Method Article

Massive Ovulation in Mammals: Techniques for Analyzing Natural Spontaneous Polyovulation and Induced Polyovulation in Lagostomus maximus

DOI:

10.3791/69653

⸱

February 6th, 2026

 ,  ,  ,  ,  , 
1Centro de Estudios Biomédicos Básicos, Aplicados y Desarrollo (CEBBAD), Universidad Maimónides, 2Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET)

Corresponding Authors: Noelia P. Leopardo <leopardo.noelia@maimonides.edu>

In This Article

Summary

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This protocol describes methods for analyzing natural and induced ovulation in Lagostomus maximus, including oocyte recovery after spontaneous, hormonal-induced, or seminal plasma-induced ovulation. It provides a comprehensive framework for studying ovarian responsiveness and reproductive physiology in a highly polyovulatory mammalian model.

Abstract

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The South American plains vizcacha, Lagostomus maximus, is recognized as the mammalian species with the highest ovulation rate, which can reach as many as 300 ova. This remarkable polyovulation phenomenon can be examined through several methodological approaches that reflect its distinctive reproductive physiology. Natural ovulation is assessed by oocyte release during the breeding season, which provides insights into the species' spontaneous ovulatory mechanisms, with oocytes recovered by flushing the oviduct and uterine horns. In addition, experimental induction of ovulation using exogenous hormones, such as gonadotropins, allows controlled evaluation of ovarian responsiveness and offers a valuable model for understanding endocrine regulation in hystricognath rodents. A third approach involves the use of autologous seminal plasma as a physiological trigger of ovulation, highlighting the coexistence of induced and spontaneous mechanisms in this species. Together, these techniques not only elucidate the dynamics of follicular recruitment and oocyte release but also establish L. maximus as a comparative model for reproductive biology, with potential translational implications for understanding ovulation control in mammals.

Introduction

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The South American plains vizcacha (Lagostomus maximus) is a hystricognath rodent distinguished by its exceptional reproductive physiology, characterized by an extraordinary ovulation rate that can exceed 300 oocytes per cycle1,2. This distinctive feature has positioned L. maximus as a natural model for studying ovarian function and ovulatory control in mammals3,4,5,6,7,8,9,10,11.

Unlike typical spontaneous ovulators, L. maximus exhibits both spontaneous and induced ovulatory mechanisms2. This duality, combined with massive follicular growth and asynchronous ovulation, challenges conventional approaches based solely on endocrine or histological endpoints4,12,13,14. Consequently, the use of direct, quantitative techniques is essential to accurately determine the timing and magnitude of oocyte release.

Adult female vizcachas were obtained from a natural resident population maintained at the Estación de Cría de Animales Silvestres (ECAS), Parque Pereyra Iraola, Berazategui, Buenos Aires Province, Argentina (34°49′57″ S, 58°06′12″ W). We acknowledge that sovereign rights over natural resources are the exclusive property of the Province of Buenos Aires. The number of captured animals was authorized by the Ministry of Agriculture of Buenos Aires Province. Females were captured using live traps placed at burrow entrances at different time points throughout the year.

Capture periods were selected according to the species natural reproductive cycle established by Llanos & Crespo15, and refined based on our fieldwork experience7,8,10,14. The species presents a main reproductive season that extends from late summer (March) until the beginning of springtime in late September16,17,18, and a secondary breeding period that occurs in October, especially in females that have lost their offspring15. Accordingly, the following working groups were determined: a) Group I (N= 27): late February-early April (non-pregnant, onset of main reproductive season); b) Group II (N= 26): mid-September-late October (non-pregnant, post-lactation estrus); Group III (N= 35): December-January (non-pregnant, hormone-induced ovulation); Group IV (N= 7): December-January (non-pregnant, seminal plasma-induced ovulation).

This study presents and validates several methodological approaches for analyzing ovulation in L. maximus: (1) assessment of natural ovulation through direct oocyte recovery; (2) induction of ovulation using exogenous gonadotropins; and (3) induction of ovulation with autologous seminal plasma. Each technique provides complementary insights into follicular dynamics, oocyte maturation, and the regulatory mechanisms governing ovulation.

Flushing of the oviducts and uterine horns enables direct recovery of oocytes and accurate quantification of ovulation, allowing correlation with follicular development and oocyte morphology19,20,21. Hormonal stimulation with exogenous gonadotropins, adapted from protocols used in laboratory rodents22, allows controlled evaluation of ovarian responsiveness and the timing of oocyte release. Meanwhile, the use of autologous seminal plasma as a physiological trigger of ovulation -previously described in induced ovulators such as rabbits and camelids23,24,25-offers an innovative means of exploring the endocrine and paracrine factors involved in ovulation.

By integrating these complementary methodologies, the present work provides a reproducible framework for investigating ovulatory mechanisms in L. maximus. The combination of natural and induced models allows differentiation between spontaneous follicular recruitment, ovulatory efficiency, and seminal plasma-mediated responses. Moreover, these approaches reveal phenomena such as spontaneous parthenogenetic oocyte activation and the coexistence of "defective" and "euovulatory" oocytes, contributing to the understanding of follicular selection and oocyte competence2.

Beyond its species-specific implications, the vizcacha model offers broader relevance for comparative reproductive biology. Its mixed ovulatory strategy bridges characteristics of induced and spontaneous ovulators, providing valuable parallels to reproductive processes in other mammals, including domestic species and humans7,14,26,27.

The methodological protocols detailed herein are designed to ensure reproducibility and to facilitate their adaptation to diverse experimental contexts involving follicular dynamics, ovulatory regulation, and hormonal manipulation.

Protocol

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The experimental protocols described herein were reviewed and approved by the Institutional Committee on the Use and Care of Experimental Animals (CICUAE, Universidad Maimónides, protocol number #6184). Trapping, handling, and euthanasia were performed by trained technical staff in compliance with all local, state, and federal regulations governing the care and use of laboratory animals. Animal husbandry followed the National Institutes of Health (NIH) Guidelines for the Care and Use of Laboratory Animals28 and the guidelines of the American Society of Mammalogists (ASM) for the use of wild mammals in research29. All experiments complied with the ARRIVE 2.0 guidelines30. Every effort was made to minimize the number of animals used. The South American plains vizcacha (Lagostomus maximus) is not considered an endangered species31.

1. Hormonal induction of ovulation

​NOTE: This section presents hormonal induction of ovulation in non-pregnant females of captured vizcachas. Three different protocol options are discussed.

  1. Protocol A:
    1. Administer 200 IU of equine chorionic gonadotropin (eCG) intramuscularly at 0 h and 24 h, followed by 500 IU of human chorionic gonadotropin (hCG) at 48 h.
    2. Rest females until euthanasia.
    3. Euthanize females 60-132 h post-hCG administration (see section 3).
  2. Protocol B:
    1. Administer 200 IU of eCG intramuscularly at 0 h and 24 h, followed by 1,000 IU of hCG at 72 h.
    2. Rest females until euthanasia.
    3. Euthanize females 60-132 h post-hCG administration (see section 3).
  3. Protocol C:
    1. Administer 250 IU of eCG intramuscularly at 0 h, 24 h, and 48 h, followed by 2 mg of porcine luteinizing hormone (pLH) at 72 h.
    2. Rest females until euthanasia.
    3. Euthanize females 60-132 h post-pLH administration (see section 3 and Table 1).
      NOTE: The animals in each group were euthanized at different time points (ranging from 60 h to 132 h post-hCG administration). This temporal distribution was essential to confirm the extended nature of the ovulation period in this species. Specifically, for experiments aiming for the exclusive collection of mature oocytes, euthanasia was consistently performed at 120 h post-hCG.

2. Induction of ovulation by seminal plasma

NOTE: Test the capacity of seminal plasma to induce ovulation by intramuscular administration in hormonally primed females.

  1. Semen collection and preparation
    1. Collect semen from reproductively active males by electroejaculation, as described by Giacchino32.
    2. Centrifuge semen at 200 × g for 10 min at room temperature to pellet sperm cells and cellular debris.
    3. Supplement supernatants (seminal plasma) with penicillin and streptomycin and store at -20 °C until use.
  2. Induction of ovulation in non-pregnant females by seminal plasma
    1. Administer 200 IU of eCG intramuscularly at 0 h and 24 h for each female.
    2. Inject seminal plasma intramuscularly 48 h after the final eCG dose.
    3. Rest females until euthanasia.
    4. Euthanize animals 72 h after seminal plasma injection (see section 3).
      NOTE: Include control females injected with saline solution instead of seminal plasma.

3. Anesthesia, euthanasia, and reproductive tract collection

NOTE: Use anesthesia and humane euthanasia methods to obtain intact reproductive tracts for oocyte and histological analyses.

  1. Induce euthanasia via a single, calculated intramuscular injection of 13.5 mg/kg ketamine chlorhydrate and 0.6 mg/kg xylazine chlorhydrate administered according to the manufacturer's recommendations and established institutional protocols.
  2. Confirm deep anesthesia by the absence of response to pain (pedal reflexes).
  3. Place the animal in the supine position and sterilize the abdomen.
  4. Surgically open the abdominal cavity to fully expose all abdominal organs.
  5. Perform a general inspection to determine the overall condition of the animal and confirm natural or hormone-induced ovulation by the presence of ovulatory stigmata.
  6. Obtain a blood sample by direct puncture of the superior vena cava for serological testing.
  7. Perform euthanasia on the animal by administering 0.5 mL/kg of sodium pentobarbital intracardially.
  8. Carefully dissect and remove the ovaries, oviducts, and uterine horns, taking care not to damage their structures.
  9. Keep tissues in ice-cold phosphate-buffered saline (PBS) until processing (≤15 min).

4. Oocyte recovery

NOTE: Recover oocytes from the oviducts and uterine horns by gentle flushing to ensure complete collection of released gametes while minimizing tissue damage.

  1. Prepare the flushing apparatus by attaching a blunted 26-G needle to a 1 mL syringe filled with 0.9% NaCl supplemented with 4 mg/mL BSA, 100 µg/mL penicillin G, and 100 IU/mL streptomycin.
  2. Gently insert the needle into the infundibulum of each oviduct and slowly inject the flushing solution.
  3. Collect the effluent from the distal oviduct and uterine horn into sterile Petri dishes.
  4. Repeat flushing until the recovered fluid appears clear and free of visible oocytes or cellular debris.
  5. Transfer oocytes to 100 µL droplets of M2 medium containing 4 mg/mL BSA and antibiotics under mineral oil.
  6. Confirm successful recovery by inspecting droplets under a stereomicroscope; expect to observe free oocytes lacking cumulus cells dispersed within the fluid.
  7. Continue flushing until the effluent appears clear and no additional oocytes are observed under the stereomicroscope (typically 3-5 flushes per oviduct).
    NOTE: Recovered oocytes should appear spherical and translucent under low magnification.
  8. Classify oocytes according to morphological integrity as described in section 6.

5. Oocyte morphological assessment

NOTE: Evaluate the quality of recovered oocytes based on cytoplasmic appearance, zona pellucida uniformity, and nuclear maturation stage.

  1. Identify oocytes as normal if they display a spherical shape, uniform zona pellucida, homogeneous translucent cytoplasm, and a visible germinal vesicle or first polar body.
  2. Identify oocytes as abnormal if the cytoplasm appears dark, granular, or fragmented, or if the zona pellucida is irregular.
  3. Record the nuclear maturation stage as germinal vesicle (GV), germinal vesicle breakdown (GVBD), or metaphase II (MII).

6. Evaluation of oocyte nuclear maturation and meiotic spindle integrity

NOTE: Assess nuclear maturation and spindle morphology by immunofluorescence staining of α-tubulin and chromatin counterstaining.

  1. Fix oocytes in 2% paraformaldehyde (PFA) for 20 min.
  2. Permeabilize in 0.2% Triton X-100 in PBS for 40 min.
  3. Post-fix in 2% PFA for 20 min and wash three times in PBS (30 min each).
  4. Block with 2% goat serum, 2% BSA, 2% milk powder, 0.1 M glycine, and 0.01% Triton X-100 in PBS for ≥ 1 h.
  5. Incubate oocytes overnight at 4 °C with mouse anti-α-tubulin antibody (1:200).
  6. Wash in PBS for 30 min and incubate for 1 h in darkness with fluorescein isothiocyanate (FITC)-conjugated goat anti-mouse IgG (1/1000).
  7. Counterstain with 10 µg/mL Hoechst 33324 for 10 min and wash in PBS containing 2% BSA.
  8. Mount oocytes in antifade medium and image using a confocal microscope.
  9. Capture and process images using EZ-C1 software (EZ-C1 version 3.90); adjust contrast and brightness in image-processing software as required.
    NOTE: Successful staining is indicated by a clear green fluorescence pattern of the meiotic spindle (FITC) and blue nuclear counterstaining with Hoechst 33324.

7. Statistical analysis

NOTE: Analyze quantitative data to compare ovulatory responses and oocyte quality among experimental groups.

  1. Express data as mean ± SD.
  2. Use Student's t-test for two-group comparisons.
  3. Apply one-way ANOVA followed by Bonferroni's post hoc test for multiple comparisons.
  4. Consider p < 0.05 statistically significant.

Results

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Natural ovulation

Twelve out of 27 females (group I) captured from late February to mid-April, coinciding with the primary mating season, were observed to be ovulating, whereas during the secondary mating season (September/October) (group II), 10 out of 26 captured females exhibited ovulation. A comparable success rate of approximately 40% of ovulating females was observed during both capture periods. No swelling of the ampullary region of the oviduct was noted in any instance, and oocytes were retrieved from females regardless of the presence of ovulatory stigmata (Figure 1A,B). Oocytes were retrieved in a naked state, meaning they were not enveloped by cumulus cells (Figure 1C, Table 2).

An overall mean of 154 (± 87) oocytes per female was documented, with a range of 29 to 326, as no statistically significant differences were identified between group I (198 ± 92 [mean + SD], range 33-326) and group II (mean 116 ± 66, range 29-204) (Table 2)

Induced ovulation

As seen in natural ovulation, hormone-induced ovulation was characterized by the absence of a clearly defined ampullary region, alongside the release of cumulus-free oocytes that were dispersed throughout the oviducts and uterine horns. The average quantity of released oocytes was found to be similar to that noted in naturally ovulating females, and no statistically significant differences were detected among the three protocols that were examined (Table 3). Successful ovulation can be accomplished through the administration of eCG and hCG or pLH; however, it is noteworthy that an increase in hCG (as seen in protocol B) or its substitution with pLH (as observed in protocol C) resulted in a more substantial release of oocytes (Table 3). Ovulation persisted over an extended temporal span. Oocytes were initially recovered at 60 hours subsequent to hCG administration, with recovery continuing until 132 hours post-hCG, and the mean number of recovered oocytes reached its maximum at 84 hours post-hCG (Table1).

Oocyte quality in natural and hormone-induced ovulation

A significantly high incidence of morphological anomalies in oocytes was observed in both naturally and hormone-induced ovulating females (Figure 1C). In the cohort of naturally ovulating females, the prevalence of morphologically abnormal oocytes reached 82.3% (2175/2676), whereas in the hormonally induced cohort, this prevalence was recorded at 45.9% (851/1855). The identified abnormalities encompassed cytoplasmic fragmentation and granulation, ooplasmic vacuolization, the presence of refractile bodies, enlarged perivitelline space, and empty zona pellucida (ghost oocytes).

The natural ovulation females exhibited abnormal oocytes in their flushed product, but in some cases the collected oocytes/embryos were identified as morphologically normal. These females, which were captured at the peak of the mating season, displayed a combination of oocytes and preimplantation embryos in their flushed products. Examples of normal oocytes and developing embryos are shown in Figure 2.

Intriguingly, in the hormonally induced females, a majority of the oocytes categorized as morphologically normal were identified as immature oocytes; specifically, oocytes exhibiting either a germinal vesicle or germinal vesicle breakdown. Nevertheless, upon analysis of oocyte nuclear maturation, it was determined that 47.6% (90/189) of those oocytes classified as normal immature oocytes manifested nuclear defects, including abortive activation and anomalies in meiotic spindle configuration, such as tri- and tetra-polar spindles (Figure 3). The detection of activated oocytes extruding the second polar body, together with early embryos, suggests parthenogenetic activation (Figure 4), as these females were isolated prior to hormone treatment and were not pregnant. So, morphology underscores the presence of oocyte abnormalities in this case.

It is noteworthy that within a single female from the natural ovulating group, a mass of cumulus-enclosed oocytes was retrieved alongside a substantial quantity of abnormal naked oocytes. Following manual cumulus dispersal using a fine needle, the recovered oocytes were assessed (Figure 5). They exhibited a normal morphology, characterized by a conserved spherical shape, a uniform zona pellucida, and a homogeneous cytoplasm (Figure 5C).

Induction of ovulation by seminal plasma

In females exposed to seminal plasma (Group IV), ovaries exhibited distinct gross morphological features, including prominent, well-vascularized hemorrhagic stigmata protruding from the surface and a characteristic ovarian rim with elongated, thinned, nearly translucent epithelium containing numerous follicular structures. Histological analysis confirmed the presence of hemorrhagic follicles at different stages of luteinisation, as well as pedunculated corpora lutea on the ovarian surface. Flushing of oviducts yielded cumulus-enclosed oocytes (COCs) (Figure 5A), comparable to those observed in a naturally ovulating female (Figure 5B). Mechanical dispersion of COCs revealed morphologically normal oocytes, characterized by a spherical shape, uniform zona pellucida, and homogeneous translucent cytoplasm (Figure 5C). In contrast, control females treated with physiological saline instead of seminal plasma did not yield oocytes or COCs upon flushing, and no hemorrhagic stigmata were observed in their ovaries.

Tissue section analysis, microscopy image showing cellular structure and distribution of red-stained cells.
Figure 1: The ovulating adult ovary of the plains vizcacha. (A) General view of an adult plains vizcacha ovary showing ovulation stigmata, product of follicular rupture, and subsequent invasion with hematic material. (B) Magnified image of the ovary showing hemorrhagic points resulting from follicular rupture. (C) View of the product of oviduct/uterine horn flushing showing the release of oocytes devoid of cumulus cells, with the majority exhibiting gross morphological abnormalities. Scale bars: (A,B) 350 µm; (C) 100 µm. Please click here to view a larger version of this figure.

Microscopy time-lapse of cellular division stages; multi-frame sequence; embryogenesis process.
Figure 2: Normal oocytes and embryos from a female plains vizcacha with natural ovulation. (A) Metaphase II mature oocyte with the first polar body. (B) Activated oocyte showing extrusion of the second polar body (green arrow); note the remnants of the first polar body (pink arrow). (C) Two-cell embryo. (D) Four-cell embryo. (E) Early blastocyst. (F) Hatching blastocyst. Scale bars: 25 µm. Please click here to view a larger version of this figure.

Cell division stages microscopy images, diagram; cellular analysis, fluorescent labeling process.
Figure 3: Spontaneously activated oocytes from plains vizcachas exhibiting morphological alterations and parthenogenetic development. (A) Spontaneous two-pronuclei (red and blue arrowheads) zygote. (B) Polarised cytoplasm in a zygote undergoing cytokinesis. (C) Two pronuclei (red and blue arrowheads) zygote, and polar body (green arrowhead) with cytoplasm retraction. (D) Abnormal two-cell embryo. (E) Unequal divided four-cell embryo. (F) Haploid zygote with one pronucleus (blue arrowhead) and extrusion of second polar body (green arrowhead). (G)Immunofluorescence detection of the normal meiotic spindle. (H) Tripolar spindle. (I) Oocyte with two metaphase plates and failure of the second polar body extrusion. Scale bars: (A-F) 40 µm; (G-I) 10 µm. Please click here to view a larger version of this figure.

Cell division stages, fluorescence microscopy, labeled chromosomes, cell biology, image set analysis.
Figure 4: Oocyte morphology and development in hormone-induced ovulating female plains vizcachas. (A) Immature oocyte. (B) Germinal vesicle breakdown. (C) Metaphase II plate. (D) Metaphase II and nuclear material in the polar body. (E) Activated oocyte with two pronuclei. (F) Abortive telophase activation. (G) Two-cell embryo. (H) Two-cell embryo with initial cytoplasmic fragmentation. The nuclear material was stained with Hoechst 33324. Scale bars: 10 µm. Please click here to view a larger version of this figure.

Microscopic image, pollen structure comparison: A, B clumped morphology; C isolated pollen grains.
Figure 5: Induced ovulation by the administration of seminal plasma in the plains vizcacha. (A) Cumulus-oocyte complex recovered after oviduct flushing in a female stimulated with seminal plasma. (B) Cumulus-oocyte complex recovered from a single female experiencing natural ovulation. (C) Morphologically normal oocytes obtained after mechanical disaggregation of the cumulus-oocyte-complex. Scale bars 40 µm; Please click here to view a larger version of this figure.

Hours after hCGNNumber of oocytes (per female)Mean ±SD
3630; 0; 0-
6047; 24; 95; 12562.75  ± 56.34
84617; 41; 115; 156; 158; 182111.50 ± 67.86
10848; 31; 45; 8041.00  ± 30.14
132439; 46; 50; 5046.25 ± 5.18
*data correspond to treatments A and B (cf. Table 3 and Protocol) analyzed together. eCG: equine chorionic gonadotropin. hCG: human chorionic gonadotropin. SD: standard deviation.

Table 1: Timeline distribution of oocyte recovery after hCG administration in females induced to ovulate via eCG/hCG stimulation*. *Data correspond to treatments A and B (cf. Table 3 and Protocol section) analyzed together. eCG: equine chorionic gonadotropin. hCG: human chorionic gonadotropin.

Experimental group (capture time)Ovulating females/captured females (%)Total Oocytes recoveredOocytes/female (mean ± SD)Range (min - max)
GROUP I (February/April)12/27 (44.44)2238186 ± 92 33–326
GROUP II (September/October)10/26 (38.36)1160116 ± 6629–204
TOTAL22/53 (41.51)3398154 ± 8729–326
 SD: standard deviation. No statistical differences were found between groups (p ≤ 0.05).

Table 2: Oocyte recovery after flushing in naturally polyovulating Lagostomus maximus females. No statistical differences between mean values (p < 0.05). SD: standard deviation.

ProtocolRf/Tf* (%)Recovered oocytes
Mean ±SDRange
A8/11 (72.7)111 ± 5450–156
B10/10 (100.0)127 ± 4285–182
C13/14 (92.8)188 ± 9560–248
*Rf: responding females; Tf: treated females; SD: standard deviation. No statistical differences were found between groups (p≤0.05).

Table 3: Hormone-induced ovulation. *Rf: responding females; Tf: treated females; SD: standard deviation. No statistical differences were found between groups (p < 0.05).

Discussion

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The protocol presented here for studying ovulation in the plains vizcacha provides a reproducible framework for assessing both natural and experimentally induced ovulatory events. A major advantage of this protocol lies in its capacity to directly quantify and characterize the extraordinarily high number of oocytes released by the plains vizcacha, a feature unmatched in other mammalian models. Unlike indirect methods based on hormonal profiles or histological sections, oviductal and uterine flushing provides immediate access to intact oocytes, enabling both quantitative and qualitative analyses. This approach also preserves the integrity of the cumulus-oocyte complexes (COCs), which is critical for assessing fertilization potential and morphological quality.

Another advantage is the methodological flexibility it offers. By combining natural ovulation with hormone-induced or seminal plasma-induced ovulation, the protocol enables comparative analyses of different ovulatory triggers within the same species. This versatility allows researchers to dissect the relative contributions of endocrine, mechanical, and seminal factors to the ovulatory process, something that cannot be easily achieved in traditional laboratory models.

The protocol is advantageous as the flushing technique yields oocytes without interfering with the normal ovulatory process and the organ. Moreover, the ovarian structure is preserved, since oocyte collection avoids the need for puncture, thereby allowing for the integration of morphological, histological, and molecular analyses within a single experimental design.

Several steps of the procedure proved to be particularly critical. The capture of females in relation to the reproductive season is essential, as ovarian activity varies throughout the year. The careful capture and handling of females to minimize stress. Precise identification of ovulatory stigmata and the careful recovery of oocytes by flushing the oviduct and uterine horn is fundamental: the use of a blunt needle together with a medium supplemented with BSA and antibiotics prevents mechanical damage and contamination, ensuring sample reliability. Finally, accurate morphological assessment is indispensable for distinguishing normal oocytes from abnormal or spontaneously activated ones, a step that strongly depends on the operator's expertise.

Modifications and troubleshooting are often required to optimize recovery, particularly when dealing with hormonally induced ovulation. For example, adjusting the timing and dosage of eCG/hCG or pLH can improve synchronization and maximize oocyte yield. The mechanical dispersal of cumulus-oocyte complexes is another critical step; excessive manipulation may damage oocytes, while insufficient dispersion can compromise morphological assessment. Troubleshooting these steps is likely to improve both the quality and quantity of oocytes recovered. The protocol allows certain modifications depending on the experimental aim. For instance, adjusting the timing or dosage of gonadotropin administration can optimize the ovulatory response, since not all hormonal regimens yield the same efficiency. Similarly, the amount of seminal plasma applied may influence the recovery of cumulus-oocyte complexes (COCs)33. Troubleshooting is often required during the flushing step, particularly when the fluid fails to circulate or when oviductal obstruction occurs. In such cases, repeating the flush or checking patency under a stereomicroscope generally resolves the problem.

The method has several limitations. First, the intricate ovarian anatomy and variability in the timing of ovulation across individuals can introduce variability in oocyte recovery. Second, spontaneous parthenogenetic activation of oocytes may complicate the interpretation of developmental competence34. Finally, while hormonal induction allows for controlled study outside the breeding season, it may not fully replicate all physiological aspects of natural ovulation.

Despite these limitations, this protocol offers distinct advantages over alternative methods. Unlike non-invasive imaging or hormonal assays alone, it allows for direct retrieval and morphological assessment of oocytes and early embryos, providing a more accurate estimate of ovulation rate and reproductive status. Compared to other polyovulatory species, the plains vizcacha model permits studies of high ovulatory output, spontaneous versus induced ovulation, and the role of seminal plasma in ovulatory mechanisms12,35,36.

The significance of this protocol extends to several research areas. It can be applied to studies of reproductive physiology, oocyte quality assessment, and the mechanisms underlying polyovulation and parthenogenesis. Furthermore, it provides a model system for testing the effects of hormonal treatments, seminal plasma components, and environmental factors on ovulation dynamics. Finally, the method may inform comparative reproductive studies across mammalian species and aid in the development of assisted reproductive technologies for polyovulatory or wild mammals.

Disclosures

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The authors declare that there is no conflict of interest that could be perceived as prejudicing the impartiality of the research reported.

Acknowledgements

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This work was supported by the intramural financial aid of Universidad Maimónides, Argentina, and the FONCYT-AMPCYT agency, Argentina (PICT-2018-04619, granted to NPL). The authors are especially grateful to the Ministry of Agriculture Authority of the Buenos Aires Province Government, Argentina, for authorizing the capture of animals; to the personnel of the Estación de Cría de Animales Silvestres (ECAS, Buenos Aires Province, Argentina) for their invaluable help in trapping and handling the animals; and to DVM Sergio Ferraris and DVM Fernando Lange from CCV-Universidad Maimónides for their assistance during the anesthesia and surgical procedures.

Materials

List of materials used in this article
NameCompanyCatalog NumberComments
Adobe PhotoshopAdobe Systems Inc., USAN/AImage processing
Bovine Serum Albumin (BSA)Sigma-Aldrich, USAA9647Used in flushing medium and blocking buffer
Centrifuge (for semen processing)Eppendorf, Germany5417RSeminal plasma preparation
Confocal Microscope (C1 Eclipse E800)Nikon Ltd., JapanN/AOocyte imaging
Electro-ejaculatorDesigned in the laboratory; adapted from domestic mammals for use in caviomorph rodentsN/ASemen collection
Euthanyl (sodium pentobarbital, sodium diphenylhydantoin)Brouwer S.A., ArgentinaN/AEuthanasia
FITC-conjugated goat anti-mouse IgG (AP124F)Chemicon, USAAP124FSecondary antibody
Hematoxylin-Eosin staining kitSigma-Aldrich, USAVariousHistology
Hoechst 33324Thermo Fisher Scientific, USAH3570Nuclear staining
Ketamine chlorhydrateHolliday Scott S.A., ArgentinaN/AAnesthesia
Laminar flow hoodThermo Scientific, USAN/AAseptic handling
Light microscopeOlympus, JapanBX51Histology
Lutropin-V (pLH)Bioniche Animal Health Care, CanadaN/AHormonal induction
M2 mediumSigma-Aldrich, USAM7167Oocyte recovery
Mineral oilSigma-Aldrich, USAM8410Oocyte culture
Novormon 5000 (eCG)Syntex S.A., ArgentinaN/AHormonal induction
Ovusyn (hCG)Syntex S.A., ArgentinaN/AHormonal induction
Paraformaldehyde (PFA, 2% and 4%)Sigma-Aldrich, USA158127Fixation
Penicillin GSigma-Aldrich, USAP3032Antibiotics for media
StereomicroscopeLeica Microsystems, GermanyM80Oocyte identification
StreptomycinGibco, Thermo Fisher Scientific, USA11860038Antibiotics for media
Syringe with blunt-end needle (1 mL)BD Biosciences, USA305219Flushing of oviducts/uterus
Triton X-100Sigma-Aldrich, USAT8787Permeabilization
VectaShield mounting mediumVector Laboratories, USAH-1000Mounting for fluorescence
Xylazine chlorhydrateRichmond Laboratories, ArgentinaN/AAnesthesia
XyleneSigma-Aldrich, USA534056Dewaxing paraffin sections
α-tubulin monoclonal antibody (mouse anti-α-tubulin, sc-5286 B7)Santa Cruz Biotechnology, USAsc-5286 B7For meiotic spindle immunostaining

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Polyovulation AnalysisNatural OvulationInduced OvulationOocyte RecoveryGonadotropin StimulationSeminal Plasma TriggerFollicular RecruitmentReproductive PhysiologyMammalian Ovulation
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