maturation, with emission of the first polar body, to become ovulated eggs (Wassarman, 1996). To obtain ovulated eggs arrested in metaphase II from superovulated adult female mice, 5 IU of PMS is injected intraperitoneally, preferably in the evening of day-1. This is followed 46–48 h later by the injection of 5 IU of hCG (day-3). In the morning of day-4 (16–18 h after injection of hCG), oviducts are excised and placed in a tissue culture dish containing 5 ml of prewarmed (37 C) M199M. The upper portion of the excised oviduct (i.e., the ampulla where ovulated eggs are located) is gently torn apart by using two pairs of tweezers (watchmaker’s forceps, # 5) under a dissecting microscope. This results in the release of ovulated eggs surrounded by cumulus cells; so-called cumulus masses. Cumulus masses are collected by using a mouth-operated, wide-bore (not drawn) glass Pasteur pipette and are treated with hyaluronidase (1 mg/ ml) in a 200 ml drop for 3–5 min at RT. Generally, such treatment results in the cumulus cells detaching and falling off the ovulated eggs, but sometimes gentle up-and-down pipetting may be required to free eggs from surrounding cumulus cells. Ovulated eggs are then transferred and washed through three 100 ml drops of M199-M. On average, 200–300 ovulated eggs can be recovered from 10 superovulated, 6–8-week-old female CD-1 mice ( 20–30 eggs/mouse) (Fig. 5.1B).
3.3. Isolation of two-cell embryos To obtain two-cell embryos from superovulated adult female mice, 5 IU of PMS is injected intraperitoneally, preferably in the afternoon of day-1, followed 46–48 h later by the injection of 5 IU of hCG (day-3). At the time of hCG injection, the female is caged overnight with a proven male breeder and checked the following morning (day-4) for a copulation plug (indicative of a successful mating, but not always visible or present). On the morning of day-5, oviducts are excised and placed in M199-M, as described above. Two methods can be used to remove embryos from excised oviducts. In one procedure, a 1-ml syringe equipped with a sterile steel needle (30G ½ in.) with a blunt (cutoff ) tip is inserted into the oviduct opening and embryos are gently flushed out with M199-M. In the other procedure, oviducts are torn apart at several points along their length with watchmaker’s forceps and embryos are released into the medium. Both procedures are done under a dissecting microscope. Isolated two-cell embryos are washed through three 100 ml drops of M199-M. Fifteen to 30 two-cell embryos can be obtained from one superovulated, 6–8-week-old female CD-1 mouse (Fig. 5.1D).
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3.4. Isolation and capacitation of sperm The cauda (tail) epididymis and the vas deferens are excised from a sexually mature (3 months old) male mouse and placed into a tissue culture dish containing 5 ml prewarmed M199-M/EGTA (EGTA is present to prevent sperm from undergoing the acrosome reaction). Any fat tissue is carefully removed with scissors and tweezers. Sperm from the cauda epididymis are released into the medium by puncturing and gently squeezing the organ with a sterile steel needle (18G ½ in.). Sperm from the vas deferens are squeezed into the medium by using the same type of needle, but bent at a 90 angle. Sperm are collected from the culture dish with 1 ml pipette tips (Pipetman) into a 10 ml conical polypropylene tube and pelleted by low-speed centrifugation (2000 rpm) for 5 min at RT. The supernatant (<5 ml) is discarded and 5 ml of fresh prewarmed M199-M, without EGTA, is added carefully on top of the sperm pellet without stirring it up. Inspecting a small sample (e.g., 50 ml) of the throw-away supernatant under a dissecting microscope helps determine the quality of sperm. In some cases, sperm density may still be high in the supernatant (indicative of high-motility sperm) and a second centrifugation cycle in a new 10 ml polypropylene tube will give another sperm pellet (often of better quality; i.e., without cellular debris, red blood cells, etc.). Only capacitated sperm can fertilize ovulated eggs (Florman and Ducibella, 2006; Yanagimachi, 1994). Sperm are capacitated in M199-M for 1 h in the incubator at 37 C. During this time, live sperm will swim up from the pellet into the medium (‘‘swim-up’’ sperm), whereas dead sperm (no motility) will remain at the bottom of the tube. Sperm remain motile for 3–4 h after which time they become less lively and will eventually sink to the bottom of the tube. The final sperm concentration will vary depending upon the particular male mouse, but on average it is about 105– 106 sperm/ml (based on 5 ml of ‘‘swim-up’’ sperm from one CD-1 male). Sperm prepared as described above cannot be stored and should be prepared fresh each time.
3.5. Storage of oocytes, ovulated eggs, and two-cell embryos Isolated oocytes, eggs, and embryos, collected in M199-M and kept at 37 C in a humidified atmosphere of 5% CO2 in air, as described above, can be maintained in viable conditions for a day or two. For storage for up to 2 weeks, oocytes, eggs, and embryos in M199-M are transferred and washed through three 100 ml drops of PBS/PVP at RT prior to fixation for 1 h in a 100 ml drop of PBS/PVP–1% formaldehyde at RT. Following fixation, cells are washed through three 100 ml drops of PBS/PVP at RT, transferred to a 100 ml drop of storage buffer, and kept at 4 C.
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4. Sperm-Egg Interactions 4.1. Incubation of sperm with fully-grown oocytes, ovulated eggs, and two-cell embryos Ovulated eggs or fully-grown oocytes and two-cell embryos, either stored in M199-M and kept at 37 C for no more than 2 days, or kept in storage buffer at 4 C, are washed through three 100 ml drops of M199-M at RT and transferred to the incubator for at least 30 min before being added to capacitated sperm (Williams et al., 2006). Ten microliter drops of prewarmed M199-M are pipetted into a tissue culture dish and covered with oil. Alternatively, if a test substance is being evaluated, the sample is dissolved in a 10-m l drop of M199-M and pipetted into a tissue culture dish, as above. Then 10 ml of capacitated ‘‘swim-up’’ sperm are added to the drop (through the oil layer; 1:1 dilution) and incubated for 15 min at 37 C. Following this period, oocytes or eggs, and two-cell embryos are added to the 20 ml drop with as little medium as possible and incubated at 37 C for an additional 35–45 min. Sperm motility is monitored under a dissecting microscope during this period and samples with sperm motility of less than 70% (i.e., less than 7 motile sperm out of 10 sperm) should be discarded. At the end of the incubation time, oocytes or eggs, and embryos with associated sperm are removed from the drops with a mouth-operated, wide-bore Pasteur pipette and washed through three 100 ml drops of M199-M. This step involves careful up-anddown pipetting (3–4 times) of groups of 5–10 cells in each drop until no more than one to two sperm remain attached to the two-cell embryos (negative control). Under these conditions, sperm bound to the oocyte or egg zona pellucida (ZP) remain bound, whereas sperm nonspecifically attached to oocyte or egg and embryo ZP are removed. Oocytes and eggs with bound sperm, as well as embryos, are then transferred to 10 ml drops of M199-M containing PBS/PVP–2% formaldehyde (1:1 by volume) covered with oil.
4.2. Analysis of sperm binding to fully-grown oocytes and ovulated eggs The number of sperm bound per oocyte, egg, or embryo is determined by counting sperm tails in the largest diameter focal plane, using dark-field microscopy and 200 magnification. Typically, there are 12 oocytes or ovulated eggs and two two-cell embryos present per sample. The presence of embryos is essential in this assay since they serve as a negative control; sperm do not bind to the ZP of fertilized eggs or embryos. After determining the number of sperm bound to each oocyte or egg in one plane of focus, the highest and lowest values are discarded, and the remaining 10 values are used to calculate the average number of sperm bound per oocyte or
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egg ( S.D.). For example, when sperm are exposed to M199-M alone (no test substance), at sperm concentrations of about 5 105 sperm/ml, and incubated with oocytes or eggs and embryos, there are 30–50 sperm bound per oocyte or egg in the largest diameter focal plane (Fig. 5.1C).
5. Analysis of the Status of the Sperm’s Acrosome The acrosome is a large lysosome-like vesicle overlying the sperm’s nucleus (Florman and Ducibella, 2006; Yanagimachi, 1994). Capacitated mouse sperm undergo the acrosome reaction (AR) shortly after binding to the egg ZP. The AR is a specialized form of cellular exocytosis. It involves multiple fusions between plasma membrane overlying the anterior region of the sperm head and the outer acrosomal membrane lying just beneath the plasma membrane. Small hybrid membrane vesicles form and the inner acrosomal membrane, overlying the sperm’s nucleus, is exposed to the egg ZP (Cohen and Wassarman, 2001). Only acrosome-reacted sperm can penetrate the ZP, reach the plasma membrane, and fuse with eggs (i.e., fertilize eggs). To assess the status of the sperm’s acrosome (i.e., whether it is acrosomeintact or acrosome-reacted), capacitated ‘‘swim-up’’ sperm (see above) are incubated at 37 C for 1 h in M199-M, either in 20–50 ml drops under oil in a tissue culture dish or in 0.5-ml Eppendorf tubes, in the presence or absence of a test substance. Sperm exposed to Ca2þ ionophore A23187 are used as a positive control since it induces sperm to undergo the AR in vitro. Capacitated sperm are pipetted into a solution of 20 mM A23187 at a 1:1 dilution to a final concentration of 10 mM (it should be noted that sperm incubated in A23187 become completely immotile within a few minutes of exposure). After incubation, sperm samples are pelleted in 0.5-ml Eppendorf tubes by centrifugation at 10,000 rpm for 5 min in a microfuge at RT and fixed with PBS–5% formaldehyde (1:1 dilution with sperm sample) overnight at 4 C. The following day, sperm are washed by centrifugation (as above) in 0.1 M ammonium acetate, pH 9.0, resuspended in 50 ml of 0.1 M ammonium acetate by gentle vortexing, pipetted onto gelatin-coated glass slides, and air-dried. Several methods have been described to assess the status of the sperm’s acrosome. Two of these methods, Coomassie staining and FITC–lectin staining of sperm, are described in the following sections.
5.1. Coomassie-staining of sperm Slides are washed with distilled H2O, methanol, H2O, for 5 min each, Coomassie-stained for 5 min, and washed extensively with H2O (Larson and Miller, 1999). Sperm can be observed directly on the glass slide or under
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a coverslip (sealed with clear nail polish for long-term storage) using PBS– 30% glycerol as mounting medium, and are scanned in the presence or absence of an acrosome by bright-field microscopy at 400 magnification.
5.2. Lectin–FITC staining of sperm Slides are washed twice with PBS for 5 min, incubated with PNA–FITC (A. hypogaea) for 30 min at RT (humid chamber), and rinsed with PBS for 15 min (Lybaert et al., 2009). Samples are mounted with antifade mounting reagent and a coverslip (sealed with clear nail polish) and examined for the presence or absence of an acrosome by fluorescent microscopy at 400 magnification. Sperm retaining an intact acrosome (Fig. 5.2A) will display a continuous blue (Coomassie)- or fluorescent green (PNA–FITC)-stained ridge overlying the sperm head. Sperm that have undergone the AR (Fig. 5.2B) will display either a patchy blue or patchy fluorescent green pattern, indicative of a partial AR, or no staining at all, indicative of a completed AR. Typically, A23187-treated sperm samples will be 60–80% acrosome-reacted, whereas sperm exposed to M199-M alone will be 15–25% acrosomereacted (due to a so-called ‘‘spontaneous AR’’).
6. Visualization of Gold-Labeled Antibody on Sperm Capacitated sperm are incubated with a sperm-binding probe (i.e., antigen of choice), fixed, washed and air-dried on gelatin-coated slides as described above (Williams et al., 2006). Samples on slides are blocked with a A
B
Figure 5.2 Light micrographs of (A) acrosome-intact and (B) acrosome-reacted mouse sperm, fixed in 2.5% formaldehyde and stained with Coomassie brilliant-blue G-250. Arrow: acrosomal cap (bar 5.5 mm).
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50 ml drop of PBS/PVP–0.05% Tween 20 containing 5% goat serum (blocking buffer) for 30 min at RT in a humid chamber, then rinsed in H2O and air-dried for 15 min at RT. A diluted gold suspension (1:10) of antibody–gold conjugate (antibody of choice, coupled to 5 nm colloidal gold particles) in blocking buffer is pipetted onto the sperm sample (50 ml drop). The slide is incubated for 2 h at RT in a humid chamber on a slow moving shaker and washed afterward with PBS by submersion in a Petri dish for 5 min. The sample is fixed again with PBS/PVP–2.5% glutaraldehyde (50 ml drop) for 1 h at RT (humid chamber) and rinsed thoroughly with H2O in a Petri dish. To enlarge colloidal gold labels for high-contrast signals that are visible by light microscopy, a 50 ml drop of silver stain solution (Silver Enhancer Kit) is applied onto the gold-labeled sperm samples for 15 min at RT in the dark (humid chamber). The slide is dipped in H2O, fixed with 2.5% sodium thiosulfate pentahydrate (50 ml drop) for 3 min at RT, rinsed with H2O, mounted with PBS/PVP–30% glycerol and a coverslip, and sealed with clear nail polish. Individual sperm are scanned by light microscopy for silver-enhanced colloidal gold–protein complexes and the number of particles (silver precipitate) associated with each sperm is determined.
ACKNOWLEDGMENTS Our research was supported in part by the National Institutes of Health (NICHD), most recently by grant HD-35105.
REFERENCES Austin, C. R., and Short, R. V. (eds.), (1982). Reproduction in Mammals. 1. Germ Cells and Fertilization, Cambridge University Press, Cambridge. Cohen, N., and Wassarman, P. M. (2001). Association of egg zona pellucida glycoprotein mZP3 with sperm protein sp56 during fertilization in mice. Int. J. Dev. Biol. 45, 569–576. Florman, H. M., and Ducibella, T. (2006). Fertilization in mammals. In ‘‘The Physiology of Reproduction,’’ ( J. D. Neill, ed.), 3rd edn. pp. 55–112. Elsevier/Academic Press. Larson, J. L., and Miller, D. J. (1999). Simple histochemical stain for acrosomes on sperm from several species. Mol. Reprod. Dev. 52, 445–449. Lybaert, P., Danguy, A., Leleux, F., Meuris, S., and Lebrun, P. (2009). Improved methodology for the detection and quantification of the acrosome reaction in mouse spermatozoa. Histol. Histopathol. 24, 999–1007. Nagy, A., Gertsenstein, M., Vintersten, K., and Behringer, R. (2003). Manipulating the Mouse Embryo: A Laboratory Manual. 3rd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Qi, H., Williams, Z., and Wassarman, P. M. (2002). Secretion and assembly of zona pellucida glycoproteins by growing mouse oocytes microinjected with epitope-tagged cDNAs for mZP2 and mZP3. Mol. Biol. Cell 13, 530–541.
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Rafferty, K. A., Jr. (1970). Methods in Experimental Embryology of the Mouse. Johns Hopkins Press. Sorensen, R. A., and Wassarman, P. M. (1976). Relationship between growth and meiotic maturation of the mouse oocyte. Dev. Biol. 50, 531–536. Wassarman, P. M. (1996). Oogenesis. In ‘‘Reproductive Endrocrinology, Surgery, and Technology,’’ (E. Y. Adashi, J. A. Rock, and Z. Rosenwaks, eds.), Vol. 1, pp. 341–357. Lippincott-Raven Press. Wassarman, P. M., and DePamphilis, M. L. (eds.), (2003). In Guide to Techniques in Mouse Development, Methods in Enzymology Vol. 225. Academic Press. Williams, Z., Litscher, E. S., Jovine, L., and Wassarman, P. M. (2006). Polypeptide encoded by mouse ZP3 exon-7 is necessary and sufficient for binding of mouse sperm in vitro. J. Cell. Physiol. 207, 30–39. Yanagimachi, R. (1994). Mammalian fertilization. In ‘‘The Physiology of Reproduction 1,’’ (E. Knobil and J. D. Neill, eds.), pp. 189–317. Raven Press.
F I V E
Isolation and Manipulation of Mouse Gametes and Embryos Eveline S. Litscher and Paul M. Wassarman Contents 1. Introduction 2. Materials, Media, and Solutions 3. Collection and Storage Conditions 3.1. Isolation of growing and fully-grown oocytes 3.2. Isolation of ovulated eggs 3.3. Isolation of two-cell embryos 3.4. Isolation and capacitation of sperm 3.5. Storage of oocytes, ovulated eggs, and two-cell embryos 4. Sperm-Egg Interactions 4.1. Incubation of sperm with fully-grown oocytes, ovulated eggs, and two-cell embryos 4.2. Analysis of sperm binding to fully-grown oocytes and ovulated eggs 5. Analysis of the Status of the Sperm’s Acrosome 5.1. Coomassie-staining of sperm 5.2. Lectin–FITC staining of sperm 6. Visualization of Gold-Labeled Antibody on Sperm Acknowledgments References
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Abstract Many experimental approaches to answer questions about mammalian development begin with the isolation of mouse gametes and/or embryos. Here, methods used to isolate and store growing and fully-grown oocytes, ovulated (unfertilized) eggs, cleavage-stage embryos, and sperm from mice are described. Procedures used to carry out capacitation of sperm, binding of sperm to oocytes and ovulated eggs, and induction and assessment of the acrosome reaction in vitro are also described. Department of Developmental and Regenerative Biology, Mount Sinai School of Medicine, New York, New York, USA Methods in Enzymology, Volume 476 ISSN 0076-6879, DOI: 10.1016/S0076-6879(10)76005-5
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2010 Elsevier Inc. All rights reserved.
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1. Introduction In mammals, oogenesis and spermatogenesis begin during early fetal development with formation of primordial germ cells from a small number of stem cells at an extragonadal site (Austin and Short, 1982). Primordial germ cells, in turn, become either oogonia in females or spermatogonia in males; cells that proliferate mitotically. Oogonia and spermatogonia then enter meiosis and become oocytes and spermatocytes, respectively. During meiosis crossing-over and recombination occur, and at the end of meiosis haploid gametes, eggs and sperm are produced. Here, methods currently used to obtain and store growing and fullygrown oocytes, ovulated (unfertilized) eggs, cleavage-stage embryos, and sperm from mice are described.1 This methodology should be useful to investigators interested in studying various aspects of oogenesis, spermatogenesis, fertilization, and embryogenesis in mice.
2. Materials, Media, and Solutions The following alphabetical list includes materials, media, and solutions used to obtain, store, and manipulate mouse gametes and cleavage-stage embryos: A23187 (Ca2þ ionophore): 40 mM stock solution in dimethylsulfoxide (DMSO), diluted 1:40 with PBS, pH 7.2, and mixed with M199-M to a concentration of 20 mM. Capacitated sperm are pipetted into this solution at a 1:1 dilution (10 mM final concentration). Used to induce sperm to undergo the acrosome reaction. Ammonium acetate: 0.1 M, pH 9.0. Used to assess the status of the sperm’s acrosome. Antibody–gold conjugate: Antibody of choice conjugated to 5 nm gold particles, colloidal suspension (Sigma). Used to label the surface of sperm. Blocking buffer: PBS/PVP with 0.05% Tween 20 and 5% goat serum. Used in gold-labeling of sperm. CD-1 mice: Randomly bred Swiss mice (Charles River). EGTA: 0.5 M ethyleneglycoltetraacetic acid (EGTA), pH 7.4 (stock solution), diluted 125 to 4 mM EGTA in M199-M. Used to prevent sperm from undergoing the acrosome reaction.
1
Three particularly relevant sources for the methodology described here are Rafferty (1970), Nagy et al. (2003), and Wassarman and DePamphilis (2003).
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Coomassie stain: 0.04% Coomassie brilliant-blue G-250 (BioRad), dissolved in 3.5% perchloric acid. Used to stain the sperm’s acrosome. Gelatin-coated glass slides: 0.3% gelatin (autoclaved), 0.01% chromium potassium sulfate, sterile filtered. Slides are dipped for several seconds and airdried. Used to assess the status of the sperm’s acrosome. Human chorionic gonadotropin (hCG; Sigma): 5 International Units (IU)/ 100 ml/female mouse (see manufacturer’s instructions for reconstitution). Used to obtain ovulated eggs. Hyaluronidase: 1 mg/ml, dissolved in M199-M and prewarmed at 37 C (from bovine testes, type VI-S; Sigma). Used to remove cumulus cells from ovulated eggs. Light mineral oil: 225 ml light mineral oil is shaken vigorously with 25 ml M199, autoclaved, allowed to separate into aqueous and oil phases at room temperature (RT), and stored in the incubator at 37 C at least 2 days prior to use. Used to culture mouse gametes and embryos. Medium 199 (M199): Containing Earle’s salts, L-glutamine, 2.2 g/l sodium bicarbonate, 25 mM HEPES buffer. Medium 199-M (M199-M): M199 supplemented with 4 mg/ml bovine serum albumin (BSA; Sigma, fraction V, A-4503) and 30 mg/ml sodium pyruvate. Used to culture mouse gametes and embryos. Mounting medium: PBS–30% glycerol (for Coomassie stain), SlowFade Antifade Kit (for PNA–FITC; Molecular Probes). Used to examine the status of the sperm’s acrosome. Pasteur pipettes: Borosilicate glass pipettes, 9 in. length, drawn to an internal diameter of 150 mm (medium-bore), 150–200 mm (wide-bore), or not drawn (large-bore). Phosphate buffered saline (PBS): 30 mM sodium phosphate, pH 7.2, 150 mM NaCl. PBS–formaldehyde: PBS with 5% paraformaldehyde. Used as a fixative for sperm. PBS/PVP: PBS containing polyvinylpyrrolidone-40 (PVP-40), 4 mg/ml, 0.02% sodium azide. Used for storage of oocytes, eggs, and embryos. PBS/PVP–formaldehyde: PBS/PVP with 1% (a fixative for oocytes, eggs, and embryos) or 2% (a fixative for sperm bound to oocytes, eggs, and embryos) paraformaldehyde. PBS/PVP–glutaraldehyde: PBS/PVP with 2.5% glutaraldehyde. A fixative used for gold-labeling of sperm. Peanut agglutinin (PNA)–FITC (Arachis hypogaea) (Sigma): 15 mg/ml in PBS. Used to stain the sperm’s acrosome. Pregnant mare’s serum (PMS; Sigma): 5 IU/100 ml/female mouse (see manufacturer’s instructions for reconstitution). Used to obtain ovulated eggs. Silver Enhancer Kit: Enlarges colloidal gold particles for signals visible by light microscopy. Contains silver enhancement solutions A, B, and sodium thiosulfate pentahydrate (SE-100, Sigma). Used to visualize gold-labeled antibodies.
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Sterile steel needles: PrecisionGlide Needles 18G 1½ in. and 30G ½ in. (Becton Dickinson) connected to 1 ml syringes. Sterile syringe filter: 0.2 mm, connected by flexible tubing and a mouthpiece. Storage buffer: M199 containing 50 mM Tris–HCl, pH 7.5, 4 mg/ml PVP40, 0.02% sodium azide. Used to store oocytes, eggs, and embryos.
3. Collection and Storage Conditions Collection of mouse gametes and their culture in vitro are carried out in sterile-filtered medium M199-M at 37 C in a humidified atmosphere of 5% CO2 in air, unless otherwise indicated. M199-M, stored at 4 C, should be used within 2 weeks of preparation. Light mineral oil saturated with M199 is used to cover drops of M199-M in sterile tissue culture dishes (60 mm diameter) and the dishes are incubated at least 30 min at 37 C prior to addition of gametes and embryos. For fixation and storage of oocytes, eggs, and embryos, drops of PBS/PVP (with or without formaldehyde) and storage buffer are covered with light mineral oil at RT. Oocytes, eggs, and embryos are transferred from one droplet to another under a dissecting microscope using a mouth-operated, medium-bore Pasteur pipette (drawn to an internal diameter of 150 mm). The pipette is connected by flexible tubing via a sterile syringe filter (0.2 mm) to a mouthpiece. It is best to prefill a medium-bore Pasteur pipette with a small amount (a few microliters) of oil and an equal small amount of M199-M (as a visible oil-M199-M border line) before taking up small batches of cells (e.g., 5–10), and carefully monitoring the uptake as well as the downloading of cells. Capacitated sperm are transferred into drops of medium by Pipetman and microtips (e.g., 10 ml tips). Samples are always transferred through the oil layer covering the drops of M199-M.
3.1. Isolation of growing and fully-grown oocytes To obtain growing oocytes, ovaries are excised from juvenile (< 3 weeks old) female mice (Qi et al., 2002; Sorensen and Wassarman, 1976). Juvenile mice possess an unusually large number of growing oocytes arrested in the Dictyate stage of first meiotic prophase (Wassarman, 1996). Fully-grown oocytes are excised from adult (>3 weeks old) females. The ovaries are placed in a tissue culture dish containing 5 ml of prewarmed (37 C) M199-M and any fat and connective tissue are removed manually by using scissors and tweezers. Ovaries are poked extensively with fine steel needles (30G ½ in.) under a dissecting microscope. Released, healthy oocytes are visible as shiny and round (sometimes oval) cells amid cellular debris and torn ovaries. Oocytes are washed by carefully transferring them through
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three 100 ml drops of M199-M. On average, 10–15 growing or fully-grown oocytes can be recovered from a single ovary (CD-1 females). Typically, ovaries excised from 13- to 16-day-old juvenile female mice harbor a relatively large number of growing oocytes that are 60 10 mm in diameter. For fully-grown oocytes (diameter 80 mm), it is best to use ovaries from young adult females (3–4 weeks old) as the yield is similar to that of growing oocytes from juvenile mice and better than that from adult mice 5 weeks of age and older (Table 5.1; Fig. 5.1A). Table 5.1 Relationship between age of juvenile female mice and diameter of their oocytes Age of female (days)
Diameter of oocytes (mm)
9 12 15 18 21
50 55 60 70 80
A
B
C
D
Figure 5.1 Light micrograph of (A) a mouse oocyte (bar 20 mm), (B) an ovulated egg, (C) mouse sperm bound by their heads to the ZP of an ovulated mouse egg (bar 20 mm) and (D) a two-cell embryo. Samples are fixed in 1% formaldehyde.
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3.2. Isolation of ovulated eggs Fully-grown oocytes undergo meiotic maturation, with emission of the first polar body, to become ovulated eggs (Wassarman, 1996). To obtain ovulated eggs arrested in metaphase II from superovulated adult female mice, 5 IU of PMS is injected intraperitoneally, preferably in the evening of day-1. This is followed 46–48 h later by the injection of 5 IU of hCG (day-3). In the morning of day-4 (16–18 h after injection of hCG), oviducts are excised and placed in a tissue culture dish containing 5 ml of prewarmed (37 C) M199M. The upper portion of the excised oviduct (i.e., the ampulla where ovulated eggs are located) is gently torn apart by using two pairs of tweezers (watchmaker’s forceps, # 5) under a dissecting microscope. This results in the release of ovulated eggs surrounded by cumulus cells; so-called cumulus masses. Cumulus masses are collected by using a mouth-operated, wide-bore (not drawn) glass Pasteur pipette and are treated with hyaluronidase (1 mg/ ml) in a 200 ml drop for 3–5 min at RT. Generally, such treatment results in the cumulus cells detaching and falling off the ovulated eggs, but sometimes gentle up-and-down pipetting may be required to free eggs from surrounding cumulus cells. Ovulated eggs are then transferred and washed through three 100 ml drops of M199-M. On average, 200–300 ovulated eggs can be recovered from 10 superovulated, 6–8-week-old female CD-1 mice ( 20–30 eggs/mouse) (Fig. 5.1B).
3.3. Isolation of two-cell embryos To obtain two-cell embryos from superovulated adult female mice, 5 IU of PMS is injected intraperitoneally, preferably in the afternoon of day-1, followed 46–48 h later by the injection of 5 IU of hCG (day-3). At the time of hCG injection, the female is caged overnight with a proven male breeder and checked the following morning (day-4) for a copulation plug (indicative of a successful mating, but not always visible or present). On the morning of day-5, oviducts are excised and placed in M199-M, as described above. Two methods can be used to remove embryos from excised oviducts. In one procedure, a 1-ml syringe equipped with a sterile steel needle (30G ½ in.) with a blunt (cutoff ) tip is inserted into the oviduct opening and embryos are gently flushed out with M199-M. In the other procedure, oviducts are torn apart at several points along their length with watchmaker’s forceps and embryos are released into the medium. Both procedures are done under a dissecting microscope. Isolated two-cell embryos are washed through three 100 ml drops of M199-M. Fifteen to 30 two-cell embryos can be obtained from one superovulated, 6–8-week-old female CD-1 mouse (Fig. 5.1D).
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3.4. Isolation and capacitation of sperm The cauda (tail) epididymis and the vas deferens are excised from a sexually mature (3 months old) male mouse and placed into a tissue culture dish containing 5 ml prewarmed M199-M/EGTA (EGTA is present to prevent sperm from undergoing the acrosome reaction). Any fat tissue is carefully removed with scissors and tweezers. Sperm from the cauda epididymis are released into the medium by puncturing and gently squeezing the organ with a sterile steel needle (18G ½ in.). Sperm from the vas deferens are squeezed into the medium by using the same type of needle, but bent at a 90 angle. Sperm are collected from the culture dish with 1 ml pipette tips (Pipetman) into a 10 ml conical polypropylene tube and pelleted by low-speed centrifugation (2000 rpm) for 5 min at RT. The supernatant (<5 ml) is discarded and 5 ml of fresh prewarmed M199-M, without EGTA, is added carefully on top of the sperm pellet without stirring it up. Inspecting a small sample (e.g., 50 ml) of the throw-away supernatant under a dissecting microscope helps determine the quality of sperm. In some cases, sperm density may still be high in the supernatant (indicative of high-motility sperm) and a second centrifugation cycle in a new 10 ml polypropylene tube will give another sperm pellet (often of better quality; i.e., without cellular debris, red blood cells, etc.). Only capacitated sperm can fertilize ovulated eggs (Florman and Ducibella, 2006; Yanagimachi, 1994). Sperm are capacitated in M199-M for 1 h in the incubator at 37 C. During this time, live sperm will swim up from the pellet into the medium (‘‘swim-up’’ sperm), whereas dead sperm (no motility) will remain at the bottom of the tube. Sperm remain motile for 3–4 h after which time they become less lively and will eventually sink to the bottom of the tube. The final sperm concentration will vary depending upon the particular male mouse, but on average it is about 105– 106 sperm/ml (based on 5 ml of ‘‘swim-up’’ sperm from one CD-1 male). Sperm prepared as described above cannot be stored and should be prepared fresh each time.
3.5. Storage of oocytes, ovulated eggs, and two-cell embryos Isolated oocytes, eggs, and embryos, collected in M199-M and kept at 37 C in a humidified atmosphere of 5% CO2 in air, as described above, can be maintained in viable conditions for a day or two. For storage for up to 2 weeks, oocytes, eggs, and embryos in M199-M are transferred and washed through three 100 ml drops of PBS/PVP at RT prior to fixation for 1 h in a 100 ml drop of PBS/PVP–1% formaldehyde at RT. Following fixation, cells are washed through three 100 ml drops of PBS/PVP at RT, transferred to a 100 ml drop of storage buffer, and kept at 4 C.
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4. Sperm-Egg Interactions 4.1. Incubation of sperm with fully-grown oocytes, ovulated eggs, and two-cell embryos Ovulated eggs or fully-grown oocytes and two-cell embryos, either stored in M199-M and kept at 37 C for no more than 2 days, or kept in storage buffer at 4 C, are washed through three 100 ml drops of M199-M at RT and transferred to the incubator for at least 30 min before being added to capacitated sperm (Williams et al., 2006). Ten microliter drops of prewarmed M199-M are pipetted into a tissue culture dish and covered with oil. Alternatively, if a test substance is being evaluated, the sample is dissolved in a 10-m l drop of M199-M and pipetted into a tissue culture dish, as above. Then 10 ml of capacitated ‘‘swim-up’’ sperm are added to the drop (through the oil layer; 1:1 dilution) and incubated for 15 min at 37 C. Following this period, oocytes or eggs, and two-cell embryos are added to the 20 ml drop with as little medium as possible and incubated at 37 C for an additional 35–45 min. Sperm motility is monitored under a dissecting microscope during this period and samples with sperm motility of less than 70% (i.e., less than 7 motile sperm out of 10 sperm) should be discarded. At the end of the incubation time, oocytes or eggs, and embryos with associated sperm are removed from the drops with a mouth-operated, wide-bore Pasteur pipette and washed through three 100 ml drops of M199-M. This step involves careful up-anddown pipetting (3–4 times) of groups of 5–10 cells in each drop until no more than one to two sperm remain attached to the two-cell embryos (negative control). Under these conditions, sperm bound to the oocyte or egg zona pellucida (ZP) remain bound, whereas sperm nonspecifically attached to oocyte or egg and embryo ZP are removed. Oocytes and eggs with bound sperm, as well as embryos, are then transferred to 10 ml drops of M199-M containing PBS/PVP–2% formaldehyde (1:1 by volume) covered with oil.
4.2. Analysis of sperm binding to fully-grown oocytes and ovulated eggs The number of sperm bound per oocyte, egg, or embryo is determined by counting sperm tails in the largest diameter focal plane, using dark-field microscopy and 200 magnification. Typically, there are 12 oocytes or ovulated eggs and two two-cell embryos present per sample. The presence of embryos is essential in this assay since they serve as a negative control; sperm do not bind to the ZP of fertilized eggs or embryos. After determining the number of sperm bound to each oocyte or egg in one plane of focus, the highest and lowest values are discarded, and the remaining 10 values are used to calculate the average number of sperm bound per oocyte or
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egg ( S.D.). For example, when sperm are exposed to M199-M alone (no test substance), at sperm concentrations of about 5 105 sperm/ml, and incubated with oocytes or eggs and embryos, there are 30–50 sperm bound per oocyte or egg in the largest diameter focal plane (Fig. 5.1C).
5. Analysis of the Status of the Sperm’s Acrosome The acrosome is a large lysosome-like vesicle overlying the sperm’s nucleus (Florman and Ducibella, 2006; Yanagimachi, 1994). Capacitated mouse sperm undergo the acrosome reaction (AR) shortly after binding to the egg ZP. The AR is a specialized form of cellular exocytosis. It involves multiple fusions between plasma membrane overlying the anterior region of the sperm head and the outer acrosomal membrane lying just beneath the plasma membrane. Small hybrid membrane vesicles form and the inner acrosomal membrane, overlying the sperm’s nucleus, is exposed to the egg ZP (Cohen and Wassarman, 2001). Only acrosome-reacted sperm can penetrate the ZP, reach the plasma membrane, and fuse with eggs (i.e., fertilize eggs). To assess the status of the sperm’s acrosome (i.e., whether it is acrosomeintact or acrosome-reacted), capacitated ‘‘swim-up’’ sperm (see above) are incubated at 37 C for 1 h in M199-M, either in 20–50 ml drops under oil in a tissue culture dish or in 0.5-ml Eppendorf tubes, in the presence or absence of a test substance. Sperm exposed to Ca2þ ionophore A23187 are used as a positive control since it induces sperm to undergo the AR in vitro. Capacitated sperm are pipetted into a solution of 20 mM A23187 at a 1:1 dilution to a final concentration of 10 mM (it should be noted that sperm incubated in A23187 become completely immotile within a few minutes of exposure). After incubation, sperm samples are pelleted in 0.5-ml Eppendorf tubes by centrifugation at 10,000 rpm for 5 min in a microfuge at RT and fixed with PBS–5% formaldehyde (1:1 dilution with sperm sample) overnight at 4 C. The following day, sperm are washed by centrifugation (as above) in 0.1 M ammonium acetate, pH 9.0, resuspended in 50 ml of 0.1 M ammonium acetate by gentle vortexing, pipetted onto gelatin-coated glass slides, and air-dried. Several methods have been described to assess the status of the sperm’s acrosome. Two of these methods, Coomassie staining and FITC–lectin staining of sperm, are described in the following sections.
5.1. Coomassie-staining of sperm Slides are washed with distilled H2O, methanol, H2O, for 5 min each, Coomassie-stained for 5 min, and washed extensively with H2O (Larson and Miller, 1999). Sperm can be observed directly on the glass slide or under
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a coverslip (sealed with clear nail polish for long-term storage) using PBS– 30% glycerol as mounting medium, and are scanned in the presence or absence of an acrosome by bright-field microscopy at 400 magnification.
5.2. Lectin–FITC staining of sperm Slides are washed twice with PBS for 5 min, incubated with PNA–FITC (A. hypogaea) for 30 min at RT (humid chamber), and rinsed with PBS for 15 min (Lybaert et al., 2009). Samples are mounted with antifade mounting reagent and a coverslip (sealed with clear nail polish) and examined for the presence or absence of an acrosome by fluorescent microscopy at 400 magnification. Sperm retaining an intact acrosome (Fig. 5.2A) will display a continuous blue (Coomassie)- or fluorescent green (PNA–FITC)-stained ridge overlying the sperm head. Sperm that have undergone the AR (Fig. 5.2B) will display either a patchy blue or patchy fluorescent green pattern, indicative of a partial AR, or no staining at all, indicative of a completed AR. Typically, A23187-treated sperm samples will be 60–80% acrosome-reacted, whereas sperm exposed to M199-M alone will be 15–25% acrosomereacted (due to a so-called ‘‘spontaneous AR’’).
6. Visualization of Gold-Labeled Antibody on Sperm Capacitated sperm are incubated with a sperm-binding probe (i.e., antigen of choice), fixed, washed and air-dried on gelatin-coated slides as described above (Williams et al., 2006). Samples on slides are blocked with a A
B
Figure 5.2 Light micrographs of (A) acrosome-intact and (B) acrosome-reacted mouse sperm, fixed in 2.5% formaldehyde and stained with Coomassie brilliant-blue G-250. Arrow: acrosomal cap (bar 5.5 mm).
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50 ml drop of PBS/PVP–0.05% Tween 20 containing 5% goat serum (blocking buffer) for 30 min at RT in a humid chamber, then rinsed in H2O and air-dried for 15 min at RT. A diluted gold suspension (1:10) of antibody–gold conjugate (antibody of choice, coupled to 5 nm colloidal gold particles) in blocking buffer is pipetted onto the sperm sample (50 ml drop). The slide is incubated for 2 h at RT in a humid chamber on a slow moving shaker and washed afterward with PBS by submersion in a Petri dish for 5 min. The sample is fixed again with PBS/PVP–2.5% glutaraldehyde (50 ml drop) for 1 h at RT (humid chamber) and rinsed thoroughly with H2O in a Petri dish. To enlarge colloidal gold labels for high-contrast signals that are visible by light microscopy, a 50 ml drop of silver stain solution (Silver Enhancer Kit) is applied onto the gold-labeled sperm samples for 15 min at RT in the dark (humid chamber). The slide is dipped in H2O, fixed with 2.5% sodium thiosulfate pentahydrate (50 ml drop) for 3 min at RT, rinsed with H2O, mounted with PBS/PVP–30% glycerol and a coverslip, and sealed with clear nail polish. Individual sperm are scanned by light microscopy for silver-enhanced colloidal gold–protein complexes and the number of particles (silver precipitate) associated with each sperm is determined.
ACKNOWLEDGMENTS Our research was supported in part by the National Institutes of Health (NICHD), most recently by grant HD-35105.
REFERENCES Austin, C. R., and Short, R. V. (eds.), (1982). Reproduction in Mammals. 1. Germ Cells and Fertilization, Cambridge University Press, Cambridge. Cohen, N., and Wassarman, P. M. (2001). Association of egg zona pellucida glycoprotein mZP3 with sperm protein sp56 during fertilization in mice. Int. J. Dev. Biol. 45, 569–576. Florman, H. M., and Ducibella, T. (2006). Fertilization in mammals. In ‘‘The Physiology of Reproduction,’’ ( J. D. Neill, ed.), 3rd edn. pp. 55–112. Elsevier/Academic Press. Larson, J. L., and Miller, D. J. (1999). Simple histochemical stain for acrosomes on sperm from several species. Mol. Reprod. Dev. 52, 445–449. Lybaert, P., Danguy, A., Leleux, F., Meuris, S., and Lebrun, P. (2009). Improved methodology for the detection and quantification of the acrosome reaction in mouse spermatozoa. Histol. Histopathol. 24, 999–1007. Nagy, A., Gertsenstein, M., Vintersten, K., and Behringer, R. (2003). Manipulating the Mouse Embryo: A Laboratory Manual. 3rd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Qi, H., Williams, Z., and Wassarman, P. M. (2002). Secretion and assembly of zona pellucida glycoproteins by growing mouse oocytes microinjected with epitope-tagged cDNAs for mZP2 and mZP3. Mol. Biol. Cell 13, 530–541.
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Rafferty, K. A., Jr. (1970). Methods in Experimental Embryology of the Mouse. Johns Hopkins Press. Sorensen, R. A., and Wassarman, P. M. (1976). Relationship between growth and meiotic maturation of the mouse oocyte. Dev. Biol. 50, 531–536. Wassarman, P. M. (1996). Oogenesis. In ‘‘Reproductive Endrocrinology, Surgery, and Technology,’’ (E. Y. Adashi, J. A. Rock, and Z. Rosenwaks, eds.), Vol. 1, pp. 341–357. Lippincott-Raven Press. Wassarman, P. M., and DePamphilis, M. L. (eds.), (2003). In Guide to Techniques in Mouse Development, Methods in Enzymology Vol. 225. Academic Press. Williams, Z., Litscher, E. S., Jovine, L., and Wassarman, P. M. (2006). Polypeptide encoded by mouse ZP3 exon-7 is necessary and sufficient for binding of mouse sperm in vitro. J. Cell. Physiol. 207, 30–39. Yanagimachi, R. (1994). Mammalian fertilization. In ‘‘The Physiology of Reproduction 1,’’ (E. Knobil and J. D. Neill, eds.), pp. 189–317. Raven Press.