Rapid Isolation of Stage I Oocytes in Zebrafish Devoid of Granulosa Cells
Summary
This protocol describes a modified procedure for rapidly isolating clean stage I oocytes in zebrafish devoid of granulosa cells, thereby providing a convenient method for oocyte-specific research.
Abstract
The study of oocyte development holds significant implications in developmental biology. The zebrafish (Danio rerio) has been extensively used as a model organism to investigate early developmental processes from oocyte to embryo. In zebrafish, oocytes are surrounded by a single layer of somatic granulosa cells. However, separating granulosa cells from oocytes poses a challenge, as achieving pure oocytes is crucial for precise analysis. Although various methods have been proposed to isolate zebrafish oocytes at different developmental stages, current techniques fall short in removing granulosa cells completely, limiting the accuracy of genome analysis focused solely on oocytes. In this study, we successfully developed a rapid and efficient process for isolating pure stage I oocytes in zebrafish while eliminating granulosa cell contamination. This technique facilitates biochemical and molecular analysis, particularly in exploring epigenetic and genome structure aspects specific to oocytes. Notably, the method is user-friendly, minimizes oocyte damage, and provides a practical solution for subsequent research and analysis.
Introduction
The zebrafish is among the most important model systems in developmental biology. In recent years, numerous studies have utilized the zebrafish as a model to study important biological events and regulatory processes from oocyte to embryo. These encompass the intricate processes of oocyte development and maturation1, the functionality of maternal genes2, the regulation of maternal-zygotic transitions3, and extensive omics analyses4.
Granulosa cells, the somatic cells enveloping and nurturing the developing oocyte within the ovarian follicle5,6, play a pivotal role in this developmental process. As primordial germ cells (PGCs) evolve into oogonia, they become surrounded by a monolayer of granulosa cells7. Together with the external thecal cells, the oocyte and its surrounding granulosa cells constitute a mature follicle8. Given the fundamental distinction between germ cells and somatic cells, obtaining a pure oocyte sample is imperative, especially for genome-related analyses.
Within the follicular structure of zebrafish, granulosa cells typically exhibit a diameter of only a few microns8, emphasizing the intimate interconnection between granulosa cells and oocytes9. This close association presents a challenge in achieving complete separation due to the considerable difference in both the number and volume of granulosa cells versus oocytes (hundreds of granulosa cells compared to a single oocyte)10,11. Even minimal contamination with a single granulosa cell can impede downstream analyses specifically targeting oocytes. Therefore, for studies focusing on genomic and epigenetic characteristics, the elimination of granulosa cells is essential.
Benefiting from well-characterized morphological criteria, oocytes at each stage can be distinguished based on diameter11. The oogenesis process in zebrafish is categorized into five stages according to morphology and karyotype11. Stage I oocytes (7-140 µm diameter) encompass oocytes from the onset of meiosis to the early stage of meiosis I. Crucially, these oocytes are transparent, allowing for the observation of the central nucleus through transmitted light (Figure 1Ai). Stage II oocytes (140-340 µm diameter) gradually become foamy and translucent. With the enlargement of follicles and the proliferation of cortical alveoli, the germinal vesicles in the center become difficult to distinguish12 (Figure 1Aii). Stage III oocytes (340-690 µm diameter) progressively accumulate vitellogenin, and fresh follicles become increasingly opaque (Figure 1Aiii). Meiosis continues in stage IV oocytes (690-730 µm diameter) as chromosomes enter the middle of meiosis II, where they stagnate (Figure 1Aiv). Stage V oocytes (730-750 µm diameter) have matured and are ready for ovulation7,11(Figure 1Av).
Based on the unique characteristics of each of the aforementioned stages, a method has been proposed to isolate oocytes from stages I to III by digesting zebrafish ovaries using a digestive solution containing collagenase I, collagenase II, and hyaluronidase, followed by filtration through a specific-sized cell strainer13. However, while this method allows for obtaining oocytes at different developmental stages, it fails to completely separate oocytes and granulosa cells. Other researchers have also suggested methods to separate granulosa cells from oocytes. However, these methods primarily rely on mechanical approaches, which can cause oocyte damage, are time-consuming, and are inadequate for obtaining a substantial number of oocytes for analysis14,15.
Given the limitations of existing methods and specific research requirements, this study aims to establish a procedure to thoroughly separate oocytes and granulosa cells and obtain a sufficient number of clean stage I oocytes for analysis. Expanding upon the referenced method13, we employ an improved digestion buffer (see Table of Materials) that is gentler and facilitates the dispersion of oocytes and dissociation of granulosa cells. Subsequently, the oocytes are passed through a cell strainer, followed by washing and further microscopic selection, enabling the acquisition of a large number of clean stage I oocytes.
Protocol
All zebrafish were handled following stringent animal care guidelines outlined by the relevant national and/or local animal welfare bodies. The maintenance and handling of fish received approval from both local authorities and the animal ethics committee of the West China Hospital of Sichuan University (approval No. 20220422003). Zebrafish ovaries contain a mixture of multi-stage oocytes, with each developmental stage being present in adult zebrafish ovaries. However, juvenile zebrafish were selected for this study due to the dominance of late-stage oocytes (stage II-III) in the ovaries of adult fish, which are large and opaque. In contrast, juveniles have flat and elongated ovaries primarily consisting of stage I oocytes (Figure 1B) (Figure 1B). Previous research has demonstrated that the morphological features of early oogenesis in adult and juvenile zebrafish are highly consistent, supporting the feasibility of utilizing juvenile zebrafish13. The procedures described below, outlined in Figure 2, provide a detailed description of the isolation process, from dissection to obtaining clean stage I oocytes. For further reference, the reagents, buffers, and equipment utilized in the study are listed in the Table of Materials.
1. Ovary dissection
- Select several female zebrafish with a standard length (SL) ranging from 10 mm to 15 mm.
NOTE: The SL, measured from the snout to the tail base in millimeters, serves as a reliable criterion for selecting appropriate fish. It is recommended to use juvenile fish between 5 and 7 weeks post-fertilization (wpf) with an SL ranging from 10 mm to 15 mm for this procedure. The ovaries of juvenile zebrafish are predominantly occupied by early oocytes, providing optimal conditions for easily obtaining stage I oocytes (Figure 1B). - Euthanize the juvenile female fish by placing them in ice-cold water (0-4 °C).
NOTE: The following procedures must be carried out to ensure euthanasia and minimize the pain experienced by zebrafish during euthanasia: confirm that the temperature of the ice-cold water is within the 0-4 °C range using a thermometer, swiftly transfer the zebrafish to the ice-cold water, and maintain them in the ice-cold water for at least 10 min after cessation of opercular movement16,17,18. - Dry excess water on the fish's surface using absorbent paper.
- Dissect the zebrafish under a stereomicroscope by following these steps:
- Cut off the head using micro-scissors along the posterior margin of the gill cover; cut off the tail along the cloaca. Transfer the middle trunk to a 35 mm dish containing 2 mL of L-15 medium (with L-glutamine).
- Dissect the middle trunk in the L-15 medium. Cut along the central axis of the abdomen and remove the viscera and swim bladder.
NOTE: The viscera of zebrafish are connected. Therefore, pinch the hindgut near the anal region and lift it to easily remove the entire visceral mass. - Detach the bilateral ovaries from the abdomen using tweezers. Remove the adipose tissue, scales, and other body tissues attached to the ovary.
NOTE: The ovaries are covered with adipose tissue. Therefore, it is best to remove them together to avoid mechanical damage to the oocytes.
CAUTION: Handle tweezers with care to avoid injury.
2. Digestion
- Transfer the ovaries to a 6-well plate containing 2 mL of Kinger's cell dissociation solution and incubate for 2-3 h at 28.5 °C. Shake the 6-well plate every 30 min to promote dispersion.
NOTE: Use tweezers to separate the ovaries into small tissue blocks to enhance digestion efficacy. The digestion time is variable; regularly monitor and terminate the process when a significant number of stage I oocytes are scattered under the stereomicroscope. The Kinger's cell dissociation solution is the working solution and can be used directly. It can be stored stably at −20 °C after packaging and thawed at a low temperature (4 °C) before use. - Add L-15 medium preheated at 28.5 °C to the digestion buffer to stop digestion.
3. Filtration
- Place a 100 µm cell strainer into another well of the 6-well plate, add L-15 medium, and ensure that the medium level is higher than the filter.
NOTE: The theoretical diameter of stage I oocytes is less than 140 µm. However, oocytes with larger diameters may still pass through the cell strainer. Therefore, a cell strainer with a smaller diameter than the target was selected at this step. Mesh sizes can be adjusted depending on the specific experimental needs. - Draw the digestive medium through the cell strainer using a pipette. Wait 1-2 min until all the stage I oocytes have passed through the strainer, then remove it. Ensure that the oocytes remain in the liquid throughout the transfer process.
NOTE: Keeping the oocytes in the medium helps maintain their integrity and optimal condition. If the oocytes are exposed to air for an extended period, they could dry out and become damaged, affecting the final separation efficiency. Keep the pipette below the liquid surface when transferring the oocytes through the cell strainer rather than adding them in a suspended manner.
4. Washing
- Remove excess medium using a pipette. Add 4 mL of fresh L-15 medium and gently resuspend the oocytes. Wait 1-2 min, then remove the supernatant.
NOTE: Use instruments with low adhesion properties to prevent damage to the oocytes. - Repeat step 4.1 5-6 times until no other impurities are present.
NOTE: Gentle handling during the washing process is crucial to avoid oocyte damage. Despite obtaining an abundance of stage I oocytes (approximately 200-400 stage I oocytes from one zebrafish) following the previous washing steps, the oocytes may still be mixed with cell fragments, oocytes from other stages, or small granules formed by broken late-stage oocytes. Additionally, some granulosa cells may still adhere to the surface of several oocytes due to incomplete enzyme digestion. Therefore, for applications demanding a higher level of purity, such as genome sequencing, additional steps are necessary to select completely clean oocytes. To address this, proceed to steps 5 and 6.
5. Selection for quality control
- Transfer the washed stage I oocytes to a 35 mm dish containing fresh L-15 medium.
- Under a microscope with a magnification equal to or exceeding 10x, remove cell fragments, other-stage oocytes, and stage I oocytes adhered by granulosa cells using a tool that can be manipulated with precision, such as a blunt injection needle.
NOTE: This step requires patience from the experimenter, but it ensures the isolation of clean and intact oocytes.
CAUTION: Handle injection needles with care to avoid injury.
6. Confirmation by nuclear staining
NOTE: For further refinement of the isolated oocytes, staining with Hoechst 33342 was employed to identify and remove individual oocytes adhered with granulosa cells that may have been missed in the previous step.
- Add a final concentration of 5 µg/mL Hoechst 33342 to the L-15 medium and incubate at room temperature for 10 min.
- Observe the Hoechst fluorescence through a fluorescence microscope under UV laser excitation.
NOTE: Hoechst is typically excited in the ultraviolet region, usually at wavelengths between 310 and 360 nm. It is recommended to use a magnification between 80x and 100x during this process. Employ the automatic exposure mode. The excitation wavelength and exposure time can be adjusted based on the specific microscope model and individual requirements. - Using a needle, carefully pick out the oocytes that do not meet the desired criteria.
NOTE: When visualizing the cells, a large nuclear chromatin is present in the center of the oocyte. If granulosa cells were adhered to the oocyte, small and bright nuclei would be visible, clinging to the oocyte's edge (Figure 3). Pick them out under the microscope. - Thoroughly and repeatedly wash the oocytes with L-15 to remove excess Hoechst.
- Use the identified stage I oocytes directly for subsequent extraction or store them at −80 °C after flash-freezing in liquid nitrogen.
CAUTION: Wear a protective face mask and cryogenic gloves when handling liquid nitrogen and ultra-low temperature freezers.
Representative Results
Figure 1 illustrates the ovarian morphology observed in both adult and juvenile zebrafish, showcasing oocytes at different developmental stages to serve as a reference. Figure 1A provides a graphical representation of oocyte morphology and size at each developmental stage, beginning with the primary growth stage (stage I) and ending with the ovulated egg (stage V). Figures 1Ai-v illustrate the diameters corresponding to stages I to V of the oocytes. Figure 1B (left panel) displays ovarian images in juvenile fish (6 wpf with SL = 1.5 cm). Notably, the juvenile ovary exhibits a distinct lobular appearance, primarily characterized by an abundance of transparent stage I oocytes, accompanied by a smaller population of stage II oocytes. This prevalence of stage I oocytes greatly facilitates their observation and retrieval. The right panel depicts the morphology of the adult fish ovary (6 months), where oocytes at various developmental stages can be observed. However, opaque late-stage oocytes (stage II-III) are predominant in the ovary.
Figure 2 illustrates the representative processes from dissection to obtaining clean stage I oocytes. Figure 3 presents stage I oocytes obtained using both the reference method and the modified method. Following Hoechst staining, the oocyte and granulosa cell nuclei exhibited blue fluorescence under UV light. In the images obtained using the reference method (Figure 3A), the oocytes' surface was densely surrounded by numerous stained granulosa cell nuclei. In contrast, stage I oocytes separated by the modified method (Figure 3B) exhibit complete exposure and cleanliness, with no granulosa cell nuclei staining present.
Our findings demonstrated the effectiveness of the proposed method for isolating stage I oocytes. The developmental process of oocytes in teleost fish exhibits several common characteristics7. To assess its cross-species applicability, further analyses were conducted on other fish species. Female pond loaches were acquired from a local market, and early-stage oocytes (diameter <100 µm) were isolated following the same procedure. The ovarian composition of the pond loach also displayed a mixture of oocytes at multiple developmental stages, characterized by a striking resemblance in oocyte morphology and size to that of zebrafish during the same stage. As illustrated in Figure 4A, the ovaries of adult female pond loach were dominated by opaque late oocytes interspersed with transparent early oocytes. A clear monolayer of granulosa cells was observed on the surface of oocytes. Consistent with our observations in zebrafish, the loach oocytes measuring less than 100 µm in diameter corresponded to the stage prior to the diplotene stage. Figure 4B depicts the clean pond loach (Misgurnus anguillicaudatus) oocytes obtained by the modified method. Compared to zebrafish ovaries, pond loach ovaries are more challenging to dissociate. After 6-7 h of digestion, more early-stage oocytes dissociate. Nonetheless, this adjusted approach yielded the desired clean and naked oocytes. Zebrafish belong to the Cyprinidae family, whereas pond loaches are classified within the Cobitidae family. These two families exhibit a distant taxonomic relationship, but their ovarian development stage and morphology are remarkably similar, indicating that this method can generally be utilized for analyzing oocytes of various fish species.
Figure 1: Representative images of zebrafish ovaries and oocytes at different stages. (A) Morphology and size of oocytes at each developmental stage (St). i-v represents the images and diameter of the oocytes from stage I to V. Scale bar = 200 µm. (B) Ovarian morphology of adult and juvenile zebrafish. Scale bars = 500 µm. Please click here to view a larger version of this figure.
Figure 2: Representative images of the isolation procedure, from dissection to the acquisition of clean stage I oocytes. Please click here to view a larger version of this figure.
Figure 3: Stage I oocytes obtained through the two methods. (A) Stage (St) I oocytes isolated by the referenced method. (B) Stage I oocytes isolated through the modified method. The oocyte and granulosa cell nuclei were stained with Hoechst and excited with UV light. Scale bars = 100 µm. BF = bright field. Please click here to view a larger version of this figure.
Figure 4: Early-stage oocytes from pond loach obtained by the modified method. (A) Representative images of ovary and oocytes of pond loach. i, ii represent the ovaries of pond loaches under different magnifications, and iii-iv represents the early-stage oocytes of pond loaches. Scale bars: i (500 µm); ii (200 µm); iii, iv (100 µm). (B) Early-stage oocytes isolated from pond loach. The oocytes and granulosa cell nuclei were stained with Hoechst and excited with UV light. Scale bars = 100 µm. BF = bright field. Please click here to view a larger version of this figure.
Discussion
In this study, we developed a method for isolating pure and clean stage I oocytes, excluding granulosa cells, for downstream analysis (particularly genomic analyses). Comparing this modified method with the referenced method13, the stage I oocytes obtained using this method are morphologically intact, sufficient in number, and free from contamination with other somatic cells, making them suitable for various subsequent studies and analyses. Furthermore, compared with other mechanical separation methods14,15, the improved method is simpler to operate, facilitates the acquisition of a larger number of oocytes, and prevents oocyte damage.
Moreover, in addition to effectively isolating stage I zebrafish oocytes, this method can also be applied to pond loaches, indicating its broad applicability among a variety of fish species. To investigate the impact of granulosa cell contamination on gene expression and epigenetic analysis, we conducted RNA sequencing (RNA-seq) and CUT&Tag analysis on oocytes obtained using the reference and modified methods. Although a few differentially expressed genes were identified by RNA-seq, some genes that were specifically expressed in granulosa cells were still identified (data not shown). The analysis also revealed substantial differences in histone modifications between oocytes isolated with different methods (data not shown). These results underscore the importance of this method in completely separating oocytes and granulosa cells for accurate gene expression and epigenetic analysis.
Several aspects require attention when implementing the proposed method: (1) Selecting the appropriate size of zebrafish is crucial. This ensures that enough oocytes are obtained and reduces subsequent washing steps. The ovaries of zebrafish with an SL <10 mm do not provide sufficient oocytes, thus requiring a larger number of fish to meet sampling requirements and increasing breeding pressure. Conversely, zebrafish with an SL greater than 15 mm exhibit the gradual formation of stage II-III oocytes, the membranes of which are easily broken during the digestion reaction. (2) When larger fish are used, the leakage of cell contents resulting from damaged oocytes may lead to contamination, thereby inhibiting the digestion reaction to some extent. In such cases, removing the old digestive buffer, washing it with L-15, and then adding a new Kinger’s cell dissociation solution for further digestion is recommended. (3) If appropriate equipment is available, shaking during digestion can enhance separation.
In step 3.1, smaller granulosa cells may accompany oocytes through the cell strainer. However, practical observations indicate that the final collected oocyte sample does not contain granulosa cells. Except in cases where incomplete digestion resulted in the presence of residual granulosa cells in some oocytes, no other Hoechst-labeled granulosa cells were detected in the system. Here, we hypothesized that granulosa cells, being smaller and lighter, might remain suspended in the culture medium and were subsequently removed during the multiple washing steps. The isolation efficiency was also quantified by calculating the proportion of oocytes with granulosa cell attachment and clean oocytes after Hoechst staining. Clean oocyte rates of up to 98% were observed in samples containing 1000 oocytes.
Despite its many advantages, the proposed approach also had some noteworthy limitations. Firstly, this method was validated for DNA or RNA extraction, sequencing, and immediate detection of intact live cells, but the feasibility and reliability of extending to other assay areas are yet to be validated. Another limitation is that the proposed method can only be applied for isolating stage I oocytes, as the isolation efficiency for other stages is unsatisfactory. Therefore, additional efforts are required to improve the efficiency and applicability of the proposed method. Additionally, future studies should focus on exploring whether the proposed method can be applied to efficiently isolate oocytes from other stages, thereby broadening its applicability.
Disclosures
The authors have nothing to disclose.
Acknowledgements
This work was supported by the National Natural Science Foundation of China (32170813 and 31871449) and Science and Technology Department of Sichuan (2024NSFSC0651), and 1·3·5 project for disciplines of excellence–Clinical Research Fund, West China Hospital, Sichuan University (2024HXFH035). The authors would like to thank Zhao Wang and Yanqiu Gao of the Laboratory of Pediatric Surgery for breeding of zebrafish related to this work. The authors would also like to thank all the reviewers who participated in the review, as well as MJEditor (www.mjeditor.com) for providing English editing services during the preparation of this manuscript.
Materials
| Kinger's cell dissociation solution | PlantChemMed | PC-33689 | Kinger's cell dissociation solution can be stored stably at -20 °C after packaging and can be used after thawing at low temperature (4 °C). It can be used directly for dissociating zebrafish ovaries. The optimal temperature is 28.5 °C, for approximately 2-3 hours. The duration can be adjusted according to the specific dissociation conditions, either shortened or extended (https://www.plantchemmed.com/chanpin?productNo=PC-33689). |
| Cell strainers (100 μm ) | Falcon | 352360 | |
| Fluorescence microscope | Zeiss | Axio Zoom.V16 | |
| Forceps | Dumont | #5 | |
| Glass capillary needle | / | / | Blunted by burning with lighter |
| Hoechst | Yesen | 40732ES03 | |
| Low adsorption pipette tips (10 μl ) | Labsellect | T-0010-LR-R-S | |
| Leibovitz’s L-15 medium medium (with L-glutamine) | Hyclone | SH30525.01 | |
| Ice bucket | / | / | Ice-cold water is used to euthanize zebrafish |
| Incubator | WIGGENS | WH-01 | |
| Juvenile fish | / | / | 5–6 weeks post-fertilization, standard length [SL] of 10–15 mm |
| Plastic dish (35 mm ) | SORFA | 230101 | |
| Stereomicroscope | Motic | SMZ-161 | |
| Tissue Culture Plate (6-wells) | SORFA | 0110006 | |
| Vannas spring scissors | Fine Science Toosl | #15000-00 |
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