Slow and steady cell shrinkage reduces osmotic stress in bovine and murine oocyte and zygote vitrification

Abstract

STUDY QUESTION

Does the use of a new cryoprotectant agent (CPA) exchange protocol designed to minimize osmotic stress improve oocyte or zygote vitrification by reducing sublethal cryodamage?

SUMMARY ANSWER

The use of a new CPA exchange protocol made possible by automated microfluidics improved oocyte and zygote vitrification with superior morphology as indicated by a smoother cell surface, higher sphericity, higher cytoplasmic lipid retention, less cytoplasmic leakage and higher developmental competence compared with conventional methods.

WHAT IS KNOWN ALREADY

The use of more ‘steps’ of CPA exposure during the vitrification protocol increases cryosurvival and development in the bovine model. However, such an attempt to eliminate osmotic stress is limited by the practicality of performing numerous precise pipetting steps in a short amount of time.

STUDY DESIGN, SIZE, DURATION

Murine meiotically competent germinal vesicle intact oocytes and zygotes were harvested from the antral follicles in ovaries and ampulla, respectively. Bovine ovaries were obtained from a local abattoir at random stages of the estrous cycle. A total of 110 murine oocytes, 802 murine zygotes and 52 bovine oocytes were used in this study.

PARTICIPANTS/MATERIALS, SETTING, METHODS

Microfluidic devices were fabricated using conventional photo- and soft-lithography. CPAs used were 7.5% ethylene glycol (EG) and 7.5% dimethyl sulfoxide (DMSO) for equilibration solution and 15% EG, 15% DMSO and 0.5 M sucrose for vitrification solution. End-point analyses include mathematical modeling using Kedem–Katchalsky equations, morphometrics assessed by conventional and confocal microscopy, cytoplasmic lipid quantification by nile red staining, cytoplasmic leakage quantification by fluorescent dextran intercalation and developmental competence analysis by 96 h embryo culture and blastomere quantification.

MAIN RESULTS AND THE ROLE OF CHANCE

The automated microfluidics protocol decreased the shrinkage rate of the oocyte and zygote by 13.8 times over its manual pipetting alternative. Oocytes and zygotes with a lower shrinkage rate during CPA exposure experienced less osmotic stress resulting in better morphology, higher cell quality and improved developmental competence. This microfluidic procedure resulted in murine zygotes with a significantly smoother cell surface (P < 0.001), more spherical cellular morphology (P < 0.001), increased cytoplasmic lipid retention in vitrified and warmed bovine oocytes (P < 0.01), decreased membrane perforations and cytoplasmic leakage in CPA-exposed murine zygotes (P < 0.05) and improved developmental competence of vitrified and warmed murine zygotes (P < 0.05) than CPA exposure using the current clinically used manual pipetting method.

LIMITATIONS, REASONS FOR CAUTION

It is necessary to design the microfluidic device to be more user-friendly for widespread use.

WIDER IMPLICATIONS OF THE FINDINGS

The theory and approach of eliminating osmotic stress by decreasing shrinkage rate is complementary to the prevalent osmotic stress theory in cryobiology which focuses on a minimum cell volume at which the cells shrink. The auto-microfluidic protocol described here has immediate applications for improving animal and human oocyte, zygote and embryo cryopreservation. On a fundamental level, the clear demonstration that at the same minimum cell volume, cell shrinkage rate affects sublethal damage should be broadly useful for cryobiology.

STUDY FUNDING/COMPETING INTEREST(S)

This project was funded by the National Institutes of Health and the University of Michigan Reproductive Sciences Program. The authors declare no conflicts of interest.

Introduction

The successful cryopreservation of embryos and oocytes has significantly expanded the scope of infertility treatment (Trounson and Mohr, 1983; Kuwayama et al., 2005). A major current concern in cryobiology is the potentially lethal mechanical stresses that cells experience as they are exposed to high osmolarity solutions that promote exchange of intracellular water with cryoprotectant agents (CPAs). Concern for such osmotic stress is particularly acute for the fast-cooling protocol called vitrification, where high concentrations of CPAs are used to prevent formation of lethal intracellular ice crystals (Rall and Fahy, 1985). Compared with mammalian cells with diameters on the order of 10 µm, oocytes and zygotes with diameters on the order of 100 µm have an ∼1000 times larger volume of water that needs to be replaced with CPAs. However, the cell surface area does not increase proportionally, making the required water–CPA exchange through the cell membrane difficult and the cell highly susceptible to osmotic stress. Early studies on osmotic stress suggested the existence of a threshold minimum cell volume where shrinkage becomes sufficient to cause lethal damage to cell membrane integrity (Meryman, 1971; Agca et al., 2000; Mullen et al., 2008). Thus, the common method for oocyte and zygote vitrification involves a 3-step equilibration process where the cells are manually pipetted into subsequently higher levels of permeable CPA concentrations (Smith et al., 2010), allowing the cell to shrink in stages, and sometimes to re-expand, to avoid the critical minimum cell volume. Exposure to impermeable solutes, which are critical for success of vitrification but does not allow for any cell re-expansion, is reserved until the final step to maintain as large a cell volume as possible. These procedures result in shriveled and wrinkled cell morphology.

In bovine, which are more difficult to cryopreserve and more representative of human compared with rodents, it has been shown that a higher number of equilibration steps increases the rate of cryosurvival (Kuwayama et al., 1992) and developmental competence (Otoi et al., 1998). However, despite improved outcome, protocols with a high number of manual equilibration steps cannot be clinically adopted due to their impracticality. The Demirci group has developed a microfluidic device for slow-rate freezing (Song et al., 2009) and recently, the Toner group pioneered the use of microfluidic CPA exchange to overcome the limitations of manual pipetting (Heo et al., 2011). They demonstrated precise control of fluid exchange to maintain a large cell volume during permeable CPA exchange. However, the study did not incorporate impermeable solutes that are highly recommended for embryo and oocyte cryopreservation (Kuleshova et al., 1999). Moreover, the device was not designed for cell removal after CPA exchange, precluding actual cooling and vitrification of the oocytes.

Early studies of osmotic stress focused primarily on minimum cell volume, particularly because cryosurvival was a major concern in cryobiology and the minimum cell volume was experimentally easy to observe. Currently, the issues of cryosurvival have largely been overcome, even for human gametes/zygotes/embryos and the field is focusing efforts on reducing sublethal damage to increase developmental competence and improve the effectiveness of single-embryo transfer (SET). Although previous studies demonstrated that there is a lethal range of minimum cell volume depending on species, cell type and individual cells, the effects of shrinkage rate are unknown. Here, we develop a novel microfluidic CPA exchange device and test the hypothesis that for a given minimum cell volume, sublethal damage during CPA exchange can be reduced by minimizing cell shrinkage rate. We evaluate the impact of different microfluidic CPA exchange procedures on cryosurvival, morphology and cell health as evaluated by lipid content and developmental competence. Our results demonstrate that for a given minimum cell volume and CPA exposure time, minimizing the cell shrinkage rate leads to improved outcomes.

Materials and Methods

Kedem–Katchalsky modeling

The device was designed to minimize osmotic stress, using the Kedem–Katchalsky equations [Equations (1 and 2)] with MatLAB (MathWorks, Natick, MA, USA).

$$J_{\mathrm{v}}=-L_{\mathrm{p}}\text{RT(}\mathrm{\Delta }{C}_{\mathrm{i}}+\sigma \mathrm{\Delta }{C}_{\mathrm{s}})$$ (1)

$$J_{\mathrm{s}}=\omega \text{RT}\mathrm{\Delta }{C}_{\mathrm{s}}+J_{\mathrm{v}}(1-\sigma ){\overline{C}}_{\mathrm{s}}$$ (2)

where J_v is the liquid volume flux through the cell membrane and J_s the solute flux through the cell membrane. The coefficients L_p, σ, and ω are related to the hydraulic conductivity, the solvent/solute interaction factor and the solute permeability, respectively. R and T are the gas constant and temperature. ${\overline{C}}_{\mathrm{s}}$ is the mean solute concentration, and ΔC_i and ΔC_s are the change in impermeable solute concentrations and change in permeable solute concentrations, respectively. The osmolarity contributions of each CPA component in the vitrification solution (VS) are assumed to be additive. L_p is 8.94×10⁻⁷ cm/s/atm, ω_DMSO is 2.54×10⁻⁵ cm/s, ω_EG is 1.0×10⁻⁵ cm/s, t_f is the user-defined duration of CPA exchange and n is the step number of computer analysis. The solvent/solute interaction σ did not have a dramatic effect on model.

Device design and fabrication

Microfluidic devices were made using conventional photolithography and soft lithography using polydimethylsiloxane (PDMS) (Dow Corning, Midland, MI, USA). The device consists of two layers: the microfluidic channel layer (150 µm) that houses the oocytes/zygotes as well as delivers the CPA solutions, and the holding pipette layer (20 µm) that provides the vacuum for the oocytes/zygotes to remain stationary under constant volumetric flow rate but changing CPA concentrations and associated viscosity and fluid density.

The device was designed with a serpentine mixing channel, an oocyte/zygote exposure chamber and a set of height-differential (10 cm) powered suction microchannels. The loading and withdrawal of oocytes or zygotes from the device was achieved through a port drilled by a 200 µm microdrill bit (Kyocera, Kyoto, Japan) into a glass slide (Fisher Scientific, Waltham, MA, USA) on which the PDMS is bonded to provide laminar flow in and out of the device. This, combined with control of vacuum, created an effective method of retrieval with no edges or corners for oocytes/zygotes to be trapped within the device. The device was characterized for efficient mixing and linear CPA exposure using fluorescence quenching and fluorescence microscopy (data not shown) and operated at 700 µl/h total flow rate from a synchronized culture media syringe pump and VS syringe pump.

Oocyte/zygote loading, trapping and withdrawal

Loading of oocyte/zygotes through the 200 µm loading/withdrawal port is achieved by activation of the suction channel powered by a negative height differential (∼10 cm) in the absence of syringe pump flow. The activation and deactivation of the suction channel is operated by a three-way valve at the end of the height differential. The exposure chamber contains 15 holding microchannels (20 by 20 µm) where up to 15 oocytes/zygotes are immobilized via the continued activation of the suction port. The cells are released out of the device through the loading/withdrawal port by the termination of suction port in the presence of syringe pump flow.

Automated and manual vitrification

Automated vitrification was achieved by two programmable syringe pumps (Chemyx, Stafford, TX, USA) capable of providing linear gradients. The VS syringe pump was programmed to provide a linearly increasing flow rate from 0 to 700 µl/h in 10 min. The culture media syringe pump was programmed to provide a linearly decreasing flow rate from 700 to 0 µl/h in 10 min. The solutions from these two syringes are passed through a serpentine fluid-mixer channel to the exposure chamber. In this chamber, cells are exposed at a constant flow rate of 700 µl/h to a gradually increasing CPA concentration starting with 100% culture media (0% CPA) and ending with 100% VS. During operation, the spent fluid escapes via the loading/withdrawal port as well as the suction port. The media that escapes from the loading/withdrawal port is discarded.

Manual vitrification for metaphase II (MII) oocytes and zygotes was performed by fully trained individuals using a previously described technique (Smith and Fioravanti, 2007) with slight modification. Mouse or bovine MII oocytes were transferred to a 20 μl washing solution drop for 1 min and then merged with two 20 μl equilibration solution drops sequentially for 2 min each merging, then oocytes were transferred to equilibration solution for 2–3 min. After equilibration, oocytes were transferred through three 20 μl VS drops for 5, 5 and 10 s and then stayed in the fourth drop of VS until cryopreservation. High security vitrification straws (CryoBio Systems, L'Aigle, France) were used for loading oocytes and the loaded straws were plunged into liquid nitrogen within a 90 s interval. Mouse zygote vitrification procedures were similar to those for oocytes, except for a longer incubation (8 min) in equilibration solution.

Oocyte and zygote harvest

All procedures described here were reviewed and approved by The University Committee on Use and Care of Animals at the University of Michigan and were performed in accordance with the Guiding Principles for the Care and Use of Laboratory Animals.

Bovine oocytes were used because their size and developmental characteristics are similar to human oocytes (Chian et al., 2004). Bovine ovaries were obtained from a local abattoir at random stages of the estrous cycle or pregnancy and transported to the laboratory in warmed phosphate-buffered saline (PBS) containing antibiotics. Cumulus–oocyte complexes were aspirated from follicles of 2–6 mm diameter and matured for 24 h in M199 medium (Sigma-Aldrich, St Louis, MO, USA) at 39°C in 5% CO₂, 5% O₂ and 90% N₂. Mature oocytes were then denuded of cumulus cells before use.

Meiotically competent germinal vesicle intact (GVI) oocytes were collected from antral follicles of 21- to 23-day-old CF-1 female mice (Harlan, Indianapolis, IN, USA) ovaries 44–46 h after 5 IU equine chorionic gonadotrophin (eCG, Sigma-Aldrich) stimulation. Denuded GVI oocytes were triple washed in human tubal fluid-HEPES (HTFH, Irvine Scientific, Santa Ana, CA, USA)/0.3% bovine serum albumin (BSA) and cultured in HTF/0.3% BSA at 37°C in a 5% CO₂ incubator for 18 h to obtain MII oocytes.

Female B6C3F1 mice at 6–8 weeks old were injected with 5 IU eCG (Sigma-Aldrich) for zygote collection. Forty-eight hours later, mice were injected with 5 IU hCG (Sigma-Aldrich). Upon hCG injection, female mice were placed with mature B6C3F1 males of known fertility until sacrificed 18 h later. Zygotes were released from the ampulla, denuded in HTFH (Irvine Scientific)/0.1% hyaluronidase, triple washed in HTFH/0.3% BSA and placed in pre-equilibrated potassium simplex optimized medium (KSOM) + amino acids (AA) (Millipore, Billerica, MA, USA) at 37°C in 5% CO₂ incubator until use.

Microscopy, roughness index and sphericity

Zygotes were analyzed at ×400 magnification for conventional microscopy. The roughness index (RI) was quantified as standard deviation of the gray values within the region of interest (ROI). The ROI was defined as the middle 50% of cell surface, so that edge effects are nullified. The zygotes were oriented such that the polar bodies do not visually obstruct the center of the zygote. Zygotes exposed to CPA by automated or manual pipetting protocols were stained by Hoechst 33342 at 5 μg/ml for 10 min but unwashed to retain cytoplasmic staining. The Hoechst-stained zygotes were analyzed using confocal microscopy. The volume and surface area were then quantified using ImageJ to calculate sphericity.

Warming, embryo culture and developmental competence analysis

Independent of CPA exposure/vitrification treatments, oocyte and zygote warmings were performed manually by fully trained individuals using a previously described technique (Smith and Fioravanti, 2007). Murine zygotes, either no CPA exposure or vitrification (NCEV; control) or vitrified and warmed, were transferred to 50 µl pre-equilibrated KSOM + AA microdrops under oil and cultured at 37°C in 5% CO₂, 5% O₂, 90% N₂ gaseous conditions. Embryo morphologies in each experimental group were assessed at 96 h of culture for blastocyst development using an inverted microscope (×200; Leica DM2RB, Wetzlar, Germany).

Cytoplasmic lipid content

Independent of CPA exposure/vitrification treatment, bovine oocytes were manually warmed and fixed separately on Poly-l-lysine-coated coverslips overnight using 2% paraformaldehyde. Fixed oocyte samples were triple washed with PBS the next day, stained with 100 nM Nile Red (Sigma-Aldrich) for 20 min at room temperature, then washed one time with PBS and stained with Hoechst 33342 (5 μg/ml) for 10 min. After Hoechst staining, oocyte samples were triple washed with PBS and mounted in 90% glycerol with anti-fading reagent. Lipid contents in bovine oocytes were visualized under confocal microscopy and were quantified with ImageJ.

Cytoplasmic leakage

Murine zygotes were exposed to CPA by automated-microfluidics and manual pipetting using VSs and equilibration solutions supplemented with 100 mg/ml 4 kDa fluorescein isothiocyanate (FITC)–dextran (Sigma-Aldrich). Entrance of FITC–dextran into cells is used as an indicator of membrane damage (Miyake et al., 2001). The fluorescence intensity of each zygote was recorded by fluorescence microscopy and normalized to NCEV murine zygotes treated in culture medium with FITC–dextran supplement for 12 min. After CPA exposure, the zygotes were carefully washed in regular VS twice to eliminate extracellular FITC–dextran. NCEV zygotes were also carefully washed in regular culture medium in the same manner. Special care was taken during sample preparation and image capture to have identical sample volume, excitation wavelength, excitation energy and exposure time.

Developmental competence analysis

To count the total number of cells (blastomeres) in blastocysts derived following no treatment (NCEV; control) or automated and manually vitrified and warming, blastocysts were placed on poly-l-lysine-coated coverslips and fixed overnight with 2% paraformaldehyde. Fixed blastocyst samples were triple washed in PBS the next day, stained with Hoechst 33342 (5 μg/ml) for 10 min, triple washed with PBS and then mounted in 90% glycerol with anti-fading reagent. Blastomere numbers were counted under fluorescent microscopy (×400) by two individuals, both blinded to knowledge of treatment and reported values were averages of the counts.

Statistical analysis

All statistical analyses were parametric and conducted first using analysis of variance followed by pair-wise comparisons using unpaired Student's t-test. Statistical significance was determined at the 95% confidence interval and performed using R (Vienna University of Economics and Business, Vienna, Austria). Statistical significance at P < 0.05 is graphically represented by different smaller case letters.

Results

Modeling cell shrinkage with the Kedem–Katchalsky equations

A cell's volumetric change upon exposure to CPA can be accurately modeled (Kedem and Katchalsky, 1958; Pfaff, 1998; Paynter et al., 1999). The Kedem–Katchalsky equations [Equations (1 and 2)] typically use step-wise input functions of CPA concentrations, as this is the experimental limitation of manual pipetting. However, with the use of microfluidics, we expand the experimental possibilities to include continuous temporal gradients of CPA concentrations using ethylene glycol (EG), dimethyl sulfoxide (DMSO) and sucrose (Fig. 1A).

Figure 1.

(A) Kedem–Katchalsky model of zygote shrinkage during CPA exchange protocols. Auto, automated-microfluidic CPA exposure; Manual, manual pipetting CPA exposure; Direct, direct exposure from culture medium to VS known to be detrimental to cryosurvival. All protocols had similar minimum cell volume but significantly different shrinkage patterns. (B) Experimental observation of murine zygote volumes matched that of predictive models. Error bars, standard error. CPA exchange profiles and representative morphology for: (C and H) NCEV, (D and I) commonly used manual pipetting protocols, (E and J) automated microfluidic protocol with gradual addition of all CPA components with smoother and rounder morphology than alternative CPA exposure protocols, (F and K) gradual exposure of permeable CPA components but abrupt exposure to the impermeable component sucrose and (G and L) direct and sudden exposure from culture media to VS. Scale bar: 50 µm. EG, ethylene glycol; DMSO, dimethylsulfoxide.

Direct exposure to VS gives the fastest maximal shrinkage rate of 7.51% volume/s and the highest osmotic shock. The current clinically used manual pipetting protocol causes a shrinkage pattern that is at times shrinking quickly, while at other times static. The sudden introductions of CPA from manual pipetting also produced fast shrinkage rates with a maximum of 4.13% volume/s. Automated microfluidic CPA exchange (auto-microfluidics), which we hypothesize to have the lowest osmotic stress, prevents instances of quick cell shrinkage by eliminating sudden changes in osmolality, producing maximum shrinkage rates of only 0.35% volume/s. If assumed spherical, the maximal surface area strain rate of the cell membrane for auto-microfluidics is 13.8 times and 22.5 times less than the maximal strain rate of manual pipetting and direct exposure, respectively. The cell membrane buckling observed under conditions of fast shrinkage rates is consistent with the known viscoelastic nature of the cell membrane–cytoskeleton complex (Ragoonanan et al., 2010; Fu and Zhang, 2012).

Zygote vitrification affecting morphology and cryosurvival

Murine zygotes with NCEV, alternatively referred to as fresh or non-treated zygotes (Fig. 1C and H), were compared with those subjected to CPA exposure regimes (Fig. 1D–G). The direct exposure (Fig. 1G and L) represents transfer of zygotes directly from culture media to VS, a protocol known to be detrimental to cryosurvival (59.6 ± 3.2%, n = 40). Experimental cell shrinkage profiles (Fig. 1B) agree well with models (Fig. 1A). Despite differences in cell shrinkage profile, murine zygotes from manual pipetting (Fig. 1D and I) and auto-microfluidic (Fig. 1E and J) CPA exposure both exhibit high cryosurvival (100 ± 0%, n = 200 and 100 ± 0%, n = 224, respectively) after vitrification and warming. Murine oocytes also had 100 ± 0% cryosurvival for both manual pipetting (n = 50) and auto-microfluidic (n = 60).

Despite shrinkage to roughly the same final cell volume, there were significant differences in cell morphology depending on the CPA exchange protocol. High magnification images reveal a significantly smoother and more spherical morphology when using a more uniform and gradual shrinkage rate enabled by microfluidic CPA exposure (Fig. 1I and J). Confocal images (see Supplementary data, Video S1) further highlight morphological differences between manual pipetting and auto-microfluidic CPA exchange. Additional morphology-focused CPA exchange studies demonstrate the importance of applying sucrose, the impermeable component of the VS gradually (Fig. 1E and J) for a smooth cell surface in comparison to abrupt sucrose addition (Fig. 1F and K). Finally, direct VS exposures (Supplementary data Video S2 and Fig. 1G and L) demonstrate the importance of avoiding abrupt permeable and impermeable CPA exposures to alleviate the large crater-like deformations on the cell membrane that are also visible in manual pipetting protocols (Fig. 1I). This suggests that inhomogeneous or incomplete mixing that can exist in manual pipetting methods generate gradients of CPA concentrations across the cell where it is exposed to more CPA on one side than the other resulting in the water volume preferentially leaving the cell from the side that is exposed to higher concentrations of CPA to form the crater-like deformations. Zygote surface smoothness can be quantified while in VS under microscopy. Rough surfaces cast dark and light areas on the cell surface. The RI is quantified as the standard deviation of the gray value of the cell. Confocal microscopy is also used to measure cell surface area and volume to calculate sphericity. Zygotes exposed to automated microfluidic CPA exposure (Fig. 1E and J; RI = 23.50 ± 0.69, n = 27; sphericity = 0.956 ± 0.004, n = 27) are significantly (P < 0.001) smoother and more spherical morphology than its manual pipetting counterpart (Fig. 1D and I; RI = 33.72 ± 0.47, n = 26; sphericity = 0.790 ± 0.008, n = 24) (Fig. 2A and B). Gradual EG and DMSO but abrupt sucrose exposure also result in rough surfaces and sphericity that is statistically comparable to manual pipetting which also consists of abrupt exposure to sucrose (RI = 34.32 ± 1.02, n = 20; sphericity = 0.783 ± 0.008, n = 16). Direct exposure of zygotes resulted in smooth surfaces despite an abrupt exposure to all CPAs: EG, DMSO and sucrose (RI = 25.08 ± 1.33, n = 20), but they develop large concave morphology that resulted in the lowest sphericity (sphericity = 0.586 ± 0.017, n = 18). NCEV zygotes had visual occlusions that cast dark regions in microscopy due to significant differences in refractive indexes in its cytoplasmic content. The lack of EG and DMSO, which homogenizes refractive indexes of cytoplasmic content, prevented the direct comparison of NCEV zygotes to other experimental groups, although NCEV zygotes clearly have a smooth surface as observed visually by the human eye and its smooth surface allowed it to have high sphericity (sphericity = 0.965 ± 0.006, n = 20).

Figure 2.

(A) Murine zygotes subjected to automated microfluidic CPA (Auto) exposure had significantly (P < 0.001) smoother cell surface than manual pipetting-based CPA (Manual) exposure. NCEV zygotes had visual occlusions that cast dark regions in conventional microscopy due to the lack of EG and DMSO that prevent direct comparison from other experimental groups. (B) Confocal microscopy reveals that murine zygotes subjected to Auto exposure have significantly higher spherical morphology (P < 0.001) than Manual. Auto exposure maintains its surface smoothness and sphericity at a level statistically comparable to NCEV zygotes. Manual and abrupt sucrose CPA exposure forms surface ripples and some large deformations in cell shape. Direct exposure, with the highest shrinkage rate, have the largest deformations in cell shape that, despite its smooth surface, have the largest impact on sphericity. Sphericity is a dimensionless measure of how spherical a three-dimensional object is. Error bars, standard error. Different letters signify statistical significance P < 0.05 by Student's t-test. RI, roughness index.

Microfluidic device design

A continuous temporal gradient of CPA concentrations is realized by two synchronized programmable syringe pumps connected to the inlets of the microfluidic device with geometry-mediated CPA mixing and exposure embedded into device design (Fig. 3A). Immobilized by holding microchannels within the exposure chamber, the microfluidic device allows for real-time monitoring by microscopy (Fig. 3B). Abnormal deformation in the zona pellucida was observed at ∼18 cmH₂O. As such, a height differential of ∼10 cm was used and the flow velocity was adjusted such that the cells would remain trapped during syringe flow. The size of the cell loading/withdrawal port allow for laminar flow at the outlet that prevented cell entrapment within the device. We demonstrated the reliability and versatility of the automated microfluidic device by vitrifying 462 murine or bovine oocytes and zygotes with 100% recovery as well as the capability of vitrifying up to 15 oocytes/zygotes simultaneously.

Figure 3.

(A) Device schematic. (B) Holding microchannels immobilize cells within the exposure chamber for real-time morphometrics during CPA exchange. CM, culture media; VS, vitrification solution.

Increased bovine cytoplasmic lipid retention

Bovine oocytes from the manual pipetting and auto-microfluidic methods were studied to demonstrate the biological advantages of lowering shrinkage rate. Confocal images of bovine oocytes with Nile Red staining were normalized to NCEV oocytes at 100 ± 0.04% (n = 17) lipid content to characterize post-warming cytoplasmic lipid retention. Vitrified bovine oocytes using manual pipetting CPA exposure have significantly lower cytoplasmic lipid retention 51.8 ± 0.05% (n = 18) than bovine oocytes vitrified using auto-microfluidic CPA exposure 76.6 ± 0.07% (n = 17; P < 0.01; Fig. 4A).

Figure 4.

(A) Vitrification causes a loss of cytoplasmic lipid in bovine oocytes. However, the use of automated-microfluidic (Auto) CPA exposure increases cytoplasmic lipid retention compared with manual pipetting (Manual) CPA exposure (P < 0.01). Inlets demonstrate a difference in lipid droplet size and distribution. (B) A lower shrinkage rate from Auto causes a significantly lower amount of cytoplasmic leakage from membrane damage than Manual as quantified by influx of FITC–dextran during CPA exposure (P < 0.05). Inlets of brightfield and fluorescent composites demonstrate representative fluorescence signal. (C) Vitrification-derived sublethal damage decreases developmental competence after 96 h of embryo culture of murine zygotes. However, Auto significantly increases the number of blastomeres per blastocyst compared with Manual (P < 0.05) and provides evidence that sublethal damage was reduced. Error bars, standard error. Different letters signify statistical significance P < 0.05 by Student's t-test. Scale bars: 50 µm.

Decreased membrane perforations and cytoplasmic leakage

Murine zygotes were exposed to CPA supplemented with 4 KDa FITC–dextran, an impermeable and fluorescent solute, by auto-microfluidic or manual pipetting treatment groups. The auto-microfluidics protocol that realizes a slower zygote shrinkage rate had significantly lower fluorescent signal (1.71 ± 0.05; n = 15) compared with cells from the manual pipetting protocol that induces higher shrinkage rates (1.99 ± 0.11; n = 17; P < 0.05; Fig. 4B). The lower FITC–dextran indicates that the auto-microfluidics CPA exchange protocol induces less cytoplasmic leakage and membrane damage. These results are consistent with the notion that more rapid membrane compression and associated buckling cause more transient membrane damage to allow impermeable solute intercalation.

Increased developmental competence

To analyze how shrinkage rates affect embryo developmental competence, the number of blastomeres per blastocyst after 96 h culture were assessed using NCEV murine zygotes as control and zygotes vitrified with manual pipetting and auto-microfluidic CPA exchange as the experimental groups. NCEV zygotes have 109 ± 4 blastomeres (n = 46) after 96 h embryo development. Both CPA exposure/vitrification methods caused sublethal damage that significantly reduced blastomeres per blastocyst versus NCEV control. Furthermore, the vitrification method with the faster shrinkage rate (manual pipetting) resulted in significantly less blastomeres per blastocyst (89 ± 3 blastomeres, n = 35) compared with the method with reduced shrinkage rate (auto-microfluidic; 98 ± 3 blastomeres, n = 46, P < 0.05). Thus, sublethal damage was significantly decreased using vitrification by auto-microfluidic CPA exposure which preserved zygotes with higher developmental competence compared with its manual pipetting counterpart (Fig. 4C).

Discussion

For the rapid cooling cryopreservation method of vitrification, the pre-cooling CPA exchange protocol plays a critical role in the overall outcome. The challenge is that there are multiple conflicting needs that must be met. Introducing a sufficient amount of CPA into the cell is critical to avoid ice formation. On the other hand, it is also important to avoid excess CPA exposure time and concentrations as the chemicals are inherently toxic to cells. This confounding factor gives rise to the desire to decrease exposure time to reduce toxicity effects which directly conflicts with the desire to increase exposure time to reduce osmotic stress. However, what all the specific mechanisms are that contribute to osmotic stress remain unclear. One of the most prevalent theories of osmotic stress is that there is a minimum cell volume below which a cell will become lethally damaged. This theory arose from early studies of osmotic stress primarily because lethality and minimum cell volume are experimentally simple to quantify. A natural conclusion from such theory that has given rise to current protocols is that permeable CPA components should be added gradually (typically in three steps) to allow cell volume re-expansions through CPA permeation into the cell, but that impermeable solutes should be added at once at the end to minimize time of exposure to the impermeable CPA components. There could, however, be factors other than lethal minimum cell volumes contributing to osmotic stress. As the field of assisted reproduction technologies focus on increasing the efficiency of SET, the focus has also changed from lethality or cryosurvival to cell health and developmental competence. Here, we demonstrate using both mathematical modeling and experimental validation that shrinkage rate can be independently controlled while keeping the minimum cell volume reached the same. Furthermore, we show that maintaining a low shrinkage rate throughout the CPA exchange procedure is critical for reducing sublethal cell damage. These experiments are enabled by precise control of cell CPA exposure using microfluidic cell immobilization, microfluidics-enabled continuous gradual increase in CPA concentrations and the use of impermeable solutes as a key component in VS. It is worthy of note that there have also been early attempts to control CPA exposure to oocytes using a microscope diffusion chamber of microliter suspensions (McGrath, 1985). The micro-chamber device demonstrated the ability to observe dynamic single cell osmotic responses to increasing concentrations of sodium chloride by a diffusive mechanism through a dialysis membrane; however, this requires that the kinetic effects of the dialysis membrane for the solute of interest be clearly defined and considered for quantitative analysis. More recently, a combined mathematical and experimental analysis was performed by Heo et al. However, they did not consider the inclusion of impermeable CPAs, such as sucrose, in the VS as is highly recommended for practical cryopreservation (Kuleshova et al., 1999) and necessary to control shrinkage rate independently of minimum cell volume.

In our microfluidic system, the oocytes or zygotes are trapped within the exposure chamber by weir-type holding microchannels powered by a negative height differential for suction (Fig. 3B). Immobilized oocytes or zygotes within the microfluidic exposure chamber enabled real-time and high magnification imaging that is often lost during manual pipetting as stereoscopes are often used for their wide and depth of view but low magnification. Mixing within the microfluidic and generation of linearly increasing CPA concentrations to the cell exposure chamber were achieved using a similar mechanism (Heo et al., 2011). We then controlled minimum cell volume to be constant for all experimental groups while varying the maximal shrinkage rate and shrinkage profile.

One challenge for comparing CPA exchange protocols and their sublethal effects is the biological readout. Unlike lethal damage, the differences are more subtle. With improved vitrification protocols that now largely avoid lethal damage, the use of vitrification has become widespread. Measurement and reduction of sublethal damage associated with vitrification is thus timely and critical. To address the readout challenge, we quantified developmental competence by cryopreserving murine zygotes and comparing cell numbers of blastocysts (Fig. 4C). To better understand the mechanics that may underlie this reduction of sublethal damage in the auto-microfluidic CPA exchange protocol, the three conditions tested for lipid content and developmental competence were analyzed for differences in cell morphology (Figs 1H–J and 2) as well as for two additional CPA exchange protocols (Figs 1K and L and 2). The surface of the oocyte/zygote/embryo comprises a cell membrane–cytoskeleton complex that must be maintained relatively intact (e.g. low roughness and high sphericity) for successful cryopreservation (Ragoonanan et al., 2010). Surfaces of cells are not elastic or rigid but viscoelastic and thus are expected to deform depending not only on total strain (i.e. minimum cell volume) but also strongly dependent on the rate of strain application (i.e. cell shrinkage rate) (Pravincumar et al., 2012). Slower rates of strain application have been shown in other cell types to be important to allow sufficient time for mechanically induced actin remodeling and prevent membrane detachment. The low roughness and high sphericity of the auto-microfluidic CPA exchange protocol (Fig. 1J) is consistent with a viscoelastic material experiencing less stress because of a slower applied strain rate (Peeters et al., 2005). The high roughness and/or low sphericity observed in the other CPA exchange protocols that exert faster compressive strain rates (Fig. 1H, I, K and L) are consistent with buckling of a viscoelastic thin shell (Fu and Zhang, 2012). Such changes in morphology have been hypothesized to cause membrane damage that allow leakage of cytoplasmic content (Ashwood-Smith et al., 1988) and that the cell undergoes an energetically costly rapid crisis exocytotic response for membrane resealing (Togo et al., 1999; Miyake et al., 2001). A comparison of two microfluidic protocols [Fig. 1J (auto) and Fig. 1K (abrupt sucrose)] also suggest the importance of gradual addition of not only permeable CPAs but impermeable CPAs (sucrose) as well. The sudden addition of sucrose, which causes large and irregular changes in cell shape, has been previously shown (Smith et al., 2011) and again observed in our studies to be the step linked with causing the buckled membrane morphology. We additionally quantified cytoplasmic lipid retention of bovine oocytes (Fig. 4A) as cytoplasmic lipid is a major energy source, inversely related to sublethal cell damage, and a contributor to developmental competence (Dunning et al., 2011; Somfai et al., 2011; Sutton-McDowall et al., 2012; Chankitisakul et al., 2013). Cytoplasmic lipid can be predictive of future developmental success (Wong et al., 2010) and the significant difference in post-warm cytoplasmic lipid between auto and manual exposure groups is evidence that the shrinkage rate and its effect on morphology are more energetically costly to the oocyte, and may be due to the energy used in the exocytotic response to membrane resealment. A higher influx of FITC–dextran during CPA exposure demonstrates that cytoplasmic content leakage occurs at a higher rate in conditions with higher shrinkage rates (Fig. 4B): suggesting that a higher shrinkage rate causes a higher degree or frequency of membrane perforations during CPA exposure. All measures demonstrate that the auto-microfluidic gradual CPA exchange protocol with lower cell shrinkage rate is superior to its manual pipetting counterpart with a higher cell shrinkage rate. Importantly, having both low roughness and high sphericity is correlated with improved lipid content, less cytoplasmic leakage and higher developmental competence (Figs 2 and 4).

Conclusions

This paper reports fundamental insights on how a slow and steady cell shrinkage rate during CPA exchange reduces sublethal cellular damage. To enable this study, we developed a user-friendly and precise microfluidic CPA exchange system that significantly reduces the variability in terms of fluid exchange timing, volume error as well as convective forces applied to cells. This contrasts with commonly used pipetting-based manual CPA exchange protocols that are difficult to reproduce exactly, even within a single operator, resulting in cryopreservation outcome variability (Tsang and Chow, 2010). By eliminating run-to-run and operator-to-operator variability, the automated microfluidic system is envisioned to be broadly useful in the discovery, validation and use of new CPA formulations and procedures. The elimination of experimental variability would allow for easier determination of patient-specific partiality to different CPA formulations in human oocyte cryopreservation where there is a large variability in permeability characteristics. As a consideration for potential clinical use of the device, it is noted that the microfluidic cell introduction and recovery rate for 462 oocytes and zygotes has been 100%. Beyond direct applicability of the described devices and protocols to enhance oocyte and zygote vitrification, we anticipate the new mechanistic insights that complement and add to the minimum cell volume theory to be generally helpful in guiding future improvements in cell, organoid and tissue cryopreservation. The benefits observed from the gradient CPA exchange may also be advantageous to an automated warming process post-vitrification, which will require future investigation.

Supplementary data

Supplementary data are available at http://humrep.oxfordjournals.org/.

Authors' roles

D.L. and J.D. performed experiments; G.W.S. made numerous intellectual contributions; G.D.S. and S.T. conceptualized the work; D.L. wrote the manuscript. Corresponding author for gamete/zygote/embryo biology and vitrification: G.D.S. Corresponding author for microfluidics and modeling: S.T.

Funding

Funding was provided by NIH (GM096040, CA072005 and CA136829) and a Reproductive Sciences Program Pilot Grant.

Conflict of interest

D.L., J.D., G.D.S. and S.T. have a patent pending for automated CPA exposure methodology.