Interactions of Hemin with Model Erythrocyte Membranes - The

Mar 17, 2010 - ... BasakA. ChakrabartiM. K. SanyalB. RungeB. M. Murphy ... Peter G. Vekilov , Jeffrey D. Rimer , Katy N. Olafson , Megan A. Ketchum...
1 downloads 0 Views 4MB Size
Download PDF
J. Phys. Chem. B 2010, 114, 4529–4535

4529

Interactions of Hemin with Model Erythrocyte Membranes Yasser Qutub,†,‡ Veselina Uzunova,†,‡ Oleg Galkin,‡ and Peter G. Vekilov*,‡,§ Department of Chemical and Biomolecular Engineering and Department of Chemistry, UniVersity of Houston, Houston, Texas 77204-4004 ReceiVed: January 21, 2010; ReVised Manuscript ReceiVed: February 11, 2010

To address the interactions of hemin with phospholipid bilayers, we introduce hemin to a solution of dimyristoylphosphatidylcholine (DMPC), a long chain phospholipid, and 3-(cholamidopropyl)(dimethylammonio)-2-hydroxy-1-propanesulfonate (CHAPSO), a detergent, in which vesicles form at around 37 °C. We show that vesicles composed of DMPC/CHAPSO form and grow, following a mechanism that does not trap solution and excludes larger solutes, such as hemin, from the vesicle interior. The existence of a limited number of patches of likely 2D crystalline hemin embedded in the phospholipid bilayer suggests that this layer is saturated with hemin molecules. We show that despite this saturation, even after prolonged contact with hemin-containing solution outside the vesicles, hemin is not released on the other side of the membrane; i.e., the phospholipid bilayer is impermeable to hemin. Comparison of the properties of the model membrane to those of the erythrocyte membrane suggests that the latter might also be impermeable to hemin and, given the absence of pores suitable to hemin in the erythrocyte membranes, that hemin might accumulate in erythrocytes after its release due to hemoglobin instability. Introduction Free hemin may be released excessively in erythrocytes from abnormal hemoglobins, and it may be released in the bloodstream from oxidized normal hemoglobin1-3 after erythrocyte breakage. Hemin is a substance with proven toxicity both in vitro and in vivo. In vitro hemin causes lysis of mouse3,4 and human erythrocytes,2 the malaria agents Plasmodium berghei5 and Plasmodium falciparum,6 neuron and neuron-like cell cultures.7,8 Hemin toxicity in vivo is a possible factor in pathological conditions with elevated levels inside erythrocytes or in plasma and tissues: hemorrhages and hemorrhagic infarctions of the central nervous system,7,8 glucose-6-phosphatedehydrogenase deficiency,9 and sickle cell disease.10 Hemin induces in vitro oxidation of low-density lipoprotein and is implicated in the development of atherosclerosis.11,12 Hemin associates to lipid bilayers,13,14 monolayers,15 and erythrocyte cytoskeletal and membrane proteins.16-18 Its interactions with different structures have been studied to explain the toxicity mechanisms. Hemin toxicity stems from two molecular properties: hydrophobicity and participation in oxidation reactions. Hydrophobicity enables interactions with cell membranes and association to hydrophobic surfaces. The presence of hemin in membranes disrupts their function, increasing disorder and permeability.17 Hemin partakes in oxidation/reduction reactions via its iron ion, resulting in production of reactive oxygen species and alteration of phospholipid and protein structure.19 There are no channels or pores in the red cell membrane that would allow active or passive transport of hemin out of the cell.20 Hemin transport across lipid bilayers during its efflux out of the mitochondriaswhere its de novo synthesis takes placesmay be facilitated by the presence of hemin binding proteins.13 Here we address a different mechanism of efflux of free hemin out of the cells: crossing the lipid part of the cell †

These authors contributed equally to this work. Department of Chemical and Biomolecular Engineering. § Department of Chemistry. ‡

membrane. For this, we study interactions of hemin with model lipid bilayers. We use unilamellar vesicles of dimyristoylphosphatidylcholine (DMPC), a long chain phospholipid, and 3-(cholamidopropyl)(dimethylammonio)-2-hydroxy-1-propanesulfonate (CHAPSO), a detergent, as model bilayers. The correspondences and differences between phospholipid bilayers of DMPC and CHAPSO and the erythrocyte membranes are discussed in the contexts of the molecular mechanism of the observed hemin-bilayer interactions. This allows the tentative conclusion that the main features of these interactions, the impermeability of the bilayer to hemin and the formation of hemin patches, might be characteristics of the erythrocyte membranes. Materials and Methods Reagents and Solutions. DMPC as lyophilized powder (Avanti Polar Lipids, Alabaster, AL) is mixed with hemin stock in 0.15 M phosphate buffer, pH ) 7.35. The solution is sonicated for 20 min and CHAPSO (Sigma, St. Louis, MO) is added. Homogenization of the final mixture is achieved by repeated vortex mixing, sonication, and heating/cooling. The molar ratio of DMPC to CHAPSO is Q(DMPC/CHAPSO) ) 3, the total lipid concentration Clp is 5% by weight, and the initial hemin concentration Ch in the aqueous phase is 0.7 mg/mL or 1.1 mM. The sample is mounted on a microscope slide prepared using Parafilm as spacer; the thickness of the solution layer in such a slide, measured by focusing the microscope on the bottom and top glass surfaces in contact with the solution, is 127 µm. Temperature control is achieved by circulating water above the slide; for this, the slide is attached to a brass frame with dimension 100 mm × 50 mm × 20 mm (length × width × height) with in and out ports. Through these ports, the brass frame is connected to a RTE-7 water bath (Neslab). The temperature is measured with an accuracy of 0.1 °C with an HH 506R thermocouple thermometer (Omega Engineering Inc.).

10.1021/jp100611n  2010 American Chemical Society Published on Web 03/17/2010

4530

J. Phys. Chem. B, Vol. 114, No. 13, 2010

Qutub et al.

Figure 2. Composition of lipids in erythrocyte membranes according to Bernhardt and Ellory.20

layer, a bandpass filter with transmission in the region from 420 to 460 nm is placed in the optical path. Hemin absorbance at 440 nm, from the data in Figure 1a, is shown at several concentrations in Figure 1b. From this dependence, the extinction coefficient of hemin at this wavelength is 24 mM-1 cm-1. Figure 1. (a) Spectra of hemin in phosphate buffer at four concentrations indicated in the plot. (b) Absorbance of hemin at 440 nm plotted as a function of hemin concentration for the determination of its extinction coefficient, indicated in the plot.

Optical Observations and Dynamic Light Scattering Characterization of Solutions. Vesicle formation and hemin behavior are observed and recorded with Leica DMR microscope. Water immersed objective lenses (20× and 63× APO L U V I) were used. Color images were taken with a Sony DKC5000 CCD camera. Black-and-white pictures were acquired with a 12-bit linear response Kodak Megaplus ES 1.0/1260 camera (1024 × 1024 pixel CCD array). Polarized light images were obtained with the same setup and recorded with the Sony camera. Dynamic light scattering (DLS) characterization was carried out on an ALV-5000/EPP static and dynamic light scattering device (ALV-Gmbh, Langen, Germany) with a 35 mW He-Ne laser operating at wavelength λ ) 632.8 nm (Uniphase). The solution samples were placed in a cylindrical cuvette with 8 mm internal diameter. The detector was positioned at 90°. DLS spectra were acquired at 6 min collection periods. Each run was repeated four times. The distribution of the sizes of the scatterers was obtained using Provencher’s Laplace inversion CONTIN algorithm for the correlation function.21 Quantification of Hemin. Since the intensity of images taken by the Kodak camera is proportional to the intensity of the light incident to the camera charge-coupled photoelement (CCD), these images were used to determine the distribution of the heme concentration in different parts of the slide. The absorption of light by heme and its derivatives in aqueous solutions has not been extensively studied: the existing measurements were carried out mostly in organic solvents or, if in aqueous solutions, used heme encapsulated in detergent micelles.22 The spectra of hemin in 0.15 M phosphate buffer at pH ) 7.35 saturated with O2 are shown in Figure 1a. These spectra are similar to those of met-Hb and indicate that, as expected, the heme iron is in Fe3+ form. These spectra reveal that the strongest absorbance of hemin is in the blue wavelength range. Hence, to quantify the hemin distribution in the solution

Results and Discussion Characteristics of the DMPC/CHAPSO Bilayers and Comparison with the Composition and Structure of Erythrocyte Membranes. The lipid composition of the erythrocyte membrane according to ref 20 is summarized in Figure 2. The two components, which comprise about 50% of the mass of the membrane, are phosphatidylcholine (PC) and cholesterol (CHOL). The molar ratio of PC to CHOL in erythrocyte membranes Q(PC/CHOL) ) 1.25.23 In our model phospholipid membrane, we substitute DMPC for PC and CHAPSO for CHOL. The molar ratio Q(DMPC/CHAPSO) ) 3, which is close to the ratio of total phospholipids to cholesterol in the erythrocyte membranes, Figure 2. With this Q, the formation of the phospholipid vesicles upon temperature increase is reliably reproducible even in the presence of hemin and the hemin interactions with the phospholipid bilayer can be monitored. Phosphatidylcholine contains a palmitoyl residue, which contains 16 carbon atoms, denoted as 16 C, at position 1 and oleoyl residue (18 C) at position 2. In DMPC, these are substituted with two residues of myristic acid (14 C), Figure 3. CHOL is an essential component of cellular membranes and affects many of their properties such as order, stability, phase transitions, thickness, fragility, microviscosity, lateral diffusion, permeability, and membrane proteins activity.24,25 It is amphiphilic: the small polar head is a 3β-hydroxy group, while the rest of the molecule, the steroid nucleus and the isooctyl tail, is nonpolar. CHOL orients in phospholipid bilayers with its hydrophobic plane parallel to the long axis of the phospholipids and the polar head toward the aqueous environment.24 In the liver CHOL is catabolized to bile acids by addition of hydroxyl groups to the steroid nucleus and conversion of the isooctyl moiety of the tail to isopentanoic acid moiety.26 CHAPSO is a bile salt analogue, in which, as in other bile acids, the hydrophobic steroid plane is functionalized by three hydroxyl groups, which orient on one side of the steroid nucleus and thus form a second, hydrophilic plane in a biplanar topology.26 In addition, an electrically neutral zwitterionic sulfobetaine moiety is attached to the tail, Figure 3.27 In phospholipid bilayers, CHAPSO has two positions, schematically illustrated in Figure 4a: some CHAPSO molecules are embedded on the surface,

Interactions of Hemin with Model Erythrocyte

J. Phys. Chem. B, Vol. 114, No. 13, 2010 4531

Figure 3. Structures of phosphatidylcholine (PC), dimyristoylphosphatidylcholine (DMPC), cholesterol (CHOL), and 3-(cholamidopropyl)(dimethylammonio)-2-hydroxy-1-propanesulfonate (CHAPSO).

Figure 4. Schematic of the positions of CHAPSO and hemin, highlighted, respectively, in dark and light gray, in the phospholipid bilayer. (a) The positions of CHAPSO in a bilayer are shown, after ref 31. (b) In bicelles, CHAPSO occupies a third position along the edges. (c) An island of hemin is shown in addition to CHAPSO in a vesicle bilayer.

with the biplanar assembly parallel to the bilayer plane and its polar plane facing to aqueous environment. In the other position, CHAPSO forms dimers, in which the polar planes face one another and are inserted between the hydrophobic tails of the phospholipids, while the zwitterion tail is in the solution.28 While CHAPSO is similar to the CHOL metabolic products, its structure and polarity are significantly different from those of CHOL. We employ it here since CHAPSO-containing

phospholipid bilayers offer two significant advantages to those containing CHOL: The formation of macroscopic phospholipid vesicles can be induced by a simple temperature increase from a fluid solution, in which hemin can be uniformly dissolved. Furthermore, as demonstrated below, these vesicles form by the assembly of bicelle disks and this excludes hemin from their inside. Since CHOL does not support the formation of bicelles, these studies would be impossible with CHOL-containing vesicles. Evolution of the System upon Temperature Increase. Figure 5 shows a series of images representing the time evolution of structures in a solution containing DMPC, CHAPSO, and hemin. Observations are made through the blue band-pass filter discussed above, so that the color intensity corresponds to the hemin concentration integrated along the slide thickness. Thus, the weaker intensity of the vesicle interior in all panels in Figure 5 indicates that hemin is excluded from these locations. Mixtures of a long chain phospholipid and a detergent have complex phase behavior dependent on temperature, total lipid concentration Clp, and the molar ratio of lipid to detergent Q. Observations of mixtures of DMPC and CHAPSO (without hemin) at Clp ) 5 wt % and Q(DMPC/CHAPSO) ) 3 reveal the following variations of macroscopic state with temperature: fluid below 20 °C, gel-like at room temperature, and fluid above 30 °C, with large vesicles visible at temperatures above 34 °C. To decide between multiwall and single wall vesicles at these temperatures, we monitored the solution in polarized light, in which multiwall vesicles form a characteristic Maltese cross pattern. Since this pattern was only observed at temperatures above 59 °C,29 we conclude that the vesicles formed at 34 °C are unilamellar (ULV).

4532

J. Phys. Chem. B, Vol. 114, No. 13, 2010

Figure 5. Vesicles in DMPC/CHAPSO solutions with total lipid concentration Clp ) 5% and DMPS tp to CHAPSO molar ratio Q ) 3 in the presence of 1.1 mM hemin at different temperatures. (a) 2 min after temperature increase from 34 to 35.2 °C: small vesicles (150 µm form; hemin is excluded from their interior. The arrow points at a patch of hemin embedded in the lipid bilayer.

Figure 6. Dynamic light scattering characterization of solutions of DMPC and CHAPSO at temperatures indicated in the plots. Total lipid concentration Clp ) 5%, ratio Q(DMPC/CHAPSO) ) 3. The correlation functions g2(τ) and the corresponding delay time distribution function G(τ) are shown.

Dynamic light scattering characterization of the fluid stable below 20 °C reveals the presence of a single scatterer with a characteristic delay time τ = 0.1 ms, Figure 6a. Applying the Einstein-Stokes relation, as in ref 30, we find that these objects have a hydrodynamic diameter of ∼23 nm. Comparing this with solid-state and solution NMR results on similar objects in binary lipid mixtures,31-33 we conclude that the scatterers are bicelles: disk-like micelles, which are magnetically orientable and exhibit liquid-crystal-like properties.28 The bicelles observed in ref 33 are ∼5 nm thick and 40-50 nm in diameter, which would yield the hydrodynamic diameter observed by DLS. In the bicelles, the long chain phospholipid forms the flat bilayer and the shortchain one lines the edge of the disk. In the system that we study, the CHAPSO molecules, in addition to their two positions in the bilayer, discussed above, line the bicelle edge and ensure the stability of the bicelles,28 Figure 4b. The same type of bicelle is present in the fluid existing above 30 °C, Figure 6b. We tentatively conclude that the gel observed in the temperature range 23-29 °C is a jammed bicelle

Qutub et al. suspension. The data in Figure 6b suggest that at temperatures above 30 °C the bicelles transform into larger objects, likely vesicles: the strong peak at τ = 100 ms corresponds to a hydrodynamic diameter of ∼20 µm, similar to those seen with optical microscopy in Figure 5a. The vesicle formation lowers the bicelle number density and leads to liquefaction of the gel. The evolution of the viscosity of the solution in the presence of hemin is very similar to that in its absence, provided that the same Q and Clp are employed. Unfortunately, the high concentration of hemin, combined with its high absorbance at all visible wavelengths, Figure 1a, prevents light scattering characterization of these solutions; for light scattering characterization of solutions containing 50 µM of hemin, see ref 34. Below 34 °C no vesicles are detected with optical microscopy and the solution appears homogeneous. In analogy to the observations around 30 °C in the absence of hemin, we conclude that the solution fluidity and apparent homogeneity is due to the low number and small size of the vesicles. Large vesicles, seen in Figure 5a, appear upon temperature increase from 34 to 35.2 °C. Again, since the Maltese cross pattern was not observed by polarized light microscopy, we conclude that the vesicles formed at 34 °C are unilamellar (ULV). These ULV’s are of initial diameter ∼15 µm and within a few seconds grow to ∼50 µm. Judging by the color intensity, the solution trapped inside them contains negligible amounts of hemin. If the temperature is raised to 36 °C, vesicles grow larger and fuse with other vesicles, Figure 5b. Further vesicle growth is observed in Figure 5c upon a temperature increase to 36.4 °C. Three hours after a temperature increase to 37.2 °C we see giant unilamellar vesicles (GUVs) whose diameter reaches well above 150 µm. They still exclude hemin from the interior but dark patches of hemin appear, which likely indicate hemin associated to the phospholipid bilayer of the vesicles. These patches have uniform intensity, Figure 5d, reproducible in multiple observations. This uniform intensity suggests that the patch thickness and the density of hemin in them are consistent in all observed patches. Below, we discuss two aspects of these observations: (i) the mechanism of vesicle formation and growth and (ii) the patches and what they suggest about the interactions of hemin with the membrane. Mechanism of Vesicle Formation and Growth. The above observations suggest a mechanism for the formation and growth of vesicles the DMPC/CHAPSO/hemin solution, schematically represented in Figure 7. Formation of Vesicles by Fusion of Bicelles. At 30 °C or below the system is fluid and contains bicelles consisting of DMPC and CHAPSO suspended in an aqueous solution of lipid molecules; equilibrium exists between the lipids in the solution and those in the bicelles, Figure 7a. At temperatures above 30 °C, vesicles form by fusion of bicelles, Figure 7b, and are visible in an optical microscope, Figure 5a, at temperatures above 34 °C. The mechanism of bicelle destabilization likely involves an increase of the number of CHAPSO dimers embedded in the phosphoric bilayer (see discussion of CHAPSO positions above); this destabilization is stronger at values of the ratio Q > 1.35 The destabilized bicelles respond readily to temperature increases by fusing into vesicles. Growth by Addition of Single Molecules and by Vesicle Fusion. The formed vesicles grow by addition of single lipid molecules from the aqueous phase. The amount of molecules that can be added by this mechanism is limited by the equilibrium with dissolving bicelles, Figure 7c. Likely, this is the only mechanism of growth while the vesicles are small and

Interactions of Hemin with Model Erythrocyte

J. Phys. Chem. B, Vol. 114, No. 13, 2010 4533

Figure 7. Schematic of the mechanism of vesicle formation and growth in the studied mixture of DMPC and CHAPSO (DMPC/CHAPSO ratio Q ) 3, total lipid concentration Clp ) 5% w/v) at different temperatures, as indicated in the panels. Small solid circles: individual DMPC and CHAPSO molecules. Ellipses: bicelle disks. Open circles: vesicles. (a) The system consists of bicelles in equilibrium with single lipid molecules. (b) Unilamellar vesicles (ULV) form by fusion of several bicellar discs. (c) ULVs grow by addition of single lipid molecules. (d) ULVs fuse and giant unilamellar vesicles (GUV) form.

away from one another. Another mechanism is growth, directly observed in Figure 5b, by vesicle fusion. Vesicle fusion has been observed in systems of long-chain phospholipids and shortchain lecithins, where small ULVs undergo temperature induced fusion due to a phase transition between a gel-like and a liquidcrystalline state of the alkyl chains of the long-chain phospholipid.36 Temperature induced fusion in our system, starting at 36 °C, may be due to the gel to liquid transition of DMPC embedded in the bilayer. This temperature is close to value of 37 °C reported for vesicle fusion in mixtures of dipalmitoylPC with short-chain lecithin.36 The CHAPSO oligomers, schematically illustrated in Figure 4 likely disfavor the high curvature of the small vesicles and thus also favor fusion. One mechanism of vesicle growth, which may be excluded by the observations in Figure 5, is by the incorporation of bicelles. It is likely that such association would lead to temporary disruption of the vesicle membranes and leakage of hemin inside the vesicle. On the other hand, Figure 5 shows that hemin in consistently excluded from the vesicle inside at all vesicle sizes. The intensity of the blue color inside the vesicles does not increase upon storage for days. This shows that the phospholipid membrane is impermeable to hemin and it is constrained on one side of the membrane. Hemin Patches and the Mechanism of Interaction of Hemin with the Membrane. The geometry of the slide in which we monitor the vesicles growing in the presence of hemin is schematically depicted in Figure 8b. The slide height along the optical path is 127 µm. GUVs appear circular in the plane of observation and their diameter in this plane surpasses the slide thickness. Since GUV’s favor spherical shape, this deviation from sphericity indicates that the GUV’s touch both upper and lower slide glasses and the light transmitted through the GUV’s center is only affected by the composition of the solution inside the vesicle. The black-and-white camera used has a 12-bit digital video output coding for 4096 levels of gray for each pixel. The linear response makes the intensity value of a pixel proportional to the intensity of transmitted light at that slide area. Custom-made software scans the images and plots pixel values as a function of position. The transmitted intensity profile along the line in Figure 8a is shown in Figure 8c. This line was chosen so that it would pass through the heme patch.

Figure 8. Quantification of hemin embedded within lipid bilayer. (a) Picture of a giant unilamellar (GUV) vesicle taken with linear response camera through a l ) 440 nm filter, at which hemin has an absorption maximum. The intensity of the image is proportional to the integral of hemin concentration along the axis perpendicular to the image plane. The arrow indicates a patch of embedded hemin of size ∼20 µm. (b) Schematic side view of GUV in slide. The heme-containing solution is shown in pink, the lipid bilayer in yellow, and the embedded heme in red. Optical pathways of light passing through different areas of the slide are indicated with black arrows. The respective light intensities are denoted as Ib, the base intensity transmitted though the hemecontaining solution, Imax, the maximal intensity transmitted through the vesicle at the point of contact of the bilayer with top and bottom glasses (this intensity is not affected by absorbance by the heme), and Ihemin, Imax lowered by absorbance of the heme embedded in the lipid bilayer. (c) Profile of transmitted intensity at wavelength l ) 440 nm along the dotted line in (a); values of Ib, Ihemin, and Imax are indicated.

In the intensity profile in Figure 8c Ib is the base intensity transmitted through regions with no vesicles and occupied entirely by the aqueous phase. Because of the high absorbance at 440 nm of hemin present in the aqueous phase, this intensity is the lowest. Imax is the intensity transmitted through regions of no hemin on the light path, where the vesicle bilayer touches the glass and the hemin-containing aqueous phase is completely excluded. To verify that this is the maximum transmitted intensity, we shifted up and down the line in Figure 8a and found that Imax stays within 50 intensity units, its natural variance in Figure 8b. Ihemin is intensity transmitted through the hemin patch. We assume that Imax equals the incident intensity corrected for absorbance by the optical elements of the microscope and the slide glasses. Since the top and bottom phospholipid bilayers

4534

J. Phys. Chem. B, Vol. 114, No. 13, 2010

of the observed vesicle are each only ∼5 nm thick, we assume that they do not attenuate Imax. Then, according to the Bouge-Lambert-Beer law log(Imax/Ihemin) ) εhemin, bilayerCheminh, where h ) 5 nm is the thickness of the phospholipid bilayer. To evaluate the relevant εhemin, we note that the hemin absorbance when interacting with phospholipid bilayers is higher than in aqueous solutions.14 The difference in molar extinction coefficient between hemin in lipid bilayers and in solution ∆ε was measured in liposomes composed of equimolar amounts of PC and CHOL. The spectra in ref 14 reveal that ∆ε440 ) 10.5 mM-1 cm-1. We assume that the same ∆ε applies to hemin embedded in phospholipid bilayers we study, and with the molar extinction coefficient of hemin in phosphate buffer at 440 nm determined in Figure 1b εhemin,soln ) 24 mM-1 cm-1, we get εhemin,bilayer ) 34.5 mM-1 cm-1. From Figure 8b, the average pixel values for Ihemin and Imax are 2050 and 2300 arbitrary units, respectively. These two numbers yield that C hemin in the patch is 1.45 × 10-3 µmol/cm2 of the patch area, or 8 or 9 hemin molecules/nm2 of the patch. Considering that hemin is a planar molecule 0.2 nm thick and 1.2 nm long, we conclude that the patches are composed almost exclusively of hemin with orientation as described in ref 13 and illustrated in Figure 4c, with carboxylate residues pointing toward the aqueous environment in both the inner and outer monolayers. Thus, the patches are akin to a two-dimensional hemin crystal, consisting of two molecular layers related by mirror symmetry. Since the patches occupy such a small fraction of the total area of the vesicle membrane, it is likely that they grow by the attachment of single molecules embedded in the bilayer and not from those in the solution around the vesicle. The obvious driving force for the hemin insertion into the bilayer and assembly into patches is its hydrophobicity. An investigation of the hemin transport on liposomes of various compositions revealed that hemin is initially incorporated in the outer monolayer and moves to the inner monolayer by a flipping process analogous to that of phospholipids.13 Chlorophyll, a molecule with a structure similar to hemin, has a flip half-time of 4 min in phospholipid bilayers.37 The low density of patches embedded in the membranes of the GUV’s in our experiments suggests that the formation of a patch is a rare event that may be triggered by a special perturbation of the phospholipid bilayer. The concentration of the single hemin molecules embedded in the bilayer must be greater than the solubility of the patches; otherwise the patches would dissolve. However, this concentration must be lower than the critical concentration needed for the nucleation of new patches. This later limitation suggests that the hemin concentration in the membrane is determined by the equilibrium with the hemin in the solution outside the vesicles. Despite the presence of a relatively high hemin concentration in the membrane, hemin is not released inside the vesicles, where its concentration is near zero for several days. Besides hemin’s hydrophobicity, another factor in the retention of hemin may be the presence of CHAPSO. CHOL retards hemin efflux out of liposomes by modulating membrane order and increasing the lipid packing density rather than by direct interaction.13 CHAPSO is likely to affect hemin transport similarly to CHOL. Relevance of Observed Heme-Bilayer Interactions to Erythrocyte Membranes. How does the permeability for hemin of the model membrane studied here compare to that of live cellular membranes? The permeability of phospholipid bilayers is described by two models: a solubility-diffusion theory, which is applicable to neutral solutes, and a transient pore mechanism, which accounts for movements of ionic solutes through hydrated

Qutub et al. membrane defects induced by thermal fluctuations.38 Because of its large hydrophobic moiety, hemin permeability likely follows the solubility-diffusion model. The permeability of phospholipid bilayers for water, ions, and small organic molecules depends on lipid bilayer thickness, packing density and fluidity. In turn, these are functions of the lipid chain length and bond types, temperature, and CHOL concentration.38,39 Since DMPC has fewer carbon atoms in its aliphatic chains than PC, the model bilayer is thinner and more permeable for hemin. On the other hand, the absence of double bonds in DMPC would increase the lipid packing density with respect to the erythrocyte membrane and thus would lower its permeability. The addition of CHOL to PC bilayers affects phospholipid chain mobility: the rigid character of the sterol nucleus increases order by imposing steric constraints on the neighboring phospholipid tails.24 The decreased fluidity reduces the permeability for hemin.13 The sterol nucleus of CHAPSO will restrict the motions of phospholipid tails in a similar manner. However, due to its lower hydrophobicity CHAPSO is present at a lower molar fraction in the model membrane than CHOL is present in erythrocyte membranes, so that the model membrane would again be more permeable. The core body temperature is regulated within a narrow margin (36.67-37.78 °C). The permeability of hemin was studied above within a temperature range of 3 °C, and only the upper limit of this range, 37.2 °C, is comparable to physiological. Upon a temperature increase, the phospholipid bilayers undergo a phase transition from the crystalline to gel state of the alkyl chains, whereby defects form, the surface area per lipid molecule increases, and the packing density decreases. The transition leads to thinning of the bilayer and increasing structural disorder, thus favoring intercalation of hemin. CHAPSO, similarly to CHOL, will possibly counteract effects of temperature increase to some extent and decrease the permeability at a given temperature. Thus, considering the three factors for membrane permeability (the nature of the lipid molecules, the molar ratio between them, and temperature), the model membrane is unlikely to have a lower permeability than the erythrocyte membrane. The found containment of the hemin on the one side of the model membrane is likely applicable to the erythrocyte membranes. In that case, the only way that hemin may exit the erythrocytes into the blood plasma is by its strong binding to albumin in contained in the blood plasma, or upon lysis of the red blood cells. The factors that influence membrane permeability determine membrane fluidity,40 which, in turn, is the main factor for the lateral mobility of molecules embedded in the membrane. Thus, the mechanism of patch formation, by the lateral motion and assembly of hemin molecules embedded in the membrane, is relevant to erythrocyte membranes and hemin patches might be expected in the latter. Conclusions We show that a solution containing DMPC and CHAPSO forms giant unilamellar vesicles at temperatures near the physiological. These vesicles form by the fusion of bicelles, disk-like micelles existing in solution of CHAPSO and DMPC, and grow by the attachment of monomers from the solution or, at sizes greater than 100 µm, by fusion of several vesicles. While some solution may be trapped during the formation of the first small vesicles from bicelles, the late-stage growth proceeds in a manner that excludes larger solutes, such as hemin, from the interior of the vesicles. In this way, a suspension of large vesicles

Interactions of Hemin with Model Erythrocyte is created that allows investigation of the interactions of the phospholipid bilayer to various solutes constrained on the one side of the vesicle membrane. We found that hemin is constrained on the outside of the studied vesicles. We found that hemin forms patches embedded in the phospholipid bilayer. The patches are akin to a twodimensional crystal consisting of two molecular layers related by mirror symmetry. Consideration of the properties of the membrane suggests that erythrocyte membranes would likely have even lower permeability than the studied model membranes and that hemin patches are also likely in erythrocyte membranes. Acknowledgment. This work was supported by the National Science Foundation (Grant MCB 0843726) and The Welch Foundation (Grant E-1641). References and Notes (1) Chui, D. T.; et al. Free Radical Biol. Med. 1996, 21, 89. (2) Kirschner-Zilber, I.; Rabizadeh, E.; Shaklai, N. Biochim. Biophys. Acta 1982, 690, 20. (3) Chou, A. C.; Fitch, C. D. J. Clin. InVest. 1981, 68, 672. (4) Chou, A. C.; Fitch, C. D. J. Clin. InVest. 1980, 66, 856. (5) Orjih, A. V.; Banyal, H. S.; Chevli, R.; Fitch, C. D. Science 1981, 214, 667. (6) Fitch, C. D.; Chevli, R.; Banyal, H. S.; Philips, G.; Pfaller, M. A.; Krogstad, D. J. Antimicrob. Agents Chemother. 1982, 819. (7) Vanderveldt, G. M.; Regan, R. F. Free Radical Res. 2004, 38, 431. (8) Goldstein, L.; Teng, Z.; Zeserson, E.; Patel, M.; Regan, R. F. J. Neurosci. Res. 2003, 73, 113. (9) Janney, S. K.; et al. Blood 1986, 67, 331. (10) Liu, S. C.; Zhai, S.; Palek, J. Blood 1988, 71, 1755. (11) Miller, Y. I.; Shaklai, N. Biochim. Biophys. Acta 1999, 1454, 153. (12) Miller, Y. I.; Felikman, Y.; Shaklai, N. Biochim. Biophys. Acta 1995, 1272, 119. (13) Cannon, J. B.; Kuo, F.; Pasternack, R. F.; Wong, N. M.; MullerEberhard, U. Biochemistry 1984, 23, 3715. (14) Tipping, E.; Ketterer, B.; Christodoulides, L. Biochem. J. 1979, 180, 327.

J. Phys. Chem. B, Vol. 114, No. 13, 2010 4535 (15) Ginsburg, H.; Demel, R. A. Biochim. Biophys. Acta 1983, 732, 316. (16) Solar, I.; Shaklai, N. Biochim. Biophys. Acta 1989, 983, 199. (17) Schmitt, T. H.; Frezzatti, W. A., Jr.; Schreier, S. Arch. Biochem. Biophys. 1993, 307, 96. (18) Liu, S. C.; et al. J. Biol. Chem. 1985, 260, 12234. (19) Kumar, S.; Bandyopadhyay, U. Toxicol. Lett. 2005, 157, 175. (20) Red Cell Membrane Transport in Health and Disease, 1st ed.; Springer: Berlin, 2003. (21) Provencher, S. W. Comput. Phys. Commun. 1982, 27, 229. (22) Falk, J. E. Porphyrins and Metalloporphyrins; Elsevier Publishing Co.: Amsterdam, 1964. (23) Broekhuyse, R. M. Clin. Chim. Acta 1974, 51, 341. (24) Crockett, E. L. Aerical Zoology 1998, 38, 291. (25) Hui, S. W.; Steward, C. M.; Carpenter, M. P.; Steward, T. P. J.Cell Biol. 1980, 85, 283. (26) Hofmann, A. F. News Physiol. Sci. 1999, 14, 24. (27) Sanders, C. R., II; Prestegard, J. H. Biophys. J. 1990, 58, 447. (28) Sanders, C. R.; Scott Prosser, R. Structure 1998, 6, 1227. (29) Qutub, Y. Thesis, Univerisity of Houston, 2006. (30) Pan, W.; Galkin, O.; Filobelo, L.; Nagel, R. L.; Vekilov, P. G. Biophys. J. 2007, 92, 267. (31) Sanders, C. R.; Prosser, R. S. Structure 1998, 6, 1227. (32) Vold, R. R.; Prosser, R. S.; Deese, A. J. J. Biomol. NMR 1997, 9, 329. (33) Harroun, T. A.; Koslowsky, M.; Nieh, M. P.; de Lannoy, C. F.; Raghunathan, V. A.; Katsaras, J. Langmuir 2005, 21, 5356. (34) Pan, W.; Uzunova, V. V.; Vekilov, P. G. Biopolymers 2009, 91, 1108. (35) Kubota, K.; Sato, T.; Kogure, H.; Wakamatsu, K. PF ActiVity Rep. 2002, 19, 147. (36) Eum, K. M.; Riedy, G.; Langley, K. H.; Roberts, M. F. Biochemistry 1989, 28, 8206. (37) Birrell, G. B.; et al. Biochim. Biophys. Acta 1980, 603, 213. (38) Paula, S.; Volkov, A. G.; Van Hoek, A. N.; Haines, T. H.; Deamer, D. W. Biophys. J. 1996, 70, 339. (39) Huster, D.; Jin, A. J.; Arnold, K.; Gawrisch, K. Biophys. J. 1997, 73, 855. (40) Rubenstein, J. L. R.; Smith, B.; McConnell, H. M. Proc. Natl. Acad. Sci. U.S.A. 1979, 76, 15.

JP100611N