Hum. Reprod. Advance Access originally published online on August 12, 2006
Human Reproduction 2007 22(1):250-259; doi:10.1093/humrep/del319
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1,2-propanediol and the type of cryopreservation procedure adversely affect mouse oocyte physiology
Colorado Center for Reproductive Medicine, Englewood, CO, USA
1 To whom correspondence should be addressed at: Colorado Center for Reproductive Medicine, 799 East Hampden Avenue, Suite 520, Englewood, CO 80113, USA. E-mail: dgardner{at}colocrm.com
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Abstract |
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BACKGROUND: The aim of this work was to examine the effect of 1,2-propanediol (PrOH) and type of cryopreservation procedure (slow freezing and vitrification) on oocyte physiology. METHODS: Intracellular calcium of mouse metaphase II (MII) oocytes was quantified by fluorescence microscopy. The effect of PrOH on cell physiology was further assessed through analysis of zona pellucida hardening and cellular integrity. Protein profiles of cryopreserved oocytes were generated by time-of-flight mass spectrometry (TOF-MS). RESULTS: PrOH caused a protracted increase in calcium, which was sufficient to induce zona pellucida hardening and cellular degeneration. Using ‘nominally calcium free’ media during PrOH exposure significantly reduced the detrimental effects. Proteomic analysis identified numerous up- and down-regulated proteins after slow freezing when compared with control and vitrified oocytes. CONCLUSIONS: Using such approaches to assess effects on cellular physiology is fundamental to improving assisted reproduction techniques (ART). This study demonstrates that PrOH causes a significant rise in intracellular calcium. Using calcium-free media significantly reduced the increase in calcium and the associated detrimental physiological effects, suggesting that calcium-free media should be used with PrOH. In addition, analysis of the oocyte proteome following cryopreservation revealed that slow freezing has a significant effect on protein expression. In contrast, vitrification had a minimal impact, indicating that it has a fundamental advantage for the cryopreservation of oocytes.
Key words: calcium/cryoprotectant/oocyte/proteome/slow freezing
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Introduction |
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TopAbstractIntroductionMaterials and methodsResultsDiscussionAcknowledgementsReferences |
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Cryopreservation of human oocytes is becoming a valuable technique in assisted reproduction. The ability to efficiently cryopreserve human oocytes will not only assist many female patient groups, including those who are preparing to undergo fertility-threatening radiotherapy or chemotherapy (Falcone et al., 2004
) but will also have an impact on practical aspects of human IVF. Examples include the inability to collect a sperm sample on the day of oocyte retrieval, permitting robust viral infection testing during oocyte donation and storage of supernumerary oocytes for patients in countries that only allow the insemination of a limited number of oocytes (Boggio, 2005; Li et al., 2005).
Despite the first birth from a frozen human oocyte occurring 20 years ago (Chen, 1986), the success of this technology has yet to reach the equivalent prevalence of embryo cryopreservation. The technique most commonly used for cryopreservation in IVF clinics is slow freezing. This process involves introducing a permeable cryoprotectant [usually 1,2-propanediol (PrOH)] into the cell and removing water via an osmotic gradient (typically involving sucrose). Further dehydration occurs during the subsequent cooling and seeding phases, which leads to extracellular ice formation and a subsequent increase in osmolarity. By reducing the water content of the cell and introducing the permeable cryoprotectant, it is proposed that the cytosol will vitrify, forming a glass-like state, and not form membrane damaging ice crystals. An alternative cryopreservation technique, vitrification, involves rapid cooling and warming rates, so that both the external medium and cytosol vitrify without ice crystal formation. For this to occur, higher concentrations of cryoprotectant are used (three to four times higher) than in slow freezing. This has raised concerns regarding the toxicity of these components. However, vitrification does offer several practical advantages over slow freezing, such as limited time out of the incubator (<10 min), no zona pellucida fracturing and reduced equipment/running costs (Kuleshova and Lopata, 2002).
The process of vitrification has been improved significantly in recent years (Vajta et al., 1998; Lane and Gardner, 2001; Liebermann et al., 2003; Isachenko et al., 2004; Kuwayama et al., 2005). Such improvements include the development of novel instruments that permit the use of sub-microlitre volumes, which significantly increases the rates of cooling and warming. Studies to date have employed electron microscope grids (Martino et al., 1996), open pulled straws (Vajta et al., 1998), solid surface vitrification (Dinnyes et al., 2000), cryotop (Hochi et al., 2004), nylon mesh (Matsumoto et al., 2001) and the cryotip (Kuwayama et al., 2005). The Cryoloop is another such instrument that has been employed to increase cooling and warming rates (Lane et al., 1999). The Cryoloop is composed of a thin nylon loop that holds a surface volume of 
10 nl. Oocytes or embryos are pipetted onto the loop in a minimal volume, making cooling and warming extremely rapid (>20 000°C/min). Previous publications have shown that Cryoloop vitrification offers a significant advantage over slow freezing of both mouse oocytes and embryos (Lane and Gardner, 2001; Lane et al., 2002). It was also shown that compared with vitrification, oocytes cryopreserved using slow freezing produced blastocysts with significantly fewer cells (including a lower inner cell mass : trophectoderm ratio) and reduced viability following embryo transfer (Lane and Gardner, 2001). Furthermore, when monitoring metabolism through pyruvate uptake, mouse oocytes and developing embryos following slow freezing were metabolically impaired compared with those that were vitrified (Lane and Gardner, 2001; Lane et al., 2002). Analysis of membrane integrity through the retention of the enzyme lactic dehydrogenase (LDH) also indicated that slow freezing induces significantly more plasma membrane damage than vitrification (Lane et al., 2002). With inconsistent reports on the survival rates following oocyte slow freezing and limited published data on current vitrification techniques, further cellular-based investigation into oocyte cryopreservation is warranted. Using such an approach, cellular stresses and detrimental effects on cell physiology can be analysed and subsequently minimized.
Recently, two cryoprotectants commonly used during vitrification (ethylene glycol and dimethylsulfoxide) have been shown to cause a large transient increase in intracellular calcium in mouse metaphase II (MII) oocytes, (Larman et al., 2006). A third cryoprotectant, PrOH, is typically used during slow freezing of oocytes and cleavage-stage embryos (Lassalle et al., 1985; Emiliani et al., 2000; Borini et al., 2004). The effects of PrOH on oocyte physiology and intracellular calcium levels are unknown. Because PrOH is almost exclusively used during slow freezing, it is imperative to understand the cellular affects it may induce.
The aim of this study was to investigate the effects of PrOH on oocyte physiology by analysing intracellular calcium. Furthermore, with the recent application of time-of-flight mass spectrometry (TOF-MS) to the analysis of the mammalian preimplantation embryo proteome (Katz-Jaffe et al., 2005), it was possible to investigate the effect of cryopreservation on the oocyte proteome.
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Materials and methods |
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Oocytes were collected from 4- to 5-week-old F1 (C57BL/6 x CBA/Ca) females (Jackson Laboratory, Bar Harbor, ME, USA). Female mice were administered 5 IU of pregnant mare’s serum gonadotrophin (PMSG) and 48–50 h later 5 IU of hCG. Oocytes were collected 12.5–13.5 h post-hCG and denuded by incubation in G-MOPS (Lane and Gardner, 2004
) [supplemented with 5 mg/ml of human serum albumin (HSA)] with 0.5 mg/ml of hyaluronidase. Oocytes were washed three times and held in droplets of G-MOPS with HSA under Ovoil (Vitrolife, Gothenburg, Sweden) at 37°C. Unless otherwise stated, all materials were obtained from Sigma Chemical CO. (St Louis, MO, USA). This work received prior approval from the Institutional Animal Care and Users Committee.
Slow freezing and thawing
All slow-freezing procedures were performed at room temperature, and the base medium for all freezing solutions was G-MOPS supplemented with 20% fetal calf serum (FCS). Oviducts from individual mice were collected, and oocytes extruding their first polar body were isolated and assigned randomly to one of the following groups: control (non-cryopreserved), slow freezing with or without calcium. Upon slow freezing, oocytes were first exposed to G-MOPS containing 1.5 mol/l PrOH for 10 min, followed by G-MOPS with 1.5 mol/l PrOH and 0.2 mol/l sucrose solution for 10 min. Oocytes from each individual mouse were loaded into 25 cc straws (Institute Medicine Vetinaire, Bicef, L’Aigle, France) and placed into a freezing machine (Freeze Control CL2000; CryoLogic, Napa, CA, USA). The starting temperature of the freezing procedure was 20°C. Oocytes were cooled at the rate of 2.0°C/min to –7°C, seeded at –7°C and held for 10 min. Further cooling was carried out at a rate of 0.3°C/min to –35°C before plunging into liquid nitrogen. Oocytes were stored in liquid nitrogen for a minimum of 24 h before thawing.
Straws containing oocytes were thawed in air for 30 s before being placed in a 30°C water bath for 10 s. The base medium for thawing was G-MOPS supplemented with 20% FCS. Oocytes were thawed at room temperature. Oocytes were expelled into 1.0 mol/l PrOH with 0.2 mol/l sucrose for 5 min and then moved through the following steps: 0.2 mol/l sucrose for 5 min and then 0.1 mol/l sucrose for 5 min in G-MOPS. Lastly, oocytes were washed in G-MOPS for 5 min before survival assessment and subsequent extraction.
Vitrification and warming
Oviducts from individual mice were collected, and oocytes extruding their first polar body were isolated and assigned randomly to the following groups: control (non-cryopreserved), vitrification with or without calcium. Vitrification and warming were carried out at 37°C in a base solution of G-MOPS supplemented with 12 mg/ml of HSA. Vitrification was performed using a two-step method and the Cryoloop, as previously described (Lane and Gardner, 2001). Oocytes from each individual mouse were placed in the initial cryoprotectant solution (8% DMSO, 8% EG v/v: equates to 1.13 mol/l DMSO, 1.43 mol/l EG) for 1 min. Oocytes were then moved to the second solution (16% DMSO, 16% EG v/v, equates to 2.25 mol/l DMSO, 2.86 mol/l EG, 0.65 mol/l sucrose and 10 mg/ml Ficol) for <30 s before being pipetted onto a Cryoloop, which had been preloaded with this second solution. The Cryoloop was then plunged into a cryovial filled with liquid nitrogen to facilitate vitrification. For warming, oocytes were moved through 1 ml serial dilutions of sucrose (0.25 mol/l for 1 min, 0.125 mol/l for 2 min and 0 mol/l for 5 min) to reduce osmotic stress (Lane and Gardner, 2001).
Intracellular calcium measurements
Intracellular calcium changes were monitored by loading the oocytes with 10 µmol/l Indo-1 AM (Molecular Probes, Eugene, OR, USA) for 30 min. The loading media included 250 µmol/l sulfinpyrazone to reduce dye compartmentalization and extrusion (Lawrence et al., 1997). Individual oocytes were held in G-MOPS (supplemented with 20% FCS) with or without calcium (using a holding pipette) in a chamber at room temperature (21–22°C) on a Nikon TE300 microscope equipped with a SFX-2 microfluorimeter (Solamere Technology Group, Salt Lake City, UT, USA). For calcium-free treatment oocytes were incubated in G-MOPS (with 20% FCS) without calcium at 37°C for 30 min before cryoprotectant exposure. Solutions without calcium were termed nominally free, because calcium was simply omitted from the stock solution. To ensure rapid exposure that would best mimic the vitrification procedure, oocytes were held in a small droplet (20 µl) of G-MOPS before 980 µl of the cryoprotectant solution (1.5 mol/l PrOH, made up in G-MOPS with 20% FCS) was added to the chamber.
Zona pellucida hardening and oocyte degeneration assessments
For zona hardening assessment, oocytes were transferred to a 1% w/v solution of chymotrypsin (in G-MOPS supplemented with 5 mg/ml of HSA) on a heated stage (37°C), and the integrity of the zona pellucida was monitored over time with the end point being when the zona was no longer clearly visible (Matson et al., 1997).
To assess the effect of exposure to PrOH on cellular integrity, we incubated oocytes in 1.5 mol/l PrOH (G-MOPS with 20% FCS) at room temperature. Following timed exposure to PrOH (2, 5 and 10 min), oocytes were washed in G-MOPS. Oocytes were then incubated in G-MOPS at 37°C for 4 h before determining which oocytes had survived the treatment.
Cation and anion exchange chromatography
Groups of five oocytes (n = 13–17 replicates per group) were extracted in 5 µl of lysis buffer (9 mol/l urea/2% CHAPS) and stored at –80°C until analysis. Cationic (CM10) and anionic (Q10) ProteinChips (Ciphergen Biosystems, Freemont, CA, USA) were equilibrated twice with binding buffers, 0.1 mol/l sodium acetate, pH 4.0 and 0.1 mol/l Tris–HCl, pH 9.0, respectively (Katz-Jaffe et al., 2005). Oocyte samples were thawed and incubated for 30 min on a shaker at room temperature. Any unbound sample was discarded, and the chips were washed three times with the respective binding buffer. Chips were then quickly rinsed with distilled water before air-drying. The energy-absorbing molecule, sinapinic acid (Ciphergen Biosystems), was prepared as a saturated solution in 50% acetonitrile/0.5% trifluoroacetic acid, and each spot was loaded twice with 1 µl. Once dry, the chips were analysed by TOF-MS.
Tof-Ms
TOF-MS was performed using the PCS-4000 Series mass spectrometer (Ciphergen Biosystems). TOF data were collected from averaged 530 laser shots per spot capturing peptides and proteins <20 kDa. Mass-to-charge ratios (m/z) were calibrated externally by the all-in-1 peptide molecular mass standard (Ciphergen Biosystems). Protein profiles were generated by Ciphergen Express Software, version 3.0 (Ciphergen Biosystems).
Statistics and bioinformatic analysis
Differences in protein expression levels were analysed with Ciphergen Express Software, version 3.0 (Ciphergen Biosystems). Peaks with a signal-to-noise ratio higher than 6 were selected, and profiles were then normalized to the same total ion current. The Mann–Whitney non-parametric test was used to calculate statistically significant differences across the groups (P < 0.05). Hierarchical and horizontal clustering was performed using the BioMarker Wizard module of the software to group samples with similar protein profiles. To test the reproducibility of the platform, samples were analysed in triplicate, and the coefficient of variance of individual peaks (dividing the standard deviation by the mean peak height multiplied by 100%) in the replicate spectra was calculated to be between 7 and 12%.
For comparison of increases in intracellular calcium, statistical difference was determined through unpaired student’s t-test. Statistical difference in levels of zona pellucida hardening and cell integrity were determined through ANOVA and subsequent Bonferroni comparison.
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Results |
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TopAbstractIntroductionMaterials and methodsResultsDiscussionAcknowledgementsReferences |
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To investigate the effect of PrOH on intracellular calcium, mouse oocytes were loaded with a fluorescent calcium indicator (Indo-1) and subjected to a PrOH exposure regime identical to that of the initial phase of slow freezing. Figure 1a shows that exposure to 1.5 mol/l PrOH at room temperature causes a protracted calcium increase. The mean base to peak ratio increase was 4.6 ± 0.6 (Figure 1c). The calcium remained elevated during the 10-min exposure. Using calcium-free medium significantly (P = <0.05) reduced the increase in the mean base-to-peak ratio observed during PrOH challenge (Figure 1b and c; 1.9 ± 0.2).
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The concentrations of dimethylsulphoxide (DMSO) and ethylene glycol used during vitrification can increase intracellular calcium to levels sufficient to induce zona pellucida hardening (Larman et al., 2006
). The state of zona hardening, after exposing the oocytes to 1.5 mol/l PrOH at room temperature, was assessed by monitoring the time it takes for zona dissolution with a 1% solution of 
-chymotrypsin (Figure 2). Oocytes not treated with PrOH had a mean zona dissolution time of 236.4 ± 27.3 s. The time for zona dissolution of these control oocytes was almost 3-fold less than those exposed to 1.5 mol/l PrOH for 10 min at room temperature. The mean zona dissolution time for PrOH-treated oocytes was 683.2 ± 63.8 s. If calcium was omitted from the medium, the time taken for zona dissolution following PrOH exposure was comparable to the untreated control (262.2 ± 35.2 s).
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A problem associated with oocyte cryopreservation, particularly slow freezing, is the low overall survival rate (50–65%: Todorow et al., 1989a
,b; Carroll et al., 1990; Stachecki et al., 1998). Prolonged increases in intracellular calcium can be cytotoxic (Berridge et al., 1998; Orrenius et al., 2003). Increasing the exposure time of oocytes to 1.5 mol/l PrOH (at room temperature) decreased the survival number. Figure 3 shows that 94.5 ± 3.6% of oocytes survived if the treatment period was 2 min. Increasing the exposure time to 10 min decreased the survival rates to 40% (Figure 3). To determine if reducing the calcium increase caused by PrOH would enhance survival, exposure was carried out in calcium-free medium. Calcium-free medium was found to dramatically enhance survival rates, with each of the timed exposures exhibiting 100% survival.
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Previous studies have revealed that slow freezing significantly reduces oocyte and embryo metabolism, as well as subsequent embryo development and viability (Lane and Gardner, 2001
; Lane et al., 2002). Using TOF-MS, the effect of slow freezing and vitrification (in the presence and absence of calcium) on the oocyte proteome was investigated. Over the molecular weight range up to 20 kDa, 17 positively charged proteins showed a significant alteration in expression levels after slow freezing (with or without calcium) (P = <0.05). Figure 4a shows the original cationic protein profiles enhanced around the 3500–5000 Da range for the following groups: control (non-cryopreserved) in vivo MII oocytes, slow frozen MII oocytes with calcium, slow frozen MII oocytes without calcium, vitrified MII oocytes with calcium and vitrified MII oocytes without calcium. Of the 17 positively charged proteins that showed a significant alteration of expression, compared with control (non-cryopreserved) oocytes, 12 were up-regulated and 5 were down-regulated. Figure 4a shows an example of two up-regulated proteins observed following slow freezing (highlighted in red box) and two of the down-regulated proteins (highlighted in green box). From the group of 12 up-regulated positively charged proteins after slow freezing, 9 showed statistically greater up-regulation when slow freezing occurred in the presence of calcium (Figure 4a, red box). In addition, several proteins appeared only in the profiles of in vivo control MII oocytes (Figure 4a, black box).
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0.05) in protein expression across the five groups; black indicates proteins only observed in controls, green indicates proteins down-regulated in slow frozen oocytes and red indicates proteins up-regulated in slow frozen oocytes. (b) CM10 heat map displaying hierarchical clustering of control oocytes with vitrified oocytes (with or without calcium) separate to slow frozen oocytes (with or without calcium). Each column of squares represents an individual sample, whereas each row of squares represents an individual protein profile. Red, up-regulated proteins; green, down-regulated proteins.Hierarchical clustering grouped control and vitrified MII oocytes (with or without calcium) in one classification separate from slow frozen MII oocytes (with or without calcium) (Figure 4b). The positively charged proteome of vitrified MII oocytes (with or without calcium) most closely resembled the positively charged proteome of control MII oocytes (Figure 4b). Some examples of these differentially expressed positively charged proteins are displayed in Figure 5. Figure 5a shows a box plot for a 4.1 kDa protein that was significantly (P = <0.00001) up-regulated after slow freezing (with or without calcium). Figure 5b displays a box plot for a 3.8-kDa protein that was significantly (P = <0.00002) down-regulated after slow freezing (with or without calcium), whereas Figure 5c shows a box plot for a 3.9-kDa protein that was further significantly (P = <0.0002) up-regulated when calcium was added to the slow-freeze medium.
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Over the molecular weight range analysed (up to 20 kDa), 21 negatively charged proteins showed a significant alteration in expression levels after slow freezing (with or without calcium) (P = <0.05). Figure 6a shows the original anionic protein profiles enhanced around the 3800–4300 Da range for the following groups: control (non-cryopreserved) in vivo MII oocytes, slow frozen MII oocytes with calcium, slow frozen MII oocytes without calcium, vitrified MII oocytes with calcium and vitrified MII oocytes without calcium. Of the 21 negatively charged proteins that showed a significant alteration of expression, after slow freezing compared with control (non-cryopreserved) oocytes, 7 were up-regulated (examples are shown in Figure 6a highlighted by the light grey boxes) and 14 were down-regulated (examples are shown in Figure 6a highlighted by the dark grey boxes).
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Hierarchical clustering once again grouped control MII oocytes and vitrified MII oocytes (with or without calcium) in one classification separate from slow-frozen MII oocytes (with or without calcium) (Figure 6b). The negatively charged proteome of vitrified MII oocytes (with or without calcium) most closely resembled the negatively charged proteome of control MII oocytes (Figure 6b).
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Discussion |
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TopAbstractIntroductionMaterials and methodsResultsDiscussionAcknowledgementsReferences |
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The ability to routinely cryopreserve human oocytes with survival, fertilization and pregnancy rates approaching those experienced with fresh oocytes, will have a significant impact on the practice of IVF. However, a major obstacle with oocyte cryopreservation is the variable survival rates, especially with slow freezing (12–90%: Chen, 1986
; Porcu et al., 1997; Boldt et al., 2003; Fosas et al., 2003). Substituting sodium with choline has been shown to dramatically improve survival rates with slow freezing of mouse MII oocytes (Stachecki et al., 1998). A survival rate of 52.5% was reported following freezing with a typical sodium-containing solution, but replacing the sodium with choline increased survival rates to 84.4%. Although this is a significant improvement, it should be noted that vitrification of mouse MII oocytes already yields even greater survival rates of 95% (Dela Pena et al., 2001; Lane and Gardner, 2001). Furthermore, the impact of choline-based slow freezing on human oocyte cryopreservation is unclear because there is yet to be a published, scientific comparison with a sodium-depleted medium (Quintans et al., 2002; Boldt et al., 2003).
Recent clinical publications have shown that vitrification may offer some improvement over slow freezing, with increasingly higher survival rates and less of an effect on cell physiology (Hong et al., 1999; Lane and Gardner, 2001; Kuleshova and Lopata, 2002; Liebermann et al., 2003; Kyono et al., 2005; Valojerdi and Salehnia, 2005). In part, this is probably because of the introduction of ultra-rapid vitrification, which employs techniques that allow the volume of the vitrification solution to be in the sub-microlitre range. This effectively increases the cooling and warming rates to >20 000°C/min (Steponkus et al., 1990; Mukaida et al., 2001). In the single largest published study to date, Liebermann et al. (2003) demonstrated an 81% survival rate of human oocytes using the Cryoloop. This is encouraging when considered alongside several other smaller studies that report 90–100% survival rates (Hong et al., 1999; Liebermann and Tucker, 2002; Katayama et al., 2003; Chian et al., 2005; Kuwayama et al., 2005; Kyono et al., 2005).
Larman et al. (2006) demonstrated that two cryoprotectants commonly used in vitrification protocols (DMSO and ethylene glycol) cause transient increases in intracellular calcium in mouse MII oocytes. The increase in calcium is sufficient to cause zona hardening, presumably through triggering cortical granule exocytosis, which is a calcium-dependent event (Kline and Kline, 1992). By removing extracellular calcium, it was demonstrated that the source of the calcium increase in response to ethylene glycol was mainly extracellular in contrast to intracellular for DMSO. Here, we report that exposing mouse MII oocytes (at room temperature) to 1.5 mol/l PrOH causes a protracted calcium increase that is also capable of causing zona pellucida hardening. This treatment protocol is similar to that of the first equilibration step exacted during slow freezing. Zona hardening can be overcome by the inclusion of FCS (George and Johnson, 1993). It is believed that the active component, fetuin, acts as a competitive substrate for proteolytic enzymes released from the cortical granules. Zona hardening assays were carried out in medium containing HSA, which has been shown not to have the same protective action of FCS or bovine serum albumin (BSA) (George and Johnson, 1993). However, the slow-freezing protocol included FCS, which may have reduced the loss of protein expression normally associated with cortical granule exocytosis and zona hardening.
Unlike DMSO and ethylene glycol, the calcium response to PrOH remains elevated as opposed to being transient in nature. Although the PrOH challenge was carried out at room temperature, a protracted calcium increase was also observed at 37°C (data not shown), indicating the difference between the cryoprotectants is not temperature dependent. Most PrOH-induced increase is extracellularly derived because removing extracellular calcium during the exposure significantly reduces the level of intracellular calcium release. This is also the case during ethylene glycol challenge (Larman et al., 2006). Exactly how these cryoprotectants induce a rise in intracellular calcium is unknown, but passive transport as the cryoprotectant crosses the plasma membrane, is probably the most likely. Removing extracellular calcium also prevents zona hardening in response to PrOH. The clinical significance of this data is that although PrOH induces a prolonged increase in intracellular calcium, this can be controlled by using a medium to which calcium has not been added. Therefore, it is recommended that oocyte slow-freezing protocols should evaluate calcium-free media.
Zona hardening is not the sole downstream effect of a rise in intracellular calcium. Calcium is a ubiquitous cellular signalling messenger; therefore, strict homeostatic regulation of calcium is vital (Berridge et al., 1998). Elevated and sustained increases in intracellular calcium can lead to inappropriate activation of proteases and phospholipases, degeneration and apoptosis (Shaw and Trounson, 1989; Gordo et al., 2000, 2002; Orrenius et al., 2003; Takahashi et al., 2004). PrOH has also been shown to cause degeneration and parthenogenetic activation of mouse MII oocytes (Shaw and Trounson, 1989; Van Der Elst et al., 1992). Exposing oocytes to 1.5 mol/l PrOH (for 10 min at room temperature) causes degeneration in 60% of oocytes, indicating a severe effect on oocyte viability. Using calcium-free media, the effect of the exposure to PrOH was eliminated, suggesting that the rise in calcium levels leads to cell degeneration. It is concerning that not only can PrOH release sufficient calcium to cause degeneration but also initiate activation and cell development of those that survive (Shaw and Trounson, 1989; Van Der Elst et al., 1992). Cell cycle progression in parthenogenetically activated mouse oocytes is dependent on the level of the rise in intracellular calcium (Vincent et al., 1992; Ducibella et al., 2002). The increase in calcium induced by PrOH is adequate to support development of the mouse embryo to the blastocyst stage (Larman, unpublished observation). Furthermore, removing calcium from the slow-freezing media reduced the level of proteome alteration. However, even in the absence of calcium, the proteome of the oocyte was affected by the slow-freezing process. These data clearly support the idea that the increase in intracellular calcium induced by PrOH exposure leads to a decrease in oocyte viability and an alteration of the oocyte proteome.
Recent advances in mass spectrometry have allowed its application in studying biological samples (Seibert et al., 2004; Xiao et al., 2005). Protein profiling has been used to identify candidate proteins or biomarkers that are altered in response to different physiological states (Wong et al., 2004; Zhang et al., 2004; Xiao et al., 2005). With the ability to generate protein profiles and identify biomarkers from individual or groups of oocytes/embryos (Ellederova et al., 2004; Katz-Jaffe et al., 2006), it is possible to utilize this technology to identify proteins affected by specific assisted reproduction technology (ART) procedures and protocols. Differential protein expression between developing and degenerating human embryos was reported by Katz-Jaffe et al. (2006), with several candidate protein identifications. This approach has also been used to demonstrate that embryos developing in a low (5%) O2 environment as opposed to atmospheric O2 (20%) have a proteome closer to that of in vivo controls (Katz-Jaffe et al., 2005), signifying the usefulness of this approach when comparing alternative treatments and techniques.
This is the first report that has analysed the proteome of mouse oocytes after cryopreservation. By monitoring the proteome of small groups of oocytes that have been slow frozen or vitrified, we have found that those vitrified most closely resemble the in vivo control. A previous study showed that slow freezing has detrimental effects on oocyte metabolism and subsequent embryo development and viability (Lane and Gardner, 2001). Disruption of the plasma membrane, possibly through free-radical production, resulted in protein efflux (Lane et al., 2002), which may explain the down-regulation of proteins following slow freezing observed in this study. The observation that specific proteins were significantly up-regulated after slow freezing could be associated with the length of time the oocytes are exposed to the toxic cryoprotectant. This exposure may induce stress-related responses and the up-regulation of stress proteins and/or apoptosis. Ongoing research is focused on the purification and identification of a select number of these differentially expressed proteins to provide a mechanistic insight into the effects of oocyte cryopreservation at the cellular level.
This study raises significant concerns with regard to the current use of PrOH during oocyte cryopreservation. PrOH causes a protracted increase in intracellular calcium that is capable of inducing cellular degeneration and parthenogenetic activation. This is also the first report that has analysed the proteome of cryopreserved oocytes. Slow frozen oocytes show a striking alteration in their proteome. Whether this is simply because of the PrOH exposure regimen and/or cooling is unclear at present, but an investigation has been initiated to determine which stage of the slow freezing procedure causes proteome alteration. However, using calcium-free media does reduce, though not completely eliminate, the level of perturbation for some proteins during slow freezing. This supports the implementation of calcium-free media during slow freezing. In contrast to the outcome with slow freezing, hierarchical cluster analysis groups the proteome of vitrified oocytes with that of controls. Performing such in-depth analyses of cellular physiology is a fundamental approach to improving and optimizing cryopreservation techniques and in this particular incidence, provides evidence that slow freezing is undesirable for oocyte cryopreservation.
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Acknowledgements |
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The authors thank Vitrolife for their support of this study.
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References |
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Submitted on May 16, 2006; resubmitted on July 6, 2006; accepted on July 11, 2006.
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