Lipid phase transitions in cat oocytes supplemented with deuterated fatty acids

Abstract

Cryopreservation of oocytes has already been used to preserve genetic resources, but this technology faces limitations when applied to the species whose oocytes contain large amounts of cytoplasmic lipid droplets. Although cryoinjuries in such oocytes are usually associated with the lipid phase transition in lipid droplets, this phenomenon is still poorly understood. We applied Raman spectroscopy of deuterium-labeled lipids to investigate the freezing of lipid droplets inside cat oocytes. Lipid phase separation was detected in oocytes cryopreserved by slow-freezing protocol. For oocytes supplemented with stearic acid, we found that saturated lipids form the ordered phase being distributed at the periphery of lipid droplets. When an oocyte is warmed to physiological temperatures after cooling, a fraction of saturated lipids may remain in the ordered conformational state. The fractions of monounsaturated and polyunsaturated lipids redistribute to the core of lipid droplets. Monounsaturated lipids undergo the transition to the ordered conformational state below −10°C. Using deuterated fatty acids with a different number of double bonds, we reveal how different lipid fractions are involved in the lipid phase transition of a cytoplasmic lipid droplet and how they can affect cell survival. Raman spectroscopy of deuterated lipids has proven to be a promising tool for studying the lipid phase transitions and lipid redistributions inside single organelles within living cells.

Significance

We proposed to expand the capabilities of Raman spectroscopy to study lipid phase transition and phase coexistence by adding deuterated lipids to analyze the state of different lipid fractions within lipid droplets. Deuterated labels open the opportunity to discern the particular lipid fraction from other lipids in Raman spectra. Thus, we investigated the transition of lipid fractions with different unsaturation degree and analyzed how these fractions distribute within lipid droplets inside a living oocyte. It is demonstrated that the conformational and phase state of particular lipid fractions can be studied in situ in a nondestructive single-cell manner using Raman spectroscopy approach.

Introduction

Cryopreservation of gametes and early embryos is a promising approach for the preservation of genetic material of laboratory (1) and farm (2) and endangered animals (3, 4, 5). However, the cryopreservation technology is not universal; in each case (for each species and type), it is necessary to find the particular cryopreservation protocols that take into account the specificity of the cells. The main difficulties in cryopreservation are associated with phase transitions in biological samples during cooling to cryogenic temperatures. These transitions radically change the cellular environment and the conditions for biological processes inside cells. Nowadays, cryopreservation protocols have been developed to reduce damage associated with water crystallization in and around cells. For this purpose, cryoprotectants, compounds that suppress the ice formation and have relatively low toxicity, are added to aqueous solutions for cryopreservation (6). At the same time, other sources of cryoinjury associated with phase transitions of materials that form biological cells are still insufficiently studied. The choice of a strategy for their suppression remains an unresolved problem.

An example of phase transitions occurring directly within biological compartments is the phase transition of lipids (LPT). This phenomenon occurs in membranes and lipid droplets (LDs). Despite the different dimensionality, two-dimensional for membranes and three-dimensional for droplets, both structures are composed of lipid molecules having two order parameters related to conformational and translational ordering. In complicated biological mixtures, the LPT is broadened, and in its vicinity, the coexistence of lipids of different phases can be observed. In addition to influencing the cryopreservation process, the coexistence of domains of different phases (or “rafts”) is an important regulatory mechanism in cell membranes under physiological conditions (7). Therefore, this phenomenon is being actively studied on model objects using various techniques, including fluorescence microscopy (8), ²H-NMR (9, 10, 11), Raman spectroscopy (12), atomic force microscopy (13), and many other techniques. However, this phenomenon remains largely unexplored in living cells, for which many methods have only recently been adapted (14,15). Moreover, phase separation of lipids within LDs remains much less investigated (16).

LDs are believed to play an important role in the cryopreservation of mammalian embryos and oocytes with high lipid content (17, 18, 19). It was demonstrated that delipidation can increase cell survival after cryopreservation (17,18,20). However, LDs are required for cell metabolism (21,22), and delipidation has a negative effect on further embryo development (23). The primary mechanism of cryoinjuries related to LDs remains a question for research. The most significant changes in LDs during cryopreservation are related to the LPT. Therefore, it is assumed that the efficiency of cell survival during cryopreservation is directly related to the LPT. In analogy with how the addition of cryoprotectants can reduce the injury caused by the ice crystallization, attempts were made to control the LPT by regulating the lipid composition (18,24, 25, 26, 27, 28).

Vibrational spectroscopy opens the possibility of studying the LPTs directly in biological cells without complex sample preparation (29). Infrared spectroscopy was applied for studying cell membranes (30,31), including mammalian oocytes (24,32). Recently, we investigated the LPT in single LDs using Raman spectroscopy and showed that the phase transition in LDs is significantly broadened compared with monocomponent lipid models (33). The phase coexistence in LDs of cat oocytes at scales of a few microns was found (16). It was shown that within one LD coexist an ordered phase formed by conformationally ordered lipids and a disordered phase formed by lipids in a disordered conformational state. This discovery raises new questions about the composition of the coexisting phases, how it depends on temperature, and whether it is possible to suppress the phase separation during cryopreservation by varying lipid content.

Unfortunately, vibrational spectroscopy approaches have a low capability to distinguish different lipids and study the lipid separation effect. Nevertheless, it is possible to highlight the molecules of interest using stable isotope labeling. The replacement of protons with deuterons leads to an isotopic shift in the vibrational spectra to the spectral range not occupied by other lines, which allows an independent study of deuterated compounds. Raman spectroscopy of deuterated compounds combines the contactless high-resolution measurements of fluorescence microscopy and sensitivity to isotopic labels of mass spectrometry. The advantage of deuterated labels over fluorescent ones is the small size of the tags. This is especially important for small molecules such as fatty acids. Using deuterium labeling, one can track lipid uptake (34, 35, 36, 37, 38) and chemical transformations (39). Moreover, Raman spectra contain information about conformational ordering (40).

Labeled fatty acids can be added to a cell after forming complexes with albumin, such as bovine serum albumin (BSA). Fatty acid transport across the cell membrane occurs with the help of membrane proteins fatty acid translocase (СD36) and membrane fatty-acid-binding protein (41). Inside the cell, fatty acids are transported by cytoplasmic fatty-acid-binding and transport proteins or as part of the long-chain fatty acyl-CoA esters. Depending on the regulatory mechanisms, the introduced acids can be used in catabolic reactions (β-oxidation) (42) or accumulate through the formation of more complex membrane lipids (43) and triglycerides (44) inside the endoplasmic reticulum. Triglycerides can also be formed directly by LD, making it possible to study new LDs and old ones formed before the introduction of fatty acids.

In this work, using phase transitions of LDs in oocytes of a domestic cat as an example, we demonstrated the possibility of Raman spectroscopy of deuterated lipids for studying phase transitions in living cells and organelles. The suggested approach made it possible to reveal how, inside LDs, different lipid fractions are spatially redistributed and undergo the change in their conformational states upon cooling. The study of oocytes cooled using different cryopreservation protocols revealed that the observed effects of lipid phase separation should be encountered in practice during slow programmed freezing.

Materials and methods

Preparation of fatty acid-BSA solutions

Deuterated stearic-d35 acid (dSA) and arachidonic-5,6,8,9,11,12,14,15-d8 acid (dAA) were obtained from Cambridge Isotope Laboratories (Andover, MA), deuterated oleic-15,15,16,16,17,17,18,18,18-d9 acid (dOA) was obtained from Avanti Polar Lipids (Alabaster, AL), and fatty-acid-free BSA was obtained from Merck (Darmstadt, Germany). Chemical formulas of these compounds are shown in Fig. 1 a. The dSA-BSA, dOA-BSA, and dAA-BSA solutions were prepared according to the protocol described previously (45), with some modifications. The saline solutions of acids with alkali were made at the first stage, then the salts were combined with fatty-acid-free BSA for the transporting of fatty acids into the cells. The solution of potassium stearate in 0.01 M concentration was prepared as described in (38), using a water bath at 90°С for 60 min and ultrasonic grinding (Sonicator Q700; Qsonica, Newtown, CT) for 60 min. The stages of heating and grinding were switched several times for 5–6 h until the required consistency of the soap solution was reached. The 0.01 M solutions of dOA and dAA with potassium hydroxide were obtained by heating in the water bath at 40 and 50°С, respectively, for 30 min. The saline complexes with BSA (3.34 mM) were prepared to obtain a 5 mM solution of dSA-BSA, dOA-BSA, dAA-BSA in 3:1 M ratios.

Figure 1.

Structural formulas of deuterated fatty acids (a) and Raman spectra of fatty acids measured above and below the main transition temperature (b). Solid lines correspond to lipids in a disordered phase state, and dashed lines correspond to the lipids in an ordered conformational state. Spectra of different lipids are vertically shifted for clarity. To see this figure in color, go online.

Oocytes collection and in vitro maturation

All the animal experiments were approved by the Bioethics Committee of the Institute of Cytology and Genetics (protocol number 22.2 of May 30, 2014; the Siberian Branch of the Russian Academy of Sciences, Novosibirsk, Russia) and were consistent with the European Convention for the Protection of Vertebrate Animals used for Experimental and Other Scientific Purposes. The domestic cat’s ovaries were obtained after routine ovariohysterectomy from the veterinary clinic of Novosibirsk and were transported to the Institute of Cytology and Genetics during 3–4 h at 4°С in collect medium consisting of Medium 199 (Sigma-Aldrich, St. Louis, MO) supplemented with HEPES (0.01 M) and gentamicin (50 μg/mL).

The ovaries were minced, and cumulus-oocyte complexes (COC) were isolated under an M80 stereomicroscope (Leica Microsystems, Wetzlar, Germany) and placed into collect medium at 38°C. The oocytes with uniformly dark ooplasm surrounded by several layers of cumulus cells were selected for the experiment. The oocytes were cultured in 20-μL drops (3–5 COCs in a drop) of modified Medium 199 (Sigma-Aldrich) containing 10 IU/mL human chorionic gonadotropin (Chorulon; Intervet International, Boxmeer, the Netherlands), 2 IU/mL equine chorionic gonadotropin (Follimag; Mosagrogen, Russia) covered with mineral oil (FertiPro, Beernam, Belgium) in the CO₂-incubator (Galaxy 48R; Eppendorf, Hamburg, Germany) at 38.5°C, 5% CO₂, and 90% humidity. COCs were exposed to dSA, dOA, and dAA complexes with BSA in 3:1 M ratio in fatty acids concentrations of 400 μM during the entire maturation period of 42 h.

Sample freezing

Mature oocytes supplemented with deuterium-labeled fatty acids were placed in the cryoprotectant solution of 0.2 M sucrose and 1.5 M ethylene glycol dissolved in potassium simplex optimized medium KSOM without phenol red (EmbryoMax; Merck). To avoid the osmotic shock, equilibration with cryoprotectant solution was performed in three steps: 1) oocytes were placed for 3 min in a drop with a cryoprotectant concentration of one-third from the final concentration, 2) 3 min in cryoprotectant solution dissolved to one-half of the final concentration, and 3) placing oocytes in the undiluted cryoprotectant solution. Finally, the drop with oocytes was transferred on the glass plate to a cavity and covered with a mica slice. To avoid drying, the sample was sealed with petroleum jelly.

The sample was installed in optical cryostat FTIR600 (Linkam Scientific Instruments, Epsom, UK) cooled by liquid nitrogen vapor flow. Freezing was performed by the standard slow-freezing protocol used for mammalian oocytes and preimplantation embryo cryopreservation (46,47). Initially, the sample was cooled to the ice nucleation temperature T_n = −7°C at the cooling rate of 1°C/min. Ice nucleation was induced by touching the coverslip with a copper wire precooled in liquid nitrogen. After ice formation, the sample was kept at T_n from 10 to 30 min to provide ice recrystallization. The sample was cooled to −40°C (in some cases −50°C) with the cooling rate of 0.3°C/min, then after 10 min delay, it was cooled to −170°C with the cooling rate of 10°C/min. Particular temperature regimes were selected depending on the experiment type:

• 1)To carry out Raman mapping experiments and temperature dependences on cooling, the protocol was performed with interruptions at specified temperatures. Raman mapping measurements started after ∼30 min of cryostat stabilization to avoid sample movements. In temperature dependence measurements, preliminary sample holding at a given temperature was 5 min.
• 2)Raman spectra on heating were measured during sample heating from −40°C (or −50°C). The warming rate was 2°C/min; the sample was kept for 5 min at the temperature of interest before Raman recording.
• 3)To verify whether the lipid phase separation occurs in oocytes after cooling in maximal accordance to the cryopreservation protocols, the samples were cooled to −170°C, as described above, without additional delays. After ∼1–2 h of the cryostat stabilization to reduce the sample movements, the Raman mapping experiment was then carried out.

We also carried out an oocyte vitrification experiment. Oocytes were vitrified in the serum-free solution (VS) of 20% ethylene glycol, 20% dimethyl sulfoxide (DMSO), 0.5 M sucrose, and 10% Ficoll PM-70 dissolved in phosphate-buffered saline (PBS). Before vitrification, oocytes were equilibrated with a two-step method. Groups of two to three oocytes were placed in a 50-μL drop of PBS, which was mixed with 50-μL drop of equilibration solution containing 15% ethylene glycol and 15% DMSO in PBS for 3 min. 50 μL of equilibration solution was then added to the drop with oocytes for another 3 min. After equilibration, the oocytes were placed in a 20-μL drop of VS and gently pipetted for 30 s. Finally, the oocytes in a small drop of VS (<1 μL) was placed on a small piece (3 × 10 mm) of 100-μm-thin coverglass. This sample was dropped on a sample holder of the optical cryostage precooled to −170°C. We observed no signatures of ice crystallization inside or around the cell using bright-field microscopy and Raman spectroscopy, which indicated that the samples were vitrified.

Raman experiment

For Raman spectra measurement, we used a laboratory-built experimental setup described earlier (48), with minor modifications. Raman scattering was excited by the emission of a single-mode solid-state laser (Excelsior; Spectra Physics, Milpitas, CA). A microscope objective (PL FLUOTAR L; Leica Microsystems) was used to focus the laser beam on a sample and collect scattered light. Irradiation power after the objective was 15 mW. The spectrum of scattered light was measured using an imaging spectrometer (SP2500i; Princeton Instruments, Trenton, NJ) equipped with a back-illuminated CCD camera (Spec-10:2K; Princeton Instruments). The spectral resolution was 2.5 cm⁻¹. Wavelengths for all measured spectra were calibrated with the absolute precision of ∼1 cm⁻¹ using a neon-discharge lamp. Raman mapping measurements were carried out using a piezo-positioning stage PXY-200 (Newport, Irvine, CA) in a closed-loop mode. Measurement time and mapping conditions were selected depending on the experiment type:

• 1)To obtain Raman maps at −25°C, we used Raman mapping with 0.6-μm resolution and 3-s spectrum accumulation time.
• 2)For the temperature dependence study, we measured ∼100 spectra from random local points within the oocyte with an accumulation time of 10 s.
• 3)Raman maps at −170°C were measured with 1-μm resolution and 6-s accumulation time.

The spectral analysis included the preliminary corrections of intensity bursts. The baseline subtraction was done by a linear function, if not otherwise stated. To evaluate the intensity of Raman lines, the mean on three pixels with locally maximal intensities was used. The frequency of Raman peak maximum was evaluated from the second derivative analysis with preliminary application of Savitzky-Golay filter with a nine-pixels window size.

Results

Deuterium-labeled fatty acids

In our study, we used fatty acids with different deuterated moieties. Thus, to better understand how the deuterated Raman lines reflect the changes in the conformational state, we studied the Raman spectra of pure fatty acids (Fig. 1). In these measurements, small drops of fatty acids were placed on glass substrate in optical cryostage and measured at temperatures above and below melting temperature (49). Stearic acid (SA) and dSA reference Raman spectra were measured at +25 and +75°C for conformationally ordered and disordered states, respectively. Oleic acid (OA) and dOA were measured at 0 and 20°C. In the case of arachidonic acid (AA) and dAA, the ordered state was measured at −170°C and the disordered state was measured at 20°C. Deuteration resulted in a slight decrease in fatty acid melting temperatures. For stearic-d35, the transition point changed after deuteration from 70 to 67°C. In the case of partly deuterated OA, the melting point was found to be the same, 12°C, as for OA within ±1°C range. Raman lines in the range above 2800 cm⁻¹ correspond to the CH stretching vibrations, and the lines in the range from 2000 to 2300 cm⁻¹ are attributed to the CD stretching modes. We used the intensity of symmetric CH₂ stretching mode (sCH₂) at 2850 cm⁻¹ to characterize the contribution of protonated lipids. For dSA and dOA, their contributions to Raman spectra were evaluated by the intensity of the symmetric CD₂ stretching mode (sCD₂) at 2105 cm⁻¹. In the case of dAA, we investigated the intensity of Raman line at 2250 cm⁻¹ corresponding to the symmetric vibration of the methine group.

In addition to the spatial distribution analysis, Raman spectra also contain information about the conformational state of hydrocarbon chains. In the case of the CH band, conformational ordering can be extracted from sCH₂ and antisymmetric CH₂ stretching (aCH₂) Raman lines at 2850 and 2880 cm⁻¹, respectively. We used two indicators widely used in vibrational spectroscopy, the intensity ratio I(aCH₂)/I(sCH₂) (33) and the maximal frequency of sCH₂ (29), ν(sCH₂). I(aCH₂)/I(sCH₂) was evaluated as described previously (33). Because the fatty acids are differently deuterated, we used different spectral characteristics to detect the conformational changes in the Raman tags. For fully deuterated dSA, the most prominent effect of phase transition in Raman spectrum is related to the width of sCD₂. The changes of this spectral feature can be detected by evaluating the full width at half maximum (FWHM) of the sCD₂ peak and the intensity ratio between the sCD₂ peak and its right shoulder at ∼2150 cm⁻¹, I₂₁₅₀/I(sCD₂). For dOA, the Raman lines of the terminal methyl group can be used for phase transition detection. Whereas, in general, CD₃ group might not be optimal for probing the conformational state of the hydrocarbon chain, we found that, for partly deuterated dOA, Raman lines of the methyl group are more useful than the deuterated methylene moiety. The use of the CD₃ group opens the possibility for simultaneous study of two different deuterated lipids. We used the intensity ratio I₂₁₃₀/I(sCD₃) to quantify the relative intensity changes between the peak at ∼2130 cm⁻¹ (Fermi resonance of symmetric CD₃ (sCD₃) stretching vibration (50)) and the symmetric CD₃ stretching mode at 2070 cm⁻¹. In the case of dAA, the Raman lines of methine CD vibrations appear to be almost insensitive to the changes in the conformational state of the hydrocarbon chain.

Lipid separation at T = −25°C

We measured the Raman maps of cat oocytes preliminary cultured within media supplemented with different deuterated fatty acids. The introduced deuterated fatty acids can be catabolized via β-oxidation or used as a substance for the formation of phospholipids and triglycerides. We did not find any differences between the Raman spectra of pure fatty acids and those introduced into oocytes. Thus, most fatty acids are expected to participate in the formation of LDs and membranes. Because the membranes and phospholipid monolayers at the surface of LDs are much thinner than the characteristic size of the volume from which Raman light scattering is collected, their contribution to measured Raman spectra is much less than the Raman signal from the bulk triglycerides inside LDs. Therefore, unless otherwise stated, under the accumulated fatty acids, we will mean the corresponding hydrocarbon chains of triglycerides within the LDs.

To study the lipid separation, we excluded the areas in which the intensity of the contribution from the CH and CD bands was below the average (38). This approach allows isolating the LDs on the maps and studying the intensity ratios of Raman lines without distortions associated with low signal-to-noise ratio for lipid Raman lines in Raman spectra of cytoplasm. Fig. 2, a, e, and j show the spatial distribution of the ratio between Raman lines of deuterated (new) and protonated (old) lipids. The average intensity ratio between dSA and protonated lipids in Raman spectra is ∼0.14 that corresponds to ∼12 mol% dSA concentration when the equal Raman scattering cross sections for deuterated and protonated compounds are assumed (see Supporting material). In this estimation, we also employ the assumption that dSA can represent an average lipid in LDs. This evaluation is in agreement with our previous measurements (38). For dOA, which is partly deuterated, the intensity ratio I(sCD₂)/I(sCH₂) in its Raman spectrum is ∼0.36 (see Fig. 1 b). Therefore, we can estimate for Fig. 2 e that the average concentration of dOA accumulated in cat oocytes is 29%. In the case of dAA, the evaluation of its concentration is problematic because this fatty acid much differs from the average hydrocarbon chain in LDs of cat oocytes. For estimation of the dAA fraction, we used intensity ratios between the Raman peak of stretching C=C related to dAA at 1633 cm⁻¹ and C=C peak related to nondeuterated lipids at 1658 cm⁻¹. We can estimate that the addition of dAA increases the concentration of double bonds by ∼12%, and its fraction was ∼2%. Different fractions of different fatty acids result from different capabilities to form complexes with BSA or different propagation through the cytoplasmic membrane (51). A detailed description of how the fractions of deuterium-labeled hydrocarbon chains were estimated can be found in the Supporting material.

Figure 2.

Raman mapping of LDs in oocytes supplemented with deuterated fatty acids and cooled to −25°C. The first row of panels shows the data corresponding to the oocyte cultured with dSA. (a) Map of I(sCH₂)/I(sCD₂) ratio in LDs; (b, c, and d) different fragments of Raman spectra corresponding to high and low values of I(sCH₂)/I(sCD₂) ratio. The second row shows the data corresponding to the oocyte cultured with dOA. (e) Map of I(sCH₂)/I(sCD₂) ratio in LDs; (f, g, and h) different fragments of Raman spectra corresponding to high and low values of I(sCH₂)/I(sCD₂) ratio. The third row shows the data corresponding to the oocyte cultured with dAA. (i) Map of I₂₂₅₀/I(sCD₂) ratio in LDs; (j, k, and l) different fragments of Raman spectra corresponding to high and low values of I₂₂₅₀/I(sCD₂) ratio. The red lines are Raman spectra of LDs with a high fraction of deuterated lipids (the highest quartile), and the blue line is a spectra with a low fraction of deuterated lipids (the lowest quartile). To see this figure in color, go online.

A comparison of deuterated fatty acid distribution indicates that dSA are located predominantly in the peripheral regions of LDs (Fig. 2 a). In contrast, monounsaturated dOA and polyunsaturated dAA tend to distribute within a core region of LDs (Fig. 2, e and j). This distribution agrees with previous results indicating that lipids in the central core of LDs contain more unsaturated lipids (16). To verify that the inhomogeneity in labeled fatty acid distribution is not related to the overlay of different cell organelles, we measured Raman spectra of isolated LDs (Fig. S3) that proved phase separation interpretation.

To evaluate the degree of lipid separation, we compared the Raman spectra corresponding to the highest and the lowest quartiles of I(sCD₂)/I(sCH₂) and I₂₂₅₀/I(sCH₂) ratio distributions for the maps in Fig. 2. Scaled by the intensity of the symmetric CH peak at 2850 cm⁻¹ (see Fig. 2, d, h, and l), these spectra reveal the changes in the contribution of deuterated lipids (Fig. 2, c, g, and k) and their relations to the C=C band intensity at 1658 cm⁻¹. The intensity of the last band is proportional to the concentration of C=C bonds and is often used to evaluate the lipid unsaturation degree. In dAA, this band is slightly shifted to 1633 cm⁻¹ (Fig. 2 j). Thus, its contribution to the intensity of the main C=C band is negligible.

From the spectra presented in Fig. 2, we can learn that the intensity of dSA contribution in the spectrum of the ordered phase is at least two times higher than in the disordered phase (15 vs. 8.5% from overall lipid content). The disordered phase contains more unsaturated lipids. For monounsaturated dOA, the concentration is 1.34 times higher for the disordered phase (33.7 vs. 25% from overall lipid content), and for polyunsaturated dAA, the concentration is at least 1.44 times higher in the disordered phase (estimated fraction ∼2.6 and ∼1.9% from overall lipid content).

The size of observed domains is the same order as the spatial resolution of our experiment. Thus, most of the measured spectra are the superposition of contributions from different phases and do not accurately determine the fraction of deuterated molecules in each phase. To overcome this problem, we applied a non-negative matrix factorization (NNMF) approach (52) that decomposed the Raman mapping data on three components corresponding to the cytoplasm and two different conformational states of lipids inside LDs (16). Fig. 3 shows the spectral components derived from NNMF decomposition. Their spatial distributions are in good agreement with the maps shown in Fig. 2.

Figure 3.

Spectral components derived from the Raman mapping data of deuterated oocytes using multivariate analysis. (a) and (b) show the spectral components in the ranges of CD and CH stretching vibrations, respectively. The black dashed line is the component attributed to cytoplasm contribution, the red line corresponds to disordered lipids, and the blue line is the contribution of ordered lipids. Spectra of differently deuterated oocytes are vertically shifted for clarity. (c) shows the CD band of dOA of different spectral contributions. To see this figure in color, go online.

For oocytes with dSA, NNMF decomposition indicates that the Raman mapping data can be described by the ordered phase containing almost all dSA (only ∼10% are attributed to other spectral components). Its fraction in conformationally ordered lipids was estimated to be ∼19%. The spectral component corresponding to cytoplasm also has a detectable lipid contribution (from both dSA at 2105 cm⁻¹ and unlabeled lipids at 2848 cm⁻¹). This is because there are many LDs in the mapping area (Fig. 2 a), and all spectra of this map contain some residual lipid contribution from off-focus lipids. No additional СD peaks in the residual spectra that could indicate the participation of dSA in cellular metabolism were found. The magnitude of observed residual deuterated lipid contribution reflects the accuracy of NNMF decomposition. In the case of dOA, we can see that oleic hydrocarbon chains are presented in both ordered and disordered phases. Its abundance is 1.4 times higher in the disordered phase (estimated fraction 38 vs. 27% in the ordered phase) (Fig. 3). dAA can be found in both phases and is 1.45 times more abundant in the disordered phase. Notably, the spectral components of dOA in ordered and disordered phases demonstrate the different spectral shapes of the CD band, confirming that, inside an LD at T = −25°C, oleic hydrocarbon chains are present in ordered and disordered conformational states.

Temperature dependence of lipids phase state

The coexistence of lipid domains with different LPT temperatures broadens the overall LPT transition. In this case, within the LPT temperature range, the coexistence of lipid domains of liquid and solid phases is expected. We observed the phase separation in LDs at different temperatures during cooling (Fig. 4). However, Raman mapping at room temperature did not reveal any phase separation, even for dSA, which is the most infusible.

Figure 4.

The bright-field microscopy and Raman mapping of LDs at different temperatures inside cat oocyte supplemented with dSA. To see this figure in color, go online.

Raman mapping experiments require time-consuming cryostage stabilization, whereas single LD measurements do not give information about the average phase state of lipids. Thus, we carried out experiments in which ∼100 Raman spectra were measured from random points within 50- × 50-μm area inside the oocyte under each temperature. Using noncentered principal component analysis for data decomposition, the representative lipid contribution was extracted (28). Because the main variance in the obtained spectra is associated with the difference between the cytoplasm and lipid contributions, it was sufficient to combine the first two principal components to separate these two spectral contributions. We used the constraint that the extracting spectrum obtained from the combination of spectral representations of the first two principal components should not have any Raman lines in the spectral range of the OH stretching band (above 3050 cm⁻¹). Further, we analyzed the temperature dependence of the extracted lipid contribution.

We studied the oocytes cultured with dSA and dOA because those Raman spectra are more sensitive to the chain conformation changes. Fig. 5 shows the temperature dependences of the main characteristics of the Raman spectrum, which provide information on the lipid phase state. Because the C=O band is multicomponent at low temperatures (33), we evaluated its frequency using centroid. It can be seen that I(aCH₂)/I(sCH₂) ratio for oocytes supplemented with dSA is always higher than for dOA containing oocytes. One also can note that for oocytes with dOA the onset of the LPT occurs between 0 and 5°C because above this temperature I(aCH₂)/I(sCH₂) ≈ 0. This temperature of LPT onset agrees well with the previous study of cat oocytes (33). In the case of dSA, I(aCH₂)/I(sCH₂) is nonzero, even at 40°C that is above the temperature of oocyte culture. Temperature dependences of ν(sCH₂) and ν(C=O) are in qualitative agreement with I(aCH₂)/I(sCH₂) ratio temperature dependence. At low temperatures, frequencies of sCH₂ and C=O modes appear to be higher for dSA samples than for dOA. This may result from the difference in the LDs composition after the addition of different fatty acids. It is noteworthy that, when measuring Raman spectra from oocyte on cooling, I(aCH₂)/I(sCH₂) ≈ 0 at 25°C, and the temperature dependence for ν(sCH₂) also demonstrates the difference between cooling/heating directions. The difference in the parameters of Raman spectra at the same temperature on cooling and then on heating indicate nonequivalence of lipid states before and after cooling.

Figure 5.

Temperature dependences of spectral characteristics of lipid contribution in Raman spectra measured from cat oocytes cultured with dSA and dOA. Temperature dependences show in order from top to bottom: (a) I(aCH₂)/I(sCH₂), (b) ν(sCH₂), (c) ν(C=O), (d) I₂₁₃₀/I(sCD₃), (e) I₂₁₅₀/I(sCD₂), and (f) FWHM of sCD₂ peak. Red dashed lines are linearly interpolated averaged data, and red triangle symbols are experimental data for oocytes cultured with dSA. For dOA, blue dashed lines are linearly interpolated average data, and blue symbols are experimental data measured at heating. Empty dark red circles are data obtained on cooling an oocyte cultured with dSA; the filled dark red circle is the datapoint measured for the same sample on heating. The blue vertical dotted line denotes the temperature of dOA rich-phase melting. The black vertical dotted line denotes the LPT onset temperature observed in dOA-supplemented oocyte. Temperature dependences were obtained on heating from low temperatures. To see this figure in color, go online.

When considering the spectral features of deuterated lipids, one can find that the lipid fraction containing dOA melts between −13 and −10°C (Fig. 5 d). This peculiarity is less prominent but also can be detected in temperature dependences of I(aCH₂)/I(sCH₂), ν(sCH₂), and ν(C=O). For the dSA fraction, we observe gradual changes in I₂₁₅₀/I(sCH₂) (Fig. 5 e) and FWHM of sCD₂ mode (Fig. 5 f). This behavior indicates that the fraction of the ordered phase gradually decreases with the temperature increase. The difference between oocytes cultured with dOA and dSA qualitatively correlates with the temperature behavior of triolein melting near −5°C (33,53), and the melting point of stearin is 54 or 72.5°C, depending on polymorph (54). The FWHM of sCD₂ mode for dSA in the liquid phase is ∼40 cm⁻¹ (see Fig. 5 f; (55)). The maximal width evaluated in our experiments after oocyte cooling was only 34 cm⁻¹. Linear extrapolation of our data estimates that 40-cm⁻¹ width could be reached between 60 and 70°C. This indicates some fraction of dSA remains in the ordered phase state after cooling, even at physiological temperatures. It is necessary to note that, before cooling, at 25°C, Raman spectra demonstrate that deuterated lipids are in the disordered conformational state (Fig. 5, e and f). Thus, these data show that saturated lipids within LDs can undergo an irreversible transition to the conformationally ordered state.

Comparison of lipid distribution for different cryopreservation protocols

The experiments described above were performed with some deviations from the standard protocol for slow freezing of oocytes. To ensure that the phase separation occurs in cells cooled with the standard freezing protocols, we carried out Raman mapping of oocytes cooled to −170°C with the program freezing and vitrification. At low temperatures, light scattering spectra of the oocytes demonstrated the high level of photoluminescence, limiting the precision of the Raman experiment. The background under sCD₂ and sCH₂ bands was interpolated using a cubic polynomial. In addition, the experimental Raman maps were slightly distorted because of instabilities in the sample position at low temperatures. Fig. 6 shows the maps of I(sCD₂)/I(sCH₂) ratio distribution. It can be noted that, for the slowly frozen oocytes, oval-like structures were observed associated with the increased concentration of dSA at the periphery of LDs. In the case of vitrified oocytes, the main differences in concentrations of dSA are related to the variation of dSA content between different LDs. Similar content variation between different LDs was observed earlier at room temperature (38).

Figure 6.

Comparison of I(sCD₂)/I(sCH₂) maps obtained at T = −170°C from the oocyte cultured with dSA and frozen using program freezing protocol (left) and vitrified oocyte (right). To see this figure in color, go online.

Thus, we conclude that the phase separation in LDs does occur during programmed freezing. Comparison with vitrified oocytes indicates that vitrification protocols can be used to overcome the cryoinjuries related to lipid phase separation, at least at the cooling stage.

Discussion

The use of deuterated lipids as Raman labels was proposed several decades ago (55,56). However, their application to living cells began only in recent years (34, 35, 36, 37, 38, 39, 40). Compared to more common fluorescence labeling, the Raman signal of deuterium-labeled compounds can be normalized to the contribution of unlabeled, protonated material. Interpretation of Raman measurements is also easier than fluorescence, especially when chemical reactions or phase states of lipids are studied. The above considerations and our experimental results indicate that the Raman spectroscopy of deuterated lipids is a method of choice for studying phase transitions and phase separation directly inside living cells. In addition, Raman approaches could be combined with fluorescence microscopy and ²H-NMR (57). Although, in this work, we used high concentrations of labeled fatty acids, which markedly affect LPT, it is possible to detect concentrations an order of magnitude lower, at which the change in the composition of LDs would be negligible.

We did not find spatial heterogeneities in the distribution of deuterated phospholipids within LDs at room temperature that agree with previous observations (38). Our data show that the uncooled oocytes at room temperature contain lipids in a liquid disordered phase. This state was observed for oocytes cultured with dSA, dOA, and dAA. Depending on the type of fatty acids added to oocytes, we found that the temperature effects on the phase state of different lipid fractions within LDs.

When oocytes were cultured with dSA and as a result were enriched with the saturated lipids, the onset of LPT occurs during cooling at temperatures between 15 and 25°C. Further cooling leads to the coexistence of two phases with conformationally ordered and disordered lipids. Temperature decrease gradually increases the fraction of lipids in the ordered state with the formation of domains a couple of microns in size. Saturated lipids demonstrate the highest difference in content between the two phases, being several times more abundant in the ordered phase. According to Raman mapping data, the ordered phase enriched in saturated fatty acids tends to be distributed at the periphery of LDs. We suggested that this distribution might result from heterogeneous nucleation of the ordered phase (16). Hypothetically, the excessive distribution of ordered phase near LD’s surface might affect the phase state and functional properties of the surface monolayer. After cooling and the subsequent warming, some portion of the saturated lipids remains in the ordered state, even at 40°C, which is slightly above the optimal physiological temperature for cat oocytes. If we assume that lipids must be in a disordered phase state to participate in biological processes, the ordered phases at physiological conditions can disrupt the cellular processes. This can be one of the sources for cryoinjuries associated with LDs.

It can be supposed that LDs of cat oocytes are in a homogeneous long-living metastable state, like a supersaturated solution. At room temperatures, some portion of saturated lipids can exist in a more energetically favorable solid ordered state, but they are dissolved in the fraction of fusible unsaturated lipids in a disordered conformational state. In this solution, saturated lipids also remain in disordered conformations that hinder the formation of nuclei of solid phases in which all lipids should be in the conformationally ordered state. During cooling, conformational ordering becomes more favorable, which triggers the transition of saturated lipids in a solid ordered state. After rewarming, the lipid phases inside the LDs become equilibrated. Most likely, some of the saturated lipids remain in the solid ordered phase. On the other hand, one cannot exclude that homogeneous state is stable, and after prolonged cell culture, ordered phase domains in LDs will melt. It is known that, at room temperatures, phospholipid mixture of 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) and 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) is homogeneous at DPPC concentrations below 40% and exhibits phase coexistence at higher concentrations (9,10).

For oocytes cultured with unsaturated fatty acids, we observed the onset of LPT between 0 and 5°C. This temperature is close to one reported earlier (33). We did not find changes for the CD band of dOA at this temperature, indicating that unsaturated hydrocarbon chains are not directly involved in the LPT onset. In the case of monounsaturated dOA hydrocarbon chains, the transition into an ordered conformational state starts between −13 and −10°C. This observation illustrates the multiplicity and complexity of LPT in biological LDs, in which the composition of ordered and disordered phases changes with temperature. In contrast to saturated lipids, unsaturated lipids are more abundant in disordered phases and tend to redistribute to the core of LDs. Raman mapping indicates that unsaturated lipids demonstrate lower concentration gradients between ordered and disordered phases than saturated lipids. It can be concluded that the addition of unsaturated lipids has little if any effect on the onset of the phase transition, which is determined by the concentration of saturated phospholipids. Instead of lowering the temperature of LPT onset, the additional unsaturated lipids can undergo a separate transition at low temperatures. On the other hand, the addition of unsaturated lipids can reduce the relative proportion of saturated lipids, which can slightly shift the temperature of the LPT onset.

We have unequivocally found that the lipid phase separation occurs during the standard programmed freezing protocol. It is important to note that in cells at −170°C all lipids are in an ordered conformational state. Using deuterated labels greatly simplified the observation of the phase separation at low temperatures. In the case of vitrification, we did not observe signs of the phase separation on a scale of a few microns, which indicates a specific advantage of this approach over the programmed freezing protocol. However, our data do not exclude the presence of submicron domains of saturated phospholipids that can persist after warming the vitrified cells to physiological temperatures. Further research may shed light on this question. It is noteworthy that the discussed phase transition occurs at relatively high temperatures and can be related not only to cryopreservation but also to cold adaptation and cryotolerance.

Many years ago, optical cryomicroscopy opened the way for the practical study of water crystallization and the changes in osmotic pressure on a cell during freezing (58,59). These studies have largely contributed to cryobiology and have become the key to the development of applied cryopreservation. Spectroscopical methods, such as infrared (31) and Raman spectroscopy (60) as well as Raman mapping (61) and coherent Raman microscopy, enforced with the usage of isotope tags, (40) and ²H -NMR (15) offer opportunities for detailed study of those phenomena that until recently have been hidden from direct observations. We believe that these methods will allow obtaining new experimental observations that form the basis of new strategies and approaches for cryopreservation.

Conclusions

Raman spectroscopy was applied to study the problems of the phase transitions and the phase coexistence in the LDs upon cat oocyte cooling. Deuterium-labeled fatty acids were used to distinguish spectral contributions from different lipids. We investigated the distributions of hydrocarbon chains that were saturated, monounsaturated, and polyunsaturated. The major results can be summarized as follows:

• 1)Raman mapping of LDs in oocytes at −25°C allowed us to evaluate the difference in lipid content of the coexisting phases. The greatest concentration gradient was found for dSA representing lipids with saturated hydrocarbon chains. The lower estimate is that the dSA concentration is two times higher within the ordered phase. The decomposition of Raman mapping data using the NNMF approach indicates that almost all dSA (>90%) can be accounted to the ordered phase. The concentration of unsaturated lipids containing dOA and dAA is ∼1.5 times higher for the disordered phase.
• 2)Temperature dependences for Raman lines of protonated and deuterated lipids allowed us to detect the phase transitions of different lipid fractions and investigate the lipidome modification effects. For oocytes cultured with dSA and underwent cooling below 5°C, a small fraction of lipids remains in the ordered conformational state even at 40°C. For oocytes cultured with dOA, one can see the partial transition involving deuterated lipids at approximately −12°C and the LPT onset between 0 and 5°C. Therefore, we can conclude that the content of saturated lipids mainly determines the LPT onset temperature.
• 3)We verified that the lipid separation occurs in standard cryopreservation protocols. Lipid separation was observed at −170°C in cat oocyte cooled with the slow-freezing protocol but not in the vitrified one.

Our results demonstrate the capability of Raman spectroscopy of deuterated lipids to dissect the LPTs in complex biological systems. Deuterated lipids can provide information not only about their spatial distribution but also about their conformational state. This approach is a powerful tool to investigate how different strategies for modifying the lipid composition affect the state and distribution of lipids at physiological temperatures and during cell cryopreservation.

Author contributions

N.V.S., S.Y.A., S.V.O., and K.A.O. designed research. S.V.O. performed sample preparation. K.A.O. performed Raman experiments. V.I.M. participated in oocyte vitrification experiments. K.A.O. and N.V.S. analyzed data. K.A.O. wrote the manuscript with contributions from all coauthors.

Editor: Georg Pabst.