Effect of Monoglyceride Structure and Cholesterol Content on Water

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Effect of Monoglyceride Structure and Cholesterol Content on Water Permeability of the Droplet Bilayer Zuzanna Michalak, Michelle Muzzio, Peter J. Milianta, Rosario Giacomini, and Sunghee Lee* Department of Chemistry, Iona College, 715 North Avenue, New Rochelle, New York 10801, United States ABSTRACT: The process of water permeation across lipid membranes has significant implications for cellular physiology and homeostasis, and its study may lead to a greater understanding of the relationship between the structure of lipid bilayer and the role that lipid structure plays in water permeation. In this study, we formed a droplet interface bilayer (DIB) by contacting two aqueous droplets together in an immiscible solvent (squalane) containing bilayer-forming surfactant (monoglycerides). Using the DIB model, we present our results on osmotic water permeabilities and activation energy for water permeation of an associated series of unsaturated monoglycerides as the principal component of droplet bilayers, each having the same chain length but differing in the position and number of double bonds, in the absence and presence of a varying concentration of cholesterol. Our findings suggest that the tailgroup structure in a series of monoglyceride bilayers is seen to affect the permeability and activation energy for the water permeation process. Moreover, we have also established the insertion of cholesterol into the droplet bilayer, and have detected its presence via its effect on water permeability. The effect of cholesterol differs depending on the type of monoglyceride. We demonstrate that the DIB can be employed as a convenient model membrane to rapidly explore subtle structural effects on bilayer water permeability.

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INTRODUCTION Many natural and synthetic systems rely on the transport of small molecules across bilayer membranes for their functional properties. Of intense interest in this realm is the permeability of water molecules across cellular membranes, important to maintain organism homeostasis and to eliminate waste. For this purpose, water flux across cellular barrier membranes is usually tightly controlled.1 A wealth of prior studies have investigated the effect of the properties of individual lipids and bilayers upon water permeability, such as thickness, fluidity, area per molecule, and lysis tension.2−5 These studies have suggested that tighter packing of lipids leads to a reduction of water permeability. However, the flux of water across cellular membranes is largely a function of both the lipid composition of the membrane, as well as the presence or absence of membrane protein channels, or pores, which can greatly facilitate the transport of water. In the absence of such pores, certain lipid mixtures can greatly retard the flow of water molecules, whereas, in the presence of pores such as those of the aquaporin family, the rate of water permeability is enhanced by orders of magnitude.1 In order to understand more completely how the lipid composition of membranes can modulate the rate of water flow across the semipermeable cellular membrane, a wide variety of different model systems have been employed. Prominent systems have included liposomal systems6 as well as the numerous types of supported lipid membranes (e.g., black lipid membranes).7,8 These have often been employed for osmotic © 2013 American Chemical Society

permeability measurements, in which a concentration gradient is imposed across the two sides of a membrane, such as the inner compartment of a vesicle or liposome versus its outer bathing solution, in order to cause an osmotically induced flow. From known values of osmotic pressure on each side of the membrane, coupled with values for the area of membrane traversed, water flux may be used to determine permeability coefficients. Such values are significant insofar as they may shed light on the underlying lipid bilayer structure responsible for both permeability and overall function of the membrane. Cholesterol is present in mammalian cellular membranes as a high fraction of total lipid (∼20−50 mol %), and plays a crucial role, principally in its organizing effect upon other lipidic components of the membrane.9 For example, cholesterol has been shown to reduce the area per molecule of phospholipids in membranes. Furthermore, it has an influence upon the level of both fluidity and rigidity of the bilayer, thus affecting membrane permeability, including permeability to water.10 Typically, the presence of cholesterol will reduce water permeability by a factor of 2−4, and often much higher. Interestingly, it has long been observed that the effect of cholesterol is more pronounced where the phospholipids of the membrane are saturated rather than unsaturated.11 However, while there have been extensive computational12−14 and Received: October 21, 2013 Revised: December 2, 2013 Published: December 4, 2013 15919

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Table 1. All Monoglycerides Used in This Study Contain a Fatty Acid, 18 Carbon Atoms Long with a cis Double Bond at Position 9 for MO (C18:1, c9), at Position 11 for MV (C18:1, c11), and at Positions 9 and 12 for ML (C18:2, c9, c12) lipids

chemical formula

abbreviations

1-(9Z-octadecenoyl)-rac-glycerol 1-(11Z-octadecenoyl)-rac-glycerol 1-(9Z,12Z-octadecadienoyl)-rac-glycerol

C21H40O4 C21H40O4 C21H38O4

MO (C18:1, c9) MV (C18:1, c11) ML (C18:2, c9, c12)

experimental studies15,16 regarding the interaction of cholesterol with phospholipids and other lipidic substances, both in natural and artificial membranes, the mechanism by which cholesterol imposes its ordering effect and concomitant effect upon permeability has not always been entirely elucidated. Great interest has been recently garnered for the use of the droplet interface bilayer (DIB) as a model membrane. These are lipid bilayers between pairs of aqueous droplets, formed by juxtaposing micrometer-sized aqueous droplets, submerged in an oil phase, each of which is stabilized by a lipid monolayer.17,18 Bilayer formation occurs at the point of contact, forming adherent droplets. DIB membranes have been employed for a wide variety of uses, such as the study of bioelectric phenomena (“biobatteries”),19 for microfluidic based devices,20 for reconstituting and interrogating membrane channel proteins by electrophysiological and other means,21 and for membrane transport studies.22−24 DIB systems have also come into prominence for their engineering applications, including for the printing of tissue-like materials25 and for systems to drive extremely rapid rates of crystallization.26 Despite the great success of DIB systems for the study of membrane-incorporated proteins, questions remain as to whether the insertion of such proteins into the droplet lipid bilayer gives rise to a model system which is truly reflective of the native environment of the protein. The native state, and thus true activity, of a membrane protein is intimately dependent on its surroundings. Therefore, it would be important to understand the structure of lipid organization in DIB systems per se, in addition to how they may act as a model for natural cellular membranes. Since natural membranes are complex mixtures, rather than simple lipids, it is important to investigate the properties of lipids in combination with important species (such as cholesterol), in order to further develop the DIB for biomimetic applications. Furthermore, improvement in the design of DIB systems for engineering and biomedical applications would also greatly benefit from a better understanding of the structure and arrangement of lipidic and other components of the droplet lipid bilayer. In this work, we contribute studies that systematically develop the droplet interface bilayer as a model membrane system for osmotically induced water permeability investigations. We employ an associated series of unsaturated monoglycerides as the principal component of droplet bilayers, each having the same chain length but differing in the position and number of double bonds. The tailgroup structure in a series of monoglyceride bilayers is seen to affect the activation energy for the water permeation process. Moreover, the addition of cholesterol to each type of monoglyceride has a differing influence.

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(MO), monovaccenin (MV), and monolinolein (ML), as shown in Table 1. These monoglycerides are naturally occurring neutral lipids that have been extensively studied from the point of view of their phase behavior in aqueous dispersion.27 For example, MO forms a rich variety of bilayer-containing phases in water, such as two inverted bicontinuous cubic phases formed in excess water. Such phases have been notable for their use in various applications such as crystallization of proteins.28 Monoglycerides generally have a simple molecular structure, which consists of a glycerol headgroup and an acyl chain, connected at the sn-1 position of the glycerol group. MO has an acyl chain consisting of 18 carbon atoms with one cis CC double bond between the 9th and 10th carbon atoms. MV also possesses a C18 acyl chain but is a positional isomer of MO where the cis CC double bond locates between the 11th and 12th carbon atoms. ML is more unsaturated, as it has two cis CC double bonds at C9 and C12. In a cis-unsaturated monoacylglyceride, the presence of the cis double bond in the chain creates a kink or a bend in the molecule. However, the magnitude of this curvature is dependent upon the position of the bond. The effect of varying the double bond position from C9 to C11 (i.e., monoolein to monovaccenin) is to make a “slimmer” molecule (since its cis double bond has been shifted away from the glycerol headgroup), which is manifested in the shape factor of the lipid.29,30 The two cis double bonds of ML provide the largest degree of curvature of the respective tailgroups, hence the most chain splay of the species studied here.

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MATERIALS AND METHODS

Materials. The monovaccenin (MV) and monolinolein (ML) were purchased from Nu Chek Prep Inc. and used as received (purity ≥99%). Monoolein (MO, purity ≥99%), cholesterol (CHOL, purity ≥99%), squalane (99%), and all other chemicals, of the highest purity available, were purchased from Sigma-Aldrich and used without additional purification. Sample Preparation. For all our experiments, the lipids employed were three members of the monoglyceride series (MO, MV, and ML), as well as cholesterol. Squalane (perhydrosqualene) was used as the immiscible organic phase. Monoglyceride-containing organic phases (lipid solutions) were prepared by dissolving the monoglyceride directly into squalane, followed by bath sonication for ∼30 min at ambient temperature. Similarly, lipid solutions containing cholesterol were also prepared by direct dissolution, but these required a longer time of sonication (∼ >1 h) to result in a clear solution. For all of the experiments conducted herein, the total lipid concentration in the oil solution (i.e., monoglyceride plus any added cholesterol) was approximately 10−12 mM. Aqueous solutions using osmolytes (NaCl at nominally 0.1 M) were prepared from purified, deionized water (18.2 MΩ·cm) using a Millipore water purification system (Direct Q-3). The osmolality (in mOsm/kg) of all solutions used was measured by a VAPRO vapor pressure osmometer (model 5600). All solutions were prepared immediately prior to use. All experiments were carried out using a custom built temperature-controlled microchamber which was thermostatted via an external circulating water bath. This allows variation and control of temperature from 16 to 65 °C within ±1 °C accuracy.

SYSTEM STUDIED

In this work, we have investigated the osmotic water permeability of a series of monoglycerides of C18 chain length, with varying degree and type of double bond unsaturation. We have chosen the following series of monoglycerides: monoolein 15920

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System. In order to visually image and manipulate droplets, we employed an inverted microscope (Nikon Eclipse Ti−S with halogen lamp) combined with two hydraulic micropipet manipulators (Narishige), supported on a vibration isolated workstation (Newport). The detailed system has been described elsewhere.31 All experiments are video recorded using a camera (Andor Zyla sCMOS) directly attached to the microscope. The recorded videos and images were post analyzed to measure the dimension of droplets and contact area using custom built image analysis software. All images were collected with a pixel size of 0.16 μm using the entire field of 1920 × 1080 pixels. Thus, the uncertainty in the diameter measurement is 0.32 μm (2 pixels × 0.16 μm/pixel). Since we generally have maintained droplets in the diameter size range of ∼100 μm, the error associated with the diameter measurement is 0.32%, which propagates to ∼0.6% relative uncertainty in volume calculation. The typical diameter of the DIB region is ∼50 μm, which will contribute to ∼0.9% relative uncertainty in the calculation of the area of the bilayer. When the uncertainties in optical resolution are taken together, these correspond to an uncertainty in permeability of ∼1.3%. Formation of Droplet Bilayer. For formation and manipulation of an aqueous microdroplet, a micropipet with a desired tip, having a typical diameter in the range 10−30 μm, was formed using a micropipet puller (Narishige PB-7) and subsequently hydrophobized by exposure to vapors of hexamethyldisilazane [(CH3)3SiNHSi(CH3)3)]. To achieve a hydrophobic coating of the micropipet, for inhibiting wetting of the micropipet glass surface by an aqueous solution, about 2−3 drops of hexamethyldisilazane was added to the center of an enclosed container having freshly pulled micropipets and held for at least 30 min. In order to form adherent droplet pairs, one aqueous microdroplet with a desired initial size is produced by applying slight positive pressure through a syringe connected to the micropipet. The tip of the micropipet is surrounded by the lipid solution in squalane. The microdroplet is allowed to fall through the lipid solution to the bottom of the oil chamber, where it is supported by a borosilicate glass slide. Another microdroplet having different osmolarity with a desired initial size is produced adjacent to the first. The two microdroplets are then brought together into adherent contact by briefly prodding with the tip of the micropipet, to form a droplet interface bilayer (DIB). Then, the adherent droplets are allowed to rest unconstrained on the glass slide, as shown in Figure 1. Droplets on the borosilicate glass slide forming the bottom of the oil solution chamber did not wet the glass, likely due to the presence of the monoglycerides in the oil solution which effectively hydrophobized the surface. We also used a plastic coverslip (Rinzl plastic coverslips, Electron Microscopy Sciences, Hatfield, PA) to form an oil solution chamber and found no difference in droplet

shape and corresponding permeability value. All droplet pairs had substantially the same initial size relative to each other, in the range of 100 ± 5 μm diameter. All droplet dimensions reported here are for diameters. In order to directly ascertain the effect upon droplet volume measurement arising from any adhesion of a water droplet to a surface, we performed the following test. First, we measured the diameter of a water droplet which was isolated and held by a micropipet tip with gentle aspiration. At this point, the held water droplet is in the middle of the oil solution and does not touch the bottom of the surface. No deformation to the spherical shape of a water droplet was observed. That is, the droplet was held in such a manner that no neck or cap of any sort was sucked into the micropipet from the spherical droplet. We measured the diameter of this water droplet. Second, we let this water droplet fall to the bottom of the chamber by gently tapping the micropipet. Once the water droplet was at rest at the bottom of the surface, this droplet came in contact with the surface. We measured the diameter of the same water droplet now at rest on the surface. This was tested for two kinds of substrates: borosilicate glass slide and plastic. We found no difference in droplet diameter within the limitations of the optical resolution (0.32 μm), regardless of whether it is held in the middle of the oil without touching any surface or whether it is resting on the surface, for both borosilicate glass and plastic substrate. Water Permeability Measurements. When two osmotically imbalanced droplets were made to adhere at a bilayer, water transport immediately commenced. Changes in droplet size due to this water transport were thus measured from the commencement of the process. Figure 2 shows examples of such a system, depicting the relative

Figure 2. Water transport rates (relative volume change for a pair of droplets over time, V0, initial volume) for swelling droplet (filled diamonds), shrinking droplet (open diamonds), and total volume (cross) over the time course of 5 min. The osmotic gradient between two droplets is set to 200 mOsm/kg. The selected corresponding image sequences are also shown. The arrow on the image indicates the direction of water transport. The scale bar on the image represents 100 μm. volume changes (V/V0) for swelling and shrinking droplet along with the corresponding images. In each case, there was DIB between adherent droplets: one is a pure water droplet, and the other droplet contains an osmolyte, NaCl. As shown in Figure 2, the leftmost droplet in the images contains NaCl (0.1 M) with an accurately known osmolality in the range of 200 mOsm/kg. The osmotic gradient drives water transport through the droplet bilayer. Any electrolyte flux is expected to be negligible compared to that of water. This resulted in a change in droplet diameter. The total volume of two droplets (cross marks in Figure 2) remains close to constant during the duration of the experiment (∼

Figure 1. Two droplets adhere to form a bilayer when a pair of aqueous droplets, each of which are stabilized by a lipid monolayer, are brought into contact in a squalane solution containing lipids (10−12 mM). Droplet pairs using lipid in a solvent of large molecular size (such as squalane) generally have a large contact area as shown, the size and area of which can be measured optically using custom-made software. The scale bar on the image represents 100 μm. 15921

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MO > ML: an increase of ∼150% for MV (9.5 to 24.5 kcal/mol), ∼100% increase for MO (9.4 to 18.8 kcal/mol), and ∼43% increase for ML (9.3 to 13.3 kcal/mol) in the presence of 4:1 mole ratio of cholesterol to monoglyceride. This signifies the importance of the degree of unsaturation and the position of the cis double bond in the monoglyceride chain on the interaction with the cholesterol in the bilayer. These studies lay the groundwork for the further development of DIB systems for the study of a wide variety of other processes involving small molecule permeation, such as nitric oxide and carbon dioxide. Moreover, this work opens up the possibility for investigation of other intercalants having a profound influence on water permeation, such as the water channel proteins, i.e., the aquaporins. Finally, the ease with which bilayers may be formed and from which permeability data may be amassed will help to situate DIB systems as useful experimental adjuncts to and complements of the plethora of ongoing molecular dynamics (MD) studies of permeability and other biophysical membrane properties.

Figure 7. Mean activation energy (kcal/mol) for a series of pure monoglycerides (white) and cholesterol mixtures. The mole ratio of cholesterol to lipid mixture was 1:1 (gray) and 4:1 (black).

and to 18.8 kcal/mol (4:1 ratio) for MO; from 9.3 to 12.2 kcal/ mol (for 1:1 ratio) and to 13.3 kcal/mol (4:1 ratio) for ML. A similar observation has been reported by Blok,37 showing that Ea is independent of fatty acyl composition of PC (∼10− 11 kcal/mol) in the absence of cholesterol, while Ea on the effect of cholesterol depended on the types of PC. It is interesting to note that the extent of the increasing Ea is in order MV > MO > ML: an increase of ∼150% for MV (9.5 to 24.5 kcal/mol), ∼100% increase for MO (9.4 to 18.8 kcal/ mol), and ∼43% increase for ML (9.3 to 13.3 kcal/mol) in the presence of 4:1 mole ratio of cholesterol content. The differential effect of cholesterol upon the activation energy for the various monoglycerides can be explained in terms of the position and effective length of the cholesterol in the bilayer. Cholesterol will orient itself in a bilayer with its polar hydroxy group proximate the aqueous phase and the hydrophobic steroid ring oriented parallel to and nested in the hydrocarbon chains of the lipid. In the particular case of monoglyceride bilayers, the cholesterol headgroup is also known to form hydrogen bonds to the glycerol portion.38 In the case of monoolein, with its double bond at the C9 position, the sterol ring of the cholesterol abuts the double bond, inhibiting the lateral interaction that cholesterol ordinarily has with saturated lipids. That is, the location of the kink or bend in MO cannot be avoided by CHOL. According to the work of Crilly and Earnshaw,36 cholesterol was seen as straddling the double bond of MO in planar lipid bilayers. In contrast, the double bond in MV is at C11, which is effectively far enough removed from the sterol ring that cholesterol “sees” only the long saturated segment of MV. There is precedent for this kind of differential interaction of CHOL with unsaturated lipids.37,39,40 Blok37 studied water permability in liposomes composed of either DOPC (unsaturated at C-9) or DEPC (1,2-dierucoyl-sn-glycero-3phosphocholine, which is cis-unsaturated at the C-13 position of the acyl chain). At a 50% CHOL concentration, Ea for water permeation in DOPC liposomes was 12.7 kcal/mol, whereas, for DEPC, the relevant value was ∼20 kcal/mol. The results were ascribed to the fact that, in DOPC, the cis double bond is at the C9 position of the fatty acid, that part of the paraffin chain that is involved in the interaction with the rigid sterol nucleus. However, in erucic acid (i.e., the acyl group in DEPC), the cis double bond is at the c13 position, i.e., in the part of the paraffin chain that does not interact with the sterol ring. As for ML, its intrinsic fluidity arising from its two kinks is apparently not quenched by intercalation of cholesterol. Wydro,41 for example, has observed that the ordering effect of cholesterol is

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AUTHOR INFORMATION

Corresponding Author

*Phone: 914-633-2638. Fax: 914-633-2240. E-mail: SLee@ iona.edu. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors would like to acknowledge the financial support from the National Science Foundation (NSF-CHE-1212967). M.M. and P.J.M. thank the Patrick J. Martin Foundation for a scholarship. We also thank Darius Fartash and Nousin Haque for their experimental contributions at the beginning of this project and Geoffrey B. Cawley for image analysis software.

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REFERENCES

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