Spindle shape and volume differ in high- and low-quality metaphase II oocytes

in Reproduction
Authors:
Monika Fluks Department of Embryology, Institute of Developmental Biology and Biomedical Sciences, Faculty of Biology, University of Warsaw, Warsaw, Poland

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Robert Milewski Department of Biostatistics and Medical Informatics, Medical University of Bialystok, Białystok, Poland

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Szymon Tamborski Institute of Physics, Faculty of Physics, Astronomy, and Informatics, Nicolaus Copernicus University in Torun, Toruń, Poland

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Maciej Szkulmowski Institute of Physics, Faculty of Physics, Astronomy, and Informatics, Nicolaus Copernicus University in Torun, Toruń, Poland

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Anna Ajduk Department of Embryology, Institute of Developmental Biology and Biomedical Sciences, Faculty of Biology, University of Warsaw, Warsaw, Poland

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https://orcid.org/0000-0002-7262-1370

Correspondence should be addressed to A Ajduk; Email: a.ajduk@uw.edu.pl
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In brief

Optical coherence microscopy non-invasively visualizes metaphase II spindles allowing for quantitative analysis of their volume and shape, which may prove useful in the assessment of the oocyte quality. Using a mouse model, we showed also that analysis of spindle length combined with morphokinetics improves the evaluation of the resulting embryos.

Abstract

The proper development of embryos strongly depends on the quality of oocytes, so the evaluation of oocytes may be a useful initial step in IVF procedures. Additionally, it enables embryologists to make more informed decisions regarding the treatments chosen for the patients and better manage patients’ expectations. Optical coherence microscopy (OCM) allows for non-invasive 3D visualization of intracellular structures, such as spindles or nuclei, which have been linked to the success of embryonic development. Here, we applied a mouse model to examine whether OCM imaging could be used in the quality assessment of metaphase II (MII) oocytes. We showed that quantitative parameters describing the shape and volume of the MII spindle were associated with the quality of the resulting embryos, including the likelihood of blastocyst formation and the embryos’ ability to differentiate the trophectoderm and primitive endoderm, but not the epiblast. We also created a multivariate linear regression model, combining OCM-based quantification of MII spindles with morphokinetic analysis of the embryos, that allowed for improved evaluation of the embryo quality. Finally, we proved that OCM does not interfere with the viability of the scanned cells, at least during the preimplantation development. Therefore, we believe that OCM-based quantitative assessment of MII spindles can improve the oocyte and embryo selection in IVF procedures.

Abstract

In brief

Optical coherence microscopy non-invasively visualizes metaphase II spindles allowing for quantitative analysis of their volume and shape, which may prove useful in the assessment of the oocyte quality. Using a mouse model, we showed also that analysis of spindle length combined with morphokinetics improves the evaluation of the resulting embryos.

Abstract

The proper development of embryos strongly depends on the quality of oocytes, so the evaluation of oocytes may be a useful initial step in IVF procedures. Additionally, it enables embryologists to make more informed decisions regarding the treatments chosen for the patients and better manage patients’ expectations. Optical coherence microscopy (OCM) allows for non-invasive 3D visualization of intracellular structures, such as spindles or nuclei, which have been linked to the success of embryonic development. Here, we applied a mouse model to examine whether OCM imaging could be used in the quality assessment of metaphase II (MII) oocytes. We showed that quantitative parameters describing the shape and volume of the MII spindle were associated with the quality of the resulting embryos, including the likelihood of blastocyst formation and the embryos’ ability to differentiate the trophectoderm and primitive endoderm, but not the epiblast. We also created a multivariate linear regression model, combining OCM-based quantification of MII spindles with morphokinetic analysis of the embryos, that allowed for improved evaluation of the embryo quality. Finally, we proved that OCM does not interfere with the viability of the scanned cells, at least during the preimplantation development. Therefore, we believe that OCM-based quantitative assessment of MII spindles can improve the oocyte and embryo selection in IVF procedures.

Introduction

Good-quality oocytes are requisite for the successful development of future embryos. Thus, reliable, and noninvasive assessment of their competence may be a valuable addition to the IVF procedures. Such assessment allows for selecting the best oocytes for fertilization, which may be crucial if the number of oocytes that can be fertilized is limited by law (e.g. in Poland only six eggs can be fertilized at once unless the patient is over 35 years old or has other medical conditions that would justify lifting the limit). Additionally, it may enable embryologists to optimize IVF treatment (including oocyte cryobanking option) for individual patients and manage patients’ expectations. Importantly, it may also provide additional information that could be useful in evaluating the quality of the resulting embryos.

Currently, visual assessment of morphology is still the standard method of metaphase II (MII) oocyte selection before IVF (Ebner et al. 2003, Rienzi et al. 2011). A variety of morphological characteristics of oocytes, such as zona pellucida thickness (Bertrand et al. 1995), cytoplasmic granularity (Kahraman et al. 2000), perivitelline space size (Xia 1997), and oocyte shape (Ebner et al. 2007), have been reported to correspond to the embryonic ability to develop and implant. Additionally, spindle shape and zona pellucida birefringence, other biomarkers of oocyte quality, can be assessed with polarized light microscopy (Montag et al. 2011). However, oocyte morphology visualized in transmitted light has been shown to have little or no correlation with the IVF success rate (Nikiforov et al. 2022). Moreover, morphology assessment is prone to intra- and interobserver bias (Paternot et al. 2009, Bormann et al. 2020). Therefore, quantifiable measurements of subcellular structures could provide a more reliable alternative.

Until recently, imaging, enabling quantitative analysis of the oocyte structures, has usually involved fluorescence microscopy and thus required fluorescent dyes or tags. Such an approach leads to the photodamage of cellular components (Magidson & Khodjakov 2013), so it cannot be used with oocytes or embryos intended for reproductive purposes. However, novel, noninvasive imaging methods, such as harmonic generation microscopy (Hsieh et al. 2008, Thayil et al. 2011) and optical coherence microscopy (OCM) (Xiao et al. 2012, Zheng et al. 2012, Raghunathan et al. 2016, Karnowski et al. 2017, Ajduk & Szkulmowski 2019, Fluks et al. 2022) have been recently developed. OCM is derived from optical coherence tomography, which is widely applicable in medicine as the primary method for tissue 3D structural and functional imaging in vivo. OCM is capable of 3D imaging at a uniform resolution on a micrometer scale in all spatial directions. Moreover, its invasiveness is negligible, as it utilizes scattered light intensity as the contrasting mechanism and therefore requires no additional staining. We have previously demonstrated that our optimized OCM protocols can shorten the imaging time, decrease the required light intensity, and reduce noise (Karnowski et al. 2017). We have also shown that OCM can be a useful tool in the quality assessment of immature oocytes (Fluks et al. 2022).

In this study, we applied a mouse model to examine whether OCM imaging could be used in the evaluation of MII oocytes’ quality. We focused on visualization and volumetric analysis of MII spindles. We were particularly interested in spindle dimensions, such as length, width, and volume, as well as the distance between the spindle and the oocyte cortex. The proper structure of the meiotic spindle is crucial for the accuracy of chromosome segregation (Bennabi et al. 2016, Thomas et al. 2021) and an appropriately short distance between the MII spindle and the cortex is fundamental for establishing and maintaining asymmetry in oocytes (reviewed in Ajduk et al. 2013). Together, they ensure the correct progression of fertilization and second meiotic division, requisite for the further development of embryos derived from MII oocytes. The OCM-derived parameters describing nuclear apparatus in MII oocytes were correlated with the ability of the resulting embryos to complete preimplantation development. Additionally, we investigated whether combining OCM-derived quantifications of nuclear apparatus with morphokinetic analysis of the embryos improves the evaluation of embryo quality. Finally, we tested whether OCM is safe for the imaged cells.

Materials and methods

Animals

Mice were maintained in the animal facility of the Faculty of Biology, University of Warsaw at 14 h light:10 h darkness cycle and provided with food and water ad libitum. Animals were sacrificed by cervical dislocation. All animal experiments were approved by the Local Ethical Committee for Experimentation on Animals No. 1 in Warsaw, Poland (approval no. 698/2018) and were performed in compliance with the national regulations (Act on the Protection of Animals Used for Scientific and Educational Purposes from January 15, 2015) and reported according to the ARRIVE guidelines (https://arriveguidelines.org).

Isolation of oocytes and zygotes

MII oocytes were obtained from 2.5- to 4.5-month-old F1 (C57Bl6/Tar × CBA/Tar) mice primed with 7.5 International Unit (IU) of pregnant mare serum gonadotropin (PMSG) (BioVendor, Brno, Czech Republic) followed 48 h later by 10 IU of human chorionic gonadotropin (hCG, Intervet, Warsaw, Poland). Oocytes were released from the oviducts 14 h after hCG into hyaluronidase solution (Merck, 150 IU/mL in PBS). After separating from cumulus cells by gentle pipetting, oocytes were transferred to M2 medium (M16 medium buffered with HEPES (Fulton & Whittingham 1978)).

Germinal vesicle (GV) oocytes were obtained from 1- to 2.5-month-old and 18- to 20-month-old F1 mice primed with 7.5 IU PMSG. GV oocytes were released from ovarian antral follicles into M2 medium 48 h after PMSG injection. Oocytes were separated from the cumulus cells by gentle pipetting. In vitro maturation was performed for 16 h in M16 medium at 37.5°C and in 5% CO2 in the air to obtain young and maternally aged MII oocytes.

Parthenogenetic activation

Parthenogenetic activation of MII oocytes was performed for 6 h in M2 medium without Ca2+ and Mg2+ supplemented with 10 mM SrCl2 (Merck; to induce Ca2+ oscillations in the oocytes) and 5 μg/mL cytochalasin D (Merck; to inhibit the second polar body extrusion and induce diploidization) in 37.5°C and in 5% CO2 in the air.

Time-lapse imaging and morphokinetic analysis

One-cell embryos were placed in 16-well dishes (Vitrolife, Gothenburg, Sweden) with 30 μL of KSOM medium (Merck) and subjected for 5 days to time-lapse imaging (every 10 min) in the PrimoVision imaging system enclosed in a standard embryo culture incubator. Images acquired by PrimoVision were analyzed in FIJI software (Schindelin et al. 2012) and morphokinetic parameters were calculated as described before (Milewski et al. 2018). The morphokinetic parameters included: (1) tNEBD – timepoint of the disappearance of pronuclei in the zygote; (2) t2 to t4 – timepoints when the embryo reaches a certain number of cells, i.e. two cells for t2, three cells for t3, etc.; (3) tM and tSB – timepoint of complete compaction and beginning of cavitation, respectively; (4) m1 – duration of the first embryonic M-phase, i.e. the period between the disappearance of pronuclei and the two-cell stage; (5) cc2a,b – duration of the cell cycle for two-cell stage blastomeres, the faster and the slower one, respectively; and (6) s2 – synchronicity of the second round of cleavage divisions (from two- to four-cell stage). The beginning of incubation with SrCl2 served as the starting point for the morphokinetic calculations.

Optical coherence microscopy setup

OCM imaging was performed with a custom-made fiber-based OCM imaging system combined with an inverted microscope (Eclipse Ti-E, Nikon) as described previously (Karnowski et al. 2017). The system was equipped with a broadband supercontinuum light source (SuperK EXTREME EXW-4 with SuperK Split spectral splitter, NKT Photonics A/S) producing a spectrum as broad as 115 nm (measured at the level of −3 dB intensity drop) and centered at 800 nm, which resulted in axial resolution of 1.9 µm (in-depth, Z axis). The OCM interferometer sample arm shared an imaging objective (Nikon Plan Fluor 20×, NA = 0.5) with an inverted microscope system, which provided 1.9 µm transverse resolution (lateral, XY axes). The average optical power of the focused probing beam at the sample plane was 1.2 mW. The light spectra modulated with interference fringes were captured with a rate of 140 kHz using a custom-designed spectrometer equipped with the 2048 pixels line-scan CMOS camera (Basler Sprint spL2048-140 km, Basler AG). The spectra were digitized with a high-bandwidth framegrabber (PCIe-1433, National Instruments). The sensitivity of the instrument was 89 dB (measured at a typical 20 µs exposure time). The imaging range in depth was 0.9 mm. The OCM probing beam scanned a sample in both lateral directions with the use of a pair of galvanometric scanners (8320 K, Cambridge Technology Inc., Bedford, MA, USA) driven by an analog I/O card (PCI-6733, National Instruments). The standard OCM data preprocessing pipeline was performed on the spectra acquired for every point on the XY plane and resulted in a tomogram line along the Z axis. It consisted of fixed pattern noise removal, resampling to the wavenumber domain, residual dispersion correction, spectrum shaping, and Fourier transformation (Szkulmowski et al. 2005). The measurements were controlled by custom-designed software (C++/C#) which assured precise synchronization between all components of the system.

OCM imaging and 3D rendering of spindles

Oocytes were transferred to a glass bottom dish filled with M2 medium. The glass bottom was coated with a thin (~100–150 μm) layer of agar (1% solution in 0.9% NaCl, Bacto agar, BD) to avoid the unwanted, strong OCM signal from the glass surface. The medium was covered with a layer of mineral oil (BioXtra, Merck) to prevent evaporation. The images were acquired as described previously (Fluks et al. 2022): fifteen 3D volumes were acquired in five series 12 s apart, each composed of three volumes 3 s apart, and then averaged. Each volume was composed of 250 OCM lines in each lateral direction. Images obtained this way were subjected to 3D subcellular structure rendering in the Imaris software (Bitplane, Oxford Instruments). Meiotic spindles in MII oocytes were visualized using the ‘Surfaces’ tool with a surface area detail level set to 1 μm and an absolute intensity threshold set to 150. In the case of touching structures, the ‘Split touching objects’ function was enabled, and the seeding points’ diameter was set to 4 μm, then blocks constituting the desirable structures were merged.

When the quality of OCM-scanned and control MII oocytes was compared, both groups of oocytes were kept in the same conditions during the time required for OCM scanning: in M2 medium under mineral oil, in 37.5°C, and in the atmospheric concentration of CO2. The only difference between those groups was that one was subjected to an OCM scan, and the other was not.

Measurements of mitochondrial activity and amount of reactive oxygen species

Oocytes were incubated in either 5 μM CellROX Orange (ThermoFisher Scientific), a fluorescent indicator of reactive oxygen species (ROS), or 2 μM JC-1 (ThermoFisher Scientific), a cationic indicator of mitochondrial activity, which at low concentrations is monomeric and emits in the green spectrum, while at high concentrations forms aggregates and shifts to red. The loading time for both dyes was 30 minutes at 37.5°C. Oocytes were imaged using a fluorescence microscope (Zeiss Axiovert) equipped with AxioCam HRm camera. For ROS imaging, single-plane images of the equatorial region were taken. Oocytes were illuminated with light passing through a 538–562 nm excitation filter, and the emitted light was collected with a 570–640 nm emission filter (exposure time 100 ms, 4 × 4 binning). ROS concentration was assessed by measuring the mean intensity of CellROX Orange fluorescence in a single equatorial plane. In each experiment, the fluorescence intensity in OCM-scanned oocytes (F) was normalized with the mean fluorescence intensity in control, unscanned oocytes dyed and imaged simultaneously (FCTRL). To visualize mitochondria, stacks of ten images along the oocyte Z-axis (5.5 μm apart) were taken using light passing through 450–490 nm and 528–562 nm excitation filters and collected using 500–550 nm and 570–640 nm emission filters (exposition time of 20 ms for green and 30 ms for red channels, 4 × 4 binning). Mitochondrial activity was assessed as a ratio of red to green JC-1 fluorescence measured for the sum of all ten stack images. Fluorescence intensity measurements were done using FIJI software (Schindelin et al. 2012).

Immunofluorescence staining and image analysis

Oocytes or embryos obtained from the activated MII oocytes were fixed individually in 4% PFA (Merck, 30 min, RT), permeabilized with 0.5% Triton X-100 (Merck, 30 min, RT), and blocked with 3% BSA (overnight in 4°C). After blocking, oocytes were incubated for 24 h in 4°C with rabbit monoclonal anti-phospho-histone H2A.X (Ser139) antibody (Cell Signaling Technology, cat. no. 9781; 1:200) and embryos – with rabbit polyclonal anti-NANOG (ReproCELL, cat. no. RCAB002P-F, 1:100; for epiblast (EPI) cells), polyclonal goat anti-SOX17 (R&D Systems, AF1924, 1:100; for primitive endoderm (PE) cells), and mouse monoclonal anti-CDX2 (BioGenex, cat. no. MU392A-UC, 1:50; for trophectoderm (TE) cells). It was followed by incubation with secondary antibodies: donkey anti-rabbit tagged with Alexa Fluor 647 (Invitrogen, cat. no. A31573, 1:200), donkey anti-goat tagged with Alexa Fluor 488 (Invitrogen, cat. no. A11055, 1:200), donkey anti-mouse tagged with Alexa Fluor 594 (Invitrogen, cat. no. A21203, 1:200) for 2 h in RT. DNA was then stained for 30 min at room temperature or overnight at 4°C with Hoechst 33342 (1 μg/mL; Merck) or chromomycin A3 (0.01 mg/mL; Merck). Samples were imaged with an inverted confocal microscope (Nikon Eclipse Ti2 Re-scan RCM or Zeiss 510 LSM Meta).

Fluorescence intensity of p(Ser139)-H2A.X staining was measured for the chromosomes in single-plane confocal images and standardized with the intensity of DNA fluorescence measured for the same region. Numbers of embryonic cells were counted in Z-stack images by Imaris software.

Statistical analysis

The datasets were tested for normal distribution with Shapiro–Wilk test. Statistical analysis involved χ2 test, parametric Student’s t-test, non-parametric Mann–Whitney U test, and Spearman's rank correlation. We also conducted logistic and linear regression analyses. The differences between groups were considered statistically significant for P < 0.05. Values in the text show mean ± s.d., but graphs display medians and quartiles.

Results

Spindle volume and shape differ in MII oocytes of high and low developmental competence

To investigate whether imaging of spindles in mouse MII oocytes allows for the selection of oocytes with high developmental potential, freshly ovulated oocytes of normal morphology (n = 107, six replicates, 11 mice in total) were individually imaged in OCM, activated parthenogenetically, and cultured in vitro for 5 days to follow their preimplantation development. The embryos were monitored throughout the culture period by time-lapse imaging (Fig. 1A). OCM imaging allowed for accurate 3D visualization of spindles in live MII oocytes and measurements of the spindle dimensions, such as length, width, and volume, as well as the distance between the spindle and the oocyte cortex (Fig. 1B and Supplementary Fig. 1, see section on supplementary materials given at the end of this article).

Figure 1
Figure 1

Relationship between MII spindle dimensions and oocytes’ ability to form blastocysts. (A) Experimental design (details in the text). OCM, optical coherence microscopy; PA, parthenogenetic activation; TLI time-lapse imaging. (B) OCM scans of a representative MII oocyte in three planes: XY, XZ, and YX with a 3D rendering of the spindle and a corresponding bright field (BF) image. Spindle in the OCM scans marked with asterisks. MII oocyte is accompanied by some cumulus cells visible in the bright field (arrows). Scale bar 20 μm. (C) Immunostaining of a representative blastocyst. DNA labeled in white, CDX2 (trophectoderm) – in green, NANOG (epiblast) – in red, and SOX17 (primitive endoderm) – in cyan. Scale bar 20 μm. (D–H) Spindle parameters describing MII oocytes that did or did not form a proper blastocyst after activation (n = 84 and 23, respectively). Violin plots show distribution of the analyzed data; the solid black line indicates median and the dashed black lines first and third quartile values. NS – no statistically significant difference. (I) Percentage of oocytes with different spindle volumes that formed blastocysts after parthenogenetic activation. (J) Percentage of oocytes with different spindle widths that formed blastocysts after parthenogenetic activation.

Citation: Reproduction 167, 4; 10.1530/REP-23-0281

We found that oocytes, which achieved the blastocyst stage after parthenogenetic activation (n = 84), had significantly smaller spindle volume (1850.3 ± 608.3 µm3 vs 2334.9 ± 540.9 µm3; P < 0.001) than those failing to form a proper blastocyst (embryos that either fail to cavitate or initiated cavitation but then decompact or did not form a single cavity or inner cell mass (ICM); n = 23; Supplementary Fig. 2). There was no statistical difference in other analyzed parameters (length: 28.3 ± 3.7 µm vs 28.9 ± 4.4 µm; length/width ratio: 3.0 ± 0.9 vs 2.8 ± 0.8; distance to the cortex: 12.1 ± 1.8 µm vs 12.6 ± 1.9 µm), although the difference in spindle width almost reached the statistical significance (9.9 ± 1.9 µm vs 11.0 ± 2.1 µm; P = 0.05) (Fig. 1D, E, F, G, H and Supplementary Table 1). Univariate logistic regression analysis confirmed these findings and indicated that an increase in spindle volume or width (within the analyzed value range) decreases the odds of blastocyst formation (Table 1). Accordingly, oocytes with smaller spindles (volume ≤ 1750 µm3) formed blastocysts almost twice more often (95.1%, 39/41) than blastocysts with larger spindles (volume ≥2500 µm3; 52.2%, 12/23; P < 0.001) (Fig. 1I). Analogically, oocytes with narrower spindles (width <11 µm) formed blastocysts more often (84.2%, 64/76) than oocytes with broader spindles (≥11 µm; 64.5%, 20/31; P < 0.05) (Fig. 1J), although here the difference was less pronounced.

Table 1

Univariate logistic regression analysis linking MII spindle-related parameters with the ability of the resulting embryos to achieve the blastocyst stage. Only statistically significant relationships are listed.

Analyzed parameters Embryos, n Odds ratio* 95% CI P
MII spindle volume 107 0.999 0.998–1.000 0.002
MII spindle width 107 0.734 0.571–0.945 0.016

*A measure quantifying the association between two variables. For a continuous predictor (like the analyzed parameters listed in the table), the odds ratio greater than 1 indicates that the outcome (i.e. achieving the blastocyst stage by the embryos) is more likely to occur as the predictor increases. The odds ratio lower than 1 indicates that the event is less likely to occur as the predictor increases.

Moreover, spindle volume, width, and distance of the spindle to the cortex, correlated negatively (P < 0.05 to P < 0.001), whereas the ratio of spindle length to width correlated positively (P < 0.001) with the total number of cells in the 5-day-old embryos derived from the oocytes (Supplementary Table 2). This observation was confirmed by univariate linear regression analysis (P < 0.05; Table 2).

Table 2

Univariate linear regression analysis linking MII spindle-related parameters with the number of cells in the resulting 5-day-old embryos.

Spindle parameters Embryos, n Coefficient* 95% CI P R2
Cells in 5-day-old embryos
 Volume 93 −0.027 −0.040 −0.013 <0.001 0.143
 Width 93 −8.05 −12.26 −3.84 <0.001 0.137
 Length/width 93 18.47 8.49 28.46 <0.001 0.129
 Spindle–cortex distance 93 −5.13 −10.19 −0.07 0.047 0.043
Cells in 5-day-old blastocysts
 Length 73 2.38 0.78 3.98 0.004 0.110
 Width 73 −4.73 −7.74 −1.72 0.003 0.122
 Length/width 73 12.59 6.28 18.90 <0.001 0.183
TE cells in 5-day-old blastocysts
 Volume 71 −0.008 −0.016 –0.0002 0.045 0.057
 Length 71 1.45 0.09 2.80 0.037 0.061
 Width 71 −3.75 −6.25 −1.24 0.004 0.114
 Length/width 71 9.43 4.15 14.71 0.001 0.156
ICM cells in 5-day-old blastocysts
 Length 71 0.51 0.09 0.93 0.017 0.080
 Length/width 71 1.80 0.07 3.52 0.041 0.059
PE cells in 5-day-old blastocysts
 Length 71 0.42 0.13 0.72 0.006 0.106
 Length/width 71 1.26 0.02 2.50 0.046 0.057

Only statistically significant relationships are listed here.

*Coefficient describes the mean change in the value of the dependent variable (e.g. number of cells in 5-day-old embryos) corresponding to the unit change in the independent variable (i.e. one of the spindle parameters listed in the table); All stages included.

Next, we examined the relationship between the spindle dimensions and the quality of blastocysts obtained from the activated oocytes after 5-day-long culture. The blastocyst quality was assessed as the number of cells (total and in the first embryonic cell lineages) (Milewski et al. 2018). To this end, 5-day-old blastocysts were fixed and immunostained for NANOG (a marker of EPI), SOX17 (a marker of PE), and CDX2 (a marker of TE) (Fig. 1C). Our analysis indicated that the spindle length and the spindle length/width ratio correlated positively with the total number of cells in blastocysts, the number of cells in TE (length/width ratio only), the ICM (i.e. EPI and PE cells taken together), and PE (P < 0.05 to P < 0.01; Supplementary Table 2). Additionally, spindle width correlated negatively with the number of all cells and TE cells in the blastocysts (P < 0.05; Supplementary Table 2). Again, univariate linear regression analysis supported these results (P < 0.05 to P < 0.001; Table 2). There were no significant relationships between the spindle dimensions and the number of EPI cells in the blastocysts.

Spindles from young oocytes are more compact than spindles from maternally aged oocytes

Oocytes from aged females are considered to possess lower developmental competence than oocytes from young females (reviewed in Cimadomo et al. 2018). Therefore, we decided to compare the spindle volume and shape in these two groups of oocytes to verify our earlier observations regarding MII spindles in oocytes of high and low developmental potential. In this experiment in vitro matured oocytes were used, as old females respond to superovulation very poorly. The experiment was conducted in three replicates (five females in total) for young and five replicates (28 females in total) for old mice.

First, we noticed that spindles in in vitro matured MII young oocytes (n = 45) differ in size and shape from their ovulated counterparts (n = 107) (Supplementary Fig. 3). They were longer (30.3 ± 4.6 µm vs 28.4 ± 3.8 µm; P < 0.01), but narrower (9.1 ± 2.1 µm vs 10.1 ± 2.0 µm; P < 0.01), which resulted in higher length to width ratio (3.6 ± 1.3 vs 3.0 ± 0.8; P < 0.01) and smaller volume (1371.4 ± 518.7 µm3 vs 1954.4 ± 624.8 µm3; P < 0.0001). The spindles in in vitro matured oocytes were also located closer to the cortex (11.0 ± 4.2 µm vs 12.2 ± 1.8 µm; P < 0.0001), which was most likely caused, at least partially, by their smaller width (we measured the spindle–cortex distance starting from the centre of the spindle).

Next, we compared spindles in in vitro matured oocytes from young and aged females. Similar to what has been observed for oocytes able and unable to form a proper blastocyst, spindles in young MII oocytes (n = 45) had significantly smaller volumes as compared to their aged counterparts (n = 44; 1371.4 ± 518.7 µm3 vs 2217.0 ± 760.2 µm3; P < 0.0001). They were also narrower (9.1 ± 2.1 µm vs 11.1 ± 1.2 µm; P < 0.0001). However, spindles in young oocytes were slightly (but significantly) shorter than the spindles in oocytes from aged females (30.3 ± 4.6 µm vs 33.2 ± 4.6 µm; P < 0.01). The ratio of length to width also differed between spindles from young and old oocytes (3.6 ± 1.3 vs 3.0 ± 0.5; P < 0.05), whereas the spindle–cortex distance was comparable (11.0 ± 4.2 vs 10.7 ± 3.1; P > 0.05) (Fig. 2A, B, C, D, and E).

Figure 2
Figure 2

Dimensions of MII spindles in oocytes from young and aged mouse females. (A–E) Parameters describing spindle volume, shape, and location in oocytes from young and aged mouse females. Violin plots show distribution of the analyzed data; the solid black line indicates median and the dashed black lines first and third quartile values. NS, no statistically significant difference.

Citation: Reproduction 167, 4; 10.1530/REP-23-0281

Combining OCM-derived information on spindle dimensions with morphokinetic data improves the quality assessment of MII oocytes

As we showed above, spindle volume and shape are clearly correlated with the developmental competence of MII oocytes. Although it would be difficult to predict the quality of the resulting embryos based solely on these parameters, they may be valuable additions to other methods of embryo evaluation. Here, we looked for multivariate regression models including both the morphokinetics and the OCM-derived MII spindle parameters. We used the data obtained in the first experiment, when MII oocytes were subjected to an OCM scan, then individually activated parthenogenetically, filmed for 5 days, and finally fixed and stained for the markers of the first embryonic cell lineages (Fig. 1A and C). Time-lapse imaging allowed us to calculate morphokinetic parameters, such as rates of the subsequent cleavage divisions, compaction, and cavitation. The spindle parameters correlated only with certain morphokinetic parameters (Supplementary Table 3), which facilitated the creation of a multivariate regression model.

Regression models created for single spindle-related parameters explained only up to approx. 18% of the variability in the analyzed outcome (according to R2 values; Table 2). However, a multivariate linear regression model combining spindle length with three morphokinetic parameters (time between activation and three-cell stage (t3), length of the cell cycle in the slower blastomere of two-cell stage embryos (cc2b), and time between activation and cavitation (tSB)) explained almost 57% of the variability in the total number of cells in 5-day-old embryos (adjusted R 2 = 0.565) (Table 3). Regression models based on the same set of parameters or on the set with a single change (cc2a, length of the cell cycle in the faster blastomere of two-cell stage embryos, instead of cc2b) explain also almost 50% of the variability in the total cell number in blastocysts (adjusted R 2 = 0.490 and 0.495, respectively) (Table 3). All these multivariate models explained higher percentage of the variability in the outcome parameter than any univariate linear regression model based solely on single morphokinetic parameters (Supplementary Table 4). In all these models, the longer the spindle is, the more cells the embryos have, which accords with the univariate regression models involving the spindle length (Table 2). As expected, the opposite relationship links tSB with the number of cells in the 5-day-old embryos: a delay in cavitation is associated with the lower cell number. Interestingly, the timings related to cleavage divisions: t3 and cc2b (or cc2a) seem to affect the embryonic cell number in the opposite way: shortening of the former and prolongation of the latter facilitate the number of cells in 5-day-old embryos.

Table 3

Multivariate linear regression models linking selected spindle-related and morphokinetic parameters with the number of cells in 5-day-old embryos.

Parameters Coefficient§ 95% CI P Adj R2*
Cells in 5-day-old embryos 0.565
 Spindle length 1.17 –0.15 2.49 0.081
t3 −1.38 −2.94 0.19 0.084
cc2b 2.54

−0.11
5.18 0.060
tSB −2.90 −3.70 −2.11 <0.001
Cells in 5-day-old blastocysts 0.490
 Spindle length 1.28 0.02 2.55 0.046
t3 −1.56 −2.95 −0.16 0.029
cc2b 1.71

−0.77
4.19 0.173
tSB −2.30 −3.22 −1.39 <0.001
Cells in 5-day-old blastocysts 0.495
 Spindle length 1.24 −0.01 2.49 0.052
t3 −2.37 −3.99 −0.75 0.005
cc2a 1.47 −0.34 3.28 0.110
tSB −1.99 −2.66 −1.32 <0.001

t3 – time between activation and three-cell stage; cc2a, cc2b length of the cell cycle in the faster and slower blastomere of two-cell stage embryos; tSB – time between activation and cavitation.

§The coefficient in a regression equation that describes the mean change in the value of the dependent variable (e.g. no. of cells in 5-day-old embryos) corresponding to the unit change in the independent variable (e.g. spindle length or one of the morphokinetic parameters) while values of other variables remain fixed; *A statistical measure determining the proportion of variance in the dependent variable (e.g. no. of cells in 5-day-old embryos) that can be explained by the multivariate linear regression model and taking into account the number of independent variables included in the model; All stages included.

Adj, adjusted.

OCM scanning does not diminish the quality of MII oocytes

To become recognized as a potentially useful tool for oocyte quality assessment, OCM cannot negatively impact the quality of the visualized cells. As imaging is often linked to photoinduced oxidative stress in cells (Pomeroy & Reed 2012), we examined whether OCM influences mitochondrial activity and the amount of reactive oxygen species (ROS) in MII oocytes (Fig. 3A). To this end, cells were incubated either with JC-1, a mitochondrial dye sensitive to mitochondria membrane potential or with CellRox Orange, a fluorescent indicator of ROS. Both experiments were conducted in two replicates (three mice in total for JC1 staining and four mice for CellRox Orange staining). We found that oocytes imaged by OCM (n = 40) displayed mitochondrial activity similar to that observed in control cells (n = 42; 1.2 ± 0.7 vs 1.0 ± 0.4, P > 0.05) (Fig. 3B and C). On the other hand, OCM-scanned oocytes (n = 50) had slightly higher levels of ROS compared to the non-imaged control group (n = 66; 1.3 ± 0.5 vs 1.0 ± 0.3; P < 0.001) (Fig. 3D and E).

Figure 3
Figure 3

OCM influence on the quality of MII oocytes. (A) Experimental design (details in the text). OCM, optical coherence microscopy; CTRL, control group. (B) Representative images of OCM-scanned and control (CTRL) oocytes labeled with JC-1, an indicator of mitochondrial activity. Scale bar 100 μm. (C) Ratios of JC-1 red to green fluorescence intensities were quantified in 40 OCM-imaged and 42 control oocytes. A higher ratio indicates more active mitochondria. (D) Representative images of OCM-scanned and control (CTRL) MII oocytes labeled for ROS with CellRox Orange dye. Scale bar 100 μm. (E) The relative intensity of CellRox Orange fluorescence was quantified for 50 OCM-imaged and 66 control oocytes. (F) Representative images of pSer139-H2A.X (white) and DNA (green) staining in OCM-scanned and control (CTRL) oocytes. ETP, oocytes treated with 50 µg/mL etoposide (ETP) for 1 h, serving as a positive control; CTRL NEG, oocytes that were incubated only with secondary antibody, serving as a negative control for the staining. Scale bar 20 µm. (G) The relative intensity of pSer139-H2A.X immunostaining was quantified for 42 OCM-scanned and 45 control oocytes. (C, E, G) Violin plots show distribution of the analyzed data; the solid black line indicates median and the dashed black lines first and third quartile values. NS, no statistically significant difference.

Citation: Reproduction 167, 4; 10.1530/REP-23-0281

As oxidative stress may lead to DNA damage (Cooke et al. 2003), we investigated whether OCM scanning leads to double-stranded DNA breaks, using phosphorylated form of histone H2A.X (pSer139-H2A.X) as a marker (Herchenröther et al. 2023) (Fig. 3A). This experiment was conducted in three replicates (three mice in total). Analysis of immunostaining fluorescence intensity showed that amount of pSer139-H2A.X is the same in OCM-scanned (n = 42) and control oocytes (n = 45; 0.3 ± 0.3 vs 0.3 ± 0.2; P > 0.05) (Fig. 3F and G) suggesting that OCM scanning does not induce DNA damage in cells.

The abovementioned difference in ROS amount did not translate to diversified developmental abilities of the OCM-imaged and control MII oocytes. The experiments testing their developmental potential were conducted in six replicates (11 mice in total) (Fig. 4A). Morphokinetic analysis indicated that embryos obtained from the OCM-imaged MII oocytes developed initially more slowly than the control group: statistically significant delay was noted for tNEBD (time between activation and disappearance of pronuclei in the zygote) and t2 (time between activation and two-cell stage) (Table 4). These differences disappeared, however, at the later cleavage stages. Five-day-old embryos obtained from the activated OCM-scanned and control MII oocytes had similar numbers of cells (93.7 ± 42.9 and 96.6 ± 50.0, respectively; n= 102 and 55, respectively; P > 0.05; Fig. 4B). Both groups reached the blastocyst stage with a similar frequency: 79.3% (96/121) for OCM vs 82.7% (62/75) for controls (P > 0.05). Importantly, immunostaining (Fig. 1C) showed that the blastocysts (n = 82 for OCM and n = 41 for the control group) were of the same quality: they had similar total numbers of cells (111.3 ± 24.1 vs 118.8 ± 21.8; P > 0.05) as well as the numbers of cells belonging to the first embryonic lineages (TE: 90.7 ± 20.5 vs 95.9 ± 17.4; ICM: 20.2 ± 6.7 vs 22.4 ± 6.6; EPI: 8.0 ± 3.4 vs 8.6 ± 4.0; PE: 12.1 ± 4.7 vs 13.8 ± 4.3; P = 0.05 for PE and P > 0.05 for the rest of the cell types) (Fig. 4C, D, E, F, and G).

Figure 4
Figure 4

OCM influence on developmental capabilities of MII oocytes. (A) Experimental design (details in the text); OCM – optical coherence microscopy, CTRL – control group, PA – parthenogenetic activation, TLI – time-lapse imaging. (B) The total number of cells in 5-day-old embryos obtained from OCM-scanned and control (CTRL) MII oocytes (n = 102 and 55, respectively). (C–G) The total number of cells and number of TE, ICM, EPI, and PE cells in 5-day-old blastocysts obtained from OCM-scanned and control (CTRL) MII oocytes (n = 82 and 41, respectively). (B–G) Violin plots show distribution of the analyzed data; the solid black line indicates median and the dashed black lines first and third quartile values. NS, no statistically significant difference.

Citation: Reproduction 167, 4; 10.1530/REP-23-0281

Table 4

Morphokinetic parameters for embryos obtained from OCM-scanned and control MII oocytes.

Morphokinetic parameters OCM scanned Control P
Embryos, n Mean ± s.d. Embryos, n Mean ± s.d.
tNEBD 119 15.93 ± 3.44 75 15.61 ± 3.42 <0.05
m1 (t2tNEBD) 119 1.85 ± 0.40 73 1.79 ± 0.28 NS
t2 119 17.78 ± 3.43 73 17.26 ± 3.32 <0.01
t3 113 38.79 ± 3.58 69 38.48 ± 4.15 NS
t4 113 39.92 ± 3.81 68 39.21 ± 4.21 NS
cc2a (t3t2) 113 21.17 ± 2.88 69 21.45 ± 2.60 NS
cc2b (t4t2) 113 22.30 ± 2.88 68 22.26 ± 2.88 NS
s2 (t4–t3) 113 1.13 ± 2.26 68 0.90 ± 0.93 NS
tM 109 57.01 ± 4.22 67 55.85 ± 4.15 NS
tSB 105 79.23 ± 8.62 66 78.84 ± 9.44 NS

tNEBD – time between activation and the disappearance of pronuclei in the zygote; t2 to t4 – times between activation and two-, three-, and four-cell stages, respectively; tM and tSB– time between activation and complete compaction or beginning of cavitation, respectively; m1 – duration of the first embryonic M-phase; cc2a,b – duration of the cell cycle for two-cell stage blastomeres, the faster and the slower one, respectively; s2 – synchronicity of the second round of cleavage divisions.

Discussion

In the present study, we showed that OCM imaging allows for 3D visualization and volumetric analysis of spindles in MII oocytes. Importantly, our results indicate that in mice the volume and shape of the spindle might be valuable markers of oocyte developmental potential, as oocytes that were able to develop into blastocysts after activation had smaller volumes and shorter widths. Moreover, spindle volume, width, and the spindle–cortex distance correlated negatively with the number of cells in 5-day-old embryos. We also found an association between certain spindle-related parameters (volume, length, width, and length/width ratio) and quality of the blastocysts, assessed by their cell numbers – total and of the first embryonic cell lineages. According to some researchers, human oocytes with longer spindles also have greater developmental capabilities (Rama Raju et al. 2007, Tomari et al. 2011), although not all reports support this observation (Kilani et al. 2009, Molinari et al. 2012, Korkmaz et al. 2014). The importance of spindle compactness has been as well indicated before by studies on human oocytes involving polarized light microscopy. Retardance is considered to be linked to the density of microtubules inside the spindle (Oldenbourg 1999), so it may actually reflect its compactness and it has been showed that human oocytes with higher retardance of the spindle have higher developmental potential (Shen et al. 2006, Rama Raju et al. 2007, Kilani et al. 2009, Tomari et al. 2011). Again, there are also publications that do not confirm this tendency (Molinari et al. 2012, Korkmaz et al. 2014).

We used parthenogenetic activation instead of in vitro fertilization of oocytes to initiate embryonic development, which may be considered as a limitation of this study. We decided for this approach, because in vitro fertilization of individual oocytes proved inefficient in our hands. However, our parthenogenetic activation protocol was chosen very carefully. We used cytochalasin D to ensure diploidization of the parthenogenotes and SrCl2 to induce oscillatory pattern of the activating Ca2+ signal, very similar to the sperm-triggered Ca2+ response (Zhang et al. 2005). Although parthenogenotes are unable to achieve full term development, they are perfectly able to complete preimplantation development and form properly build blastocysts (Niimura & Futatsumata 1999). Therefore, we believe that the relationships we observed here for parthenogenotes are also true for embryos derived from fertilization.

Interestingly, our results indicate that conditions, in which oocyte undergo maturation (in vivo vs in vitro), affect the spindle size and shape. We noticed that spindles in in vitro matured oocytes are longer than in their ovulated counterparts, which accords with the previously published data (Ibáñez et al. 2005). However, in our experiment, as opposed to the study by Ibáñez et al. (2005), spindles in in vitro matured oocytes were also narrower, and in consequence had smaller volumes. This discrepancy may be caused by differences between mouse strains used in experiments, or by the imaging method. Previously, spindles were analyzed mainly by immunostaining of microtubules followed by confocal microscopy. On the other hand, in OCM we identify the region occupied by the spindle based on its optical properties, different than for the rest of the cytoplasm. This difference is caused most likely by the high density of microtubules and microtubule-associated proteins in the spindle, as depolymerization of microtubules with nocodazole led to the disappearance of this structure in OCM scans (Karnowski et al. 2017) Therefore, if microtubules at the edges of the spindle are less dense, e.g. due to suboptimal conditions during in vitro maturation, the visualized region may be narrower.

Importantly, according to our results, spindles in in vitro matured old oocytes are generally larger than in young ones. It accords with the previously published data on mouse and horse oocytes indicating that maternal aging leads to thicker metaphase plates (Pan et al. 2008, Rizzo et al. 2019). Some authors have noticed that in the case of chromosome misalignment, spindles tend also to be elongated (Rizzo et al. 2019); however, others have not observed aging-related elongation of the MII spindles (Pan et al. 2008, Al-Zubaidi et al. 2021). The differences in spindle volume and width between young and old oocytes mimic differences between oocytes able and unable to form a proper blastocyst. This tendency corresponds to the decreased developmental potential of maternally aged oocytes (Cimadomo et al. 2018). Unexpectedly, young, so potentially more developmentally capable, oocytes had shorter spindles than their aged counterparts. Even though it may look like this result contradicts the associations indicated by our regression analysis (longer spindles were associated with a higher total number of cells and higher numbers of TE, ICM, and PE cells in the blastocysts), the issue is not so unequivocal. Our earlier data obtained for the same mouse cross (F1 (C57Bl6/Tar × CBA/Tar)) indicated that although embryos derived from in vitro fertilized old oocytes had some problems with achieving the blastocyst stage (progressing to two-cell stage was the main hurdle), the number of cells in blastocysts was the same (Czajkowska & Ajduk 2023). Therefore, we cannot expect that the parameter related to the cell number in blastocysts is higher in young oocytes than in old ones; instead, it should be similar. However, as spindles in aged oocytes are on average longer, we must consider a possibility that spindle size may be affected in old oocytes by some physiological processes that do not occur in young oocytes.

The molecular mechanism underlying the spindle enlargement observed in our experiments in oocytes of lower quality or originating from aged females is not fully elucidated and requires further research. Based on our current knowledge, we can speculate that it could be related to decreased number of actin filaments, which participate in chromosome segregation (Mogessie & Schuh 2017, Dunkley & Mogessie 2023). Alternatively, it could be also associated with higher levels of acetylated (i.e. stable) microtubules – such an increase in acetylated tubulin accompanied by larger MII spindles has been actually reported for maternally aged mouse oocytes (He et al. 2019). Both the abovementioned mechanisms could affect chromosome segregation and, as a result, decreased the developmental potential of oocytes. We cannot also exclude a possibility that the differences in spindle dimensions observed between young and old oocytes are related to their different susceptibility to in vitro maturation conditions.

To improve the assessment of embryo developmental potential, we combined OCM-based spindle measurements with morphokinetic analysis. We have shown before that combining morphokinetic data with parameters obtained by other imaging methods may significantly strengthen regression models (Milewski et al. 2018). The multivariate regression model described in the current paper associated spindle length and three morphokinetic parameters (time between parthenogenetic activation and three-cell stage (t3), length of the cell cycle in the slower blastomere of two-cell stage embryos (cc2b), and time between activation and cavitation (tSB)) with the total number of cells in 5-day-old embryos and the total number of cells in 5-day-old blastocysts. In the case of the total number of cells in 5-day-old blastocysts, cc2b parameter could be replaced with length of the cell cycle in the faster blastomere of two-cell stage embryos (cc2a). Similarly to our models, increased cavitation time (tSB) has been associated before with poorer human embryo quality, particularly with the higher risk of chromosomal abnormalities in embryos (Campbell et al. 2013, 2014,; Tvrdonova et al. 2021, Shulman et al. 2022).

Interestingly, two other morphokinetic parameters, t3 and cc2b, exert opposing effects in our model, which may reflect the need for a balance between the efficiency and accuracy of cell divisions. It has been shown that both too fast and too slow cleavage divisions reflect the poor developmental potential of the human embryo (Cruz et al. 2012, Meseguer et al. 2012, Chamayou et al. 2013, Milewski et al. 2015). It is likely that timely divisions reflect good quality of the cytoplasmic component, including functional mitochondria and cytoskeleton, and high-quality nuclear apparatus. Divisions that are too fast may result in incorrect segregation of the genetic material and lead to aneuploidy. On the other hand, divisions that are too slow may be a sign of DNA damage or chromosomal aberrations that activate one of the cell cycle checkpoints that halt the cell cycle progression (Ajduk & Zernicka-Goetz 2013, Milewski & Ajduk 2017).

It needs to be emphasized that to become a potentially useful tool for oocyte/embryo quality assessment, OCM cannot negatively impact the quality of the visualized cells. Although OCM scanning increased the amount of ROS in MII oocytes, there was no change in mitochondrial activity, nor did it affect DNA integrity or oocytes’ ability to successfully complete their preimplantation development. The fact that OCM scanning increased the amount of ROS in MII oocytes contradicts our earlier results obtained for oocytes arrested in prophase of the first meiotic division (Fluks et al. 2022). We hypothesized then that infrared radiation used in OCM may have a photobiomodulatory effect in cells, i.e. stimulate mitochondrial activity and ROS turnover (Lubart et al. 2006, Sommer, 2019). However, photobiomodulation depends not only on light wavelength and intensity but also on the physiological (particularly metabolic) state of the cell (Peplow et al. 2010, Hamblin 2018). It has been reported that cellular metabolism depends on the cell cycle stage (Duan & Pagano 2011, Kaplon et al. 2015, Zylstra & Heinemann 2022), which could explain the different susceptibility of MII and prophase I oocytes to OCM-induced photomodulation.

In summary, we demonstrated in a mouse model that OCM provides valuable information on the volume and shape of MII spindles that can be used to determine the developmental capabilities of the embryos derived from the MII oocytes. Additionally, we showed that factors such meiotic maturation conditions (in vivo vs in vitro) and maternal aging should be always considered when spindle dimensions are analyzed, as they tend to affect the spindle structure. Importantly, OCM seems to be safe for oocytes: imaged cells developed to the blastocyst stage as efficiently as their control, unimaged counterparts. Although to fully prove the non-invasiveness of OCM scanning, one should follow also the postimplantation development of the embryos obtained from imaged oocytes, we believe that their unaltered ability to develop into blastocysts containing all three embryonic cell lineages strongly suggests that OCM is indeed non-invasive. Experiments involving animal models are just the first step on a bumpy road leading to introducing a novel technique to clinical practice, but they are crucial, as they often determine the direction of further studies. Models presented here are not meant to be directly transferred to other species; they must be optimized separately for each target species, including humans. We recommend treating our study as a proof of concept, showing that OCM can be used to visualize and measure metaphase spindles and that associations between spindle size or shape and developmental potential of oocytes can be found. Our results indicate that OCM-based quantitative analysis of the MII spindles has the potential to become a valuable addition to oocyte/embryo selection protocols.

Supplementary materials

This is linked to the online version of the paper at https://doi.org/10.1530/REP-23-0281.

Declaration of interest

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

Funding

This work was financed by OPUS grant (2017/27/B/NZ5/00405) from the National Science Centre (Poland) to AA. The cost of Open Access publication was covered by the Society for Biology of Reproduction in Poland.

Data availability statement

The data underlying this article will be shared on reasonable request to the corresponding author.

Author contribution statement

MF planned and conducted the experiments, analyzed the data, and prepared the first version of the manuscript. AA secured the funding, designed the general experimental outline, conducted some of the experiments, analyzed the data, and prepared the final version of the manuscript. RM conducted the regression model analysis. ST and MS provided expertise and assistance with OCM technology. All authors revised the manuscript.

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  • Figure 1

    Relationship between MII spindle dimensions and oocytes’ ability to form blastocysts. (A) Experimental design (details in the text). OCM, optical coherence microscopy; PA, parthenogenetic activation; TLI time-lapse imaging. (B) OCM scans of a representative MII oocyte in three planes: XY, XZ, and YX with a 3D rendering of the spindle and a corresponding bright field (BF) image. Spindle in the OCM scans marked with asterisks. MII oocyte is accompanied by some cumulus cells visible in the bright field (arrows). Scale bar 20 μm. (C) Immunostaining of a representative blastocyst. DNA labeled in white, CDX2 (trophectoderm) – in green, NANOG (epiblast) – in red, and SOX17 (primitive endoderm) – in cyan. Scale bar 20 μm. (D–H) Spindle parameters describing MII oocytes that did or did not form a proper blastocyst after activation (n = 84 and 23, respectively). Violin plots show distribution of the analyzed data; the solid black line indicates median and the dashed black lines first and third quartile values. NS – no statistically significant difference. (I) Percentage of oocytes with different spindle volumes that formed blastocysts after parthenogenetic activation. (J) Percentage of oocytes with different spindle widths that formed blastocysts after parthenogenetic activation.

  • Figure 2

    Dimensions of MII spindles in oocytes from young and aged mouse females. (A–E) Parameters describing spindle volume, shape, and location in oocytes from young and aged mouse females. Violin plots show distribution of the analyzed data; the solid black line indicates median and the dashed black lines first and third quartile values. NS, no statistically significant difference.

  • Figure 3

    OCM influence on the quality of MII oocytes. (A) Experimental design (details in the text). OCM, optical coherence microscopy; CTRL, control group. (B) Representative images of OCM-scanned and control (CTRL) oocytes labeled with JC-1, an indicator of mitochondrial activity. Scale bar 100 μm. (C) Ratios of JC-1 red to green fluorescence intensities were quantified in 40 OCM-imaged and 42 control oocytes. A higher ratio indicates more active mitochondria. (D) Representative images of OCM-scanned and control (CTRL) MII oocytes labeled for ROS with CellRox Orange dye. Scale bar 100 μm. (E) The relative intensity of CellRox Orange fluorescence was quantified for 50 OCM-imaged and 66 control oocytes. (F) Representative images of pSer139-H2A.X (white) and DNA (green) staining in OCM-scanned and control (CTRL) oocytes. ETP, oocytes treated with 50 µg/mL etoposide (ETP) for 1 h, serving as a positive control; CTRL NEG, oocytes that were incubated only with secondary antibody, serving as a negative control for the staining. Scale bar 20 µm. (G) The relative intensity of pSer139-H2A.X immunostaining was quantified for 42 OCM-scanned and 45 control oocytes. (C, E, G) Violin plots show distribution of the analyzed data; the solid black line indicates median and the dashed black lines first and third quartile values. NS, no statistically significant difference.

  • Figure 4

    OCM influence on developmental capabilities of MII oocytes. (A) Experimental design (details in the text); OCM – optical coherence microscopy, CTRL – control group, PA – parthenogenetic activation, TLI – time-lapse imaging. (B) The total number of cells in 5-day-old embryos obtained from OCM-scanned and control (CTRL) MII oocytes (n = 102 and 55, respectively). (C–G) The total number of cells and number of TE, ICM, EPI, and PE cells in 5-day-old blastocysts obtained from OCM-scanned and control (CTRL) MII oocytes (n = 82 and 41, respectively). (B–G) Violin plots show distribution of the analyzed data; the solid black line indicates median and the dashed black lines first and third quartile values. NS, no statistically significant difference.

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