Factors Affecting the Membrane Permeability Barrier Function of Cells

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Biological and Environmental Phenomena at the Interface

Factors affecting the membrane permeability barrier function of cells during preservation technologies Willem Wolkers, Harriette Oldenhof, Fengrui Tang, Jiale Han, Judith Bigalk, and Harald Sieme Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.8b02852 • Publication Date (Web): 02 Dec 2018 Downloaded from http://pubs.acs.org on December 6, 2018

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FACTORS

AFFECTING THE MEMBRANE PERMEABILITY BARRIER FUNCTION OF CELLS

DURING PRESERVATION TECHNOLOGIES

Willem F. Wolkers1, Harriëtte Oldenhof2, Fengrui Tang1, Jiale Han1,2, Judith Bigalk2, Harald Sieme2

1Institute

of Multiphase Processes, Leibniz Universität Hannover, Germany, 2Unit for

Reproductive Medicine, Clinic for Horses, University of Veterinary Medicine Hannover, Germany

*corresponding

author: Willem F. Wolkers, Institute of Multiphase Processes, Leibniz

Universität Hannover, Germany, Callinstrasse 36, 30167 Hannover, Germany; phone: +49 511 762 19353, fax: +49 511 762 19389, e-mail address: [email protected]

keywords: membrane phase behavior, membrane permeability, membrane hydraulic permeability,

cryopreservation,

freeze-drying,

porcine

oocytes,

phosphatidylcholine

liposomes, fibroblasts

ACS Paragon Plus Environment

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ABSTRACT

Cellular membranes are exposed to extreme conditions during the processing steps involved in cryopreservation (and freeze-drying) of cells. The first processing step involves adding protective agents. Exposing cells to protective agents causes fluxes of both water and solutes (i.e. permeating cryoprotective agents) across the cellular membrane, resulting in cell volume changes and possibly osmotic stress. In addition, protective molecules may interact with lipids which may lead to membrane structural changes and permeabilization. After loading with protective agents, subsequent freezing exposes cells to severe osmotic and mechanical stresses, caused by extra and/or intracellular ice formation, and a drastically increased solute concentration in the unfrozen fraction. Furthermore, cellular membranes undergo thermotropic and lyotropic phase transitions during cooling and freezing, which drastically alter the membrane permeability and its barrier function. In this review, it is shown that membrane permeability to water and solutes are dependent on temperature, medium osmolality, types of solutes present, cell hydration level, as well as the absence or presence of ice. Freezing most drastically alters the membrane permeability barrier function, which is reflected as a change in the activation energy for water transport. In addition, membranes become temporarily leaky during freezing-induced fluid-to-gel membrane phase transitions resulting in the uptake of impermeable solutes.

1. INTRODUCTION

Cryopreservation can be used to store cells, tissues and biological materials that are used in therapeutic applications until they are needed. The processing steps that are involved in these


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preservation methods, however, expose cells to severe stresses and protective agents are needed to minimize damage. Causes of stress include osmotic imbalances, mechanical forces and cytotoxicity for solutes for which cells have distinct tolerance limits. Cells encounter severe hypertonic conditions upon addition of high concentrations of protective agents as well as during freezing. During thawing or rehydration the reverse process takes place, and cells experience hypotonic conditions resulting in water uptake and swelling. Especially the membrane permeability to water and solutes determine the ability of cells to cope with cryopreservation-induced osmotic stresses. In addition, thermotropic and lyotropic lipid phase transitions and lipid-solute interactions may alter membrane organization and structure as well as its permeability characteristics. Development of cryopreservation methods for cells and tissues requires insights in such membrane features. In this review, we will discuss (1) membrane permeability to water and solutes derived from osmotically induced cell volume responses, (2) interactions between membranes and protective solutes, and (3) membrane phase and permeability behavior of cells under extreme conditions. Furthermore, we will discuss how insights in factors determining and affecting the membrane permeability barrier function can be used to develop cell preservation technologies.

2. MEMBRANE PERMEABILITY TO WATER AND SOLUTES

2.1 Cell volume response in hypo or hypertonic solutions containing non-permeating solutes to determine membrane permeability to water: cell shrinkage and swelling under anisotonic conditions, osmosis, Boyle van ’t Hoff behavior, isotonic and osmotically inactive volume, membrane hydraulic permeability, rectification


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Cells respond to changes in the external medium osmolality by water transport across the cellular membrane to maintain equilibrium between the intra and extracellular solute concentration. As a consequence, the cell volume changes and cells shrink and swell in hyper and hypotonic media, respectively (figure 1A−C). The cell volume in isotonic medium (of ~300 mOsm) is referred to as Vo. The relationship between cell volume and medium osmolality gives a linear correlation in a so-called Boyle-van ’t Hoff plot, from which the osmotically inactive volume (Vb) can be derived by extrapolating to infinite osmolality (figure 1C). Vb exhibits differences amongst different cell types and developmental stages. The osmotically inactive volume of red blood cells, for example, comprises 47% of the isotonic cell volume (i.e. Vb equals 0.47×Vo).1 For oocytes, Vb values around 20% have been reported,2−4 whereas for sperm Vb ranges from 50 to 80%.5−7 Non-viable cells typically do not respond to changes in medium osmolality, observed as deviation from linearity in a Boyle-van ’t Hoff plot.8 Inclusion of such data points affects calculated Vb values. Boyle-van ’t Hoff behavior only gives a static picture of the cell volume response to anisotonic media. The rate of the change in cell volume versus incubation time can be used to determine the cell membrane permeability to water (Lp). Cell volume response measurements can be done using a microscope with a micromanipulator setup including a holding pipette,9 microfluidic devices,10 or by electronic particle size (Coulter counter) measurements.11,12 Lp governs the kinetics of cell volume changes when a cell is subjected to conditions that depart from isotonic conditions. An established model can be used to fit experimental data on cell volume responses of exposed to anisotonic solutions containing only non-permeating solutes to derive Lp values.13,14 The membrane permeability to water primarily depends on the cellspecific membrane composition and on the temperature. However, Lp is also affected by the presence of solutes and their concentration.12,15 Figure 1D illustrates how medium osmolality affects the Lp of porcine oocytes.


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Differences between Lp values that have been determined under hypotonic conditions compared to those determined under hypertonic conditions have been described as rectification, which refers to an apparent difference in water permeability depending on the direction of water flow (i.e., into the cell vs out of the cell). Rectification has been reported for a variety of cell types.16,17 For IVM oocytes, it appears as if rectification is observed when comparing Lp values obtained at 240 mM to those obtained at 450 mM and higher. However, at 180 mM, the Lp values for both immature germinal vesicle (GV) stage in vitro matured (IVM) oocytes are not significantly different compared to those at 450 mM or higher. Therefore, we are cautious to interpret the effects of medium osmolality on the Lp of porcine oocytes in terms of rectification. On the one hand, the increased salt concentration under hypertonic conditions alters membrane structure and permeability.18 It has been reported that sodium and potassium monovalent cations interact with carbonyl oxygen groups of phosphatidylcholine lipids resulting in altered lipid diffusivity.19 In addition, the further from equilibrium, the more the cell is exposed to stress causing cells to lose their membrane integrity. If such cell volume measurements are done in either hypertonic or hypotonic media close to the osmotic tolerance limits of the cell this will affect the observed Lp.

2.2 Cell volume response during loading with permeating cryoprotective agents to simultaneously determine membrane permeability to water and solutes: water and solutes move with different velocities across membranes, cell volume excursions, interaction between solute and water transport

Cryopreservation requires loading of cells with protective agents to preserve intracellular structures. Permeating CPAs like ethylene glycol, dimethyl sulfoxide and


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propylene glycol can pass the cellular membrane, whereas disaccharides typically cannot. The diffusivity of permeating CPAs across cellular membranes, however, is slower than that of water.14 This is evident from the cell volume response during exposure to a solution with permeating CPAs, which shows two phases (figure 2A−C). Exposing cells to a cryopreservation solution, which typically contains molar concentrations of CPAs, creates an osmotic gradient causing water to move out of the cells. After this initial cell volume decrease, both permeating CPAs and water move into the cell until equilibrium is reached between the extra and intracellular osmolality. A number of formalisms are available that can be used to fit this biphasic volume response, using transport equations describing the water and solute/CPA flux, to determine the cell membrane permeability for water (Lp) and solutes (Ps). These include the so-called one parameter (1P) or solute permeability model,20 the two parameter (2P) model in which water and solute transport are considered as independent processes,21 and a three parameter (KK) model which takes solute-solvent interactions during transport across the cellular membrane into account.13 Interactions typically occur during co-transport through pores like aquaporins. However, despite the fact that some members of the aquaporin protein family are permeated by glycerol, typically only water is selectively transported through aquaporins while co-transport of CPAs is negligible.22 All models mentioned above are thermodynamic models which assume a homogeneous membrane, thus ignoring the microdomain structure of the cellular membrane. Whereas in earlier studies predominantly the KK model was applied to derive membrane transport parameters of cells, later studies have shown that the 2P model is simpler to use and equally accurate.14 The 2P model can be used in case there is absence of co-transport; passive diffusion of water and solute across the lipid bilayer, passive solute transport and water transport through selective channels, and transport of water and solute through separate channels.


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Oocyte volume measurements were done using a micropipette perfusion technique.9 Oocytes were held in a droplet using a micromanipulator setup, and subsequently flushed with CPA solution using a syringe while acquiring a video. The data presented Figure 2 A-C have been fitted using the 2P formalism.14 The change in cell volume during the perfusion process (dV/dt), at a particular temperature (T (297.15 K)) is described as a function of both the water volume (VW): [1]

𝑑𝑉𝑤 𝑑𝑡

= ― 𝐿𝑃𝐴𝑅𝑇[𝑀𝑒 ― 𝑀𝑖]

and the solute volume (VS): [2]

𝑑𝑉s 𝑑𝑡

= 𝑃𝑆𝐴 (𝑀𝑠𝑒 ― 𝑀𝑠𝑖)𝑉𝑠

here, Vs is the partial molar volume of the CPA, M the osmolality, w refers to water, s to permeating, while e and i refer to external and internal cellular locations. R is the universal gas constant. Cell specific parameters including the area (A), isotonic cell volume (V0) and osmotically inactive cell volume (Vb) were used to derive intracellular concentrations. Vb values for IVM and GV oocytes were derived from a Boyle van ‘t Hoff plot (see inset Figure 1C). Differences in solute permeability of CPAs are reflected in the shape of the cell volume response, i.e. the extent of dehydration prior to returning to the original cell volume as well as the rate at which this occurs. Lp and Ps values of different CPAs are dependent on the cell developmental stage (compare GV and IVM), whereas Lp values can be affected by the solutes present in the formulation (figure 2D−E). These factors make it difficult to compare values of membrane transport parameters in the literature.10,23−25

2.3 Factors affecting membrane permeability to water and solutes: cell stage and membrane composition, hypo-/hypertonic conditions, solute type(s) and concentration, temperature


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Membrane permeability to water and solutes determine the ability of cells to cope with cryopreservation-induced osmotic stresses. Therefore, quantitative understanding of the membrane transport properties is important in order to rationally design preservation protocols to avoid large volume excursions.26 The temperature dependency of the membrane permeability to water and solutes exhibits Arrhenius behavior. The Arrhenius equation for Lp is given by:

 ELp  1 1   Lp  Lpg exp       R  T TR  

[3]

An Arrhenius plot of experimentally determined Lp values can be used to derive the activation energy for water transport (ELp) and Lp at a reference temperature, typically 0°C (Lpg). Water transport directly through the lipid bilayer via solubility-diffusion is characterized by relatively high ELp values, whereas transport through aquaporins decrease ELp.27,28 Membrane transport parameters can be used to predict cell volume responses during the various steps of cryopreservation processing: CPA loading, freezing, thawing, and CPA unloading. This can help to avoid large volume excursions. Ideally, prediction of the cell volume response during an entire cryopreservation process requires knowledge of Lp values determined under hypertonic, hypotonic, supra and subzero conditions, as well as knowledge on the temperature dependence of Lp. Lp values determined under hypertonic conditions (i.e. during CPA loading) cannot necessarily be used to predict cell volume responses under hypotonic conditions (i.e. during thawing or CPA unloading). In addition, Lp values determined at suprazero temperatures cannot necessarily be used to predict the cell volume response during freezing (see section 3).

3. INTERACTIONS

BETWEEN MEMBRANES AND PROTECTIVE AGENTS AFFECT MEMBRANE


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PERMEABILITY

3.1 Cell membrane permeabilization upon exposure to vitrification solutions: concentration -and temperature- dependency, pre-/post-freeze toxicity/leakiness, multi-step vitrification protocols

As indicated above, membrane permeability of a cell to water is not a constant value. Solutes (e.g. salts, CPAs) may alter membrane structure and permeability by interacting with the lipid headgroups or by partitioning in the hydrophobic bilayer. In addition, cell volume changes upon CPA addition may cause leakage. Addition of high CPA concentrations, as typically used in case of vitrification, can disturb membrane integrity causing leakage of intracellular constituents and/or uptake of impermeable solutes (i.e. dependent on the osmotic driving force). Cell membrane permeabilization, inferred from uptake of membrane impermeable solutes/dye, takes place during incubation in solution containing permeating CPAs in a dose- and time-dependent manner (figure 3A). Membrane permeabilization upon exposure to vitrification solution (VS) is reduced after pre-incubation in equilibration solution (ES) (Figure 3B). Thus, the extent of membrane permeabilization taking place when oocytes are directly exposed to a vitrification solution can be reduced by gradually increasing the CPA concentration. This reduces the osmotic shock and limits the total volume excursion compared to cells that are directly exposed to full strength vitrification solution.26 Exposure times to full strength vitrification solutions, however, should be limited in order to minimize membrane permeabilization, maintain cytoskeleton features3 and physiological functions (figure 3B-inset). In a typical vitrification protocol, cells are first equilibrated in solution(s) with relatively low CPA concentrations. After this, cells are transferred to full strength vitrification


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solution, to dehydrate the cells, and to avoid intracellular ice formation and to facilitate extraand intra-cellular glass formation during cooling.29 Mixtures of permeating CPAs are used to avoid crystallization of single components and to minimize cytotoxic effects. Nonpermeating disaccharides are typically added to vitrification and/or warming solutions, to counteract hypotonic stress during thawing.

3.2

Liposome

model

studies

for

investigating

lipid-solute

interactions:

dye-

retention/leakage studies, phospholipid hydration, preferential exclusion, partitioning

Liposomes with entrapped carboxyfluorescein/dye can be used as a model system to study membrane protection and leakiness during handling in different CPA formulations.30,31 Liposome stability is evident as dye retention within the liposomes. Adding CPAs eventually causes leakage, while distinct differences in the concentration dependency can be seen amongst glycerol, ethylene glycol, and dimethyl sulfoxide (Figure 3C). Membrane permeabilization effects of dimethyl sulfoxide have been studied in detail,32−34 and appear to be different from other CPAs.35 Different behavior amongst CPAs is related to differences in CPA-lipid interactions; which can be studied using infrared spectroscopy.30 Dimethyl sulfoxide tends to dehydrate phospholipid headgroups while raising the membrane phase transition temperature (Tm), whereas glycerol causes an increase in the hydration level of the lipid headgroups while decreasing Tm (Figure 3E−F). Also changes in liposome volume may cause leakage. Glycerol, which causes the most dye leakage, is expected to be the least permeable and cause the largest volume changes. Increasing the cholesterol/lipid ratio affects membrane fluidity and phase behavior (i.e. increases and decreases fluidity above and below Tm, respectively),36 and increases membrane stability.37 Furthermore, in case of phosphatidylcholine liposomes, exposure to higher CPA concentrations is tolerated without


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membrane permeabilization. Interactions between solutes and membranes have been classified into alcohol- and sugar-like mechanisms.38 The alcohol-like mechanism is related to the hydrophobic properties of the solute. Hydrophobic molecules preferably partition into the lipid bilayer. The sugar-like mechanism is related to the extent to which the solute accumulates on the surface of the membrane. Solute effects on membranes have also been divided into kosmotropic and chaotropic effects depending on the water structuring properties of the solute.39 Most CPAs are kosmotropic, which means that they tend to be excluded from the membrane surface, which is also described in the preferential exclusion theory.40

4. MEMBRANE

PHASE AND PERMEABILITY BEHAVIOR OF CELLS UNDER EXTREME

CONDITIONS

(figure 4)

4.1 Membrane phase behavior during cooling, freezing and drying of cells: membrane structure and lipid composition, thermotropic and lyotropic phase transitions, freezinginduced fluid-to-gel phase transition, effects of salts and protective agents

Membrane lipid bilayers can exist in a variety of conformations, including lamellar liquid crystalline (Lα) and lamellar gel (Lβ) phases.41,42 Transitions between these phases can be induced by temperature or dehydration (i.e. thermotropic and lyotropic phase transitions, respectively), or by adding solutes that interact with lipids. Under normal physiological conditions, cellular membranes are in a fluid phase, exhibiting large conformational disorder. In this phase more ordered domains designated as ‘rafts’ are embedded, which are enriched in


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cholesterol and sphingolipids.43 Membrane fluidity is primarily determined by its composition and the structure of the acyl chains. Shorter chain lengths and double bonds in unsaturated lipids increase the fluidity of the bilayer.44 Cooling alters membrane organization and structure, which in some cases is irreversible and impairs cell function.45−47 Cholesterol plays an important role in modulating membrane phase behavior during cooling.48 Membrane fluidity of cells can be studied using Fourier transform infrared (FTIR) spectroscopy by inspecting the position of the symmetric CH2-stretching vibration absorbance band (CH2) arising from membrane lipids. A shift of CH2 to lower wavenumbers denotes a decrease in the rotational freedom of the lipid acyl chains.41 Membrane phase transitions can be inferred from discontinuities in CH2 versus temperature plots. In cells, at suprazero temperatures, typically a broad non-cooperative phase transition is observed, whereas freezing and ice formation induces a sharp fluid-to-gel-phase transition (Figure 4A). This is the result of both thermotropic as well as lyotropic factors; since cooling causes a general decrease in rotational freedom of the acyl chains, whereas ice formation additionally causes dehydration and removal of water normally surrounding the phospholipid head groups.49,50 The extent and rate of the freezing-induced membrane phase transition is dependent on the ice nucleation temperature, the cooling rate, as well as the presence of CPAs.49−53 Higher ice nucleation temperatures (Tn) causing more severe cellular dehydration result in strong and cooperative phase transitions, whereas phase transitions are absent in case of supercooling or at low Tn (e.g. −10°C). In the FTIR based method that was used in these studies, the cooling rate was kept constant at 1°C/ min and the nucleation temperature was varied by touching the sample with a pre-cooled copper wire. Interestingly, CPAs do not prevent the occurrence of a freezing-induced fluid-to-gel membrane phase transition, but they merely decrease the cooperativity of the phase transition and rate of dehydration (Figure 4B).


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This is the case both for permeating CPAs as well as disaccharides.53,54 Disaccharides have been described to replace water bound to the phospholipid head group in case of air drying thereby preventing lyotropic phase transitions.55,56 Apparently this does not occur during freezing-induced dehydration. Whereas air drying itself results in a decrease in acyl chain rotational freedom (at 20°C), dried cells do not exhibit cooperative phase transitions during cooling. Moreover, at low subzero temperatures (at −30°C), CH2 is similar as that of hydrated cells subjected to supercooling without freezing-induced dehydration. Only if cells are dried in the presence of 3 M NaCl, prior to cooling, a similar CH2 value is found (Figure 3B). This indicates that, at high Tn (of −3°C), the osmotic driving force generated by extracellular ice formation causes membrane lipids to become more tightly packed compared to air-dried samples.57−59 Cells frozen with high Tn have more time for water transport than cells frozen with a lower Tn. This leads to the sharp phase transition observed at high Tn, whereas at lower Tn cells do not have enough time to fully dehydrate.

4.2 Ice formation and protective agents affect membrane water transport: Arrhenius behavior, supra-/subzero temperatures, protectants, modeling freezing induced dehydration for determining optimal cooling rates

Subzero membrane permeability to water can be determined from cell volume images acquired using a cryomicroscope.60 In addition, FTIR studies on freezing-induced membrane phase behavior can be used to derive Lp at subzero temperatures by assuming that the freezing-induced shift in CH2 is proportional to the cell volume response during freezing.49,50,51,53,59 Models and equations are available for fitting plots on the cellular volume


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decrease during freezing, at a particular Tn and cooling rate, and describing the temperature dependency (i.e. Arrhenius behavior) of Lp at subzero temperatures.60,61,62 The derived parameters describing membrane water permeability at reference temperature 0°C (Lpg) and the activation energy for water transport (ELp) in turn can be used to model cell volume responses and predict optimal cooling rates for cryopreservation.63,64 Arrhenius behavior of Lp is different at supra and subzero temperatures, because ice formation increases ELp (Figure 4C). This is explained by an increased lipid packing at lower temperatures and presence of a gel phase, causing water molecules to encounter a more hydrophobic environment when crossing the membrane.49,50 CPAs appear to counteract the effects of ice on membrane permeability to water, and decrease ELp while increasing Lp at a given temperature (Figure 4C). Thus, experimental data from our group indicates that CPAs facilitate cellular dehydration to continue down to low subzero temperatures, which in turn decreases the incidence of intracellular ice formation.53

4.3 Freezing alters the membrane permeability function of cells for membrane impermeable solutes: loading cells with membrane impermeable protectants simply by freezing, cryopreservation, freeze-drying

In addition to altering the membrane permeability to water, freezing causes membranes to become permeable for molecules for which they are normally impermeable.54,65,66,67 This is explained by the fact that membrane imperfections are being formed during freezing-induced membrane phase transitions, which facilitate leakage or uptake of solutes along concentration gradients (Figure 4D). Actually, this allows incorporation of the disaccharide trehalose into cells with high loading efficiencies.54,68


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Loading efficiencies of 50 % have been observed for fibroblasts; i.e. freezing cells with 200 mM trehalose, results in 100 mM intracellular trehalose after freezing and thawing. Also charged molecules such as lucifer yellow, which is similar in size as trehalose and normally membrane impermeable, is taken up after freezing and thawing. It needs to be investigated if larger molecules are also taken up. Freezing-induced trehalose uptake facilitates cryosurvival without the need to make use of permeating CPAs such as dimethyl sulfoxide which have cytotoxic features.68 Disaccharides have both cryo- and lyoprotective properties and therefore can be used as protective agents during freeze-drying. Freezing-mediated loading of cells with disaccharides has been shown to be beneficial for freeze-drying because they stabilize intracellular biomolecular structures (e.g. DNA/nuclei) during dried storage.67,69

5. SUMMARY: DEVELOPMENT OF CELL PRESERVATION TECHNOLOGIES REQUIRES INSIGHTS IN MEMBRANE PHASE AND PERMEABILITY BEHAVIOR

Membrane structure and permeability characteristics determine how a cell copes with osmotic imbalances and solute interactions that are encountered during cryopreservation. Experimentally determined permeability parameters can be used to predict and model cell volume responses at supra- as well as subzero temperatures, which in turn can be used to design CPA loading/removal protocols, and to predict optimal cooling rates for cryopreservation. As is discussed here, however, it should be noted that membrane permeability to water and solutes/CPAs are not fixed parameters. They are dependent on temperature, medium osmolality, types of CPAs and other solutes present, cell hydration level, as well as the absence or presence of ice. Furthermore, differences amongst CPAs exist in how they interact with membranes and eventually confer permeabilization and


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cytotoxicity. Freezing most drastically alters the membrane permeability barrier function, which is reflected as an increase in the activation energy for water transport (counteracted by cryoprotective agents). In addition, membranes become temporarily leaky during freezinginduced fluid-to-gel membrane phase transitions resulting in the uptake of impermeable solutes. In this case, one can take advantage of the distorted membrane permeability barrier function and incorporate trehalose into cells stabilizing them during cryopreservation or freeze-drying.


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ACKNOWLEDGMENTS

This work was financially supported by the German Research Foundation (DFG: Deutsche Forschungsgemeinschaft) via the Cluster of Excellence ‘From regenerative biology to reconstructive therapy’ (REBIRTH) and grants WO1735/6-1 and SI1462/4-1. Bulat Sydykov and Arielle Keller Brunatto are acknowledged for their help with fitting data and modeling using MATLAB. All previous and current members of the laboratories of WW and HS are acknowledged for their contributions.

REFERENCES

(1) Alshalani, A.; Acker, J.P. Red blood cell membrane water permeability increases with length of ex vivo storage. Cryobiology 2017, 76, 51–58. (2) Newton, H.; Pegg, D.E.; Barrass, R.; Gosden, R.G. Osmotically inactive volume, hydraulic conductivity, and permeability to dimethyl sulphoxide of human mature oocytes. J. Reprod. Fertil. 1999, 117, 27–33. (3) Mullen, S.F.; Rosenbaum M.; Critser, J.K. The effect of osmotic stress on the cell volume, metaphase II spindle and developmental potential of in vitro matured porcine oocytes. Cryobiology 2007, 54, 281–289. (4) Liu, J.; Mullen, S.; Meng, Q.; Critser, J. Dinnyes, A. Determination of oocyte membrane permeability coefficients and their application to cryopreservation in a rabbit model. Cryobiology 2009, 59, 127–134. (5) Sieme, H.; Harrison, R.A.; Petrunkina, A.M. Cryobiological determinants of frozen semen quality, with special reference to stallion. Anim. Reprod. Sci. 2008, 107, 276–292.


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(6) Si, W.; Benson, J.D.; Men, H.; Critser, J.K. Osmotic tolerance limits and effects of cryoprotectants on the motility, plasma membrane integrity and acrosomal integrity of rat sperm. Cryobiology 2006, 53, 336–348. (7) Gilmore, J.A.; McGann, L.E.; Liu, J.; Gao, D.Y.; Peter, A.T.; Kleinhans, F.W.; Crister, J.K. Effect of cryoprotectant solutes on water permeability of human spermatozoa. Biol. Reprod. 1995, 53, 985–995. (8) Oldenhof, H.; Blässe, A.K.; Wolkers, W.F.; Bollwein, H.; Sieme, H. Osmotic properties of stallion sperm subpopulations determined by simultaneous assessment of cell volume and viability. Theriogenology 2011, 76, 386–391. (9) Gao, D.Y.; McGrath, J.J.; Tao, J.; Benson, C.T.; Critser, E.S.; Critser, J.K. Membrane transport properties of mammalian oocytes: a micropipette perfusion technique. J. Reprod. Fertil. 1994, 102, 385–392. (10) Zhao, G.; Zhang, Z.; Zhang, Y.; Chen, Z.; Niu, D.; Cao, Y.; He, X. A microfluidic perfusion approach for on-chip characterization of the transport properties of human oocytes. Lab Chip 2017, 17, 1297–1305. (11) McGann, L.E.; Turc, J.M. Determination of water and solute permeability coefficients. Cryobiology 1980, 17, 612–613. (12) Agca, Y.; Mullen, S.; Liu, J.; Ward, J.J.; Gould, K.; Chan, A.; Critser J. Osmotic tolerance and membrane permeability characteristics of rhesus monkey (Macaca mulatta) spermatozoa. Cryobiology 2005, 51, 1–14. (13) Kedem, O.; Katchalsky, A. Thermodynamic analysis of the permeability of biological membranes to non-electrolytes. Biochim. Biophys. Acta 1985, 27, 229–246. (14) Kleinhans, F.W. Membrane permeability modeling: Kedem-Katchalsky vs a twoparameter formalism. Cryobiology 1998, 37, 271–289. (15) Lahmann, J.M.; Benson, J.D.; Higgins, A.Z. Concentration dependence of the cell


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membrane permeability to cryoprotectant and water and implications for design of methods for post-thaw washing of human erythrocytes. Cryobiology 2018, 80, 1–11. (16) Toupin, C.J.; Le Maguer, M.; McGann, L.E. Permeability of human granulocytes to water: rectification of osmotic flow. Cryobiology 1989, 26, 431–44. (17) Peckys, D.B.; Kleinhans, F.W.; Mazur, P. Rectification of the water permeability in COS-7 cells at 22, 10 and 0°C. PLoS One 2011, 6, e23643. (18) Muldrew, K.; Schachar, J.; Cheng, P.; Rempel, C.; Liang, S.; Wan, R. The possible influence of osmotic poration on cell membrane water permeability. Cryobiology 2009, 58, 62–68. (19) Böckmann, R.A.; Hac, A.; Heimburg, T.; Grubmüller, H. Effect of sodium chloride on a lipid bilayer. Biophys. J. 2003, 85, 1647–1655. (20) Mazur, P.; Leibo, S.P.; Miller, R.H. Permeability of the bovine red cell to glycerol in hyperosmotic solutions at various temperatures. J. Membr. Biol. 1974, 15, 107–136. (21) Jacobs, M.H. The simultaneous measurement of cell permeability to water and to dissolved substances. J. Cell Comp. Physiol. 1932, 2, 427–444. (22) Borgnia, M.; Nielsen, S.; Engel, A.; Agre, P. Cellular and molecular biology of the aquaporin water channels. Annu. Rev. Biochem. 1999, 68, 425–458. (23) Leibo, S.P. Water permeability and its activation energy of fertilized and unfertilized mouse ova. J. Membr. Biol. 1980, 53, 179–188. (24) Agca, Y.; Liu, J.; McGrath, J.J.; Peter, A.T.; Critser, E.S.; Critser, J.K. Membrane permeability characteristics of metaphase II mouse oocytes at various temperatures in the presence of Me2SO. Cryobiology 1998, 36, 287–300. (25) Agca, Y.; Liu, J.; Critser, E.S.; Critser, J.K. Fundamental cryobiology of rat immature and mature oocytes: hydraulic conductivity in the presence of Me2SO, Me2SO permeability, and their activation energies. J. Exp. Zool. 2000, 286, 523–533.


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(26) Benson J.D. Modeling and Optimization of Cryopreservation. In: Wolkers W.F.; Oldenhof, H. (eds.) Cryopreservation and Freeze-Drying Protocols. Methods in Molecular Biology (Methods and Protocols) 2015, 1257, 83–120, Springer, New York, NY. (27) Elmoazzen, H.Y.; Elliott, J.A.W.; McGann, L.E. The effect of temperature on membrane hydraulic conductivity. Cryobiology 2002, 45, 68–79. (28) Verkman, A.S.; van Hoek, A.N.; Ma, T.; Frigeri, A.; Skach, W.R.; Mitra, A.; Tamarappoo, B.K.; Farinas J. Water transport across mammalian cell membranes. Am. J. Physiol. 1996, 270, C12-C30. (29) Fahy G.M.; Wowk B. Principles of Cryopreservation by Vitrification. In: Wolkers W.F.; Oldenhof, H. (eds.) Cryopreservation and Freeze-Drying Protocols. Methods in Molecular Biology (Methods and Protocols) 2015, 1257, 163–167, Springer, New York, NY. (30) Sydykov, B.; Oldenhof, H.; de Oliveira Barros, L.; Sieme, H.; Wolkers, W.F. Membrane permeabilization of phosphatidylcholine liposomes induced by cryopreservation and vitrification solutions. Biochim. Biophys. Acta, 2018, 1860, 467–474. (31) Sydykov, B.; Oldenhof, H.; Sieme, H.; Wolkers, W.F. Storage stability of liposomes stored at elevated subzero temperatures in DMSO/sucrose mixtures. PLoS One 2018, 13, e0199867. (32) de Menorval, M.A.; Mir, L.M.; Fernandez, M.L.; Reigada, R. Effects of dimethyl sulfoxide in cholesterol-containing lipid membranes: a comparative study of experiments in silico and with cells. PLoS One 2012, 7, e41733. (33) Anchordoguy, T.J.; Carpenter, J.F.; Crowe, J.H.; Crowe, L.M. Temperaturedependent perturbation of phospholipid bilayers by dimethylsulfoxide. Biochim. Biophys. Acta 1992, 1104, 117–122. (34) Gordeliy, V.I., Kiselev, M.A.; Lesieur, P.; Pole, A.V.; Teixeira, J. Lipid membrane structure and interactions in dimethyl sulfoxide/water mixtures, Biophys. J. 1998, 75, 2343–


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2351. (35) Schrader, A.M.; Cheng, C.Y.; Israelachvili, J.N.; Han, S. Communication: contrasting effects of glycerol and DMSO on lipid membrane surface hydration dynamics and forces. J. Chem. Phys. 2016, 145, 041101. (36) Hung, W.C.; Lee, M.T.; Chen, F.Y.; Huang, H.W. The сondensing effect of сholesterol in lipid bilayers. Biophys. J. 2007, 92, 3960–3967. (37) de Meyer, F.; Smit, B. Effect of cholesterol on the structure of a phospholipid bilayer. Proc. Natl. Acad. Sci. U.S.A. 2009, 106, 3654–3658. (38) Pereira, C.S.; Hunenberger, P.H. The influence of polyhydroxylated compounds on a hydrated phospholipid bilayer: a molecular dynamics study. Mol. Simul. 2008, 34, 403–420. (39) Koynova, R.; Brankov, J.; Tenchov, B. Modulation of lipid phase behavior by cosmotropic and chaotropic solutes. xperiment and thermodynamic theory. Eur. Biophys. J. 1997, 25, 261–274. (40) Arakawa, T.; Timasheff, S.N. The stabilization of proteins by osmolytes. Biophys. J. 1985, 47, 411–414. (41) Mantsch, H.H.; McElhaney, R.N. Phospholipid phase transitions in model and biological membranes as studied by infrared spectroscopy. Chem. Phys. Lipids 1991, 57, 213–226. (42) Caffrey, M.; Hogan, J. LIPIDAT: A database of lipid phase transition temperatures and enthalpy changes. DMPC data subset analysis. Chem. Phys. Lipids 1992, 61, 1–109. (43) Brown, D.A.; London, E. Structure and function of sphingolipid- and cholesterolrich membrane rafts. J. Biol. Chem. 2000, 275, 17221–17224. (44) Israelachvili, J.N.; Marcelja, S.; Horn, R.G. Physical principles of membrane organization. Quaterly Rev. Biophys. 1980, 13, 121–200. (45) Steponkus, P.L. Role of the plasma membrane in freezing injury and cold


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acclimation. Ann. Rev. Plant Physiol. 1984, 35, 543–584. (46) Drobnis, E.Z.; Crowe, L.M.; Berger, T.; Anchordoguy, T.J.; Overstreet, J.W.; Crowe, J.H. Cold shock damage is due to lipid phase transitions in cell membranes: a demonstration using sperm as a model. J. Exp. Zool. 1993, 265, 432–437. (47) Gousset, K.; Wolkers, W.F.; Tsvetkova, N.M.; Oliver, A.E.; Field, C.L.; Walker, N.J.; Crowe, J.H.; Tablin, F. Evidence for a physiological role for membrane rafts in human platelets. J. Cell. Physiol. 2002, 119, 117–128. (48) Stoll, C.; Stadnick, H.; Kollas, O.; Holovati, J.L.; Glasmacher, B.; Acker, J.P.; Wolkers, W.F. Liposomes alter thermal phase behavior of red blood cell membranes. Biochim. Biophys. Acta 2011, 1808, 474–481. (49) Akhoondi, M.; Oldenhof, H.; Stoll, C.; Sieme, H.; Wolkers, W.F. Membrane hydraulic permeability changes during cooling of mammalian cells. Biochim. Biophys. Acta 2011, 1808, 642–648. (50) Oldenhof, H.; Friedel, K.; Sieme, H.; Glasmacher, B.; Wolkers, W.F. Membrane permeability parameters for freezing of stallion sperm as determined by Fourier transform infrared spectroscopy. Cryobiology 2010, 61, 115–122. (51) Wolkers, W.F.; Balasubramanian, S.K.; Ongstad, E.L.; Zec, H.; Bischof, J.C. Effects of freezing on membranes and proteins in LNCaP prostate tumor cells. Biochim. Biophys. Acta 2007, 1768, 728–736. (52) Oldenhof, H.; Friedel, K.; Akhoondi, M.; Gojowsky, M.; Wolkers, W.F.; Sieme, H. Membrane phase behavior during cooling of stallion sperm and its correlation with freezability. Mol. Membr. Biol. 2012, 29, 95–106. (53) Oldenhof, H.; Gojowsky, M.; Wang, S.; Henke, S.; Yu, C.; Rohn, K.; Wolkers, W.F.; Sieme, H. Osmotic stress and membrane phase changes during freezing of stallion sperm: mode of action of cryoprotective agents. Biol. Reprod. 2013, 88, 68:1–11.


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(54) Zhang, M.; Oldenhof, H.; Sieme, H.; Wolkers, W.F. Freezing-induced uptake of trehalose into mammalian cells facilitates cryopreservation. Biochim. Biophys. Acta 2016, 1858, 1400–1409. (55) Crowe, J.H.; Carpenter, J.F.; Crowe, L.M.; Anchordoguy, T.J. Are freezing and dehydration similar stress vectors? A comparison of modes of interaction of stabilizing solutes with biomolecules. Cryobiology 1990, 27, 219–231. (56) Crowe, J.H.; Hoekstra, F.A.; Crowe, L.M. Anhydrobiosis. Annu. Rev. Physiol. 1992, 54, 579–599. (57) Akhoondi, M.; Oldenhof, H.; Sieme, H.; Wolkers, W.F. Freezing-induced removal of water from phospholipid head groups in biomembranes. Biomed. Spectrosc. Imaging 2012, 1, 293–302. (58) Akhoondi, M.; Oldenhof, H.; Sieme, H.; Wolkers, W.F. Freezing-induced cellular and membrane dehydration in the presence of cryoprotective agents. Mol. Membr. Biol. 2012, 29, 197–206. (59) Oldenhof, H.; Akhoondi, M.; Sieme, H.; Wolkers, W.F. Use of Fourier transform infrared spectroscopy to determine optimal cooling rates for cryopreservation of cells. Biomed. Spectrosc. Imaging 2013, 2, 83–90. (60) Mazur, P. Kinetics of water loss from cells at subzero temperatures and the likelihood of intracellular freezing. J. Gen. Physiol. 1963, 47, 347–369. (61) Levin, R.L.; Cravalho, E.G.; Huggins, C.E. A membrane model describing the effect of temperature on the water conductivity of erythrocyte membranes at subzero temperatures. Cryobiology 1967, 13, 415–429. (62) Toner, M.; Cravalho, E.G.; Armant, D.R. Water transport and estimated transmembrane potential during freezing of mouse oocytes. J. Membr. Biol. 1990, 115, 261– 272.


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(63) Devireddy, R.V.; Swanlund, D.J.; Olin, T.; Vincente, W.; Troedsson, M.H.T.; Bischof, J.C.; Roberts, K.P. Cryopreservation of equine sperm: optimal cooling rates in the presence and absence of cryoprotective agents determined using differential scanning calorimetry. Biol. Reprod. 2002, 66, 222–231. (64) Mazur, P.; Rall, W.F., Leibo, S.P. Kinetics of water loss and the likelihood of intracellular freezing in mouse ova. Influence of the method of calculating the temperature dependence of water permeability. Cell. Biophys. 1984, 6, 197–213. (65) Stoll, C.; Holovati, J.L.; Acker, J.P.; Wolkers, W.F. Synergistic effects of liposomes, trehalose and hydroxyethyl starch for cryopreservation of human erythrocytes. Biotechnol. Prog. 2012, 28, 364–371. (66) Gläfke, C.; Akhoondi, M.; Oldenhof, H.; Sieme, H.; Wolkers, W.F. Cryopreservation of platelets using trehalose: the role of membrane phase behavior during freezing. Biotechnol. Prog. 2012, 28, 1347–1354. (67) Oldenhof, H.; Zhang, M.; Narten, K.; Bigalk, J.; Sydykov, B.; Wolkers, W.F.; Sieme H. Freezing-induced uptake of disaccharides for preservation of chromatin in freeze-dried sperm during accelerated aging. Biol. Reprod. 2017, 97, 892–901. (68) Zhang, M.; Oldenhof, H.; Sieme, H.; Wolkers W.F. Combining endocytic and freezing-induced trehalose uptake for cryopreservation of mammalian cells. Biotechnol. Prog. 2017, 33, 229–235. (69) Zhang, M.; Oldenhof, H.; Sydykov, B.; Bigalk, J.; Sieme, H.; Wolkers, W.F. Freezedrying of mammalian cells using trehalose: preservation of DNA integrity. Sci. Rep. 2017, 7, 6198. (70) Yoshioka, K.; Suzuki, C.; Tanaka, A.; Anas, I.M.-K.; Iwamura S. Birth of piglets derived from porcine zygotes cultured in a chemically defined medium. Biol. Reprod. 2002, 66, 112–119.


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(71) Yoshioka, K.; Suzuki, C.; Onishi, A. Defined system for in vitro production of porcine embryos using a single basic medium. J. Reprod. Dev. 2008, 54, 208–213. (72) Somfai, T.; Men, N.T.; Noguchi, J.; Kaneko, H.; Kashiwazaki, N.; Kikuchi, K. Optimization of cryoprotectant treatment for the vitrification of immature cumulus-enclosed porcine oocytes: comparison of sugars, combinations of permeating cryoprotectants and equilibration regimens. J. Reprod. Dev. 2015, 61, 571–579.


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FIGURE LEGENDS

Fig. 1. Cell volume response and membrane hydraulic permeability of porcine oocytes exposed to anisotonic solutions. Hypo- and hypertonic phosphate buffered saline (PBS) was prepared by diluting PBS with water or by adding sucrose, respectively. Oocytes were collected from ovaries obtained at a local slaughter, and handled as described elsewhere.70,71 For volume measurements, the micropipette perfusion technique was used.9 Oocytes were held in a 10 L droplet using a micromanipulator setup, and subsequently flushed with 2 mL anisotonic solution using a syringe while acquiring a video. Frames at distinct time intervals were extracted (A), volumes were analyzed using ImageJ software by assuming a spherical shape (yellow circles), and relative volume changes were calculated and plotted as a function of the exposure duration (B). Cell volumes determined after 10 min exposure were plotted as a function of the medium osmolality (C). Cells exhibit swelling and shrinking under hypoand hypertonic conditions, respectively. The cell volume in medium with an osmolality of 300 mOsm was taken as the isotonic volume (Vo), and the osmotically inactive volume (Vb) was found by extrapolating the normalized cell volume data in the Boyle-van ’t Hoff plot (Cinset) to infinite osmolality. Volume changes were analyzed for both immature germinal vesicle stage (GV; open symbols and bars) as well as in vitro matured (IVM; closed symbols and bars) oocytes. The membrane permeability for water (Lp) was determined by fitting cell volume response data (D) with equation 1. Mean values ± standard deviations were determined from 6−9 oocytes, originating from 3−4 different batches of ovaries. For both the GV and the IVM group, separate one-way ANOVA tests were conducted to determine statistical significant differences. Different letter represent significant differences in the GV group (p≤0.05), whereas different capital letters represent significant differences in the IVM group (p≤0.05).


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Fig. 2. Cell volume response of porcine oocytes exposed to solutions supplemented with different permeating cryoprotective agents to determine water and solute permeability. Oocytes were exposed to 1.5 M ethylene glycol/EG (circles, white bars), dimethyl sulfoxide/DMSO (triangles, bars with upward diagonal lines), and propylene glycol/PG (squares, bars with downward diagonal lines). Cell volumes initially decrease, due to water leaving the cell, after which they return to their original volume coinciding with the uptake of permeating cryoprotective agents/CPAs. The membrane hydraulic permeability (Lp) and permeability for permeating CPAs (Ps) were determined by fitting cell volume versus time (plots

(A−C)) with equations 1 and 2. Permeability towards water (D) and solutes (E) was

determined both for immature (GV) as well as in vitro matured (IVM) oocytes. Mean values ± standard deviations were determined from 7−9 oocytes, originating from 3−4 different batches of ovaries.

Fig. 3. Membrane leakiness upon exposure to permeating agents studied using porcine oocytes as well as a liposome model system. Porcine oocytes can be vitrified using a twostep vitrification protocol, in which cells are first exposed to equilibration solution (ES: containing EG/PG, 2% each), followed by transfer in vitrification solutions (VS; containing EG/PG, 17.5% each) and plunging in liquid nitrogen.72 Oocyte membrane permeabilization and leakiness upon exposure to 4−35% EG/PG was studied by adding propidium iodide (PI; membrane impermeable fluorescent dye) and monitoring PI-staining as a function of the exposure duration (A). Membrane permeabilization upon exposure to VS is reduced after preincubation in ES (B). The capability of oocytes to mature in vitro, however, rapidly decreases with increasing exposure duration to VS after ES pre-treatment (B-inset). Values were calculated from 30−75 oocytes; originating from 3−6 different batches of ovaries. Liposomes


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prepared from phosphatidyl choline (PC) with(out) cholesterol (CHO) and entrapped carboxyfluorescein (CF), can be used as a model system to study CF retention/leakage and protectant-lipid interactions (C). Liposomes were incubated with varying concentrations of DMSO (circles), EG (triangles), or glycerol/GLY (squares) after which the extent of leakage of CF was determined fluorometrically (D). In addition, using infrared spectroscopy, the phase shift in the absorbance band arising from the phospholipid headgroup (PO4) was inspected to study protectant/lipid interactions (E). Effects on acyl chain interactions and lipid phase behavior were studied by following the wavenumber position of the CH2 stretching vibration band (CH2) as a function of the temperature (F). Plots and data presented in panel D−F were adapted from a previous study by our group.30

Fig. 4. Membrane phase behavior and water permeability of mammalian cells at supra and subzero temperatures; effects of cryoprotective agents and dehydration. Membrane phase behavior of fibrobrasts was studied using infrared spectroscopy by monitoring the temperature dependency of CH2. Cellular membranes typically exhibit a broad noncooperative phase transition at supra-zero temperatures. In addition, at sub-zero temperatures ice nucleation induces cellular dehydration coinciding with a sharp fluid-to-gel membrane phase transition (A). Effects of freezing, drying and osmotic stress on membrane phase behavior were investigated (B). It can be seen that the freezing-induced phase transition is not prevented by addition of CPAs (closed and open circles). Furthermore, it is especially the osmotic stress and not drying per se that results in such a phase shift (downward and upward triangles). In the Arrhenius plot, it can be seen that the membrane hydraulic permeability is affected by the temperature as well as the presence of ice and CPAs (C). Presence of DMSO decreases the activation energy for water transport (ELp) and increases the rate of water transport allowing cellular dehydration to continue at low subzero temperatures. Plots and


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data presented in panel A−C were adapted from previous studies by our group.49,57,58,53,54 Panel D presents a schematic presentation of the membrane phase state at physiological temperatures as well as after ice nucleation and freezing-induced dehydration. Fluid and gel phase lipid domains are indicated, as well as ice formation and fluxes (arrows) of water (circles) and impermeable solutes (squares).


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TABLE OF CONTENTS GRAPHIC


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Fig. 1. Cell volume response and membrane hydraulic permeability of porcine oocytes exposed to anisotonic solutions. Hypo- and hypertonic phosphate buffered saline (PBS) was prepared by diluting PBS with water or by adding sucrose, respectively. Oocytes were collected from ovaries obtained at a local slaughter, and handled as described elsewhere.70,71 For volume measurements, the micropipette perfusion technique was used.9 Oocytes were held in a 10 L droplet using a micromanipulator setup, and subsequently flushed with 2 mL anisotonic solution using a syringe while acquiring a video. Frames at distinct time intervals were extracted (A), volumes were analyzed using ImageJ software by assuming a spherical shape (yellow circles), and relative volume changes were calculated and plotted as a function of the exposure duration (B). Cell volumes determined after 10 min exposure were plotted as a function of the medium osmolality (C). Cells exhibit swelling and shrinking under hypo- and hypertonic conditions, respectively. The cell volume in medium with an osmolality of 300 mOsm was taken as the isotonic volume (Vo), and the osmotically inactive volume (Vb) was found by extrapolating the normalized cell volume data in the Boyle-van ’t Hoff plot (C-inset) to infinite osmolality. Volume changes were analyzed for both immature germinal vesicle stage (GV; open symbols and bars) as well as in vitro matured (IVM; closed symbols and bars) oocytes. The membrane permeability for water (Lp) was determined by fitting cell volume response data (D) with equation 1. Mean values ± standard deviations were determined from 6−9 oocytes, originating from 3−4 different batches of ovaries. For both the GV and the IVM group, separate one-way ANOVA tests were conducted to determine statistical significant differences. Different letter represent significant differences in the GV group (p≤0.05), whereas different capital letters represent significant differences in the IVM group (p≤0.05). 954x897mm (96 x 96 DPI)


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Fig. 2. Cell volume response of porcine oocytes exposed to solutions supplemented with different permeating cryoprotective agents to determine water and solute permeability. Oocytes were exposed to 1.5 M ethylene glycol/EG (circles, white bars), dimethyl sulfoxide/DMSO (triangles, bars with upward diagonal lines), and propylene glycol/PG (squares, bars with downward diagonal lines). Cell volumes initially decrease, due to water leaving the cell, after which they return to their original volume coinciding with the uptake of permeating cryoprotective agents/CPAs. The membrane hydraulic permeability (Lp) and permeability for permeating CPAs (Ps) were determined by fitting cell volume versus time (plots (A−C)) with equations 1 and 2. Permeability towards water (D) and solutes (E) was determined both for immature (GV) as well as in vitro matured (IVM) oocytes. Mean values ± standard deviations were determined from 7−9 oocytes, originating from 3−4 different batches of ovaries. 1077x577mm (120 x 120 DPI)


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Page 33 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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Fig. 3. Membrane leakiness upon exposure to permeating agents studied using porcine oocytes as well as a liposome model system. Porcine oocytes can be vitrified using a two-step vitrification protocol, in which cells are first exposed to equilibration solution (ES: containing EG/PG, 2% each), followed by transfer in vitrification solutions (VS; containing EG/PG, 17.5% each) and plunging in liquid nitrogen.72 Oocyte membrane permeabilization and leakiness upon exposure to 4−35% EG/PG was studied by adding propidium iodide (PI; membrane impermeable fluorescent dye) and monitoring PI-staining as a function of the exposure duration (A). Membrane permeabilization upon exposure to VS is reduced after pre-incubation in ES (B). The capability of oocytes to mature in vitro, however, rapidly decreases with increasing exposure duration to VS after ES pre-treatment (B-inset). Values were calculated from 30−75 oocytes; originating from 3−6 different batches of ovaries. Liposomes prepared from phosphatidyl choline (PC) with(out) cholesterol (CHO) and entrapped carboxyfluorescein (CF), can be used as a model system to study CF retention/leakage and protectant-lipid interactions (C). Liposomes were incubated with varying concentrations of DMSO (circles), EG (triangles), or glycerol/GLY (squares) after which the extent of leakage of CF was determined fluorometrically (D). In addition, using infrared spectroscopy, the phase shift in the absorbance band arising from the phospholipid headgroup (PO4) was inspected to study protectant/lipid interactions (E). Effects on acyl chain interactions and lipid phase behavior were studied by following the wavenumber position of the CH2 stretching vibration band (CH2) as a function of the temperature (F). Plots and data presented in panel D−F were adapted from a previous study by our group.30 1074x712mm (120 x 120 DPI)


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Fig. 4. Membrane phase behavior and water permeability of mammalian cells at supra and subzero temperatures; effects of cryoprotective agents and dehydration. Membrane phase behavior of fibrobrasts was studied using infrared spectroscopy by monitoring the temperature dependency of CH2. Cellular membranes typically exhibit a broad non-cooperative phase transition at supra-zero temperatures. In addition, at sub-zero temperatures ice nucleation induces cellular dehydration coinciding with a sharp fluidto-gel membrane phase transition (A). Effects of freezing, drying and osmotic stress on membrane phase behavior were investigated (B). It can be seen that the freezing-induced phase transition is not prevented by addition of CPAs (closed and open circles). Furthermore, it is especially the osmotic stress and not drying per se that results in such a phase shift (downward and upward triangles). In the Arrhenius plot, it can be seen that the membrane hydraulic permeability is affected by the temperature as well as the presence of ice and CPAs (C). Presence of DMSO decreases the activation energy for water transport (ELp) and increases the rate of water transport allowing cellular dehydration to continue at low subzero temperatures. Plots and data presented in panel A−C were adapted from previous studies by our group.49,57,58,53,54 Panel D presents a schematic presentation of the membrane phase state at physiological temperatures as well as after ice nucleation and freezing-induced dehydration. Fluid and gel phase lipid domains are indicated, as well as ice formation and fluxes (arrows) of water (circles) and impermeable solutes (squares). 1050x711mm (120 x 120 DPI)


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Factors Affecting the Membrane Permeability Barrier Function of Cells

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